AN ABSTRACT OF THE THESIS OF

Aaron J. Hartz for the degree of Master of Science in Oceanography presented on May 26, 2010.

Title: Investigating the Ecological Role of in Free‐Living Marine Heterotrophic .

Abstract approved:

Barry F. Sherr

Phagotrophic protists are major consumers of microbial in aquatic ecosystems. However, biochemical mechanisms underlying prey recognition and by protists are not well understood. We investigated the potential roles of cell signaling mechanisms in chemosensory response to prey, and in capture of prey cells, by a marine (Uronema sp.) and a heterotrophic ( marina). Inhibition of kinase signal transduction caused a decrease in both chemosensory response and . Inhibition of G protein coupled receptor signaling pathways significantly decreased chemosensory response but had no effect on prey ingestion. Inhibitor compounds did not appear to affect general cell health, but had a targeted effect. In addition, upon finding rhodopsin in protein extracts from the plasma membrane of O. marina, an

investigation into the photosensory behavior of O. marina was made. This organism has a positive phototactic response to moderate levels of white light and a wavelength‐specific phototactic response. O. marina response to light was affected by culture conditions and feeding state. In the presence of blue light, emit red fluorescence; results indicate that in the presence of blue light, O. marina exhibited a positive phototactic response to a simulated “patch” of phytoplankton suggesting the use of the photoreceptor to detect the red phytoplankton fluorescence. These results support the idea that cell signaling pathways known in other eukaryotic organisms are involved in feeding behavior of free‐living protists. As this field of study makes advances and more information relating to biochemical signaling in free‐living protists becomes available, it will give researchers new tools and approaches in the study of feeding behavior in .

© Copyright by Aaron J. Hartz

May 26, 2010

All Rights Reserved

Investigating the Ecological Role of Cell Signaling in Free‐Living Marine Heterotrophic Protists.

by

Aaron J. Hartz

A THESIS

submitted to

Oregon State University

in partial fulfillment of the requirements for the degree of

Master of Science

Presented May 26, 2010

Commencement June 2011

Master of Science thesis of Aaron J. Hartz presented on May 26, 2010.

APPROVED:

Major Professor, representing Oceanography

Dean of the College of Oceanic and Atmospheric Sciences

Dean of the Graduate School

I understand that my thesis will become part of the permanent collection of Oregon State University libraries. My signature below authorizes release of my thesis to any reader upon request.

Aaron J. Hartz, Author

ACKNOWLEDGEMENTS

The author expresses appreciation to the following people:

My major professor and committee, Barry Sherr, Evelyn Sherr, Rick Colwell, Theresa Filtz, and Mario Magaña. I would also like to thank Ricardo Letellier for the assistance he provided. My office mates Wiley Evans, Brandon Briggs, Maria Kavanaugh, and Brie Lindsay. Robert Allan, Lori Hartline, and Robert Duncan from the student programs office. Additional friends and family, Norman Hartz, Karen Hartz, Jeremy Hartz, Johnny Anderson, Amy Simmons, Chris Holm, Paul Walczak, Jon Ellenger, Faron Anslow, John Stevenson, and Nate Meehan.

CONTRIBUTION OF AUTHORS

Dr. Barry Sherr and Dr. Evelyn Sherr were involved in the design and writing of manuscripts in Chapters 2 and 3.

TABLE OF CONTENTS

Page 1 General Introduction…………………………………………………………………… 1 1.1 Overview…………………………………………………………………………………. 1 1.2 Receptor activation/signal transduction…………………………………….. 2 1.3 Behavioral response of phagotrophic protists to 4 environmental stimuli……………………………………………………………………… 1.3.1 Chemosensory behavior………………………………………………………….. 5 1.3.2 Photosensory behavior…………………………………………………………… 7 1.4 Organisms used in this study……………………………………………………… 9 1.5 Goals of this study……………………………………………………………………… 9 2 Using Inhibitors to Investigate the Involvement of Cell Signaling in 11 Predation by Marine Phagotrophic Protists……………………………………… 2.1 Abstract…………………………………………………………………………………… 12 2.2 Introduction……………………………………………………………………………… 13 2.3 Materials and methods……………………………………………………………… 14 2.4 Results and discussion……………………………………………………………… 17 2.5 Acknowledgements……………………………………………………………………. 24 2.6 References………………………………………………………………………………… 24 3 Photoresponse in the colorless marine dinoflagellate Oxyrrhis 27 marina 3.1 Abstract…………………………………………………………………………………… 29 3.2 Introduction……………………………………………………………………………… 31 3.3 Materials and Methods……………………………………………………………… 32 3.3.1 Cultures………………………………………………………………………………… 32 3.3.2 Flow cytometric analysis………………………………………………………… 32 3.3.3 Extraction and characterization of O. marina membrane 33 3.3.4 Phototaxis experiments ………………………………………………………… 35 3.3.5 Evaluation of cAMP induction by light …………………………………….. 37 3.3.6 Inhibition of Rhodopsin ………………………………………………………… 38

TABLE OF CONTENTS (Continued) Page 3.4 Results ……………………………………………………………………………………… 39 3.4.1 Characterization of O. marina membrane‐associated proteins ….. 39 3.4.2 O. marina response to light ……………………………………………………… 39 3.4.3 O. marina response to Chl‐a fluorescence ………………………………… 40 3.4.4 Light stimulation of intracellular downstream signaling …………… 40 3.4.5 Inhibition of rhodopsin function by hydroxylamine ………………….. 41 3.5 Discussion …………………………………………………………………………………. 41 3.6 Conclusions ……………………………………………………………………………….. 45 3.7 References ………………………………………………………………………………… 47 4 General Conclusions …………………………………………………………………… 59 4.1 Overview ………………………………………………………………………………… 59 4.2 Chemosensory/inhibitor study ………………………………………………… 59 4.3 Photosensory behavior study …………………………………………………… 61 4.4 What does this mean for the field of aquatic microbial ? … 65 4.5 Future studies ………………………………………………………………………… 67 4.6 Closing remarks ……………………………………………………………………… 69 5 Bibliography ……………………………………………………………………………… 71

LIST OF FIGURES

Figure Page 1 Flow cytometry plot ………………………………………………………………… 52 2 Configurations for phototaxis experimental setup …………………… 53 3 Maximum likelihood tree of rhodopsin proteins ………………………. 54 4 O. marina phototaxis to white light …………………………………………… 55 5 O. marina phototaxis to specific wavelengths …………………………… 56 6 O. marina photoresponse to phytoplankton auto‐fluorescence … 57 7 Intracellular cAMP …………………………………………………………………… 58

1 Introduction

1.1 Overview

Phagotrophic protists are important components of the . They are significant grazers of and phytoplankton (Jurgens and Matz 2002; Quevedo and Anadon 2001). Predation by bacterivorous protists can be sufficient to control bacterial biomass (Gasol et al. 2002) and to play a role in shaping the genotypic and phenotypic composition of planktonic bacteria (Jurgens and Matz 2002). Herbivorous protists, including and heterotrophic , also consume a significant fraction of phytoplankton standing stocks and biomass production in the ocean (Strom et al. 2001, Calbet & Landry 2004, Calbet 2008). This has implications for the marine ecosystem since bacteria and phytoplankton influence the major biogeochemical cycles in the ocean (Azam 1998, Falkowski and Raven 2007). In addition, heterotrophic protists excrete nitrogen and phosphorus compounds, as well as trace elements such as iron, that can be utilized by primary producers (Caron and Goldman 1990, Barbeau et al. 1996) and contribute to the flow of energy to higher trophic levels by serving as a food source for metazooplankton (Sherr and Sherr 2007; Stoecker and Capuzzo 1990; Calbert and Saiz

2005). Most heterotrophic protists ingest prey via phagocytosis, an ancient mode of nutrition, most likely predating eukaryotic (Dyer & Obar 1994).

Phagocytosis is widespread among the major phylogenetic branches of protistans (Sherr and Sherr 2002). Currently the study of heterotrophic protists in aquatic food webs is a frontier in marine research (Wolfe 2000, Boenigk and Arndt 2002,

Sherr and Sherr 2002, Weiss 2002, Caron et al. 2009). Questions of current interest in marine microbial ecology are: What determines prey location and selection by 2 phagotrophic protists? How do environmental stimuli affect swimming and feeding behavior?

Heterotrophic protists respond to surrounding stimuli and motile cells can change their position to find optimal conditions. Protistan response to stimuli can be complex and may be affected by the physiological state of the cells (Fenchel 2002). In addition, physiological and behavioral responses to signal molecules via their binding to transmembrane receptor molecules on the cell surface is a ubiquitious characteristic of microbes (Hunter 2000, Gomperts et al. 2004, Rosales 2005). These mechanisms consist of ligand-receptor molecule binding on the cell surface, which initiates an intracellular signal transduction cascade resulting in a change in behavior as such as engulfment and digestion of a prey cell, or a change in swimming velocity and direction in response to a signal (i.e. chemical cues).

Ligand-receptor binding and intracellular signal transduction mechanisms have been studied extensively in metazoan cells and in a small number of protist

(Greenberg 1995, Hunter 2000, Dodd and Drickamer 2001, Alberts et al. 2002, Renaud et al. 2004, Rosales 2005). Although the biochemical basis of the complex feeding behavior of marine protists has not yet been fully explored, there is good background evidence that the mechanisms described below are involved in prey ingestion by phagotrophic protists.

1.2 Receptor activation/signal transduction

Various types of receptor molecules are associated with cell membranes. The largest family of cell-surface receptors is that of the G protein-coupled receptors (GPCRs;

3 a class of seven-transmembrane receptors) (Alberts et al. 2002). G-protein coupled receptors are a ubiquitous part of the cellular machinery across a range of , including protists (Van Haastert and Devreotes 2004) and have the ability to transduce a variety of signals into the cell including light, amino acids, peptides, sugars, and large proteins. GPCR’s have been shown to be part of the chemosensory machinery used by immune cells in mammalian systems (Devreotes and Janetopoulos 2003; Haribabu et al.

1999) and in the protist Dictyostelium discoideum (Manahan et al. 2004; Traynor et al.

2000).

These trans-membrane receptors are activated when stimulated by the binding of specific signal compounds. This induces a conformational change in the GPCR molecule that sets off guanine-triphosphate (GTP)-activated phosphorylation pathways in the cell, leading to changes in cell and behavior. GPCRs are the basic biochemical machinery for chemosensory and other stimulus-linked responses in eukaryotes (Fawzi et al. 2001, Alberts et al. 2002). There is evidence that the fundamental elements for GPCR- mediated signal transduction are present in phagotrophic protists (Renaud et al. 2004).

Another important class of receptor molecules are lectins, which bind to sugar moieties of larger biomolecules on the surfaces of prey cells. Such binding can initiate prey ingestion in phagotrophic cells. There is a large body of work on biochemical control of phagocytosis in laboratory cultured protists, such as Tetrahymena and

Paramecium (Almagor et al. 1981; Hellung-Larsen et al. 1986; Renaud et al. 1995, 2004;

Leick et al. 1996, 1997; Bell et al. 1998), euglenoid in the genera and

Astacia (Levandowsky 2001), parasitic protists such as trypanosomes and Leishmania

(Mosser &, Rosenthal 1993, Ratcliff et al. 1996, Basseri et al. 2002), and in metazoan

4 white blood cells (Greenberg 1995, Ballarin et al. 2002). These studies include demonstrations of the effect on cell behavior of ligand binding by membrane-bound receptor molecules, and of signal transduction mechanisms within the cell that are switched on by ligand-receptor binding (Greenberg 1995, Alberts et al. 2002, Renaud et al. 2004).

As mentioned above, ligand-receptor binding at the cell surface activates intracellular signal transduction, in which phosphorylation of specific proteins leads to molecular processes in the cell that can result in changes in cell behavior, e.g. change in or phagocytosis of a prey cell. The signal transduction activator molecules are protein kinases (Stoka 1999), which facilitate phosphorylation (adding phosphate molecules) of compounds in cells. Phosphorylation of specific proteins is known to be a widely used molecular mechanism involved in switch-like outputs in cell behavior

(Rasmussen et al. 1996, Hunter 2000, Alberts et al. 2002, Gomperts et al. 2004). Tyrosine kinase proteins in particular are important in phagocytosis in both and protist cells

(King et al. 2003, Renaud et al. 2004, Rosales 2005). Components of tyrosine kinase signaling pathways appear highly conserved from to vertebrates, and have been found in two (King and Carroll 2001, King et al. 2003).

1.3 Behavioral response of phagotrophic protists to environmental stimuli

Examples of stimuli that can result in a change in protist behavior include: detection and orientation with respect to chemical stimuli (chemotaxis, Fenchel and

Blackburn 1999); magnetic fields (magnetotaxis, Bazylinski et al 2000); gravity

(geotaxis, Fenchel and Finlay 1984; Noever, Cronise, and Matsos 1994), and light

5

(phototaxis, Song 1983). To date, the study of chemotaxis has probably received most attention among the sensory systems in phagotrophic protists.

Protistan response to potential prey can be complex, resulting in selective ingestion or avoidance of some prey items (Verity 1991, Boenigk and Arndt 2000, Weiss

2002). Previous studies suggest several factors that may be involved in response to potential prey, including prey size (Fenchel 1987, Gonzalez et al. 1990, Simek and

Chrzanowski 1992, Strom 2000), prey motility (Monger and Landry 1992, Gonzalez et al. 1993, Matz and Jurgens 2002) and growth state (Gonzalez et al. 1993, Flynn et al.

1996), prey cell surface hydrophobicity (Monger et al. 1999, Matz and Jurgens 2001,

Matz et al. 2002), and chemical composition which affects ‘taste’ of prey cells (Verity

1991, Wolfe 2000). A combination of these factors is likely involved. Prey selection or rejection is also likely to involve cell signaling events which are initiated by binding of specific ligand compounds to transmembrane receptor molecules (Hartz et al. 2008,

Wooten et al. 2007).

1.3.1 Chemosensory behavior

Several studies have shown heterotrophic protists to utilize chemosensory mechanisms to move towards an attractant as a proxy for prey cells (e.g. Fenchel and

Blackburn 1999; Fenchel and Jonsson 1988; Snyder 1991; Manahan et al. 2004).

However, these studies do not provide details on what type of receptors are involved or the resulting intracellular signaling pathways that lead to a change in motility (speed and/or direction) or initiation of prey ingestion (phagocytosis).

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A variety of chemicals have been shown to affect the movement and, in some cases, the rate, of phagotrophic protists, both positively and negatively (Wolfe

2000). For example, chemicals that have higher concentrations in patches of decomposing organic matter such as aggregates or decaying fecal pellets could signal a bacterial food patch (Sibbald et al. 1987, Fenchel and Blackburn 1999).

Previously demonstrated chemo-attractants for protists include , amino acids, fatty acids, peptides, and bacterial and phytoplankton prey (Hellung-Larsen et al. 1986;

Sibbald et al. 1987; Bennett et al. 1988; Van Houten and Preston 1988; Verity 1988,

1991; Fenchel and Blackburn 1999, Martel 2006). Repellant molecules can also cause changes in protistan behavior. Several studies show that compounds associated with phytoplankton sulfur chemistry, DMS and DMSP, act as sublethal chemical defenses against protist grazers and can decrease prey ingestion rates (Wolfe and Steinke 1996,

Wolfe et al. 1997, Strom et al. 2003 a&b).

A study by Wootton et al. (2007) has shown that a cell surface mannose-binding lectin is directly involved in prey recognition by the heterotrophic dinoflagellate Oxyrrhis marina. Furthermore, Strom et al. (2007) suggested that dissolved free amino acids (L- enantiomers) serve as signal molecules that can differentially affect feeding responses in phagotrophic protists. This finding is quite possibly related to GPCR signaling; GPCR’s are known to be effective receptors for a variety of peptides (Cooper and Hausman 2007) and L-amino acids (Conigrave et al 2000).

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1.3.2 Photosensory behavior

Light is an important physical property of aquatic ecosystems, and has diverse effects on microbes. Light is of fundamental importance to photosynthetic microbes.

Heterotrophic microbes have also been shown to benefit from light. It is now well known that many have , some of which may act as a light-activated proton pump, a process termed phototrophy (Beja et al. 2001, Furhman et al. 2008).

Rhodopsin-based proton pumps are not limited to prokaryotes; they have also been found in the marine macro-alga acetabulum (Tsunoda et al. 2006) and in the Leptosphaeria maculans (Waschuk et al. 2005). In addition, sensory rhodopsins are found in many species in Domains , Bacteria, and Eukarya (Sharma et al.

2006, Spudich, 2006) and are involved in photosensory behavior such as phototaxis.

Photosensory behavior in is relatively well studied (e.g. review of Hegemann

2008), but is less well known for heterotrophic protists.

Phototaxis has been investigated in only a handful of heterotrophic protist species, and even less attention has been given to the ecological advantages phototaxis might entail for a non-photosynthetic protist. Responses to light by heterotrophic protists are diverse and can vary even among species of the same genus. Some examples of response to light include the following observations: proteus locomotion can be oriented in the direction of a light gradient (Mast 1910) and the rate of movement of the amoeba is dependent on light intensity (Mast and Stahler 1937). The ciliate

Chlamydodon mnemosyne exhibits positive phototaxis during starvation and negative phototaxis when well fed (Selbach, Hader, and Kuhlmann 1999). The ciliate Stentor coeruleus moves away from light while Stentor niger moves toward light and Stentor

8 polymorphus exhibits no response to light. Another ciliate japonicum is positively phototactic to blue light and photophobic to red light (Song 1983). Zoospores of Ulkenia sp., which consume bacteria, exhibit positive phototaxis to produced by the bacteria fischeri (Amon and French 2004).

In order for phototaxis to occur, a photoreceptor must receive the initial light stimuli, leading to an intracellular signal transduction cascade, and ending in the photoresponse (Song 1983). Classes of photoreceptors suggested to be involved in protist photoresponse include flavins (the ciliates Loxodes sp., Finlay and Fenchel 1986, and Chlamydodon mnemosyne; Selbach and Kuhlmann 1999), hypericins ( ciliates; Lobban et al. 2007), and rhodopsins (in the ciliates ,

Stentor coeruleus; Fabczak et al. 2008, Fabrea salina; Podesta et al. 1994, Paranema trichophorum; Saranak and Foster 2005).

Rhodopsin is a seven-transmembrane photoreceptor in the class of G-protein couple receptors (GPCR) (Palczewski 2006). Rhodopsin consists of an opsin (trans- membrane protein) and a chromophore (retinal). The 11-cis-retinal is located in the transmembrane region of the opsin and is covalently bound by a protonated Schiff base.

When the chromophore absorbs a photon, it changes form to all-trans-retinylidene, leading to a conformational change in the opsin (i.e. inactive to active). The active opsin then binds to intracellular G-proteins and a signal transduction cascade begins

(Palczewski 2006); in the case of a photosensitive protist this would allow the cell to

‘perceive’ the light and could lead to a change in behavior.

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1.4 Organisms used in this study

The marine phagotrophic organisms used in these studies include a small (15 µm long) marine hymenostome ciliate, Uronema sp., and the marine dinoflagellate Oxyrrhis marina. These protists were chosen because they are robust and can be grown to high density in laboratory culture. The ciliate was isolated from Oregon coastal seawater and grown on a bacterial isolate cultured from the same seawater. The heterotrophic dinoflagellate (CCMP 604), obtained from the national marine phytoplankton collection at the Bigelow Laboratory for Ocean Sciences, West Boothbay Harbor, ME, was originally isolated from Friday Harbor, WA, and was cultured using an algal prey,

Dunaliella tertiolecta (CCMP 1320) grown in f/2 medium.

1.5 Goals of this study

The goals of this thesis changed throughout the course of the study. This was partly due to the availability of existing protocols, the ability to modify those protocols, and the development of new methods. Originally a large portion of the proposed study was aimed toward an investigation of predator/prey attachment and what cell surface molecules on the predator and prey were involved. The study quickly changed to an investigation of intracellular signal transduction following a chemotactic response.

Subsequently, an attempt was made to isolate chemo-receptor molecules from the cell surface of O. marina, but the only receptors recovered were rhodopsins. After finding a report by Zhang et al. (2007) that Oxyrrhis marina (strain: CCMP 1795; Long Island sound, CT, USA) had at least 7 encoding for rhodopsins, and realizing that the capacity of O. marina to respond to light stimuli had not yet been explored, the focus of

10 this study shifted to the photosensory behavior of O. marina as a model for how light can be used by heterotrophic marine protists to orient in their environment.

The studies reported in the following manuscripts seek to bridge the gap between cell and molecular biology and ecology. Much is known, for example, about how signals are transferred from one molecule to the other at the receptor and intracellular level.

However, relatively little is understood about how this relates to actual physiological and behavioral response at the microbial level in marine ecosystems. This is especially true in the context of feeding behavior by marine heterotrophic protists. Results from the following studies provide evidence that sensory responses are involved in the behavior of both O. marina and Uronema sp., and that O. marina exhibits photosensory response that may aid in detection of algal prey.

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Using Inhibitors to Investigate the Involvement of Cell Signaling in Predation by Marine Phagotrophic Protists.

Aaron Hartz, Barry Sherr, Evelyn Sherr

Journal of Eukaryotic Blackwell Publishing Inc. 350 Main Street

Malden, MA 02148 USA

Volume 55, Issue 1, 2008

12

Using Inhibitors to Investigate the Involvement of Cell Signaling in Predation by Marine Phagotrophic Protists

AARON J. HARTZ, BARRY F. SHERR and EVELYN B. SHERR

College of Oceanic and Atmospheric Sciences, Oregon State University, Corvallis,

Oregon 97331

Journal of Eukaryotic Microbiology, 55(1), 2008 pp. 18–21

2.1 Abstract

Phagotrophic protists are major consumers of microbial biomass in aquatic ecosystems. However, biochemical mechanisms underlying prey recognition and phagocytosis by protists are not well understood. We investigated the potential roles of cell signaling mechanisms in chemosensory response to prey, and in capture of prey cells, by a marine ciliate (Uronema sp.) and a heterotrophic dinoflagellate

(Oxyrrhis marina). Inhibition of protein kinase signal transduction biomolecules caused a decrease in both chemosensory response and predation. Inhibition of G protein coupled receptor signaling pathways significantly decreased chemosensory response but had no effect on prey ingestion. Inhibitor compounds did not appear to affect general cell health, but had a targeted effect. These results support the idea that cell signaling pathways known in other eukaryotic organisms are involved in feeding behavior of free‐living protists.

Key Words. Cell signaling, chemosensory response, heterotrophic protist, Oxyrrhis marina, prey ingestion, Uronema sp.

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2.2 Introduction

Response of free‐living phagotrophic protists to potential prey can be complex, resulting in selective ingestion or avoidance of prey cells (Boenigk and

Arndt 2000a, b; Fenchel 1987; Strom 2000; Verity 1991; Wolfe 2000). Phagotrophic protists also exhibit chemosensory response to prey cells or to attractant/repellant compounds (Fenchel and Blackburn 1999; Fenchel and Jonsson 1988; Sibbald,

Albright, and Sibbald 1987). However, little is known regarding specific biochemical mechanisms underlying protistan feeding behavior.

At the cellular level, prey selection or chemosensory response is likely to be based on cell signaling events initiated by binding of specific ligand compounds to receptor molecules associated with cell membranes. Ligand–receptor binding at the cell surface can activate intracellular signal transduction, in which phosphorylation of specific proteins (e.g. G proteins or protein kinases) leads to processes in the cell that can result in changes in cell behavior, such as change in motility or phagocytosis of a prey cell. Phosphorylation of specific proteins is known to be a widely used molecular mechanism involved in switch‐like outputs in cell behavior

(Alberts et al. 2002; Gomperts, Kramer, and Tatham 2004; Hunter 2000; Rasmussen et al. 1996). Tyrosine kinase proteins in particular are important in phagocytosis in both animal and protist cells (King, Hittinger, and Caroll 2003; Renaud et al. 2004;

Rosales 2005).

G‐protein coupled receptors (GPCRs) are a ubiquitous part of the cellular machinery in a range of eukaryotes, including protists (Van Haastert and Devreotes

14

2004). GPCRs have been show to be part of the chemosensory machinery used by immune cells in mammalian systems (Devreotes and Janetopoulos 2003; Haribabu et al. 1999) and in the Dictyostelium discoideum (Manahan et al. 2004;

Traynor et al. 2000). Components of G‐protein and tyrosine kinase signaling pathways appear highly conserved from sponges to vertebrates, and have been found in choanoflagellates (King and Carroll 2001; King et al. 2003). The of two ciliates (Tetrahymena: www.ciliate.org; : http://paramecium.cgm.cnrs gif.fr/db/index) include genes involved in G‐protein pathway signaling.

We used specific inhibitor compounds to investigate the extent to which cell signaling mechanisms may be involved in chemosensory response and predation by two free living marine protists ‐ a bacterivorous ciliate, Uronema sp., and a herbivorous dinoflagellate, Oxyrrhis marina.

2.3 Materials and Methods

Cultures.

Cultures were grown and maintained in autoclaved 0.2 µm filtered seawater

(FSW) at ~20 °C. A small (15 µm long) hymenostome ciliate, Uronema sp. was isolated from Oregon coastal seawater and grown on a bacterial isolate cultured from the same seawater. The bacterial isolate was maintained on agar slants

(marine agar; 2216, BD Difco, Franklin Lakes, NJ) and grown in marine broth (5.0 g peptone11.0 g extract, BD Difco, L‐1 FSW) before experiments. The heterotrophic dinoflagellate O. marina (CCMP 604) was cultured using an algal prey,

15

Dunaliella tertiolecta (CCMP 1320) grown in f/2 medium.

Chemical inhibitors.

Inhibitors of specific cell signaling mechanisms were: (1) genistein—an inhibitor of protein tyrosine kinase (Sigma‐Aldrich, St. Louis, MO); (2) staurosporine—an inhibitor of phospholipid/calcium‐dependent protein kinase

(Sigma‐ Aldrich); (3) NF023—an antagonist for G‐protein a‐subunits (Calbiochem,

San Diego, CA); (4) guanosine—a GDP analog that competitively inhibits G‐protein activation by GTP (Calbiochem); (5) GP­ant—an inhibitor of G‐protein activation by

GPCRs (Tocris Bioscience, Ellisville, MO); and (6) SCH­202676—an inhibitor of ligand binding to G protein coupled receptor (Tocris Bioscience). Initial concentrations (see Table 1) were chosen based on manufacturers recommendation of IC50 (concentration required for 50% inhibition). Compounds (1), (2), and (6) were dissolved in DMSO and (3), (4), and (5) were dissolved in 0.2 µm FSW. Ranges of concentrations of inhibitors were added to 2ml of protist cells in late log phase of growth, and equivalent volume of FSW or DMSO (final concentration <1 µl/ml) were added to control treatments. All protist treatments were preincubated for 30 min before testing for chemosensory or predation response. Cells were observed at

200X magnification for irregularities in swimming behavior as a qualitative assessment of general cell health with and without inhibitors.

Chemosensory assay.

Based on the protocols of Sjoblad, Chet, and Mitchell (1978), triplicate 1.5‐ml microcentrifuge tubes (Fisher Scientific, Waltham, MA) containing 0.5‐ml pre incubated protist cells (104 cells/ml) were laid horizontally and a 15‐µl glass

16 capillary (Drummond Scientific Co., Broomall, PA) filled with either FSW or

FSW+attractant was inserted into each tube. Attractants used were 25‐mg peptone

(BD Difco) l‐1 FSW for Uronema sp. and D. tertiolecta lysate (concentrated by centrifugation, 350 g for 5 min, lysed by sonication, vortexed, and filtered through a

0.2‐µm filter) for O. marina. After 1 h, the capillary contents were dispensed into a microcentrifuge tube containing 0.75 ml FSW, fixed with paraformaldehyde (final concentration ~1% w/v), stained with DAPI (Sigma‐Aldrich; 25 µg/ml; Sherr,

Caron, and Sherr 1993), and filtered onto a black membrane filter (0.8‐µm pore size). Filters were mounted on slides and protist cells on the whole filter enumerated via epifluorescence microscopy. Results were reported as a chemosensory index (cells in capillaries/cells in control capillaries) in which values < 1.0 indicate reduced chemosensory response.

Predation assay.

Prey cells were concentrated by centrifugation (bacteria: 6,400 g for 5min; D. tertiolecta: 350 g for 5min), re‐suspended in 1ml of FSW, and added to duplicate subsamples of protist cells at concentrations of 107 ml‐1 for bacteria, and 105 ml‐1 for

D. tertiolecta and incubated at ~ 20 °C. Protist cells were allowed to graze on prey items for 3 h. At time 0 and 3 h, 0.5 ml samples were transferred to 2.0‐ml cryovials, fixed with paraformaldehyde (final concentration ~ 1% w/v) for 10 min, and stored at ‐80 °C. Thawed samples were analyzed using a FACSCalibur flow cytometer (BD

Bioscience, Franklin Lakes, NJ) using protocols of Sherr, Sherr, and Longnecker

(2006). Remaining prey items were counted at time 0 and 3 h to evaluate number of prey consumed. Bacteria were stained with SYBR Green I (Invitrogen Corp.,

17

Carlsbad, CA; Marie et al. 1997) before flow cytometric analysis; bacteria counts were based on SYBR Green fluorescence and D. tertiolecta were detected based on

Chl‐a fluorescence. Results were reported as an index of prey survival (prey in treatment/prey in control) in which values >1 indicate decreased prey ingestion by protists.

Apoptosis test.

The Vybrant Apoptosis Kit #2 (Invitrogen Corp.) containing propidium iodide, which stains DNA in membrane‐ compromised cells, and AnnexinV‐

AlexaFluor488 (Invitrogen Corp.), which binds to externalized phosphatidylserine and is indicative of early stages of programmed cell death (van Engeland et al. 1998) were used to evaluate the possible artifact of the protein kinase inhibitors initiating programmed cell death (Elliott, Scarpello, and Morgan 2002). Protist cells were incubated with either 1 µl/ml DMSO, 50–100 µM genistein, 250–500 nM staurosporin, or 1–2mM hydrogen peroxide, a known inducer of apoptosis (Das,

Mukherjee, and Shaha 2001). Flow cytometry was used to analyze protist cells for cell‐specific change in green and red fluorescence indicating Annexin and PI binding, respectively.

2.4 Results/Discussion

Effect of inhibitors on general cell health.

We evaluated whether the inhibitor compounds used might have non‐ specific effects on the physiology of phagotrophic protists. Cell swimming behavior did not appear to be qualitatively affected by incubation with DMSO or inhibitor

18 compounds, except for 1 µM SCH‐ 202676, which resulted in complete cessation of movement. Genistein and staurosporine did not induce programmed cell death‐like symptoms in O. marina and Uronema sp. at the concentrations used in this study based on AnnexinV/PI binding (data not shown).

Effects of inhibitors on chemosensory response and prey ingestion.

The protein kinase inhibitor staurosporine decreased chemosensory response in Uronema sp. (t‐test; n=3; P<0.005) and O. marina (t‐test; n=3; P<0.05) and predation in both protists (Table 1). In the ciliate, genistein inhibited the chemosensory response (t‐test; n=3; P<0.005) and prey ingestion, but not in O. marina (Table 1A). SCH‐202676, an inhibitor of GPCRs effectively reduced the chemosensory response in O. marina (t‐test; n=3; P<0.05) and Uronema sp. (t‐test; n=3; P<0.02) (Table 1). Two inhibitors of G proteins (NF023 and guanosine) resulted in a moderate decrease in chemosensory ability in both protists, but had little effect on predation. GPant‐2 (25 µM) decreased the chemosensory response in the ciliate by ~ 70% (t‐test; n=3; P<0.005) but had no effect on the chemosensory ability in O. marina and no effect on predation in either protist. Results from predation experiments (n=2) were not assigned a statistical comparison.

Our study is the first to use selective inhibitor compounds to examine the role of cell signaling pathways in chemosensory behavior and prey ingestion by free‐living of marine phagotrophic protists and provides evidence that biochemical mechanisms involving cell membrane‐bound GPCR molecules and intracellular signal transduction molecules are likely to be involved in feeding behavior of phagotrophic protists. Other studies provide additional evidence suggesting

19 biochemical signaling mechanisms are involved in chemosensory response and feeding behavior of free‐living aquatic protists. Leick et al. (1997) found that protein kinase inhibitors eliminated chemosensory behavior in the freshwater hymenostome ciliate Tetrahymena sp. In addition, a cell surface mannose‐binding lectin appears to be directly involved in prey recognition by O. marina (Wootton et al. 2007) and Strom, Wolfe, and Bright (2007) suggested that dissolved amino acids

(L‐enantiomers) may serve as signal molecules that can affect feeding responses in a marine ciliate.

Ligand receptor binding at the cell surface, and signal transduction pathways are likely to control grazing behavior of marine phagotrophic protists. These mechanisms have been shown to operate in ‘‘model’’ cells ranging from protists to mammalian white blood cells (Alberts et al. 2002; Devreotes and Janetopoulos

2003; Gomperts et al. 2004; Haribabu et al. 1999; Hunter 2000; King and Carroll

2001; Manahan et al. 2004; Rasmussen et al. 1996; Renaud et al. 2004; Rosales

2005; Traynor et al. 2000; Van Haastert and Devreotes 2004). It is likely that such cellular processes are vitally important in determining prey selection, capture success, and rates of ingestion among heterotrophic protists in the sea.

Understanding the extent to which cell surface ligand– receptor binding and consequent signal transduction pathways operate in protist prey selection, ingestion rates, and other aspects of feeding behavior may result in the development of novel approaches to study of predator prey interactions in microbial communities.

Our study suggests at least two phagotrophic marine protists are utilizing

20

GPCRs to detect molecules that evoke a chemosensory response. Once a stimulus is encountered, intracellular signaling, mediated by protein kinase phosphorylation, is probably involved in producing a physiological response leading the protist to move toward an attractant or ingesting a prey item. Although the results of this study are preliminary, they do demonstrate that signaling pathways studied in other cell lines are involved in the chemosensory ability and phagocytosis in marine as indicated by experiments utilizing inhibitors. Additional studies are needed to further elucidate the role of cell signaling in protist feeding behavior.

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Table 1. Summary of results for signal transduction inhibition of prey ingestion and chemosensory response in: A) the heterotrophic ciliate Uronema sp. feeding on a marine bacterial isolate, and B) the heterotrophic dinoflagellate Oxyrrhis marina feeding on the marine alga Dunaliella tertiolecta. Data for prey survival index and chemotaxis index are given as mean values ± 1 standard error (SE). A prey survival index > 1 indicates decreased protist grazing on prey cells; a chemotaxis index < 1 indicates decreased chemosensory response, compared to control treatments without the inhibitor. Control values = 1.

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(A) Ciliate, Uronema sp.

Predation response Chemosensory response

Inhibitor Inhibitor Prey Inhibitor Chemotaxis Cellular survival Index (SE) target Conc. index (SE) Conc.

Staurosporine 10 nM 1.4 (0.03) 10nM 0.65 (0.21)

Protein 50 nM 4.3 (0.02) 50nM 0.54 (0.22) kinase (general) 100 nM 4.5 (0.2) 100nM 0.04 (0.014) 250 nM 4.3 (0.4) ‐‐‐‐‐‐‐‐ ‐‐‐‐‐‐‐‐‐‐‐‐

Genistein 25 µM 2.2 (0.03) 25 µM 0.10 (007) Protein kinase (general) 100 µM 4.0 (0.1) ‐‐‐‐‐‐‐‐‐‐ ‐‐‐‐‐‐‐‐‐‐‐‐

Guanosine 50 nM 0.84 (0.03) 250 nM 1.1 (0.15) G‐protein 100 nM 1.6 (0.5) 500 nM 0.84 (0.05)

150 nM 1.4 (0.2) 60 µM 0.61 (0.048)

G‐protein GPant‐2 25 µM 1.0 (0.01) 25 µM 0.31 (0.02)

NF023 500nM 0.82 (0) 500 nM 0.81 (0.04) G‐protein 1 µM 0.83 (0.05) 1 µM 0.89 (0.06)

SCH‐202676 ‐‐‐‐‐‐‐‐‐‐ ‐‐‐‐‐‐‐‐‐‐‐ G‐protein 100 nM 0.92 ((0.15) coupled receptor 500 nM 1.18 (0.1) 500 nM 0.10 (0.04)

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B. Heterotrophic dinoflagellate, Oxyrrhis marina

Predation response Chemotaxis response

Inhibitor Inhibitor Prey Inhibitor Chemotaxis Cellular survival Index (SE) target Conc. index (SE) Conc.

Staurosporine 100 nM 1.2 (0.1) 100 nM 0.62 (0.2) Protein kinase 250 nM 1.4 (0.1) 250 nM 0.20 (0.07) (general) 500 nM 1.9 (0.2) 500 nM 0.16 (0.09)

Protein Genistein 25 µM 1.1 (0.2) 25 µM 1.0 kinase (general)

G‐protein Guanosine 50 µM 1.1 (0.1) 100 µM 0.76 (0.28)

NF023 500 nM 1.1 (0.1) 10 µM 0.77 (0.08) G‐protein 50 µM 1.0 (0.04) 50 µM 0.75 (0.28)

G‐protein GPant‐2 25 µM 1.0 (0.1) 25 µM 1.0 (0.05)

G‐protein SCH‐202676 ‐‐‐‐‐‐‐ ‐‐‐‐‐‐‐‐‐‐‐ 100 nM 0.93 (0.17) coupled receptor 500nM 1.0 (0.1) 500 nM 0.40 (0.01)

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2.5 Acknowledgements

Funding for this study was provided by NSF grant OCE‐0647593 to B. and E. Sherr.

2.6 References

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Boenigk, J. & Arndt, H. 2000b. handling during interception feeding of four species of heterotrophic nanoflagellates. J. Eukaryot. Microbiol., 47:350–358.

Das, M., Mukherjee, S. & Shaha, C. 2001. Hydrogen peroxide induces apoptosis‐like death in Leishmania donovani promastigotes. J. Cell Sci., 114:2461–2469. Devreotes, P. & Janetopoulos, C. 2003. Eukaryotic chemotaxis: distinctions between directional sensing and polarization. J. Biol. Chem., 278: 20445–20448.

Elliott, J., Scarpello, J. & Morgan, N. 2002. Differential effects of genistein on apoptosis induced by fluoride and pertussis toxin in human and rat pancreatic islets and RINm5F cells. J. Endocrinol., 172:137–143.

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Strom, S., Wolfe, G. & Bright, K. 2007. Inhibition of marine planktonic protists by amino acids. I. Feeding and behavior in the ciliate Favella sp. Aquat Microbiol. Ecol., 47:107–121.

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3 Photoresponse in the Colorless Marine Dinoflagellate Oxyrrhis Marina.

Aaron Hartz, Barry Sherr, Evelyn Sherr

TO BE SUBMITTED

Journal of Eukaryotic Microbiology Blackwell Publishing Inc. 350 Main Street

Malden, MA 02148 USA

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Photoresponse in the colorless marine dinoflagellate Oxyrrhis marina

Aaron Hartz*

Barry Sherr

Evelyn Sherr

College of Oceanic and Atmospheric Sciences Oregon State University

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3.1 Abstract

As phagotrophic protists are the major consumers of phytoplankton and in the sea, it is important to understand the capabilities of these unicellular predators to respond to environmental stimuli which may aid in orientation in the environment and in detecting prey cells. In an investigation of potential signal receptor proteins in the cell membrane of a strain of the marine colorless dinoflagellate Oxyrrhis marina, we discovered two expressed rhodopsins which matched two of seven rhodopsin‐coding genes reported in the NCBI database for another strain of O. marina. A BLASTp search based on the rhodopsins of O. marina showed them to be most closely related to a rhodopsin of the autotrophic dinoflagellate Pyrocystis lunula, and to rhodopsins of Bacteria rather than of Archaea or other eukaryotic microbes. We inferred these to be sensory rhodopsins, a type of

G‐protein coupled receptor (GPCR) trans‐membrane signaling molecules. Since phototaxic behavior based on sensory rhodopsins has been reported in other protists, we investigated the photosensory response of O. marina. The dinoflagellate exhibited strongest positive phototaxis at low levels (2 – 3 µE m‐2 s‐1) of white light when the cells were previously light adapted and/or starved. Positive phototaxis was also found for blue (450 nm), green (525 nm) and red (680 nm) wavelengths.

In a further test, O. marina showed significantly greater phototaxis toward concentrated algal food, compared to bleached algae, illuminated by blue light to stimulate red chlorophyll‐a (Chl‐a) autofluorescence. Concentration of a cytoplasmic downstream messenger molecule, cyclic adenosine monophosphate

30

(cAMP), a component of the signaling pathway of GPCRs, increased in O. marina cells after exposure to white light. In addition, treatment with hydroxylamine, a rhodopsin signaling inhibitor, significantly decreased phototaxic response in the dinoflagellate. Our results demonstrate that a colorless marine dinoflagellate can orient to light based on rhodopsin present in the outer cell membrane and may be able to use photosensory response to detect algal prey based on Chl‐a autofluorescence.

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3.2 Introduction

How phagotrophic protists orient in their environment and locate prey is a research topic of longstanding interest. Chemosensory response to prey by marine heterotrophic protists is well known (Sibbald et al. 1987, Verity 1988, Fenchel and

Blackburn 1999). Studies of microbial response to light stimuli have an even longer history, dating to the 19th century. Verworn (1889) noted: “we find that among unicellular organisms, so far as they are irritable at all to light, phototaxis is a wide‐ spread phenomenon.” Since that time, significant advances have been made in understanding the biochemical basis of photosensory response in unicellular algae

(Kateriya et al. 2004, Hegemann 2008). Photosensory capability in heterotrophic protists has received less attention. Prior studies have found variable photosensory response in an amoeba (Mast and Stahler 1937), a colorless euglena (Saranak and

Foster 2005), and several species of ciliates (Song 1983, Fenchel and Finlay 1986,

Podesta et al. 1994, Selbach et al 1999, Cadetti et al. 2000, Sobierajska et al. 2006,

Lobban et al. 2007, Fabczak et al. 2008).

The marine heterotrophic dinoflagellate Oxyrrhis marina is widely distributed in coastal habitats, ranging from tidal pools to open coastal water, exhibits a high degree of genetic and functional diversity, and has been used as a model marine protist in laboratory experiments (Lowe et al. 2005, Menden‐Deuer and Grumbaum 2006, Wooten et al. 2007, Hartz et al. 2008). The findings we report here: that sensory rhodopsins present in the outer cell membrane of a strain of O. marina may be the basis of observed photosensory response, expands the scope of

32 behavioral abilities of this species and of heterotrophic dinoflagellates in general.

Although sensory rhodopsins have been described in a few species of ciliates, fungi, and algae (Ridge 2002, Ruiz‐Gonzalez & Marin 2004, Sharma et al. 2006, Fabczak et al. 2008, Klare et al. 2008), our study is the first to report phototaxic behavior due to a putative sensory rhodopsin expressed by a colorless dinoflagellate.

3.3 Materials and Methods

3.3.1 Cultures:

Algal food, Dunaliella tertiolecta (CCMP 1320), was grown on f/2 medium made with 0.2 µm filtered autoclaved seawater (FASW) obtained from the Oregon coast. Oxyrrhis marina (CCMP 604; originally isolated from Friday Harbor, WA, USA) was grown in FASW on D. tertiolecta as the sole food. All cultures were maintained at room temperature (20 ‐ 25 oC) with a 12 hour light/dark cycle under natural daylight fluorescent lamps. For photosensory experiments that tested response based on light history, light adapted O. marina were grown with a 12 hour light/dark cycle, while dark adapted O. marina were grown for 3 transfer cycles, with approximately 4 days between transfers, in the dark at room temperature.

3.3.2 Flow cytometric analysis:

The abundances and cell‐specific Chl‐a fluorescence of the dinoflagellate and algal prey were determined with a Becton‐Dickinson FACSCaliber Flow Cytometer using a 488 nm laser. Depending on cell abundances, from 100 to 300 μl of sample were processed during run times of 3 ‐5 minutes. BD CellQuest software was used to set flow cytometer photomultiplier tubes to optimize detection of O. marina and

33

D. tertiolecta cells based on side scatter and FL‐3 (red) fluorescence properties

(Figure 1). In preliminary tests, we determined that preservation of O. marina with buffered formaldehyde resulted in cell loss, and that preservation even with low final concentration of acid Lugol solution interfered with flow cytometric analysis, so for experiments live cells were promptly assayed.

The relative cell‐specific red (FL‐3) fluorescence of O. marina was used to assess feeding state. Dinoflagellate cells with ingested algal prey had FL‐3 fluorescence signatures equivalent to amount of fluorescence observed in one to several algal cells (upper right box in Figure 1). Cultures that had consumed algal prey to undetectable abundance, but still had traces of prey in food vacuoles, based on average cell‐specific FL‐3 fluorescence 2‐3 fold higher than that of completely starved cells, were considered to be well fed. Two days after all prey were consumed, O. marina cells had negligible FL‐3 fluorescence and were considered to be starved.

3.3.3 Extraction and characterization of O. marina membrane proteins:

Initial protein extractions were done using the Mem-PER protein extraction kit (Pierce, Thermo Fisher Scientific, Rockford, IL). As this detergent‐based protocol lyses cells, all membrane proteins, including those from intracellular , are extracted. One‐liter cultures of O. marina were grown under light until negligible red fluorescence of ingested prey remained. The entire liter of culture was concentrated via centrifugation at 2200 RPM (~500 x g) for a yield of approximately 1‐2 x 107 O. marina cells. After membrane extraction, detergent was cleaned from proteins with

34 a SDS-PAGE sample preparation kit (Pierce, Thermo Fisher Scientific). Extracted proteins were concentrated using a Model SPD111V Savant SpeedVac. Proteins in aliquots of the concentrated sample were separated via SDS-PAGE gel electrophoresis (Ready Gel Tris-HCL 4-15%; Bio-Rad, Hercules, CA), and resulting protein bands were stained with BioSafe Coomassie stain (Bio-Rad) or Coomassie

Fluor Orange (Invitrogen Corp., Carlsbad, CA). Individual gel bands were excised and digested using an In-Gel Tryptic Digestion Kit (Pierce, Thermo Fisher Scientific.

Digested protein samples were concentrated by vacuum centrifugation and then analyzed on an LC‐MS/MS mass spectrometer at the Oregon State University Mass

Spectrometry Facility.

Since the Mem-PER extraction protocol non‐specifically targets all membrane proteins, an additional protein extraction was done with the Hook Cell Surface

Protein Isolation Kit (G-Biosciences, Maryland Heights, MO). This protocol uses a biotin-tag/streptavidin affinity column to extract only proteins exposed on the outer plasma cell membranes of intact cells. O. marina culture conditions, cell concentration, and protein separation and analysis were as described above.

Protein data were searched against databases of known proteins using

Mascot software (Matrix Science, Boston, MA.) The rhodopsin proteins identified in our study were used as a query in a BLAST search (NCBI, BLASTp using preset parameters). From the BLASTp output we selected the top 30 sequences (E values ranging from 3e-145 to 2e-20). The protein sequences were imported into the

Geneious software (Biomatters, Ltd., Auckland, NZ) based on an algorithm

35 developed by Guindon and Oliver (2003). The sequences were aligned and a tree produced using maximum likelihood (bootstrapped 1000 times).

3.3.4 Phototaxis experiments:

To determine O. marina response to light, rectangular four-well polystyrene dishes (Nunc, Rochester, NY) were cut and separated into individual rectangular dishes. In a darkroom, light emitting diodes (LED’s) were placed 6 cm away from individual dishes and connected to a 13.8 volt power supply (RadioShack

Corporation, Fort Worth, TX) (Figure 2‐A). For each experiment, 20 ml of well mixed

O. marina culture were added to each of four replicate dishes, and each dish was exposed to a LED light for 30 minutes. Various phototaxis experiments used either white light LED’s (DigiKey Corp., Thief River Falls, NM) or red (680 nm), green (525 nm) or blue (430, 450 nm) LED’s (Roithner Lasertechnik, Vienna, Austria). Light level (μE m2 s‐1) was varied by placing neutral density film (GamProducts, Inc.,

Hollywood, CA) in front of the LED lights. Light levels were measured using a PAR sensor (Heinz Walz GmbH, Effeltrich, Germany).

After 30 minutes of exposure of O. marina cells to light, samples were taken from the proximal (P) and distal (D) ends of the dish relative to the light. Plastic transfer pipettes cut diagonally at the tip were used to draw sample from the entire depth of the water column (about 1cm) at the P and D ends of the dish. Cell per ml abundances were determined immediately via flow cytometry. A phototaxis index:

(P-D)/(P+D), with values ranging from 0 to 1, was calculated from the cell abundance data. Positive index values indicated that the cells moved towards the

36 light, values near 0 indicated no photoresponse, and negative values indicated that the cells moved away from the light.

Additional phototaxis experiments were carried out to investigate the possibility that O. marina can detect red Chl‐a fluorescence induced by blue light.

The dishes described above were used in preliminary experiments. 200 ml of late log phase D. tertiolecta culture was centrifuged at 350 g for 5 minutes and re- suspended in 10 ml FASW. Three 2 ml glass vials were filled with concentrated D. tertiolecta and sealed to prevent potential chemosensory cues. The sealed vials were placed at one end of three experimental dishes. As a control, sealed vials filled with concentrated algae photo‐bleached under intense white light for approximately 90 minutes were positioned similarly in three other dishes. All dishes were illuminated with blue 450 nm light under the sealed vials of algae (Figure 2‐B).

The rest of the experimental protocol was as described for the initial phototaxis experiments. At the end of the experiment, photo‐bleached cells were examined using an Olympus BX61 epifluorescence microscope to confirm that the algal cells remained intact. The relative cell‐specific FL‐3 (red) fluorescence in bleached and in unbleached algal cells used in the experiments was assessed by flow cytometry.

In order to further evaluate O. marina photoresponse to chlorophyll autofluorescence of algal prey, an additional linear tube experimental design was devised (Figure 2‐C). Two 6 cm sections of clean 10 mm I.D. glass tubing were connected with tubing. A sealed glass vial containing either concentrated unbleached or bleached D. tertiolecta, prepared as described above, was connected

37 to one end of each tube with silicon tubing. The opposite end contained a reservoir of ~2 x 104 cells ml‐1 of O. marina open to the glass tube. The glass tubing was filled with FASW, allowing the dinoflagellate cells to swim freely through the connected glass tubes. The sealed vial with concentrated algae was illuminated with a 450 nm

LED positioned at a 90° angle with respect to the glass tubing. The suspension of concentrated unbleached algae was sufficiently dense to be visibly red when illuminated with blue light. Each treatment was done in triplicate. After 1.5 hours, the 6 cm sections of glass tubing adjacent to the vials of algae were removed and abundances of O. marina in these tube sections were immediately analyzed by flow cytometry.

3.3.5 Evaluation of cAMP induction by light:

Cyclic adenosine monophosphate (cAMP) is a downstream cytoplasmic component of membrane‐bound receptor signaling. If the photoresponse of O. marina were due to a membrane bound GPCR such as rhodopsin, exposure to light should result in a detectable increase in intracellular cAMP concentration. The cAMP-Glo assay (Promega Corp., Madison, WI) was used to investigate the potential for such a response. Dark‐cultured O. marina cells were either left in the dark or stimulated by exposure to a white light LED for periods of 1 to 90 seconds.

Following light exposure, cells were lysed and the cAMP‐Glo reaction mix added. In this assay, reaction with cAMP results in a decrease in concentration of ATP, assessed by decreased luciferase light production. Thus there is an inverse relationship between the luminescent output of the assay and cAMP concentration.

38

Sample aliquots were placed in white 96 well plates (Greiner Bio-One North

America, Monroe, NC) and relative luminescence units recorded using a microplate luminometer (SpectraMaxL; Molecular Devices, Sunnyvale, CA.).

3.3.6 Inhibition of Rhodopsin:

Hydroxylamine, an inhibitor of rhodopsin GCPR signaling, was used to investigate whether O. marina photoresponse could be based on rhodopsin. Sensory rhodopsin consists of a retinal chromophore covalently linked to the rhodopsin protein by a protonated Schiff base bond (Palczewski 2006). Hydroxylamine cleaves the retinal from the protein, rendering the sensory rhodopsin disfunctional

(Zadok et al. 2005). Hydroxylamine (Sigma-Aldrich Inc., Saint Louis, MO) was dissolved in autoclaved 0.2 μm filtered seawater. To check whether hydroxylamine might have a non‐specific effect on cell physiology, aliquots of O. marina light adapted/well fed cells were incubated with hydroxylamine at final concentrations of

0.0, 0.25, 0.5, 0.75, 1, and 2 mM for 2 hours at room temperature in the dark.

Approximately 104 cells ml‐1 of algal prey were then added to all treatments. After a further two hour incubation in the dark, all treatments were sampled for O. marina cell abundance and cell‐specific prey ingestion using flow cytometry.

To test the effect of hydroxylamine on O. marina phototactic behavior, the dinoflagellate was incubated with hydroxylamine at final concentrations of 0.0, 0.1,

0.12, 0.5, and 1 mM for 30 minutes in bright light, followed by 90 minutes in the dark prior to the start of the phototaxis experiments. Phototaxis experiments were set up in rectangular dishes with white light LEDs as previously described (Figure 2‐

39

A). The phototaxis index of cells incubated with hydroxylamine was compared to control cells with no hydroxylamine exposure.

3.4 Results

3.4.1 Characterization of O. marina membrane­associated proteins

Mascot search with protein analysis data from the Mem‐PER membrane protein extraction kit samples resulted in the identification of an expressed rhodopsin (O. marina; gi|157093543). A similar search of proteins from gel bands from the Hook outer membrane extraction kit identified two expressed rhodopsins

(O. marina; gi|157093543 and gi|157093545). No other putative signal receptor proteins were detected in our samples. A maximum likelihood tree of the top 30 matches to the two O. marina rhodopsins recovered showed that the rhodopsins found in this study were most closely related to other O. marina rhodopsins and to the rhodopsin of an autotrophic dinoflagellate, Pyrocystis lununa. These dinoflagellate rhodopsins grouped with rhodopsins present in Bacteria,

Phylum Proteobacteria (Figure 3).

3.4.2 O. marina response to light

O. marina showed variable response to light depending on light level and wavelength, as well as on growth history of the dinoflagellate cells. Based on a phototaxis index, the strongest positive photoresponse was observed at low light intensity, 2 or 3 μE m‐2 s‐1 (Figure 3). The phototaxis index decreased at the upper and lower light levels and became negative in the case of light adapted‐well fed cells

40 at the highest light level tested. However, cells that were dark adapted‐well fed showed negative photoresponse to all light levels tested. In experiments to test photoresponse to individual light wavelengths, significant positive phototaxis occurred in treatments with 450 nm, 525 nm, and 680 nm light (Figure 4). In general, the phototaxis index values for individual light wavelengths were within the range of those from experiments with white light.

3.4.3 O. marina response to Chl­a fluorescence

In the initial dish experiment, results were suggestive of phototaxis toward

Chl‐a fluorescence, but not definitive. In the linear tube experimental configuration,

O. marina phototaxis was significantly greater (P=0.006; t test,) in response to concentrated unbleached algae compared to photo‐bleached algae illuminated with

450 nm light (Figure 5). Microscopic inspection showed that the photo‐bleaching process left algal prey cells intact. However, after photo‐bleaching, the algal cells visually changed color from green to white, and based on relative cell‐specific red

(FL3) fluorescence units as measured by flow cytometry, they emitted significantly less Chl‐a fluorescence per cell (average of 7 FL3 units) compared to normal algal cells (average of 854 FL3 units, P < 0.001; t‐test).

3.4.4 Light stimulation of intracellular downstream signaling

A one second exposure of intact O. marina cells to light resulted in a sharp intracellular increase (measured as a decrease in relative luminescence units, RLU) in cAMP concentration (Figure 6). With further light exposure up to 90 seconds,

41 cAMP levels in O. marina cells were also elevated (significantly lower RLU) compared to cAMP concentrations in control cells with no light exposure.

3.4.5 Inhibition of rhodopsin function by hydroxylamine

After exposure to hydroxylamine concentrations of 0.12, 0.5, and 1.0 mM, photosensory response was significantly decreased (P <0.0003 to 0.028; t‐test) compared to controls with no exposure to hydroxylamine. However it was observed that treatment of O. marina with as little as 0.25 mM hydroxylamine resulted in a significant decrease in O. marina cells (t‐test; p=0.014). Treatment with hydroxylamine did not affect prey ingestion.

3.5 Discussion

Although heterotrophic dinoflagellates are ubiquitous and important in marine food webs (Sherr and Sherr 2007), to date there has been little study of the behavior of these protists. Buskey (1997) investigated the feeding behavior of a heterotrophic armored dinoflagellate, Protoperidinium pellucidum, which captures and digests prey extracellularly via a feeding veil. He found that the dinoflagellate used chemoreception to judge the location of a prey cell, and fed preferentially on over dinoflagellate prey. Menden‐Deuer & Grünbaum (2006) reported that

O. marina could detect and aggregate in patches of algal prey. Wooten et al. (2007) showed that chemoreception of algal prey by O. marina was due in part to a mannose‐binding lectin present on the outer cell membrane of the dinoflagellate.

We have previously reported that intracellular signal transduction pathways are involved in the chemosensory response of O. marina (Hartz et al. 2008).

42

While the heterotrophic dinoflagellate O. marina was previously noted by

Droop (1954) to move toward light in culture, our study is the first to report details of photosensory behavior and a potential biochemical basis of light response in this species. Our results showed that O. marina has the ability to detect white light as well as individual wavelengths of light. The direction and strength of phototaxic response in O. marina varied with light intensity. The strongest positive phototaxis occurred at light levels of 2‐3 μE m‐2 s‐1 for white light and 4 μE m‐2 s‐1for 450 nm light. This may represent light intensities preferred by O. marina in its natural habitat, the coastal ocean. These values approximate the light level found at the base of the euphotic zone, about 1 % of incident light. For example, 1 % of the average surface photosynthetically active radiation of the Oregon coastal ocean, ~

35 mol quanta m‐2 day‐1 (Wetz et al. 2004), is equivalent to ~ 4 μE m‐2 s‐1.

O. marina that were grown in the dark and well fed exhibited negative phototaxis to all levels of white light tested, while dark adapted and starved cells were positively phototaxic (Figure 4). These results mirror those of Selbach et al.

(1999), who found that the ciliate Chlamydodon mnemosyne exhibited positive phototaxis during starvation and negative phototaxis when well fed. Ability to sense a change in light level may aid phagotrophic protists in orienting in their environment, i.e. as a cue to the direction of a different location if food is scarce, or to remain in a location where food conditions have been favorable.

O. marina exhibited positive phototaxis to low light levels of at least three distinct wavelengths of light: blue (450 nm), green (525 nm) and red (680 nm).

43

Blue and green wavelengths of light penetrate farthest in coastal ocean water, while red wavelength radiation is rapidly diminished in the upper few meters of the water column. However, chlorophyll pigments of phytoplankton produce red fluorescence when illuminated with blue light. This suggested the possibility that the capacity to detect red light might be used by O. marina to locate patches of phytoplankton prey.

We did find that O. marina exhibited significantly greater photoresponse toward unbleached compared to bleached algae illuminated by blue light. In these experiments, the sealed vials of unbleached algae were visibly dark red due to Chl‐a autofluorescence when illuminated with 450 nm light, while no visible red color was emitted from bleached algae. Flow cytometric analysis confirmed a more than 100‐ fold decrease in cell‐specific red fluorescence of algae after bleaching. This result indicated that O. marina was attracted to the red fluorescence emitted from algal cells. While the idea of photosensory prey detection by marine protists needs further study, we note that Amon and French (2004) showed that phagotrophic zoospores of the marine thraustochitrid Ulkenia sp. were attracted to bioluminescence from the bacterial prey species Vibrio fisheri. Furthermore, they found Ulkenia sp. zoospore phototaxis toward blue and white light peaked at 3.5 μE m‐2 s‐1, similar to light levels found by us to induce maximum phototaxic response in

O. marina.

Several types of biomolecules have been shown to serve as photosensory receptors in marine heterotrophic protists, including flavins, hypericins, and rhodopsins (Selbach and Kuhlmann 1999, Cadetti et al. 2000, Saranak and Foster

2005, Lobban et al. 2007). While the first two classes of sensory pigments are

44 mainly found in ciliates, genes coding for rhodopsins are widespread in Archaea,

Bacteria, and Eukarya. Rhodopsins are known to have photosensory functions in microbes in all three Domains (Sharma et al. 2006, Fuhrman et al. 2008, Klare et al.

2008). We found two different rhodopsin proteins expressed in the outer cell membrane of the strain of O. marina (CCMP 604) used in this study. Seven rhodopsin‐coding genes of O. marina (strain CCMP 1795) are posted in the NCBI database (www.ncbi.nlm.nih.gov). The strain of O. marina we used might have a different complement of rhodopsin genes, although the proteins we isolated matched two of the O. marina rhodopsin genes in the database. It is also possible that other expressed O. marina rhodopsins were not detected, as the dinoflagellate responded to three distinct wavelengths of light.

A BLAST search using the rhodopsin protein sequences identified in our study as a query revealed that these and the other O. marina rhodopsins in the database had a closest match to a type I (microbial) rhodopsin‐related sequence discovered in the marine autotrophic dinoflagellate, Pyrocystis lunula (Figure 3).

These dinoflagellate rhodopsins are related to Bacterial rhodopsins (Figure 3), as has previously been reported for the P. lunula rhodopsin (Ruiz‐Gonzalez and Marin

2004; Sharma et al. 2006). In contrast, sensory type 1 rhodopsins present in other eukaryotic microbes, including fungi, , and cryptophyte algae, are most closely related to Archaeal rhodopsins (Ruiz‐Gonzalez and Marin 2004; Sharma et al. 2006). The scattered distribution of eukaryotic rhodopsin‐related genes has been suggested by these authors as indicative of early and diverse origins, including

45 lateral transfer between species in separate domains, for this universal class of proteins.

Internal cell signal cascades triggered by activation of membrane‐bound

GPCR molecules such as rhodopsin have been well described in eukaryotic cells.

Cyclic adenosine monophosphate (cAMP) is an important downstream signal transduction messenger and is catalyzed by adenylyl cyclases which are activated by GPCRs (Gomparts et al. 2004). After exposure to white light for one second or longer, cAMP concentration increased in O. marina cells. We also investigated the role of rhodopsin in O. marina phototaxis by exposing the dinoflagellate to hydroxylamine, which prevents the retinylidene chromophores from re‐binding to the opsin component of sensory rhodopsin (Hegemann et al. 1988, Saranak and

Foster 1997). We found that phototaxis in O. marina was significantly decreased by treatment with hydroxylamine. It was also observed that, although hydroxylamine caused a decrease in O. marina cell count at concentrations of 0.25 mM, hydroxylamine did not affect prey ingestion or normal swimming behavior suggesting the remaining cells were relatively healthy. These results support the idea that rhodopsin may be the basis of the observed photosensory behavior of O. marina.

3.6 Conclusions

To date there has been little investigation of the ecological role of photoreceptors in predatory protists. We found that the colorless dinoflagellate O. marina responded both to white light and specific wavelengths of light and

46 exhibited both positive and negative phototaxis. O. marina may be able to locate patches of algal prey by detection of Chl‐a autofluorescence. Finding of expressed rhodopsins in the cell membrane, as well as results of experimental tests, indicated that rhodopsin is likely to be a sensory photoreceptor in this marine dinoflagellate.

The capacity to respond to light as well as to chemical cues in the environment expands the behavioral repertoire of phagotrophic protists; this research topic deserves increased attention.

47

3.7 References

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Buskey, E.J. 1997. Behavioral components of feeding selectivity of the heterotrophic dinoflagellate Protoperidinium pellucidum. Mar. Ecol. Prog. Ser 153: 77‐89. Droop, M. 1954. A note on the isolation of small marine algae and flagellates for pure cultures. J. Mar. Biol. Asso. U.K. 33:511‐514. Fabczak, H., Sobierajska, K., Fabczak, S. 2008. A rhodopsin immunoanalog in the related photosensitive protozoans Bleparisma japonicum and Stentor coeruleus. Photochem. Photobiol. Sci. 7:1041‐1045. Falkowski, P., Raven, J. 2007. Aquatic Photosynthesis, Second Edition. 484 pp. Princeton University Press. Princeton, NJ.

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Gomparts, B., Kramer, I., Tatham, P. 2004. Signal Transduction. Elsevier Academic Press, San Diego.

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Hartz, A., Sherr, B. F. & Sherr, E. B. 2008. Role of receptor/ligand binding and intracellular‐signal transduction in controlling predation behavior by marine phagotrophic protists. J. Euk. Microbiol. Hegemann, P. 2008. Algal sensory photoreceptors. Ann. Rev. Biol. 59:167‐ 189.

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Lobban, C., Hallam, S., Mukherjee, P., & Petrich, J. 2007. Photophysics and multifunctionality of hypericin‐like pigments in heterotrich ciliates: A phylogenetic perspective. Photochem. Photobiol. 83:1074‐1094.

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Mast, S. & Stahler, N. 1937. The relation between luminous intensity, adaptaion to light, and rate of locomotion in Amoeba proteus (Leidy). Biol. Bull. 73:126‐133. Menden‐Deuer, S. & Grünbaum, D. 2006. Individual foraging behaviors and population distributions of planktonic predators aggregating to phytoplankton layers. Limnol. Oceanogr. 51: 109‐116 Palczewski, K. 2006. G‐protein coupled receptor rhodopsin. Ann. Rev. Biochem. 75:743‐767. Podesta, A., Marangoni, R., Villani, C., & Colombetti, G. 1994. A rhodopsin‐like molecule on the plasma membrane of Fabrea salina. J. Euk Microbiol. 41(6): 565‐ 569. Ridge, K.D. 2002. Algal rhodopsins: phototaxis receptors found at last. Curr. Biol. 12: R588‐R590. Ruiz‐Gonzalez, M., & Marin, I. 2004. New insights into the evolutionary history of type 1 rhodopsins. J. Mol. Evol. 58:348‐358. Saranak, J., & Foster, K. 2005. Photoreceptor for curling behavior in Peranema trichophorum and evolution of eukaryotic rhodopsin. Eukaryotic Cell 4(10):1605‐ 1612.

Selbach, M., & Kuhlmann, H. 1999. Structure, fluorescent properties and proposed function in phototaxis of the stigma apparatus in the ciliate Chlamydodon mnemosyne. J. Experiment. Biol.. 202:919–927. Selbach, M., Hader, D., & Kuhlmann, H. 1999. Phototaxis in Chlamydodon mnemosyne: determination of the illuminance‐response curve and the action spectrum. J. Photochem. Photobiol. 49:35‐40.

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Sharma, A., Spudich, J., Doolittle, W. 2006. Microbial rhodopsins: functional versatility and genetic mobility. TRENDS in Microbiol. 14:463‐469. Sherr, E., & Sherr, B. 2002 Significance of predation by protists in aquatic microbial food webs. Antonie Van Leewenhoek Internat. J. Gen. Mol. Microbiol. 81:293‐308. Sherr, E., & Sherr, B. 2007. Heterotrophic dinoflagellates: a significant component of microzooplankton biomass and major grazers of diatoms in the sea. Mar. Ecol. Prog . Ser. 352: 187‐197. Sibbald, M, J., Albright L. J. & Sibbald, P. R. 1987. Chemosensory responses of a heterotrophic microflagellate to bacteria and several nitrogen compounds. Mar. Ecol. Prog . Ser. 36:201‐204. Song, P. 1983. Protozoan and related photoreceptors: molecular aspects. Ann. Rev. Biophys. Bioengineer. 12:35‐68.

Spudich, J. 2006. The multitalented microbial sensory rhodopsins. TRENDS in Microbiol. 14:480‐487.

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Verworn, M. 1899. General Physiology. 615pp. Macmillan and Co. New York. Wetz, M., Wheeler, P., & Letelier, R. 2004. Light‐induced growth of phytoplankton collected during the winter from the benthic boundary layer off Oregon ,USA. Mar. Ecol. Prog . Ser. 280:95‐104. Wootton, E., Zubkov, M., Jones, D., Jones, R., Martel, C., Thornton, C., & Roberts, E. 2007. Biochemical prey recognition by planktonic protozoa. Env. Microbiol. 9:216‐ 222. Zadok, U., Klare, J., Engelhard, M., & Sheves, M. 2005. The hydroxylamine reaction of sensory rhodopsin II: light‐induced conformational alterations with C13=C14 nonisomerized pigment. Biophysical Journal 89:2610‐2617.

Zhang, H., Hou, Y., Miranda, L., Campbell, D., Sturm, N., Gaasterland, T., & Lin, S. 2007. Spliced leader RNA trans‐splicing in dinoflagellates. PNAS. 104:4618‐4623.

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Figure Legends

Figure 1. Flow cytometer dot plot of red fluorescence versus side scatter parameters showing locations of the algal prey, D. tertiolecta, O. marina cells with ingested algal prey, and O. marina cells without ingested algal prey.

Figure 2. Phototaxis experiment configurations: A) Experimental configuration used to measure O. marina photoresponse to white and specific wavelengths of light, using LED’s to create a light gradient through a rectangular dish. B) As in A), but a sealed glass vial of concentrated algae, or a bleached algal control at one end of the rectangular dish were illuminated with a 450 nm LED placed under the dish to stimulate Chl‐a autofluoresence. C) Linear tube configuration used in an additional test of response to Chl‐a autofluorescence.

Figure 3. Maximum likelihood tree of rhodopsins most closely related to those found in O. marina. Numbers on nodes represent bootstrap values.

Figure 4. Photoresponse of O. marina to a range of intensities of white light, quantified as a phototaxis index comparing cell abundances in the proximal and distal ends of a rectangular dish after a 30 min exposure to each light level. Phototaxis index values represent the average of three replicates, error bars = standard deviation. The dotted line indicates an index value of 0, which represents no response to light.

Figure 5. Photoresponse of O. marina to a range of intensities of red, green and blue wavelengths of light in dish experiments as in Figure 4. Phototaxis index values represent average of four replicates, error bars = one standard deviation. The dotted line indicates an index value of 0, which represents no response to light.

Figure 6. Comparison of O. marina photoresponse to unbleached and bleached concentrated algae illuminated with 450 nm light in the linear tube experiment. Data represent average of triplicate samples, error bars are one standard deviation.

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Figure 7. Cyclic adenosine monophosphate (cAMP) activation in response to a light stimulus. cAMP concentration has an inverse relationship to Relative Luminiscence Units (RLU). Data points represent average of duplicate measurements and error bars are one standard error.

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4 Conclusions

4.1 Overview

The manuscripts in this thesis add new information to the scant literature on how membrane receptors and intracellular signaling are linked to behavior in marine phagotrophic protists. The results of these studies are supported by previous work on biochemical control of phagocytosis in laboratory cultured protists such as ciliates in the genera Tetrahymena and Paramecium (Almagor et al.

1981; Hellung‐Larsen et al. 1986; Renaud et al. 1995, 2004; Leick et al. 1996, 1997;

Bell et al. 1998), and euglenoid flagellates in the genera Euglena and Astacia

(Levandowsky 2001); in parasitic protists such as trypanosomes (cause of malaria) and Leishmania (cause of skin lesions) parasites (Mosser &, Rosenthal 1993, Ratcliff et al. 1996, Basseri et al. 2002); and in metazoan white blood cells (Greenberg 1995,

Hill & Rowley 1998, Ballarin et al. 2002). These studies include demonstrations of the effect on cell behavior of ligand binding by membrane‐bound receptor molecules, and of signal transduction mechanisms within the cell that are switched on by ligand‐receptor binding (Greenberg 1995, Alberts et al. 2002, Renaud et al.

2004).

4.2 Chemosensory/inhibitor study

Data reported in Hartz et al. (2008) provided evidence that signaling pathways found in other types of eukaryotic cells are, in fact, involved in swimming and feeding behavior of marine heterotrophic protists. The effects of the two protein kinase signal transduction inhibitors varied between the heterotrophic

60 dinoflagellate O. marina and the Uronema sp. ciliate. One of these inhibitors, staurosporine, decreased chemosensory response and predation in both protists, suggesting that protein kinase signaling pathways are involved in at least two behavioral responses. The second protein kinase inhibitor, genistein, inhibited chemotaxis and prey ingestion in the ciliate, but not in the dinoflagellate. This could reflect a difference in intracellular signaling pathways in the two protists, or in the ability of the cells to take up the chemical inhibitor. An inhibitor of G‐protein coupled receptors (GPCR’s) effectively reduced the chemosensory response in O. marina and Uronema sp., but had no affect on prey ingestion. Two inhibitors of intracellular G‐proteins resulted in a moderate decrease in chemosensory ability in both protists, but had little effect on predation. In the natural environment, protist feeding would most likely be affected by inhibition of chemosensory ability. The conditions used for the prey ingestion experiments in this study provided the protists with prey abundances one or two orders of magnitude greater than what would be found in situ. It is possible that protists may not need chemosensory signals to locate prey at high abundance. These findings suggest that while G‐ protein signaling systems are involved in chemotaxis in these two marine protozoa, they are probably not directly involved in prey ingestion (phagocystosis).

This study was the first to use selective inhibitor compounds to examine the role of cell signaling pathways in chemosensory behavior and prey ingestion by free‐living marine phagotrophic protists and provided evidence that biochemical mechanisms involving cell membrane‐bound GPCR molecules and intracellular signal transduction molecules are likely to be involved in feeding behavior of

61 phagotrophic protists. Many enzymes involved in signal transduction cascades are regulated by protein kinase phosphorylation/dephosphorylation. Knowing this, it comes as no surprise that general inhibitors of protein kinases had a significant effect on the chemosensory ability and grazing in both protists.

This work supports the results of other studies which indicated that biochemical signaling mechanisms are involved in chemosensory response and feeding behavior of ‘model cell’ protists. Leick et al. (1997) found that protein kinase inhibitors eliminated chemosensory behavior in the freshwater hymenostome ciliate Tetrahymena sp. In addition, Strom, Wolfe, and Bright (2007) suggested that dissolved amino acids (L‐enantiomers) may serve as signal molecules that can affect feeding responses in a marine tintinnid ciliate. It is likely that such cellular processes are important in determining prey selection, capture success, and rates of ingestion among heterotrophic protists in the sea.

Understanding the extent to which cell signaling pathways operate in protist prey selection, ingestion rates, and other aspects of feeding behavior may result in the development of novel approaches to study of predator prey interactions in microbial communities.

4.3 Photosensory behavior study

It has been over half a century since Droop (1954) noted that O. marina moved toward light during culturing studies. However Hartz et al. (ms in preparation) is the first study to provide details of O. marina photoresponse. The heterotrophic dinoflagellate Oxyrrhis marina was clearly able to respond

62 behaviorally to white, blue, green, and red light. The strongest positive phototaxis occurred between 2‐3 µE/m2/s for white light and 4 µE/m2/s for 450 nm light. This may represent an optimum light level for O. marina in its natural habitat, the coastal ocean. Although this value would most likely fluctuate, our results suggest that O. marina seeks out low light levels that may be similar to those found at the base of the euphotic zone.

Culture conditions and physiological state affected O. marina photo‐behavior.

For example, the dark adapted starved cells may have used light as a cue to find more favorable conditions. The dark adapted well fed cells were already experiencing optimum conditions with respect to food availability, and may have perceived light as a cue to less favorable conditions (i.e. they may not gain an advantage by moving from their current location if they already have a sufficient food source). Additionally, our study showed that O. marina has the ability to respond to individual wavelengths of light; this ability may enable O. marina to use light cues to find optimum conditions no matter what habitat the cell is in.

The results from this study are also the first to suggest a herbivorous protist has the ability to detect phytoplankton auto‐fluorescence. These results showed that O. marina was attracted to the induced red fluorescence emitted from

Dunaliella cells illuminated with blue light. This could have ecological implications since in the ocean most light below 10 m is blue due to attenuation of other wavelengths. Dense patches of phytoplankton in the blue‐lit water column may emit sufficient numbers of photons to serve as a signal for photoreceptive

63 herbivorous protists. This could serve as an additional sensory response used in finding photosynthetic prey and could aid the predator in finding patches of prey when chemical signals are insufficient. Chemical cues from prey will be limited by the rate of diffusion, whereas fluorescence emitting from prey would be almost instantaneous.

These results are supported by Amon and French (2004) who showed that

Ulkenia sp. zoospores were attracted to bioluminescence from Vibrio fisheri.

Furthermore, they found Ulkenia sp. phototaxis toward blue and white light to peak at 3.5 µE/m2/s, a similar finding to results reported in our current study. The Amon and French study along with results from our study could point toward a new area of focus within the broader topic of feeding behavior at the microbial level. The results from these two studies suggest photo detection systems in predatory microbes may be used for more than sensing the surrounding environment, and could be used to aid in prey location.

Two different rhodopsin proteins were recovered from the O. marina strain used in this study. There are seven rhodopsin genes from O. marina in the NCBI database (www.ncbi.nlm.nih.gov). This suggests either that we were not successful in isolating all O. marina rhodopsin proteins, or that O. marina differentially expresses rhodopsin proteins depending on conditions. Or perhaps different strains of O. marina contain different suites of rhodopsin genes. The rhodopsin genes available in the NCBI database are all from strain CCMP 1795, whereas in our study

O. marina strain CCMP 604 was used Further studies are needed to valuate if

64 variations in culture conditions (i.e. light levels, light cycles, and wavelength) contribute to expression of different rhodopsins.

Rhodopsins are found in eukaryotic and prokaryotic microbes (Spudich

2006), therefore it is not suprising that rhodopsin genes are found in O. marina. A

BLAST search using the rhodopsin proteins from O. marina identified in our study as a query revealed they were closely related to rhodopsin from the dinoflagellate

Pyrocystis lunula. However, unlike other rhodopsins, the next most closely matched proteins were rhodopsins from prokaryotes rather than rhodopsins present in other eukaryotes. There is evidence that Pyrocystis lunula rhodopsin is a result of lateral transfer from Bacteria (Ruiz‐Gonzalez and Marin 2004;

Sharma et al. 2006). Since the rhodopsin genes of O. marina are also within the

Bacterial rhodopsin , it maybe that the acquisition of the rhodopsin gene occurred early in dinoflagellate evolution. If so, then other dinoflagellate rhodopsins should also fall into this group.

Results from this study suggest that cAMP signaling is involved in transmitting a light stimulus signal in O. marina, which supports the involvement of rhodopsin signaling. Most likely there are multiple signal transduction pathways involved in downstream signaling following a light stimulus in O. marina.

It has been well over a hundred years since researchers began seriously studying microbial photoresponse. Verworn (1889) wrote, “we find that among unicellular organisms, so far as they are irritable at all to light, phototaxis is a wide‐ spread phenomenon.” Since then study of light detection in photosynthetic protists

65 has made large advances (for reviews: Hegemann 2008, Kateriya et al 2004), however photosensory mechanisms and behavior in heterotrophic protists have received less attention. Light is an important factor affecting the marine ecosystem and will have an affect on the behavior of any organism containing a photo‐ detection system. This will likely be linked to behavioral changes including circadian rhythms, position in the water column, and possibly feeding behavior.

Although evidence from this study points to O. marina using rhodopsin receptors to respond to light, other photoreceptors could also be involved.

4.4 What does this mean for the field of aquatic microbial ecology?

The results presented in this thesis build upon previous studies. However there is still a lack of information in this area of research. The biochemical signaling occurring within a single celled predator is complex and is no doubt integrated into the behavior of the cell. Until the cellular processes controlling protist behavior are better comprehended there will be gray areas in the understanding of aquatic microbial ecology. Bulk sampling and counting of field samples, and incubation experiments can provide information on species composition, cell abundances, and grazing rates, but they offer little or no information on why certain species are being grazed or why some species are found in higher abundances at specific depths. To make sense of field measurements, it is necessary to have a better understanding of how protist cells (and other microbes) sense and respond with behavioral changes to environmental stimuli and potential prey. This gap in information has existed for

66 too long and only relatively recently has available technology allowed for this to become widely studied.

Prior efforts to elucidate the role of biochemistry in prey selectivity by protists in marine systems have also included: evaluation of the role of cell surface electrostatic charge or hydrophobicity (Hammer et al. 1999, Monger et al. 1999,

Matz and Jurgens 2001); role of prey nutritional status (Gonzalez et al. 1993, Flynn et al. 1996, Martel 2006); evaluation of relative ingestion rates of artificial particles coated with various chemicals (Pace and Bailiff 1987; Matz et al. 2002); and the phylogenetic affiliation or phenotypic characteristics of bacterial cells (Van Hannen et al. 1999, Jurgens et al 1999, Simek et al. 1999, Suzuki 1999, Jurgens and Matz

2002, Matz et al. 2002). Martel (2006) found that chemosensory response and prey ingestion in the heterotrophic dinoflagellate Oxyrrhis marina did not appear to be closely coupled, and suggested that the biochemical mechanisms of chemosensory behavior and of prey capture may differ. Studying the response of a herbivorous protist to thin layers of phytoplankton, Menden‐Deuer and Grumbaum (2006) reported that the swimming speed of O. marina increased significantly in response to intact cells, but not in response to I. galbana exudates.

Behavioral responses elicited by receptor‐ligand binding on the cell surface will depend on signal transduction mechanisms in the protistan cell, analogous to the better‐known biochemical signal mechanisms of bacteria and of the cells of multicellular organisms (Alberts et al. 2002, Rogales 2005). As an example of the universality of this mechanism, chemical signal receptors on the surface of protistan

67 membranes are known to bind compounds, including insulin and opiates, which act as signaling hormones in metazoans (Renaud et al. 1995). Once a signal molecule binds to a receptor site, the release of secondary messenger compounds within the cell is triggered, which then may result in a change in motility, or in initiation of phagocytosis (Van Houten 1994, Renaud et al. 1995). As a demonstration of such a signaling mechanism in protists, Leick et al. (1997) were able to eliminate chemosensory behavior of Tetrahymena by exposing the ciliates to protein kinase inhibitors. Leick et al. (1997) showed that addition of inhibitors of tyrosine kinase did not disrupt swimming in the ciliates, but did block chemosensory response of the ciliates to a chemo‐attractant, proteose peptone. Limited evidence already exists that free‐living flagellated protists have such signaling systems: there is molecular genetic evidence for the presence of a protein kinase in a photosynthetic (Kiriyama et al. 1999), and for a receptor for tyrosine kinase in choanoflagellates (King & Carroll 2001).

4.5 Future studies

As more genetic information becomes available for free living phagotrophic protists, it will be possible to see what specific genes, encoding for signaling pathway components, are included in each species’ . This will also allow researchers to design experiments that make use of gene knockout or knockdown techniques using RNA interference. In this way, an experiment could be designed that either changes or reduces the expression of a protein involved in cell signaling, either at the receptor level or downstream signal transduction. Following gene

68 knockdown the behavioral response of a marine protist could be evaluated in response to environmental stimuli (e.g. light), or in response to potential prey items.

More work needs to be done on predator/prey contact, since this is likely to be the point when prey will be accepted or rejected by the predator. Wootton et al.

(2007) have shown a cell surface mannose‐binding lectin to be directly involved in prey recognition by O. marina. It is possible that the matchup of lectins and sugars on the cell surface of predators and prey determines if the prey is accepted or rejected. Lectins present in the cell membranes of phagotrophic protists could be investigated using the same approach used in Hartz et al. (ms in preparation) to detect rhodopsins in the cell membranes of O. marina: extraction of cell membrane proteins,isolation by SDS‐Page gel electrophoresis, and mass spectroscopic identification. This could allow for a ‘survey’ approach to screening protists and their prey for lectin molecules.

Additional experiments could be developed to better simulate “field” conditions. For example, studies could be designed to determine if the phytoplankton autofluorescence under blue light conditions results in a higher prey ingestion rate by O. marina. Experiments could be set up in large tanks (or columns) to investigate if O. marina is orienting to an optimum light level in the water column to find prey that might also be attracted to a similar light level.

O. marina appears to be widely distributed in coastal habitats ranging from tidal pools to open coastal water and exhibits a high degree of genetic and functional diversity (Lowe et al. 2005). Future studies could use strains of O. marina collected

69 from different habitat types and determine if they respond differently to a variety of stimuli and prey type.

More work can be done to extract and identify chemical and photoreceptors from O. marina. A range of O. marina strains could be used to determine if the expression of receptors changes when the cells are grown in different culture conditions and on different prey. These experiments could also be done with other protist species. In general, there is much work to be done to investigate the role of cell signaling in the ecological behavior of phagotrophic protists.

4.6 Closing remarks

Phagotrophic protists have essential roles in marine systems, as consumers of biomass at the base of food webs and as regenerators of inorganic nutrients from the organic matter of their prey. Previous laboratory studies using model cells have provided limited information on how biochemical receptor binding and intracellular signaling mechanisms might regulate protistan behavior and predator‐prey interactions in microbial food webs under natural conditions. In addition, very little is known about the role of photosensory mechanisms in heterotrophic protists and how they might play a part in ecological interactions and more specifically feeding behavior. Understanding the importance of these mechanisms, and to what extent they affect feeding behavior in natural microbial food webs is essential to understanding the functional ecology of phagotrophic protists. As this field of study makes advances and more information relating to biochemical signaling in free‐ living protists becomes available, it may give researchers new tools and approaches

70 in the study of in situ grazing rates and prey selectivity by marine protists.

71

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