Elucidation of Functional and Regulatory Aspects of Sulfate Transport in Chlamydomonas Reinhardtii

Home , Sabeeha Merchant

ELUCIDATION OF FUNCTIONAL AND REGULATORY ASPECTS OF SULFATE TRANSPORT IN CHLAMYDOMONAS REINHARDTII

A DISSERTATION

SUBMITTED TO THE DEPARTMENT OF BIOLOGY

AND THE COMMITTEE ON GRADUATE STUDIES

OF STANFORD UNIVERSITY

IN PARTIAL FULFILLMENT OF THE REQUIREMENTS

FOR THE DEGREE OF

DOCTOR OF PHILOSOPHY

Wirulda Pootakham

July 2010

This work is licensed under a Creative Commons Attribution- Noncommercial 3.0 United States License. http://creativecommons.org/licenses/by-nc/3.0/us/

This dissertation is online at: http://purl.stanford.edu/jq238dx7130

ii I certify that I have read this dissertation and that, in my opinion, it is fully adequate in scope and quality as a dissertation for the degree of Doctor of Philosophy.

Arthur Grossman, Primary Adviser

I certify that I have read this dissertation and that, in my opinion, it is fully adequate in scope and quality as a dissertation for the degree of Doctor of Philosophy.

Martha Cyert, Co-Adviser

I certify that I have read this dissertation and that, in my opinion, it is fully adequate in scope and quality as a dissertation for the degree of Doctor of Philosophy.

Mary Mudgett

I certify that I have read this dissertation and that, in my opinion, it is fully adequate in scope and quality as a dissertation for the degree of Doctor of Philosophy.

Zhiyong Wang

Approved for the Stanford University Committee on Graduate Studies. Patricia J. Gumport, Vice Provost Graduate Education

This signature page was generated electronically upon submission of this dissertation in electronic format. An original signed hard copy of the signature page is on file in University Archives.

iii ABSTRACT

Chlamydomonas reinhardtii (Chlamydomonas) exhibits a suite of responses to

2- sulfur (S) deprivation, including an elevation of sulfate (SO4 ) uptake, synthesis of extracellular arylsulfatase, down-regulation of photosynthesis and cessation of growth and cell division. One of the earliest responses to S starvation is an increase in the

2- 2- SO4 transport rate. Aspects of SO4 transport during S-replete and S-depleted

2- conditions were previously studied, although the SO4 transporters had not been functionally identified. In this study, both forward and reverse genetic approaches

2- were employed to identify SO4 transporters in Chlamydomonas. SULTR2, SLT1, and

SLT2 transcripts and polypeptides increased markedly in S-starved wild-type cells,

2- suggesting that these genes encode high-affinity SO4 transporters that function when the cells experience S limitation. Mutant strains defective for acclimation to S starvation, sac1 and snrk2.1, exhibited much less of an increase in the level of

SULTR2, SLT1, and SLT2 transcripts and their encoded proteins during S deprivation compared to wild-type cells. Consequently, these two strains were unable to induce

2- SO4 uptake to the same extent as in the wild-type strain. The SULTR2, SLT1 and

SLT2 polypeptides were localized to the plasma membrane and their rates of turnover were significantly impacted by S availability; the turnover for SLT1 and SLT2 (but not for SULTR2) was demonstrated to be dependent on proteasome function. Mutants identified for each of the S-deprivation-responsive transporters were used to establish

2- their critical role in the transport of SO4 into S-starved cells.

I have also discovered a sequential, temporal regulation of the S-starvation

2- responsive genes. The primary responses (e.g. the induction of high-affinity SO4

iv transporters) are not dependent on protein synthesis occurring on 80S ribosomes. In contrast, the secondary responses (the induction of ARS, ECP76 and ECP88) require de novo protein synthesis. ARS73a, a putative transcriptional activator, is directly or indirectly involved in the regulation of the expression of the second tier genes, most of which encode proteins associated with the scavenging of extracellular S or the redistribution of internal S. Genetic analysis has shown that ARS73a and SAC3 kinase act in the same pathway; ARS73a is epistatic to SAC3. These new discoveries are incorporated into a model that describes S-deprivation elicited regulation in

Chlamydomonas.

v ACKNOWLEDGEMENTS

My PhD project was like a roller coaster ride. There were lots of ups and downs, and I cherished every moment of it. There was a rush of joy, self-satisfaction, and excitement when I got fascinating data. There were also times when my scientific world was filled with disappointment and frustration from failed experiments. I would like to thank a number of people who’ve helped me throughout my graduate career.

First and foremost, I want to thank my advisor, Arthur Grossman. You are a great scientist and a wonderful teacher. Thank you so much for your support and encouragement. You always gave me great advice and never told me what to do.

That’s one of the greatest qualities any advisor could have. You’ve given me freedom to make my own decisions about the project, and I’m very grateful for that. Most importantly, thank you for encouraging me during tough times. If you hadn’t convinced me that this project would turn out to be a beautiful story, I might have given up halfway through.

There are three very important people without whom I couldn’t have completed this PhD: Nakako Shibagaki, Jeffrey Moseley and David Gonzalez-

Ballester.

Nakako, you are one of the most amazing people I’ve ever met. I remember doing my first rotation project with you, and I didn’t know a lot about molecular biology. You were kind and very patient with me. You taught me a lot of techniques and little tricks that turned out to be really useful. Thanks for all the great discussions we had about my project and keeping me company on the weekends.

vi Jeff, you rock! You are my personal “Chlamydomonas sourcebook,” really.

Pretty much everything I know about Chlamy, I learned from you. Thanks for letting me share my scientific passion with you. It’s good to know that there is another person who cares about sulfate transporters as much as I do. I miss all the discussions we had

– you always motivated me.

David, lab would not be as fun without you. Thank you for helping me set up the mutant screen. I couldn’t have done it without you. I also want to thank you for cheering me up when I had bad days. I enjoyed making spaghetti noodles from TAP medium and washing glassware with you!

I also want to thank other Grossman/Bhaya lab members, past and present:

Devaki Bhaya, Florence Mus, Anne Steunou, Fariba Fazeli, Claudia Catalanotti,

Leonardo Magneschi, Matt Prior, Ariana Afshar, Blaise Hamel, Mark Heinnickel,

Wenqiang Yang, Sussi Wisén, Eva Nowack, Michelle Davidson, David Dewez,

Rosario Gomez, Mine Berg, Kate Mackey, Shaun Bailey, Chao-Jung Tu, and

Chungsoon Im. You guys create such a fun working environment. Thanks for your friendship and intellectual scientific discussions.

Thanks to my committee members: Martha Cyert, Mary Beth Mudgett and

Zhiyong Wang, for their support, encouragement and most importantly for their helpful suggestions. I would also like to thank my undergraduate advisor, David Stern, for jump-starting my research career.

Last but certainly not least, I want to thank my family: Mom, Dad, Thanyakarn

(my brother), and my partner and best friend, Metha Jeeradit. Mom and Dad, I couldn’t have done this without you. I don’t think I could ever thank you enough for

vii your endless love and support. I’m incredibly lucky to have you both and I would like to dedicate this thesis to you. Thanyakarn, you’ve always been a great friend and a wonderful brother. Thanks for keeping me sane while I was in graduate school.

Metha, thanks for being with me every step of the way. You are nothing but supportive, and I would never get this far without you. I’m really grateful for the love and support you’ve shown me. Thanks for enduring my relentless complaints about my experiments. Finally, thanks for being an amazing hiking/rock- climbing/bouldering partner. Those activities really helped maintain my sanity during my graduate career. ☺

viii TABLE OF CONTENTS

ABSTRACT iv ACKNOWLEDGEMENTS vi TABLE OF CONTENTS ix LIST OF TABLES xi LIST OF FIGURES xii

CHAPTER 1: Introduction 1 Sulfur in the environment 2 Chlamydomonas as a model organism 3 Sulfate acquisition and assimilation 5 Sulfate transport in Chlamydomonas 5 Arabidopsis sulfate transporters 7 Putative sulfate transporters in Chlamydomonas 10 Sulfate assimilation 11 Synthesis of cysteine and methionine 13 Synthesis of glutathione 14 Recycling of sulfur compounds 15 Sulfur starvation responses in Chlamydomonas 16 General and specific responses 16 Genes responsive to sulfur deprivation 18 Identification of genes controlling sulfur deprivation responses 20 Model for acclimation of Chlamydomonas to sulfur starvation 23 Thesis aims 25 References 37

CHAPTER 2: Selenate resistant mutant selection in Chlamydomonas 44 reinhardtii Abstract 45

ix Introduction 46 Results 50 Discussion 52 Materials and methods 56 References 67

CHAPTER 3: The sulfate transporters of Chlamydomonas reinhardtii; 69 From regulation to functionality Abstract 70 Introduction 72 Results 76 Discussion 91 Materials and methods 101 Acknowledgements 112 References 146

CHAPTER 4: ARS73a is involved in a tiered regulation of sulfur 151 starvation responses Abstract 152 Introduction 154 Results 159 Discussion 165 Future directions 169 Materials and methods 171 References 192

CHAPTER 5: Concluding remarks and future directions 195 References 206

x LIST OF TABLES

CHAPTER 1 2- Table 1-1. Arabidopsis SO4 transporters: their subcellular 34 locations, expression patterns, and functions. 2- Table 1-2. Genes involved in SO4 assimilation in Chlamydomonas. 36

CHAPTER 2 2- Table 2-1. Characteristics of SO4 transport in wild-type cells (D66) 66 and sr mutants in S-depleted conditions.

CHAPTER 3 Table 3-1. Growth rates of the CP60-1C strain harboring genes 142 encoding Arabidopsis SULTR1;2 or various Chlamydomonas 2- SO4 transporters. Table 3-2. List of SULTR2-, SLT1- and SLT2-specific primers 143 used for PCR screening of the insertion library. 2- Table 3-3. Characteristics of SO4 transport in wild-type cells, 145 2- single, double and triple SO4 transporter mutants after 24 h of S deprivation.

xi LIST OF FIGURES

CHAPTER 1 2- Figure 1-1. SO4 transporter structure. 27 2- Figure 1-2. SO4 assimilation pathway. 28 Figure 1-3. Proposed pathway for recycling of S. 30 Figure 1-4. Accumulation of ARS activity in wild-type 31 and sac1 strains. Figure 1-5. Model for S startvation response signaling pathways 32 in Chlamydomonas during S-replete and S-depleted conditions.

CHAPTER 2 Figure 2-1. Schematics showing steps in protocol use to select 59 for selenate resistant mutants. Figure 2-2. Secondary screening of sr mutants. 60 Figure 2-3. DNA gel blot analyses of insertional mutants. 63 Figure 2-4. Liquid culture assays for ARS activities in 65 TAP and after 18, 24, 42, and 72 h of growth in TAP-S medium for wild type (WT), sr111, and sr116.

CHAPTER 3 2- Figure 3-1. Amino acid sequence alignments of SO4 113 transporter proteins. 2- Figure 3-2. SO4 uptake in wild-type (WT) Chlamydomonas 116 cells and the sac1 and snrk2.1 mutant strains. Figure 3-3. Real time quantitative PCR analysis of the levels of 117 various transcript. 2- Figure 3-4. Changes in transcript abundances for various SO4 118 transporters following imposition of S deprivation.

xii Figure 3-5. SULTR2, SLT1, and SLT2 polypeptide abundances 120 in wild-type (WT), sac1, and snrk2.1 strains. Figure 3-6. Accumulation of SULTR2, SLT1, and SLT2 121 transcripts and proteins and their decay following the addition 2- of SO4 to S-deprived cells. Figure 3-7. Cycloheximide inhibition of accumulation of the 123 2- SO4 transporter protein during S deprivation. Figure 3-8. Determination of half-lives of SULTR2, SLT1, and 124 SLT2 proteins and the role of the proteasome in degradation. Figure 3-9. Localization of the Chlamydomonas SLT1, SLT2, 126 and SULTR2 transport proteins. 2- Figure 3-10. Expression of SO4 transporter-GFP fusion 128 proteins in S. cerevisiae cells. Figure 3-11. The copy number and position of the insertion of 130 2- the AphVIII marker gene in the SO4 transporter mutants. Figure 3-12. Amino acid sequence alignments of wild-type and 132 the SLT1 and SULTR2 mutant gene products. Figure 3-13. Real-time quantitative PCR for analysis of transcript 135 accumulation in wild-type and mutant strains. Figure 3-14. SULTR2, SLT1, and SLT2 polypeptide abundances 137 2- in SO4 transporter mutants. 2- Figure 3-15. Characteristics of SO4 transport in wild-type cells 138 and single, double and triple mutants in S-replete medium or medium devoid of S. 2- Figure 3-16. Characteristics of SO4 transport in wild-type 140 cells and single, double and triple mutants (progeny of a cross between a wild-type D66 strain and an slt1slt2sultr2 triple mutant) deprived of S for 24 h.

xiii CHAPTER 4 Figure 4-1. Effects of transcriptional and translational inhibitors 177 on the accumulation of S-deficiency responsive transcripts. Figure 4-2. Accumulation of ARS activity in wild-type and 180 ars73a strains. Figure 4-3. DNA gel blot analysis of genomic DNA from 181 wild-type and ars73a strains. Figure 4-4. Location of AphVIII marker in the ars73a mutant 182 and a predicted amino acid sequence of ARS73a. Figure 4-5. Real-time quantitative PCR for analysis of transcript 183 accumulation in wild-type (WT) and mutant strains. 2- Figure 4-6. Accumulation of SO4 transporter and ARS 186 polypeptides in wild-type and ars73a strains. Figure 4-7. Accumulation of ARS activity in the wild-type 187 (WT), sac3, ars73a and sac3ars73a double mutants. Figure 4-8. Model of S-deficiency responsive gene regulation. 189

CHAPTER 5 Figure 5-1. Diagram showing promising lines of future 205 investigations.

xiv

CHAPTER 1

INTRODUCTION

1 I. SULFUR IN THE ENVIRONMENT

Sulfur (S) is an essential element present in proteins, lipids, and polysaccharides. It is also required for the production of molecules that help organisms cope with reactive oxygen species (ROS) and heavy metals. In the aerobic atmosphere

2- of the Earth, S exists predominantly in its fully oxidized form, sulfate (SO4 ). Plants

2- and microorganisms reduce SO4 to sulfide through the process of assimilative reduction; the sulfide is used exclusively for the synthesis of cysteine, methionine,

2- glutathione and other metabolites. In contrast, certain anaerobic bacteria use SO4 as a terminal electron acceptor for respiration. Since animals do not have the enzymes

2- required for the reduction of SO4 to sulfide, the substrate required for the synthesis of cysteine, methionine and glutathione, the ingestion of S-containing amino acids are essential in mammalian diets (Tabe and Higgins, 1998).

Because most organisms do not store significant levels of S, their growth and development depend on an external source of this nutrient. In the past, soils had accumulated high S level as a consequence of the use of fertilizers contaminated with

2- SO4 salts or exposure to acid rain (Cole and Johnson, 1977; Johnson et al., 1982).

However, with increased purity of fertilizers and decreased occurrence of acid rain, S can be limiting in the environment. It may even limit plant productivity in certain agricultural areas (Mahler and Maples, 1987), which can result in reduced crop quality and seed yields. It was recently shown that the seeds of wheat plants grown in fields where S was limiting produced less storage protein and accumulated more glutamine and asparagine, which can be converted to the toxic compound acrylamide during the heating of the wheat flour (Muttucumaru et al., 2006).

2 II. CHLAMYDOMONAS AS A MODEL ORGANISM

Chlamydomonas reinhardtii (Chlamydomonas throughout) is a unicellular, soil-dwelling green alga that has been developed as a model organism for analyzing a number of different physiological processes. It has been particularly useful for the dissection of photosynthetic function and regulation since it can grow heterotrophically (in the dark on acetate) as well as photoautotrophically; mutant strains that are completely deficient for photosynthetic function are still viable. Since

Chlamydomonas possesses two anterior flagella (cilia in animals) and a basal body

(centrioles in animals), this alga has also served as a strong model for elucidating the composition, architecture and function and of these cellular structures. This model algal system has been especially valuable for elucidating pathological effects of cilia dysfunction (Keller et al., 2005) and is currently being developed as agents for bioremediation and the generation of biofuels (Ghirardi et al., 2000; Ghirardi et al.,

2007).

Another advantage of using Chlamydomonas is the availability of a range of genetic and molecular techniques that can be employed to study this organism.

Chlamydomonas can be maintained as a haploid or a diploid that can be studied by classical genetic manipulation, making it convenient to identify recessive mutations and to assign complementation groups. Genetic crosses are relatively simple and tetrad analyses can be performed within a three-week period. Several molecular techniques can be applied to Chlamydomonas and cDNA libraries, along with BAC and cosmid genomic libraries are available. There are a number of strategies to introduce DNA into Chlamydomonas, and it is one of the few organisms for which transformation of

3 the nuclear, mitochondrial and chloroplast genomes can be achieved (Kindle, 1990;

Lefebvre and Silflow, 1999). There are several methods of transforming

Chlamydomonas cells with exogenous DNA, which include vortexing the cells in the presence of glass beads and electroporation (Kindle, 1990; Shimogawara et al.,

1999a). The ability to transform cells with a frequency of well over 3,000 transformants per microgram of DNA allows for the generation of a mutant library.

Furthermore, the Chlamydomonas genome sequence has recently been completed and it has been invaluable for examining genome-wide expression patterns in this alga under various environmental conditions (Merchant et al., 2007).

Chlamydomonas is also suitable for studies of the responses of photosynthetic eukaryotes to nutrient limitation. Because it is a single celled organism, the analyses are not complicated by complex developmental patterns and a diversity of tissue types

2- (characteristic of vascular plants). For example, there are twelve genes encoding SO4 transporters in Arabidopsis; some are expressed in roots and others are expressed in

2- leaves (Takahashi et al., 2000; Shibagaki et al., 2002; Kataoka et al., 2004a). SO4 is transported from the soil into root cells by high-affinity transport systems, from the roots to the xylem, from the xylem to the shoots and then is remobilized from the leaves when the plants begin to senesce. Different factors may affect the accumulation and function of these transporters, rendering the study of the regulation of these transporters in vascular plants difficult. Chlamydomonas also represents an excellent system for dissecting the ways in which photosynthetic organisms respond to S deprivation. Furthermore, while many genes encoding regulatory components of the S starvation response pathway have been identified in Chlamydomonas (Davies et al.,

4 1996; Davies et al., 1999; Gonzalez-Ballester et al., 2008), few have been identified in vascular plants (Maruyama-Nakashita et al., 2006) (see below).

2- III. SO4 ACQUISITION AND ASSIMILATION

2- Much of the SO4 in soils may not be readily available to plants or microbes.

2- The SO4 anion can be adsorbed onto the surface of soil particles, and a large

2- proportion may be covalently bonded to organic molecules in the form of SO4 esters and sulfonates (Grossman and Takahashi, 2001). Chlamydomonas can utilize

2- esterified SO4 in the soil by secreting extracellular arylsulfatases (ARS). The

2- synthesis of ARS is regulated by the availability of SO4 in the environment. When

2- the concentration of free soil SO4 is low, Chlamydomonas synthesizes ARS proteins

2- that are exported into the extracellular space where they hydrolyze esterified SO4

2- from organic molecules, releasing free SO4 for uptake and assimilation (Lien and

Schreiner, 1975; de Hostos et al., 1988).

2- A. SO4 transport in Chlamydomonas

2- Eukaryotic organisms have multiple systems for SO4 uptake. High-affinity uptake is often detected following S deprivation (Yildiz et al., 1994; Cherest et al.,

2- 1997). The characterization of SO4 transport in Chlamydomonas during S-limited and S-sufficient growth has been reported (Yildiz, Davies et al. 2004). Both the

2- maximum velocity (Vmax) and the substrate concentration at which SO4 transport was at half maximum velocity (K1/2) were altered when S-replete cells were transferred to medium devoid of S; the Vmax increased by ~10 fold while the K1/2 decreased by ~7

5 fold (Yildiz et al., 1994). High-affinity transport activity can be detected within an hour of S deprivation. The increase in transport activity following S starvation was blocked by the addition of cycloheximide (an inhibitor of cytosolic translation) but not by the addition of chloramphenicol (an inhibitor of chloroplast translation). These results demonstrate that the development of new transport activities that accumulate in

2- response to S deprivation requires cytosolic protein synthesis. Furthermore, SO4 transport in Chlamydomonas is energy-dependent and may be driven by a proton gradient generated by a plasma membrane ATPase (Yildiz et al., 1994).

2- In photosynthetic organisms, the reductive assimilation of SO4 occurs in

2- plastids, which means that the SO4 anion must be transported across both the plasma membrane and the double membrane of the plastid envelop prior to reduction and

2- incorporation into organic molecules. Genes encoding plasma membrane SO4 transporters in plant were first cloned from Stylosanthes hamata by functional

2- complementation of Saccharomyces cerevisiae SO4 transporter mutants (Smith et al.,

2- 1997). This breakthrough was followed by identification of SO4 transporters from other plant species, including Arabidopsis thaliana (Arabidopsis throughout) and

Brassica olereacae (Takahashi et al., 1997; Takahashi et al., 2000; Buchner et al.,

2- + 2- 2004). SO4 transporters found in vascular plants are in the H /SO4 cotransporter

+ 2- family (SLC26 family) while those present in mammals are in the Na /SO4 transporter family (SLC13 family).

6 2- B. Arabidopsis SO4 transporters

2- Arabidopsis has twelve putative SO4 transporters that have been classified into four groups. These transporters, their functions, subcellular locations and the patterns of expression of their genes are presented in Table 1-1. The AtSULTR1 group represents the high affinity transporters and the AtSULTR2 group represents

2- transporters with lower affinity for SO4 . All members of AtSULTR1 and AtSULTR2

2- groups were able to rescue the growth of a yeast SO4 transporter mutant (CP154-7B; this strain is disrupted for the genes encoding both of its two high-affinity transporters

2- and is unable to grow in medium containing less than 1 mM SO4 ). The kinetic properties of the transporters were also determined in the complemented yeast strain.

The affinity of the AtSULTR1 transporters is in the micromolar range (~4-7 µM) and that of the AtSULTR2 group is in the millimolar range (Takahashi et al., 2000;

Shibagaki et al., 2002; Yoshimoto et al., 2002; Yoshimoto et al., 2003). Expression patterns of AtSULTR1;1 and AtSULTR1;2 suggest that high affinity transporters are synthesized in cells present in the outer layer of roots and that their activity helps

2- 2- maximize the capacity of plants for SO4 uptake when there is little available SO4 in the environment. AtSULTR1;1 is present in root cortical cells and the epidermis of root hairs, and the gene is highly induced in response to S deprivation (Takahashi et al., 2000). In contrast, the AtSULTR1;2 protein, also located in root cortical cells,

2- appears to be primarily responsible for the uptake of SO4 under conditions of S sufficiency (Shibagaki et al., 2002; Yoshimoto et al., 2002). Studies with T-DNA

2- knockout lines have demonstrated that the contribution of AtSULTR1;2 to total SO4 uptake into roots is predominant over the function of AtSULTR1;1, although either of

7 2- the two transporters is sufficient for the acquisition of environmental SO4

(Yoshimoto et al., 2007).

Unlike AtSULTR1;1 and AtSULTR1;2, AtSULTR1;3 exhibited a phloem- specific localization. The AtSULTR1;3-green fluorescent protein fusion was found in sieve element-companion cell complexes in roots and cotyledons. Movement of 35S-

2- labeled SO4 from cotyledons to sink organs was restricted in the T-DNA insertion mutant of AtSULTR1;3, suggesting that the AtSULTR1;3 transporter plays a role in

2- the loading of SO4 into the sieve tube, initiating source-to-sink translocation of S in

Arabidopsis (Yoshimoto et al., 2003).

Two low-affinity transporters, AtSULTR2;1 and AtSULTR2;2, are localized in the vasculature of roots and shoots. They likely play a role in the translocation of

2- SO4 from root to leaves (Takahashi et al., 2000). Not much is known about the

AtSULTR3 group of transporters; AtSULTR3;1, AtSULTR3;2 and AtSULTR3;3 are expressed exclusively in leaves (Takahashi et al., 2000). One member of this group,

AtSULTR3;5, has been shown to work in conjunction with AtSULTR2;1 to facilitate

2- the root-to-shoot transport of SO4 in the vasculature (Kataoka et al., 2004a).

AtSULTR3;5 colocalizes with the AtSULTR2;1 low-affinity transporter in xylem

2- parenchyma and pericycle cells in root. In a yeast (heterologous system), SO4 uptake was barely detectable in the yeast CP154-7B mutant that was expressing AtSULTR3;5

2- alone. However, coexpression of AtSULTR3;5 and AtSULTR2;1 resulted in SO4 transport activity that was significantly higher than the activity observed in CP154-7B

2- only expressing AtSULTR2;1. Furthermore, root-to-shoot transport of SO4 was restricted in a sultr3;5 mutant. Together, these results suggest that although

8 2- AtSULTR3;5 may not function independently to facilitate SO4 transport, it may

2- enhance the capacity of AtSULTR2;1 to distribute SO4 to aerial parts of the plant

(Kataoka et al., 2004a).

2- Two members of group 4 SO4 transporters accumulate specifically in the tonoplast membrane and are localized to xylem parenchyma cells in roots and hypocotyls. Vacuoles isolated from the sultr4;1sultr4;2 double mutant showed excess

2- accumulation of SO4 , suggesting that AtSULTR4;1 and AtSULTR4;2 facilitate the

2- efflux of SO4 from vacuoles, which would help optimize the internal distribution of

2- SO4 within plants. Comparisons of single and double mutants have suggested that

2- AtSULTR4;1 plays a major role in facilitating the efflux of SO4 from vacuoles while

AtSULTR4;2 has a supplementary function (Kataoka et al., 2004b).

+ 2- The H /SO4 cotransporters usually have 10-12 transmembrane domains that form their catalytic moiety. Like a number of anion transporters of the SLC26 family, they contain a carboxy-terminal STAS domain that resembles the bacterial anti-sigma factor antagonist SPOIIAA. SPOIIAA is a small protein that interacts with the antisigma factor SPOIIAB, freeing the sigma factor to activate the sporulation in Bacillus subtilis (Aravind and Koonin, 2000; Ho et al., 2003). The STAS domain is thought to have a regulatory function and recent work has shown that it is critical for the function

2- of the SO4 transporters and for efficiently localizing them to the plasma membrane

(Shibagaki and Grossman, 2004, 2006). A number of mutations in the AtSULTR1;2

STAS domain, including lesions in the conserved Thr that is thought to be a site of phosphorylation, render the transporter nonfunctional in yeast cells (Rouached et al.,

2005; Shibagaki and Grossman, 2006). The structure and function of the STAS

9 domain has also received some attention from the medical community since lesions in the STAS domain can lead to serious diseases in humans (Everett and Green, 1999).

2- C. Putative SO4 transporters in Chlamydomonas

2- Based on whole genome analysis and EST information, seven putative SO4 transporter genes were initially identified on the Chlamydomonas genome. Four of

+ 2- these are similar to plant type transporters (H /SO4 cotransporters); they have been designated SULTR1, SULTR2, SULTR3 and SULTR4 (sulfate transporter). The

2- + 2- remaining SO4 transporters are of the animal type (Na /SO4 transporters) and have been designated SLT1, SLT2, SLT3 (SAC1-like transporter). The SULTR transporters encode polypeptides with strong sequence similarity to the SLC26A anion family of

+ 2- transporter polypeptides. Like other plant H /SO4 cotransporters, Chlamydomonas transporters have 10-12 predicted transmembrane domains followed by a carboxy- terminal STAS domain (Figure 1-1). The deduced amino acid sequences of SULTR1

+ 2- and SULTR2 exhibit strong similarity with H /SO4 cotransporters from vascular plants like Arabidopsis, while the deduced amino acid sequence of SULTR3 is more

2- similar to SO4 transporters from bacteria. The gene that was originally designated

SULTR4 was recently shown to encode a molybdate transporter and was renamed

MOT1 (Tejada-Jimenez et al., 2008).

The three SLT genes of Chlamydomonas (SLT1, SLT2, and SLT3) encode

+ 2- polypeptides with strong sequence similarity to mammalian Na /SO4 transporters in the SLC13 family. These transporters have 10-12 predicted transmembrane domains,

+ 2- but unlike the H /SO4 transporters, they do not possess a STAS domain. Instead, all

10 + 2- three putative Na /SO4 transporters contain a TrkA-C domain that has been associated with potassium transport and may bind NAD+. Currently, the TrkA-C domain has no defined physiological or regulatory function (Tucker and Fadool, 2002;

Kraegeloh et al., 2005).

2- D. SO4 assimilation

2- Once SO4 is in the cytoplasm, it has to be translocated into plastids where it is

2- reduced. The pathway for the uptake and reductive assimilation of SO4 is given in

Figure 1-2 and a list of genes involved in this pathway is shown in Table 1-2. A gene

2- encoding a subunit of a Chlamydomonas chloroplast SO4 transporter was initially isolated and designated SULP1; the SULP1 protein appears to be localized to the chloroplast envelop (Chen and Melis, 2004). SULP1 is a transmembrane protein with

2- strong similarity to the CysT subunit of the ABC-type SO4 transporters that are found in bacteria (Chen and Melis, 2004). The SULP1 mRNA increases during S starvation of Chlamydomonas (Chen and Melis, 2004) and a sulP antisense strain

2- shows reduced photosynthesis and lower overall SO4 uptake capacity (Chen et al.,

2- 2005). Other components of the chloroplast SO4 transporter were subsequently identified; these include a second transmembrane protein (SULP2), a nucleotide binding protein (SABC) and a substrate binding protein (SBP) (Lindberg and Melis,

2008). Genes encoding these components (SULP2, SABC, SBP) are also upregulated during S deprivation (Lindberg and Melis, 2008; Gonzalez-Ballester et al., 2010).

2- Prior to reduction, intracellular SO4 is activated by ATP sulfurylase (ATS) to form adenosine 5’-phosphosulfate (APS). ATS isoforms are located in both the plastid

11 and cytosol of plant cells (Rotte and Leustek, 2000). Chlamydomonas appears to have two genes for putative ATP sulfurylases, ATS1 and ATS2, and both of the encoded proteins have potential chloroplast transit peptides (Yildiz et al., 1996; Allmer et al.,

2006). Biochemical work is required to verify the predicted subcellular locations of the two isozymes. Furthermore, the transcripts encoded by ATS1 and ATS2 have been shown to increase during S starvation (Yildiz et al., 1996).

2- The APS generated by ATP sulfurylase can serve as a substrate for SO4 reduction or can be phosphorylated by APS kinase to yield 3’-phosphoadenosine 5’- phosphosulfate (PAPS) (Arz et al., 1994; Lee and Leustek, 1998). While Arabidopsis has four isoforms of APS kinases, the Chlamydomonas genome contains a single gene encoding this enzyme (Kopriva, 2006). PAPS is used in sulfotransferase reactions for the sulfation of various metabolites including flavonols, choline, and glucosides

2- (Varin et al., 1997). In the pathway leading to the reduction of SO4 , APS is reduced to sulfite by APS reductase. APS reductases are located in the plastid and the source of reductant for the enzyme is reduced glutathione (Bick et al., 1998). Like APS kinase,

APS reductase is encoded by a single gene in Chlamydomonas (Kopriva, 2006).

Sulfite generated by the APS reductase reaction is reduced to sulfide by plastid sulfite reductase (SIR), which uses electrons from reduced ferredoxin (Yonekura-Sakakibar et al., 2000). In Chlamydomonas, there are two ferredoxin-type SIR genes (SIR1 and

SIR2) and one bacterial-type SIR gene (SIR3); the latter probably uses NADPH or

NADH as a source of reductant.

12 E. Synthesis of cysteine and methionine

The sulfide formed in the sulfite reductase reaction combines with O- acetylserine (OAS) to generate cysteine. This reaction is catalyzed by O- acetylserine(thiol)lyase (OASTL). OASTL is present in the cytosol, chloroplast, and mitochondrion in Arabidopsis (Lunn et al., 1990). Genes encoding a number of

OASTL isoforms have been identified in various plants (Saito et al., 1994; Gotor et al., 1997; Hesse et al., 1999), with four OASTL genes in Chlamydomonas. The level of transcripts from OASTL4 of Chlamydomonas increase in response to S deprivation

(Ravina et al., 1999; Zhang et al., 2004); transcript levels from the other three putative

OASTL genes have not been characterized.

The formation of OAS, one of the substrates used in the OASTL reaction, is catalyzed by the enzyme serine acetyltransferase (SAT), which is found in the cytosol, chloroplast, and mitochondrion (Ruffet et al., 1995). Cytosolic SAT, which is feedback inhibited at micromolar concentrations of cysteine (Noji et al., 1998), is associated with OASTL and four molecules of pyridoxal 5’-phosphate in the cysteine synthase complex (Bogdanova and Hell, 1997). The isolated complex is inefficient in synthesizing cysteine. Complex formation/disassociation may be a way in which the activities of SAT and OASTL are regulated (Droux et al., 1998). Two SAT genes,

SAT1 and SAT2, both encoding proteins with predicted chloroplast targeting sequences or transit peptides, are present on the Chlamydomonas genome. The levels of the SAT1 transcript and SAT catalytic activity increase in response to S starvation (Ravina et al.,

2002; Zhang et al., 2004).

13 Methionine is synthesized from cysteine and O-phosphohomoserine through three consecutive reactions catalyzed by cystathionine γ-synthase, cystathionine β- lyase and methionine synthase. Cystathionine γ-synthase, which is localized in the chloroplast, catalyzes cystathionine formation (Ravanel et al., 1998; Ravenel et al.,

1998). Cystathionine β-lyase generates homocysteine from cystathionine (Droux et al.,

1995). Both cystathionine γ-synthase and cystathionine β-lyase genes are single copies in Chlamydomonas. The last step of methionine synthesis is catalyzed by methionine synthase, which methylates homocysteine to form methionine using N5- methyltetrahydrofolate as a methyl group donor (Hesse et al., 2004). The

Chlamydomonas genome appears to have two genes encoding methionine synthase,

METE and METH. METE is a cobalamin (vitamin B12)-independent enzyme while

METH is a cobalamin-dependent enzyme. The expression of METE may be controlled by a riboswitch in which cobalamin would binds the RNA and stops its translation

(Croft et al., 2005; Grossman et al., 2007).

F. Synthesis of glutathione

Glutathione is a dominant non-protein thiol in plants and it serves as a major antioxidant. It is a tripeptide composed of glutamate, cysteine, and glycine. Upon oxidation, one reduced glutathione (GSH) can react with another to produce the disulfide form (GSSG). Glutathione serves as a redox buffer and is a substrate of glutathione-S-transferase, which is involved in the detoxification of xenobiotics

(Marrs, 1996). Glutathione is also a precursor of phytochelatins, peptides that play an important role of detoxification of heavy metals (Rauser, 1995). Glutathione synthesis

14 takes place in chloroplasts and is catalyzed by γ-glutamylcysteine synthase and glutathione synthase. Arabidopsis γ-glutamylcysteine synthase and glutathione synthase were isolated by complementation of E. coli mutants (May and Leaver, 1994;

Rawlins et al., 1995). Both γ-glutamylcysteine synthase and glutathione synthase are encoded by single genes in Chlamydomonas.

G. Recycling of S compounds

A fine control of intracellular redistribution of S among different metabolites is important for cells experiencing S deprivation. Recycling of S from proteins requires proteolysis and reutilization of the released cysteine and methionine for de novo protein synthesis; the pathway for S recycling is shown in Figure 1-3. Certain extracellular polypeptides containing very few cysteine and/or methionine are synthesized during S starvation (see below) and can substitute for S-rich proteins that function when the cells are not limited by S (Takahashi et al., 2001). The S-rich proteins may then be degraded and used for the generation organic S metabolites and

2- SO4 , allowing for more effective use of the limiting nutrient. Cysteine dioxygenase

(CDO) catalyzes the conversion of cysteine to 3-sulfinoalanine, a reaction that likely

2- represents the first step in the conversion of cysteine to SO4 . Another enzyme associated with the recycling of cysteine is taurine dioxygenase (TAUD) which catalyzes the conversion of taurine to sulfite and aminoacetaldehyde. While no TAUD homologs have been identified in plants or animals, Chlamydomonas has two TAUD genes (TAUD1 and TAUD2), and the encoded proteins have significant similarity to bacterial TauD (Gonzalez-Ballester et al., 2010). The sulfite generated in the TAUD

15 2- 2- reaction can be oxidized to SO4 by sulfite oxidase (SUOX), and the SO4 redirected toward sulfation through the PAPS pathway (Gonzalez-Ballester et al., 2010).

IV. S STARVATION RESPONSES IN CHLAMYDOMONAS

A. General and Specific responses

Chlamydomonas exhibits a suite of responses when nutrients become limited.

These responses have been described as “general” and “specific.” General responses are associated with deprivation for any essential nutrient and include the cessation of cell growth and division, accumulation of starch and a decline in the rate of photosynthesis. The specific responses are those that are associated only with deprivation of a single nutrient. In the case of S deprivation, the specific responses are mostly associated with activities that promote scavenging of extracellular S and the recycling of intracellular S. These processes include both the synthesis of new high

2- affinity SO4 transport systems and an increase in extracellular ARS activity, which is

2- involved in hydrolyzing SO4 from organic compounds that are in the neighborhood of the cell (de Hostos et al., 1988; Yildiz et al., 1994). Degradation of proteins and lipids that are not essential during S deprivation can also supply cells with a limited amount of S (Ferreira and Teixeira, 1992). It has been shown that S-starved

Chlamydomonas cells degrade 85% of the chloroplast sulfolipid, sulfoquinovosyl diacylglycerol (SQDG), to redistribute the S for protein synthesis and other processes.

The degradation of SQDG precedes that of proteins such as ribulose biphosphate carboxylase/oxygenase and occurs as early as 6 hours after the imposition of S starvation (Sugimoto et al., 2007). In parallel, the synthesis of phosphatidylglycerol,

16 the other acidic lipid in chloroplasts, increases and may compensate for the loss of

SQDG from the membranes; this potential substitution would likely preserve the activities of specific membrane-associated protein complexes, including photosystem I

(Sugimoto et al., 2008).

Furthermore, cells may conserve S by replacing proteins rich in S-containing amino acids with functionally equivalent proteins that have fewer S amino acids

(Gonzalez-Ballester et al., 2010). Examples of this are represented by a group of extracellular proteins (ECPs) and light harvesting proteins (especially the LHCBM) of

Chlamydomonas, and the β-conglycinin storage protein of soybean (Naito et al., 1994;

Takahashi et al., 2001). ECP56, ECP61, ECP76 and ECP88 are extracellular polypeptides synthesized in response to S deprivation. They have features of cell wall proteins, and the mature polypeptides either have one or no S-containing amino acid.

These proteins likely replace some of the S-rich cell wall proteins as the cells experience S deprivation (Takahashi et al., 2001; Gonzalez-Ballester et al., 2010); the cell wall proteins present in S-replete cells may be degraded and the S-containing amino acids redistributed during S deprivation. Similarly, a gene enconding a specific light harvesting protein LHCBM9 is strongly upregulated during S deprivation while there is a loss in transcripts encoding LHCBM1-8 (Nyugen et al., 2008; Gonzalez-

Ballester et al., 2010). These results suggest that the reduction in the synthesis of S- rich LHCBM1-8 polypeptides may be balanced by an increase in S-poor LHCBM9 synthesis. Since LHCBM polypeptides are very abundant in cells, substitution of

17 LHCBM1-8 with LHCBM9 polypeptide would allow significant conservation of S resources (Gonzalez-Ballester et al., 2010).

B. Genes responsive to S deprivation

There are a number of genes in Chlamydomonas that respond to S deprivation.

2- The expression of genes encoding enzymes involved in SO4 acquisition and assimilation, including ATS, SIR, OASTL, and SAT, are induced when cells experience

S limitation (Zhang et al., 2004; Nyugen et al., 2008; Gonzalez-Ballester et al., 2010).

Following the increase in their transcript levels, S-starved Chlamydomonas cells also exhibit elevated SAT and OASTL enzymatic activies (Ravina et al., 2002). Transcripts from ARS and ECP genes also increase in response to S deprivation, and they are rapidly degraded once S-deprived cells are provided with adequate levels of S

(Takahashi et al., 2001). Redistribution and recycling of S among different metabolites is of vital importance to cells experiencing S deprivation. A number of transcripts encoding proteins that may be related to the recycling of S, including CDO1 (encoding cysteine dioxygenase) and TAUD1 (encoding taurine dioxygenase), are induced under

S limitation (Gonzalez-Ballester et al., 2010). S-deprived Chlamydomonas cells exhibit a marked decline in photosynthetic electron transport. Many transcripts encoding proteins associated with photosystem I, photosystem II, light harvesting complexes, and chlorophyll synthesis decline when cells are exposed to S deprivation conditions (Zhang et al., 2004; Gonzalez-Ballester et al., 2010).

The responses of Chlamydomonas to S starvation require a mechanism for sensing S availability and activating a signaling pathway that facilitates survival of the

18 cells in a nutrient-limited environment. We are just beginning to learn about regulatory elements that control S acclimation processes in plants. Recently a transcription factor and miRNAs involved in controlling transcript accumulation when the plants are deprived of S have been identified (Maruyama-Nakashita et al., 2006; Kawashima et al., 2009). SLIM1, an EIL family transcription factor, functions as a central regulator that controls many of the S-deprivation-responsive Arabidopsis genes. Even though expression of SLIM1 itself is not modulated by S availability, SLIM1 regulates the

2- genes encoding proteins required for SO4 acquisition as well as the degradation of glucosinolates (Maruyama-Nakashita et al., 2006). SLIM1 also controls the induction of microRNA-395, the targets of which include APS1, APS3, and APS4 (encoding

2- ATP sulfurylases) and SULTR2;1 (encoding a SO4 transporter) (Kawashima et al.,

2009).

For Chlamydomonas, the screening of thousands of insertional mutants has led to the identification of several regulatory elements required for the proper acclimation of cells to S deprivation (Davies et al., 1994; Davies et al., 1996; Davies et al., 1999;

Pollock et al., 2005). The screen identified Chlamydomonas colonies that were either unable to synthesize ARS during S deprivation or that constitutively expressed ARS activity in S-replete medium. The assay for ARS activity exploited the chromogenic

2- 2- substrate 5-bromo-4-chloro-3-indolyl SO4 (XSO4); cleavage of SO4 from the substrate by ARS results in a blue precipitate. Figure 1-4 shows wild-type

Chlamydomonas colonies grown under S-starved and S-replete conditions, as well as a mutant (designated sac1; see below) that is unable to synthesize ARS in response to S deprivation. Only the wild-type, S-starved cells generate high levels of extracellular

19 ARS and therefore, they are the only colonies on the plates that develop a blue halo within a few hours of being sprayed with XSO4.

C. Identification of genes controlling S deprivation responses

The sac1 (sulfur acclimation) mutant was one of the first mutants identified using the XSO4 screening procedure described above. The SAC1 protein appears to play a central role in controlling S-deprivation responses; this strain exhibits abnormal

2- SO4 uptake activity and is unable to synthesize ARS and other extracellular proteins in response to S deprivation. Essentially no induction of ECP76, ECP88, ATS1 and a

2- number of other genes involved in SO4 acquisition and assimilation was observed in the mutant strain (Yildiz et al., 1996; Takahashi et al., 2001; Zhang et al., 2004). A number of genes associated with photosynthetic electron transport and the amelioration of the damaging effects elicited by accumulation of ROS also change during S deprivation and are under SAC1 control (Zhang et al., 2004). Modification of photosynthetic electron transport during nutrient limitation is critical for cell survival.

Mutants unable to perform these modifications may generate high levels of ROS that could lead to extensive cellular damage. The sac1 mutant became chlorotic and died in the light within 3 days of resuspension in medium devoid of S (Davies et al., 1996).

+ 2- The SAC1 gene product has homology to Na /SO4 transporters (SLT proteins). Its deduced polypeptide sequence and the sac1 mutant phenotype suggest a similarity with the Snf3 system of control in yeast. Snf3 is a transporter-like protein that controls expression of genes involved in hexose utilization (Ozcan et al., 1996;

Ozcan et al., 1998). The similarity between SAC1 and Snf3 raises the possibility that

20 polypeptides whose original function was to bind and transport various substrates into cells may have evolved into regulatory elements that sense intracellular and/or extracellular nutrient concentrations. Like the SLT proteins, the SAC1 polypeptide has

TrkA-C domains positioned in a predicted cytosolic loop that separates two sets of transmembrane helices, as shown in Figure 1-1. The TrkA-C domains of SAC1 may

2- play a role in transducing signals that are triggered by changes in the levels of SO4 in the environment.

Another mutant, designated sac3, exhibits constitutive, low level ARS activity in S-replete medium, but still exhibits an increase in ARS activity upon S starvation.

SAC3 functions either directly or indirectly in maintaining repression of ARS activity when S is available (Davies et al., 1999). Other S-deprivation induced genes may also be negatively regulated by SAC3 (Ravina et al., 2002). Moreover, unlike wild-type cells, the sac3 mutant does not exhibit a decrease in chloroplast transcriptional activities during S starvation, suggesting that SAC3 is required for the inactivation of the chloroplast RNA polymerase sigma factor Sig1 when cells experience S deprivation (Irihimovitch and Stern, 2006). SAC3 encodes a putative serine-threonine kinase of the plant-specific SNRK2 family (SAC3 has also been designated as

SNRK2.2). Thus far, protein targets of the kinase activity of SAC3 have not been identified.

Recently, a number of mutants (ars mutants) that were unable to acclimate properly to S starvation were identified (Pollock et al., 2005). Two allelic mutant strains, ars11 and ars44, have insertions in a gene that encodes a protein which, like

SAC3, is a member of the SNRK2 serine-threonine kinase family; this protein has

21 been designated SNRK2.1 (Gonzalez-Ballester et al., 2008). The ars11 and ars44

2- strains have sac1-like phenotypes: no ARS activity, defective for SO4 uptake, a more rapid decrease in cellular chlorophyll content than wild-type cells during S

2- deprivation, and the inability to induce expression of genes involved in SO4 assimilation. All of these phenotypes are more pronounced in the ars11 mutant than in sac1, which suggests a more central role of SNRK2.1 in controlling the responses of the cells to S starvation. The SNRK2.1 kinase is also critical for S-sparing responses

(replacing S-rich cell wall proteins with ECPs, and light harvesting complex proteins

LHCBM1-8 with LHCBM9) and for restructuring the photosynthetic apparatus under

S-limited conditions. The inability of the ars11 strain to properly manage absorbed excitation energy during S deprivation may result in elevated ROS, which ultimately results in bleaching and rapid death of the mutant strain in the light (Gonzalez-

Ballester et al., 2008; Gonzalez-Ballester et al., 2010).

Some members of the Arabidopsis SNRK2 kinase family also appear to function in controlling S limitation responses. Arabidopsis plants with T-DNA insertions in SNRK2.3 exhibit a decreased induction of SULTR2;2, which encodes a

2- low-affinity SO4 transporter (Kimura et al., 2006).

Another Chlamydomonas mutant with essentially no induction of ARS activity during S starvation is interrupted for a gene encoding a putative adenylyl/guanylyl cyclase (ars401) (Shibagaki, unpublished), one of more than 50 putative proteins of this family in Chlamydomonas (Merchant et al., 2007). This putative adenylyl/guanylyl cyclase may generate cyclic nucleotides that function to amplify responses to stress signals. Another Chlamydomonas mutant that is currently being

22 analyzed may be involved in controlling a staged response of the cells to S deprivation

(see Chapter 4).

D. Model for acclimation of Chlamydomonas to S starvation

A model showing S-dependent signal transduction pathway in

Chlamydomonas during S-replete and S-depleted conditions is shown on Figure 1-5.

This model is based on the information from the analyses of single and double mutants

2- and how each lesion alone or in combination affect ARS and SO4 uptake activities, the expression patterns of S-deprivation-induced genes and the physiological responses of the cells. It is known that SAC1 positively enhances expression of many

S-deprivation-regulated genes when Chlamydomonas is transferred to medium devoid of S. On the other hand, the SAC3 kinase negatively regulates some S starvation- induced genes. A double mutant sac1sac3 exhibits low level constitutive ARS activity in S-rich medium, like the sac3 single mutant, but during S deprivation, unlike sac1, there is some increase in ARS expression although lower than in either wild-type or the sac3 mutant strains (Davies et al., 1994). These data suggest that the epistatic relationship between SAC1 and SAC3 is not clear and that there may be other proteins involved in controlling ARS expression. SAC1, which has significant homology to

+ 2- Na /SO4 transporters, may act as a sensor that resides on the plasma membrane. The

TrkA-C domains present in SAC1 could be involved in protein-protein interactions that trigger a cascade of biochemical changes that modulate the expression of S- regulated genes.

23 The SNRK2.1 protein kinase was also shown to be required for normal manifestation of ARS activity and is a key regulatory component likely to be in the

2- same signaling cascade as SAC1. It has been proposed that SO4 availability sensed by SAC1 initiates a phosphorylation cascade through SNRK2.1 which activates

2- expression of S-responsive genes, including those encoding proteins involved in SO4 uptake and assimilation, and modifications of the photosynthetic apparatus and extracellular space. However, SNRK2.1 activation is not completely SAC1-dependent since the sac1sac3 double mutant still possesses low ARS activity in S-replete and S- depleted conditions. The analysis of the snrk2.1sac3 double mutant demonstrated that

SNRK2.1 is epistatic to SAC3; the phenotype of the snrk2.1sac3 mutant is similar to that of the snrk2.1 single mutant (Gonzalez-Ballester et al., 2008).

Studies that have explored interactions between regulators of S and phosphorus

(P) deprivation responses have also provided new insight into regulatory mechanisms that control S starvation responses. A mutant defective in P deprivation responses, psr1, is unable to synthesize extracellular phosphatases or accumulate high-affinity phosphate transporters during P starvation (Shimogawara et al., 1999b). While wild- type cells do not exhibit ARS activity in P-depleted medium, the P-starved psr1 mutant shows significant level of ARS activity (Moseley et al., 2009). The induction of ARS expression and ARS activity in the psr1sac3 double mutant is greater than that observed in the psr1 single mutant, suggesting that SAC3 inhibits full activation of

ARS in P-starved psr1 cells. These results are consistent with the model that SAC3 acts as a repressor of S-deficiency responsive genes.

24 Moreover, a potential role of SAC1 in post-transcriptional regulation of S deprivation responses was revealed in the same genetic study. While the abundance of the ARS transcript is comparable between the psr1sac3 and the psr1sac1sac3 mutants, the triple mutant exhibits significantly less ARS activity than the psr1sac3 strain. The discrepancy between ARS transcript accumulation and ARS activity suggests that

SAC1 is essential for post-transcriptional regulation of ARS synthesis, activity, or both (Moseley et al., 2009). SAC3 may be critical for modulating the activation state of SNRK2.1, either directly or indirectly. The specific transcription factor (DNA binding protein) that interacts directly with the promoter region of responsive genes has not yet been identified.

V. THESIS AIMS

2- Overall, little is known about how cells regulate their SO4 uptake activity in response to environmental conditions. One of the earliest responses to S deprivation in

2- Chlamydomonas is the induction of high-affinity SO4 transporters. To better

2- understand both the function and regulation of the putative SO4 transporters of

Chlamydomonas, I used both forward and reverse genetic approaches. Chapter two describes a selection for selenate-resistant mutants and their preliminary

2- characterization. In chapter three, I identified genes encoding putative SO4 transporters of Chlamydomonas and isolated mutants that disrupted those putative

2- SO4 transporter genes that were induced in response to S deprivation. The work in

2- this chapter demonstrates the functionality of the transporters in SO4 uptake, their subcellular locations, and the ways in which they are regulated (both at transcriptional

25 and post-transcriptional levels). Chapter four discusses novel aspects of the regulation of S starvation responses and the identification of a regulatory element that may not be critical for the initial responses of the cell to S deprivation, but that may control a second tier of responses. These new discoveries are incorporated into a model that describes S-deprivation elicited regulation in Chlamydomonas and help orient a discussion of the most promising future directions for the research.

Cytoplasm STAS

TrkA-C TrkA-C Cytoplasm

2- Figure 1-1. SO4 transporter structure. (A) Representation of the structure of a

+ 2- H /SO4 transporter (SULTR family) in the membrane, showing 12 transmembrane domains and the carboxy-terminal STAS domain that protrudes into the cytoplasm.

+ 2- (B) Representation of a Na / SO4 transporter (SLT family) with the intracellular

TrkA-C domains.

ARS Ester 2- 2- periplasmic space Ester SO4 SO 4

SLT SULTR plasma membrane

2- SO4 APS ATScyt SBP

SULP1 SULP2

SABC Ser 2- SO4 AcetylCoA ATScp SATcyt GSH APS APK APR OAS GSSG 2- PAPS SO3 Fdred S2- Cys SIR OASTLcyt Sulfation of flavanol,

choline, glucoside Fdox S2-

SATcp Ser OAS OASTLcp

AcetylCoA OASTLm Cys OPH S2- Cys GCS CGS OAS γ-GluCys Glu Cyst

SATm AcetylCoA Gly GSHS CBL hCys Ser GSH mitochondrion MS

Met

chloroplast

28 2- + 2- Figure 1-2. SO4 assimilation pathway. ARS, arylsulfatase; SLT, Na / SO4

+ 2- 2- transporter; SULTR, H /SO4 transporter; SULP1, SULP2, SO4 permease subunit;

2- SBP, SO4 binding protein; SABC, ATP-binding protein; ATS, ATP sulfurylase;

APS, adenosine 5’-phosphosulfate; PAPS, 3’-phosphoadenosine 5’-phosphosulfate;

2- APK, APS kinase; APR, APS reductase; SIR, SO3 reductase; SAT, serine acetyltransferase; Ser, serine; OAS, O-acetylserine; OASTL, O-acetylserine

(thiol)lyase; Cys, Cysteine; OPH, O-phosphohomoserine; CGS, cystathionine γ- synthase; Cyst, cystathionine; CBL, cystathionine β-lyase; hCys, homocysteine; MS, methionine synthase; Met, methionine; Glu, glutamate; GCS, γ-glutamylcysteine synthase; γ-GluCys, γ-glutamylcysteine; Gly, glycine; GSHS, glutathione synthetase;

GSH, glutathione (reduced form); GSSG, glutathione (oxidized form); Fdred, reduced ferridoxin; Fdox, oxidized ferridoxin. Possible localizations are indicated by subscripts: cytoplasm (cyt), chloroplast (cp), mitochondrion (m).

CDO CSAD? Cys 3-sulfinoalanine hypotaurine

non-enzymatic?

taurine

TAUD

2- SO3

SUOX

SO 2- 4

assimilation sulfation

Figure 1-3. Proposed pathway for recycling of S. Cys, cysteine; CDO, Cys dioxygenase; CSAD, sulfoalanine decarboxylase (no gene encoding this enzyme was

2- identified on the Chlamydomonas genome); TAUD, taurine dioxygenase; SO3 ,

2- 2- sulfite; SUOX, SO3 oxidase; SO4 , sulfate.

wild-type sac1

TAP

TAP-S

Figure 1-4. Accumulation of ARS activity in wild-type and sac1 strains. Cells were grown on TAP or TAP –S medium for 7 days and sprayed with XSO4. The color was allowed to develop for 4 hours.

A +S

SAC1

SAC3

SIG1 SNRK2.1

Chloroplast X transcription

S starvation-induced genes

-S B SAC1

P SAC3 SIG1 P SNRK2.1

Chloroplast

transcription P

S starvation-induced genes

32 Figure 1-5. Model for S startvation response signaling pathways in Chlamydomonas during S-replete and S-depleted conditions. A, Under S-replete condition, SNRK2.1 is not active. B, SAC1 likely acts as a sensor for S starvation and activates SNRK2.1 by derepressing SAC3, which is a negative regulator of SNRK2.1. Once SNRK2.1 kinase is activated, it may phosphorylate a transcription factor, X, which subsequently turns on the expression of S deprivation-induced genes, including those encoding enzymes

2- involved in the assimilation of SO4 , restructuring of cell wall and modification of photosynthetic apparatus. The activity of SNRK2.1 kinase might not be completely

SAC1-dependent.

2- SO4 Subcellular Expression Function Reference transporter location pattern SULTR1;1 Plasma Root cortex, High-affinity [A], [B] membrane epidermis, root transporter hairs. Weak responsible for 2- expression in SO4 uptake from hydathodes and soil into roots; Km auxiliary buds of = 3.6 µM leaves. SULTR1;2 Plasma Root epidermis High-affinity [B], [C] membrane and cortex, root transporter hairs. Weak responsible for 2- expression in SO4 uptake from guard cells and soil into roots; Km hydathodes. = 6.9 µM SULTR1;3 Plasma Phloem sieve Source-to-sink [D] membrane element and translocation of 2- companion cell SO4 complexes in roots and hypocotyls. SULTR2;1 Plasma In leaves, xylem Root-to-shoot [A]

membrane parenchyma and transport; Km = phloem cells. In 0.41 mM roots, xylem parenchyma and pericycle cells. 2- SULTR2;2 Plasma Phloems in roots, SO4 distribution [A] membrane vascular bundle from root to shoot. sheath cells in Km > 1.2 mM leaves. SULTR3;1 Plasma Expressed in Cannot [A], [G] membrane leaves complement yeast 2- (based on cells null for SO4 localization transport activity in yeast cells) SULTR3;2 N/A Expressed in Cannot [A] leaves complement yeast 2- cells null for SO4 transport activity SULTR3;3 N/A Expressed in Cannot [A], [I] leaves complement yeast 2- cells null for SO4 transport activity

34 2- SO4 Subcellular Expression Function Reference transporter location pattern SULTR3;4 N/A N/A N/A [H] SULTR3;5 Plasma Xylem Colocalized with [E] membrane parenchyma and and enhance pericycle cells in uptake capacity of roots. SULTR2;1. Facilitating root- to-shoot transport 2- of SO4 . SULTR4;1 Tonoplast Pericycle and Main transporter [F] membrane parenchyma cells facilitating efflux 2- of the vascular of SO4 from tissues in roots; vacuoles vasculatures in hypocotyls. SULTR4;2 Tonoplast Pericycle and Facilitating efflux [F] 2- membrane parenchyma cells of SO4 from of the vascular vacuoles tissues in roots; (supplementary vasculatures, role supporting epidermis, cortical SULTR4;1 cells in function) hypocotyls.

2- Table 1-1. Arabidopsis SO4 transporters: their subcellular locations, expression

2- patterns, and functions. Genes encoding SO4 transporters in bold are induced upon S starvation. References are as follows: [A] Takahashi et al., 2000; [B] Yoshimoto et al.,

2002; [C] Shibagaki et al., 2002; [D] Yoshimoto et al., 2003; [E] Kataoka et al.,

2004a; [F] Kataoka et al., 2004b; [G] Shibagaki and Grossman, 2004; [H] Maruyama-

Nakashita et al., 2006; [I] Hirai et al., 2003. N/A, not available.

Activity Gene Arylsulfatase ARS1 ARS2 + 2- Na /SO4 transporter SLT1 SLT2 SLT3 + 2- H /SO4 transporter SULTR1 SULTR2 SULTR3 2- Chloroplast SO4 transport system SULP1 SULP2 SBP1 SABC ATP sulfurylase ATS1 ATS2 APS kinase APK1 APS reductase MET16 2- SO3 reductase SIR1 SIR2 SIR3 O-acetylserine (thiol)lyase OASTL1 OASTL2 OASTL3 OASTL4 Serine acetyl transferase SAT1 SAT2 Cystathionine γ-synthase CGS1, METB Cystathionine β-lyase METC Methionine synthase METE METH γ-Glutamylcysteine synthase GSH1 Glutathione synthetase GSH2

2- Table 1-2. Genes involved in SO4 assimilation in Chlamydomonas.

36 REFERENCES

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CHAPTER 2

SELENATE RESISTANT MUTANT SELECTION IN CHLAMYDOMONAS

REINHARDTII

44 ABSTRACT

Upon S starvation of wild-type Chlamydomonas reinhardtii (Chlamydomonas) cells, one of the earliest responses observed is an increase in efficiency of sulfate

2- 2- anion (SO4 ) import. Features of SO4 transport during S-replete and S-depleted conditions have been reported, although there has been little molecular

2- characterization of the genes and proteins encoding putative SO4 transporters. An

2- approach often employed for identification of SO4 transporters is the selection of

2- mutants that are resistant to selenate, a toxic analog of SO4 . This strategy has been

2- successfully used to identify strains containing lesions in SO4 transporter genes from

Saccharomyces cerevisiae and Arabidopsis thaliana. A similar approach was used to select for Chlamydomonas mutants that were capable of growing on medium containing levels of selenate that are toxic to wild-type cells. Approximately 60,000 primary transformants were selected and 12 strains showed a significantly higher level of selenate resistance compared to wild-type cells. Unfortunately, none of the selenate-

2- resistant strains exhibited aberrant SO4 transport activity, suggesting that there could

2- be some genetic redundancy in genes encoding components of the SO4 uptake system. The selenate-resistant mutants that were obtained could have acquired the ability to detoxify or sequester selenate, thereby preventing this anion from being

2- nonspecifically incorporated into amino acids and proteins. Furthermore, the SO4 transporters of Chlamydomonas may be extremely inefficient in importing selenate since Chlamydomonas synthesizes selenoproteins and may have evolved a selenate- specific transporter.

45 INTRODUCTION

Sulfur (S) is an essential nutrient for all organisms including plants and algae.

It is incorporated into proteins (as cysteine and methionine), lipids, and

2- polysaccharides. The dominant, fully oxidized form of S, sulfate (SO4 ), is the most

2- 2- stable and the preferred form of S. SO4 is taken into cells by SO4 transporters on the plasma membrane and subsequently translocated into plastids where it is reduced and incorporated into cysteine and methionine. The first step in reductive assimilation is

2- the activation of SO4 by ATP sulfurylase to form adenosine 5’-phosphosulfate (APS)

(Leustek et al., 2000). Chlamydomonas reinhardtii (Chlamydomonas throughout) appears to have two genes encoding ATP sulfurylases based on the genome sequence, and at least one isoform is likely to be targeted to the chloroplast (Gonzalez-Ballester

2- 2- and Grossman, 2009). The SO4 moiety of APS is reduced to sulfite (SO3 ) by APS

2- reductase using glutathione as an electron donor. Subsequently, SO3 is reduced to

2- sulfide by plastid ferredoxin-dependent SO3 reductase. The reduction of APS to

2- 2- SO3 and of SO3 to sulfide takes place exclusively in plastids. The sulfide is then combined with O-acetylserine to form cysteine in a reaction catalyzed by O- acetylserine(thiol)lyase (Leustek et al., 2000).

Chlamydomonas exhibits a suite of responses when S becomes limiting. These responses include the synthesis of extracellular arylsulfatases (ARS) and increased

2- 2- SO4 uptake. S-deprived Chlamydomonas cells synthesize new, high-affinity SO4 transport systems that can be detected in cells within an hour of S deprivation (Yildiz

2- et al., 1994). While early work focused on the characterization of SO4 transport in

46 2- Chlamydomonas, none of the SO4 transporters had been identified and characterized at the molecular level.

2- A genetic approach that is often used to identify SO4 transporters involves the

2- selection of mutants that are resistant to selenate, a toxic analog of SO4 . It has been

2- 2- proposed that both SO4 and selenate anions are taken up by SO4 transporters.

Selenate uptake in many organisms, including Escherichia coli and Saccharomyces

2- cerevisiae, was shown to be mediated by a SO4 transporter (Lindblow-Kull et al.,

2- 1985; Cherest et al., 1997). Both of the high-affinity SO4 transporters in S. cerevisiae

(SUL1 and SUL2) were demonstrated to facilitate selenate uptake (Cherest et al.,

2- 1997). E. coli employs the same transport system for SO4 , selenite and selenate

2- uptake, but there is a greater specificity and higher affinity for SO4 than for either of the two selenium (Se) compounds (Lindblow-Kull et al., 1985).

Se is an oxygen group element with chemical property similar to S although the Se atom is slightly larger. Like S, Se exists in several different oxidation states as selenide (Se2-), elemental selenium (Se), selenite (Se4+), and selenate (Se6+). Se is an essential trace element for animals and bacteria, but whether it is essential for plants remains controversial (Minorsky, 2003). Several Se-dependent enzymes have been identified in which an integral selenocysteine (SeCys) residue is inserted into the catalytic site (Stadtman, 1990). Mutagenesis studies have shown that SeCys residues play a critical role as replacement of the active site SeCys by a cysteine (Cys) residue greatly reduces catalytic activity (Axley et al., 1991). The incorporation of the active site SeCys into these essential selenoproteins is a cotranslational process directed by a

UGA codon. UGA normally functions as a universal termination codon; in order for

47 UGA to function as a SeCys codon, a special incorporation system is required (Boeck et al., 1991).

All known eukaryotic SeCys-containing proteins are animal proteins, and so far selenoproteins have not been found in yeast and plants. When the genomes of S. cerevisiae and Arabidopsis thaliana (Arabidopsis throughout) were sequenced, their analyses revealed neither selenoprotein genes nor any of the components of the SeCys incorporation pathway. Recently, the presence of selenoproteins in Chlamydomonas has been demonstrated by a radioactive labeling experiment (with 75Se). Se was demonstrated to be essential for optimal growth of Chlamydomonas, however the requirement for Se is very low, possibly in the nanomolar or subnanomolar range

(Novoselov et al., 2002). Ten selenoproteins were identified in Chlamydomonas; some are common selenoproteins found in other eukaryotes (such as glutathione peroxidase) and some are unique to this organism (such as methionine-S-sulfoxide reductase).

Interestingly, the SeCys incorporation sequences found in selenoprotein genes in

Chlamydomonas are similar to those of animals, suggesting that selenoproteins evolved early and were independently lost in yeast, land plants, and some animals (Fu et al., 2002; Novoselov et al., 2002).

Se is metabolized via the S assimilation pathway. Once inside the cells, selenate is activated by ATP sulfurylase to adenosine phosphoselenate (APSe), an activated form of selenate. ATP sulfurylase has been shown to activate selenate, as

2- well as SO4 both in vitro and in vivo (Shaw and Anderson, 1972; Pilon-Smits et al.,

1999). APSe is subsequently reduced to selenite and selenide by APS reductase and

2- SO3 reductase, respectively (Dilworth and Bandurski, 1977; Pilon-Smits et al., 1999).

48 SeCys is formed by the action of O-acetylserine(thiol)lyase, which couples selenide with O-acetylserine (Ng and Anderson, 1978). SeCys is metabolized to selenomethionine (SeMet) via three enzymatic steps catalyzed by cystathionine γ- synthase, cystathionine β-lyase and methionine synthase forming selenocystathionine, selenohomocysteine and SeMet, respectively (Terry et al., 2000).

The major mechanism whereby high intracellular selenium concentrations induce toxicity is associated with the incorporation of SeCys and SeMet into proteins in place of cysteine and methionine. The difference in size and ionization properties of

S and Se may result in significant alterations in protein structure. The bond between two Se atoms is approximately one seventh longer and one fifth weaker than the disulfide bond (Brown and Shrift, 1982). Therefore, the incorporation of SeCys in place of cysteine into proteins could interfere with the disulfide bridge formation, resulting in altered polypeptide structure, that could in turn impact the catalytic activity of the protein (Brown and Shrift, 1982).

Identification of selenate-resistant strains has been invaluable in elucidating

2- SO4 transport and assimilation mechanisms in S. cerevisiae, Neurospora crassa, and

Arabidopsis (Cherest et al., 1997; Marzluf, 1970; Shibagaki et al., 2002). In order to

2- identify SO4 transporters in Chlamydomonas, I performed insertional mutagenesis to generate a mutant library and selected for strains capable of growing on selenate-

2- containing medium. Some of these strains were expected to be defective in SO4 uptake.

49 RESULTS

Generation of a mutant library and selection for selenate resistant strains: To generate a pool of insertional mutants, a Chlamydomonas cell-wall-deficient D66 strain was transformed by electroporation with a 1.7 kb PCR fragment containing the

AphVIII gene (encoding an aminoglycoside 3'-phosphotransferase conferring paromomycin resistance) under the control of the PSAD promoter (Figure 2-1). After an overnight recovery of the electroporated cells, transformants were plated directly

2- onto medium devoid of SO4 , but supplemented with 2 mM thiocyanate, a poor S source, 6 µM sodium selenate and 5 µg mL-1 paromomycin. The selenate concentration was the minimum required to kill paromomycin-resistant strains in the

D66 background grown on TAP-S medium. To assess the transformation efficiency, transformants were also plated onto TAP-S supplemented with 2 mM thiocyanate and

5 µg mL-1 paromomycin. Colonies capable of growing on selenate-containing medium became visible on the plates after 2-3 weeks. Thirty-one such colonies were identified from approximately 60,000 primary transformants.

To confirm the selenate resistant phenotype of these 31 sr (selenate resistant) mutants, a spot test was performed on TAP-S medium supplemented with 10 µM sodium selenate (the selenate concentration was increased from 6 µM to 10 µM because this medium did not contain paromomycin). Twelve of the sr mutants from the primary selection showed varying degrees of selenate resistance and the rest appeared to be false positives (their selenate resistant level was not significantly different from that of the parental strain) (Figure 2-2). The sr mutants were observed at a frequency of ~1 in 5000. sr103, sr111, and sr125 mutants exhibited high level of

50 selenate resistance while other mutants such as sr138 and sr140 showed a lower degree of resistance. DNA gel blot analysis was carried out to determine the number of copies of AphVIII that were inserted into the genomes of the mutant strains.

Approximately 60% of the mutants contained a single insertion locus and the rest have two AphVIII insertion loci (Figure 2-3). It should be noted that this analysis would not identify mutants in which there were multiple insertions at a single locus.

2- SO4 uptake by the sr mutant strains: Mutants capable of growing on the selenate-

2- containing medium are expected to be defective for SO4 uptake. To assess whether

2- 2- the lesions in sr mutants affected the rate of SO4 transport, the SO4 uptake rates in

2- S-starved strains were measured at an external SO4 concentration of 250 µM (the uptake rate measured at this concentration is close to the maximum velocity).

2- Unfortunately, none of the sr mutants exhibited aberrant SO4 transport activity

2- during S deprivation, suggesting that the inducible SO4 uptake systems in these mutants were not defective (Table 2-1). These sr mutants might have acquired an enhanced ability to detoxify selenate. Alternatively, Chlamydomonas may have

2- transporters that are specific for selenate; it does not get into cells via SO4 transporters.

Other S-deprivation responses in the sr mutants: S-starved Chlamydomonas cells

2- normally synthesize extracellular ARS, which cleaves SO4 from organic molecules,

2- which makes the SO4 available for uptake. The level of ARS activity in the sr mutants was measured in both S-replete and S-depleted conditions. Like in wild-type cells, ARS activity was not detected in any of the mutants maintained in S-rich medium. sr116 exhibited a significantly higher ARS activity than the wild type while

51 other sr mutants (for example, sr111) showed a normal level of ARS activity during S deprivation (Figure 2-4). The sr116 mutant was crossed to a wild-type strain of an opposite mating type and the progeny showed co-segregation of the paromomycin resistant phenotype with elevated ARS activity when grown on –S medium. However,

2- the sr116 mutant did not appear to be selenate-resistant and its SO4 uptake rate under

S-depleted condition was normal (Figure 2-2 and Table 2-1). Due to time limitation, I did not continue working on the sr116 mutant to identify the lesion that was responsible for the high ARS activity phenotype.

DISCUSSION

2- Selenate readily competes with the uptake of SO4 and it has been shown that

2- in some organisms both anions are taken up via SO4 transporters (Lindblow-Kull et al., 1985). Selenate resistant selection has successfully been used to identify mutants

2- containing lesions in SO4 transporter genes in S. cerevisiae and A. thaliana (Cherest

2- et al., 1997; Shibagaki et al., 2002). In an attempt to identify SO4 transporters in

Chlamydomonas, a similar approach was employed to select for selenate resistant mutants. Mutants capable of growing on the selection medium are expected to be

2- defective for SO4 uptake. Twelve selenate resistant mutants were identified from

~60,000 primary transformants. The level of selenate resistance of each mutant was quantified semi-quantitatively by a spot test (Figure 2-2). Some sr mutants are more resistant to selenate than others, but unfortunately, all of mutants identified exhibit

2- normal SO4 transport during S deprivation (Table 2-1), suggesting that decreased

2- SO4 (and presumably selenate) uptake is not the mechanism by which these mutants

52 cope with high selenate concentration in the medium. It is possible that some sr mutants have acquired the ability to detoxify selenate (i.e. conversion to a non-toxic form) while others may efficiently sequester and transport selenate into vacuoles to prevent it from being assimilated into SeCys and SeMet and subsequently incorporated into proteins. Alternatively, some of the mutants may become more effective at transporting selenate out of the cell than the parental strain.

The rationale for performing the selenate resistant mutant selection was based

2- on the assumption that selenate is taken up by SO4 transporters. Even though this has been demonstrated for S. cerevisiae and A. thaliana (Cherest et al., 1997; Shibagaki et al., 2002), it might not be the case in Chlamydomonas. While there is no requirement for Se in yeast and higher plants, there is evidence that Se is required for the growth of

Chlamydomonas (Novoselov et al., 2002). Despite the fact that this Se requirement is very low, Chlamydomonas might have developed a set of membrane proteins specific for selenate uptake and, unlike the circumstances in yeast or plants, selenate may not

2- be transported into cells through SO4 transporters. Alternatively, selenate could be

2- taken up by constitutive SO4 transporters (which may be down-regulated during S deprivation; see Chapter 3) as opposed to the inducible transporters that function primarily under S starvation. Mutants harboring lesions in the constitutive transporters

2- would not be identified in this selection as most of the SO4 transport that occurs under S-deprivation conditions (Yildiz et al., 1994), conditions used to select for the mutants in this study, would be performed by the inducible transporters.

SeCys-containing proteins have been identified in bacteria, archaea and eukaryotes. They have not been found in yeast and plants. A number of selenoproteins

53 (including glutathione peroxidase 1 and the components of the selenocysteine insertion system) have been identified in Chlamydomonas (Novoselov et al., 2002).

Because Se is normally required in the nanomolar range, high concentrations of selenate are still toxic even to Se-requiring organisms. Chlamydomonas might have evolved a strategy to regulate the amount of selenate transported into cells as well as a means to cope with excess intracellular selenate. A number of sr mutants isolated may contain lesions in genes encoding proteins that are involved in these pathways. For example, in the Chlamydomonas genome there is a gene encoding a putative selenobinding protein that may be important for sequestering selenate for detoxification. If this is the case, enhanced expression of this gene might allow cells to tolerate a higher level of intracellular selenate. Interestingly, none of the sr mutants exhibited an increase in accumulation of the SBDP transcript (encoding a selenobinding protein) compared to wild-type cells.

Genetic redundancy could also explain why none of the selenate resistant

2- 2- mutants exhibit aberrant SO4 uptake activity. There are six putative SO4 transporters in the Chlamydomonas genome and the mutation in a single transporter may not affect

2- 2- the SO4 transport. The probability of having lesions in multiple SO4 transporter genes is extremely low given that most mutants carried one or two AphVIII insertions

(Figure 2-3). Mutants harboring mutations in genes encoding regulators controlling

2- the expression of SO4 transporter genes during S deprivation may be able to grow on selenate-containing medium. However, such mutants may also be sensitive to S- limited condition and fail to survive on –S medium supplemented with selenate. For example, a mutant bearing a lesion in SNRK2.1, which encodes a central regulator of S

54 2- deprivation responses, shows no increase in the rate of SO4 transport under S limitation (see Chapter 3). One would expect this mutant to be able to survive on selenate containing S-deficient medium. However, besides its inability to induce high-

2- affinity SO4 transport, snrk2.1 mutant is also defective in other aspects of S deficiency responses (such as S scavenging and S sparing) and as a consequence, the mutant cannot survive for a long period of time on –S medium.

2- Some selenate resistant mutants may carry lesions in genes encoding SO4 assimilation enzymes. An inability of such mutants to incorporate selenate into SeCys and SeMet may protect the cells from the deleterious effects of intracellular selenate accumulation. In S. cerevisiae, a number of strains that are resistant to selenate have lesions in the MET3 gene, which encodes ATP sulfurylase and catalyzes the first step

2- of SO4 reduction (Smith et al., 1995; Cherest et al., 1997). As previously shown, yeast cells bearing mutations in genes encoding APS reductase and APS kinase are also capable of growing in the presence of relatively high levels of selenate compared to wild-type cells (Cherest et al., 1997). The sac1 mutant is unable to activate genes

2- associated with the reduction and assimilation of SO4 and it was shown to be considerably more resistant to selenate than the parental strain (Pollock et al., 2005).

The primary goal of the selection of selenate resistant strains was to identify

2- mutants defective for SO4 uptake. Since none of the sr mutants demonstrated reduced

2- SO4 uptake activity as expected, I did not identify the genes altered in those strains even though it would be interesting to investigate the mechanisms that allow the mutants to grow in the presence of high selenate concentration.

55 MATERIALS AND METHODS

Strains and growth conditions: Strain D66 (nit2-, cw15, mt+) (Pollock et al., 2003) was used as a parental strain for the mutagenesis. For transformation experiments, cells were grown to mid-logarithmic phase (2-4 x 106 cells mL-1) in Tris-acetate- phosphate (TAP) medium under continuous light (80 µmol photon m-2 s-1). TAP–S medium was prepared as described previously (Davies et al., 1994).

Generation and selection of selenate resistant mutants : To generate a library of insertional mutants, a 1.7-kb PCR fragment containing a selectable marker AphVIII

(conferring resistance to paromomycin) (Sizova et al., 2001) under the control of

PSAD promoter and terminator was used for transformation. This fragment was amplified with a forward primer (5’-ACCAATCGTCACACGAGC-3’) and a reverse primer (5’-CTTTCCATCGGCCCAGCAAC-3’) using the pSL18 plasmid as a template (Sizova et al., 2001). PCR was performed as follows: pre-incubation at 95°C for 5 min, followed by 35 cycles of denaturation at 95°C for 30 s, annealing at 52°C for 30 s, elongation at 72°C for 2 min. The PCR product was purified with a QiaQuick

PCR Purification Kit (Qiagen). The cell wall-deficient strain D66 was transformed by electroporation (Shimogawara et al., 1999) using a modification procedure reported by

Colombo et al. (2002). Briefly, the D66 strain was grown to a density of 4 x 108 cells mL-1, concentrated by centrifugation (2500 X g, 5 min) and then resuspended in TAP medium supplemented with 40 mM sucrose to a density of 4 x 108 cells mL-1. 200 µl of concentrated cells was transferred into an electroporation cuvette with 0.4 cm gap

(VWR), mixed with 200 ng of AphVIII DNA, and incubated at 16°C for 10 min before electroporation (set at 10 µF; Biorad). Electroporated cells were allowed to recover

56 overnight under low light in 10 mL of TAP medium containing 40 mM sucrose and plated the following day onto solid TAP-S medium containing 2 mM sodium thiocyanate, 6 µM selenate and 5 µg mL-1 paromomycin (Pollock et al., 2005).

Transformation efficiency was evaluated by selecting transformants on TAP-S medium supplemented with 5 µg mL-1 paromomycin. The level of selenate resistance of each mutant was confirmed by a spot test.

Southern Blot Analyses: Genomic DNA was isolated from 50 mL liquid cultures of the D66 parental strain and selenate resistant mutants using a standard phenol- chloroform extraction protocol (Sambrook et al., 1989). 10 µg of genomic DNA was digested for 20 h with 10 u of restriction endonucleases (PstI, PvuII, or SacI; New

England Biolabs). The fragments were separated by gel electrophoresis (0.7% agarose) at 30V for 20 h. The digested DNA was blotted overnight onto nylon membranes (GeneScreen, DuPont-New England Nuclear) using 20X SSC as a transfer buffer, and the transferred DNA was cross-linked to the membrane by UV illumination. A fluorescein-labelled probe was synthesized by random primer labeling in the presence of fluorescein-conjugated-dCTP. Probe synthesis and hybridization were performed using the Amersham AlkPhos DirectTM Labeling and Detection

Systems following the manufacturer’s protocol.

2- 2- SO4 uptake assays: SO4 transport assays were performed as previously described

(Yildiz et al., 1994). Exponentially growing cells were harvested and washed once with TAP-S medium, and resuspended in the same medium to a concentration of 4 x

106 cells mL-1. An aliquot of 1.5 mL was placed in a water-jacketed chamber

-2 -1 2- maintained at 27ºC and illuminated at an intensity of 250 µmol photon m s . SO4

57 uptake was initiated by the addition of MgSO4 (final concentration 250 µM)

35 2- containing 50 µCi of SO4 (Perkin Elmer). At various times following the addition

35 2- of SO4 (20, 40, 60, 80, and 100 s), 100-µL aliquots were withdrawn from the uptake mixture and filtered onto a 45 µm nitrocellulose membrane (Millipore). Cells were washed with ice-cold TAP-S medium supplemented with 1 mM MgSO4.

Following the wash, the filters was transferred to a scintillation vial and dissolved in 4

2- mL of EcoLume scintillation cocktail (MP Research). SO4 uptake was plotted as a function of time and the rate was calculated from the slope of the regression line.

Assay for ARS activity: Liquid assays for ARS activities were performed as described in de Hostos et al., 1988. 50 µL of S-starved samples were added to 500 µL of 0.1 M glycine-NaOH, pH 9, 10 mM imidazole, 4.5 mM p-nitrophenylsulfate

(Sigma, St.Loius, MO) and incubated for 10 min at 27ºC. The reaction was stopped by the addition of 20 µL of 10 M NaOH and the 410 nm absorbance measured. This assay was performed on whole cultures, and cells were sedimented prior to determining the absorbance.

Transform D66 with AphVIII construct

PSAD promoter AphVIII

Select for selenate resistant (sr) mutants Assess transformation efficiency on on TAP-S + paromomycin + selenate TAP-S + paromomycin

Confirm/quantify the selenate resistance of each mutant by a spot test

2- Check SO4 Determine the copy uptake rate in the number of AphVIII sr mutants insertion

Figure 2-1. Schematics showing steps in protocol use to select for selenate resistant mutants.

59 TAP –S TAP –S + 10 µM selenate

D66 D66

sr101 sr101

sr102 sr102 sr103 sr103

D66 D66

sr101 sr101

sr104 sr104

sr106 sr106

D66 D66 sr107 sr107 sr108 sr108

sr109 sr109

sr110 sr110 sr114 sr114

D66 D66 sr111 sr111

sr113 sr113

sr114 sr114 sr123 sr123

60 TAP –S TAP –S + 10 µM selenate

D66 D66 sr112 sr112

sr113 sr113

sr115 sr115 sr116 sr116

sr120 sr120

D66 D66

sr122 sr122

sr123 sr123

sr124 sr124

sr125 sr125 sr126 sr126

D66 D66

sr127 sr127

sr129 sr129

sr130 sr130

sr131 sr131

sr134 sr134

D66 D66

sr135 sr135

sr137 sr137

sr138 sr138

sr139 sr139

sr140 sr140

61 Figure 2-2. Secondary screening of sr mutants. D66 and sr mutants (from the primary selection) were grown to mid-logarithmic phase and washed with TAP-S medium.

Subsequently, cells were serially diluted and grown on TAP-S containing 0 or 10 µM sodium selenate for 7 and 21 days, respectively.

62 A

10 8

6 5

4 3 2 Endogenous PSAD gene 1

10 8

6 Endogenous 5 PSAD gene

4 3 2

10 8

6 5

4 3

2 Endogenous PSAD gene 1

63 Figure 2-3. DNA gel blot analyses of insertional mutants. Arrows indicate bands corresponding to endogenous PSAD sequence. Molecular weights (in kilobases) are shown to the left. Genomic DNA from wild-type (D66) and sr mutants was digested with PstI (A, C) or PvuI (B) and the probe was the PSAD::AphVIII fragment amplified from pSL18.

0.45 WT sr111 sr116 0.4

0.35

0.3 0.25

0.2

0.15

0.1

0.05

Arylsulfatase activity (arbitrary units) units) activity (arbitrary Arylsulfatase 0 1h -S 18h -S 24h -S 42h -S 72h -S

Figure 2-4. Liquid culture assays for ARS activities in TAP and after 18, 24, 42,

and 72 h of growth in TAP-S medium for wild type (WT), sr111, and sr116. Data

points are average of triplicate measurements and error bars represent one standard

deviations.

2- SO4 uptake rate Strain 2- 5 (fmol SO4 per 10 cells) D66 389 ± 24 (parental strain) sr103 336 ± 46 sr104 391 ± 67 sr106 315 ± 39 sr111 307 ± 27 sr112 367 ± 12 sr114 348 ± 19 sr116 345 ± 37 sr121 376 ± 35 sr125 320 ± 27 sr129 314 ± 82 sr130 401 ± 49 sr138 351 ± 41 sr140 320 ± 14

2- Table 2-1. Characteristics of SO4 transport in wild-type cells (D66) and sr mutants in

S-depleted (24 h starved) conditions. Values are averages and standard errors of 2-4 replicates.

66 REFERENCES

Axley, M.J., Boeck, A., and Stadtman, T.C. (1991). Catalytic properties of an Escherichia coli formate dehydrogenase mutant in which sulfur replaces selenium. Proc. Natl. Acad. Sci. U.S.A. 88. Boeck, A., Forchhammer, K., Heider, J., and Baron, C. (1991). Selenoprotein synthesis: an expension of the genetic code. Trends Biochem. Sci. 16, 463-467. Brown, T.A., and Shrift, A. (1982). Selenium: toxicity and tolerance in higher plants. . Biol. Rev. 57, 59-84. Cherest, H., Davidian, J.-C., Thomas, D., Benes, V., Ansorge, W., and Surdin-ker Jan, Y. (1997). Molecular characterization of two high affinity sulfate transporters in Saccharomyces cerevisiae. Genetics 145, 627-637. Davies, J.P., Yildiz, F., and Grossman, A.R. (1994). Mutants of Chlamydomonas with aberrant responses to sulfur deprivation. Plant Cell 6, 53-63. Dilworth, G.L., and Bandurski, R.S. (1977). Activation of selenate by adenosine 5'- triphosphate sulfurylase from Saccharomyces cerevisiae. . Biochem. J. 163, 521-529. Fu, L.H., Wang, X.F., Eyal, Y., She, Y.M., Donald, L.J., Standing, K.G., and Ben- Hayyim, G. (2002). A selenoprotein in the plant kingdom. Mass spectrometry confirms that an opal codon (UGA) encodes selenocysteine in Chlamydomonas reinhardtii gluththione peroxidase. J Biol Chem 277, 25983-25991. Gonzalez-Ballester, D., and Grossman, A.R. (2009). Sulfur: From acquisition to assimilation. (Amsterdam, The Netherlands: Elsevier). Leustek, T., Martin, M.N., Bick, J.A., and Davies, J.P. (2000). Pathways and regulation of sulfur metabolism revealed through molecular and genetic studies. Annu. Rev. Plant Physiol. Plant Mol. Biol. 51, 141-165. Lindblow-Kull, C., Kull, F.J., and Shrift, A. (1985). Single transporter for sulfate, selenate, and selenite in Escherichia coli K12. . J. Bacteriol. 163, 1267-1269. Minorsky, P.V. (2003). SELENIUM IN PLANTS. Plant Physiol 133, 14-15. Ng, B.H., and Anderson, J.W. (1978). Synthesis of selenocysteine by cysteine synthases from selenium accumulator and non-accumulator plants. Phytochemistry 17, 2069-2074. Novoselov, S.V., Rao, M., Onoshko, N.V., Zhi, H., Kryukov, G.V., Xiang, Y., Weeks, D.P., Hatfield, D.L., and Gladyshev, V.N. (2002). Selenoproteins and selenocysteine insertion system in the model plant cell system, Chlamydomonas reinhardtii. Embo J 21, 3681-3693. Pilon-Smits, E.A.H., Hwang, S., Lytle, C.M., Zhu, Y., and Tai, J.C. (1999). Overexpression of ATP slufurylase in Indian mustard leads to increased selenate uptake reduction, and tolerance. . Plant Physiol 119, 123-132. Pollock, S.V., Colombo, S.L., Prout, D.L., Godfrey, A.C., and Moroney, J.V. (2003). Rubisco activase is required for optimal photosynthesis in the green alga Chlamydomonas reinhardtii in a low-CO2 atmosphere. Plant Physiology 133, 1854-1861.

67 Pollock, S.V., Pootakham, W., Shibagaki, N., Moseley, J.L., and Grossman, A.R. (2005). Insights into the acclimation of Chlamydomonas reinhardtii to sulfur deprivation. Photosynth Res 86, 475-489. Sambrook, J., Fritsch, E.F., and Maniatis, T. (1989). Molecular Cloning: A Laboratory Manual. (Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press). Shaw, W.H., and Anderson, J.W. (1972). Purification, properties and substrate specificity of adenosine triphosphate sulphurylase from spinach leaf tissue. Biochem. J. 127, 237-247. Shibagaki, N., Rose, A., McDermott, J.P., Fujiwara, T., Hayashi, H., Yoneyama, T., and Davies, J.P. (2002). Selenate-resistant mutants of Arabidopsis thaliana identify Sultr1;2, a sulfate transporter required for efficient transport of sulfate into roots. Plant J 29, 475-486. Shimogawara, K., Fujiwara, S., Grossman, A.R., and Usuda, H. (1999). High- efficiency transformation of Chlamydomonas reinhardtii by electroporation Genetics 148, 1821-1828. Sizova, I., Fuhrmann, M., and Hegemann, P. (2001). A Streptomyces rimosus aphVIII gene coding for a new type phosphotransferase provides stable antibiotic resistance to Chlamydomonas reinhardtii. Gene 277, 221-229. Smith, F.W., Hawkesford, M.J., Prosser, I.M., and Clarkson, D.T. (1995). Isolation of a cDNA from Saccharomyces cerevisiae that encodes a high affinity sulfate transporter at the plasma membrane Mol. Gen. Genet. 247, 709- 715. Stadtman, T.C. (1990). Selenium Biochemistry. Annu. Rev. Biochem. 59, 111-128. Terry, N., Zayed, A.M., de Souza, M.P., and Tarun, A.S. (2000). Selenium in higher plants. Annu. Rev. Plant Biol. 51, 401-432. Yildiz, F.H., Davies, J.P., and Grossman, A.R. (1994). Characterization of sulfate transport in Chlamydomonas reinhardtii during sulfur-limited and sulfur- sufficient growth. Plant Physiol 104, 981-987.

CHAPTER 3

THE SULFATE TRANSPORTERS OF CHLAMYDOMONAS REINHARDTII;

FROM REGULATION TO FUNCTIONALITY

69 ABSTRACT

Chlamydomonas reinhardtii (Chlamydomonas throughout), a unicellular green alga, exhibits several responses following exposure to sulfur (S)-deprivation conditions.

These responses include the synthesis of extracellular arylsulfatases (ARS), potential restructuring of the S-rich cell wall and photosynthetic apparatus, and an increased

2- 2- efficiency of import and assimilation of the sulfate anion (SO4 ). Aspects of SO4 transport during S-replete and S-depleted conditions have previously been studied,

2- although the transporters responsible for the passage of SO4 across the plasma membrane and into the cytoplasm of the cell have not been functionally identified.

2- Here, we employed a reverse genetic approach to identify putative SO4 transporters, establish their subcellular localizations, characterize their biosynthetic/regulatory

2- features and demonstrate their functionality. Six putative SO4 transporters were identified based on sequence similarity to known transporters from both plants and animals. Under conditions in which the cells were starved for S, transcripts encoding three of these transporters, SLT1, SLT2, and SULTR2, markedly increased.

Furthermore, increased accumulation of the SLT1, SLT2, and SULTR2 transcripts during S deprivation was closely followed by de novo synthesis of the encoded

2- proteins, suggesting that the putative transporters function as high-affinity SO4

2- transporters that allow the cells to efficiently scavenge SO4 from the environment.

All three transporters were shown to be localized to the plasma membrane. The half- lives of the transporter polypeptides were significantly impacted by S availability, and degradation of the two SLT transporters was markedly inhibited by a reagent that blocks the action of the proteasome; SULTR2 showed no sensitivity to this inhibitor.

70 The Chlamydomonas sac1 and snrk2.1 mutant strains (defective for acclimation to S deprivation) exhibited much less of an increase in levels of SLT1, SLT2 and SULTR2 transcripts and polypeptides in response to S deprivation compared to wild-type cells.

Finally, mutants disrupted for the individual transporter genes were identified. The slt1, slt2, and sultr2 single mutants were barely affected in their ability to transport

2- SO4 as the cells became S deprived (possibly a consequence of functional redundancy among the gene products). However, S-deprived strains deficient for two of the transporters (slt1 and slt2, slt1 and sultr2, slt2 and sultr2) were significantly

2- 2- reduced for their rates of SO4 uptake, while there was essentially no increase in SO4 uptake following S-deprivation of the slt1slt2sultr2 triple mutant.

71 INTRODUCTION

Sulfur (S) is an essential element for all organisms and is present in proteins,

2- lipids, carbohydrates, and several metabolites. Sulfate (SO4 ) is the preferred S source

2- for most organisms. In photosynthetic organisms, the reductive assimilation of SO4 occurs in plastids, which means that this ion must traverse both the plasma membrane and plastid envelope prior to reduction and incorporation into organic molecules.

2- SO4 is relatively inert and must be activated by the enzyme ATP sulfurylase before being reduced to sulfide and incorporated into the amino acids cysteine and methionine (Leustek et al., 2000), which can be used for the synthesis of proteins or converted into other metabolites including glutathione and dimethyl sulfide.

2- Much of the SO4 in the soil is not readily available to plants or microbes. The

2- SO4 anion can be adsorbed onto the surface of the soil particles, and a large

2- proportion may be covalently bonded to organic molecules in the form of SO4 esters

2- and sulfonates. When experiencing low SO4 availability, the unicellular, soil- dwelling alga Chlamydomonas exhibits a suite of responses that include the synthesis

2- of extracellular arylsulfatases (ARS), elevated SO4 transport activity, and an increase in the levels of transcripts encoding ATP sulfurylase and other enzymes associated with S assimilation. Many of these responses allow the alga to more efficiently

2- scavenge and assimilate available SO4 in the environment (Davies and Grossman,

1998; Grossman and Takahashi, 2001; Zhang et al., 2004; Gonzalez-Ballester et al.,

2008; Shibagaki and Grossman, 2008; Gonzalez-Ballester et al., 2010).

Specific polypeptides involved in regulating Chlamydomonas S deprivation responses have been identified. The SAC1 (sulfur acclimation) protein is required for

72 many S-limitation-induced responses. Chlamydomonas mutants with lesions in the

2- SAC1 gene exhibit abnormal SO4 uptake, are unable to synthesize extracellular ARS and show little increase in many S-deprivation-responsive transcripts, including those encoding ARS, ATP sulfurylase, serine acetyltransferase, and the ferredoxin- dependent sulfite reductase. Furthermore, sac1 mutants cannot suppress photosynthetic electron transport activity and rapidly die when placed in S-deficient medium in the light. Even though the SAC1 gene encodes a protein similar to anion

+ 2- transporters from a number of different organisms, including the Na /SO4 transporter from mammals, the phenotypes of sac1 mutants strongly suggest that SAC1 functions in regulating cellular responses to S deprivation (Davies et al., 1996). A second polypeptide that plays a central role in the acclimation of Chlamydomonas to S deprivation is SNRK2.1, a member of the Snf1-related protein kinase 2 family. Like

SAC1, SNRK2.1 is required for most responses associated with the acclimation of

Chlamydomonas to S deprivation. An snrk2.1 mutant (initially designated ars11) is defective for expression of several S-responsive genes and the mutant bleaches and dies more rapidly than wild-type cells during S starvation; the responses of snrk2.1 to

S deprivation are generally more severe than those of the sac1 mutant (Gonzalez-

Ballester et al., 2008).

2- Increased SO4 uptake in response to S limitation has been extensively documented for prokaryotic and eukaryotic organisms, including Escherichia coli,

Neurospora crassa, Saccharomyces cerevisiae and Arabidopsis thaliana (Breton and

Surdin-Kerjan, 1977; Ketter and Marzluf, 1988; Shibagaki et al., 2002; Gyaneshwar et

2- al., 2005). In prokaryotic organisms, SO4 is often transported into the cell by a single

73 2- transport system, whereas eukaryotes often have multiple SO4 transporters. In S.

2- cerevisiae and A. thaliana, there are both high and low affinity SO4 transporters

(Breton and Surdin-Kerjan, 1977; Leustek et al., 2000). Increased accumulation of

2- transcripts encoding SO4 transporters upon S starvation has also been noted for S. cerevisiae, N. crassa, and A. thaliana (Ketter and Marzluf, 1988; Cherest et al., 1997;

Yoshimoto et al., 2002).

2- A. thaliana has 12 putative SO4 transporters that have been classified into four groups. The AtSULTR1 group represents high affinity transporters responsible

2- for the initial uptake of SO4 into root cells, while the AtSULTR2;1, AtSULTR2;2 and AtSULTR3;5 are low affinity transporters likely to play important roles in the

2- translocation of SO4 from roots to leaves (Takahashi et al., 2000; Hawkesford, 2003;

Kataoka et al., 2004a). Furthermore, two isoforms of the AtSULTR4 group have been

2- localized to the tonoplast and facilitate SO4 efflux from vacuoles (Kataoka et al.,

+ 2- 2004b). The plant-associated H /SO4 co-transporters usually contain twelve transmembrane domains followed by a linking region that connects to a carboxy-

2- terminal STAS (SO4 transporter and anti-sigma antagonist) domain. There is significant interest in the function of the STAS domain since lesions in this domain can lead to serious human diseases (Everett and Green, 1999). While the precise function of the STAS domain is not clear, it is required for the activity and proper

2- assembly of the SO4 transporters (Shibagaki and Grossman, 2004; Rouached et al.,

2005; Shibagaki and Grossman, 2006) and may help regulate interactions of the transporter with partner proteins that function downstream in the S assimilation pathway (Shibagaki and Grossman, 2010).

74 2- Features of SO4 transport in Chlamydomonas during S-replete and S-depleted conditions have been reported, although there has been little molecular

2- characterization of the genes and polypeptides encoding the putative SO4 transporters. Initial studies demonstrated that both the maximum velocity (Vmax) and

2- substrate concentration at which SO4 transport is at half maximum velocity (K1/2) were altered in S-deprived cells. The Vmax increased approximately 10-fold and the

2- K1/2 decreased roughly 7-fold. The enhancement of SO4 transport upon S deprivation is prevented by cycloheximide, but not by chloramphenicol, demonstrating that protein synthesis on 80S cytoplasmic ribosomes is required for the synthesis of the high-affinity transport systems (Yildiz et al., 1994).

Analysis of the Chlamydomonas genome sequence has led to identification of

2- several genes encoding proteins with high sequence similarity to known SO4

2- + 2- transporters. Three of the putative SO4 transporters (SULTR1-3) are in the H /SO4 family (characteristic of vascular plants) while another three (SLT1-3; SAC1-like

+ 2- transporter) are in the Na /SO4 transporter family that is present in bacteria, non- vascular plants and mammals; these transporters have not been identified in vascular plants. Another SULTR-type transporter, originally designated SULTR4, was recently shown to encode a functional molybdate transporter and the gene has been renamed

MOT1 (Tejada-Jiménez et al., 2007). Transcripts from some of the transporter genes increase significantly during S deprivation; increased accumulation of SLT1 and

SULTR2 transcripts during S starvation was previously reported (Gonzalez-Ballester et

2- al., 2008). Chlamydomonas also possesses SO4 permeases (SulP1 and SulP2) that resemble the permeases of bacteria (Laudenbach and Grossman, 1991; Chen et al.,

75 2003; Lindberg and Melis, 2008). These transport proteins have been localized to the

2- chloroplast envelope and function in the transport of SO4 from the cytosol into chloroplasts where reductive assimilation of the anion occurs.

Here we identified, localized and examined the regulation of Chlamydomonas

2- SO4 transporters. We defined the kinetics of accumulation of both the RNA encoding the transporters and the transporter polypeptides following the imposition of S

2- deprivation, and monitor their decay after SO4 is added back to starved cells. We also evaluated the impact of the regulatory elements SAC1 and SNRK2.1 on the accumulation of SULTR and SLT transcripts in S-replete and S-depleted cells and show that the proteasome is involved in the turnover of SLT but not SULTR transporters. Finally, various methods were used to localize the different transporters to specific cellular membranes and to identify and characterize mutants that were specifically defective for the function of the individual transporter polypeptides. These analyses provide a comprehensive view of the function, biogenesis and regulation of

2- the S-responsive SO4 transporters in Chlamydomonas.

RESULTS

2- Chlamydomonas SO4 transporter genes and proteins: Previous work identified

2- full-length cDNA clones encoding the A. thaliana and Stylosanthes hamata SO4

2- transporters. These genes were characterized and used to identify potential SO4 transporters from other species (Takahashi et al., 1997). Similarly, analysis of the entire Chlamydomonas genome sequence (Merchant et al., 2007) allowed us to

2- identify genes encoding putative SO4 transporters in this alga. Six candidate genes

76 (SULTR1-3 and SLT1-3) were identified based on homologies to plant, animal and

2- bacterial SO4 transporters. The deduced amino acid sequences of SULTR1 and

+ 2- SULTR2 are highly similar (60%) to the H /SO4 cotransporters (SLC26 family) from vascular plants, including those of A. thaliana and S. hamata (Figure 3-1A), although both of the Chlamydomonas transporter proteins have an insertion of 17 amino acids starting at amino acid 199 of SULTR1. The deduced amino acid sequence of SULTR3

2- is more similar to SO4 transporters from bacteria. SULTR1 and SULTR2 of

Chlamydomonas are most similar to A. thaliana AtSULTR1;2 and AtSULTR4

(Figure 3-1A). The carboxy-terminal STAS (sulfate transporter and antisigma factor antagonist) domain of SULTR transporters (Figure 3-1A, marked by red bar) is present on various anion transporters and extends into the cytoplasm of the cell.

In contrast to the SULTR-type transporters, the deduced amino acid sequences of SLT1, SLT2, and SLT3 of Chlamydomonas exhibit strong sequence similarity to

+ 2- Na /SO4 transporters (SLC13 family). All three members of the SLT family in

Chlamydomonas have 10-12 predicted transmembrane domains and an intracellular loop containing a TrkA-C domain (Figure 3-1B, marked by orange bar). While the precise function of the TrkA-C domain is not known, it may be involved in the

2- regulation of SO4 transporter activity, potentially through interactions with partner proteins. The similarity among the Chlamydomonas SLTs and between the SLTs and

2- putative SO4 transporter of the moss Physcomitrella patens is shown in Figure 3-1B.

2- Induction of SO4 transport and accumulation of SULTR2, SLT1 and SLT2 transcripts during S deprivation: When organisms are deprived of S, they often

77 2- exhibit an increase in their affinity for SO4 (decrease in the K1/2) and in the maximal

2- rate of SO4 transport (elevated Vmax). As shown in Figure 3-2, the maximal rate of

2- SO4 transport into wild-type Chlamydomonas cells (strain 21gr) increases approximately 13 fold after 24 h of S deprivation. To identify those transporters likely

2- to be responsible for the change in kinetics of SO4 transport of cells acclimated to S deprivation, we monitored changes in the abundance of transcripts encoding the

2- putative SO4 transporters of Chlamydomonas by RT-qPCR. RNAs from wild-type cells placed in medium devoid of S, nitrogen (N) or phosphorus (P) for 12 h. The elevated levels (note the log scale) for transcripts from ARS2 (encoding an extracellular ARS) during S deprivation, from PHOX (encoding an alkaline phosphatase) during P deprivation, and from NIT1 (encoding a nitrate reductase) during N deprivation (Figure 3-3), suggest that the cells were responding normally to

2- each of the nutrient starvations. Transcripts encoding the SO4 transporters showed remarkable changes in abundance when the cells were starved for S, but not when starved for P or N (Figure 3-3). SULTR2 and SLT2 transcript levels increased by >100 fold, while the level of the SLT1 transcript exhibited an increase of ~1,000 fold after

12 h of S deprivation. In contrast, SULTR1 and SLT3 transcript levels decreased roughly 1,000- and 10-fold, respectively, specifically during S deprivation, while the abundance of the SULTR3 transcript exhibited little change (Figure 3-3). The accumulation of SULTR2, SLT1, and SLT2 transcripts following the imposition of S deprivation suggests that these three genes may encode the dominant, potentially high

2- affinity SO4 transporters, while the decrease in SULTR1 and SLT3 transcript levels

78 2- suggest that they may encode low-affinity SO4 transporters (not needed in an S- limited environment).

2- The induction of SO4 transport activity observed in wild-type

Chlamydomonas cells during S limitation is reduced in the sac1 mutant and completely abolished in the snrk2.1 mutant (Figure 3-2). To understand the relationship between transporter activity and the levels of transcripts encoding the

2- different transporters, we quantified SO4 transporter transcripts in wild-type and mutant strains. While the level of the ARS2 transcript increased approximately 3 orders of magnitude in wild-type cells (21gr) within 2 h of S deprivation (Figure 3-

4A), the sac1 or the snrk2.1 mutants showed a marked reduction in the increase in

ARS2 mRNA abundance (Figure 3-4B and 3-4C, respectively). Furthermore, the elevation (SULTR2, SLT1, SLT2) and depression (SULTR1, SLT3) of levels of

2- transcripts encoding the various putative SO4 transporters following S-deprivation was generally not nearly as marked in the sac1, and especially in the snrk2.1 mutants relative to wild-type cells. The levels of accumulation of the SULTR2, SLT1 and

SLT2 polypeptides were consistent with their transcript abundances during S starvation (Figure 3-5).

Interestingly, some transcripts encoding transporters were still elevated in the mutants following S deprivation; the most notable examples are the SLT1 transcript in the sac1 mutant, and the SULTR2 transcript in the snrk2.1 mutant (Figure 3-4B and 3-

4C); the SULTR2 transcript is also elevated to some extent under S-replete conditions in the sac1 mutant. Furthermore, while the SULTR2 transcript still increases significantly in snrk2.1, the absolute level of the transcript is much lower in the mutant

79 than in the wild-type strain (2-3 orders of magnitude) under both S-replete and S- deprivation conditions. Also, the increase in the SLT1 transcript in the sac1 mutant is only observed after 24 h of S deprivation (it is observed 2 h after wild-type cells are exposed to S deprivation). The SLT1 transcript in the snrk2.1 strain is essentially not detected under S-replete or S-deprivation conditions. Finally, while the level of the

SULTR1 transcript markedly declines in wild-type cells, its decline is diminished in the sac1 mutant, and it actually increases by 5-10 fold in the snrk2.1 strain. The

SULTR3 transcript, which also decreases to some extent in the wild-type strain during

S deprivation, may increase slightly in the snrk2.1 mutant (compare Figure 3-4A and

3-4C).

These results suggest that there are some complex relationships linking the mutant phenotypes to transcript levels, although regulation in the mutant strains is aberrant for essentially all transporter transcripts that were examined; we can conclude that the SAC1 and SNRK2.1 regulators are required for both increases and decreases in transcript abundances that are observed following the transfer of Chlamydomonas cells to S-deprivation conditions. Furthermore, SNRK2.1 appears to be important for maintaining moderate levels of SLT1 and SULTR2 transcripts when the cells are growing in nutrient-replete medium. Finally, some of the transcripts encoding transporters that normally sharply decline in wild-type cells during S deprivation can become elevated in the mutant strains (SULTR1 in the snrk2.1 mutant). These findings suggest that there may be both direct and indirect consequences of elimination of the

SAC1 and/or SNRK2.1 regulatory proteins during S deprivation on the level of transporter transcripts; the increase in the SULTR3 transcript in snrk2.1 might reflect

80 the inability of the strain to suppress expression of the SULTR3 gene, which might be the result of a compensatory effect elicited in cells severely compromised in their ability to scavenge S from external and probably internal sources.

SULTR2, SLT1, and SLT2 protein levels: We analyzed wild-type cells for levels of

SULTR2, SLT1 and SLT2 transcripts and polypeptides, both before and during S

2- deprivation and following the addition of SO4 back to starved cells. Figure 3-6A shows the levels of these transcripts in cells collected at various times from 1-32 h

2- after removal of S from the medium, and at 1, 2, 4 and 8 h after the addition of SO4 to cells that had experienced 24 h of S starvation. The same samples that were used to quantify transcript abundance by RT-qPCR were also used for microsomal membrane preparations to monitor the levels of the transporter proteins by Western blot analyses, as shown in Figure 3-6B.

As presented in Figure 3-6A, levels of SULTR2, SLT1 and SLT2 transcripts are high after 1 h of S deprivation and remain at approximately the same level after 24 h of S deprivation. However, these mRNAs declined at different rates following the

2- addition of SO4 to the medium. The SLT1 transcript declined most rapidly, although none of the transcripts dropped to the +S levels, even after 8 h following the re-

2- addition of SO4 to the medium.

For monitoring transporter protein abundances, polyclonal antibodies were raised against a peptide unique to SLT2 (Figure 3-1B, marked by magenta bar), a peptide unique for SULTR2 (Figure 3-1A, marked by blue bar) and a peptide present in both SLT1 and SLT2 (SLT ‘general’ antibodies; Figure 3-1B, marked by green

81 bar) (see MATERIALS AND METHODS). The SLT general antibodies (SLT in

Figure 3-6B) detected a specific band of approximately 95 kDa, which corresponds to the predicted molecular mass of SLT1 and SLT2 (93 kDa and 96 kDa, respectively).

The specific SULTR2 antibodies recognized a polypeptide of molecular mass of ~80 kDa, which corresponds to the predicted molecular mass of SULTR2 (~84 kDa).

Generally, all of the transporter polypeptides appear to migrate as doublets (the most well resolved doublet is for SULTR2), which could be a consequence of protein modification, proteolytic cleavage of a short amino or carboxyl terminal region, and/or the recognition of more than one polypeptide by the antibodies (although the two proteins would have to show the same kinetic changes with respect to the S status of the medium). The marked, rapid accumulation of the transcript for the SLT and

SULTR2 transporters (after 1 h of S deprivation, Figure 3-6A) is followed by a dramatic increase in the levels of the transporter polypeptides, which is first detected 2

2- h after the initiation of S deprivation (Figure 3-6B). Following administration of SO4 to S-starved cultures, the level of SULTR2, SLT2 and probably SLT1 (detected by the general SLT antibody and confirmed by analysis of transporter polypeptides in strains

2- defected for specific SO4 transporters, shown in Figure 3-14) polypeptides declined

2- and became barely detectable 8 h after the SO4 content of the medium had been replenished (Figure 3-6B). We also observed some decline in polypeptide levels at 32 h relative to 24 h of S deprivation, although the decline was much less severe than in cultures administered S.

82 Turnover of SULTR2, SLT1, and SLT2 polypeptides: We analyzed whether the

2- 2- decrease in abundance of the SO4 transporters following the addition of SO4 was a consequence of enhanced degradation. Cells were placed in medium devoid of S for

2- 10 h to allow accumulation of SO4 transporter polypeptides prior to the administration of cycloheximide (CHX), an inhibitor of protein synthesis on 80S

2- ribosomes, with or without the addition of 1 mM SO4 . The concentration of CHX used in these experiments was previously shown to specifically block cytoplasmic protein synthesis (Kawazoe et al., 2000); it also completely blocked accumulation of

2- the SO4 transporter proteins in cells transferred to medium lacking S (Figure 3-7).

2- The levels of the SO4 transporters were monitored 20, 60 and 240 min following the

2- 2- addition of the inhibitor and/or SO4 to the cultures. As expected, the SO4 transporters accumulated after 10 h of S starvation (Figure 3-8A, lane 2).

Interestingly, these polypeptides were turned over significantly more rapidly in

2- cultures that received SO4 along with CHX (Figure 3-8A; lanes 10-12) compared to cultures that were only administered CHX but remained S-starved (Figure 3-8A; lanes

4-6). The band intensity was quantified, normalized to the level of the FOX1 protein and the half-life of each polypeptide was derived by fitting the values to the decaying

2- exponential equation. These results indicate that administration of SO4 enhances degradation of the SULTR2, SLT1, and SLT2 polypeptides, although the kinetics of the turnover varies among the transporters. Since the SLT general antibodies recognize both SLT1 and SLT2 polypeptides, the half-life estimated from the signals detected with these antibodies reflects the turnover rate of both of the SLT proteins.

2- Interestingly, when SO4 was added back to the cultures, the addition of CHX made

83 little difference to the kinetics of the loss of SLT and SULTR2 polypeptides (compare

2- Figure 3-8A, lanes 7-9 with 10-12), raising the possibility that the addition of SO4 to starved cultures both rapidly blocked the synthesis of these protein (which in part also declines because of transcript degradation) as well as stimulating their degradation.

2- To determine whether the proteasome was involved in degradation of the SO4 transporters, we performed a similar experiment in which cells were S-starved for 10 h

2- prior to the addition of CHX and SO4 to the cultures, in the presence and absence of the proteasome inhibitor MG132; samples were collected 30, 90 and 240 min

2- following the addition of the inhibitors plus SO4 . Figure 3-8B shows that degradation of SLT1 and SLT2 polypeptides was completely blocked by MG132, suggesting that proteasome activity is required for the turnover of these proteins.

Intriguingly, SULTR2 turnover was not affected when proteasome function was inhibited (repeated on three separate samples with similar results), suggesting that there are different mechanisms for eliminating the transporter proteins once S becomes sufficient in the environment.

2- Transporter localization: To localize the SO4 transporters, total membranes were prepared from S-starved cells and separated by a two-phase aqueous polymer system into a plasma membrane (polyethylene glycol) fraction and a fraction containing the rest of the cellular membranes. The polypeptides from the plasma membrane and

‘other membrane’ (non-plasma membrane) fractions were resolved by SDS-PAGE and immunoblot analyses were performed using antibodies to proteins of known localization. As shown in Figure 3-9A, the plasma membrane fraction was largely

84 free of contamination from thylakoid membrane and chloroplast envelope, based on the distribution of CRD1 (Allen et al., 2008), and from mitochondrial membrane, based on the distribution of COX2b (Page et al., 2009). As expected, two plasma membrane markers, H+ ATPase (Norling et al., 1996) and FOX1 ferroxidase (Herbik et al., 2002), partitioned preferentially to the phase with the plasma membrane

(polyethylene glycol). SLT2 and SULTR2 were detected almost exclusively in the plasma membrane fraction, however, a weak signal with the general SLT antibodies was also observed in the non-plasma membrane fraction. This latter partitioning pattern is similar to that of a tonoplast V ATPase. In order to clearly establish whether

2- or not these SO4 transporters are localized to the plasma membrane, proteins from the polyethylene glycol phase were separated on a 15-45% sucrose gradient (Figure 3-

9B). While the tonoplast V ATPase was mainly in fractions 7-9, SLT1, SLT2, and

SULTR2 migrated to the bottom of the gradient and were clearly localized with the plasma membrane marker proteins FOX1 ferroxidase and the H+ ATPase (fractions 1-

3).

Expression in heterologous sytems: To demonstrate that SULTR2, SLT1, and SLT2

2- encode functional SO4 transporters, their cDNA clones were used to functionally complement the S. cerevisiae (yeast) uracil auxotrophy strain CP60-1C, which harbors

2- mutations in SUL1 and SUL2 SO4 transporter genes. CP60-1C grows very slowly on

2- medium containing 100 µM SO4 (or less) as a sole S source (Cherest et al., 1997).

2- cDNAs encoding the full-length Chlamydomonas SO4 transporters were cloned into the yeast expression vectors pDR196-GW or pDR196-GW-GFP, both of which

85 contain the promoter region from PMA1 (encoding a plasma membrane H+ ATPase), which drives constitutive expression of the inserted genes. The stop codons from the

2- SO4 transporter cDNAs were removed when cloned into pDR196-GW-GFP to create a fusion protein with the C-terminus of GFP. Table 3-1 shows that expression of

Chlamydomonas SULTR2, SLT1, and SLT2 did not rescue the slow-growth phenotype

2- of the CP60-1C strain in liquid medium containing 100 µM SO4 as a sole S source; the same growth rate was observed for yeast cells carrying the vector alone and those

2- carrying plasmids with Chlamydomonas SO4 transporter genes. Both the GFP-tagged and untagged forms of transporters yielded the same results (failure to complement the yeast mutant). In contrast, CP60-1C cells transformed with Arabidopsis SULTR1;2 cDNA exhibited a significantly shorter doubling time. AtSULTR1;2 has previously

2- been shown to rescue the yeast SO4 transporter mutant (Shibagaki et al., 2002).

While it was possible that the difference in Chlamydomonas and yeast codon usage could result in heterologous expression problems, this is probably not the case

2- since, as shown in Figure 3-10A, the Chlamydomonas SO4 transporter-GFP fusion proteins accumulated in the yeast transformants. We speculated that the recombinant transporters are retained in the endoplasmic reticulum (ER), Golgi apparatus, or secretory vesicles and not enough was localized to the plasma membrane to rescue the

CP60-1C phenotype. Indeed, in yeast cells expressing Chlamydomonas SULTR2-,

SLT1-, and SLT2-GFP fusion proteins, the fluorescence signal appeared to accumulate within intracellular membranes, probably a proliferation of the ER (Figure

3-10B, panels I, II, and III). In contrast, in CP60-1C cells harboring AtSULTR1;2 fused to GFP, much of the fluorescence signal is distributed over the cell surface

86 (Figure 3-10B, panel IV). These results suggest that our inability to complement the yeast mutant is a consequence of inefficient localization of the Chlamydomonas transporters to the yeast plasma membrane.

Generation of an insertional mutant library: Our inability to confirm the

2- functionality of the putative SO4 transporters using the heterologous system prompted us to screen for Chlamydomonas strains bearing mutations in those transporter genes. Insertional mutagenesis has been widely used in Chlamydomonas to generate mutant strains affecting various biological processes. To obtain mutants in

2- putative SO4 transporter genes, we generated a library containing approximately

52,000 insertional mutants using D66 as the parental strain. Cells were transformed with a 1.7 kb PCR fragment containing the AphVIII gene, which confers resistance to the antibiotic paromomycin, and transformants were plated on selective medium. The steps in the construction of the mutant library are described in the MATERIALS

AND METHODS. Genomic DNA isolated from transformants was screened by PCR

2- to identify specific gene disruptions. Mutants with an insertion in the SO4 transporter genes were identified by PCR using a target gene-specific primer and a primer specific to the inserted DNA (Table 3-2 shows the list of primer pairs that were used). PCR products were sequenced to confirm that the disruption was in the targeted transporter gene. The specific transformant harboring the insertion was then identified, backcrossed (with strain 21gr) to generate a homogeneous genetic background and then more fully characterized.

87 Figure 3-11A shows the positions of the AphVIII gene in the strains disrupted for SLT1, SLT2, SULTR2. The Southern blot analyses illustrated that slt1, slt2, and sultr2 mutants carry a single AphVIII insertion (Figure 3-11B). These results were also confirmed by a genetic linkage analysis (data not shown). In a slt1 mutant, the

AphVIII construct was inserted in the eighth exon, interrupting the translation of a full- length product by introducing an incorrect splice site that caused a frame shift and the appearance of a premature stop codon (Figure 3-12A shows the alignment of the wild- type and truncated SLT1 gene products). The aberrant SLT1 transcript in the slt1 mutant does not accumulate to the same extent as does the normal transcript in wild- type cells starved for S (the accumulation in the mutant is approximately one order of magnitude less than in the parental strain; Figure 3-13A), suggesting that the mutation may also cause transcript instability. The lesion also abolished SLT1 protein accumulation during S deprivation, although a signal is observed when the general

SLT antibody is used (Figure 3-14A). Since the general SLT antibody recognizes both

SLT1 and SLT2 polypeptides, the signal observed in the lane with polypeptides from the slt1 mutant is likely to result from the presence of SLT2 as there is essentially no detectable SLT signal in a slt1slt2 double mutant (Figure 3-14B).

The integration site of the AphVIII gene in the slt2 mutant is in the 5’ untranslated region (Figure 3-11), 38 bp upstream of the translation start codon. Even though this disruption is not in the coding sequence, the accumulation of both the

SLT2 transcript and polypeptide are severely impacted by the mutation (Figure 3-13A and 3-13A); essentially no SLT2 protein is detected in the mutant. We also observed a novel phenotype of the slt2 mutant: the SLT3 transcript, which decreased during S-

88 limited growth of wild-type cells and the slt1 and sultr2 mutants, consistently increased in the slt2 mutant (Figure 3-13A).

In the sultr2 mutant, a fragment of the AphVIII construct was inserted in the ninth exon of SULTR2, resulting in the generation of a chimeric transcript with a premature stop codon. The SULTR2-AphVIII transcript would be translated into a truncated polypeptide missing the last 115 amino acids of SULTR2 (Figure 3-12B).

This truncated polypeptide appears to be unstable and does not accumulate when the sultr2 mutant is starved for S (Figure 3-14A).

2- SO4 uptake by the mutant strains: Prior to performing phenotypic analyses, each of the single mutants was backcrossed several times to the wild-type 21gr strain to eliminate other background mutations. The single mutants were then crossed to each other to generate double and triple mutants. The transcript and protein analyses for the strains with the multiple mutations are shown in Figure 3-13 and Figure 3-14. There did not appear to be significant compensation for the absence of two S deprivation- induced transporters in the double mutants; the level of transcript and polypeptide accumulation for the third transporter was similar to that observed in wild-type cells

(Figure 3-13B and Figure 3-14B), based on normalization to the level of the CBLP transcript and COX2b protein, respectively. In order to assess whether the absence of

2- 2- specific SO4 transporters impacted the rate of SO4 transport, we measured the

2- 2- uptake rate of SO4 over a range of external SO4 concentrations (0.02 – 200 µM) in the wild-type and the various mutant strains following S deprivation. As shown in

2- Figure 3-15A and 3-15B, the rates of SO4 uptake in the slt1, slt2, and sultr2 single

89 mutants were not markedly depressed relative to that of wild-type cells. In contrast, the three double mutants all showed a significant decline (by ~50%) in their rate of

2- SO4 transport relative to wild-type cells. Most extreme was the phenotype of the triple slt1slt2sultr2 mutant. This strain, which lacks all of the inducible transporters,

2- exhibited a drastic decrease in the rate of SO4 uptake. Indeed, the triple mutant

2- exhibited essentially no increase in its SO4 uptake capacity when deprived of S; the

2- rate of SO4 transport was the same in starved and unstarved cells (Figure 3-15A and

3-15B). Furthermore, wild-type cells and the slt1slt2sultr2 triple mutant showed a

2- similar rate of SO4 transport under nutrient-replete conditions. The K1/2 was also

2- calculated from the initial rates of SO4 uptake for single, double and the triple mutants; we were unable to detect a significant difference in the transport affinity of wild-type cells relative to any of the mutant strains (Table 3-3). This may not be too surprising for the single and double mutants, but the triple mutant also showed a decrease in the K1/2 that closely resembled that of wild-type cells. This might arise from the fact that slt2 mutation only affects transcript accumulation during S deprivation and does not disrupt the coding region; a small amount of SLT2 protein can still be translated from the basal level of SLT2 transcript. In fact, a very weak signal using the SLT antibodies was detected in the slt1slt2sultr2 mutant in S-depleted medium, which is likely from a low level of SLT2 polypeptide (Figure 3-14B).

Alternatively, post-translational modifications of the transporters that function during

S-replete growth may alter their kinetic characteristics, although they remain at low levels in the plasma membrane (so the Vmax for uptake remains very low). In order to confirm that the phenotype observed in the double and triple mutants is caused by the

90 2- lesions in the SO4 transporter genes, wild-type alleles of each transporter gene were introduced into the slt1slt2sultr2 mutant background by a genetic cross to wild-type

2- strain (D66). The SO4 uptake rates of the progeny correlated perfectly with the

2- number of SO4 transporter genes present in each strain (Figure 3-16A and 3-16B).

Note that the relative uptake rates of the single and double mutants are similar to the results presented in Figure 3-15, even though the absolute transport rates are lower in this experiment (due to the background difference between 21gr and D66 strains). In sum, these results demonstrate that SLT1, SLT2, and SULTR2 are the prominent,

2- high-affinity transporters involved in SO4 uptake when the cells are deprived of S, while other transporters function during S-replete growth. Furthermore, there appears to be some functional redundancy among the activities of these transporters.

DISCUSSION

2- SO4 transporters in Chlamydomonas: One of the early responses of most

2- organisms to S deprivation is the generation of high-affinity SO4 transport systems.

2- In this study, we identified genes encoding SO4 transporters and demonstrated that the activation of these genes occurred specifically during S deprivation. The elevated transcript accumulation was detected 1-2 h after transferring cells to medium devoid

2- of S (Figure 3-3). It is noteworthy that the transcripts encoding the high-affinity SO4 transporters accumulate significantly more rapidly than transcripts encoding other S- responsive genes, such as ARS2 and ECP76, following the transfer of cells to –S medium (Takahashi et al., 2001). These results suggest that there is a tiered regulation

2- of the S-responsive genes and that the induction of the high-affinity SO4 transport

91 system is one of the earliest responses to S starvation. In addition, S-starved cells

2- exhibit increased SO4 uptake activity within 1 h of the onset of S deprivation (Yildiz et al., 1994). This initial increase in uptake rate precedes accumulation of newly

2- synthesized SO4 transporters (Figure 3-6B), suggesting that there might also be post-

2- translational modifications of the existing SO4 transporters. This suggestion is in

2- accord with the finding that the K1/2 for SO4 transport in the slt1slt2sultr2 triple mutant still rapidly changes following the imposition of S deprivation; the Vmax remains low because the three S-responsive transporters are not synthesized. Finally,

2- transcripts for two putative SO4 transporters, SULTR1 and SLT3, decline during S deprivation. The encoded proteins may have low-affinity transporter activity that is

2- likely to primarily function in the uptake of SO4 under S-replete conditions.

+ 2- Intriguingly, Chlamydomonas has both H /SO4 -related cotransporters

+ 2- (SULTR) and Na /SO4 -related transporters (SLT) while vascular plants such as A.

+ 2- thaliana have only retained the H /SO4 cotransporters. This finding suggests that

Chlamydomonas diverged from the plant lineage (~1 billion years ago) prior to the

+ 2- 2- loss of the Na /SO4 -related transporters. Furthermore, some SO4 transporters in

Chlamydomonas appear to have arisen from a recent duplication since the closely related organisms such as Volvox carteri and the marine chlorophyte Ostreococcus tauri appear to have fewer transporters. For example, Volvox appears to have one

+ 2- + 2- H /SO4 -related and two Na /SO4 -related cotransporters and Ostreococcus has only

+ 2- two Na /SO4 -related transporters; Chlamydomonas has three members for each of

+ 2- those protein families. Retention of the Na /SO4 -related transporters in Ostreococcus may reflect the fact that it would not be energetically favorable to use H+ coupled

92 2- + SO4 transport in the oceans where Na concentrations are high. In addition, Volvox

2- may not require a large set of SO4 transporter proteins since it has an extensive extracellular matrix that could potentially store significant amounts of S. The inability of Chlamydomonas to store S and perhaps other nutrients may have favored the selection of strains in which there was an expansion of families of genes encoding

+ 2- nutrient transport proteins. Furthermore, retaining both the Na /SO4 -related and

+ 2- H /SO4 -related transporter types may allow Chlamydomonas to survive under

+ 2- diverse environmental conditions. The H /SO4 -related transporters may be primarily

+ 2- used when the pH of the milieu around the cell is low while the Na /SO4 -related

2- cotransporters may be responsible for the majority of SO4 uptake when the pH is high and when it is more efficient to use Na+ as a counter ion.

The SLT2 and SLT3 genes are tandemly arranged on chromosome 10 in a head- to-tail orientation; the 3’ untranslated region of SLT2 overlaps with the 5’ untranslated region and the first exon of SLT3. In wild-type cells, SLT2 is heavily transcribed during S deprivation and this likely interferes with the transcription of the downstream

SLT3 gene, resulting in a decrease in SLT3 mRNA abundance (Figure 3-13). In contrast, transcriptional activity of SLT2 in the slt2 mutant is decreased due to the insertion in the 5’ untranslated region, which probably accounts for an unusually high rate of SLT3 transcription during S deprivation. Interestingly, SLT2 and SLT3 (76% identity, 85% similarity to each other) have a similar exon-intron organization, suggesting that they arose from a gene duplication but, based on their unique expression patterns, the genes have subsequently specialized (Figure 3-4A). Finally, the SLT and SULTR protein families each likely contain both high- and low-affinity

93 transporters, allowing Chlamydomonas to tailor uptake processes to the features of the environment under both high and low S conditions.

2- 2- Transcriptional regulation of SO4 transporters: Several isoforms of SO4

2- transporters play important roles in facilitating SO4 uptake during S deprivation in A. thaliana. The -S-responsive induction of these transporter genes (AtSULTR1;1,

AtSULTR1;2, AtSULTR4;2) has been shown to be under the control of SLIM1, the

ETHYLENE-INSENSITIVE-LIKE3 (EIL3) transcriptional regulator (Maruyama-

Nakashita et al., 2006). As already mentioned, a number of factors associated with S- responsive regulation in Chlamydomonas have also been identified; these include

SNRK2.1, SNRK2.2 and SAC1. Mutants devoid of these regulatory elements show aberrant responses to S deprivation (Gonzalez-Ballester and Grossman, 2009). We

2- examined accumulation of SO4 transporter transcripts in wild-type cells and the sac1

2- and snrk2.1 mutants; these mutant strains do not elevate SO4 transport activity nearly as much as wild-type cells when they are deprived of S (Figure 3-2). In contrast to wild-type cells, the sac1 mutant only accumulates SLT1, SLT2, and SULTR2 transcripts 24 h after the imposition of S starvation. This is consistent with the model proposed in Moseley et al. (2009) in which SAC1 is a sensor that resides on the plasma membrane and acts as a negative regulator of the SNRK2.2 repressor protein

2- kinase when environmental SO4 levels fall below a certain threshold. This causes de-

2- repression of the S-responsive genes, including those encoding SO4 transporters.

2- After a prolonged period of S starvation, intracellular SO4 levels also decline and further stimulate transcription of S-responsive genes as a consequence of activation of

94 the SNRK2.1 activator protein kinase; such activation can occur in the sac1 mutant,

2- leading to elevated accumulation of SO4 transporter mRNA 24 h after the onset of S starvation (Figure 3-4B), which then leads to an increase in the level of transporter protein (Figure 3-5) and uptake activity (Figure 3-2), although both are still lower than in wild-type cells.

Similarly, the snrk2.1 mutant is severely compromised in its ability to increase

2- the rate of SO4 uptake during S starvation (Figure 3-2); its phenotype is significantly more severe than that of the sac1 mutant. The abundance of SLT1, SLT2, and SULTR2 transcripts in the snrk2.1 mutant, under both S-replete and S-depleted conditions, is much lower than the levels observed in wild-type cells (Figure 3-4). For SULTR2, expression is induced in the snrk2.1 mutant during S deprivation, but the basal and fully induced transcript levels are both much lower than observed in wild-type cells.

The SLT1 transcript does not accumulate in this mutant. These results suggest that

SNRK2.1 is critical for maintaining both basal level expression and accumulation of

SLT1 and SULTR2 during acclimation of the cells to S deprivation. Similar to the

2- situation in sac1 mutant, the inability of snrk2.1 to induce SO4 transport activity

2- during S-deprivation is associated with its failure to accumulate SO4 transporter

2- transcripts. The capacity for SO4 transport in wild-type, sac1 and snrk2.1 mutant cells starved for S generally correlates with the levels of transcript and transporter polypeptide accumulation (Figure 3-2, Figure 3-5, and Figure 3-6A and 3-6B).

Together, these results indicate that SLT1, SLT2, and SULTR2 encode high-affinity

2- SO4 transporters that are controlled by the S status of the environment at the level of

95 2- transcript abundance and that are responsible for the majority of SO4 uptake when the cells become S-deprived.

In contrast, the SULTR1 and SLT3 genes appear to be down-regulated during S starvation (the transcripts are reduced by 1000-fold and 10-fold, respectively; Figure

2- 3-4A), suggesting that the proteins encoded by these genes are low-affinity SO4 transporters that function primarily under S-sufficient conditions. Furthermore, S deprivation does not cause a significant reduction in SULTR1 and SLT3 transcript abundance in the sac1 mutant, while these transcripts are elevated to some extent in the snrk2.1 strain (Figure 3-4B and 3-4C). Hence, SAC1 and SNRK2.1 are important not only for stimulating the activity of the SLT1, SLT2, and SULTR2 genes, but are also involved in depressing the activities of the SULTR1 and SLT3 genes during S limitation.

Even though the SULTR3 transcript does not appear to be regulated by the S status of the cells, its level accumulates during S starvation in a snrk2.1 background

(Figure 3-4A and 3-4C). The inability of the snrk2.1 mutant to upregulate expression of the high-affinity transporters and other genes involved in the acclimation program may result in severe internal S starvation that elicits secondary effects, including an increase in SULTR3 transcript accumulation. Alternatively, SNRK2.1 may play a direct role in regulating SULTR3 expression.

2- Accumulation and turnover of SO4 transporters: SLT1, SLT2 and SULTR2 polypeptides accumulate in S-starved cells 1-2 h after the transcripts peak (Figure 3-

6A and 3-6B). These proteins are synthesized de novo upon imposition of S

96 deprivation since administration of CHX at the time of transferring cells from S- replete to S-depleted medium blocked their accumulation (Figure 3-7). Moreover,

2- these SO4 transporters are turned over rapidly when S becomes available; they are

2- almost undetectable 8 h after SO4 is added to S-starved cultures (Figure 3-6B). The

2- half-lives of SLT1, SLT2, and SULTR2 are 2-3 times longer in the absence of SO4 ; the rate of turnover increases when the cells are placed in S-rich medium (Figure 3-8).

These data strongly suggest that de novo synthesis of SLT1, SLT2, and SULTR2

2- specifically facilitates SO4 transport when cells experience S starvation. Once starvation conditions are relieved, these high-affinity transporters are rapidly

2- degraded, with S (either SO4 or a reduced S metabolite) stimulating the degradation process. The proteasome is involved in degrading SLT1 and SLT2, although it is not known whether the involvement is direct or indirect. Surprisingly, proteasome activity does not appear to be required for SULTR2 degradation. There are several examples of degradation of plasma membrane transporter polypeptides by monoubiquitination and subsequent proteasome-independent proteolysis in vacuoles (Eguez et al., 2004;

Wolf, 2004). Tight regulation of high-affinity transporter synthesis and turnover that depends on S availability would allow for an economy of energy utilization by the cells, balancing the transport process with intracellular demand, and might also help

2- control the uptake of selenate, a toxic analog of SO4 , under all environmental

2- conditions. In addition, the high-affinity SO4 transporters may have a lower capacity

2- for SO4 uptake than the low-affinity transporters (SULTR1 and SLT3), making them

2- less suitable for SO4 uptake when there is an abundance of the anion in the environment.

Localization of SLT1, SLT2 and SULTR2 on the plasma membrane and their

2- 2- role in SO4 uptake: To facilitate SO4 uptake from the environment, algal and plant

2- cells must have the capacity to synthesize high affinity SO4 transporters, some of which must reside on the plasma membrane. Using two-phase aqueous polymer separation coupled with sucrose density gradient centrifugation, we demonstrated that all three of the S-deprivation induced transporters, SLT1, SLT2, and SULTR2, reside in the plasma membrane (Figure 3-9). Expression of the cDNAs encoding these

2- transporters in the yeast SO4 transporter mutant CP60-1C failed to rescue its growth phenotype (Table 3-1). The heterologous transporter polypeptides accumulated in the yeast cells, but appeared to be primarily localized to intracellular membrane structure

(Figure 3-10); the proteins never reached the plasma membrane in sufficient quantity to complement the CP60-1C phenotype. Perhaps, plasma membrane localization of

2- Chlamydomonas SO4 transporters requires a post-translational modification (e.g. phosphorylation) that does not occur in yeast cells. Recently, phosphorylation of the yeast nitrate transporter Ynt1 has been shown to be essential for its delivery to the plasma membrane during N limitation (Navarro et al., 2008). Similarly, phosphorylation of the yeast plasma membrane ATPase is critical for its proper maturation and routing to the cell surface (DeWitt et al., 1998). Alternatively,

Chlamydomonas transporters may require an accessory protein in the secretory pathway for proper targeting. The PHOSPHATE TRANSPORTER TRAFFIC

FACILITATOR 1 (PHF1), a plant-specific SEC12-related protein, has been

98 demonstrated to play a critical role in facilitating ER exit of a high-affinity phosphate transporter in Arabidopsis (Gonzalez et al., 2005).

2- We also attempted to express Chlamydomonas SO4 transporters in Xenopus

2- oocytes, but never succeeded in obtaining oocytes with an increased rate of SO4 uptake (data not shown). These results, along with those of the experiments described for heterologous expression of the Chlamydomonas transporters in the yeast system, suggested that there are biogenesis-specific processes that are required for proper maturation/localization of Chlamydomonas transporters. Intriguingly, while none of

2- the high-affinity SO4 transporters from Chlamydomonas was able to complement the

2- yeast CP60-1C strain, SO4 transporters of Arabidopsis (SULTR1;1, SULTR1;2, and

SULTR1;3) do rescue the yeast mutant phenotype (Table 3-1; (Takahashi et al., 2000;

Shibagaki et al., 2002; Kataoka et al., 2004a)). These results suggest that

Chlamydomonas and Arabidopsis, which diverged over a billion years ago (Merchant et al., 2007), have different post-translational modifications, stabilities and/or mechanisms by which the transporters are localized to the plasma membrane.

Since we were unable to confirm the functionality of SULTR2, SLT1, and

SLT2 in the heterologous systems, Chlamydomonas mutants with insertions in the

2- SO4 transporter genes were identified by a PCR-based screen (Figure 3-11) and

2- characterized at the physiological level by measuring SO4 uptake rates in the various strains (Figure 3-15). The results clearly show that the transporters are critical for the

2- uptake and assimilation of SO4 in S-deprived cells and that there is some functional redundancy among the transporters. Intriguingly, the slt1slt2sultr2 triple mutant and wild-type cells exhibited a similar level of sensitivity to selenate when grown on –S

99 medium (data not shown), suggesting that selenate may be taken up by a different

2- transporter or the residual SO4 uptake activity of the triple mutant is sufficient to transport selenate into cells. If selenate is transported into cells through other uptake

2- systems, it would explain why mutants with defective SO4 transport were not identified in the selenate-resistant mutant selection (Chapter 2).

2- In the S-starved sultr2 mutant, the uptake rates at low external SO4 concentrations (0.02 – 2 µM) are comparable to those of wild-type cells while the

2- transport rates at higher SO4 concentrations (20 – 200 µM) are much lower than the rates in the wild-type strain, suggesting that SULTR2 may be responsible for much of

2- the SO4 uptake into S-deprived cells. Remarkably, increased uptake capacity normally observed during S deprivation is completely abolished in the triple mutant

2- (Figure 3-15B); the maximal SO4 uptake rates of the slt1slt2sultr2 mutant during S- replete and S-depleted conditions are essentially the same. These data suggest that

2- SLT1, SLT2, and SULTR2 are responsible for essentially all induced SO4 uptake that is associated with S-deprivation.

100 MATERIALS AND METHODS

Strains and growth conditions: The following Chlamydomonas strains were used:

21gr (available from the Chlamydomonas Center), D66 (nit2-, cw15, mt+) (Pollock et al., 2003), sac1 (Davies et al., 1996) and snrk2.1 (ars11 allele) (Gonzalez-Ballester et al., 2008). Cells were cultured in either S-replete (+S) or S-depleted (-S) Tris-acetate- phosphate (TAP) medium under continuous illumination (80 µmol photon m-2 s-1) on a rotating platform (200 rpm) at 25°C. TAP–S medium was prepared as described previously (Davies et al., 1994). For S starvation experiments, cells were grown to mid-logarithmic phase (2-4 x 106 cells mL-1) in TAP medium, washed once with TAP-

S medium (2500 X g for 5 min) and resuspended in TAP-S to the original cell density.

For transformation experiments, Chlamydomonas cells were grown in TAP medium to a density of 2-4 x 106 cells mL-1. Transformants were selected on TAP plates supplemented with 5 µg mL-1 paromomycin, as previously described (Davies et al.,

1996; Pollock et al., 2005).

Isolation of SLT1, SLT2 and SULTR2 cDNAs: Total RNA was isolated from 12 h S- starved 21gr (wild-type) cells using a standard phenol-chloroform extraction protocol

(Sambrook et al., 1989). Reverse transcription and PCR were performed using the

Sensiscript RT Kit (Qiagen, Valencia, CA) and Pfu Turbo DNA Polymerase

(Stratagene, La Jolla, CA). Amplified PCR products were cloned into pENTR-D-

TOPO (Invitrogen, San Diego, CA) and sequenced.

2- 2- SO4 uptake assays: SO4 transport assays were performed as previously described

2- (Yildiz et al., 1994). The rate of SO4 uptake was measured at indicated external concentrations of the anion over a 2 min time series.

101 RNA isolation and quantification: Total RNA was extracted from frozen cell pellets using the RNeasy Mini Kit (Qiagen, Valencia, CA) and treated with RNase-free

DNase I (Qiagen, Valencia, CA) to remove residual genomic DNA. First-strand cDNA was synthesized from 3-5 μg of total RNA using oligo-(dT)12-18 for priming the

SuperScript III reverse transcriptase reaction, as described in the manual (Invitrogen,

San Diego, CA). Real-time quantitative PCR (RT-qPCR) was performed with a Roche

LightCycler 480. PCR reactions were in a final vol of 20 µL comprised of 10 µL of

LightCycler 480 SYBR Green Master Mix (Roche, Nutley, NJ), 5 µL of a 1:50 cDNA dilution, 400 nM of each primer, and distilled water to make up the remainder of the

20 µL vol. Conditions used for amplification in the thermocycler were: pre-incubation at 95°C for 5 min followed by 50 cycles of denaturation at 95°C for 10 s, annealing at

60°C for 20 s, elongation at 72°C for 20 s, and measurement of fluorescence after

80°C for 5 s (the last step was incorporated into the protocol to avoid background signals resulting from the formation of primer dimers). A melt-curve analysis program

(60°C-99°C, heating rate of 2.2°C s-1 and continuous florescence measurements) was used to evaluate the specificity of the amplification reactions. All reactions were performed in triplicate with at least two biological replicates. The CBLP gene was used as a housekeeping gene control (Chang et al., 2005). The primer pairs used for

RT-qPCR analysis were: 5’-CTTCTCGCCCATGACCAC-3’ and 5’-

CCCACCAGGTTGTTCTTCAG-3’ for CBLP; 5’-

CTTAATTGCATGCGCGCCGTCA-3’ and 5’-

TCAGAACACCAACGCAAGTTTCCAG-3’ for ARS2; 5’-

TGGCCATGCTTATCGTCATCTATG-3’ and 5'-TCGATGCGCATGACCAGGAT-

102 3' for SULTR1; 5’-ACGTGGCATGCAGCTCAT-3’ and 5'-

CTTGCCACTTTGCCAGGT-3' for SULTR2; 5'-AAGCTGGACCGGTGGTGAGAA-

3' and 5'-TCACATGTCAGTGCACGCCAG-3' for SULTR3; 5’-

ACGGGTTCTTCGAGCGAATTGC-3’ and 5'-

CGACTGCTTACGCAACAATCTTGG-3' for SLT1; 5’-

GTACGGAGTTCCTTACGCGC-3’ and 5'-TTCTTCGCCACCGATGAGC-3' for

SLT2; 5’-CCCAGTCTTTTGGCGGCAAG-3’ and 5'-

GGCCTACTCGCTACCGTACC-3' for SLT3; 5’-

GCGCTGCCCTCCGTCACCTTCC-3’ and 5’-CAGCCGCACGCCCGTCCAGTAG-

3’ for NIT1; 5’-TTCCGTTTCCGTTCTCTGAC-3’ and 5’-

CCCTGCATCTTGTTCTCCAG-3’ for PHOX.

Peptide synthesis and antibody production: Specific antibodies against FOX1,

CRD1 and COX2b were kind gifts from Dr. Sabeeha Merchant, University of

California, Los Angeles. The antibodies to SULTR2 and SLT2 were prepared by

Agrisera (Vännäs, Sweden), while the SLT general antibody was prepared by Covance

Research Products (Princeton, NJ). Monospecific antibodies for SLT2 were generated in rabbits against a peptide region (DGKYLSKPDPNW) unique to this transport protein (Figure 3-1B). This peptide was conjugated to keyhole limpet hemocyanin via a terminal cysteine and injected into rabbits. Resulting antibodies were affinity purified by column chromotography using an UltraLink Iodoacetyl Gel (Pierce,

Rockford, IL) covalently bound to the unique SLT2 peptide used for antibody production. SLT2 peptide synthesis, peptide conjugation, antibody production and antibody purification were all performed by Agrisera (Vännäs, Sweden). To generate

103 polyclonal antibodies that specifically recognize SULTR2, a cDNA encoding a unique region in the SULTR2 protein (Glu677 to Gln764; Figure 3-1A) was cloned in frame into the 3’ terminus of the 10XHis-encoding sequence of the expression vector pET-

16b (Novagen, Madison, WI) and introduced into E. coli for IPTG-inducible expression. E. coli cells expressing the fusion protein were grown to an A600 of 0.5 and then induced with IPTG for 4 h at 37°C. The 10XHis-SULTR2 protein was purified using a Ni-NTA Superflow column (Qiagen, Valencia, CA) and then subjected to preparative SDS-PAGE (15% polyacrylamide gel). The region of the gel containing the fusion protein was excised and used for antiserum production in rabbits. To generate general antibodies that would react with both SLT1 and SLT2 (and potentially with SLT3), cDNAs encoding the region of SLT1 and SLT2 from Thr337 to Val584 (Figure 3-1B) were cloned into pET-16b (the SLT1 and SLT2 amino acid sequences from this region are 91% identical and 97% similar). These expressed

10XHis-SLT1 and 10XHis-SLT2 tagged polypeptides were resolved by preparative

SDS-PAGE, as described above, and regions of the gel containing the fusion proteins were excised and used as antigens for antibody production.

Protein isolation, SDS-PAGE and immunoblot analysis: Chlamydomonas cells (2-4 x 106 cells mL-1 in 100 mL) were collected by centrifugation (3000 X g, 5 min) and resuspended in 0.1 M sodium phosphate buffer (pH 7.0). Chl was extracted from the cells into 80% acetone-20% methanol and its concentration determined spectrophotometrically (Arnon, 1949) after removal of cell debris and denatured proteins by centrifugation. Quantities of cells with equal chl content (200–300 µg chl) were pelleted, resuspended in an ice-cold homogenization buffer (0.25 M sucrose, 0.1

104 M HEPES pH 7.5, 15 mM EGTA, 5% glycerol, 0.5% PVP) containing a protease inhibitor cocktail (Sigma, St. Loius, MO), and then disrupted by agitation with glass beads (425-600 µm). After removal of cell debris by a brief centrifugation (2000 X g,

5 min), the supernatant was centrifuged at 100,000 X g for 50 min to obtain a microsomal pellet, which was then resuspended in the homogenization buffer containing 1% Triton X-100. An equal vol of loading buffer (6.25 mM Tris-HCl, pH

6.8, 5% SDS, 6 M urea, 500 mM dithiothreitol, 10% glycerol and 0.002% bromophenol blue) was added to the samples prior to an incubation at 42ºC for 15 min. Solubilized polypeptides were resolved by SDS-PAGE on a 10% polyacrylamide gel and transferred to Polyvinylidene Difluoride (PVDF) membranes using a wet transfer method. Blots were blocked in 5% milk in Tris-buffered saline solution with

0.1% Tween-20 prior to 1 h incubation in the presence of primary antibodies. The dilutions of the primary antibodies used were: 1:2,500 anti-SULTR2, 1:3,000 anti-

SLT, 1:1,000 anti-SLT2, 1:1,000 anti-FOX1, 1:1,000 anti-H+ ATPase (Agrisera,

Sweden), 1:1,000 anti-V ATPase (Agrisera, Sweden), 1:3,000 CRD1 and 1: 40,000 anti-COX2b. A 1:10,000 dilution of horseradish peroxidase-conjugated anti-rabbit IgG

(Promega, Madison, WI) was used as a secondary antibody. The peroxidase activity was detected by an enhanced chemiluminescence assay (Amersham Biosciences,

Sweden).

Cell treatment: Cycloheximide (CHX), which inhibits protein synthesis on 80S ribosomes, was used at a final concentration of 10 µg mL-1. This concentration effectively inhibits protein synthesis in Chlamydomonas (Kawazoe et al., 2000). The

105 proteasome inhibitor MG132 (carbobenzoxy-leucyl-leucyl-leucinal; Sigma, St. Loius,

MO) was added to a final concentration of 10 µM (Smalle and Vierstra, 2004).

Plasma membrane isolation: The D66 strain was used for plasma membrane isolations. 10 L of cells were grown to mid-logarithmic phase (3 x 106 cell mL-1) in a stirred bottle and starved for S for 24 h. Batches of resuspended cells (from 10 g pellet wet weight) were sonicated on ice using a Fisher Scientific Sonic Dismembrator model 550 with a microtip probe, power setting 3.5, and 15 cycles of 1 min of sonication, each followed by 1 min of cooling. The total microsomal fraction was separated into plasma membranes and a fraction containing the rest of the membranes using a two-phase aqueous polymer system described by Herbik et al. (Herbik et al.,

2002). To fractionate membrane protein from the top (polyethylene glycol) phase of the two-phase aqueous polymer system, 600 µg of protein was loaded onto a 4.5 mL,

15-45% sucrose gradient and centrifuged at 4°C for 16 h at 31,000 rpm in SW60Ti rotor (Beckman Coulter, CA). The gradient was separated into 11 fractions of equal vol, diluted five-fold in a dilution buffer (5 mM MOPS pH 7.0, 2 mM dithiothreitol, 1 mM EDTA, and 1 mM PMSF), and centrifuged at 4°C for 1 h at 50,000 rpm in the

TLA100.3 rotor (Beckman Coulter, CA). Pellets were resuspended in 50 µL of the dilution buffer and an equal vol of the loading buffer was added to the samples prior to an incubation at 42°C for 15 min. SDS-PAGE and immunoblot analyses were performed as described above.

Yeast strain, media and growth conditions: The strain of Saccharomyces cerevisiae used in this study was CP60-1C (MATa his3 leu2 ura3 trp1 sul1-1 sul2-1), which

2- harbors mutations in both high-affinity SO4 transporters (SUL1 and SUL2) (Cherest

106 et al., 1997). CP60-1C transformants were grown at 30ºC in synthetic defined (SD) –

Met –Ura liquid medium supplemented with 100 µM MgSO4. Cell growth was evaluated by measuring optical density of the cultures at 600 nm (A600) in a DU640 spectrophotometer (Beckman Coulter, CA).

2- Construction of GFP-tagged SO4 transporters, yeast transformation and

2- microscopy: Full-length cDNAs encoding SO4 transporters were cloned into pDR196-GW (no tag) or pDR196-GW-GFP (GFP was fused in-frame to the transporters at the carboxyl terminus). The pDR196-GW and pDR196-GW-GFP plasmids were kind gifts from Dr. Dominique Loque. The transporters were expressed in yeast under the control of the constitutive PMA1 promoter. The constructs carrying

2- SO4 transporters were transformed into the yeast mutant CP60-1C, using the lithium- acetate procedure (Rose et al., 1990). Transformants were selected on SD medium lacking uracil and then grown in liquid medium to mid-logarithmic phase. For microscopy, a drop of cell suspension was mounted onto slides and GFP fluorescence was detected as previously described (using a Nikon TMD200 inverted fluorescence microscope equipped with a Nikon 60X 1.2-numerical aperture water immersion objective and a Biorad MRC 1024 confocal head) (Shibagaki and Grossman, 2004).

Yeast protein isolation and immunoblot analysis: Cells in mid-logarithmic phase

(A600 ~ 0.2) were harvested by centrifugation (3000 X g for 5 min), washed once with ice-cold STE10 buffer (10% w/v sucrose, 5 mM Tris pH 7.4, 10 mM EDTA) and resuspended in the same buffer containing protease inhibitor cocktail (Calbiochem,

Gibbstown, NJ). Cells were disrupted by agitation with glass beads (425 – 600 µM)

107 and the cell debris was removed by a brief centrifugation (3000 X g for 5 min). The supernatant was centrifuged at 100,000 X g for 50 min to obtain a microsomal pellet, which was then resuspended in the STE10 buffer. An equal vol of loading buffer (6.25 mM Tris-HCl, pH 6.8, 5 % SDS, 6 M urea, 500 mM dithiothreitol, 10 % glycerol and

0.002 % bromophenol blue) was added to the samples prior to an incubation at 42ºC for 15 min. Solubilized polypeptides were resolved by SDS-PAGE and the immublot performed as described earlier. Dilutions of primary antibodies used were: 1:1000 anti-PMA1 (ABCam, Cambridge, MA) and 1:1000 anti-GFP (Roche, Nutley, NJ). A

1:10,000 dilution of horseradish peroxidase-conjugated anti-rabbit IgG (Promega,

Madison, WI) or 1:10,000 dilution of horseradish peroxidase-conjugated anti-mouse

IgG (Sigma, St. Louis, MO) was used as a secondary antibody. The peroxidase activity was detected by an enhanced chemiluminescence assay (Amersham

Biosciences, Sweden).

Generation of Chlamydomonas mutants: To generate a library of insertional mutants, a 1.7 kb PCR fragment containing the selectable marker gene AphVIII

(conferring resistance to paromomycin) (Sizova et al., 2001) under the control of the

PSAD promoter was used for transformation. The cell wall-less strain D66 was transformed by electroporation (Shimogawara et al., 1998) using a modified procedure reported by Colombo (Colombo et al., 2002). After transformation, cells were incubated in TAP medium for 16-18 h to allow for the accumulation of the AphVIII protein. Transformants were subsequently selected on TAP medium supplemented with 5 µg mL-1 paromomycin.

108 Generation and screening of the mutant library: The mutant screen was performed according to Gonzalez-Ballester (Gonzalez-Ballester, 2005). In brief, transformants were generated as described above and transferred to 96-well microtiter plates containing approximately 100 µL of liquid TAP medium and grown for 5 d before they were used for genomic DNA isolation; the plates were wrapped in parafilm to minimize evaporation. Each microtiter plate represents a pool of 96 insertional mutants. Genomic DNA was isolated from a pool of 96 transformants using a standard phenol-chloroform protocol (Sambrook et al., 1989). An equal concentration of genomic DNA from 10 pools was combined to create a “superpool” of DNA at a concentration of 100 ng/µL. To screen for mutants of interest, the superpool DNA was used as a template for PCR. Each PCR reaction was performed using one gene- specific primer (gene for which a disruption is sought; Table 3-2) and one AphVIII- construct-specific primer (RB2 5’-TACCGGCTGTTGGACGAGTTCTTCTG-3’).

Multiple gene-specific primers were paired with the AphVIII primer to increase our chances of identifying a disrupted target gene. PCR reactions were performed in a final vol of 25 µL containing 0.2 µL Taq DNA Polymerase (Qiagen, Valencia, CA),

2.5 µL 10X PCR buffer, 2 µL dNTPs (2.5 mM each), 4 µL Q solution (Qiagen,

Valencia, CA), 1 µL DMSO (Sigma, St. Loius, MO), 1 µL template, 400 nM of each primer, and distilled water to make up the remainder of the 25 µL vol. Conditions used for amplification in the thermocycler were: pre-incubation at 95°C for 5 min followed by 35 cycles of sequential denaturation at 95°C for 30 s, annealing at 60°C for 30 s, and amplification at 72°C for 120 s. PCR products were separated by gel electrophoresis using 0.8% agarose gels. To identify DNA regions adjacent to the right

109 border of the AphVIII construct (downstream from PSAD promoter), the PCR product amplified with the gene-specific primer and RB2 was excised from the gel, purified using the QIAquick PCR Purification Kit (Qiagen, Valencia, CA), and sequenced.

Primers used for this screening are listed in Table 3-2.

Southern blot analyses: Genomic DNA was isolated from 50 mL liquid cultures of the wild-type strain 21gr, slt1, slt2, and sultr2 mutants using a standard phenol- chloroform extraction protocol (Sambrook et al. 1989). 10 µg of genomic DNA was digested for 20 h with 10 u of restriction endonucleases (PstI; New England Biolabs).

The fragments were separated by agarose (0.8%) gel electrophoresis, blotted overnight in 20X SSC onto nylon membranes (GeneScreen, DuPont-New England Nuclear), and the transferred DNA cross-linked to the membrane by UV illumination. An alkaline phosphatase-labeled probe was synthesized by chemical cross-linking of a thermostable alkaline phosphatase to the nucleic acid template. Probe synthesis and hybridization were performed using the Amersham AlkPhos DirectTM Labeling and

Detection Systems following the manufacturer’s protocol (Amersham Biosciences,

Sweden).

Rescue of the mutant phenotype: The phenotype of the slt1slt2sultr2 mutant was rescued by introducing wild-type alleles encoding the individual transporters into the triple mutant background. The slt1slt2sultr2 strain was crossed to D66 wild-type strain

2- and the SO4 transport rate of the progeny containing one (slt1, slt2, or sultr2) or two

2- (slt1slt2, slt1sultr2, slt2sultr2) mutations was determined by SO4 uptake assays.

Accession numbers: Sequence data from this article can be found in the

EMBL/GenBank data libraries under accession numbers NM_179568 (AtSULTR1;2),

110 2- CAA57710 (ShSHST1), XP_001766939 (putative SO4 permease from

Physcomitrella patens), GU181275 (SLT1), GU181276 (SLT2), GU181277

(SULTR2).

111 ACKNOWLEDGEMENTS: We thank Dr. Nakako Shibagaki, Dr. Jeffrey Moseley and Dr. Florence Mus for valuable discussions of the results, and all members of the

Grossman and Bhaya laboratories for support and advice. We thank Ariana Afshar,

Matthew Prior and Leonardo Magneschi for their help with the mutant screen. We would also like to thank Dr. Sheng Luan and Dr. Wenzhi Lan for their help with the

Xenopus oocyte experiment, and Dr. Mark Dudley Page for his advice on the plasma membrane isolation. The antibodies for FOX1, CRD1 and COX2b were generously provided by Dr. Sabeeha Merchant. The plasmids pDR196-GW and pDR196-GW-

GFP were kindly provided by Dr. Dominique Loque.

112 A

CrSULTR1 MRRNPSAGSAVAQADGSMAPSTSLLHPDGAGAALELSSSMMMPTDCSVINGRHGAAGASSGGAQMEFPWE CrSULTR2 MKRNTSN------VDTGGVPAPLNS---TPSTRLIQNG-YGDSKYET--ERMEFPFP AtSULTR4;1 MSYASLS------VKDLTS--LVSRSGTGSSSSLKPPGQTRPVKVIPLQHP AtSULTR1;2 ------MSSRAHPVDGSPATDGGHVPMKPSPTRHKVGIPPK ShSHST1 MSQRVSD------QVMADVIAETRSNSSSHRHGGGGGGDDTTSLPYMHKVGTPPK

CrSULTR1 NTNGNADRSSVHGTLQKVWERSKSSYQLKMSTYSALDWLAFFLPCVRWLRTYKIREYLFADIVAGISVGF CrSULTR2 EDPRYHPRDSVKG----AWEKVKEDHHHRVATYNWVDWLAFFIPCVRWLRTYR-RSYLLNDIVAGISVGF AtSULTR4;1 DTSNEARPPSIP------FDDIFSGWTAKIKRMRLVDWIDTLFPCFRWIRTYRWSEYFKLDLMAGITVGI AtSULTR1;2 QNMFKDFMYTFKET--FFHDDPLRDFKDQPKSKQFMLGLQSVFPVFDWGRNYTFKKFRG-DLISGLTIAS ShSHST1 QTLFQEIKHSFNET--FFPDKPFGKFKDQSGFRKLELGLQYIFPILEWGRHYDLKKFRG-DFIAGLTIAS

CrSULTR1 MVVPQGMSYANLAGLPSVYGLYGAFLPVITYALVGSSRQLAVGPVAVTSLIIGSSLKELVPGAETISNPN CrSULTR2 MVVPQGLSYANLAGLPSVYGLYGAFLPCIVYSLVGSSRQLAVGPVAVTSLLLGTKLKDILPEAAGISNPN AtSULTR4;1 MLVPQAMSYAKLAGLPPIYGLYSSFVPVFVYAIFGSSRQLAIGPVALVSLLVSNALGGIA------AtSULTR1;2 LCIPQDIGYAKLANLDPKYGLYSSFVPPLVYACMGSSRDIAIGPVAVVSLLLGTLLRAEIDP------ShSHST1 LCIPQDLAYAKLANLDPWYGLYSSFVAPLVYAFMGTSRDIAIGPVAVVSLLLGTLLSNEIS------

CrSULTR1 Q-LTPDQEVIQEKYNMLAIQLSLLVAILYTSVGVFRLGFLTNFLSHSVIGGFTSGAAITIGLSQVCHVWL CrSULTR2 IPGSPELDAVQEKYNRLAIQLAFLVACLYTGVGIFRLGFVTNFLSHAVIGGFTSGAAITIGLSQV----- AtSULTR4;1 ------DTNEELHIELAILLALLVGILECIMGLLRLGWLIRFISHSVISGFTSASAIVIGLSQI----- AtSULTR1;2 ------NTSPDEYLRLAFTATFFAGITEAALGFFRLGFLIDFLSHAAVVGFMGGAAITIALQQL----- ShSHST1 ------NTKSHDYLRLAFTATFFAGVTQMLLGVCRLGFLIDFLSHAAIVGFMAGAAITIGLQQL-----

CrSULTR1 KYILGISIP----RTERLHDQVSTYIEFIRNLKWQEFIMGSTFLVLLVTMKEVGKRSHRFRWLRPLGPIS CrSULTR2 KYILGISIP----RQDRLQDQAKTYVDNMHNMKWQEFIMGTTFLFLLVLFKEVGKRSKRFKWLRPIGPLT AtSULTR4;1 KYFLGYSIA----RSSKIVPIVESIIAGADKFQWPPFVMGSLILVILQVMKHVGKAKKELQFLRAAAPLT AtSULTR1;2 KGFLGIKK--FTKKTDIISVLESVFKAAHHGWNWQTILIGASFLTFLLTSKIIGKKSKKLFWVPAIAPLI ShSHST1 KGLLGISNNNFTKKTDIISVMRSVWTHVHHGWNWETILIGLSFLIFLLITKYIAKKNKKLFWVSAISPMI

CrSULTR1 VCIIALLAVYIGHVDKKGIKIIGAIKKGLPTP-TVGWWAPMPDFVDLIPIAIVVMLVDLLESTSIARALA CrSULTR2 VCIIGLCAVYVGNVQNKGIKIIGAIKAGLPAP-TVSWWFPMPEISQLFPTAIVVMLVDLLESTSIARALA AtSULTR4;1 GIVLGTTIAKVFHPPS--ISLVGEIPQGLP---TFSFPRSFDHAKTLLPTSALITGVPILESVGIAKALA AtSULTR1;2 SVIVSTFFVYITRADKQGVQIVKHLDQGINPSSFHLIYFTGDNLAKGIRIGVVAGMVALTEAVAIGRTFA ShSHST1 SVIVSTFFVYITRADKRGVSIVKHIKSGVNPSSANEIFFHGKYLGAGVRVGVVAGLVALTEAIAIGRTFA

CrSULTR1 NKNKYELVANQEIVGLGLANFAGAAFHCYSTTGSFSRSAVNNESGAKTGLAGFVTAWVVGFVLLFLTPVF CrSULTR2 RKNKYELHANQEIVGLGLANFAGAIFNCYTTTGSFSRSAVNNESGAKTGLACFITAWVVGFVLIFLTPVF AtSULTR4;1 AKNRYELDSNSDLFGLGVANILGSLFSAYPATGSFSRSAVNNESEAKTGLSGLITGIIIGCSLLFLTPMF AtSULTR1;2 AMKDYQIDGNKEMVALGMMNVVGSMSSCYVATGSFSRSAVNFMAGCQTAVSNIIMSIVVLLTLLFLTPLF ShSHST1 AMKDYALDGNKEMVAMGTMNIVGSLSSCYVTTGSFSRSAVNYMAGCKTAVSNIVMSIVVLLTLLVITPLF

CrSULTR1 EKLPYCTLGAIVCSSVTGLLEYEQAIYLWKVNKLDFLVWMASFLGTLFISIEIGLGIAIGLAMLIVIYES CrSULTR2 AHLPYCTLGAIIVSSIVGLLEYEQAIYLWKVNKLDWLVWMASFLGVLFISVEIGLGIAIGLAILIVIYES AtSULTR4;1 KYIPQCALAAIVISAVSGLVDYDEAIFLWRVDKRDFSLWTITSTITLFFGIEIGVLVGVGFSLAFVIHES AtSULTR1;2 KYTPNAILAAIIINAVIPLIDIQAAILIFKVDKLDFIACIGAFFGVIFVSVEIGLLIAVSISFAKILLQV ShSHST1 KYTPNAVLASIIIAAVVNLVNIEAMVLLWKIDKFDFVACMGAFFGVIFKSVEIGLLIAVAISFAKILLQV

CrSULTR1 AFPHTAMLGRIPGSGVYRNVKQYPQSQLTPGILVMRIDSPIYFANVQWIKDRLRVYEDRHRDWS--GEHG CrSULTR2 AFPNTALVGRIPGTTIWRNIKQYPNAQLAPGLLVFRIDAPIYFANIQWIKERLEGFASAHRVWS--QEHG AtSULTR4;1 ANPHIAVLGRLPGTTVYRNIKQYPEAYTYNGIVIVRIDSPIYFANISYIKDRLREYEVAVDKYTNRGLEV AtSULTR1;2 TRPRTAVLGNIPRTSVYRNIQQYPEATMVPGVLTIRVDSAIYFSNSNYVRERIQRWLHEEEEKVK-AASL ShSHST1 TRPRTAVLGKLPGTSVYRNIQQYPKAAQIPGMLIIRVDSAIYFSNSNYIKERILRWLIDEGAQRT-ESEL

CrSULTR1 TKLEFAILDMSPVTHIDATGVHALEGWIEHFAHVGTQLVLCNPSVKVIRELETAGVPDMLGRDWIFVTVH CrSULTR2 VPLEYVILDFSPVTHIDATGLHTLETIVETLAGHGTQVVLANPSQEIIALMRRGGLFDMIGRDYVFITVN AtSULTR4;1 DRINFVILEMSPVTHIDSSAVEALKELYQEYKTRDIQLAISNPNKDVHLTIARSGMVELVGKEWFFVRVH AtSULTR1;2 PRIQFLIIEMSPVTDIDTSGIHALEDLYKSLQKRDIQLILANPGPLVIGKLHLSHFADMLGQDNIYLTVA ShSHST1 PEIQHLITEMSPVPDIDTSGIHAFEELYKTLQKREVQLILANPGPVVIEKLHASKLTELIGEDKIFLTVA

CrSULTR1 DAVSFCSRQLAEAGMAVT------PLSMTQQPSSTSDE------CrSULTR2 EAVTFCSRQMAERGYAVKEDNTSSYPHFGSRRTPGALPAPSSQLDSSPPTSVTESTSGTPAAGTYSSIGG AtSULTR4;1 DAVQVCLQYVQSSNLEDK------HLSFTRRYGGSNNNSSSSNALL AtSULTR1;2 DAVEACCPKLSNEV------ShSHST1 DAVATYGPKTAAF------

CrSULTR1 ------CrSULTR2 AVPAVAGHTAAGNGGSHSPSAQPGVQLTTTGSQRQQ AtSULTR4;1 KEPLLSVEK------AtSULTR1;2 ------ShSHST1 ------

113 B

CrSLT1 ------CrSLT2 ------CrSLT3 ------XP_001766939 MTRSMPLYRGEQEEMWFSHTESIKTTPSATTNAPLSDGIRIPRFHGVRGGPDPMHRNPDLRNVAVLLSCS

CrSLT1 ------MAALSWQGIV CrSLT2 ------MGFGWQGSV CrSLT3 ------MAAIGWPGIV XP_001766939 VQGGEVLDLGVVPGAKPALYCWFGFMISSLLNCVMNCLFEFDFVESAENSGRELRRESDKMVQLGWESYL

CrSLT1 AVTFTALAFVVMAADWVGPDITFTVLLAFLTAFDGQIVTVAKAAAGYGNTGLLTVVFLYWVAEGITQTGG CrSLT2 SIAFTALAFVVMAADWVGPDVTFTVLLAFLTAFDGQIVTVAKAAAGYGNTGLLTVIFLYWVAEGITQTGG CrSLT3 AIISVAISFIIMAADWVGPDITFTILLSWLTAFDGKIITVAKAAAGYGNTGLLTVIFLYWVAEGVTQTGG XP_001766939 VLATLIAGLVVMAGDWVGPDFVFALMVGFLTACR--VITVKESTEGFSQNGVLTVVILFVVAEGIGQTGG

CrSLT1 LELIMNYVLGRSRSVHWALVRSMFPVMVLSAFLNNTPCVTFMIPILISWGRRCGVPIKKLLIPLSYAAVL CrSLT2 LELIMNFVLGRSRSVHWALARSMFPVMCLSAFLNNTPCVTFMIPILISWGRRCGVPIKKLLIPLSYASVL CrSLT3 LELVMNYVLGRSRSVHWALVRSMFPVMVLSAFLNNTPCVTFMIPILMSWARRCGVPPKKLLIPLSYAAVL XP_001766939 MEKALNLLLGKATSPFWAITRMFIPVAITSAFLNNTPIVALLIPIMIAWGRRNRISPKKLLIPLSYAAVF

CrSLT1 GGTCTSIGTSTNLVIVGLQDARYAKSKQVDQAKFQIFDIAPYGVPYALWGFVFILLAQGFLLPGNSSRYA CrSLT2 GGTCTSIGTSTNLVIVGLQDARYTKAKQLDQAKFQIFDIAPYGVPYALWGFVFILLTQAFLLPGNSSRYA CrSLT3 GGTCTSIGTSTNLVIVGMQDTRYNKQNKEDEAKFGMFDIAPYGVPYALMGFVFIILTQRFLLPGNSSRYA XP_001766939 GGTLTQIGTSTNFVISSLQEKRYTQLKRPGDAKFGMFDITPYGIVYCIGGFLFTVIASHWLLPSDETKRH

CrSLT1 KDLLLAVRVLPSSSVVKKKLKDSGLLQQNGFDVTAIYRNGQLIKISDPSIVLDGGDILYVSGELDVVEFV CrSLT2 KDLLIAVRVLPSSSVAKKKLKDSGLLQQSGFSVSGIYRDGKYLSKPDPNWVLEPNDILYAAGEFDVVEFV CrSLT3 KDLLIARLLVTT------IQQS----SNEPTTRRACRQVDPDTVLEANDILYCAGELDVVEFV XP_001766939 SDLLLVARVPPESPVANNTVREAGLKGMERLFLVAVERQGRVTHAVGPQYLLEPEDLLYFCGELEQAHFY

CrSLT1 GEEYGLALVNQEQE--LAAERPFGSGEEAVFSANGAAPYHK---LVQAKLSKTSDLIGRTVREVSWQGRF CrSLT2 GEEFGLGLVNADAE--TSAERPFTTGEESVFTPTGGAPYQK---LVQATIAPTSDLIGRTVREVSWQGRF CrSLT3 GEEFGLGLVTAETERALTDGQAVGDSEATAFHDTGASPYKK---LVQVTMTKTADLVGRTVREVSWQGRF XP_001766939 SKAFSLELLTNEAISGSKRANFQGEKHPSALENGSCGSVEDSTLIMQASVRKGADIIGKTLDQIDFRKRF

CrSLT1 GLIPVAIQRGNGREDGRLSDVVLAAGDVLLLDTTPFYDEDREDIKTNFDGKLHAVKDGAAKEFVIGVKVK CrSLT2 GLIPVAIQRGNGREDGRLNDVVLAAGDVLILDTTPFYDEEREDSKNNFAGKVRAVKDGAAKEFVVGVKVK CrSLT3 GLIPVAIQRGNGREDGRLNDVVLAAGDVLILDTTPHFDEARDDFKINFE-KLRFVKDGAAKEFVIGVKVK XP_001766939 DVAVLGLKRGETHQPGPLSEMVVNANDVLVLLGDNEEVLQKPEVKAVFK-DVEKLDEALEKEYLTGMKVT

CrSLT1 KSAEVVGKTVSAAGLRGIPGLFVLSVDHADGTSVDSSDYLYKIQPDDTIWIAADVAAVGFLSKFPGLELV CrSLT2 KSSEVVNKTVSAAGLRGIPGLFVLSVDRADGSSVEASDYLYKIQPDDTIWIATDIGAVGFLAKFPGLELV CrSLT3 KNSEVVNKTVTAAGLRGVPGLFVLSVDRADGSSVDASDYLYKIQPGDTLWLAADVGAVGFLSKFPGLELV XP_001766939 NRFKGVGKTVYDAGLRGINGLTLLAIDRQSGEHLKFIEDDTVVELGDTLWFAGGVQGVHFLLKISGLEHS

CrSLT1 QQEQVDKTGTSILYRHLVQAAVSHKGPLVGKTVRDVRFRTLYNAAVVAVHRENARIPLKVQDIVLQGGDV CrSLT2 QQEQVDKTGTSILYRHLVQAAVSHKGPIVGKTVRDVRFRTLYNAAVVAVHREGARVPLKVQDIVLQGGDV CrSLT3 QQEQVDKTGTSILYRHLVQAAVSHKGPLVGKTVRDVRFRTLYNAAIVAVHREGVRVPLKVQDIVLQGGDV XP_001766939 QAPQVSKLRADILYRQLVKASVASESPLVGNTVREAHFRNKYDAVVLAIHRQGERLSMDVRDVKLRAGDV

CrSLT1 LLISCHTNWADEHRHDKSFVLVQPVPDSSPPKRSRMIIGVLLATGMVLTQIIGG-LKNKEYIHLWPCAVL CrSLT2 LLISCHTNWADEHRHDKSFVLLQPVPDSSPPKRSRMVIGVLLATGMVLTQIVGG-LKSREYIHLWPAAVL CrSLT3 LLISCHTKWAEEHRMDKAFVLVQAVPDSSPPKRGRMAIGVLLVVGMVLTQIVGG-LKEKEYIHLWPAAVL XP_001766939 LLLDTGSNFGHRYRNDAAFSLISGVPESSPVKKSRMWVALFLGAAMIATQIVSSSIGGTELINLFTAGIL

CrSLT1 TAALMLLTGCMNADQTRKAIMWDVYLTIAAAFGVSAALEGTGVAAKFANAIISIGKGAGGTGAALIAIYI CrSLT2 TSALMLLTGCMNADQARKAIYWDVYLTIAAAFGVSAALEGTGVAASFANGIISIGKNLHSDGAALIA--- CrSLT3 TAALMLLTGCMNADQARKAIMWDVYLTIAAAFGVSAALENTGVAGKVANAIISIGKSIGGDGPALIAIYV XP_001766939 TSGLMLLTRCLSADQARNSIDWRVYTTIAFAIAFSTCMEKSKLARAIADIFIKISESIGGMRASYVAIYI

CrSLT1 ATALLSELLTNNAAGAIMYPIAAIAGDALKITPKDTSVAIMLGASAGFVNPFSYQTNLMVYAAGNYSVRE CrSLT2 -TAMLSELLTNNAAGAIMYPIAAIAGDALKISPKETSVAIMLGASAGFINPFSYQCNLMVYAAGNYSVRE CrSLT3 ATAVMSELLTNNAAGAIMYPIAAIAGDQLKIPAVDISVAIMLGASAGFINPFSYQTNLMVYAAGNYSVRE XP_001766939 ATALLSELVSNNAAAAIMYPIAADLGDALGVVPTRMSVVVMLGASAGFTLPYSYQTNLMVYAAGDYRFME

CrSLT1 FAIVGAPFQVWLMIVAGFILVYRNQWHQVWIVSWICTAGIVLLPALYFLLPTRIQIKIDGFFERIAAVLN CrSLT2 FAIIGAPFQIWLMIVAGFILCYMKEWHQVWIVSWICTAGIVLLPALYFLLPTKVQLRIDAFFDRVAQTLN CrSLT3 FATIGAPFQIWLMVVASFILCYMKQWKQVWIATWSITAFIVFVPALLTLLPHTVQNRMEAFFDRIAEAIN XP_001766939 FAKFGLPCQCFMIITVILIFLLDN---RIWVAVGLGFALMLVVLGWHLVW------EFVPASIR

CrSLT1 PKAALERRRSLRRQ------VSHTRTDDSGSSGSP---LPAPKIVA- CrSLT2 PKLIIERRNSIRRQ------ASRTGSDGTGSSDSPR-ALGVPKVITA CrSLT3 PRAALQRRRSARAQSFGGKAMSVGSTESRTDGSSTPDVALTFIEMPKMGVR XP_001766939 SKFSPGRKE------KTEKIEQ------

114 2- Figure 3-1. Amino acid sequence alignments of SO4 transporter proteins. A.

Predicted CrSULTR1 and CrSULTR2 amino acid sequences were aligned with each other and with representative Arabidopsis thaliana and Stylosanthes hamata high-

2- affinity SO4 transporters using BioEdit version 7.0.9.0 software. B. Predicted

2- CrSLT1, CrSLT2, CrSLT3 proteins were aligned with each other and a putative SO4 permease from Physcomitrella patens (PpST). Black and grey shadings indicate identical and similar amino acid residues, respectively. The red bar (A) highlights the

C-terminal STAS domain and the blue bar (A) represents the region of CrSULTR2 used as an antigen for antibody production. The magenta bar (B) indicates the peptide sequence in SLT2 that is recognized by the SLT2 antibody; the orange bar indicates the TrkA-C domain, and the green bar (B) shows the region recognized by the general

SLT antibody.

115

600

)

-1 +S 500 -S 400 cells) 5 300 (10 uptakerate -1 2-

4 200

SO 100 (fmol s

0 WTWT sac1sac1 snrk2.1ars11

2- Figure 3-2. SO4 uptake in wild-type (WT) Chlamydomonas cells and the sac1 and snrk2.1 mutant strains grown in S-replete (+S) and 24 h in S-depleted (-S) media.

2- -1 Uptake rates were determined at 250 μM SO4 (specific activity 330 Ci mol .) The

2- initial SO4 uptake rate was calculated from the slope of the line defining the transport

2- of SO4 into cells as a function of time. The values are averages of three experiments with a single biological replicate, although similar results were obtained in independent experiments, and the bars represent one standard deviation.

116

replete -S -N -P

Figure 3-3. Real time quantitative PCR analysis of the levels of various transcripts.

Levels of SULTR2, SLT1, SLT2, SULTR1, SLT3, SULTR3, ARS2, PHOX and NIT1 transcripts in cells deprived of S, N or P (samples were collected 12 h after transferring cells from nutrient-replete medium to medium devoid of S, N or P).

Levels of individual transcripts are represented as a relative abundance with respect to the housekeeping control gene (CBLP).

117

118 2- Figure 3-4. Changes in transcript abundances for various SO4 transporters following imposition of S deprivation. Levels of transcripts encoding ARS2, SULTR2, SLT1,

SLT2, SULTR1, SLT3 and SULTR3 were measured by RT-qPCR for wild-type (WT) cells (A), and the sac1 (B) and snrk2.1 (C) mutants. RNA samples were isolated from cells 2, 6, and 24 h after they were transferred from TAP to TAP–S medium. The expression of ARS2 was used a positive control for proper S-starvation responses in wild-type cells.

119

WT sac1 snrk2.1

kDa 0 4 24 0 4 24 0 4 24 (h)

95 SLT

95 SLT2

SULTR2 72 17 0 FOX1 13 0

Figure 3-5. SULTR2, SLT1, and SLT2 polypeptide abundances in wild-type (WT), sac1, and snrk2.1 strains. The time courses show accumulation of SLT1, SLT2, and

SULTR2 polypeptides following transfer of cells from S-replete to S-deficient medium. Samples were taken prior to, and 4 and 24 h after the cells were transferred.

The ferroxidase, FOX1, protein served as a loading control.

120

121 Figure 3-6. Accumulation of SULTR2, SLT1, and SLT2 transcripts and proteins and

2- their decay following the addition of SO4 to S-deprived cells. Time course of accumulation of SULTR2, SLT1, and SLT2 mRNAs (A) and polypeptides (B) upon transfer of cells from S-replete to TAP-S medium. After 24 h in TAP-S, 1 mM MgSO4 was added to starved culture. Samples were taken at 1, 2, 4, 8, 24 and 32 h after the

2- starvation (from +S to –S) and 1, 2, 4 and 8 h after the addition of SO4 back to cultures that had been S-starved for 24 h. RNA and protein isolations are described in

MATERIALS AND METHODS. A ferroxidase, FOX1, served as a loading control

(accumulation of FOX1 is S-independent).

122

2- Figure 3-7. Cycloheximide inhibition of accumulation of the SO4 transporter protein during S deprivation. Chlamydomonas cells were grown in TAP and then transferred to TAP-S medium. At the time of S removal, a transcriptional inhibitor actinomycin D

(ActD), a cytosolic translational inhibitor cycloheximide (CHX), or an organellar translational inhibitor chloramphenicol (Cm), was added to the cultures. Samples were taken prior to starvation as well as 2 and 6 h after the removal of S. The ferroxidase protein, FOX1, served as the loading control (accumulation of FOX1 is S- independent).

123

Figure 3-8. Determination of half-lives of SULTR2, SLT1, and SLT2 proteins and the role of the proteasome in degradation. A. Chlamydomonas cells were grown in TAP and transferred to TAP-S. After 10 h of starvation, the culture was split into three

2- aliquots: CHX was added to aliquots 1 and 3 and 1mM SO4 was added to aliquots 2 and 3. Samples were taken prior to starvation (lane 1), after 10 and 14 h of starvation

(lanes 2 and 3, respectively), and 20, 60, and 240 min after the addition of CHX (lanes

124 2- 2- 4-6), SO4 (lanes 7-9), and SO4 + CHX (lanes 10-12). B. Identical cultures were used for this analysis, except that the cells starved for S for 10 h were administered

2- 2- SO4 + CHX (lanes 4-6) or SO4 + CHX + MG132 (lanes 7-9). The experiment was repeated three times, yielding essentially identical results, with representative data shown. The signal strength of each band was quantified by ImageJ software

(Abramoff et al., 2004) and fitted to the exponential decay equation to obtain half- lives (t1/2). A cytochrome c oxidase, COX2b, served as a loading control.

125

Figure 3-9. Localization of the Chlamydomonas SLT1, SLT2, and SULTR2 transport proteins. A. Chlamydomonas strain D66 was grown in TAP medium to mid- logarithmic phase and then grown in TAP-S for an additional 24 h. Total membranes

(TM) were isolated and partitioned into plasma membrane enriched fraction

(polyethylene glycol fraction; PM) and a fraction containing all other membranes

126 (dextran fraction; OM). 20 µg of protein from each fraction was resolved by SDS-

PAGE and transferred to PVDF membrane for immunoblot analyses with anti-SLT, anti-SLT2, anti-SULTR2, anti-plasma membrane H+-ATPase, anti-FOX1 ferroxidase

(plasma membrane), anti-CRD1 (thylakoid membrane and chloroplast envelope), anti-

COX2b (mitochondrial inner membrane), and anti-V ATPase (tonoplast). B. To separate plasma membrane from the tonoplast, 600 µg of protein from the polyethylene glycol phase was loaded onto a 15-45% sucrose gradient and separated into 11 fractions of equal vol (fraction 1 is at the bottom (B) of the gradient and fraction 11 is at the top (T) of the gradient). The proteins of each fraction were resolved by SDS-PAGE and transferred to PVDF membrane for immunoblotting, as described in MATERIALS AND METHODS.

127

B I II

III IV

128 2- Figure 3-10. Expression of SO4 transporter-GFP fusion proteins in S. cerevisiae cells. A, CP60-1C cells transformed with an empty pDR196-GW-GFP, plasmids carrying SLT1, SLT2, or SULTR2 were grown to mid-logarithmic phase and the microsomal fraction was isolated, separated by SDS-PAGE and the immunoblot performed to detect the chimeric transporters tagged with GFP and a plasma membrane ATPase, PMA1. B, Confocal images of CP60-1C expressing the fusion proteins SULTR2-GFP (I), SLT1-GFP (II), SLT2-GFP (III) or AtSULTR1;2-GFP

(IV). The bar on the image represents 3 µm.

129

130 Figure 3-11. The copy number and position of the insertion of the AphVIII marker

2- gene in the SO4 transporter mutants. A. Diagram showing the site of AphVIII insertion in the SLT1, SLT2 and SULTR2 genes. Black bars, grey bars and hatched bars represent exons, introns and untranslated regions, respectively. B. DNA gel blot analysis of wild-type, slt1, slt2, and sultr2 strains. Genomic DNA was digested with

PstI, separated by agarose gel electrophoresis, transferred to a nitrocellulose membrane and hybridized with the 1.7-kb PSAD::AphVIII PCR fragment. Asterisks designate the newly introduced PSAD::AphVIII gene sequences. Size standards (in kilobases) are shown to the left.

131

132 133 Figure 3-12. Amino acid sequence alignments of wild-type and the SLT1 (A) and

SULTR2 (B) mutant gene products. The amino acid sequence of the proteins encoded by the slt1 and sultr2 mutant genes was deduced. In both cases, the insertions caused a frame-shift mutation generating a premature stop codon and the production of truncated proteins. The regions of amino acid sequences in the slt1 and sultr2 strains deviated from the corresponding wild-type polypeptides are underlined. Predicted proteins from wild-type (WT) and mutant (m) strains were aligned using BioEdit version 7.0.9.0 software.

134

135 Figure 3-13. Real-time quantitative PCR for analysis of transcript accumulation in wild-type and mutant strains. The accumulation of ARS2, SLT1, SLT2, SLT3, SULTR1,

SULTR2, and SULTR3 transcripts in wild-type cells and in the single (slt1, slt2, sultr2)

(A), double (slt1slt2, slt1sultr2, slt2sultr2) and triple (slt1slt2sultr2) mutants (B). RNA samples were collected prior to, and 4 and 24 h after the cells were transferred from

TAP to TAP–S medium. Levels of individual transcripts are given as relative abundance with respect to the housekeeping control gene (CBLP).

136

2- Figure 3-14. SULTR2, SLT1, and SLT2 polypeptide abundances in SO4 transporter mutants. The wild-type and single mutants (A); the double and triple mutants (B). The time courses show accumulation of SLT1, SLT2, and SULTR2 polypeptides following transfer of cells from S-replete to S-deficient medium. Samples were taken prior to, and 4 and 24 h after the cells were transferred. The COX2b protein served as a loading control.

137

2- Figure 3-15. Characteristics of SO4 transport in wild-type cells and single, double and triple mutants in S-replete medium or medium devoid of S (deprived of S for 24

2- h). Transport assays were performed as a function of external SO4 concentrations: A,

2- -1 0.02-0.2 µM; B, 2-200 µM. Initial rates of uptake are expressed as fmol of SO4 s

138 (105 cells)-1. Values are averages of 2-4 biological replicates; each biological experiment was performed in duplicate. Error bars represent one standard deviation.

139

2- Figure 3-16. Characteristics of SO4 transport in wild-type cells and single, double and triple mutants (progeny of a cross between a wild-type D66 strain and an slt1slt2sultr2 triple mutant) deprived of S for 24 h. Transport assays were performed

2- as a function of external SO4 concentrations: A. 0.02-0.2 µM; B. 2-200 µM. Initial

140 2- -1 5 -1 rates of uptake are expressed as fmol of SO4 s (10 cells) . Values are averages of

2-3 technical replicates, and error bars represent one standard deviation.

141

Table 3-1. Growth rates of the yeast CP60-1C strain harboring genes encoding

2- Arabidopsis SULTR1;2 or various Chlamydomonas SO4 transporters. Doubling time

(in h) is measured as a change in A600.

142

Primer Sequence SULTR2-5’UTR-F1 5'- TAACGGGCCTCCGCAAGACA -3' SULTR2-F8 5'- TTCCCTCCGTGTACGGCCTGTA -3'

SULTR2-F9 5'- GCCGCCTTAGCCGAAGCTTAGT -3' SULTR2-D 5'- GCTTCCTTCCTGGGAGTGCT -3' SULTR2-E 5'- AGGATGCGGCTCTACCCAAT -3' SULTR2-K 5'- CTGTGAGGCGAGGCGATAGATG -3'

SULTR2-G 5'- AGCAGGCAGTGGGATTGAAA -3' SULTR2-R5 5'- GCCCAGACCGATCTCCACACTGA -3'

SULTR2-I 5'- CGAAGTGCGGATAGGAGGAG -3' SULTR2-J 5'- TGCAGGGAACTGCACTGGTA -3'

SLT1-5’UTR-F1 5’-TGCTCACTTACATAGTCAGGCGCG-3’ SLT1-SEQ-F3 5’-AAGTCCAAGCAGGTCGACCA-3’ SLT1-SEQ-F5 5’-GCAAGACCAGTGACCTGATCG-3’

SLT1-SEQ-F7 5’-GCAGGAGCAGGTGGAC AAGA-3’ SLT1-SEQ-F10 5’-ACCTCCGTCGCCATCATGCT-3’ SLT1-3’UTR-R1 5’-GCTTCTGTTCACAGGATTACATTCAA-3’ SLT1-SEQ-R3 5’-TGGGTAGCAGGAAGTACAGCGC-3’

SLT1-SEQ-R4 5’-TGTCCTGGACCTTGAGCGGGAT-3’ SLT1-SEQ-R5 5’-TGTCAAGCAGCAGCACATCGCC-3’

SLT1-SEQ-R6 5’-TCCTGCAGACCCACGATGACCA-3’ P-SLT2-F1 5'- CGCTGCTGGAAAAGCATATGCAATTC -3'

SLT2-SEQ-F2 5'- CAACCTGGTCATCGTCGGTC -3' SLT2-F8 5'- GCGGGTACTGGACAGTTGGACACA -3' SLT2-F6 5'- TTGCCCCTATCACACAGGATGACAC -3'

SLT2-F2 5'- CAACCGGGTTGCAACTTCCTGAT -3' SLT2-R10 5'- GAAACCCGTTCCCCTGCTGCAGT -3'

SLT2-R8 5'- TGGTGTCCAGGATGAGCACGTC -3' SLT2-R13 5'- CTGTGTGATAGGGGCAACGACAAT -3'

SLT2-R12 5'- TTGAATTGCGGCAGATGGTGTAAC -3' SLT2-3’UTR-R2 5'- TCGGTCCGCGCAACTTCTTTGT -3'

143

Table 3-2. List of SULTR2-, SLT1- and SLT2-specific primers used for PCR screening of the insertion library.

144

2- Table 3-3. Characteristics of SO4 transport in wild-type cells, single, double and

2- triple SO4 transporter mutants after 24 h of S deprivation. K1/2 (in µM) is calculated from the initial rates using a Michaelis-Menten equation. Values are averages of 2-4 biological replicates with each experiment performed in duplicate. Error bars represent one standard deviation.

145 REFERENCES

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150

CHAPTER 4

ARS73a IS INVOLVED IN A TIERED REGULATION OF SULFUR

STARVATION RESPONSES

151 ABSTRACT

During sulfur (S) deprivation, the unicellular alga Chlamydomonas reinhardtii exhibits elevated expression of numerous S-deficiency responsive genes. These genes

2- encode proteins associated with sulfate (SO4 ) acquisition and assimilation, alterations in cellular metabolism and internal S recycling. Administration of the cytoplasmic translational inhibitor cycloheximide prevents the S starvation triggered accumulation of transcripts encoding arylsulfatases (ARS), the extracellular polypeptide ECP76, a light harvesting protein LHCBM9 and a haloperoxidase (PAP2); the induction of

2- genes encoding the high-affinity SO4 tranporters is not affected. These results suggest that there are two tiers of regulation associated with S deprivation responses; the first tier (primary response) is protein synthesis-independent while the second tier

(secondary response) requires de novo protein synthesis. A strain harboring an insertion of a drug resistant (paromomycin) cassette in a gene designated ARS73a, which encodes a putative transcriptional regulator, fails to show the protein synthesis- dependent increase in the ARS, ECP76, LHCBM9 and PAP2 transcripts in response to

2- S deprivation, although the increase in transcripts encoding the SO4 transporters is similar to that of wild-type cells. These results suggest that the ARS73a protein, which may function as a regulator of transcription, is responsible for up-regulation of those genes associated with the second tier of regulation. Furthermore, the ars73a strain is unable to synthesize the ARS polypeptide and exhibits low ARS activity in following exposure to S-limited condition. A genetic cross between ars73a and sac3 strains yielded a double mutant with a phenotype that is similar to that of ars73a strain (low

152 ARS activity), demonstrating that ARS73a is epistatic to SAC3. These results are critical for generating models to explain S-deprivation responses in Chlamydomonas.

153 INTRODUCTION

Sulfur (S) is an essential macronutrient that can be found in proteins, lipids, electron carriers and redox controllers (Leustek et al., 2000). Most organisms, including Chlamydomonas reinhardtii (Chlamydomonas throughout), have a limited capacity to store S and thus depend on a continual supply of an external source of this nutrient. The majority of S that is taken up from the soil solution by microbes and

2- 2- plants is the sulfate anion (SO4 ). Assimilation of SO4 involves reduction to sulfide and subsequent incorporation into the amino acids cysteine and methionine (Grossman and Takahashi, 2001).

The ability of microorganisms to acclimate to nutrient deprivation is critical to their survival in the natural environment. Chlamydomonas exhibits both general and specific responses when experiencing S deprivation. The general responses are common to a number of stress conditions and they include the cessation of cell division, accumulation of storage starch, and a decrease in photosynthetic rate. The specific responses are those that are associated with the deprivation of a single

2- nutrient. The specific S-deprivation responses include an elevated rate of SO4 uptake, synthesis of extracellular arylsulfatases (ARS), and an increased capacity to assimilate

2- SO4 by increasing levels of enzymes for cysteine biosynthesis (de Hostos et al.,

1988; Yildiz et al., 1994; Ravina et al., 2002).

ARS is secreted into the periplasmic space of Chlamydomonas cells where it

2- 2- hydrolyzes soluble SO4 esters in the medium, releasing free SO4 for uptake and assimilation. ARS activity is first detected 3 h after the transfer of cells into S- deficient medium (de Hostos et al., 1988). Identification and characterization of ARS

154 polypeptides led to the cloning and characterization of two genes, ARS1 and ARS2.

Transcripts from both of these genes accumulate in response to S deprivation (de

2- Hostos et al., 1989; Ravina et al., 2002; Zhang et al., 2004). The increase in SO4 uptake during S limitation of Chlamydomonas is a consequence of the synthesis a

2- new, high-affinity SO4 transport systems (Yildiz et al., 1994). Expression of genes

2- encoding three SO4 transporters, SULTR2, SLT1, and SLT2, are strongly upregulated at both the transcript and protein levels, almost immediately after the imposition of S

2- starvation. The initial rate of SO4 uptake increases as early as 1 h after removal of S from the medium and becomes maximum after approximately 6 h. An increase in the

2- affinity of the transport system for SO4 could also be detected within an hour of S

2- deprivation (Yildiz et al., 1994). Interestingly, S-starved cells exhibit increased SO4 uptake prior to the detection of ARS activity, suggesting that the controls of these two processes are differentially sensitive to the level of S in the environment.

Chlamydomonas also has mechanisms to conserve and recycle intracellular S during S-limiting conditions. Degradation of proteins and lipids that are not essential under S-deficient conditions can supply cells with a limited amount of S (Ferreira and

Teixeira, 1992). S-starved Chlamydomonas cells degrade most of the chloroplast sulfolipid to redistribute S for protein synthesis and other processes (Sugimoto et al.,

2007). Four prominent extracellular polypeptides, ECP56, ECP61, ECP76 and ECP88 are synthesized in response to S deprivation. The ECP genes are activated following the elimination of S from the medium and their mRNAs are rapidly degraded once an

S source is resupplied to S-deprived cells (Takahashi et al., 2001). Even though the function of these ECPs is not known, they exhibit features similar to those of cell wall,

155 hydroxyproline rich glycoproteins and the mature polypeptides have either one or no

S-containing amino acid. The cell wall proteins present during S-replete growth may be replaced by the ECP polypeptides and their S-containing amino acids can be redistributed during S deprivation (Takahashi et al., 2001).

S deprivation also triggers a potential change in the polypeptide composition of light harvesting complexes, favoring the synthesis of complexes containing polypeptides with that have few S amino acids. A gene encoding a specific light harvesting protein, LHCBM9, was strongly up-regulated under S-limited conditions while mRNAs encoding the LHCBM1-8 polypeptides exhibited a marked decline in their accumulation. Interestingly, LHCBM9 has a low content of S-containing amino acids. Because LHCBMs are very abundant proteins, substitution of S-rich LHCBM1-

8 with LHCBM9 would allow cells to conserve a significant amount of S (Gonzalez-

Ballester et al., 2010).

A number of S starvation-elicited responses appear to be controlled at the level

2- of gene expression. Genes encoding SO4 transporters, ARS, ECPs, LHCBM9, and

2- enzymes involved in SO4 assimilation [e.g. ATP sulfurylase, serine O-acetyl transferase, O-acetyl serine (thiol)lyase] are upregulated in cells deprived of S (de

Hostos et al., 1989; Yildiz et al., 1996; Takahashi et al., 2001; Ravina et al., 2002;

Zhang et al., 2004). Screens for mutants that are unable to acclimate properly to S limitation have led to an identification of three key regulators of S deprivation responses (Davies et al., 1994; Pollock et al., 2005).

The SAC1 (Sulfur Acclimation 1) gene encodes an integral membrane protein

+ 2- similar to a Na /SO4 cotransporter. SAC1 is a positive regulator critical for activation

156 of genes involved in scavenging and assimilating S from the environment. The sac1

2- mutant strain exhibit aberrant SO4 uptake and is unable to synthesize ARS, as well as other extracellular polypeptides in response to S deprivation. Induction of genes

2- associated with SO4 acquisition and assimilation was abolished in the sac1 mutant

(Davies et al., 1996; Ravina et al., 2002; Zhang et al., 2004). SAC1 also regulates several genes involved in the restructuring of the photosynthetic apparatus and the amelioration of damaging effects elicited by the accumulation of reactive oxygen species. The light sensitivity of the S-starved sac1 strain probably reflects it inability to downregulate photosynthetic electron transport (Davies and Grossman, 1994;

+ 2- Wykoff et al., 1998). The similarity between SAC1 and the Na /SO4 transporter raises the possibility that polypeptide whose original function was to bind and transport substrates into cells may have evolved into a regulatory element that senses extracellular nutrient concentrations. This is analogous to the sensing and regulation of glucose by yeast cells; Snf3 of yeast is a glucose sensor that has features of a high affinity glucose transporter (Ozcan et al., 1998).

The SAC3 (Sulfur Acclimation 3) gene, also designated as SNRK2.2 (Snf1- related protein kinase 2.2), encodes a serine/threonine kinase that acts as a negative regulator of S deficiency-responsive genes. The sac3 mutant exhibits a low constitutive level of ARS activity and expresses elevated basal levels of S deficiency responsive genes during S-replete growth (Davies and Grossman, 1994; Davies et al.,

1999; Ravina et al., 2002; Gonzalez-Ballester et al., 2008). SAC3 is also required for the proper downregulation of chloroplast transcriptional activity in S-starved cells

(Irihimovitch and Stern, 2006). There is no clear epitasis relationship between SAC1

157 and SAC3. The sac1sac3 double mutant has lower constitutive ARS activity than sac3 cells when grown under nutrient-replete conditions. However, in contrast to the sac1 single mutant, the double mutant exhibits partial up-regulation of both ARS enzymatic activity and transcript levels in response to S deficiency (Davies and Grossman, 1994).

2- Furthermore, based on transcript abundances, the genes encoding SO4 transporters,

ECP76, LHCBM9 and OASTL4 are induced to the same extent in the sac1sac3 double mutant as in the sac3 single mutant; SAC3 appears to be epistatic to SAC1.

However, there is a significant discrepancy between the accumulation of ARS transcripts and the level of ARS activity in the sac1sac3 double mutant. While the

ARS1 and ARS2 transcripts accumulated to the same extent in the sac3 and sac1sac3 strains, the double mutant displayed only 10% of the ARS activity associated with the single mutant (our unpublished data). These data along with results from Moseley et al. (2009) supports the hypothesis that SAC1 participates in controlling the activities of some S-responsive genes at the post-transcriptional level.

Another serine/threonine kinase, SNRK2.1, which is related to SAC3, has recently been shown to play a central role in controlling S deprivation responses. The snrk2.1 mutant fails to activate expression of most S-deprivation responsive genes and cannot maintain normal basal levels of expression of some genes during nutrient- replete condition. Furthermore, the snrk2.1 mutant is much more sensitive to light during S starvation compared to the sac1 mutant, suggesting that SNRK2.1 also plays an important role in modulating photosynthetic activities when the cells are deprived of S (Gonzalez-Ballester et al., 2008). The phenotypes of the snrk2.1sac3 double

158 mutant are indistinguishable from those of the snrk2.1, indicating that SNRK2.1 is epistatic to SAC3 (Gonzalez-Ballester et al., 2008).

2- During studies of SO4 transporter biogenesis in response to S starvation

(Chapter 3), I discovered that cycloheximide, an inhibitor of translation on 80S cytosolic ribosomes, blocked the induction of a subset of S-deficiency responsive genes, most of which are associated with S scavenging and S recycling. This finding suggested sequential temporal regulation of subsets of S-responsive genes, and that this tiered regulatory response was based on accumulation of a specific protein(s) synthesized during the early stages of S deprivation. In a previous screen for mutants defective in the synthesis of active ARS (Pollock et al., 2005), a specific mutant, ars73a, was identified that exhibited a pattern of regulation congruent with its involvement in the tiered regulatory responses; the ARS73a protein may be a transcription factor (or influence the activity of a regulatory factor).

RESULTS

Cycloheximide affects expression of some S-starvation responsive genes: During S

2- deprivation, several genes including those encoding high-affinity SO4 transporters,

2- ARS, enzymes in SO4 assimilation pathway are up-regulated in Chlamydomonas

(Figure 4-1A). As expected, the accumulation of those transcripts was prevented by the administration of a transcriptional inhibitor actinomycin D to the culture at the time of S removal (Figure 4-1B). Surprisingly, a subset of S-deprivation responsive genes was not induced in the presence of cycloheximide, which inhibits translation on

80S cytoplasmic ribosomes. The activation (increased transcript abundance) of ARS1

159 and ARS2 (encoding ARS) along with ECP76 (encoding a putative cell wall polypeptide), LHCBM9 (encoding a light harvesting protein), SBDP (encoding a selenobinding protein), and PAP2 (encoding a haloperoxidase) was abolished when the cytosolic protein synthesis was inhibited (Figure 4-1C). In contrast, the induction

2- of the high-affinity SO4 transporter genes (SULTR2, SLT1 and SLT2) and a putative amino acid transporter gene (AOT4) was not altered in the presence of cycloheximide.

These results suggest that there are at least two tiers of transcriptional regulation in response to S deprivation and that the second tier requires cytoplasmic protein synthesis.

Note that the induction of some genes, including OASTL4 (encoding O-acetyl serine (thiol)lyase) and TAU (encoding taurine dioxygenase), was only partially blocked by cycloheximide addition. Although cycloheximide did not impact the expression of SULTR2, SLT1 and SLT2, the accumulation of SULTR1 and SLT3 transcripts increased in the presence of cycloheximide (Figure 4-1C). This could be due to the general effect of translational inhibitor-mediated stabilization of the transcripts (possibly by sequestering them in ribosomal complexes). The mRNA level of the house keeping gene CBLP was shown to remain constant during S deprivation

(Chang et al., 2005), however, an increase in CBLP transcript accumulation was observed when cycloheximide was administered to cells upon removal of S from the medium (data not shown), supporting the argument that cycloheximide can affect mRNA stability. Finally, a translational inhibitor of 70S ribosome (in mitochondria and plastids), chloramphenicol, appeared to have no significant effect on expression of

S-deficiency responsive genes (Figure 4-1D).

160 Identification and preliminary characterization of the ars73a mutant: Proper acclimation to S deprivation can be monitored by assaying for extracellular ARS activity. Wild-type Chlamydomonas cells (cell wall minus strains) grown on solid medium with limiting S secrete the ARS enzyme into the agar. This activity can be

2- detected using the chromogenic substrate, 5-bromo-4-chloro-3-indolyl SO4 (XSO4); hydrolysis of this substrate by ARS results in the formation of a blue precipitate. This assay was employed in a screen to identify insertional mutant strains that cannot synthesize ARS following S starvation (Pollock et al., 2005). One mutant identified in this screen, designated ars73a, exhibited a very low level of ARS activity, as shown in

Figure 4-2.

To determine whether the ars (lacking ARS activity) phenotype co-segregates with the AphVIII gene (confers paromomycin resistance to the cells) that was introduced during the transformation, the ars73a mutant was crossed to a wild-type strain. Approximately two hundred random progeny were scored for both ARS activity on S-deficient medium and growth on paromomycin-containing medium; co- segregation of the mutant phenotype (low ARS activity) and paromomycin-resistance

(data not shown) was absolute, suggesting that the mutation was tightly linked to the

AphVIII insertion.

A Southern blot analysis shown in Figure 4-3 illustrated that the ars73a mutant contains a single insertion, in accord with the results from the genetic analysis that showed an approximate 1:1 ratio of paromomycin resistant and sensitive progeny

(data not shown). Adaptor-mediated PCR was used to identify one of the genomic regions flanking the AphVIII marker gene (Pollock et al., 2003). The diagram in

161 Figure 4-4A shows that the AphVIII gene integrated into the 6th intron of the ARS73a predicted gene model. The sequence of the translation product (1443 amino acids) predicted from the model does not have a signal peptide or any known motif/domain besides a glutamine-rich region (highlighted in Figure 4-4B), which is a common feature of transcriptional activators (Pabo and Sauer, 1992). Since the full-length cDNA has not been cloned (although partial ESTs are available), the predicted

ARS73a amino acid sequence may not be accurate. The level of ARS73a transcript does not appear to be regulated by the S status of the cells (Gonzales-Ballester and

Grossman, unpublished data). Interestingly, ARS73a is immediately downstream of

AOT4 gene, which is strongly upregulated during S deprivation (Gonzalez-Ballester et al., 2010) (Figure 4-1A). The accumulation of the AOT4 transcript was induced normally in the ars73a strain, suggesting that the mutation does not affect expression of AOT4 (Figure 4-5C). Moreover, a mutant harboring a lesion in AOT4 gene does not exhibit a low level of ARS activity (Gonzales-Ballester and Grossman, unpublished data).

Expression profile of S-deficiency responsive genes in ars73a mutant: To study the responses of the ars73a strain to S deprivation, levels of transcripts from several genes that are known to be controlled by the S status of the cells were analyzed. These genes

2- included SULTR1, SULTR2, SLT1, SLT2, SLT3 for the SO4 transporters, ARS1 and

ARS2 for ARS, ECP76 for an extracellular polypeptide that is probably associated with cell wall, LHCBM9 for a light harvesting protein, SBDP for a putative selenobinding protein, PAP2 for a haloperoxidase, OASTL4 for O-acetylserine

162 2- (thiol)lyase, an enzyme important for SO4 assimilation, and AOT4 for a putative amino acid transporter. As expected from the mutant phenotype, ARS1 and ARS2 transcripts were not up-regulated to the same extent as in wild-type cells (a 10-fold increase in ars73a strain compared to a 1000-fold increase in wild-type strain). The magnitude of ECP76, LHCBM9, PAP2 and SBDP induction was also markedly diminished in the ars73a mutant (Figure 4-5C).

2- Surprisingly, expression of three high-affinity SO4 transporters (SULTR2,

SLT1, SLT2) and the putative amino acid transporter AOT4 was completely unaffected by the lesion in ARS73a (Figure 4-5C). Even more astonishing, the pattern of S- deprivation responsive transcript accumulation in the ars73a strain closely resembled that of wild-type cells treated with cycloheximide (compare Figure 4-5B and Figure

4-5C). Taken together, these results suggest that: 1) there are two tiers of S- deprivation responses, one independent of (first tier) and one requiring (second tier) protein synthesis, and that 2) the synthesis of the ARS73a protein is required for activating expression of genes in the second tier.

In accord with measurements of their transcript abundances, SULTR2, SLT1, and SLT2 polypeptides accumulated to approximately the same extent in both wild- type and the ars73a strains during S deprivation (Figure 4-6). In contrast, while the amount of ARS protein in S-starved wild-type markedly increased, the ars73a mutant could not accumulate the ARS polypeptide under the same conditions (Figure 4-6), resulting in very low ARS activity in –S medium (Figure 4-2).

Furthermore, ARS73a appears to be involved in down-regulating the putative

2- low-affinity SO4 transporters SULTR1 and SLT3 under S-limited conditions; their

163 transcript levels did not decline in the S-starved ars73a mutant (Figure 4-5C). In both wild-type and ars73a cells, SULTR1 and SLT3 mRNA abundances increased in the presence of cycloheximide (Figure 4-5), which could be due to a nonselective transcript-stabilizing effect of cycloheximide. For some S-deficiency induced genes

(e.g. ARS1, ARS2, SBDP, LHCBM9), the addition of cycloheximide to the ars73a mutant further reduced their mRNA abundances during S starvation (compare Figure

4-5C and Figure 4-5D), implying that another unknown protein may play some role in activating those –S-induced genes and that this protein must also be synthesized de novo.

Epistasis analysis of ARS73a and SAC3: To investigate the relationship between the

ARS73a and SAC3 genes, the ars73a mutant (in 21gr genetic background) was crossed to the sac3 mutant in the same genetic background. As mentioned earlier, SAC3 encodes a serine/threonine kinase that acts as a negative regulator, repressing expression of S-deprivation responsive genes under nutrient-replete conditions

(Davies et al., 1999). An assay for ARS activity was used to assess the acclimation response of wild-type, sac3, ars73a, and sac3ars73a mutants to S starvation. While wild-type and ars73a cells displayed no ARS activity on S-replete medium, the sac3 strain exhibited low level of ARS activity (Davies et al., 1994; Davies et al., 1999)

(Figure 4-7A and Figure 4-7C). The ARS phenotype of the sac3ars73a double mutant is similar to that of the ars73a single mutant under nutrient-replete and nutrient-depleted conditions. Both wild-type and sac3 strains accumulated ARS in S- deficient medium whereas the ars73a and sac3ars73a strains had almost no detectable

164 ARS activity (Figure 4-7B and Figure 4-7C). The identical ARS activity phenotypes of ars73a and sac3ars73a strains in both +S and –S medium support the conclusion that ARS73a is epistatic to SAC3.

DISCUSSION

Chlamydomonas exhibits a suite of responses to S deprivation and several of

2- those responses are regulated at a transcriptional level. Genes encoding SO4

2- transporters, enzymes participating in SO4 assimilation, extracellular ARS, and proteins involved in S recycling (e.g. ECP76, LHCBM9) are induced when cells become S limited (Ravina et al., 2002; Zhang et al., 2004; Gonzalez-Ballester et al.,

2010). The transcriptional regulation of these genes appears to be staged. The first tier responses take place within an hour of the onset of S deprivation and do not require de

2- novo protein synthesis. They include the up-regulation of the high-affinity SO4 transporters (SULTR2, SLT1, SLT2) and the putative amino acid transporter AOT4

2- (Figure 4-1). The high-affinity transport systems facilitate efficient uptake of SO4 at low extracellular concentrations. Even though only arginine transport activity has been demonstrated in Chlamydomonas (Kirk and Kirk, 1978), it has recently been shown that cells can grow slowly with methionine or cysteine as a sole S source, raising the possibility that AOT4 transporter could participate in the uptake of S-containing amino acids from the environment.

The second tier responses are dependent on protein synthesis. The induction of

ARS1, ARS2, ECP76, LHCBM9, SBDP, and PAP2 was blocked in S-starved cells in the presence of cycloheximide (Figure 4-1C). While the initial phase of S deficiency

165 responses focuses on the synthesis of more efficient, higher-affinity transport systems, most of the genes controlled by second tier regulation are involved in scavenging extracellular S and redistributing and recycling intracellular S. These processes take place when the S supply is severely limited and the cells have to optimize S utilization in order to remain viable. The expression of second tier genes is likely controlled by a protein that is synthesized as a part of the first tier response.

Interestingly, during S deprivation, ars73a exhibited a similar pattern of gene expression as wild-type cells treated with cycloheximide (Figure 4-5B and Figure 4-

5C). This result implies that de novo synthesis of the ARS73a protein in a S-limited environment is crucial for expression of genes in the second tier. The sequence flanking the AphVIII insertion in the ars73a strain was identified and the marker was integrated in the 6th intron of the predicted model of ARS73a. Although the genetic analyses shows co-segregation between paromomycin-resistance (conferred by the

AphVIII marker) and the ars phenotype, it is essential to show that introduction of the

ARS73a sequence into the mutant strain rescues the ars phenotype.

The first S-deficiency response regulator recently identified in Arabidopsis is

2- an EIL family transcription factor, SLIM1. It controls the activation of SO4 acquisition and assimilation as well as the degradation of glucosinolates under S deprivation conditions (Maruyama-Nakashita et al., 2006). A major regulator of phosphorus (P)-starvation responses in Chlamydomonas, PSR1, is also a transcriptional activator controlling numerous genes involved in acclimation of cells to low levels of P in the environment (Wykoff et al., 1999). A few regulators of the S- starvation responses have been identified in Chlamydomonas (e.g. SAC1, SNRK2.1,

166 and SAC3), and they appear to control the responses primarily at the transcriptional level (Davies et al., 1994; Davies et al., 1996; Davies et al., 1999; Zhang et al., 2004;

Gonzalez-Ballester et al., 2008). So far, no transcription factor that plays a specific role in regulating expression of S-deprivation-responsive genes has been identified.

Although ARS73a does not possess a DNA-binding domain or contain a nuclear localization signal, it has a glutamine-rich region, which is often associated with transcription factors (Pabo and Sauer, 1992). The phenotype of the ars73a strain also suggests that this polypeptide functions as a transcriptional activator (or co- activator) of the second tier responses. De novo synthesis of ARS73a (or a protein that activates ARS73a) may be a part of the first tier response since prevention of protein synthesis (by cycloheximide) in S-starved wild-type cells blocked the induction of genes in the second tier. The accumulation of the ARS73a transcript is not regulated by the S status of the cells, raising the possibility that post-transcriptional control is required for the production of active ARS73a. Translation of the ARS73a polypeptide may be induced by S limitation. Alternatively, the ARS73a protein level may remain unchanged but its activity may be modulated by an unidentified protein (for example, a kinase) that is synthesized de novo following S deprivation.

It is worth noting that the addition of cycloheximide to S-deprived ars73a mutant cells results in a further repression of some genes of the second tier (e.g. ARS1,

ARS2, LHCBM9) (Figure 4-5C and Figure 4-5D). This result indicates that there may be other unknown proteins that control expression of second tier genes. In addition to its function in the activation of genes, ARS73a also participates in the down-regulation of SULTR1 and SLT3 transcript levels when the cells are S-limited. Multiple roles of

167 other regulatory elements have been noted. Yeast Rap1p protein has been shown to function as both a transcriptional activator and repressor depending on the sequence context (Drazinic et al., 1996), and the adenovirus E1A transcriptional repressor can function as an activator when tethered to a promoter (Bondesson et al., 1994).

To study the relationship between ARS73a and other genes that encode regulators of the S-deficiency acclimation pathway, I crossed the ars73a strain to the sac3 mutant. The double mutant, sac3ars73a, exhibits an identical phenotype to that of ars73a (Figure 4-7), demonstrating that SAC3 and ARS73a act in the same pathway and that ARS73a is epistatic to SAC3. The model presented in Figure 4-8 illustrates the regulation of S-deficiency responses. During nutrient-replete growth, the

2- sensor SAC1, which resides on the plasma membrane, binds SO4 and remains inactive. The negative regulator, SAC3, represses expression of the S-starvation responsive genes. Upon sensing S limitation, SAC1 deactivates SAC3 and at the same time the serine/threonine kinase SNRK2.1 becomes active (Moseley et al., 2009).

Activation of transcription is initiated by SNRK2.1, which phosphorylates other proteins and possibly a transcription factor (either directly or indirectly) that activates expression of genes associated with the first tier (SULTR2, SLT1, SLT2 among other genes) of S-deprivation responses. During this early stage in the S deprivation

2- program, the synthesis of high-affinity SO4 transporters facilitates the efficient

2- uptake of SO4 into cells. Also, as a part of this first tier response, the putative transcription factor ARS73a is either synthesized de novo or activated by an unknown protein designated Y (which is synthesized during the first tier response).

Subsequently, ARS73a, either through direct or indirect transcriptional activation,

168 elicits expression of several genes associated with the second tier response, most of which encode proteins involved in scavenging S from the environment and recycling and redistributing internal S. Evolving a staged response would allow for a finer tuning of the system, with each metabolic and physiological change occurring as the need for such change becomes imperative.

Studies of the interaction between regulators of P and S deprivation responses has provided support for a model in which SAC1 inhibits SAC3 (rather than activating

SNRK2.1) in S-deprived cells. Chlamydomonas PSR1 gene is required for proper acclimation to P starvation. PSR1 controls expression of genes encoding proteins that function in P scavenging which includes extracellular phosphatases, high-affinity phosphate transporters, and proteins involved in mobilizing internally stored polyphosphate (Shimogawara et al., 1999; Moseley et al., 2006). In addition to its inability to synthesize extracellular phosphatase or induce high-affinity phosphate transporter in a P-limited environment, the P-starved psr1 mutant also activates a number of S-deprivation responsive genes (this is not observed in P-starved wild-type cells). Interestingly, this aberrant activation of S-starvation responses in P-deficient psr1 cells does not require an input from SAC1 (Moseley et al., 2009). These data are consistent with the model in which SAC1 inactivates SAC3 kinase when S is limiting in the environment.

FUTURE DIRECTIONS

A large number of genes exhibit changes in their expression level during S deprivation (Zhang et al., 2004; Gonzalez-Ballester et al., 2010), and only a small

169 subset of those S-starvation responsive genes was investigated in this work. A genome-wide transcript analysis (RNA-seq technology) should be employed to define each tier of the regulation. Treatment with protein synthesis inhibitor and the use of the ars73a mutant will help reveal the hierarchy of S-deprivation responsive genes.

Once the genes are categorized into different tiers, the promoter regions of genes from each grouping could be studied in detail. Genes in the same tiers are likely to be regulated by the same transcription factors and cis acting elements associated with the specific S-responsive sets of genes could be identified using bioinformatic approaches.

Since ARS73a has certain features of a transcriptional activator, it would be interesting to clone the gene and over-express the protein for DNA-binding studies.

ARS73a seems to function as both a transcriptional activator and repressor in S- deprived cells. Moreover, the predicted polypeptide does not have any known domains or motifs except a glutamine-rich region. It will be informative to identify regions of the ARS73a protein that is responsible for gene activation and repression.

170 MATERIALS AND METHODS

Strains and growth conditions: The following strains were used in this work: 21gr

(available from the Chlamydomonas Center), ars73a (nit2-, cw15, mt+) (Pollock et al.,

2005), sac3 (Davies et al., 1994; Davies et al., 1999). Cells were cultured in either S- replete (+S) or S-depleted (-S) Tris-acetate-phosphate (TAP) medium under continuous illumination (80 µmol photon m-2 s-1) on a rotating platform (200 rpm) at

25°C. TAP–S medium was prepared as described previously (Davies et al., 1994). For

S starvation experiments, cells were grown to mid-logarithmic phase (2-4 x 106 cells mL-1) in TAP+S medium, washed once with TAP-S medium (2500 X g for 5 min) and resuspended in TAP-S to the original cell density.

Southern blot analyses: Genomic DNA was isolated from 100 mL liquid cultures of the wild-type strain 21gr and the ars73a mutant using a standard phenol-chloroform extraction protocol (Sambrook et al., 1989). Approximately 10 µg of genomic DNA was digested overnight with 10 u of restriction endonucleases (PstI; New England

Biolabs). The fragments were separated by agarose (0.8%) gel electrophoresis, blotted overnight in 20X SSC onto nitrocellulose membranes (GeneScreen, DuPont-New

England Nuclear), and the transferred DNA cross-linked to the membrane by UV illumination. An alkaline phosphatase-labeled probe was synthesized by chemical cross-linking of a thermostable alkaline phosphatase to the nucleic acid template.

Probe synthesis and hybridization were performed using the Amersham AlkPhos

171 DirectTM Labeling and Detection Systems following the manufacturer’s protocol

(Amersham Biosciences, Sweden).

RNA isolation and quantification: Total RNA was extracted from frozen cell pellets using the RNeasy Mini Kit (Qiagen, Valencia, CA) and treated with RNase-free