By Sarah Jaffri the Psychrophilic Green Alga, Chlorella BI Sp. Was

ABSTRACT

CHARACTERIZATION OF THE PHOTOSYNTHETIC APPARATUS OF CHLORELLA BI SP., AN ANTARCTICA MAT ALGA UNDER VARYING TROPHIC GROWTH STATES

by Sarah Jaffri

The psychrophilic green alga, Chlorella BI sp. was isolated from a transient Antarctic pond as part of a mat consortium. Previous research on Chlorella BI sp. showed that the organism was able to utilize inorganic and organic forms of carbon and alter its photosynthetic apparatus in response to varying trophic growth states. Based on these early results, the goals of this thesis project were to: (1) characterize the photosynthetic apparatus of Chlorella BI sp. under different trophic states in comparison to the mesophilic species, Chlorella vulgaris; and, (2) determine the effect on the photosynthetic apparatus of Chlorella BI sp, when it is shifted from dark to light conditions. Chlorella BI sp. grew exponentially under the three tested trophic states. The photosynthetic apparatus exhibited functional and structural alteration. It is concluded Chlorella BI sp. has retained the ability to alter its photosynthetic apparatus in response to adaptation to a variable habitat.

CHARACTERIZATION OF THE PHOTOSYNTHETIC APPARATUS OF CHLORELLA BI

SP., AN ANTARCTICA MAT ALGA UNDER VARYING TROPHIC GROWTH STATES

A Thesis

Submitted to the

Faculty of Miami University

in partial fulfillment of

the requirements for the degree of

Master of Science

Department of Microbiology

Sarah Jaffri

Miami University

Oxford, OH

2011

Advisor Rachael Morgan-Kiss Reader D.J. Ferguson Reader Gary R. Janssen TABLE OF CONTENTS

Chapter One: Introduction P. 1-11

Chapter Two: Functional and structural Analysis of the different trophic states of Chlorella BI sp. in comparison to the mesohphilic species, Chlorella vulgaris P.12-53

Chapter Three: Alterations in the photosynthetic apparatus of Chlorella BI sp. in response to a mimicked shift from polar winter to summer P.54-67

References P.68-72

LIST OF TABLES

Table 1: Effect of the addition of various organic carbon sources on the growth of Chlorella BI sp. in the presence or absence of light P. 33

Table 2: Growth kinetics of Chlorella BI sp. in the presence of different trophic conditions P. 35

Table 3: Low temperature (77K) fluorescence emission ratios of Chlorella BI sp. cells grown under different trophic conditions P. 38

Table 4. Steady state Chl a fluorescence parameters of Chlorella BI sp. under the different trophic states P. 40

Table 5: Effect of addition of various organic carbon sources on the growth of C. vulgaris in the presence or absence of light P. 45

Table 6: Growth kinetics of C. vulgaris under the different trophic states P. 46

iii LIST OF FIGURES

CHAPTER ONE

Figure 1: Transmission Electron micrographs of Chlorella BI sp., P. 9

Figure 2: Photosynthetic apparatus in oxygenic photosynthesis P. 10

Figure 3: Pulse Amplitude Modulation induction curve of Chlorella BI sp. P. 11

CHAPTER TWO

Figure 1A: Neighbor joining phylogenetic trees were constructed using the nucleotide sequence of genes rbcL (A) and psbA (B) for Chlorella BI sp. P.32 Figure 2: Growth physiology of Chlorella BI sp. under different trophic conditions. B. Glucose consumption for Chlorella BI sp. under variable trophic growth states P. 34 Figure 3: Chlorophyll a: b ratio and total chlorophyll (ng/cell) for Chlorella BI sp. under variable trophic growth states P. 36 Figure 4: Chl a fluorescence emission spectra at 77K of whole cells of Chlorella BI sp. under the different trophic states states P. 37 Figure 5: Representative room temperature Chl a fluorescence induction curves of whole cells of Chlorella BI sp. grown under variable trophic conditions P. 39

Figure 6: Representative immunoblot of soluble fraction polypeptides isolated from Chlorella BI sp. grown under each trophic state P. 41 Figure 7: Densitometry of Chlorella BI sp. soluble fraction, Rubisco (A) and ferredoxin (B) P. 42 Figure 8A: Representative thylakoid SDS-PAGE gel of Chlorella BI sp., P. 43 Figure 8B: Representative immunoblot of thylakoid polypeptide D1 P. 43 Figure 9: Densitometry of Chlorella BI sp. thylakoid polypeptide, D1 P. 44 Figure 10: Generation time of Chlorella BI sp. compared with C. vulgaris under the different trophic conditions. P. 47 Figure 11: Chlorophyll a: b ratio for C. vulgaris under the different trophic conditions. P.48 Figure 12: Total chlorophyll (ng/cell) for C. vulgaris under the different trophic conditions P. 49

iv Figure 13: Representative room temperature Chl a fluorescence induction curves of whole cells of C. vulgaris grown under variable trophic conditions P. 50

Figure14: Steady state chlorophyll a fluorescence parameters Fv/Fm, qP and qN of C. vulgaris compared to Chlorella BI sp. grown under autotrophic (A), mixotrophic (B) and heterotrophic (C) conditions P. 51

Figure 15: Densitometry of C. vulgaris immunoblots of Rubisco compared to Chlorella BI sp P. 52

Figure 16: Non-denaturing gradient gel for Chlorella BI and C. vulgaris. 2nd dimension gel analysis of the autotrophic states of C. vulgaris (A) and Chlorella BI sp (B) P. 53

CHAPTER THREE

Figure 1A: Growth of Chlorella BI sp. during the shift from dark to light P.62

Figure1B: Glucose consumption during the shift from dark to light P.62

Figure 2: Low temperature (77K) fluorescence for Chlorella BI sp. cells shifted from dark to light P.63

Figure 3: Low temperature (77K) emission ratios of Chlorella BI sp. cells shifted from dark to light P.64

Figure 4: Chlorophyll a: b ratios and total chlorophyll (ng/cell) for Chlorella BI sp. during the shift from dark to light P. 65

Figure 5: Steady state Chl a fluorescence trace analysis for Chlorella BI sp. cells shifted from dark to light P. 66

Figure 6: Steady-state chlorophyll fluorescence quenching parameters for Chlorella BI sp. during the shift dark to light P.67

ACKNOWLEDGMENTS

I thank the Department of Microbiology at Miami University and the National Science Foundation (NSF) for supporting this research. I would also like to thank my advisor, Dr. Rachael Morgan-Kiss for her support and guidance throughout this project. To my committee, Dr. Gary Janssen and Dr. D.J. Ferguson, I would like to extend my complete gratitude. I would like to thank my current and former lab mates Jenna Dolhi, Patrick Feasel, Nicholas Ketchum, Scott Bielewicz, Donald Holter, Triratana Sanguanbun, Rocky Patil, Weidong Kong, Audrey Lloyd and Alex Loomis for providing a fun working environment and making our lab a more memorable experience. I thank my parents, Saleem and Ghzala Jaffri, and siblings (Ali, Mariam and Heena) for their constant love and support throughout my Masters. As a family, we have faced a number of uphill battles but that has inevitably made us stronger. I firmly believe that everything we have given up will eventually reap us even bigger rewards. I would also like to thank Dr. Christine Weingart for believing in me as an undergraduate and inspiring me to attend graduate school. Lastly, I would like to thank Zulfiqar Haider for his constant amusement, love and support throughout my time at Miami.

DEDICATION

This thesis is dedicated to my parents, Saleem and Ghzala Jaffri. It is only a small token of appreciation for everything my parents sacrificed to give their children a proper education. For that, we will always be grateful.

-Sarah

vii CHAPTER ONE

INTRODUCTION

1.1 POLAR ENVIRONMENTS About three-fourths of the Earth’s biosphere consists of environments that are exposed to extremely low or permanently frozen temperatures; such habitats include high alpine snowfields, deep oceans, polar sea ice, ice-covered lakes and transitory ponds (Morgan-Kiss, et al., 2006). Permanently cold environments are often dominated by microorganisms such as gram negative and gram positive bacteria, Archaea, yeast, cyanobacteria and algae. These organisms are adapted to not only surviving but thriving in cold ecosystems (Morgan-Kiss, et al., 2006).

1.2 EXTREME ENVIRONMENTS Extreme environments are considered inhospitable for most life forms because of conditions such as high temperature, ionizing radiation, pressure, low nutrients and varying pH (Rothchild and Mancinelli, 2001). Low temperature environments are examples of one extreme habitat; however, despite permanent cold, these environments vary greatly in their chemical and physical properties, influencing the survival strategies of organisms in such habitats (Morgan-Kiss, et al., 2006; Neale and Priscu, 1995). It is important to determine how organisms are able to adapt and survive to such ecosystems because it will provide insight into one of the most prevalent types of habitats in the world which are being significantly impacted by climate change (Morgan-Kiss, et al., 2006). Many cold-adapted microorganisms residing in polar habitats are psychrophilic (i.e. optimal temperature < 15°C) (Morita, 1975). Photopsychrophiles are a class of psychrophiles that are able to fix carbon by using light energy and require low temperatures for growth (Morgan-Kiss, et al., 2006). These organisms have been isolated from varying permanently cold environments such as microbial mats (Hawes and Howard-Williams, 2003), the underside of sea ice, and within the water column of

1 permanently ice-capped lakes. Like all photosynthetic organisms, photopsychrophiles use the photochemical apparatus to convert light energy into chemically-stored energy products which are ultimately used to fix inorganic carbon.

1.3 MICROBIAL MATS IN POLAR ENVIRONMENTS Microbial mats are abundant in polar environments and consist of stratified organically rich layers of microbes found on the surfaces of rocks, soil or aquatic sediment (De los Rios, et al., 2004). They are dominated by cyanobacteria and other photosynthetic organisms, which form a biological refuge for a diverse collection of other heterotrophic and chemotrophic microorganisms (Ramsing, et al., 1993; Risatti, et al., 1994). Microbial mats are vertically stratified at the level of biology and physical features (light, nutrients, redox potentials, oxygen)(Paerl and Pinckney, 1996) . Primary producers reside on the mat surface, which is considered the oxic zone of the mat, contributing to primary production and nutrient cycling in the mat. Below the photosynthetic layer, there is a steep decline in oxygen and light levels leading to anoxic conditions (Paerl, et al., 2000) Mats are considered “complete ecosystems” because of the diversity of biochemical processes (Canfield and Des Marais, 1993). To date, physiological studies on photopsychrophiles isolated from mats in polar environments have been limited. In contrast, there has been extensive work on a psychrophilic green alga isolated from an ice-covered lake in the McMurdo Dry Valleys, Antarctica. Chlamydomonas raudensis UWO241 was isolated by Neale and Priscu (Neale and Priscu, 1995) from the deepest biotic zone of the dry valley lake Lake Bonney. Due to a permanent ice-cover, algal communities in Lake Bonney are adapted to extreme shade conditions (Pocock, et al., 2004; Morgan-Kiss, et al., 1998). The organism is able to effectively use PSII for photosynthesis and has acclimated to growing at low temperature and light. However, as a consequence of the adaptation to constant shade conditions and low temperatures, C. raudensis has a limited ability to acclimate to variable environments, and appears to be an obligate photoautotroph.

1.4 CHLORELLA BI SP., A MAT ALGA Recently a green algal species was isolated from a microbial mat in “Fresh Pond”, a transitory pond on Bratina Island located on the McMurdo Ice Shelf (Morgan-Kiss, et al., 2008). This organism was initially isolated on a dilute nutrient media in the dark and was identified by producing green colonies on the agar plates, which suggested that photosynthesis is dispensable in this alga. Phylogenetic analyses of the 18S nuclear and 16S plastidal rRNA sequences identified this alga as belonging to the Chlorella clade and the Chlorellaceae family (Morgan-Kiss, et al., 2008). This organism appears to be a new species; however, more detailed phylogenetic analyses are necessary before it can be confirmed as a new species. It is a single celled microbial eukaryote, non-flagellated and 2 to 10 μm in diameter. Prominent starch granules were observed in the cells with photosynthetic membranes transversing the granules, morphological characteristics typical of Chlorella species (Fig. 1A) (Morgan-Kiss, et al., 2008). Interestingly, it also shares morphological features such as fibrils with a closely related genus, Micractinium (Morgan-Kiss, et al., 2008) (Fig. 1B). Unlike the invariable environment inhabited by C. raudensis, Chlorella BI sp. faces capricious environmental extremes such as high photosynthetically active radiation (PAR), ultraviolet light (UV), and desiccation events (Hawes, et al., 2001). In addition to extreme seasonal variation between permanent winter darkness and 24 hour sunlight in the summer, the pond microbiota are also exposed to freeze and thaw cycles at the beginning and ends of winter and summer seasons, respectively. In addition, during the growing season, multiple environmental factors can vary including light, UV, nutrients, organic carbon, and water availability. These additional environmental extremes play a role in restricting growth of phototrophic microorganisms in Antarctic environments. It is likely that phototrophic microorganisms residing in the mat community have adapted unique strategies to survive and reproduce in this extreme niche; however, detailed physiological studies are currently lacking. One way that the pond psychrophile Chlorella BI sp. may survive its highly variable environment is by its ability to utilize alternative carbon and energy sources when light is limiting. In a recent study, Morgan-Kiss et al. (Morgan-Kiss, et al., 2008) reported that Chlorella BI sp. was able to grow under variable

3 trophic growth regimes (i.e. the ability of an organism to utilize variable sources for energy and carbon acquisition). Under autotrophic conditions, the organism is able to produce organic carbon by fixing inorganic carbon. This process utilizes light as the energy source which drives photosynthetic electron transport creating NADH and ATP. Under a mixotrophic regime, the alga supplements photosynthesis with oxidation of organic carbon compounds. Most studies suggest that the two processes of photosynthesis and glucose metabolism work independently of one another (Ogawa and Aiba, 1981). However, the addition of an alternative organic carbon source can inhibit the synthesis of the photosynthetic apparatus if there is accumulation of the organic carbon within the cell (Usada and Edwards, 1982). Lastly, heterotrophic ability refers to relying solely on an organic carbon source for its carbon and energy supply. At the level of carbon and energy acquisition, Chlorella BI sp. exhibits the ability to grow under autotrophic, mixotrophic and heterotrophic modes of growth (Morgan-Kiss, et al., 2008). This ability to grow under variable trophic growth regimes is in contrast with the Antarctic lake alga C. raudensis, which is a strict photoautotroph (Morgan-Kiss, et al., 1998). Thus, while both organisms are photopsychrophiles, it is evident that there are physiological differences between the two Antarctic algae that reflect adaptation to different natural Antarctic habitats. An ability to utilize variable carbon and energy sources would be an adaptive advantage in the extreme seasonal light environment of Antarctica and may be accompanied by alterations in the composition of its photosynthetic apparatus when it is shifted from darkness (i.e. the polar winter) to light (i. e. the polar summer). One phenomenon associated with plants and some algae is the process of “greening” which involves massive restructuring of the photosynthetic apparatus when organisms shift from dark to light conditions. Greening under low temperatures (i.e. shifting from darkness to light at temperatures below 10°C) is particularly stressful for photosynthetic organisms that are adapted to moderate temperatures due to the production of ROS and subsequent oxidative damage. Presumably, Antarctic photosynthetic organisms may go through a greening process yearly during the transition from Antarctic winter to summer; however, there are currently no studies documenting this phenomenon in cold-adapted algal species. The physiological effect of greening under low temperatures in temperate organisms can cause major alterations in

4 photosynthetic electron transport components, xanthophyll accumulation, LHCII polypeptides (Maxwell, et al., 1994; Wilson and Huner, 2000). The transition mimicked by the different greening conditions has significant impacts on the redox state of the intersystem electron transporter, plastoquinone (PQ), which generates the trans-thylakoid pH gradient. The PQ pool is located between photosystem I and photosystem II and acts as an important sensor in electron transport. It shuts down the photosynthetic apparatus when it senses an imbalance in the electron pool (Huner, et al., 1998) Wilson and Huner et al., demonstrated that Chlorella vulgaris, when shifted from low to moderate temperatures turns from yellow to green due to the accumulation of more chlorophyll and light harvesting antenna polypeptides. This change was the result of the oxidation of the PQ pool and the decrease in the trans-thylakoid pH gradient (Wilson and Huner, 2000) allowing in the organism to absorb light energy and use the products of photosynthesis for growth.

1.5 LIGHT ENVIRONMENT Polar algae are often subjected to highly variable light regimes, depending on their habitat. For example, organisms isolated from under ice communities are exposed to only 1.8% to 3.3% surface irradiation (Lizotte and Priscu, 1992). Levels of Photosynthetic Active Radiation (PAR) and ultra violet radiation become greatly reduced in water columns of under ice communities. Algae in such habitats have evolved their photosynthetic apparatuses to efficiently capture light energy under these extreme shade conditions (Morgan-Kiss, et al., 2006). However, algae found in seasonally open water lakes, ponds, sea ice and coastal regions can experience variable periods of high levels of irradiation during their growth season. They have adapted different mechanisms in their natural habitats such as surface mucilage sheaths and internal UV absorbing compounds or pigments which function to protect the cytoplasm and chloroplast from the intense light. For example, the unicellular freshwater green alga, Micrasterias denticulata, produces large amounts of mucilage which protects the cells from UV radiation and extreme light (Oertel, et al., 2004). Another Antarctic algae Scenedemus sp. found in fresh lakes and ponds has protective UV screening compounds which protect the photosynthetic apparatus. It further copes with extreme light by having enhanced

5 replacement of the damaged D1 protein or Rubisco and repair of DNA damage (Lesser, et al., 2002).

1.6 PHOTOSYNTHESIS Photosynthesis is a process of converting light energy into chemical energy and takes place in plant chloroplasts, eukaryotic algae, and photosynthetic prokaryotes. The overall process is the light-driven splitting of a water molecule which is used in an electron transport chain to produce reducing equivalents (NADPH) and to build up a proton motive force to synthesize ATP. These energy products are then used to fix inorganic carbon in the Calvin Cycle. In oxygenic photosynthesis, there are two types of light-driven reaction centers within the photochemical apparatus, Photosystems II and I (PSII and PSI, respectively). These heterodimeric complexes are embedded in the photosynthetic membranes called thylakoids (Fig. 1). Electron transport between PSII and PSI are connected by the cytochrome b6f complex and two mobile electron carriers, plastoquinone and plastocyanin (Fig. 1) A system of light-harvesting polypeptides is associated with each photosystem respectively which gather solar energy and transfer it to the reaction centers in order to drive electron transport. The complexes are made of an outer antenna consisting mainly of chlorophyll a/b binding proteins, which bind to chlorophyll a and b (Durnford, 2003). The pigments harvest light energy and transfer it to an inner antenna (CP43 and CP47), which connects the outer antenna to the reaction center. The proteins in the reaction center are primarily responsible for charge separation and electron transfer to the primary quinone accepter QA. (Bacon, 2001. Kluwer Academic Publishers; Bacon, 2001. Kluwer Academic Publishers).

1.8 CHLOROPHYLL FLUORESCENCE Chlorophyll fluorescence can be used as a tool to measure the photosynthetic performance of an organism. A variety of tools have been developed that measure various aspects of Chl fluorescence. Pulse amplitude modulated (PAM) fluorescence monitors in vivo changes in photochemical function at the level of Chl-a fluorescence associated with PSII (Butler, 1978). On the other hand, low temperature fluorescence (77K) fluorescence

6 can be utilized to monitor both PSII and PSI fluorescence yields and is useful for determining the energy distribution. With regarding to the theory of PAM fluorometer, in a dark adapted state, the PSII reaction centers are “open” indicating that they are able to perform photochemistry (specifically they are able to reduce the quinine, QA). The minimal level of fluorescence

(F0) is established when the dark adapted state are exposed to a weak modulated measuring beam of light (Fig.3). After reaching F0, the “open” PSII reaction centers are exposed to a short high intensity beam of light which theoretically “closes” all reaction centers and gives a maximal level of fluorescence (Fm) (Fig.3). The difference between

Fm and F0 (ΦFm-ΦF0) is defined as variable fluorescence, Fv. The maximum quantum yield of PSII photochemistry is given by the parameter Fv/Fm (Fig.3) (Butler, 1978). Quenching is any process that competes with fluorescence for energy emitted from excited Chl molecules. It is expressed as a coefficient (q) and is measured in the light adapted state which occurs when actnic light (light that drives photosynthesis) is turned on. qP is the photochemical quenching coefficient and determines the relative proportion absorbed energy that is transferred to the photochemical apparatus. In contrast, qN represents the loss in fluorescence due to alternative quenching process such as heat (Butler, 1978).

7 1.10 THESIS OBJECTIVES Field-based studies regarding the ecosystems of Antarctica and the microbial communities that dominate them are relatively well studied for some habitats such as the dry valley lakes of the Taylor Valley. However, isolates of organisms residing in such environments have been poorly characterized, especially the primary producers like the photopsychrophiles. Understanding cold adaptation of photopsychrophiles such as Chlorella BI sp. will provide novel information on how the photosynthetic apparatus have evolved to deal with the combination of light and permanent low temperatures. Furthermore, the ability of this photopsychrophile to exploit alternative trophic abilities such as phototrophy, mixotrophy and heterotrophy will provide a better understanding of how microorganisms balance carbon and energy acquisition in extreme environments. The overall goal of this project is to characterize alterations in the photosynthetic apparatus of a psychrophilic microalga under different trophic states and to understand the effect of low temperature adaptation on major structural and functional changes of the photochemical apparatus.

Two aims will be addressed:

Aim 1: Effect of trophic mode on the function of the photosynthetic apparatus in the psychrophilic Chlorella BI sp. compared with the temperate alga, Chlorella vulgaris.

Aim 2: Kinetics of functional changes in the photochemical apparatus in response to a shift from heterotrophy to mixotrophy under low temperatures in Chlorella BI sp.

Figure 1. Transmission electron micrographs of Chlorella BI sp. A. Cells were grown under mixotrophic conditions. B. Cell wall of Chlorella BI sp., decorated with fibrils.

Figure 2. Photosynthetic apparatus in oxygenic photosynthesis (Morgan-Kiss unpublished). Membrane bound light harvesting complexes (LHCs) absorb light and transfer energy to major reaction centers PSII and PSI. NADPH and ATP are the final products of the light dependent reactions.

Figure 3. Pulse amplitude induction curve for Chlorella BI sp.,

CHAPTER TWO

FUNCTIONAL AND STRUCTURAL ANALYSIS OF THE DIFFERENT TROPHIC STATES OF CHLORELLA BI SP. IN COMPARISON TO THE MESOHPHILIC SPECIES, CHLORELLA VULGARIS

12 INTRODUCTION Trophy or trophic state indicates the nutritional requirements for an organism in a given environment. This state can be variable depending on fluctuations in the organism’s natural habitat, such as light, CO2 and organic carbon availability. Microbial mats have strong fluctuations in a variety of environmental parameters, including nutrient and light availability (Hawes, et al., 2001). Since Chlorella BI sp. is a mat-dwelling species, it seems likely that this photopsychrophile has developed an adaptive advantage by possessing multiple trophic abilities. In support of this prediction, Morgan-Kiss et al. showed that Chlorella BI sp. exhibited the ability to utilize various sources of carbon and energy which allowed cultures to grow under autotrophic, mixotrophic and heterotrophic conditions (Morgan-Kiss, et al., 2008). This trophic versatility differentiates Chlorella BI sp. from C. raudensis UWO 241 which is a strict photoautotroph and thus requires light for growth (Morgan-Kiss, et al., 1998). Other Chlorella species such as Chlorella vulgaris, a mesophile, can also grow under varying trophic growth conditions (Griffiths, et al., 1960) Karlander and Krauss, 1966). In addition, unlike many temperate algae, C. vulgaris can growth at low temperatures (Maxwell, et al., 1994), making C. vulgaris a mesophile algal candidate for comparative studies with Chlorella BI sp.

The objective of this chapter was to characterize the growth of the mat alga Chlorella BI sp. under three trophic growth conditions and to determine the effect of these growth regimes on the structure and function of the photosynthetic apparatus. In addition, some comparative experiments were performed using the mesophilic Chlorella vulgaris.

METHODS Growth conditions. Cultures of the Antarctic mat green alga Chlorella BI sp. (Morgan- Kiss, et al., 2008) and the mesophilic alga Chlorella vulgaris were grown axenically in

250 mL Pyrex tubes and continuously aerated under ambient CO2 conditions. Chlorella BI sp. was grown in Bolds Basal Medium (BBM) (Nicholas and Bold, 1965) supplemented with vitamins (B12, cobalamine, thiofolate) in autotrophic conditions (BBMv). For mixotrophic and heterotrophic growth conditions, BBMv was supplemented with 0.1 % glucose. All three cultures were grown in thermo-regulated aquaria at 8° C and autotrophic and mixotrophic cultures were grown in low light (20 µmol m-2s-1). Growth kinetics were monitored as the change in optical density at 750 nm.

The doubling times (tgen) were calculated as ln2/µ, where µ is the pseudo-first order rate constant for growth (Guillard, 1973). Glucose consumption. 50 µL from autotrophic, mixotrophic and heterotrophic cultures was collected every day for a period of two weeks and analyzed using the Glucose (HK) Assay kit (SIGMA). The decrease in absorbance at 340 nm was directly proportional to the reduction of NAD coupled to Glucose 6 Phosphate (G6P). G6P was then oxidized to 6 phosphogluconate in a reaction catalyzed by G6PDH; NAD was reduced to NADH.

Glucose+ ATP Hexokinase Glucose-6-Phosphate (G6P) +ADP G6P+NAD G6PDH 6-Phosphogluconate+NADH . Chlorophyll analysis. 1.5-2 mL samples were collected during mid log phase of the different trophic states. Cultures were centrifuged at 10,000 rpm for 20 min. Cells were resuspended in 1 mL of 90 % acetone and then bead beated 3 x 30 s. The supernatant was measured for Chlorophyll (Chl) a (chlorophyll-protein complexes associated with the reaction center) and Chlorophyll b (chlorophyll-protein complexes associated with accessory pigments) using methodology developed by Jeffery & Humphrey (Jeffery and Humphrey, 1975).

14 Thylakoid isolation. Cultures were harvested in mid-log phase by centrifugation at 3000 x g for 20 min. The pellets were flash frozen and stored at -80 ºC until required for use. The pellet was resuspended in Tricine-NaOH (pH 7.8) buffer containing 0.3 M sorbitol, 10 mM NaCl, 5 mM MgCl2, 1 mM benzamidine and 1 mM caproic acid. The suspension was passed through a chilled French Pressure Cell twice at 10,000 lb/in2. The broken cells were centrifuged at 272 x g for 5 min. The supernatant was then centrifuged at 23,700 x g for 30 min and then washed twice in a buffer containing 50 mM Tricine-

NaOH (pH 7.8), 10 mM NaCl and 5 mM MgCl2 by centrifugation at 13, 300 x g for 15 min. The pellet was then suspended again in 200 µl of the above wash buffer supplemented with 0.1 M sorbitol and 10 % glycerol and stored at -80 °C until required for SDS-PAGE and immunoblotting (Morgan-Kiss, et al., 1998). Low temperature Chl fluorescence. Low temperature (77K) Chl-a fluorescence emission spectra were collected on whole cells using the Perkin Elmer Luminescence Spectrometer (LS 50 B) (Buckinghamshire, England). Samples were collected and dark adapted at 8 ºC for 10 min prior to the measurement. Samples were flash frozen at 77 K with liquid nitrogen and scanned at the excitation wavelength of 435 nm for chlorophyll a (Morgan- Kiss, et al., 1998). A natural fluorophore, Fluoroscein (0.7 µM), (514nm) was used as an internal standard Room temperature chlorophyll fluorescence. Steady state Chl-a fluorescence was generated in live cells by using pulse amplitude modulated Chl a fluorescence detection system (Dual PAM-101, Chlorophyll Fluorescence & P700 Photosynthesis Analyzer Walz). Samples were collected from all three cultures and dark adapted for 10 min. The cultures were incubated in the dark in the presence of 4 mM NaHCO3 for 5 min prior to all measurements. Fluorescence parameters (FV/FM, qP, qN) (Butler, 1978) were calculated from the induction curve using proprietor software. Fv/Fm measured the maximum efficiency of PSII photochemistry. qP measured the oxidation state of the primary quinone acceptor QA of PSII. qN estimated the loss of fluorescence due to non- photochemical quenching. SDS-PAGE and immunoblotting. The solubilized membrane or soluble fractions were loaded on equal chlorophyll or protein basis (3 μg of either total protein or chlorophyll per lane) and separated on a SDS-PAGE gel. Gels was transferred to methanol activated

15 PVDF membranes and blocked for 1 h in a Tris-Buffered-Saline (TBS)/Tween/5 % non fat milk solution. The membranes were then probed with 1º antibodies raised against the thylakoid polypeptide (Agrisera Vännäs, Sweden) PsbA (1:5000 dilution) or soluble fraction polypeptides RbcL (1:5000) or Ferredoxin (1:1000) (Agrisera Vännäs, Sweden). After incubation with the primary antibody, PVDF membranes were washed four times with TBS/Tween solution at 15 min intervals at room temperature by shaking vigorously. The blots were challenged with Protein A conjugated to HRP (1:10,000 dilution) (SIGMA) for 1 h. The PVDF membranes were washed 4 times with TBS/Tween and then incubated with ECL chemiluminescent detection agents. The complex of the secondary + cleaved ECF substrate was detected on X-Ray film (Kodak) (Morgan-Kiss, et al., 1998). Blue Native PAGE gels Mid-log phase cultures (3 L) of Chlorella BI and C. vulgaris were harvested by centrifugation. These pellets were washed with cell wash buffer (0.3 M sucrose, 1 mM MgCl2, 25 mM Hepes, pH 7.5 KOH) and centrifuged again at 2500 x g for 5 min and resuspended in 50 mL of the same buffer. The cells were broken by passing through a chilled French Press at 12,000lb/in2 three times. The homgenate was centrifuged at 20,000 x g for 20 min and the pellets were resuspended in 30 mL of thylakoid unstacking buffer (0.3 M sucrose, 10 mM EDTA, 5 mM Hepes, pH 7.5 KOH). Broken cells were centrifuged at 10,000 x g for 10 min and resuspended in 4.8 mL of 1.8 M sucrose resuspension buffer (1.8 M sucrose, 10 mM EDTA, 5 mM Hepes, pH 7.5 KOH). The total volume (4.8 mL) was separated into ultracentrifuge tubes and overlaid with 2 mL of 1.3 M sucrose middle layer buffer (1.3 M sucrose, 10 mM EDTA, 5 mM Hepes, pH 7.5 KOH). This layer was then overlaid with 5 mL of 0.5 M sucrose top layer buffer (0.5 M sucrose, 5 mM Hepes, pH 7.5 KOH) and centrifuged at 100,000 x g for one hour at 4 °C. Chlorophyll bands were collected and pooled. The layers were diluted with 3 volumes of thylakoid membrane buffer (10 mM EDTA, 5 mM Hepes, pH 7.5 KOH) and centrifuged at 20,000 x g for 10 min at 4°C. The pellet was dried and 400 µL of Amino Caproic Acid (ACA) buffer was added. For each sample, chlorophyll was measured and the final concentration was adjusted to 1 mg Chl/mL. The membranes were solubilized by adding 1 % (w/v) β- dodecyl maltoside (DDM) detergent and stirred gently on ice for 25 min. The samples

16 were centrifuged at 16,000 x g for 15 min to remove unsolubilized materials and the supernatant was collected. DDM was added to a final concentration of 1 % (10 µL [10 % DDM]/100 µL sample), 10 µL of sample buffer and 5 µL of coomassie blue additive was added to the samples. 23 µL of the samples were run on a gradient gel (Invitrogen) with 10 µL of the high range marker (Fermentas SM 0671) at 200 V for 45 min and then stained with Coomassie overnight. For the second dimension, lanes were cut out of unstained gradient gels and incubated in 5 mL denaturing solution (20 % SDS and β-mercaptanol) for 10 min at room temperature and for 3 min at 50 °C. The strips were rinsed in water to remove excess β- mercaptoethanol, placed horizontally across 15 % SDS-PAGE gels and then covered with a layer of stacking gel. The gels were run at 90 V for 1 h and then stained with Coomassie overnight.

17 RESULTS

Phylogenetic analysis of photosynthetic genes of Chlorella BI sp. Phylogenetic trees were constructed of rbcL and psbA genes for Chlorella BI sp. Neighbor joining analysis indicated that the rbcL gene clusters with uncultured organisms isolated from Lake Bonney (Fig. 1A). The psbA gene clustered well with that of Chlorella elliposida, a psychrotolerant chlorophyte (Fig. 1B) species which prefers to grow at low temperatures of 15 ºC and 20 ºC and low salinity (Cho, et al., 2007).

Growth of Chlorella BI sp. on different carbon sources Preliminary characterization of Chlorella BI sp. on BBMv plates indicated that the organism could grow on plates supplemented with different organic carbon sources in the presence of light (Table 1). Highest growth was observed in the presence of light (mixotrophic) or dark (heterotrophic) in the presence of glucose. The alga was also able to grow in the absence of an organic carbon supply if light was present, but had comparatively better growth in the presence of an organic carbon source (Table 1). Cultures of Chlorella BI sp. grew exponentially in liquid culture under all three trophic growth regimes (i.e. photoautotrophic-light/CO2; mixotrophic-light/glucose; heterotrophic-dark/glucose) (Fig. 2A). However, dark grown cultures exhibited a significant reduction in final growth yield and rate (Table 2) compared with photoautotrophic and mixotrophic conditions. At the level of glucose consumption, heterotrophic and mixotrophic-grown cultures exhibited comparable rates of glucose removal from the growth medium, with all measurable organic carbon being consumed in the first 100 hrs of growth (Fig. 2B). Once glucose had been exhausted in mixotrophic conditions, there was a shift in growth mode from mixotrophy to autotrophy. This allowed the organism to have a comparable growth phenotype to that of photoautotrophic conditions (Fig.2A). This shift was not present in heterotrophic conditions as once the glucose had been depleted, the organism entered stationary phase.

18 Chlorophyll analysis of Chlorella BI sp. in each trophic state

Chlorophyll a: b and total chlorophyll (ng/cell) were measured in cultures of Chlorella BI sp. grown under each trophic state. The Chl a: b ratio increased in the mixotrophic (●) and heterotrophic cultures through out growth (▲) compared to the autotrophic state (■) which remained relatively unchanged (Fig. 3A) Total chlorophyll (ng/cell) increased throughout growth in the autotrophic conditions and decreased in both the mixotrophic and heterotrophic conditions (Fig. 3B). In mixotrophic conditions, once the glucose had been exhausted from the culture, there was an increase in total chlorophyll (ng/cell). Trophic mode and functional adjustments in the photochemical apparatus

As the presence of an organic carbon supply can alter the requirement for photochemically derived energy (Ogawa & Aiba, 1981) energy partitioning between Photosystem II (PSII) and Photosystem I (PSI) was measured by Chl-a low temperature (77 K) fluorescence spectral analysis in cultures grown under the three different trophic states. This analysis was used to determine the in vivo light energy distribution between the photosynthetic complexes which reflects the functional organization of the photochemical apparatus (Krause and Weis, 2001). Regardless of growth regime,

Chlorella BI sp. exhibited well defined fluorescence peaks at F683 and F715 corresponding to the pigment protein complexes, PSII and PSI respectively (Fig. 3). The intensity of peaks varied between each trophic state; most notably, fluorescence yield was greatly reduced in mid log phase of heterotrophically-grown cells compared with either mid log phase of mixotrophic or autotrophic cultures. On the contrary, LHCII: PSII fluorescence

(F683/F695) ratio for mixotrophic and heterotrophic cultures was 6% and 20% higher compared with autotrophically grown cultures (Table 3). In addition, the ratio of the PSI:

PSII (F715/F695) was 24% and 22% higher for autotrophic and mixotrophic cells compared to heterotrophic cells (Table 3).

19 Steady state Chl a fluorescence parameters for Chlorella BI sp. under the different trophic conditions Light energy absorbed by chlorophylls associated with PSII is released as energy when excited chlorophyll molecules relax through various processes, including photochemistry, heat loss or chlorophyll fluorescence (Butler, 1978). The relative state of the PSII photochemistry can be monitored by the Chl-a fluorescence parameters, Fv/Fm, qP and qN. Fv/Fm (φFm-φFo/φFm) is the measure of the maximum efficiency of PSII photochemistry. Higher plants and mesophilic algae have Fv/Fm around 0.8 (Bjorkman and Demming, 1987). qP is the measure of the oxidation state of QA; an estimation of how many “open” PSII reaction centers are present in the cell. qN (Fm/Fm′-1) is the estimation of non photochemical quenching. This measurement indicates how much energy is being lost to energy dependent quenching (qE), photo inhibitory quenching (qI) and state transition quenching (qT) (Krause and Jahns, 2004). Multiple environmental stresses and changes in growth regime have been shown to affect all of the above measures of PSII photochemistry. Typical steady state Chl a fluorescence traces are shown for each trophic state in Figure 4. Autotrophic and mixotrophically-grown cells exhibited a typical induction curve, whereas heterotrophically-grown cells exhibited a strong quenching phenomenon following the saturation pulse flash (which generates Fm).

In mid log phase (100-150h) steady state Chl a fluorescence parameter, Fv/Fm was 0.592 and 0.608 respectively for autotrophic and mixotrophic cultures; however Fv/Fm was significantly lower in heterotrophically grown cells (Table 4). In contrast, QA was comparable between cells grown under each trophic state indicating the oxidation state is relatively similar for each state (Table 4). qN values for autotrophic and mixotrophic conditions were higher compared to the heterotrophic state Effects of Trophic Growth Regime on Photosynthetic Protein Abundance Functional analyses of Chlorella BI sp. cultures indicated alterations to the photochemical apparatus that were dependent on the trophic growth regime. Subsequently, structural analyses were conducted to determine if the functional changes correlated with alterations in abundance of major photosynthetic proteins. Proteins isolated from the soluble fractions of Chlorella BI sp. grown under the different trophic states were separated with SDS-PAGE. The abundance of RbcL, the large subunit of the

20 enzyme RubisCO, and ferredoxin, the terminal electron acceptor in the photosynthetic electron transport chain were quantified by western blot analysis (Fig. 5). Relative abundance (estimated by densitometric analyses) indicated that autotrophically- grown cultures exhibited ~30% higher level of RubisCO compared with cultures grown under either mixotrophic or heterotrophic conditions (Fig. 6A). However, heterotrophically- grown cultures exhibited 1.55 to 2-fold higher ferredoxin levels compared with light- grown cultures (ie. autotrophic and mixotrophic; Fig. 6B). Major membrane-bound photosynthetic proteins were also quantified from membrane fractions isolated from cultures grown under the three trophic growth regimes. Proteins isolated from the thylakoid fractions were loaded on the basis of chlorophyll or protein and were separated by SDS-PAGE to monitor any changes in the thylakoid membrane protein profiles (Fig. 7A). Cultures grown under mixotrophic conditions exhibited reductions in several membrane proteins compared with those grown under either heterotrophic or photoautotrophic conditions. The presumed protein, D1, because of the respective mass (31kdA) was less abundant in the mixotrophic state compared to the other two states. There is a notable absent band at ~40 kDa in the mixotrophic state. Further experiments must be conducted to determine the identity of the band. D1 was quantified by western blot analysis (Fig. 7B) and exhibited a reduction in levels in both mixotrophic and heterotrophic relative to photoautotrophically-grown cells (Figs. 7B and 8).

Trophic growth regime effects in a mesophilic alga The effect of growth regime under low growth temperatures was tested in the mesophilic alga, Chlorella vulgaris, to determine whether the responses observed in Chlorella BI sp. were associated with its psychrophilic nature. Similar to Chlorella BI sp., growth of C. vulgaris on plates in the presence of various organic carbon sources confirmed that it was able to grow heterotrophically at low temperatures and glucose is a preferred carbon source (Table 5). However, the response of the mesophile to low growth temperature in liquid culture revealed differences in the growth physiology between the two Chlorella spp. C. vulgaris mixotrophic cultures exhibit a higher growth rate compared to autotrophic cultures (Table 6). The generation times for autotrophic and mixotrophic cultures for both algal species were relatively similar; however, generation

21 times of the mesophilic alga were approximately 80% slower in heterotrophically-grown cultures (Fig. 10).

Chlorophyll analysis of C. vulgaris in each trophic state

Unlike Chlorella BI sp. the autotrophic state of C. vulgaris varied in the concentration ratio of Chl a: b (Fig. 11A). In the mixotrophic state, the ratio increased about 50% in the first 20 h of growth, however it remains relatively constant afterwards (Fig. 11B). In heterotrophy, the ratio was highest at 48h and then subsequently decreased (Fig. 11C). The total chlorophyll (ng/cell) was also variable for C. vulgaris. It did not change significantly in the autotrophic state (Fig. 12A) but fluctuated in mixotrophic cells (Fig. 12B). In heterotrophic cells, total chlorophyll was highest at 48h and then remained constant thereafter (Fig. 12C).

Steady state Chl a fluorescence parameters for C. vulgaris under each trophic state

Room temperature fluorescence induction curves for C. vulgaris indicated that cells grown under either autotrophic or mixotrophic conditions exhibited typical induction curves (Figs. 13 A and B). However, heterotrophic cultures exhibited an a typical induction curve with an extremely reduced fluorescence yield (Fig. 13C). In mid log phase, steady state Chl a fluorescence parameter, maximum PSII efficiency in cells of C. vulgaris grown under either autotrophic or mixotrophic conditions was comparable with that of Chlorella BI sp. grown under the same growth conditions (Fig 14A). In contrast, Fv/Fm for heterotrophically-grown cultures of C. vulgaris was 2.5-fold lower than that of Chlorella BI sp. (Fig. 14A). At the level of the oxidation state of intersystem electron transport (qP), autotrophically-grown cultures of either Chlorella species exhibited a relatively high qP, indicating that the electron pool was relatively oxidized under an autotrophic growth regime (Fig. 14B). In contrast, mixotrophically- and heterotrophically-grown cells of the mesophilic C. vulgaris exhibited a 1.5- and a 3.1- fold decrease in qP, indicating that the addition of an organic carbon source induced an

22 increase in the reduction state of the electron pool (Fig. 14B). Non photochemical quenching (qN) exhibited the opposite trend as qP in the mesophilic alga, that is qN increased in response to both mixotrophic and heterotrophic growth conditions. This trend was contrary to the response of qN in the psychrophilic Chlorella BI sp. which exhibited a reduction in qN in response to addition of organic carbon (Fig. 14C).

Structural changes in C. vulgaris under different trophic states Abundance of the major photosynthetic protein, RbcL was compared in cultures of the mesophilic C. vulgaris and the psychrophilic Chlorella BI. sp. grown under variable trophic conditions. Both species responded to the addition of glucose by decreasing RubisCO levels, relative to autotrophically-grown cells (Fig. 15).

Blue Native PAGE analysis The abundance of chlorophyll-protein holo complexes was investigated by Blue native PAGE (Fig. 16), followed by 2nd dimension SDS-PAGE. While thylakoids extracted from all three trophic states from both Chlorella species were subjected to native PAGE, only membrane complexes collected from autotrophically-grown cultures were sufficiently separated to allow for analyses in the 2nd dimension. Several major chlorophyll-protein complexes were resolved in thylakoids isolated from either species grown under autotrophic conditions, including PSI holocomplex, PSII core complex, as well as several isomers of LHCII (Fig. 16). Interestingly, Chlorella BI sp. appears to have relatively reduced levels of PSI, a phenomenon which has also been observed in the psychrophilic C. raudensis (Morgan-Kiss, et al., 1998).

23 DISCUSSION

Antarctic microbial mats are subjected to a wide variety of environmental extremes (Hawes, et al., 2001). As a result, resources within the mat greatly fluctuate causing organisms to utilize multiple nutrients for growth. In particular, Chlorella BI sp. has developed the ability to grow in different trophic states and alter its photochemical apparatus as an adaptation to the dynamic environment.

Chlorella BI sp. exhibited the ability to utilize organic carbon and displayed an increased growth in the first 100hrs of growth (Fig.1A) suggesting the alga prefers to grow mixotrophically and once it has consumed the available carbon (Fig. 1B), it switches to an autotrophic state. As a result, there are comparable growth rates in the two states; however, there was a significantly reduced growth rate in the heterotrophic state (Table 2). Chlorophyll synthesis was monitored in both Chlorella BI sp. and C. vulgaris to determine if there were changes in chlorophyll content in the different trophic states. Mixotrophic and heterotrophic cultures of Chlorella BI sp. and C. vulgaris had increased Chlorophyll a: b ratios (Figs. 2 & 11) with a substantial increase in the Chlorophyll a: b ratio in heterotrophic cultures for C. vulgaris (Fig. 11C). An increased ratio suggests decreased amounts of Chlorophyll b, an accessory protein that is found in the peripheral LHCII (Eggink, et al., 2001) and plays an important role in the insertion of LHCII proteins (LHCPs) in the chloroplast. LHCPs are imported more substantially when sufficient Chl b is available suggesting there is less LHCII present in the mixotrophic and heterotrophic cultures of the alga’s as there is less Chl b present. However, further studies need to be conducted to determine the abundance of LHCII in each trophic state.

Mixotrophic and heterotrophic growth of Chlorella BI sp. indicated it can assimilate glucose in the light as well as the dark. The rate of glucose consumption is similar in both states. However, the growth rate and final yield of mixotrophic cultures (Table 2) is higher than heterotrophic cultures due to the added presence of light suggesting the presence of two energy systems, photosynthesis and oxidative glucose metabolism increasing the growth rate of the alga in mixotrophic cultures. Unlike the psychrophilic species, C. vulgaris exhibited optimal growth in mixotrophy compared to the autotrophic and heterotrophic states (Table 6). It is unable to shift to autotrophic

24 conditions once organic carbon has been depleted like Chlorella BI sp. suggesting that the psychrophile developed the ability to grow and switch between different trophic states as an adaption to the dynamic cold mat environment compared to the ubiquitous freshwater environment of C. vulgaris. The organic rich/highly shaded interior of Antarctic microbial mats (Stal, 1995) provides the selective force for such adaptations, creating a significant advantage for Chlorella BI sp. when irradiance levels are low during the transition from polar summer to polar winter. On the contrary, C. raudensis is found in a very stable Antarctic environment. The absence of turbulence allows the organism to keep its position in the water column of Lake Bonney. As a result, it is not adapted to utilizing different carbon sources in its growth regime and subsequently cannot alter its photosynthetic apparatus in the presence of organic carbon (Morgan-Kiss, et al., 1998) illustrating a key difference between the two psychrophiles. Chlorella BI sp. exhibited the ability to alter its photosynthetic apparatus in the presence or absence of organic carbon. At the level of dynamic energy distribution between photosystems, 77K emission spectra clearly demonstrates that the light energy distribution between PSII and PSI is different when the alga is grown under mixotrophic conditions where there is preference for the excitation of PSII and PSI (Table 3). Furthermore, the energy of PSI/PSII in the heterotrophic state is greatly reduced compared to the other two states. The PSI/PSII (F715/F695) ratio being lower in this state compared to autotrophic and mixotrophic conditions (Table 3). These results suggest that PSI is likely down regulated under dark conditions. In addition, LHCII/PSII 77K fluorescence ratio is highest under heterotrophic conditions (Table 3), which may indicate that while PSII is present, LHCII might be functionally disconnected from PSII. The detachment of LHCII enables plants and algae to functionally adapt to the changes in light quality. This process requires that a pool of LHCII exist which can detach from the PSII reaction center. It is hypothesized that two populations of LHCII are present in the thylakoid membrane, one that is permanently bound to PSII and the other that is able to alter its functional association (Kyle, et al., 1983) indicating the high ratio of LHCII/PSII 77K ratio in Chlorella BI sp., is due to LHCII detaching itself from LHCII proteins that are permanently bound to the reaction center as a result of growing in the dark.

25 Steady state fluorescence parameters of Chlorella BI sp. indicated functional differences of PSII photochemistry in response to variable trophic growth regimes. The

Fv/Fm of the alga is highest in the mixotrophic state (Table 4) with reduced ratios in the autotrophic and heterotrophic state. However, the Fv/Fm of the mesophilic species is lower in each trophic state than Chlorella BI sp., (Fig. 14), implying the maximum PSII photochemistry is lower at cold temperatures for C. vulgaris and was most profoundly reduced under heterotrophic conditions. The induction curve for heterotrophic cultures of C. vulgaris (Fig. 13C) further suggests photochemistry is severely down regulated as the

Fv/Fm is the lowest in this state for C. vulgaris. This possibly is the result of the stress from growing at sub-optimal temperatures. Transient quenching where F′ dips below Fo′ in the light, suggests an enhanced ability for reduced PSII reaction centers to act as quenchers of fluorescence in Chlorella BI sp. This phenomenon is present in C. raudensis and Chlorella BI sp. cultures supplemented with glucose (Figs.4B & 4C) However, it is absent from such cultures in C. vulgaris (Figs.13B & 13C).

The absence of light in the heterotrophic culture of Chlorella BI sp. prevents reduction of the electron pool indicated by high qP values (Table 4). This suggests QA is relatively oxidized and a low qN (Table 4) indicates energy is not being dissipated through excess energy. However, C. vulgaris has decreased capacity to maintain an oxidized qP in the presence of organic carbon (Fig. 14) suggesting that the mesophile has a relatively low capacity to maintain an oxidized electron transport pool under these conditions, relative to the psychrophile. The assimilation of organic carbon for growth at low temperatures by C. vulgaris is a possibility for over reduction of the electron pool, where the pathways that metabolize glucose may use the electron pool to dispense excess electrons. Also, because C. vulgaris is more commonly found in stable fresh water environments, it may not have alternative routes of electron transport. In addition to the main route of electron flow during photosynthesis which is called linear flow or Z scheme, there are auxiliary routes that interact with the main pathway. These systems help algae to adapt to different environments and prevent the over reduction of the electron pool (Peltier et al, 2010). However, because the electron pool is more reduced in C. vulgaris than Chlorella BI sp., in heterotrophic conditions, this may suggest that the mesophile does not possess such alternate mechanisms. Low qP values for C. vulgaris

26 further suggest over excitation of PSII caused by growing at suboptimal temperatures and assimilating organic carbon simultaneously causing a modification in the redox state of the intersystem electron transport of the apparatus (Savitch, et al., 1996). The low qP values also correlate with the high chlorophyll a: b ratios for C. vulgaris (Fig. 11) in mixotrophic and heterotrophic cultures. The decrease in chlorophyll b suggests the mesophilic species down regulates its chlorophyll content as a consequence of a reduced electron pool which occurs due to growth at cold suboptimal temperature and the assimilation of organic carbon simultaneously.

Chlorella BI sp. exhibited structural differences in the mixotrophic state as it has reduced abundance of the large subunit of RubisCO (Fig. 6A), ferredoxin (Fig. 6B) and D1 (Fig. 8) compared to the other two states. The decreased abundance of photosynthetic proteins may suggest the photosynthetic apparatus is less prominent in the mixotrophic state as there might be competition between the two energy systems, photosynthesis and oxidative glucose metabolism (Marquez, et al., 1993). The decreased abundance of the photosynthetic proteins might be further attributed to the reduced number of chloroplast in the presence of glucose (Xie, et al., 2001). The two pathways might compete for electrons and as a result, there might be reduced numbers of chloroplasts in the presence of glucose. Similar results were observed in Chlorella protothecoides (Ahmed and Hellenbust, 1990) where chloroplast development and RubisCO activity were repressed in the presence of glucose. It remains to be determined whether a decrease in Rubisco activity correlates with a decrease in RubisCO abundance in Chlorella BI sp. Reduced protein abundance could further be attributed to sugar repression of photosynthetic gene expression which causes decreased protein levels (Jang and Sheen, 1994) Organic carbon such as glucose may attribute to inhibiting photosynthesis by decreasing the level of Calvin cycle enzymes and increasing the level of glycolytic enzymes (Stitt, 1991) which are important for carbohydrate metabolism and other metabolic pathways in higher plants and algae (Koch, 1996). A number of higher plant model systems such as maize (Sheen, 1990), Arabidopsis (Cheng, et al., 1992) and tobacco (von Schaewen, et al., 1990) plants all indicated repression of photosynthetic genes in the presence of such sugars as sucrose and glucose. The decrease in total chlorophyll (ng/cell) for mixotrophic and heterotrophic cultures of Chlorella BI sp. and

27 C. vulgaris (Figs. 2 & 12) further suggest chlorophyll synthesis genes and other key photosynthetic genes may be repressed in the presence of organic carbon suggesting sugar can act as physiological signals in order to control essential processes such as photosynthesis in higher plants and algae. Future experiments must be conducted in order to determine if the abundance of photosynthetic proteins increases after sugar has been depleted Also, the specific glucose uptake mechanism for Chlorella BI sp. should be investigated to further characterize the role of glucose in the mixotrophic state. Like Chlorella BI sp. there was decreased abundance of Rubisco in the mixotrophic state of C. vulgaris (Fig. 15). The low protein abundance may be attributed to the presence of an organic carbon, however, it can also be due to high excitation pressure the alga faces when it is grown in the presence of light and organic carbon implied by the low qP values compared to Chlorella BI sp. Low qP levels indicated high excitation pressure which is measured as 1-qP (Dietz, et al., 1985) and suggest a reduction of the electron pool. The PQ pool which acts as a sensor of the electron pool, could down regulate photosynthesis because of the reduced electron pool, eventually causing the inhibition of key processes such as carbon fixation. C. vulgaris may down regulate carbon fixation suggested by the decrease in Rubisco (Fig. 15) as a mechanism to protect the photosynthetic apparatus from the excess excitation pressure (Savitch, et al., 1996) Although, both Chlorella BI sp and C. vulgaris experience a decrease in RubisCO in the mixotrophic state, it is due primarily to the fact that Chlorella BI sp. is adapted to living in permanently cold environments and utilizing organic carbon in its growth regime. On the contrary, C. vulgaris is a mesophile and not adapted to growing at low temperatures, it has to acclimate itself to growing in such environments and assimilating organic carbon and as a result causes inhibition of key processes such as carbon fixation. The heterotrophic state displayed the most reduced function of the photosynthetic apparatus. Structurally it did have similar protein abundance to the autotrophic state. There was equivalent abundance of proteins Rubisco and D1. A similar Chlorella species Chlorella pyerondoisa, was analyzed in the same trophic states as Chlorella BI sp. Metabolic flux analysis indicates that 18% of the energy supplied in the heterotrophic state was converted into ATP compared to 1.5% and 1% for autotrophic and mixotrophic

28 cultures respectively. Most of the energy harnessed by heterotrophic cultures was used for structural support and maintenance of the photosynthetic apparatus (Yang, et al., 2000). Furthermore, a functioning apparatus is necessary for Chlorella BI sp., in heterotrophy. The alga might encounter other a biotic factors such as light that would require up regulation of the photosynthetic apparatus, which would explain the comparable abundance of major photosynthetic proteins found in the heterotrophic state. Another source of energy for Chlorella BI sp. in heterotrophic growth could be degradation of starch, an insoluble polymer that is produced by glucose residues. Transitory starch is one type of starch produced during photosynthesis in the chloroplasts of many higher plants. It serves as an important store of carbohydrate for periods of darkness when photosynthesis is not possible and degraded for carbon metabolism (Smith and Martin, 1993). This carbon is then transported from the chloroplast to meet the various metabolic demands of the organism. It can be hypothesized that Chlorella BI sp. could degrade transitory starch as an energy source once it has depleted the organic carbon. However, future experiments such as growth analysis and determining what types of starch are present in Chlorella BI sp are needed. Starch is also found in amyloplasts, plastids which are found in heterotrophic tissues of plants, and involved in a wide range of biosynthetic reactions including the synthesis and storage of abundant quantities of starch (Balmer, et al., 2006). The ferredoxin/Trx system was initially thought to be restricted to chloroplast of the cell. However, Amyloplasts isolated from wheat starchy endosperm have a complete ferredoxin/Trx system (Balmer, et al., 2006). Ferredoxin has been further isolated and purified from Amyloplasts. Although the function must further be elucidated, Amyloplasts have enzymes such as glucose- 6- phosphate dehydrogenase and 6- phosphate dehydrogenase that are capable of generating NADPH which then reduce ferredoxin. Compared to Rubisco and D1, ferredoxin in Chlorella BI sp. is the most abundant protein in the heterotrophic state. One possible explanation for the increased abundance of the protein in heterotrophy could be that NADPH generated enzymatically (Neuhaus, et al., 1993) could favor the transfer of electrons from NADPH to ferredoxin, allowing the protein to distribute electrons to several biosynthetic and regulatory pathways (Hanke, et al., 2004). There are other isoforms of ferredoxin besides the most

29 abundant ferredoxin (PetF) found loosely associated with PSI in C. reinhardii. These isoforms may be involved in starch synthesis (Hanke, et al., 2004) and sulfate reduction (Nakayama, et al., 2000) indicating the isoforms of FDX are differentially regulated in response to different environmental cues. The green alga, C. reinhardii has at least six different isoforms of ferredoxin including PetF and ferredoxins FDx2-FDx6 (Merchant, et al., 2006; Terauchi, et al., 2009). Further experiments must be conducted with Chlorella BI sp., to determine if there are different isoforms of ferredoxin present and under what trophic conditions these isoforms are found. Blue native PAGE is a method that was used to isolate the PSI super complex. This technique was used to isolate the PSI complex in the different trophic states of Chlorella BI sp., and C. vulgaris. The 1st dimension analysis of Chlorella BI sp. (Fig. 16) indicated a diminished green band running (~130kDA) suggesting the PSI super complex. The alga like C. raudensis has low PSI however, Chlorella BI sp. still has a PSI peak in low temperature fluorescence which is absent from C. raudensis. One explanation can be attributed to the different environments of the two organisms. The fluctuating environment of the microbial mat, in particular high irradiation in the summer months perhaps caused Chlorella BI sp. to retain PSI in order to adjust to the high irradiation. However, C. raudensis is exposed to a very stable environment year round. The absence of turbulence allows the alga to maintain its position in the water column, resulting in their exposure to a relatively stable light regime. It has increased efficiency of light absorption and utilization particularly within the blue green region of the visible spectrum. This allows C. raudensis an adaptive advantage to this extreme low light regime. However, adaptation to such an environment resulted in the loss of the PSI complex (Morgan-Kiss, et al., 1998). Of additional interest is the insolublization of the mixotrophic and heterotrophic samples for each of the respective algae preventing 2nd dimension analysis. As native PAGE gels are non-denaturing, the right concentration of DDM detergent is needed to accurately dissolve the samples. Glucose could cause insolublization of thylakoids as there is an increase in starch and lipid content in the presence of an organic carbon (Sheen, 1990) (Jang and Sheen, 1994). Future experiments must be conducted to determine the right solubilization conditions for mixotrophic and heterotrophic samples to determine protein differences in each state.

30 Chlorella BI sp. has adapted to the variable environmental conditions of the microbial mat by growing in three distinct trophic modes. It is able to grow well in all three states; unlike C. vulgaris which was able to grow in the three states but grew poorly in heterotrophic conditions as it was unable to acclimate to in the dark at cold temperatures. It can be postulated that Chlorella BI sp. acquired its ability to utilize organic carbon given its position on the microbial mat. Because of the large diversity of organisms, there is likely a release of organic byproducts by the different organisms. Cyanobacteria are the most likely producers of the organic carbon which become available to the rest of the mat through cell lysis or excretion. The alga has adapted to utilizing such byproducts in its growth regime and more importantly organic carbon may aide in the alga surviving the harsh extremes of mats. Further experiments need to be conducted to determine if it has the ability to grow on other substrates besides organic carbon when light is absent. The versatility of Chlorella BI sp. indicated by the different trophic ability and subsequent functional and structural alteration of its photochemical apparatus suggests these properties are essential for survival on Antarctic microbial mats.

Figure 1. Neighbor joining phylogenetic trees were constructed using the nucleotide sequence of genes rbcL (A) and psbA (B) for Chlorella BI sp. Bootstrap values are indicated on the left side of the trees.

Table 1. Growth of Chlorella BI sp. on solid media in the presence or absence of a light source and a variety of organic carbon sources. (+)=growth. (-)=no growth

Trophic state Organic carbon

Control Glucose Acetate Glycerol Maltose (No sugar) (1mM) (1mM) (1mM) (1mM)

Growth in light ++ ++++ +++ +++ ++ (mixotrophy)

Growth in dark - ++++ ++ + - (heterotrophy)

Figure 2. A. Growth physiology of Chlorella BI sp. cultured under different trophic conditions. Growth was measured as the change in optical density (OD750). Cultures were grown at optimal growth temperature (8° C) under autotrophic (■) mixotrophic (●) or heterotrophic (▲) conditions. Mixotrophic and heterotrophically grown cultures were supplemented with 0.1 % glucose (n=3). B. Glucose consumption for Chlorella BI sp. in autotrophic (■), mixotrophic (●) and heterotrophic (▲) states. Glucose consumption was monitored by measuring the remainder of glucose in the spent medium (n=2).

Table 2. Growth kinetics of Chlorella BI sp. grown under different trophic conditions (n=3). *Statically significant P-value<0.05 (autotrophic vs. heterotrophic) ** statically significant P-value<0.05 (mixotrophic vs. heterotrophic)

Trophic state Growth rate Generation time Final Yield (day-1) (days) (cells/mL)

Autotrophic 0.332±0.021* 2 days ±0.168 2.4x108±1.6x108

Mixotrophic 0.334±0.015** 2 days±0.121 2.3x108±7.8x107

Heterotrophic 0.289±0.021 2.3 days±0.214 5.5x107±3.7x107

Figure 3. Chlorophyll a:b ratios (A) and total chlorophyll (ng/cell) (B) in cultures of Chlorella BI sp. during growth under different trophic states (n=3).

Figure 4. Representative Chl a fluorescence emission spectra at 77K of whole cells of Chlorella BI sp. grown in autotrophic (─ ─), mixotrophic (·····) and heterotrophic (−−) conditions. The excitation wavelength was 435 nm and the emission maxima are indicated in nm at the top of each peak. The spectra were normalized to the emission peak of the internal standard, Fluorescein Sodium at 513 nm (n=2).

37 Table 3. Ratios of major 77K fluorescence emission maxima in cells of Chlorella BI sp. grown under variable trophic growth regimes (n=2)

Trophic state LHCII/PSII PSI/PSII

Autotrophic 1.28 0.99

Mixotrophic 1.35 1.01

Heterotrophic 1.6 0.77

Figure 5. Representative room temperature Chl-a fluorescence induction curves of whole cells of Chlorella BI sp. grown under variable trophic conditions.

39 Table 4. Steady state fluorescence parameters of Chlorella BI sp. during mid log phase in each trophic state (n=2).

Trophic state Pulse Amplitude Modulation (PAM) parameters for Chlorella BI sp.

Fv/Fm qP qN

Autotrophic 0.592 0.697 0.201

Mixotrophic 0.608 0.692 0.144

Heterotrophic 0.352 0.690 0.072

Figure 6. Representative immunoblots of soluble fraction polypeptides isolated from Chlorella BI sp. cultures grown under different trophic growth conditions (1-autotrophic, 2-mixtrophic 3-heterotrophic). SDS-PAGE samples were loaded on an equal protein basis of 3 µg. The gels were probed with primary antibodies raised against the large subunit of RubisCO, RbcL (A) and ferredoxin (B). Numbers on the left represent the molecular (kD) masses of markers. (n=4)

Figure 7. Quantification of RbcL (A) and ferredoxin (B) in cultures of Chlorella BI sp. grown under variable trophic conditions. Blots were loaded on equal protein basis and relative to the autotrophic state. Blots were quantified by densitometry using Scion Image software. Values were normalized to autotrophic conditions (n=4).

Figure 8. A. Representative SDS-PAGE gel separating thylakoid membrane proteins isolated from cultures of Chlorella BI sp grown under variable trophic conditions (Lanes, 1-autotrophic; 2-mixotrophic; 3-heterotrophic). Lanes were loaded on either an equal (3 µg per lane) chlorophyll or protein level. Gels were then probed with primary antibodies raised against PSII reaction center protein (31 kDA), D1 (B). Numbers on the left represent the molecular (kD) masses of markers. (n=3).

Figure 9. Quantification of D1 protein from cultures of Chlorella BI sp. grown under variable trophic conditions. Blots loaded on protein basis were quantified by Scion Image software and were relative to autotrophic state (n=3).

Table 5. Effect of addition of various organic carbon sources on BBM plates on low temperature (4°C)-grown Chlorella vulgaris in the presence or absence of a light source. Control indicates no exogenously added organic carbon source. (-) = no growth (+) = growth.

Trophic state Organic carbon

Control Glucose Acetate Glycerol Maltose (No sugar) (1mM) (1mM) (1mM) (1mM)

Growth in ++ ++++ +++ +++ ++ light (mixotrophy)

Growth in - ++++ + ++ ++ dark (heterotrophy)

45 Table 6. Growth kinetics of C. vulgaris in the different trophic states. Like Chlorella BI sp. growth rate and generation time (LN 2/µ) were calculated in the logarithmic phase of growth. Final yield (cells/mL) was calculated from the standard curve generated from Chlorella MGD1, another Chlorella species. n=3 *statistically different P-value≥0.05 One tailed t-test (Autotrophic vs. Heterotrophic) **statistically different P-value≥0.01 One tailed t-test (Mixotrophic vs. Heterotrophic)

Trophic state Growth Rate Generation Time Final Yield (days) (day-1) (cells/mL)

Autotrophic 0.319±0.011* 2.22 days ±0.079 1.0x10 8±4.63x10 8

Mixotrophic 0.328 ±0.059** 2.18 days±0.403 1.25x108±8.53x106

Heterotrophic 0.053±0.0029 13.11 days±0.727 3.25x107±9.79x107

Figure 10. Generation time (days) of Chlorella BI sp. compared with C. vulgaris under the different trophic conditions. (n=3). *Statistically different P-value≥0.05 one tailed t- test (heterotrophic state of Chlorella BI sp. vs. heterotrophic state of C. vulgaris)

Figure 11. Chlorophyll a: b ratio for C. vulgaris compared to Chlorella BI sp whole cells grown under autotrophic (A), mixotrophic (B) and heterotrophic (C) conditions. (n=3)

Figure 12. Total chlorophyll (ng/cell) for C. vulgaris compared to Chlorella BI sp. whole cells grown under autotrophic (A), mixotrophic (B) and heterotrophic (C) conditions. (n=3)

49 Figure 13. Representative room temperature Chl-a fluorescence induction curves of whole cells of C. vulgaris grown under variable trophic conditions.

50 Figure 14. Steady state chlorophyll a fluorescence parameters Fv/Fm, qP and qN for whole cells of C. vulgaris compared to Chlorella BI sp grown under autotrophic (A), mixotrophic (B) and heterotrophic (C) conditions. (n=2).

Figure 15. Quantification of Chlorella BI sp. vs. C. vulgaris immunoblot of RubisCO. Blots were loaded on protein basis and quantified with Scion image software and relative to the autotrophic state. (n=2).

Figure 16. A. Stained non-denaturing gradient gel for Chlorella BI sp. and C. vulgaris. The samples were loaded on an equal chlorophyll basis of 1 mg/mL Standard molecular weights (kDa) are indicated on the left. Major proteins are designated on the gel. Chlamydomonas UWO 241 and C. reinhardii were run as controls. 2nd dimension gel analysis of the autotrophic states of C. vulgaris (B) and Chlorella BI sp. (C).

CHAPTER THREE

ALTERATIONS IN THE PHOTOSYNTHETIC APPARATUS OF CHLORELLA BI SP. IN RESPONSE TO A MIMICKED SHIFT FROM POLAR WINTER TO SUMMER

54 INTRODUCTION

All photosynthetic organisms possess a variety of mechanisms to maintain a balance between energy production and energy consumption. Adequate pools of photochemically derived NADPH and ATP are required for energy-consuming metabolic processes. The process of balancing energy produced with energy consumed is called photostasis, and environmental conditions that perturb this balance are often compensated for by structural and functional changes in the photochemical apparatus (Hüner, et al., 1998). In addition, plants and algae encounter significant environmental stresses on a regular basis in their natural habitats (limited nutrient or water status, fluctuating light and temperature intensities) (Mittler, 2006) and have evolved a wide range of response mechanisms to survive and reproduce in the changing environment. For example, one seasonal environmental extreme faced by Chlorella BI sp., living in a microbial mat is variable light regimes. During polar summer, there are extreme levels of sunlight and UV radiation, while the transition between summer to winter is marked by a rapid decline in photosynthetic active radiation (PAR) and eventually complete darkness for four to six months in the polar night (Hawes, et al., 2003). Because light availability is inconsistent as it varies from very high, inhibitory levels down to below the compensation point (i.e. where the rate of photosynthesis matches the rate of respiration) for photosynthesis. It seems probable that algae residing in polar mats should be able to acclimate to broad ranges of PAR, and possess mechanisms to maintain photostasis. The ultrastructural rearrangement of the photosynthetic apparatus (also termed “greening” in plants) is well characterized in plants and algae and represents the ability to alter the photosynthetic apparatus in variable environmental conditions. The objective of this chapter was to investigate how the photosynthetic apparatus of Chlorella BI sp. responds to a mimicked winter to summer transition by monitoring functional alterations during a shift from the absence of light energy (dark) to the presence of light energy (light).

55 METHODS Cultures of Chlorella BI sp. were grown heterotrophically until they reached early log phase (optical density ~ 0.2) and were then switched to the light which mimicked the transition between polar winter and summer. Samples were collected every 6 hours for up to 14 days. Samples were measured for growth, chlorophyll, glucose consumption low temperature fluorescence emission (77K) and steady state Chl a fluorescence parameters (See Chapter 2 for a detailed description of the methodology).

56 RESULTS

Growth and glucose consumption of Chlorella BI sp. during the shift from heterotrophy (dark) to light (mixotrophy)

Growth of Chlorella BI sp. in heterotrophy reached an optical density of ~ 0.2 at 0h of the shift, 72h post inoculation, comparable to the optical density of the alga in the heterotrophic cultures of the trophic state experiments at 72h (see figure 2A in Chapter 2). Once shifted to light, the organism had a growth rate of 0.011 (day-1) in log phase (50h-100h post shift) (Fig.1A). The steady increase in growth was attributed to the presence of light energy with the rate of glucose consumption being 1.8µg/mL per hour (Fig 1B). However, the increase in glucose consumption occurred after a significant lag in growth compared to the rapid growth and glucose consumption of mixotrophic cultures in Chapter Two. The final yield of the shifted mixotrophic cells was 1.09x108 approximately 50% lower than the final yield of mixotrophic cells observed in the trophic state experiments. Additionally, the generation time for shifted mixotrophic cells was 2.59 days, which was also 20% higher than the generation time of mixotrophic cells in the trophic state experiments.

Functional adjustment in Chlorella BI sp. photochemical apparatus from heterotrophy (dark) to mixotrophy (light)

As alterations in light availability can alter the requirement for photochemically derived energy, energy partitioning between Photosystem II (PSII) and Photosystem I (PSI) was measured by Chl a low temperature (77K) fluorescence spectra in cultures that were shifted to light after growing in dark conditions. The cultures were measured every 24hrs after the shift to light conditions. Subsequently, the ratios of LHCII: PSII and PSI: PSII were calculated by measuring the spectra peak at 683nm for PSII, 695nm for LHCII and 715nm for PSI. The photochemical apparatus of Chlorella BI sp. experienced functional changes in response to the shift in light quality (i.e. the presence or absence of light). The light energy distribution between PSII and PSI was relatively similar in the first 75h post shift (Fig. 2). Subsequently, there was a 20% increase in the LHCII: PSII

57 (F683/F695) ratio approximately around 100h compared to a 14% increase in PSI: PSII ratio (F715/F695) (Fig. 3). After the first 100hrs, the LHCII: PSII ratio was still higher compared to the PSI:PSII ratio, however, both ratios steadily declined.

Effect of the shift from dark to light on chlorophyll synthesis for Chlorella BI sp.

The chlorophyll a:b ratio fluctuated when Chlorella BI sp. was shifted to light. Initially, there was an initial decrease as the ratio was 1.95 at 0h and 0.35 at 24h. Subsequently there was approximately an 80% transient increase in chlorophyll a: b ratio around 30h post shift. Around 72hrs post shift, there about a 40% decrease in the ratio and afterwards the ratio remained stable (Fig. 4A). Total chlorophyll (ng/cell) did not vary as prominently as the chlorophyll a:b ratio. There was a 33% decrease from 0h to 75h post shift, however it remained relatively unchanged thereafter (Fig. 4B).

Steady state Chl a fluorescence parameters for Chlorella BI sp. in the absence or presence of light energy. Room temperature induction curves which were measured in heterotrophic cultures indicated there was less relative fluorescence in Chlorella BI sp. (Fig. 5A) compared to shifted cultures (OD~0.6) where the relative fluorescence increased almost 80% in light conditions (Fig. 5B). Upon the shift to light, the steady state Chl a fluorescence parameter, Fv/Fm, is 0.255 at 0h post shift and then increased to 0.443, a 47% increase in the maximum PSII photochemistry in the first 50h post shift (Fig. 6A). qP is 1 at 0h indicating QA is fully oxidized but subsequently decreased to 0.491 (Fig. 6B). qN remained low around 0.180 throughout the shift (Fig. 5B).

DISCUSSION

The harsh condition of a microbial mat exposes organisms to a variety of environmental extremes which can change dramatically over short periods of time. While research regarding acclimation to environmental shifts in photopsychrophiles is still in its infancy, it seems likely that cold adapted algae require a variety of mechanisms to acclimate to environmental variability. In this study, results showed that Chlorella BI sp. was able to adjust its photochemical apparatus in response to a mimicked winter to summer transition, indicating that this organism possesses the capacity to acclimate to environmental variability. Growth and glucose consumption in Chlorella BI sp. prior to the shift were similar to that of the heterotrophic cultures in the trophic state experiments. Once the organism was shifted to light, there was an increase in growth (Fig. 1A) with a concomitant decrease in glucose (Fig. 1B), which implied the organism utilized glucose for growth. However, there was a significant lag in glucose consumption in the shifted cells compared to the mixotrophic cultures in Chapter Two. The decrease in consumption can perhaps be attributed to changes induced by the shift as there was also a decrease in growth rate, generation time and final yield of shifted cells. The shift might have delayed transcription of key cellular genes needed for rapid adjustment from the absence of light to the presence of light. However, further experiments need to be conducted to determine if the transcription of key cellular genes such as psbA, rbcL and lhc is altered during the shift. Chlorella BI sp. exhibits the ability to alter its photochemical apparatus. At the level of dynamic energy distribution between the respective photosystems, however, 77K spectra analysis suggests energy partitioning does not change significantly in response to the dark to light shift (Fig. 2). The ratio of LHCII:PSII (F685/F695) at 168h for PSII was

76% higher than PSI: PSII (F715/F695) (Fig. 3) suggesting Chlorella BI sp. LHCII is not closely associated with PSII. Furthermore, the 77K spectral analysis ratios for mixotrophic cultures (Chapter 2, Table 3) are relatively similar to the ratios of the shifted cultures. This implies that in response to the change in environmental conditions (absence of light energy to the presence of light energy); Chlorella BI sp does not change

59 the relative quantities (stoichiometry) of LHCII, PSI and PSII. The increased ratio of LHCII: PSII is more likely due to an increase in optical density, which increased as the shifted cultures entered log phase at approximately 100 hrs. In further studies, the spectral analysis of shifted cultures must be repeated and normalized to the internal standard Fluoroscein (514nm) to determine the accuracy of the ratios. Chlorophyll synthesis indicated that the Chl a: b ratio was variable throughout the first 100h of the shift (Fig. 4A). The increase in the Chl a: b ratio within the first 50h of the shift, suggest Chl a is developed first. This type of chlorophyll is associated with the core complex where Chl a/b protein complexes binds to the peripheral antennae. The inner antennas, CP43 and CP47, are associated with Chl a and transfer energy from the peripheral antenna to the reaction center (Green and Durnford, 1996) indicating Chl a must be synthesized immediately after the shift for proper energy transfer. Chlorophyll b is a ubiquitous accessory pigment and is associated with adjustment to variable light conditions for most green plants and algae (Vernotte, et al., 1992). Because it is not required for immediate adjustment for the organism once it has been shifted and is dependent on Chl a for its development (von Wettstein, et al., 1995) Chl b is most likely synthesized after the first 100h of the shift to mixotrophy indicated by the drop in Chl a: b ratios. Total chlorophyll (ng/cell) does not fluctuate as prominently in the shifted cells compared to the heterotrophic and mixotrophic cultures of Chapter Two. Instead it remains relatively stable throughout the shift. As the Chl a:b ratio suggests, Chl a might be developed first when the cells are shifted to light. This may imply that as a consequence of shifting from heterotrophy to mixotrophy, Chlorella BI sp., might alter other components of total chlorophyll and create necessary pigments such as Chl a first. Further studies must be conducted to verify if Chl a is synthesized first and to determine if other pigments and components are altered during the shift. Room temperature induction curves indicated there was less relative fluorescence in the heterotrophic cultures (Fig. 5A) of Chlorella BI sp., however upon the shift to the presence of energy (Fig. 5B), there was a rapid increase in relative fluorescence attributed to the increase in the amount of chlorophyll present. Steady state Chl a fluorescence parameters further indicate functional differences of Chlorella BI sp. cells when shifted.

Fv/Fm stabilized at 0.450 (Fig. 6A) around 50h post shift and remains relatively constant

60 throughout the shift, suggesting the maximum PSII photochemistry does not fluctuate when shifted. At 0.450, the Fv/Fm was 26% lower for shifted Chlorella BI sp. mixotrophic cells compared to the Fv/Fm of mixotrophic cells during the trophic experiments, implying there is less maximum PSII photochemistry once the organism is shifted. qP of mixotrophic cells begins at 1 to 0h post shift, indicating QA is fully oxidized. The drastic difference between the qP of shifted cells compared to the qP of mixotrophic cultures in trophic state experiments can be attributed to the low density of the shifted cells which had an optical density of approximately 0.2 whereas measurements for qP were taken for the trophic state experiments during mid log phase which was approximately at an OD of 1. Furthermore, qP steadily decreased throughout the shift (Fig. 6B), suggesting that the intersystem electron transport pool became reduced in response to light exposure and was more oxidized at the beginning of the shift. qN remained relatively low throughout the shift (Fig. 6B) indicating that Chlorella BI sp. did not alter non photochemical quenching capacity during light acclimation.

Chlorella BI sp. is able to adjust its photosynthetic apparatus to the change in light quality. Future studies would be to determine the specific effect of the shift on PSI. It remains to be determined what specific aspects Chlorella BI sp. has modified for PSI in order to transition from heterotrophy to mixotrophy. Additional experiments such as characterizing protein abundance of major photosynthetic proteins and mRNA expression studies of important genes involved in the development of the photochemical apparatus are needed to give a better understanding of the change Chlorella BI sp. photochemical apparatus endures when shifted.

Figure 1A. Growth of Chlorella BI sp. during the shift from dark to light. Growth was measured as a change in optical density (OD750) as a function of time before and after the shift. The cultures were grown at 8°C/20 μmol photons m-2 s-1 (n=3). B. Glucose consumption of Chlorella BI sp. during the shift experiment. Glucose (HK) assay kit (SIGMA) was used for the enzymatic determination of glucose consumption during the shift. Consumption was measured as µg glucose/mL consumed during the shift (n=3).

Figure 2. Low temperature (77K) analysis of Chlorella BI sp. cells shifted from dark to light. Fluorescence yield was calculated for Chlorophyll a fluorescence emission (435nm) for Photosystem II (PSII) (■) and Photosystem I (PSI) (●) (n=2).

Figure 3. Low temperature (77K) fluorescence emission ratios of Chlorella BI sp. cells in the shift from dark to light (n=2).

Figure 4. Changes in chlorophyll during the shift from dark to light. Samples were collected every 6 hours after the shift to light and then analyzed for Chlorophyll a: b (A) and Total chlorophyll (ng/cell) (B) (n=4).

Figure 5. Representative room temperature chlorophyll fluorescence induction curves for Chlorella BI sp. cells shifted from dark to light.

Figure 6.Steady-state chlorophyll a fluorescence quenching parameters. A. Maximum photochemical efficiency of dark adapted cells was measured as Fv/Fm. B. Photochemical efficiency of energy transferred to the photosynthetic apparatus was measured as qP (■) and non photochemical quenching was measured as qN (▲). Measurements were taken directly before the shift to mixotrophy and at regular intervals thereafter (n=3).

67 REFERENCES

Ahmed I, Hellenbust JA (1990) Chloroplast development by nitrogen source and growth conditions in Chlorella protothecoides strain. Plant Physiol 944-949

Bacon K (2001. Kluwer Academic Publishers) Photosynthesis. Photochemistry and photobiophysics. In Govindjee, Amese, J.; Aro, E.M.; Barber, J.; Blankenship, R.E.; Murata, N.; & Ort, D.R.; (Eds). Advances in Photosynthesis 10: 420-430

Balmer Y, Vensel WH, Cai N, Manieri W, Schurmaan P, Hurkman WJ, Buchanan BB (2006) A complete ferredoxin/thioredoxin system regulates fundamental processes in amyloplasts. PNAS 103: 2988-2993

Baker NR, Rosenqvist E (2004) Applications of chlorophyll fluorescence can improve crop production strategies: an examination of future possibilites. J Exp Bot 1607-1621

Bjorkman O, Demmig B (1987) Photon yield of O2 evolution and chlorophyll fluorescence characteristics at 77K among vascular plants of diverse origins. Planta 170: 489-504

Butler WL (1978) Energy distribution in the photochemical apparatus of photosynthesis. Annu Rev Plant Physiol 29: 345-358

Canfield DE, & Des Marais DJ (1993) Biogeochemical cycles of carbon, sulfur and free oxygen in a microbial mat. Geochmicia et Cosomochimica 57: 3971-3984

Cheng CL, Acedo GN, Cristinsin M, & Conkling MA (1992) Sucrose mimics the light induction of Arabidopsis nitrate reductase gene transcription. Proc Natl Acad Sci USA 89: 1861-1864

Cho SH, Ji SC, Hur SB, Bae J, Park IS, & Song YC (2007) Optimum temperature and salinity for growth of green alga Chlorella ellipsoidea and nannochloris oculata. Fisheries Science 73: 1050-1056

Constant S., Perewoska I, Alfonso M., & Kirilovsky D (1997) Expression of the psbA gene during photo inhibition and recovery in Synechocystis PCC 6714: inhibition and damage of transcriptional and translational machinery prevent the restoration of Photosystem II activity. Plant Mol Biol 32: 1-13

De los Rios A, Ascaso C, Wierzchos J, Fernandez-Valiente E, & Quesada A (2004) Microstructural characterization of cyanobacterial mats from the McMurdo Ice Shelf, Antarctica. Appl. Environ. Microbiol. 70: 569-580

68 Dietz KJ, Schreiber U, Heber U (1985) The relation between the redox state of QA and photosynthesis in leaves at various carbon dioxide, oxygen and light regimes. Planta 166: 219-226

Durnford DG (2003) Structure and regulation of Algal light-harvesting complex genes. In Photosynthesis in Algae. Kluwer Academic Publishers, Netherlands, pp 63-82

Eggink LL, Park H, Hoober JK The role of chlorophyll b in photosynthesis. BMC Plant Biology 1:1-10

Green BR, Durnford DG (1996) The chlorophyll-carotenoid proteins of oxygenic photosynthesis. Annu Rev Plant Physiol Plant Mol Biol 685-714

Griffiths DJ, Thresher CL, & Street E (1960) The heterotrophic nutrition of Chlorella vulgaris (Brannoni No. 1 strailn). Ann Botany 24: 1-11

Guillard RRL (1973) Divison rates. ln: Stein, JE (ed) Handbook of Phycological Methods. Cambridge University Press, Cambridge, UK

Hanke GT, Kurisu G, Kusunoki M, & Hase T (2004) Photosynth Res 317-327

Hawes I, Smith R, Howard-Williams C, Schwarz AM (2001) Environmental conditions during freezing and response of microbial mats in ponds on the McMurdo Ice Shelf, Antarctica. Antarc Sci 198-208

Hawes I, & Howard-Williams C (2003) Pond life on the McMurdo Ice Shelf, one of the world’s strangest ecosystems. NIWA 11: 21-22

Huner NPA, Oquist G, Sarhan F (1998) Energy balance and acclimation to light and cold. Trends in Plant Science 3: 224-230

Jang JC, & Sheen J (1994) Sugar sensing in higher plants. Plant Cell 6: 1664-1676

Jeffery SW, Humphrey GF (1975) New spectrometric equations for determining chlorophyll a, b, c1, c2 in higher plants, algae and natural phytoplankton. Biochem Physiol Pflanz 167: 191-194

Karlander EP, & Krauss RW (1966) Responses of Heterotrophic Cultures of Chlorella vulgaris Beyerinck to Darkness and Light. I1. Action Spectrum for and Mechanism of the Light Requirement for Heterotrophic Growth. Plant Physiol 7-14

Knaff DB (1996) Ferredoxin and ferredoxin-dependent enzymes

Koch KE (1996) Carbohydrate-modulated gene expression in plants. Annu Rev Plant Physiol Plant Mol Biol 47: 509-540

69 Krause GH, Weis E Chlorophyll fluorescence and photosynthesis: the basics. Ann Rev Physiol Plant Mol Biol 313-349

Krause GH, Jahns P (2004) Non-photochemical quenching energy dissipation determined by chlorophyll fluorescence quenching: characterization and function. In Chlorophyll a Fluorescence: A signature of Photosynthesis, ed. , pp 463-495

Kyle DJ, Staehelin LA, Arntzen CJ (1983) Lateral mobility of the light harvesting complex in chloroplast membranes controls excitation energy distribution in higher plants. Arch Biochem Biophys 527-541

Lesser MP, Barry TM, Banaszak. A.T. (2002) Effects of UV radiation on a chlorophyte alga (Scenedesmus sp.) isolated from the furmarole fields of Mt. Erebus, Antarctica. J Phycol 38: 473-481

Lizotte MP, Priscu JC (1992) Spectral irradiance and bio-optical properties in perennially ice-covered lake (Lake Bonney, Antarctica). J Phycol 281: 179-185

Marquez FJ, Sasaki K, Kakizono T, Nishio N, Nagai S (1993) Growth character of Spirulina platensis in mixotrophic and heterotrophic conditions. J Ferment Bioeng 74: 408-410

Maxwell DP, Falk S, Trick CG, Hüner N (1994) Growth at low temperature mimics high-light accumulation in Chlorella vulgaris. Plant Physiol 105: 535-543

Merchant S, Allen MD, Kropat J, Moseley JL, Long JC, Tottey S, Terauchi AM (2006) Between a rock and a hard place: trace element nutrition in Chlamydomonas. Biochim Biophys Acta 1763: 578-594

Mittler R (2006) Abiotic stress, the field environment and stress combination. Trends Plant Sci 11: 15-19

Morgan-Kiss RM, Ivanov AG, Modla S, Czymmek K, Huner NPA, Priscu JP, Lisle JT, Hanson TE (2008) Identity and physiology of a new psychrophilic eukaryotic green alga, Chlorella sp., strain BI, isolated from a transitory pond near Bratina Island, Antarctica. Extremophiles 12: 701-711

Morgan-Kiss RM, Ivanov AG, Priscu JC, Maxwell DP, Hüner N (1998) Structure and composition of the photochemical apparatus of the Antarctic green alga, Chlamydomonas subcaudata. Photosynthesis Research 56: 314

Morgan-Kiss RM, Priscu JP, Pocock T, Gudynaite-Savitch L, Hüner N (2006) Adaptation and acclimation of photosynthetic microorganisms to permanently cold environments. Microbiol Molec Biol Rev 70: 222-252

Morita RY (1975) Psycrophilic bacteria. Bacteriol Rev 39: 144-167

70 Neale PJ, Priscu JC (1995) The photosynthetic apparatus of phytoplankton from a perennially ice-covered Antarctic lake: acclimation to an extreme shade environment. Plant Cell Physiol 36: 253-263

Nymark M, Valle K, Brembu T, Hancke K, Winge P (2009) An Integrated Analysis of Molecular Acclimation to High Light in the Marine Diatom Phaeodactylum tricornutum. PLoS ONE 11

Oertel A, Aichinger N, Hochreiter R, Thalhamer J, Lutz-Meindl U (2004) Analysis of mucilage secretion and excretion in Micrasterias (Chlorophyta) by means of immunoelectron microscopy and digital time lapse video microscopy. J Phycol 40: 711- 720

Ogawa T, & Aiba S (1981) Bioenergetic analysis of mixotrophic growth Chlorella vulgaris and Scenedesmus acutus. Biotechnol Bioeng 23: 1121-1132

Paerl HW, Pinckney JL (1996) Microbial consortia: Their roles in aquatic production and biogeochemical cycling. Microb Ecol 31: 225-247

Paerl HW, Pinckney JL, Steppe TF (2000) Cyanobacterial bacterial mat consortia: examining the functional unit of microbial survival and growth in extreme environments. Environmental Microbiology 2: 11-26

Pocock T, Lachance MA, Proschold T, Priscu JC, Kim S, Huner NPA (2004) Identification of a psychrophilic green alga from Lake Bonney antarctica: Chlamydomonas raudensis ETTIL. (UWO 241) (Chlorophyceae). J. Phycol 1138-1148

Ramsing N, KuÈ hl M, and Jùrgensen BB (1993) Distribution of sulfate-reducing bacteria, O2, and H2S in photosynthetic biofilms determined by oligonucleotide probes and microelectrodes. Appl Environ Microbiol 59: 3840-3849

Risatti JB, Capman WC, and Stahl DA (1994) Community structure of a microbial mat: The phylogenetic dimension. Proc Natl Acad Sci 91: 10173-10177

Rothchild LJ, & Mancinelli RL (2001) Life in extreme environments. Nature 409: 1092-1101

Usada H, Edwards GE (11982) Influence of Varying CO, and Orthophosphate Concentrations on Rates of Photosynthesis, and Synthesis of Glycolate and Dihydroxyacetone Phosphate by Wheat Chloroplasts. Plant Physiol. 469-473

Vernotte C, Picaud M, Kirlovsky D, Olive J, Ajlani G, Astier C (1992) Changes in the photosynthetic apparatus in the cyanobacterium Synechocystis sp. PCC 6714 following light to dark and dark to light transitions. Photosynthesis Research 32: 45-57 von Wettstein D, Gough S, & Kannangara CG (1995) Plant cell 7: 1039-1057

71 Wilson KE, Huner NPA (2000) The role of growth rate, redox-state of the plastoquinone pool and the trans-thylakoid membranes. Biochim Biophys Acta 1411: 21- 85

Xie J, Zhang Y, Li Y, Wong Y (2001) Mixotrophic cultivation of Platymonas subcordiformis. Journal of Applied Phycology 343-347

Yang C, Hua Q, Shimizu K (2000) Energetic and carbon metabolism during growth of microalgal cells under photoautotrophic, mixotrophic and cyclic light-autotrophic/dark- heterotrophic cultures conditions. Biochemical Engineering Journal 6: 87-102