STUDIES OF CELLULAR QUIESCENCE IN PHOTOSYNTHETIC EUKARYOTES USING
CHLAMYDOMONAS REINHARDTII AS A REFERENCE MODEL
By
Chia-Hong Tsai

A DISSERTATION
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of
Plant Biology—Doctor of Philosophy
2014

ABSTRACT
STUDIES OF CELLULAR QUIESCENCE IN PHOTOSYNTHETIC EUKARYOTES USING
CHLAMYDOMONAS REINHARDTII AS A REFERENCE MODEL
By
Chia-Hong Tsai
Cellular quiescence, defined as reversible cell cycle arrest, is a fundamental mechanism that
organisms use to maintain tissue homeostasis or to overcome adverse conditions. In fact, most
cells spend the majority of their life span in quiescence. Despite the importance, remarkably little
is known about the regulation of quiescence. For photosynthetic organisms, it requires additional
effort to maintain the state of quiescence, that is, to halt the photosynthetic machinery in a way
that it can restart momentarily when conditions improve. Thus far, how plant cells exit
quiescence, and regain competence to divide is entirely unknown.
The unicellular green alga Chlamydomonas reinhardtii was chosen as a reference model
to study quiescence in photosynthetic eukaryotes for several reasons: First, quiescence and cell
division can be discretely defined and controlled by manipulating nutrient availability. Second,
microalgae tend to accumulate triacylglycerols only under the growth-limiting, quiescent state,
which has long hampered efforts toward the efficient generation of biofuel feedstocks from
microalgae. Mechanistic insights into the cellular quiescence hold great potential to uncouple
this inverse relationship.
In this dissertation, I present the discovery of the protein Compromised Hydrolysis of
Triacylglycerols 7 (CHT7), a repressor of cellular quiescence in Chlamydomonas that ensures
the reversibility of quiescence. Moreover, I conducted comparative transcriptomics using the
cht7 mutant, which is unable to orderly progress from quiescence, to uncover the CHT7 regulon
and transcriptional programs unique to the exit of quiescence. In addition, I found that lipid

droplets coordinate lipid and protein dynamics during quiescence and are important for the
timeliness of quiescence exit. Overall, my findings make substantial progress towards the
comprehensive understanding of cellular quiescence in photosynthetic cells, and promises to
provide important insights into the regulation of cellular behavior in multicellular organisms as
well.

TABLE OF CONTENTS

LIST OF TABLES……………………………………………………………………………...viii
LIST OF FIGURES…………………….………………………………………………………..ix
CHAPTER 1 The Essence of Cellular Quiescence: a Literature Review…………………………1
Introduction to quiescence…………………………………….................................................2
Mechanistic insights into cellular quiescence………………....................................................3
Signaling cascades that regulate the entry into quiescence………………...............................5
Microbes accumulate reducing compounds during quiescence……………….........................7
Chlamydomonas reinhardtii as a reference model to study cellular quiescence…………….10
Aims of the dissertation research………………………………………….............................11
REFERENCES………………………………………………………………………………13
CHAPTER 2 Protein Compromised Hydrolysis of Triacylglycerols 7 (CHT7) Acts as a
Repressor of Cellular Quiescence in Chlamydomonas…………………………………………..21
ABSTRACT………………………………………………………………………………….22
INTRODUCTION……………………………………..……………………….……………23
MATERIALS AND METHODS…………………………………………………………….25
Strains, Genetic Analysis, and Growth Conditions……………………………………...25
Generation of Antibodies, Immunoblotting, Subcellular Fractionation, and BN-PAGE..26
Mutant Screen……………………………………………………………………………28
Confocal Microscopy and Construction of the CHT7-GFP Fusion……………………...29
Phenotyping Assays……………………………………………………………………...30
Standard DNA and RNA Procedures…………………………………………………….32
Illumina RNA Sequencing and Bioinformatics………………………………………….37
RESULTS..…………………………………………………………………………………..39
Isolation of compromised hydrolysis of triacylglycerols Mutants………………………39
CHT7 Encodes a CXC Domain DNA Binding Protein Present in the Nucleus…………41
Absence of CHT7 Affects the Exit from Quiescence but Not Cell Viability……………46
Absence of CHT7 Partially Derepresses Quiescence-Associated Transcriptional
Programs…………………………………………………………………………………51
CHT7 Levels Remain Constant in Response to the N Supply, and CHT7 Is in a Large
Complex………………………………………………………………………………….55
DISCUSSION……………………………………………………………………..…………58
REFERENCES………………………………………………………………………………61
CHAPTER 3 Comparative Transcriptomic Analysis of Nitrogen Resupply-induced
Modifications in a Chlamydomonas Quiescence-exit Mutant…………………………………...71
ABSTRACT…………………………………………………………….................................72
INTRODUCTION………………………………………………...…………………………73
MATERIALS AND METHODS…………………………………………………………….76
Strains and Growth Conditions…………………………………………………………..76
iv

Illumina RNA Sequencing and Bioinformatics………………………………………….76
Metabolite Measurements………………………………………………………………..76
Fatty Acid and Lipid Analysis…………………………………………………………...77
Standard RNA Techniques………………………………………………………………78
RESULTS…………………………………………………………………...……………….79
Transcriptomes of Parental Line during different N regimes……………………………79
Comparative Transcriptomics of cht7 and the Parental Line following N-resupply…….82
Metabolite Analyses Confirm the Effect of Transcriptional Changes on Lipid Metabolism
……………………………………………………………………………………………86
Two Classes of Lipases Affect TAG Accumulation in opposite ways…………………..91
The cAMP-Dependent Protein Kinase A Pathway Is Involved in Quiescence Exit……..93
DISCUSSION………………………………………………………………………………..97
The CHT7 regulon and possible influence of cAMP-PKA signaling……………………98
REFERENCES……………………………………………………………………………100
CHAPTER 4 The Role of Lipid Droplet in Coordinating Lipid Dynamics to Ensure the
Reversibility of Cellular Quiescence in Chlamydomonas……………………………………...106
ABSTRACT……………………………………………………………...............................107
INTRODUCTION………………………………………………………………………….108
MATERIALS AND METHODS…………………………………………………………...110
Strains and Growth Conditions…………………………………………………………110
Fatty Acid and Lipid Analysis………………………………………………………….110
Lipid Droplet Isolation, HDN-PAGE, and Co-immunoprecipitation…………………..110
Immunofluorescence, Confocal, and Transmission Electronic Microscopy…………...114
RESULTS…………………………………………………………………………………..116
Transmission Electronic Microscopy Reveals Two Stages of LD Mobilization……….116
MLDP Recruits Different Sets of Proteins to LDs……………………………………..119
MLDP Is Indispensible for α-tubulin Association with LDs…………………………...122
The Surface of Lipid Droplet Has Unique Lipid and Fatty Acid Composition………...124
Proper Surface Area of Lipid Droplet Ensures Orderly Progression out of Quiescence.126
DISCUSSION………………………………………………………………………………129
The role of lipid droplet in cellular quiescence………………………………………...129
A new model of lipid droplet formation………………………………………………..131
REFERENCES……………………………………………………………………………..135
CHAPTER 5 Conclusions and Perspectives……………………………………………………139
Regulatory processes by which CHT7 maintains the reversibility of quiescence…….........140
Potential modification of CHT7 activity during the entry into quiescence………………...141
The regulatory logic underlying the CHT7 and Rb complexes to enable transitions between
quiescence and cell division………………………………………………………………142
Functional interplay between CHT7 and other quiescence regulators……………………143
Lipid droplets are important for the survival during quiescence and for the quiescence exit
………………………………………………………………………………………………144
Biotechnological applications of cellular quiescence and lipid droplet biology…………145
REFERENCES……………………………………………………………………………..146

v

LIST OF TABLES

Table 2.1. Oligonucleotide primers used in this study…………………………………………...33

vi

LIST OF FIGURES

Figure 1.1. Rb and cell cycle machinery...………………………………………...........................4

Figure 1.2. Suppression of irreversible routes…………………………………………………….5

Figure 1.3. Chlamydomonas life cycles………………………………………………………….11

Figure 2.1. Isolation of cht mutants…………………………………………...............................40

Figure 2.2. Phenotypes of cht7…………………………………………......................................41

Figure 2.3. Meiotic progeny phenotyping and cht7 complementation…………………………..43

Figure 2.4. CHT7 is a CXC domain-containing protein present in the nucleus…………………44

Figure 2.5. Nuclear localization of CHT7……………………………………………………….45

Figure 2.6. Growth of cht7 affected by different treatments…………………………………….47

Figure 2.7. Growth, RNA, and protein of cht7 affected by different treatments………………...48

Figure 2.8. Viability and colony formation of cht7 during N deprivation……………………….50

Figure 2.9. Global gene expression comparison of N-replete and -deprived cells of PL dw15 and
cht7……………………………………………………………………………………………….53

Figure 2.10. Confirmation of transcriptional changes by qPCR and analysis of photosynthesis
and flagellum-related genes…………………………………………...........................................54

Figure 2.11. Abundance of CHT7 and hypothesis for CHT7 function…………………………..56

vii

Figure 2.12. Immunodetection of CHT7 and 2D BN-SDS-PAGE………………………………56

Figure 3.1. Cell Cycle and Proposed Functions for CHT7 in Quiescence………………………75

Figure 3.2. Experimental Design of RNA-Seq Experiments…………………………………….79

Figure 3.3. Summary Scheme of Transcriptomic Analyses in the Parental Line………………..80

Figure 3.4. Comparative Transcriptomics during Quiescence Exit……………………………...83

Figure 3.5. Changes in Gene Expression and Metabolite of Tetrapyrrole Pathway and
Peroxisomal Redox Homeostasis………………………………………………………………...85

Figure 3.6. Confirmation of Transcript Changes at Quiescence Exit……………………………86

Figure 3.7. Changes in Gene Expression and Metabolite Levels Related to MGDG and Fatty
Acid Syntheses…………………………………………………………………………………...88

Figure 3.8. Detailed Lipid Analysis at Quiescence Exit…………………………………………89
Figure 3.9. Changes in Gene Expression of Putative Lipases and β-oxidation………………...93
Figure 3.10. Possible Role o cAMP-PKA Signaling in Quiescence Exit and Transcript Profiles of
Related Genes…………………………………………………………………………………..95

Figure 3.11. Fatty Acid Profiles of PKI-Treated Samples……………………………………...96

Figure 4.1. Ultrastructure of Cells during Quiescence Exit…………………………………….117

Figure 4.2. Ultrastructure of Cells during Quiescence Exit…………………………………….118

Figure 4.3. Ultrastructure of Cells during Quiescence Exit…………………………………….119
viii

Figure 4.4. Lipid Droplet Localization and MLDP Complex Formation………………………122

Figure 4.5. Lipid Droplet Protein Quantification in the MLDP RNAi Lines…………………124

Figure 4.6. Lipid Profiles of Lipid Droplet Surface Membrane………………………………..126

Figure 4.7. Reduction of MLDP Affects Quiescence Exit, TAG Turnover, and Lipid Droplet
Homeostasis…………………………………………………………………………………….128

Figure 4.8. A model for lipid droplet-specialized deacylation/reacylation cycle……………..131

Figure 4.9. A model for lipid droplet formation and deformation……………….……………..133

Figure 5.1. Summary of pathways thought to control transitions between growth/division and
quiescence in Chlamydomonas reinhardtii……………………………………………………..144

ix

CHAPTER 1
The Essence of Cellular Quiescence: a Literature Review

1

Introduction to quiescence
The eukaryotic cell cycle is divided into different phases: G1, characterized by cell growth; S,
during which synthesis of DNA occurs; G2, when cells prepare for division; and M, which is
defined by mitosis. Other types of cell cycle control, especially the ability to temporarily
withdraw from cell division, are essential for survival during adverse conditions, particularly in
unicellular organisms, and for the maintenance of tissue homeostasis in multicellular organisms.
This non-dividing state is called quiescence, and is distinguished from senescence or terminal
differentiation by its reversibility (Gray et al., 2004; Valcourt et al., 2012). Whether prokaryotic
or eukaryotic, unicellular or multicellular, virtually all cells are capable of entering quiescence in
order to ensure the continuity of life. Nevertheless, the signals that promote quiescence vary
across different cell types. For example, bacteria and yeast enter stationary phase when their
carbon source is exhausted or in response to the deprivation of specific nutrients, e.g. nitrogen
(N), sulfur, or phosphate (Thevelein et al., 2000). In mammals, quiescence is seldom induced by
starvation because to maintain homeostasis cells are continuously supplied with glucose, amino
acids and other nutrients provided by nearby capillaries (Longo, 1999). Cells such as fibroblasts,
lymphocytes, and stem cells, typically quiesce, unless they are exposed to proliferative signaling
molecules or situational cues (Valcourt et al., 2012). Quiescence also occurs in the context of
development. In the Arabidopsis root meristem, stem cells surround a small group of organizing
cells, referred to as the quiescent center (Wildwater et al., 2005). Despite these differences, many
quiescent responses appear to be universal, including the changes in transcriptional and
translational regulations, in metabolic states, as well as in other physiological aspects (Gray et al.,
2004; Wu et al., 2004; Coller et al., 2006; Miller et al., 2010; Valcourt et al., 2012; Gifford et al.,
2013). Quiescence is relevant to many fields of study ranging from stem cell and cancer research

2

to stress biology in photosynthetic organisms. A more in-depth understanding of the conserved
mechanisms underlying the entry into, maintenance of, and exit from quiescence promises to
provide important insights into the developments of anticancer therapies and the optimization of
algal biofuel feedstocks. This chapter will summarize current knowledge about quiescence, with
a special focus on unicellular organism as it is more closely related to my work on
Chlamydomonas reinhardtii.

Mechanistic insights into cellular quiescence
Cell cycle arrest and reversibility comprises cellular quiescence. On the molecular level, cell
cycle arrest is established through a complex interplay between proteins. The human Rb tumor
suppressor protein and its homologs, also referred to as pocket proteins, are components of
multiprotein transcriptional complexes in most eukaryotic organisms with fundamental roles in
regulating cell cycle progression (Figure 1.1) (Knudsen and Knudsen, 2006; Burkhart and Sage,
2008). Briefly, Rb interacts with DNA-binding proteins such as E2F and DP to repress gene
expressions required for cell cycle progression. Acute mutation of Rb gene function in mouse
embryo fibroblasts is sufficient to reverse cell cycle arrest (Sage et al., 2003). The transition from
quiescence is stimulated by mitogenic signals, following through the sequestration of cyclindependent kinase (CDK) inhibitors p21 and p27, and the activation of CDK4 and CDK6,
allowing consecutive phosphorylation of Rb. The hyperphosphorylated Rb dissociates from E2F
transcription factors and releases them to activate respective genes (DeGregori et al., 1995).
Although yeast cells are lacking in Rb and E2F proteins (Ohtani and Nevins, 1994), they employ
a similar regulatory process. A cell size regulator WHI5 inhibits the G1/S transcription by
binding to the transcription factors SBF/MBF on the cis-regulatory elements. This inhibition can

3

be reversed by CDK-dependent phosphorylation of WHI5 when the CDK inhibitor SIC1 is
withdrawn (Costanzo et al., 2004). Deletion of WHI5 promotes cell cycle progression without
the upstream activation of SBF/MBF.

Figure 1.1. Rb and cell cycle machinery. Rb and Rb-p represent the unphosphorylated and the
phosphorylated forms of the retinoblastoma protein. In G0 and early G1, Rb physically
associates with E2F factors and blocks their transactivation domain. In late G1, Rb-p releases
E2F, allowing the expression of genes that encode products necessary for S-phase progression
(Giacinti and Giordano, 2006).

Reversibility is the defining nature of cellular quiescence. In contrast to other arrested
states such as senescence, apoptosis, and terminal differentiation, only in quiescence this block to
proliferation can be reversed (Coller, 2011). One mechanism by which the reversibility is insured
is the suppression of irreversible cell fates. Transcriptional profiling of quiescent human
fibroblasts uncovered the regulation of genes that inhibit apoptosis or cellular differentiations
(Coller et al., 2006). Later on, it became known that a transcriptional repressor HES1 blocks the
alternative routes to premature senescence and myogenic differentiation in quiescent fibroblasts
by modifying histone tails and thus affecting chromatin conformation (Sang et al., 2008; Sang
4

and Coller, 2009). In some cancers, the HES1 pathway is hijacked, thus allowing cancer cells to
escape from irreversible cell cycle arrest and undergo tumorigenesis (Figure 1.2).

Figure 1.2. Suppression of irreversible routes. Irreversible routes to senescence and terminal
differentiation are suppressed by HES1 in quiescent cell. (Modified from Cheung and Rando,
2013).

Signaling cascades that regulate the entry into quiescence
The immunosuppressant rapamycin has provided a great tool in the discovery of the mechanisms
that control the entry into quiescence. Treatment of rapamycin inhibits growth and division in
both yeast and mammalian cells. Rapamycin-treated cells acquire many features characteristic of
quiescence: G1 phase DNA content, repression of ribosomal protein genes and induction of
genes encoding protein chaperones, reduced rates of protein synthesis, and high levels of
autophagy (Barbet et al., 1996; Noda and Ohsumi, 1998). In addition, rapamycin-treated yeast
accumulates storage compounds such as glycogen and trehalose. The target of rapamycin,
namely TOR, and its directly and indirectly interacting proteins translate extracellular nutrient
signals into information that helps decide the extent of cell growth (Rohde and Cardenas, 2004).
When nutrients are abundant, TOR kinases, in association with its cooperating enzymes,

5

phosphorylate downstream targets that enhance protein synthesis and cell growth. When the
cellular AMP/ATP ratio is high or amino acids are scarce, the TOR pathway is suppressed, thus,
allowing the activation of autophagy to reclaim metabolites, to spare ATP, and to eliminate
degraded proteins and organelles. In yeast, there are two TOR kinase proteins, TOR1 and TOR2,
but only the TOR1 complex is sensitive to rapamycin. Disruption of the TOR1 complex mimics
rapamycin treatment and a quiescence-like state (Loewith et al., 2002). Therefore, the TOR
pathway has generally been considered to be a negative regulator of the transition into
quiescence. The TOR kinases were first identified in Saccharomyces cerevisiae, and are highly
conserved in animals and plants (Zhang et al., 2000; Russell et al., 2011; Xiong et al., 2013).
Components of the TOR pathway have also been identified in Chlamydomonas by screening
mutants insensitive to rapamycin as well as by in silico studies (Perez-Perez et al., 2010).
Intriguingly, the growth of Chlamydomonas cells is not completely abolished by rapamycin as is
the case for yeast. Even using a saturated concentration of rapamycin, Chlamydomonas cells can
still grow albeit they take twice as long to reach the same density.
The cyclic AMP (cAMP)-dependent protein kinase A (PKA) pathway is well-studied in
yeast, and in spite of some unanswered questions, it is presumably another element that prevents
the entry into quiescence. The enzyme adenylyl cyclase catalyzes the conversion of ATP into
cAMP and pyrophosphate. When cAMP concentration increases, for example in the presence of
high glucose, cAMP binds to the inhibitory domains of PKA which results in the dissociation
and activation of the catalytic domains. Yeast cells deficient in basal cAMP synthesis gradually
enter the quiescent state; this can be rescued by providing external cAMP (Boutelet et al., 1985;
van Aelst et al., 1991). Subcellular localization of PKA in yeast is under nutritional control, as it
shuttles from nucleus to cytoplasm when entering quiescence (Griffioen et al., 2000). The

6

nuclear targets of PKA are the nutrient-regulated transcription factors MSN2 and MSN4 (Smith
et al., 1998; Beck and Hall, 1999). Interestingly, PKA activity is dispensable in the msn2 msn4
double mutant, suggesting that in proliferative cells,

MSN2 and MSN4 counteract PKA-

dependent growth by stimulating gene expression that inhibits growth, consistent with the fact
that MSN2/MSN4 activity is required for expression of YAK1, previously shown to antagonize
PKA-dependent growth (Garrett and Broach, 1989). MSN2 and MSN4 are also targets of the
TOR pathway (Beck and Hall, 1999), providing evidence that signaling pathways thought to
prevent entry into quiescence comprise an interconnecting network.
Activators of quiescence are inherently required for the establishment of viable quiescent
cells; without them, cells frequently die rapidly after the transition. The yeast protein kinase C
encoded by PKC1 is one such regulator. The pkc1 mutant undergoes cell lysis on carbon or N
deprivation due to a deficiency in cell wall construction—a unique feature of quiescent yeast—
and thus is vulnerable to osmotic changes (Levin and Bartlett-Heubusch, 1992). A mitogenactivated protein (MAP) kinase cascade involving the MAP kinase MPK1 is thought to mediate
the signaling by PKC1. Mutants defective in MPK1, and its targets MKK1, MKK2 (MAP kinase
kinase), and BCK1 (encoding MAP kinase kinase kinase) all show a similar phenotype as pkc1
(Lee and Levin, 1992; Irie et al., 1993; Lee et al., 1993). The PKC1 pathway is transiently
activated by the inhibition of TOR to remodel the cell surface and to ensure viability during the
entry into quiescence. The pkc1 and the related mutants in the pathway die in rapamycincontaining, nutrient-replete medium, therefore, the PKC1 pathway seems not act as a nutrient
sensor (Krause and Gray, 2002).
Microbes accumulate reducing compounds during quiescence

7

A myriad of metabolic adjustments takes place to meet the changing demands of cells during
quiescence (Gray et al., 2004; Valcourt et al., 2012). The rule of thumb is: a lower anabolism
such as the shift away from TCA (tricarboxylic acid) cycle to protect against the superoxide
produced by oxidative phosphorylation, and a higher catabolism that includes, for example, the
increased abundance of glycolysis enzymes. A remarkable exception in microbial quiescence is
the accumulation of carbon stores, even though the chemical structure of the storage differs.
Quiescent yeast cells accumulate glycogen, a disaccharide trehalose, and triacylglycerols (TAGs)
as the main forms of metabolizable carbon storage (Lillie and Pringle, 1980; Hosaka and
Yamashita, 1984). The bacterial pathogen Vibrio cholera produces glycogen when living in
nutrient-poor aquatic environments (Bourassa and Camilli, 2009). Mycobacterium tuberculosis
stores TAGs and wax esters in large cytosolic inclusions in preparation for a dormancy-like state
(Daniel et al., 2004; Sirakova et al., 2012). For bacteria living in soil and the rhizosphere,
intracellular deposition of polyhydroxyalkanoates, a group of bioplastic carbon polymers, is
critical for enduring suboptimal conditions (Kadouri et al., 2005). In addition, C. reinhardtii
accumulates starch granules and TAGs during starvation for N, phosphorus, sulfur, zinc or iron,
and in response to high salt or heat stress (Hu et al., 2008; Matthew et al., 2009; Kropat et al.,
2011; Siaut et al., 2011; Hemme et al., 2014).
The importance of storage compounds to quiescence is an unresolved question. In yeast,
trehalose and glycogen decrease slowly with time during long-term stationary phase. However,
these carbohydrates are probably not metabolized for energy, at least not during quiescence, as
they do not always correlate with viability (Sillje et al., 1999). The primary function of trehalose
is thought to protect proteins from damage by oxygen radicals, and to act as a chemical
chaperone to assist protein folding (Singer and Lindquist, 1998; Benaroudj et al., 2001). Mutants

8

unable to synthesize trehalose exit quiescence much more slowly (Sillje et al., 1999; Shi et al.,
2010). Hydrolyzing one molecule of trehalose releases two molecules of glucose, whereas
cleavage of a single glycosidic bond in glycogen produces just one, making trehalose the obvious
carbohydrate of choice to fuel quiescence exit. However, trehalose is more osmotically active
than glycogen, which could be a problem. TAG has the highest energy output per weight of
material in cellular metabolism, and is rapidly hydrolyzed when supplemented with fresh
medium (Kurat et al., 2006). TGL3 and TGL4 are the major TAG lipases in yeast; the tgl3 tgl4
double mutant completely loses lipolytic activity and exhibits delayed daughter cell formation
and cell cycle progression (Kurat et al., 2006; Kurat et al., 2009). In Chlamydomonas, TAGs
accumulated during quiescence function at least in part as energy sink to accommodate the
reducing power generated by the photosynthetic machinery (Li et al., 2012). A mutant producing
less than half of the wild-type TAG content loses viability during the N deprivation-induced
quiescence, likely due to a detrimental overreduction of the photosynthetic electron transport
chain. It bleaches during N deprivation, a phenotype reversed by inhibition of the photosynthetic
electron transport chain (Li et al., 2012).
Notably, there is an inverse relationship between growth and TAG production. Growthlimiting stresses in Mycobacterium, e.g. antibiotic treatment, lead to the induction of TAG
biosynthesis. Mycobacterium mutants whose dominant TAG synthase is disrupted are unable to
arrest their growth and continue to proliferate in the conditions that normally induce quiescence
(Baek et al., 2011). It appears to result from redirecting the acetyl-CoA from the TCA cycle,
where it is used to generate chemical energy during aerobic respiration, to the synthesis of fatty
acids, which are subsequently stored in TAGs. The growth-limiting effect is not restricted to
Mycobacterium. For example, yeast mutants unable to produce glycogen or trehalose persist with

9

high TCA fluxes during stationary phase (Sillje et al., 1999). This inverse relationship is also
well known for microalgae, and has long been hampering the progress in developing microalgae
into sustainable biofuel feedstocks. The almost universal propensity of microorganisms to
channel acetyl-CoA into TAG storage in response to quiescence may act as a common strategy
for reducing growth and altering the metabolic state.

Chlamydomonas reinhardtii as a reference model to study cellular quiescence
The unicellular green alga Chlamydomonas reinhardtii has a short life cycle, a sequenced
genome (Merchant et al., 2007), and a growing molecular toolbox for forward and reverse
genetic studies, including insertional mutagenesis (Kindle, 1990), gene silencing (Kim and
Cerutti, 2009; Molnar et al., 2009), and fluorescent protein-tag (Rasala et al., 2013). In addition,
a genome-wide mutant library is under construction and will offer a powerful resource
comparable to the Arabidopsis collections. Among the principle areas using this model system
are eukaryotic flagellar structure and function, basal bodies (centrioles), cell-cell recognition, cell
cycle control, chloroplast biogenesis, phototaxis, nonphotochemical quenching, and especially
photosynthesis for Chlamydomonas can grow in the dark on an organic carbon (e.g. acetate), and
thus provides advantages over land plants (Harris, 2001; Peers et al., 2009).
The cell size of C. reinhardtii ranges between 5 to 10 μm; intracellular space is mostly
occupied by a single chloroplast. The life cycle of C. reinhardtii is very simple. Under vegetative
growth cells maintain haploidy; when N levels fall below threshold, they differentiate into
sexually active gametes. Contact between opposite mating types (mt+ and mt-) initiates a rapid
fertilization and formation of a diploid zygote that is astonishingly resistant to environmental
insults. When conditions improve, the dormant zygote undergoes meiosis and gives rise to four

10

recombinant haploid products (Sager and Granick, 1954; Goodenough et al., 2007). The cell
cycle of C. reinhardtii is a variation of the eukaryotic cell cycle, with additional rapid cell
divisions bypassing G1 until a specific daughter cell size has been reached (Figure 1.3) (Bisova
et al., 2005).

Figure 1.3. Chlamydomonas life cycles. The cell division cycle with G1, S, G2, and M phases,
and the quiescence cycle (G0) are depicted. -N, N deprivation, +N, N resupply.

The unicellular green alga Chlamydomonas reinhardtii was chosen as a reference model
organism to investigate life cycle transitions in photosynthetic eukaryotes for several reasons:
First, the two life cycle states of interest—quiescence and cell division—can be discretely
defined and controlled. Second, an extensive number of system-level studies of N deprivation
has been completed in recent years in Chlamydomonas, and mutants with unique phenotypes
during N-deprivation are available, setting a solid foundation for investigating cellular
quiescence in the context of N deprivation (Bolling and Fiehn, 2005; Li et al., 2010; Miller et al.,
2010; Moellering and Benning, 2010; Work et al., 2010; Nguyen et al., 2011; Li et al., 2012;
Blaby et al., 2013; Schmollinger et al., 2014).

Aims of the dissertation research
The research of this dissertation is dedicated to the understanding of cellular quiescence in
photosynthetic organism, using C. reinhardtii as a reference model. Chapter 2 describes the
11

discovery of CHT7, a repressor of cellular quiescence. Chapter 3 presents the transcriptomic
regulation of the quiescence cycle. Chapter 4 reports the role of lipid droplets in coordinating
lipid dynamics to ensure the reversibility of quiescence.

12

REFERENCES

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CHAPTER 2
Protein Compromised Hydrolysis of Triacylglycerols 7 (CHT7) Acts as a Repressor of
Cellular Quiescence in Chlamydomonas1

_____________________________
1

This research was published in: Tsai, C.H., Warakanont, J., Takeuchi, T., Sears, B.B.,
Moellering, E.R., and Benning, C. (2014). The protein Compromised Hydrolysis of
Triacylglycerols 7 (CHT7) acts as a repressor of cellular quiescence in Chlamydomonas. Proc.
Natl. Acad. Sci. U S A. 111, 15833-15838. I contributed to the data shown in all of the figures.

21

ABSTRACT
Microalgae are prolific photosynthetic organisms that have the potential to sustainably produce
high-value chemical feedstocks. However, an industry based on microalgal biomass still is faced
with challenges. For example, microalgae tend to accumulate valuable compounds, such as
triacylglycerols, only under stress conditions that limit growth. To investigate the fundamental
mechanisms at the base of this conundrum—the inverse relationship between biomass
production and storage compound accumulation—I applied a combination of cell biological and
genetic approaches. Conceptually, nutrient deprivation, which commonly is used to induce the
accumulation of triacylglycerol in microalgae, leads to a state of cellular quiescence defined by a
halt of cell divisions that is reversible upon nutrient resupply. To identify factors that govern
cellular quiescence, we screened for mutants of the model alga Chlamydomonas reinhardtii that,
in contrast to wildtype cells placed under conditions of nitrogen deprivation, were unable to
degrade triacylglycerols following nitrogen resupply. One of the mutants described here in detail,
compromised hydrolysis of triacylglycerols 7 (cht7), was severely impaired in regrowth
following removal of different conditions inducing cellular quiescence. The mutant carries a
deletion affecting four genes, only one of which rescued the quiescence phenotype when
reintroduced. It encodes a protein with similarity to mammalian and plant DNA binding proteins.
Comparison of transcriptomes indicated a partial derepression of quiescence-related
transcriptional programs in the mutant under conditions favorable to growth. Thus, CHT7 likely
is a repressor of cellular quiescence and provides a possible target for the engineering of highbiomass/high-triacylglycerol microalgae.

22

INTRODUCTION
Nutrient deprivation of microalgal cultures provides a facile experimental tool to induce and
study triacylglycerol (TAG) accumulation in lipid droplets and is used in biotechnological
settings for the production of high-value oils (Hu et al., 2008; Liu and Benning, 2013). In
particular, responses to the withdrawal of nitrogen (N) have been studied widely in the model
unicellular green alga Chlamydomonas reinhardtii, and a comprehensive picture of N-sparing
mechanisms during N deprivation is emerging through integrated global analysis of transcripts,
proteins, and metabolites (Miller et al., 2010; Blaby et al., 2013; Schmollinger et al., 2014).
Mechanisms of lipid droplet formation following N deprivation and proteins associated with
lipid droplets are being explored (Moellering and Benning, 2010; Goodson et al., 2011; Nguyen
et al., 2011), and mutants have become available that provide mechanistic insights in vivo into
specific aspects of the lipid biosynthetic machinery of C. reinhardtii required for TAG
accumulation (Boyle et al., 2012; Li et al., 2012; Nguyen et al., 2013).
From a cell biological viewpoint, N deprivation induces cellular quiescence, a reversible
state of the cell cycle during which cell divisions temporarily cease and cells are reprogrammed
to adjust metabolism for survival of the adverse condition (Valcourt et al., 2012). In C.
reinhardtii, metabolic changes during N deprivation-induced quiescence include the partial
degradation and reorganization of the photosynthetic apparatus and the protein biosynthetic
machinery, induction of lipase and autophagy genes, and accumulation of carbon storage
compounds (Miller et al., 2010; Blaby et al., 2013; Schmollinger et al., 2014). N deprivation also
induces gametogenesis (Beck and Acker, 1992), allowing cells of opposite mating types to fuse
and form thick-walled zygospores that can survive temporary harsh conditions. The goal of this
study was to gain mechanistic insights into the regulation of N deprivation-induced quiescence in

23

C. reinhardtii and to answer the question of how changes in the metabolic status of the algal cell
induced by N deprivation affect progression through the cell cycle or the entry and exit into and
out of quiescence. Toward this end, I searched for mutants of C. reinhardtii unable to rapidly
exit quiescence, readjust their metabolism, and resume growth following N resupply after a
period of N deprivation. As proxy for the metabolic status of the cell, I focused on TAG
degradation following N resupply and identified a mutant and corresponding protein that met the
criteria for a repressor of quiescence in C. reinhardtii.

24

MATERIALS AND METHODS
Strains, Genetic Analysis, and Growth Conditions
Chlamydomonas reinhardtii cell wall-less strain dw15 (cw15, nit1, mt+) was obtained from A.
Grossman, Department of Plant Biology, Carnegie Institution for Science, Stanford, CA, and is
referred to as the wild type (with regard to CHT7) parental line (PL) throughout. CC-198 (er-u37, str-u-2–60, mt-) and CC-110 (spr-u-1–6-2 mt+) were obtained from the Chlamydomonas
Resource Center (www. chlamycollection.org). CC-198 was crossed to cht7 as described
previously (Li et al., 2012). Progeny were analyzed for the hygromycin B resistance marker aph7,
mating type, and cell wall traits. A hygromycin-resistant, mt−, cw+ meiotic product (2-5-1) was
selected and crossed to CC-110, producing zygospores that were used for tetrad analysis to test
for cosegregation of the aph7 marker and the cht7 phenotypes.
Complemented cht7 lines were generated with two similar methods. In method 1,
cotransformation, a 7,873-bp genomic fragment containing the intact CHT7 gene with 1-kb
flanking sequences on both ends was excised from BAC 21K10 (Clemson University Genomics
Institute) by using EcoRI and HindIII. Plasmid pMN24 (Fernandez et al., 1989) carrying the C.
reinhardtii nitrate reductase gene NIT1 as selection marker was linearized with EcoRV. Both
DNA fragments were cotransformed into cht7. In each transformation, 0.5 μg of linearized
pMN24 and 0.5 μg gel-purified BAC fragment were used. In method 2, transformation of the
selectable marker linked to the BAC fragment, the 7,873-bp fragment was inserted into the
EcoRI and HindIII sites of pBR322. The construct then was digested by EcoRV and SspI to
release an 8,220-bp fragment that included the 7,873-bp fragment. The 8,220-bp fragment was
inserted into EcoRV-linearized pMN24 to produce pMN24-CHT7. EcoRV-linearized pMN24CHT7 was used for transformation of the cht7 mutant. TAP plates (see below) containing 10 mM

25

nitrate instead of 10 mM ammonium were used for selection. Colonies were picked into 96-well
plates and grown N deprived and N resupplied following the same protocol described below.
Cell growth after N resupply was measured with a FLUOstar Optima 96-well plate reader (BMG
Labtech). Those able to resume growth after N resupply likely were complemented. Strains
having pMN24 alone were used as empty vector control. Complementation lines C1 and C2 were
produced by method 1; C3 and C4 were by method 2. Primers of APH7-F and APH7-R and
primers of CHT7-F and CHT7-R were used to validate the retention of the aph7 insertion and the
reintroduction of CHT7 gene, respectively (Figure 2.3). All primer sequences are listed in Table
2.1.
In general, cells were grown in TAP medium (Harris, 1989) under continuous light (70–
80 μmol·m−2·s−1) at 22 °C or ambient room temperature (∼22 °C) for solid media, which
contained 0.8% agar (Phytoblend; Caisson Labs). Concentration of cells was monitored with a
Z2 Coulter Counter (Beckman Coulter) or a hemocytometer.

Generation of Antibodies, Immunoblotting, Subcellular Fractionation, and BN-PAGE
To generate antibodies against MLDP, the full-length coding sequence of MLDP was amplified
using primers (MLDPcds-F/MLDPcds-R) and total cDNA as the template, and was inserted into
the BamHI and XhoI sites of pET28B(+) (Novagen EMD Chemicals). For the development of
antibodies against CHT7, the full-length coding sequence of CHT7 was synthesized (Life
Technologies) with codons optimized for expression in Escherichia coli. The synthesized CHT7
sequence was inserted into the EcoRI and HindIII sites of pET28B(+). Both constructs were
introduced into E. coli BL21 (DE3). Recombinant proteins (6xHis-MLDP and 6xHis-CHT7)
were purified using Ni-nitrilotriacetic acid agarose (Qiagen) and separated by SDSPAGE to

26

examine their purity. Few other proteins were copurified with 6xHis MLDP, yielding a fairly
clean substrate for the generation of antibodies. In contrast, the elution of 6xHis-CHT7 contained
many other proteins. 6xHis-CHT7 was excised from the SDS-PAGE gel and eluted. Roughly 2
mg of each protein was sent for antibody production in rabbits by Cocalico Biologicals, Inc.
Immunoblots using MLDP antiserum are shown in Figure 2.1A; those using CHT7 antiserum are
shown in Figure 2.12A.
For immunoblot analysis, SDS-PAGE gels were blotted onto PVDF membranes in
transfer buffer [25 mM Tris, 192 mM Gly, and 10% (vol/vol) methanol] for 60 min at 100 V.
Membranes were blocked for 60 min in TBST [50 mM Tris, 150 mM NaCl, 0.05% (vol/vol)
Tween 20, pH 7.6] with 5% (wt/vol) nonfat dry milk and probed with primary antibodies
overnight at 4 °C. The primary antibodies (MLDP antiserum and CHT7 antiserum) were used at
1:1,000 dilutions. Goat anti-rabbit secondary antibodies coupled to horseradish peroxidase were
used at 1:10,000 dilution and incubated with blots at room temperature for 30 min. Blots then
were washed six times with TBST at room temperature for 10 min each. Antigen was detected
by chemiluminescence (Bio-Rad Clarity Western ECL substrate) using a charge-coupled device
(CCD) imaging system. For samples containing detergent or reducing agent, the RC DC Protein
Assay Kit (Bio-Rad) was used for protein quantification.
C. reinhardtii nuclei were isolated as described previously (Winck et al., 2011) using a
CelLytic PN kit (Sigma-Aldrich). Chloroplast fractions were prepared according to (Mason et al.,
2006). Organelle pellets were solubilized in 2× SDS sample buffer, boiled at 95 °C for 5 min,
and centrifuged at 20,000 × g for 10 min to remove insoluble portions. Supernatants were
collected and tested for the enrichment of CHT7 protein.

27

For BN-PAGE analysis, 50 mL of cells were pelleted and resuspended in PBS with
protease inhibitor (P9599; Sigma-Aldrich) and phosphatase inhibitor (Thermo Scientific) at a
concentration of 109 cells/mL and then lysed by sonication on ice with 0.6 s on and 0.5 s off for
20 s per cycle with 20 cycles and 20 s between each cycle. The sonication was repeated 20 times.
Insoluble materials were removed by centrifugation at 20,000 × g for 30 min at 4 °C. Protein
content in this case was determined using the Quick Start Bradford protein assay (Bio-Rad).
Supernatants containing 25 μg of proteins were loaded onto each lane of BN-PAGE Novex 4–
16% (wt/vol) Bis-Tris gels (Life Technologies), and BN electrophoresis was performed as
described (Wittig et al., 2006). After denaturation, gels were either blotted directly or run in a
second dimension on a 10% (wt/vol) SDS-PAGE gel followed by immunoblotting.

Mutant Screen
Insertional mutagenesis was done as described previously (Li et al., 2012) except for the use of a
shorter, 2,012-bp PvuII fragment of the pHyg3 plasmid that contains only the hygromycin B
resistance gene aph7. Hygromycin B-resistant colonies were picked into 96-well cell culture
plates (Corning) with 200 μL of TAP medium and grown for 3 d. The plates were replicated
before the cultures were subjected to 48 h N deprivation followed by 24 h N resupply as
described above. For processing, the cultures were transferred to a 96-well PCR plate (Life
Technologies) and centrifuged at 2,000 × g. The cell pellets were resuspended with 50 μL of 2×
SDS sample buffer [0.1 M Tris, pH 6.8; 2.75% (wt/vol) SDS; and 5% (vol/vol) βmercaptoethanol] and boiled at 95 °C for 5 min in a Bio-Rad iCycler thermocycler. Lysates from
each well then were blotted onto an Amersham Hybond-ECL nitrocellulose membrane (GE
Healthcare) placed on top of one layer of Whatman filter paper by using a MilliBlot- D 96-well

28

filtering system (Sigma-Aldrich). Immunodetection of MLDP was done as described above.
During the primary screen of 1,760 insertional lines, 13 putative mutants were selected for
secondary (standard immunoblot) and tertiary tests (lipid analysis), which led to the confirmation
of eight mutants with the cht phenotype.

Confocal Microscopy and Construction of the CHT7-GFP Fusion
For the detection of lipid droplets stained by Nile red, cultures of cells were incubated with the
fluorescent dye Nile red in the dark at a final concentration of 2.5 μg/mL (from a stock of 50
μg/mL in methanol). Cells were immobilized by spotting on poly-L-lysine–coated slides
(Electron Microscopy Sciences). Images were captured with a FluoView FV10i confocal
microscope (Olympus America). The 488-nm argon laser was used in combination with a 560–
615-nm filter; for chlorophyll autofluorescence, a filter for far-red was used.
To observe subcellular localization of CHT7, pMN24-CHT7- GFP was constructed to
express the translational fusion under the control of the endogenous promoter and terminator.
Plasmid pMN24-CHT7 was digested with AvrII to remove a 2,829-bp fragment containing the 3'
end of CHT7 genomic sequence. The 2,829-bp fragment was inserted into pBR322 linearized
with NheI to obtain pBR322-2829. C. reinhardtii codon-optimized EGFP was amplified by
Phusion DNA polymerase (New England BioLabs) from pJR38 (Neupert et al., 2009) using
primers GFP-F (SmaI cut site) and GFP-R (SpeI cut site), which also introduced an N-terminal
linker (GAAAAAAAAA). This PCR product was inserted into pPCR-Blunt using the Zero Blunt
PCR Cloning Kit (Life Technologies) to produce pPCRBlunt-GFP and sequenced. The GFP
fragment was excised by SmaI and SpeI and inserted into the PmlI and SpeI sites of pBR3222829 to obtain pBR322- GFP, which then was digested with FseI and SpeI to obtain a 1,486-bp

29

fragment. This fragment is composed of the last 701 bp of the CHT7 gene (before the stop
codon), glycine/alanine linker, GFP, and the first 44 bp of CHT7 3' UTR. The larger DNA
product (∼25 kb) from the FseI/SpeI double digestion of pMN24-CHT7 was gel purified with
the QIAEX II Gel Extraction Kit (Qiagen) and served as the vector for the 1,486-bp insert to
complete pMN24-CHT7-GFP. The insert from this plasmid was sequenced to confirm that GFP
was in frame with CHT7. Confocal images were collected sequentially with the Olympus
FluoView 1000 Confocal Laser Scanning Microscope (Olympus America) using a 100×
UPlanSApo oil objective (N.A. 1.4). Hoechst 33342 fluorescence was excited with the 405-nm
diode laser while the fluorescence emission was collected from 430 to 470 nm. GFP fluorescence
was excited with a 488-nm argon gas laser while the fluorescence emission was collected from
500 to 530 nm. Far-red autofluorescence was excited with a 559-nm solid-state laser while the
fluorescence emission was collected at 655–755 nm.

Phenotyping Assays
To induce N deprivation, midlog-phase cells grown in TAP were collected by centrifugation
(2,000 × g, 4 °C, 2 min), washed twice with TAP-N (NH4Cl omitted from TAP), and
resuspended in TAP-N at 0.3 OD550. N was resupplied with either of two methods that gave the
same outcome in all of the physiological and biochemical experiments tested. In method 1, after
48 h of N deprivation, cells were pelleted by centrifugation (2,000 × g, 4 °C, 2 min) and
resuspended with the same volume of TAP. In method 2, 1% culture volume of 1 M NH4Cl
(100×) was added to the N-deprived culture. Except for the primary mutant screen, method 2 was
used to avoid physical damage to cells during centrifugation. To conduct P deprivation or

30

rapamycin treatment, cells were washed and resuspended in medium without phosphate or with 1
μM rapamycin. Conditions were reversed by washing and resuspending cells in regular TAP.
Cell viability was assessed by using SYTOX Green (Molecular Probes Inc.), to which
living cells are impermeable but which binds the nucleic acids of dead cells, yielding intense
fluorescence. Cells were grown in TAP-N. At the times indicated (Figure 2.8B), 1 μL of 5 mM
SYTOX Green was added to 1 mL culture and incubated for 5 min in the dark. The cells were
observed by fluorescence microscopy (Leica DRMA2) with excitation at 488 nm using a FITC
filter for SYTOX Green and a Texas Red filter for chlorophyll autofluorescence. Data were
collected with FV1000 ASW software (Olympus).
To assess the ability of cells to divide, a set volume of culture deprived of N for the time
indicated (Figure 2.8C) was diluted in warm TAP medium with 0.4% agar (PhytoBlend) that was
not yet solidified and poured evenly over the solid TAP medium. Colony-forming units (CFUs)
and the diameter of individual colonies were measured 11 d later. CFUs were monitored for
another 2 wk, with few additional colonies forming. Cells from a second aliquot were counted
with a hemocytometer to provide a denominator to calculate the fraction (%) of CFUs per cells
plated.
For lipid analysis, extraction, TLC of neutral lipids, fatty acid methyl ester (FAME)
preparation, and gas–liquid chromatography were conducted as previously described in
(Moellering and Benning, 2010). Cell pellets were extracted into methanol and chloroform (2:1
vol/vol). To the extract, 0.5 volume of 0.9% KCl was added and mixed, followed by phase
separation at low-speed centrifugation. For TAG quantification, lipids were resolved by TLC on
Silica G60 plates (EMD Chemicals) developed in petroleum ether-diethyl ether-acetic acid
(80:20:1 by volume). After visualization by brief iodine staining, FAME of each lipid or total

31

cellular lipid was processed and quantified by gas chromatography as previously described
(Rossak et al., 1997).

Standard DNA and RNA Procedures
C. reinhardtii genomic DNA was isolated as previously described (Newman et al., 1990). For
Southern blotting, DNA digested with BamHI was separated by agarose gel electrophoresis (10
μg DNA per lane). DNA was transferred to a nylon membrane (Amersham Hybond-N+; GE
Healthcare) and UV cross-linked. The probe was labeled by digoxigenin through PCR
amplification of a 234-bp region within the hygromycin B resistance cassette with primers S-F
and S-R. Prehybridization, hybridization, and chemiluminescent detection were performed using
a kit from Roche following the manufacturer’s instructions.
To identify the locus disrupted by insertional mutagenesis, SiteFinding-PCR (Tan et al.,
2005) was used with minor modifications and with primers designed for the pHyg3 plasmid. The
primers used for finding the insertion in cht7 were SiteFinder1 in combination with GSP3-3 and
GSP3-4 and SiteFinder3 in combination with GSP8-02 and GSP8-0. In addition, nested primers
SFP1 and SFP2 were used for both combinations. To confirm the results of SiteFinding-PCR,
standard PCR was performed using primers amplifying regions across the ends of the insertion
and the flanking sequences (CHT7-1-F/CHT7-1-R; CHT7-2-F/CHT7-2-R; CHT7-3-F/CHT7-3-R;
CHT7-4-F/CHT7-4-R). All primers may be found in Table 2.1.

32

Table 2.1. Oligonucleotide primers used in this study
Name

Sequence

MLDPcds-F

GGATTCATGGCCGAGTCTGCTGGAAA

MLDPcds-R CTCGAGGCATCATAGCACAAGGCATT
S-F

ACCAACATCTTCGTGGACCT

S-R

CTCCTCGAACACCTCGAAGT

SiteFinder1

SiteFinder3

CACGACACGCTACTCAACACACCACCTCGCACAGCGTCCTCAAGCGG
CCGCNNNNNNGCCT
CACGACACGCTACTCAACACACCACCTCGCACAGCGTCCTCAAGCG
GCCGCNNNNNNGCCG

SFP1

CACGACACGCTACTCAACAC

SFP2

ACTCAACACACCACCTCGCACAGC

GSP3-3

ACTGCTCGCCTTCACCTTCC

GSP3-4

CTGGATCTCTCCGGCTTCAC

GSP8-0

CGCCCTACCTTTTGCTGGA

GSP8-02

GGTCGAAGCATCATCGGTGT

CHT7-1-F

ACACTTAGACCCGTGGCTTC

CHT7-1-R

TGGAAGTGTCATAGCGCAAG

CHT7-2-F

AAACCATGCAAAAGGTGCAC

CHT7-2-R

CGCTCACAATTCCACACAAC

CHT7-3-F

CTCAAGTGCTGAAGCGGTAG

CHT7-3-R

TGGTGGTCGACAAACTCTTG

CHT7-4-F

TTTACAACGTCGTGACTGGG

CHT7-4-R

CATGAGGTATGGTGGTCGAC

γ-tubulin -F

CGCCAAGTACATCTCCATCC

γ-tubulin -R

TAGGGGCTCTTCTTGGACAG

Gene1-F

TTCTCGGGTCAGATATTGGG

Gene1-R

TTGAGGCAGAACGACTTCTTG

Gene2-F

GTGTCCTACACAGTTCGA

Gene2-R

GTAGTACACCTTGCTTCG

Gene3-F

AATTCTCAAGTAAGTAAG
33

Table 2.1 (cont’d)
Gene3-R

TATATACAAAACAACGAATA

Gene4-F

TATTCCTCTCAAAGTCGATT

Gene4-R

TTGCATCTTTGATGTAGAAC

APH7-F

CTCAAGTGCTGAAGCGGTAG

APH7-R

TGGTGGTCGACAAACTCTTG

CHT7-F

TTCTCGGGTCAGATATTGGG

CHT7-R

TTGAGGCAGAACGACTTCTTG

GFP-F

CCCGGGAGCGGCAGCGGCTGCGGCAGCAGCCGCGATGGCCAAGGGC
GAGGAGCT

GFP-R

ACTAGTTTACTTGTACAGCTCGTCCA

Gene Name

Gene IDa

CBLP

g6364

CHT7

Cre11.g481800

APG3

Cre02.g102350

APG8

Cre16.g689650

ACCCCCGACATTCAAGCA / TGCGGCCTCCGCTTT

LHCA3

g11502

GGGTGTGTCGTGCTTGCA / CGCACGCAGGCACAGA

LHCB4

Cre17.g720250

LHCBM1

g1276

LHCBM4

Cre06.g283950

NAB1

Cre06.g268600

PSAD

g5492

qPCR Primer Sequence (Forward / Reverse)
GCCACACCGAGTGGGTGTCGTGCG /
CCTTGCCGCCCGAGGCGCACAGCG
TCGCGGTGTCTAAAATTGTACTG /
GGTTGTTAAAGCACTGCATGCA
GACGACATTCCCGACATCACT /
GCCGCAGCCTCATCATCT

CGTGGTGATTGGCGTTGAC /
GACGGAGCCCTTGTTGTTCTT
CCAAGTTCACCCCCCAGTAA /
TGCATCCCTATTGGTACATCAAA
GAAACCCCGCAGAAGTGAAG /
CACGGTAAGGGCCATTTGC
CGACTGGTGCCCACGACTA /
CACATACACTCCTCCTGCATTTTC
TTGCGACCCCGCAGTT /
CACGCACACGCTAACATCTACA

34

Table 2.1 (cont’d)
CCGGTTGCGCATCGA /

MCA1

Cre08.g358250

PSBS1/

Cre01.g016600/

TGTGCAACACCCTTCAAAATG /

PSBS2

Cre01.g016750

GCGCTGGGCGAAAGC

FAP113

Cre07.g321400

FAP138

Cre14.g632350

FAP139

g9599

FAP292

Cre03.g162850

IFT46

g5332

IFT57

Cre10.g467000

FA1

Cre06.g257600

FA2

Cre07.g351150

CHLD

Cre05.g242000

CHLI2

Cre12.g510800

CPX1

Cre02.g085450

CPX2

Cre02.g092600

POR

Cre01.g015350

AAAACCAAGCATTAACCGATCAA

GCGGAGGCATGTCATATGG /
CTGTCTCTGCAAATGTATGTCAGTGT
CGCATGCACCGAGAACAC / CACGCCCGGGCCTAA
GCCGCGGAGATGGCTAA /
CTTATTGCACGAGGCTAACCAA
TGGCTGCGCCATGGTT /
GAGTCCTGCGGTCAGGGTAA
CCAGCTGCGCCTAGAGATG /
TGGCCCCTCGTCCATGT
TGAGGCAGGCATGGCATA /
GGATACCGCTGCCTGCTTT
TCTTGGCTTTCTGCTGTGTGTAC /
GTGCACAAGACCGGTCCTTT
GGCGATTGACGGACTATGAAC /
TTAAAGACCGCGCCAAAGG
CGAGCTCAAGAAGATTTGGTTGA /
CCGAAGATGCAAGCCATAATATG
CATGTGTTGCGGTGTGCTTT /
TCCCACCCGCTCAGTCA
TGGTCGTCGACTTCCTGTGTAA /
ATGGGTACGCAGGACAGTAACA
TACGGCGGTGGCTGTGA /
ACTGCGAGTCCTCCACGTCTA
GCTGCTGGATGACCTGAAGAA /
TGATGGAGCCGACGATGAT

35

Table 2.1 (cont’d)

a

UROD1

Cre11.g467700

BCC1

Cre17.g715250

BCC2

Cre01.g037850

BCR1

Cre08.g359350

HAD1

Cre03.g208050

KAS2

Cre07.g335300

MCT1

Cre14.g621650

APX2

g10003

CAT1

Cre09.g417150

HPR1

Cre06.g295450

MAS1

g2904

MDH2

Cre10.g423250

PXN1

Cre07.g353300

CGCTTCGGCTGAGTGTTTAGA /
AGGCTCCCCGTCCATCA
CCACAGTCGCGAATCACAAT /
GGTGGCGGCGCACTT
CTGGTGGTGTGCTGCTGACT /
CGGCTCAGAACCTGAATAACAA
AACCGCGTGCTGATCAATG /
TGAAAGTGCCCTAAAACCAAACA
CTGACCCACATGAGACATGACA /
GCGGCGCATTCGTTGT
TGGGCCGGCTGTACGA /
CTCAGCTGTATCGAAGCTTTAAGATC
GGTGTCTAGGCGCATCACTTTT /
TGAGCATGGTGGCCATCTT
CTGCGCGAGGTGTTTGG /
GCGCCACAATGTCCTTGTC
GGAGGCTGCAGGAAAACTGA /
TCTCCAGCCTGGGCTACCT
TGCATCTTGCATTGGTTACATG /
CGCGTTCCCTGGCTCAT
GCCCCTTGTGCCATATGC / CCCCGATGCTGCTGTCA
GAACCGCATTCAAAAGATTGC /
TGAACTTCCGCGGTTTCG
GCTGCAAATGCATTGGATCA /
CCGGCGTGAGTTAAGAACCA

Corresponding to the version 5.3.1 genome. All primer sequences are written 5’ to 3’. N

indicates a random nucleotide. Except CBLP, all other qPCR primers were designed by Primer
Express Version 3.0 (Life Technologies).

36

To produce cDNA templates used for qPCR, total RNA was purified with the RNeasy
PlantMini Kit (Qiagen) including the oncolumn DNase digestion and then subjected to reverse
transcription by using SuperScript III reverse transcriptase (Life Technologies). qPCR was
performed on an ABI Prism 7000 (Applied Biosystems) using SYBR Green PCR Master Mix
(Life Technologies) with an equivalent cDNA template and 0.25 μM of each primer. The amount
of cDNA input was optimized after serial dilutions. In all qPCR experiments, expression of the
target gene was normalized to the endogenous reference gene CBLP, a gene commonly used for
normalization in C. reinhardtii (Allen et al., 2007), using the cycle threshold (CT) 2−ΔΔCT
method. All experiments were done using at least two biological replicates, and each reaction
was run with technical replicates. qPCR procedures and analysis followed the minimum
information for publication of quantitative real-time PCR experiment guidelines (Bustin et al.,
2009). For quantification, TRIzol reagent (Life Technologies) was used for RNA isolation, and
the concentration of RNA was measured with a NanoDrop instrument (Thermo Scientific).
qPCR primers are listed in Table 2.1.

Illumina RNA Sequencing and Bioinformatics
For Illumina RNA sequencing, total RNA was isolated by using the Qiagen RNeasy Plant Mini
Kit including the on-column DNase digestion. RNA integrity was examined with a 2100
Bioanalyzer (Agilent Technologies), and every sample had an RNA integrity number greater
than 7.0. Three biologically independent sets of samples were prepared at different times and
submitted directly to the Michigan State University Research Technologies Service Facility
(rtsf.natsci.msu.edu/) for sequencing on an Illumina Genome Analyzer II (Illumina). The cDNA
libraries were prepared for single-end sequencing. Default parameters were used to pass reads by

37

using Illumina quality control tools. The filtered sequence data were deposited at the National
Center

for

Biotechnology

Information

Sequence

Read

Archive

(www.ncbi.nlm.nih.gov/Traces/sra/) with the BioProject ID PRJNA241455 for the Illumina
dataset.
RNA abundance in the samples was computed using the CLC Genomics Workbench
(www.clcbio.com/corporate/about-clcbio/), version 5.5.1. Reads were trimmed and filtered based
on

quality

with

the

Trim

Sequences

algorithm

clcgenomicsworkbench/650/index.php?manual=Trimming.html;
ambiguities:

2).

Genome

sequence

and

annotations

(www.clcsupport.com/

Limit:

were

0.05,

downloaded

Maximum
from

JGI

(www.phytozome.net/ chlamy.php). C. reinhardtii version 5.3.1 was used. Differential
expression was determined by using the numbers of mapped reads overlapping with annotated C.
reinhardtii genes as inputs to DESeq, version 1.10.1 (Anders and Huber, 2010). GO analysis of
RNA-Seq data was performed using Goseq, version 1.10.1 (Young et al., 2010). I used the
default Wallenius approximation to calculate the over- and underrepresented GO categories
among differentially expressed genes using a P value of 0.05. The hierarchical clustering and
heat maps were calculated and drawn using Qlucore Omics Explorer (qlucore.com/).

38

RESULTS
Isolation of compromised hydrolysis of triacylglycerols Mutants
The amount of TAG in C. reinhardtii coincides with the abundance of the major lipid dropletassociated protein (MLDP) (Moellering and Benning, 2010) (Figure 2.1A). This correlation was
used to identify mutants of C. reinhardtii with altered TAG degradation after the resupply of N
to N-deprived cells. An immuno-dot blot-based assay for MLDP allowed us to screen indirectly
for TAG abundance in a mutant population generated by random insertion of a selectable marker
(Figure 2.1B). Insertion lines were cultured in N-replete medium for 48 h, followed by 48 h of N
deprivation, and then resupplied with N. Typically, 24 h after N resupply, most of the TAG
accumulated during N deprivation was hydrolyzed and MLDP was degraded in the parental line
(PL), dw15 (Figure 2.2A). Putative mutants with a persistent MLDP immunosignal after 24 h N
resupply were designated compromised hydrolysis of triacylglycerols (cht). Among the initial
1,760 insertion lines, eight putative cht mutants were identified, with cht7 showing the greatest
delay in MLDP degradation (Figure 2.2A). Importantly, TAG content also decreased with a
severe delay in response to N resupply in cht7 (Figure 2.2B). Lipid droplets observed following
Nile red staining of cells of the PL started to decrease in size after 12 h of N resupply, and by 24
h, these cells were virtually devoid of lipid droplets (Figure 2.2C). In contrast, cht7 cells retained
lipid droplets even after 24 h.

39

Figure 2.1. Isolation of cht mutants. (A) Immunoblot quantification of MLDP (top panel) and
TAG accumulation (Bottom) in dw15 [depicted as the ratio of TAG fatty acids (FA) over total
FAs] after N deprivation (−N) followed by N resupply (NR) for the indicated times (hours). (B)
Overview of the mutant isolation procedure using immunodot blots with MLDP antibodies.

40

Figure 2.2. Phenotypes of cht7. (A) Immunoblot of MLDP in the PL (dw15) and cht7 mutant
following N resupply (NR) at times indicated (hours). (B) TAG degradation in dw15 and cht7
following N resupply. (C) Confocal microscopy images (Top) and lipid droplet quantification
(Bottom) of Nile red-stained dw15 and cht7 cells following N resupply. Chlorophyll
fluorescence (C) and Nile red fluorescence of lipid droplets (LD); scale bar, 5 μm. Lipid droplets
of 5–20 cells, depending on the sample, were counted and their area was quantified with imaging
software and their volume calculated. No lipid droplets were observed in dw15 cells at NR24.
SD is indicated.

CHT7 Encodes a CXC Domain DNA Binding Protein Present in the Nucleus
The cht7 mutant was crossed to a line of the opposite mating type. Abundance of TAG in meiotic
progeny following N resupply cosegregated with the antibiotic marker, suggesting that a single
nuclear mutation was responsible for the lipid phenotype (Figure 2.3A). DNA/DNA
hybridization blots confirmed the presence of a single insertion (Figure 2.3B). With SiteFindingPCR (Tan et al., 2005), the flanking sequences on both ends of the inserted hygromycin B
marker were mapped and a 18,087-bp deletion affecting four predicted genes was discovered
(Figure 2.4A). Complementation of the defect was accomplished following transformation with a
genomic fragment containing gene 1 bracketed by 1,000 bp on either side (Figure 2.3 C and D).
Expression of this gene in the complemented lines was confirmed by the presence of the CHT7

41

protein (Figure 2.5B). Therefore, loss of gene 1 (Cre11.g481800, C. reinhardtii genome v5.3.1;
CHT7) is the cause for the delayed TAG degradation following N resupply in cht7.
The predicted CHT7 protein contains two cysteine-rich motifs comprising CXC domains
(Pfam 03638) (Punta et al., 2012), initially defined in human tesmin (Sugihara et al., 1999) and
Arabidopsis TSO1 (Hauser et al., 2000) (Figure 2.4B). TSO1 and other CXC proteins, e.g.,
soybean CPP1 (Cvitanich et al., 2000), human LIN54 (Schmit et al., 2009), and Caenorhabditis
elegans Lin54 (Tabuchi et al., 2011), have been shown to bind zinc (Andersen et al., 2007) and
specific DNA sequences through their CXC domains. To determine whether CHT7 is located in
the nucleus, I added a C-terminal green fluorescent protein (GFP) to CHT7, thereby increasing
its size by 30 kDa, and introduced it into the cht7 background (Figure 2.5B). Phenotypes of cht7
were rescued in CHT7-GFP:cht7 transgenic lines (Figure 2.5C), indicating that CHT7-GFP is
functional and present in its correct location. The nucleus was visualized using the DNA-binding
dye Hoechst 33342 (Figure 2.4C, yellow arrows; Figure 2.5A). Aside from strong chlorophyll
fluorescence delineating the chloroplast, in N-replete CHT7- GFP:cht7 transgenic lines I
observed only GFP-specific signals associated with the nucleus (Figure 2.4C, white arrows).
After 48 h of N deprivation, the cells were lysed when exposed to the Hoechst dye. Therefore, I
examined the location of CHT7 in N-deprived cells without nuclear staining. GFP signal still
was observed in the nucleus (Figure 2.4C). Because of the ambiguities related to chlorophyll
fluorescence, I used cell fractionation in combination with markers and confirmed the nuclear
location of CHT7 and its absence from chloroplasts (Figure 2.4D).

42

Figure 2.3. Meiotic progeny phenotyping and cht7 complementation. (A) TAG content of
three complete meiotic tetrads (1–11, 2–5, 2–7) after 24 h N resupply. Hr and Hs indicate
hygromycin B-resistant and -sensitive lines, respectively. The ratio of fatty acids (FA) in TAGs
over total FAs in the lipid extracts is shown. (B) Southern blot probed against the pHyg3
insertion indicates single insertion in the genome of cht7. cht1, cht2 and cht3, mutants isolated in
the same mutant screen; dw15, the parental line; un, unrelated mutant. (C) Genetic background
of complementation lines. (Top) Amplicon targeting genomic fragment of gene 1 (refer to Fig.
2A). (Bottom) Amplicon targeting the aph7 gene of the pHyg3 insertion. C1–4, complemented
cht7 mutant lines; EV, empty vector control. (D) Restoration of TAG degradation in four
independent transgenic cht7 lines carrying a genomic wild-type copy of gene 1. Cells were N
deprived for 48 h, followed by 24 h N resupply. Averages (n = 3) and SDs are given.

43

Figure 2.4. CHT7 is a CXC domain-containing protein present in the nucleus. (A) Structure
of the cht7 genomic locus with gene 1 corresponding to CHT7. Black boxes, exons; white boxes,
untranslated regions; arrow points, 3'-ends of ORFs. (B) Different CXC-domain proteins from
different species: At, Arabidopsis thaliana; Ce, Caenorhabditis elegans; Cr, C. reinhardtii; Dm,
Drosophila melanogaster; Gm, Glycine max; Hs, Homo sapiens. Black boxes, CXC domains.
(C) Nuclear localization of CHT7-GFP by confocal microscopy in N-replete (+N) or N-deprived
(−N) cells. The PL (dw15) and a transgenic line expressing EGFP-tagged CHT7 under the
regulation of the endogenous CHT7 promoter in the cht7 mutant background (CHT7-GFP:cht7)
are shown. The nuclear CHT7-GFP signal is marked by white arrows. The nuclei were stained
with Hoechst 33342 (marked by yellow arrows). (D) Enrichment of CHT7 in nuclear extracts
shown by immunoblot. Five micrograms of protein was loaded into each well. Markers:
chloroplast (CP), OE33; nucleus (N), histone 3 (H3). CBB, Coomassie Brilliant Blue; WCL,
whole-cell lysate.

44

Figure 2.5. Nuclear localization of CHT7. (A) Confocal images showing the nuclear
localization of CHT7 in N replete (+N) or N deprived (−N) cells. Fluorescence was detected in
the parental line (dw15) and a transgenic line expressing EGFP-tagged CHT7 under the
regulation of the endogenous CHT7 promoter in the cht7 mutant background (CHT7-GFP:cht7).
The nuclear CHT7-GFP signal is marked by white arrows. The nuclei were stained with Hoechst
33342 (marked by yellow arrows). (B) CHT7 protein levels at midlog growth detected by
immunoblot using CHT7 antiserum (α-CHT7). Equal total protein was loaded as shown by
Coomassie Brilliant Blue (CBB) staining. G1 and G2, two independent lines of CHT7-GFP:cht7.
(C) Restoration of the growth phenotype of two cht7 lines producing a CHT7-GFP fusion. Cells
were N deprived for 48 h, followed by 24 h of N resupply.

45

Absence of CHT7 Affects the Exit from Quiescence but Not Cell Viability
Growth of the cht7 mutant was normal in N-replete medium under standard conditions (Figure
2.6A). However, when cht7 cells in liquid cultures had been deprived of N, which then was
resupplied, growth was severely delayed (Figure 2.6B). This delay in growth and the delay in
TAG degradation following N resupply (Figure 2.2B) suggested that the cht7 mutant struggles to
reverse quiescence either because of its inability to correctly perceive the N status of cells during
resupply of N, because cht7 cells simply lose viability following N deprivation, or because of a
general defect in the regulation of quiescence. To rule out the possibility of a specific N-sensing
defect in cht7, I tested induction of quiescence by phosphate deprivation (Figure 2.6C and Figure
2.7A). The cht7 mutant also showed delayed regrowth when phosphate was resupplied to a
deprived culture, suggesting a more general defect than N sensing. In yeast and many other
organisms, the nutrient status of the cell to a large extent is integrated with progression through
the cell cycle by the TOR (target of rapamycin) signaling pathway (Gray et al., 2004). Typically,
rapamycin treatment induces quiescence, and I tested the effect of its removal in cht7. A
saturating concentration of rapamycin (1 μM) as established for C. reinhardtii (Perez-Perez et al.,
2010) doubled the time needed for division of the PL (23.9 h) and slightly more so for cht7 (30.1
h) (Figure 2.7B). A striking difference was seen when rapamycin was removed: the cht7 mutant
was much slower to regrow, similar to the effect of nutrient resupply (Figure 2.6D and Figure
2.7C). Thus, the defect in cht7 affects the ability of cells to exit quiescence, regardless of how it
is induced.

46

Figure 2.6. Growth of cht7 affected by different treatments. (A) Growth of cht7 and PL
(dw15) on N-replete medium (OD at 550 nm). Doubling times are given in parentheses. The gray
bar indicates the OD at which N-replete samples were taken for all experiments in this study.
Averages of three biological replicates are given, with SDs for all points being smaller than 3%.
(B) Growth of dw15, cht7, and four independent complementation lines (C1–4) during N
deprivation (−N) followed by N resupply (+N) measured at the times indicted. (Inset) Cultures at
24 h following N resupply. (C) Growth during phosphate deprivation (−P) followed by P
resupply (+P). (D) Growth in the medium with 1 μM rapamycin (+R) followed by the removal of
rapamycin (−R). Cultures were diluted twice (a) with medium containing 1 μM rapamycin to
avoid reaching stationary phase.

47

Figure 2.7. Growth, RNA, and protein of cht7 affected by different treatments. (A) Cultures
of dw15, cht7, and four complementation lines (C1–4) at 48 h following P resupply. (B) Cultures
after 120 h of rapamycin treatment. (C) Cultures at 48 h following the removal of rapamycin. (D
and E) RNA (D) and protein (E) contents of dw15 and cht7 cells in the presence (+N) and
absence (−N) of N or following N resupply (NR) at the times (hours) indicated. For all
quantitative data, averages (n = 3) and SD are indicated.

One trivial explanation for the growth phenotype is that most cht7 cells cannot survive N
deprivation. The number and integrity of cells were assessed by using a hemocytometer and
SYTOX Green, which does not penetrate living cells but stains the nuclear DNA of nonviable or
partially lysed cells (Sato et al., 2004). Parallel cultures of PL and cht7 had a similar number of
intact cells (Figure 2.8A), and cells were similarly viable (Figure 2.8B) during the 5 d following
N deprivation. Moreover, following N deprivation, cht7 cells changed their metabolism to
accumulate TAG (Figure 2.2, NR0) and they decreased RNA and protein synthesis (Figure 2.7 D
and E). In addition, cht7 cells increased RNA and protein synthesis following N resupply in
parallel to the PL (Figure 2.7 D and E). It should also be noted that cht7 cells were capable of
normal mating following N deprivation. Therefore, the mutant had no defect in gametogenesis.
Thus, I concluded that cht7 cells had not lysed, were viable, and were metabolically active and
48

mating competent following N deprivation. However, when plated on agar-solidified N-replete
medium at different times of N deprivation, the efficiency of cht7 colony formation (observed 11
d after plating and again after an additional 14 d with no further increase in numbers) decreased
during the first 24 h following N deprivation to ∼20%, compared with 80% for the PL (Figure

2.8C), and remained there. Presumably, during the first 24 h as cht7 cells become increasingly
N-deprived, they enter quiescence, after which only 20% of the cells pass an apparent threshold
or checkpoint, allowing for colony formation. These cht7 colonies had a decreased diameter
(Figure 2.8D) consistent with a delay in resumption of cell division. When the cht7 colonies
were transferred to N-replete liquid medium, they grew at a rate similar to that of PL but again
showed a delay in regrowth after N deprivation followed by resupply, recapitulating the original
cht7 phenotype (Figure 2.8C, Inset). Together, these observations are consistent with a
regulatory defect in cht7 preventing most cht7 cells from orderly progression out of quiescence
to resume normal growth, even though they are viable and metabolically active during N
deprivation.

49

Figure 2.8. Viability and colony formation of cht7 during N deprivation. (A) Cell
concentration of PL dw15 and cht7 during N deprivation (−N) at times (h) indicated, determined
by using a hemocytometer. The averages of three to six measurements and SDs are indicated. (B)
SYTOX Green viability staining of dw15 and cht7 cells shown in A at times indicated.
Approximately 10–20 images showing 5–10 cells each were examined and averaged per time
point. SD between images is indicated. (C) Colony formation of N-deprived (−N) dw15 and cht7
plated on N-replete solid medium at times indicated and incubated for 11 d. The average
percentage of plated cells forming a colony on three plates is given. SD is indicated. (Inset)
Three liquid cultures derived from cht7 colonies from this experiment (−N48), the original cht7
mutant, and dw15 PL first grown in N-replete medium then N deprived, followed by N resupply
and growth for 24 h, recapitulating the original phenotype. (D) Size of the colonies formed by
the N-deprived cells after plating on N-replete medium. All colonies on one plate were analyzed,
with n ranging from 6 to 70 (usually fewer for cht7). Mean, maximum, minimum, and quartile
values are indicated. Data points in all panels (A–D) were obtained in the same experiment.

50

Absence of CHT7 Partially Derepresses Quiescence-Associated Transcriptional Programs
Because CHT7 resembles known DNA binding proteins, I asked whether a change in global
transcriptional profiles during or even before entering quiescence might explain the observed
phenotypes of cht7. Global transcript profiles of cht7 and the PL were compared by RNA
sequencing (RNA-Seq) during midlog phase of an N-replete culture and after 48 h of N
deprivation [Figure 2.10A; for complete data sets and enriched Gene Ontology (GO) categories,
refer to Online Supplemental Data Sets 1 and 2]. To confirm the findings obtained by RNA-Seq
and to test for effects specific to the loss of CHT7, the expression of selected genes was tested by
quantitative PCR (qPCR) in the PL, cht7, and multiple complementation lines. The expression of
selected genes observed by RNA-Seq (three independent biological repeats) was comparable to
that measured by qPCR (correlation coefficient R2 = 0.8065; Figure 2.10B).
In line with previously reported transcriptional changes for C. reinhardtii (Miller et al.,
2010; Blaby et al., 2013; Schmollinger et al., 2014) following N deprivation, 2,647 genes were
upregulated and 3,346 down-regulated in PL (dw15) N-deprived cells compared with PL Nreplete cells (Figure 2.9A, blue circles; twofold cutoff, P < 0.05). Comparing cht7 N-replete with
PL N-replete cells, 1,477 genes were found up-regulated and 1,491 down-regulated in cht7
(Figure 2.9A, yellow circles). Most strikingly, there was a substantial overlap in genes upregulated (573) and down-regulated (894) between the two comparisons (Figure 2.9A,
intersecting blue and yellow circles; Online Supplemental Data Set 3). In other words, a subset
of genes, i.e., 49% of all genes that were misregulated in cht7 during N-replete conditions, were
expressed as if the cells had already entered quiescence. Examples are genes involved in
photosynthesis, such as PSBS1, MCA1, NAB1, and LHCBM4, and genes related to flagellum
assembly, such as FA1, FA2, FAP139, and IFT46 (Figure 2.9B; Figure 2.10 C–E for qPCR

51

confirmation). Autophagy is a hallmark of quiescence, and autophagy markers APG8
(Cre16.g689650) (Perez-Perez et al., 2010; Klionsky et al., 2012) and APG3 (Cre02. g102350)
were constitutively expressed in cht7, although at lower levels than in the PL following N
deprivation. To further explore this pattern of transcriptional alterations in N-replete cht7, I
asked whether these misregulated genes represent meaningful biological functions. In the PL Ndeprived versus PL N-replete comparison (Figure 2.9A, blue circles), differentially expressed
genes were enriched in 68 GO categories, with 18 GO categories associated with flagellum
assembly and 21 with photosynthesis (Online Supplemental Data Set 2). Differentially expressed
genes in the cht7 N-replete versus PL N-replete comparison (Figure 2.9A, yellow circles) fell
into six enriched GO categories, one of which was associated with flagellum assembly and two
with photosynthesis. Indeed, virtually every gene involved in photosynthesis and 171 of 320
genes associated with flagellum assembly tended to be differentially regulated in the same
manner in both comparisons (Figure 2.10 A, F, and G), but not necessarily to the same extent. In
most cases, differential gene expression was less pronounced in the cht7 N-replete versus PL Nreplete comparison than in the PL N-deprived versus PL N-replete comparison (Figure 2.9B;
Figure 2.10 A, F, and G; and Online Supplemental Data Set 3).

52

Figure 2.9. Global gene expression comparison of N-replete and -deprived cells of PL dw15
and cht7. (A) Large circles: total number of genes changed in expression in a comparison of
dw15 N deprived for 48 h and dw15 N replete (never deprived of N). Small circles: total number
of genes changed in expression in a comparison of the cht7 mutant N replete and dw15 N replete.
Circle size is proportional to the number of genes. (B) RNA-Seq log2 (fold change) of
representative genes encoding the indicated proteins related to photosynthesis (PS), flagellum
(FL), or autophagy (AP).

53

Figure 2.10. Confirmation of transcriptional changes by qPCR and analysis of
photosynthesis and flagellum-related genes. (A) Scatter plot of all genes in the RNASeq
comparisons (dw15 N-deprived/dw15 N-replete against cht7 N-replete/dw15 N-replete). Genes
involved in photosynthesis are shown in red, those involved in flagellum assembly in green. (B)
Comparison of fold change in abundance of transcripts for the comparison cht7 N-replete and
dw15 N-replete obtained by quantification of RNA levels by qPCR analysis (x-axis) or RNA-Seq
54

Figure 2.10 (cont’d)
experiments (y-axis). (C) qPCR analysis of autophagy marker genes (APG, autophagy-related
protein; dw15, parental line; cht7 mutant; C1–4, complemented cht7 mutant lines) in N-replete
medium. Averages and SD are indicated. At least two biological replicates were included in the
analysis. (D and E) qPCR analysis of selected photosynthetic genes (D) and flagellum-related
genes (E) in N-replete medium. Genes down-regulated in cht7 are shown in the left graph, genes
up-regulated in cht7 in the right graph. Averages and SD are indicated. At least two biological
replicates were included in the analysis. (F and G) Cluster analysis showing transcript profiles of
photosynthetic genes (F) and flagellum-related genes (G). The set of genes was clustered
hierarchically into groups on the basis of the similarity of their expression profiles. The
expression pattern of each gene is displayed here as a heat map according to the color scale at the
bottom. Heat map beneath “1” represents the ratio of mRNA levels in N-deprived dw15 to Nreplete dw15. Heat map beneath “2” represents the ratio of mRNA levels in N-replete cht7 to Nreplete dw15. Included here are all relevant genes, including those that fall below the twofold
threshold.

CHT7 Levels Remain Constant in Response to the N Supply, and CHT7 Is in a Large
Complex
One may hypothesize that to exert its effects on gene expression related to quiescence, CHT7
abundance changes in response to the N supply. However, immunoblotting indicated that CHT7
protein abundance (see Figure 2.12A for purity of CHT7 antiserum) was relatively constant
during the conditions tested, which included N deprivation and several time points following
resupply of N (Figure 2.11A). Using blue native (BN) gel electrophoresis, I also determined that
CHT7 is part of a larger protein complex that does not change in apparent size or abundance
following N deprivation (Figure 2.11B; see Figure 2.12B for 2D electrophoresis).

55

Figure 2.11. Abundance of CHT7 and hypothesis for CHT7 function. (A) Anti-CHT7
immunoblot showing CHT7 protein levels in the presence (+N) and absence of N (−N) or after N
resupply (NR) at times (hours) indicated. Equal amounts of protein were loaded. (B)
Immunodetection of a high molecular weight CHT7 complex by BN-PAGE. Whole-cell lysates
were loaded at equal protein, and cht7 was used as a negative control to discriminate against
nonspecific signals. dw15, PL of cht7; +N, N replete; –N, N deprived. (C) We hypothesize that
CHT7 acts as a repressor in safeguarding against the premature activation of global
transcriptional changes associated with quiescence (Q) during N-replete growth and also to fully
revert quiescence after N resupply. Activators of quiescence are postulated to fully turn on
quiescence following N deprivation, but their inactivation after N resupply is insufficient in the
absence of CHT7 to restore growth.

Figure 2.12. Immunodetection of CHT7 and 2D BN-SDS-PAGE. (A) Immunoblot of wholecell lysates from parental line dw15 or cht7 mutant probed with CHT7 antibodies (α-CHT7). The
migration of CHT7 at its predicted molecular weight is indicated by an asterisk. Nonspecific
binding to other proteins recognized by the antibodies is shown but did not interfere with specific
56

Figure 2.12 (cont’d)
CHT7 detection, as the CHT7 signal is missing in cht7 extracts. (B) Cell lysates of N-replete
parental line dw15 were analyzed in the first dimension by BN-PAGE (1D BN-PAGE, marker at
top) and then denatured and run on SDS-PAGE in the second dimension (2D SDS-PAGE,
marker at left) followed by anti-CHT7 immunoblot. CHT7 is indicated by the arrow.

57

DISCUSSION
Regulatory proteins that participate in the integration of the metabolic status of the cell with cell
cycle activity are of fundamental biological importance and also are potential targets to
maximize algal biomass and its TAG content by engineering. The unicellular alga C. reinhardtii
provides an excellent genetic model for a photosynthetic eukaryotic cell, in which nutrient status
may be manipulated readily to induce and reverse cellular quiescence. Hence, I isolated a mutant,
cht7, affected in the reversal of N deprivation-induced quiescence, i.e., the degradation of lipid
droplet protein MLDP and TAG, and the regrowth of algal mutant cultures when N is resupplied.
I have ruled out several trivial explanations for this phenotype, including loss of viability of the
mutant during N deprivation or a specific deficiency in an N-signaling pathway. The delay in
regrowth in response to phosphate refeeding and after removal of rapamycin-induced quiescence
of cht7 (Figure 2.6 C and D) points to a more general defect in cht7 in the integration of the
nutrient status of the cell with the cell cycle. CHT7 likely is localized in the nucleus and contains
a putative CXC DNA binding motif (Figure 2.4) consistent with its possible role as a regulator of
gene expression. Thus, I hypothesized and subsequently showed that CHT7 affects
transcriptional programs associated with N deprivation-induced quiescence. A subset of genes
normally up- or down-regulated is misregulated in cht7 under N-replete conditions, consistent
with a partial derepression of transcriptional programs characteristic for quiescence. The extent
of these changes in the expression of individual genes, as well as the number of genes affected, is
smaller than observed following full induction of quiescence in response to N deprivation in the
PL, which may explain the apparent lack of a growth phenotype of cht7 under the N-replete
conditions tested (Figure 2.6A). Thus, although quiescence programs are fully off under Nreplete conditions in the PL, they are partially on in cht7, as summarized in the model in Figure

58

2.11C. Based on this observation, CHT7 appears to act as a repressor of a subfraction of the
transcriptional program associated with quiescence. During N deprivation, activators likely come
into play to establish full quiescence to the same extent in cht7 and the PL, as they behave
similarly: for example, accumulating TAG and ceasing to divide without loss of viability.
However, the delayed growth of cht7 following N resupply suggests that CHT7 is needed to turn
off quiescence-associated programs to reestablish growth rapidly (Figure 2.11C). One may
postulate that any number of repressors and activators of quiescence have to be balanced out
during quiescence exit and that the absence of CHT7 in the mutant shifts this balance. Assuming
the involvement of multiple inputs and regulatory components besides CHT7 to govern
quiescence, the apparent threshold phenomenon documented in the ability of ∼20% of N-

deprived cht7 cells to “escape deep quiescence” (Figure 2.8C) seems plausible.

The delay in regrowth of the cht7 mutant when resupplied with N following deprivation
is similar to the phenotype of the mat3 mutant (Armbrust et al., 1995). MAT3 is a C. reinhardtii
ortholog of the mammalian retinoblastoma tumor suppressor protein (Rb) (Sadasivam and
DeCaprio, 2013). Both CHT7 and Rb/MAT3 are present in the C. reinhardtii nucleus throughout
the cell cycle (Olson et al., 2010). However, the absence of Rb/MAT3 leads to drastically
reduced cell size, an essential cue in C. reinhardtii for decisions made during cell cycle
progression, whereas cht7 cell size is normal (Figure 2.4C and Figure 2.5A). In general, Rb
interacts directly with DNA-binding proteins, such as members of the E2F and DP protein
families, to repress genes required for cell cycle progression during quiescence (Olson et al.,
2010; Sadasivam and DeCaprio, 2013). CXC domain proteins in animals have been found in
large multiprotein complexes involved in transcriptional regulation that also contain Rb
(Sadasivam and DeCaprio, 2013). Thus, the fact that CHT7 is a CXC domain protein present in a

59

large complex (Figure 2.11B) leads to intriguing questions regarding the precise regulatory
function of CHT7, such as whether CHT7 and Rb/MAT3 might cooperate in the regulation of
quiescence.
The persistence of CHT7 before, during, and after quiescence (Figure 2.11A) suggests
that fluctuation of its abundance likely is not part of the regulatory mechanism determining entry
and exit into and out of quiescence. Changes in abundance or size of the CHT7 complex during
quiescence also were not observed (Figure 2.11B). Movement of CHT7 in and out of the nucleus
also seems an unlikely mechanism to modulate CHT7, as I observed the protein in the nucleus
before and during N deprivation (Figure 2.4C). However, I currently cannot rule out
posttranslational modifications of CHT7 depending on the nutritional status of the cell that may
modulate its possible DNA binding preferences.
As summarized in Figure 2.11C, all current data point toward a role of CHT7 as a
repressor of a subset of transcriptional programs associated with nutrient deprivation-induced
quiescence. Its activity likely is balanced by other regulatory factors, such as activators of
quiescence. Its loss causes partial derepression of quiescence-associated transcriptional programs
during N-replete conditions; CHT7 apparently is not needed for full establishment of quiescence
during N deprivation, but its absence prevents the orderly and rapid exit from quiescence
following N refeeding. As such, CHT7 is a candidate regulatory factor involved in the
integration of the metabolic status of the cell and cell division, and its discovery provides a
unique entry point for a more in-depth study of the regulation of cellular quiescence and the
discovery of additional factors involved.

60

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CHAPTER 3
Comparative Transcriptomic Analysis of Nitrogen Resupply-induced Modifications in a
Chlamydomonas Quiescence-exit Mutant

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ABSTRACT
In response to environmental cues such as nutrient deprivation, cells can temporary cease
divisions and enter a reversible state of quiescence. The regulatory processes by which
photosynthetic organisms preserve the reversibility are largely unknown. Therefore, I used
Chlamydomonas reinhardtii as a reference model and conducted comparative transcriptomics of
cht7, a mutant slow to resume growth from quiescence. My data uncover transcriptional
programs unique to the exit of quiescence induced by N deprivation, and the CHT7 regulon
including tetrapyrrole pathway and peroxisomal redox homeostasis, which potentially interplays
with the cAMP-PKA pathway. Focusing on lipid metabolism for detailed analysis, I found that
the metabolite state is closely associated with the way transcripts are regulated. And in some
cases, e.g. monogalactosyldiacylglycerol and fatty acid syntheses, the transcription is subjected
to a feedback control by metabolite levels. These data provide rich resources for future
investigation of any biological process in the context of cellular quiescence.

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INTRODUCTION
The ability of cells to temporarily halt divisions is essential for survival during adverse
conditions, particularly for unicellular organisms, and for the maintenance of tissue homeostasis
in multicellular organisms. This non-dividing state is called quiescence, and is distinguished
from senescence or terminal differentiation by its reversibility (Gray et al., 2004; Coller, 2011).
Yeast offers a well-studied example of quiescence for unicellular eukaryotes: upon reaching
stationary phase, yeast cells cease proliferation due to the exhaustion of the main carbon source
or specific nutrients such as nitrogen (N) (Gasch et al., 2000). Growth typically resumes in fresh
medium. The reversibility of quiescence in mammalian cells often requires cues other than the
availability of nutrients. Examples are T lymphocytes that become activated to mount an immune
response when exposed to antigens, or dermal fibroblasts that are activated during wound healing
(Valcourt et al., 2012). In Chlamydomonas reinhardtii, a unicellular microalga on which this
study is based, N deprivation-induced cell cycle arrest has all the hallmarks of quiescence
including reversibility, change in metabolism, reduced transcription and translation, reduced
synthesis of rRNA and ribosomal proteins, and activation of autophagy (Miller et al., 2010;
Goodson et al., 2011; Schmollinger et al., 2014). In facing quiescence, a plethora of metabolic
adjustments has to take place to meet the changing demands (Gray et al., 2004; Valcourt et al.,
2012). For example, because quiescent cells cannot dilute out reactive oxygen species (ROS)
and replace damaged protein or other macromolecules by rapid synthesis, they require
specialized ROS-dissipating mechanisms to maintain redox homeostasis, such as catalase and
superoxide dismutase. Autophagy in most conditions is reduced to a very low level, but
drastically elevated during quiescence, which allows degradation and recycling of cellular
components. For photosynthetic organisms, there is an additional challenge, that is, to quench

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energy passing through photosynthetic electron transport chain in a way that it can be restored
momentarily when conditions improve. Beyond transcriptional and translational modifications,
such as down-regulation of photosynthetic genes or degradation of light-harvesting complexes,
lipid metabolism is also an important contributor. Shortly following N deprivation, acetate
typically provided in the medium under laboratory conditions to Chlamydomonas is no longer
incorporated into biomass through the glyoxylate cycle and gluconeogenesis; instead acetate is
directly funneled into biosynthesis of fatty acids and triacylglycerols (TAGs) (Miller et al., 2010;
Boyle et al., 2012). This process relieves the overreduction of photosynthetic electron transport
chain, which otherwise produces harmful ROS and subsequent cell death (Li et al., 2012b).
Meanwhile, chloroplast thylakoid lipids, especially monogalactosyldiacylglycerol (MGDG),
undergo major turnover hence contributing to the dismantling of the structural base of the
photosynthetic membrane.
The discovery of CHT7 in Chlamydomonas expands the landscape of quiescence
research. It demonstrates that the exit of quiescence is a highly controlled process, not just a
passive response to nutrient availability (Figure 3.1) (Tsai et al., 2014). I am thus interested to
understand the transcriptional modifications in response to quiescence exit, and ask whether
programmed responses unique to quiescence exit exist. In addition, while it is evident that CHT7
acts as a repressor of cellular quiescence, and is hypothesized to ensure the reversibility by
resuppressing quiescence, I wondered whether CHT7 has additional, more direct impacts on
quiescence exit. Therefore in this study, I undertook comparative transcriptomics spanning the
entire quiescence cycle of cht7 and its parental line (PL). Although I place an emphasis on lipid
metabolism when describing the data, the overall datasets provide rich resources for exploration
in any biological process in the context of quiescence.

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Figure 3.1. Cell Cycle and Proposed Functions for CHT7 in Quiescence. The cell division
cycle with G1, S, G2, and M phases, and the quiescence cycle (G0) are depicted. CHT7 is
proposed to play distinct roles at quiescence entry (- repression) and exit (+ stimulation) as
discussed in the text. -N, N deprivation, +N, N resupply.

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MATERIALS AND METHODS
Strains and Growth Conditions
Chlamydomonas reinhardtii cell wall-less strain dw15 (cw15, nit1, mt+) was obtained from A.
Grossman and is referred to as the wild-type (with regard to CHT7) parental line (PL) throughout.
21gr (wild-type mt+; the parental line of mat3-4) was obtained from the Chlamydomonas
Resource Center (http://www.chlamycollection.org). Complemented strains of cht7 and the
MLDP RNAi lines were generated as described in previous work (Moellering and Benning, 2010;
Tsai et al., 2014). Cells were grown in Tris-acetate-phosphate (TAP) medium (Harris, 1989),
under continuous light (70 to 80 μmol m-2 s-1) at 22°C or ambient room temperature (~22°C) for
solid media, which contained 0.8% agar (Phytoblend (Caisson Labs), North Logan, UT). To
induce N deprivation, mid-log phase cells grown in TAP were collected by centrifugation (1500
g, 4°C, 2 min), washed twice with TAP-N (NH4Cl omitted from TAP), and resuspended in TAPN at 0.3 OD550. N resupply was done by refeeding 1% culture volume of 1 M NH4Cl (100x) to
the N-deprived culture. In some experiments, Protein Kinase A Inhibitor (P9115, Sigma-Aldrich)
was added to the culture along with N resupply. The cell concentration of cultures was
determined using a Z2 Coulter Counter (Beckman).

Illumina RNA Sequencing and Bioinformatics
As described in (Tsai et al., 2014).

Metabolite Measurements
Chlorophylls were extracted from fresh cell pellets using 80% acetone, and concentrations were
calculated from the absorbance values at 647 and 664 nm according to (Zieger and Egle, 1965).

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For the TBARS assay 5 mL of culture was centrifuged and analyzed immediately. Cell pellets
were resuspended in 1 mL of thiobarbituric acid/trichloroacetic acid solution (0.3 and 3.9%
respectively) and heated at 95°C for 15 min. The solution alone was also heated to serve as the
blank for spectrophotometric measurements. Samples and blank were measured after no further
gas bubbles were released. TBARS were determined by absorbance at 532 and 600 nm as
previously described (Baroli et al., 2003). The extinction coefficient used was 155 mM-1 cm-1.
Quantification of cellular cyclic AMP (cAMP) was conducted using Cyclic AMP Competitive
ELISA kit (Thermo Scientific) according to manufacturer instruction. The concentration of cells
in all assays was monitored by Z2 Coulter Counter.

Fatty Acid and Lipid Analysis
For lipid analysis extraction, TLC of neutral and polar lipids, fatty acid methyl ester (FAME)
preparation, and gas–liquid chromatography were conducted as previously described in
(Moellering and Benning, 2010). Cell pellets were extracted into methanol and chloroform (2:1
vol/vol, for neutral lipids) or methanol, chloroform, and 88% formic acid (2:1:0.1 vol/vol/vol, for
polar lipids). To the extract, 0.5 volume of 0.9% KCL (neutral lipids) or 1 M KCl and 0.2 M
H3PO4 (polar lipids) was added and mixed, followed by phase separation at low-speed
centrifugation. For TAG quantification, lipids were resolved by TLC on Silica G60 plates (EMD
Chemicals, Gibbstown, NJ) developed in petroleum ether-diethyl ether-acetic acid (80:20:1 by
volume). Polar lipids were separated on the same plate using chloroform-methanol-acetic aciddistilled water (75:13:9:3 by volume) as solvent. After visualization by brief iodine staining,
FAME of each lipid or total cellular lipid was processed and quantified by gas chromatography
as previously described (Rossak et al., 1997). Staining with α-naphtol and Dragendorff reagent

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was used for the detection of galactoglycerolipids and diacylglyceryl-trimethylhomoserine
(DGTS) (Benning et al., 1995).

Standard RNA Techniques
To produce cDNA templates used for qPCR, total RNA was purified using the RNeasy plant
mini kit (Qiagen) including the on-column DNase digestion, and then subjected to reverse
transcription using Superscript III reverse transcriptase (Life Technologies). qPCR was
performed on an ABI Prism 7000 (Applied Biosystems, Grand Island, NY) using the SYBR
green PCR master mix (Life Technologies) with an equivalent cDNA template and 0.25 μM of
each primer. The amount of cDNA input was optimized after serial dilutions. In all qPCR
experiments, expression of the target gene was normalized to the endogenous reference gene
CBLP, a gene commonly used for normalization in C. reinhardtii (Allen et al., 2007), using the
cycle threshold (CT) 2-ΔΔCT method. All experiments were done using at least two biological
replicates and each reaction was run with technical replicates. qPCR procedures and analysis
followed the MIQE guidelines (Bustin et al., 2009). For quantification, Trizol reagent (Life
Technologies) was used for RNA isolation and the concentration of RNA was measured using a
NanoDrop instrument (Thermo Scientific).

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RESULTS
Transcriptomes of Parental Line during different N regimes
Previously, we conducted RNA-Seq experiments on the cht7 mutant and the parental line (PL)
grown in N-replete (mid-log phase of a culture in standard Tris-acetate phosphate (TAP) medium)
and N-deprived (48 h in TAP lacking N) conditions, and found that the transcriptional program
characteristic for quiescence was partially induced in the absence of CHT7, suggesting CHT7 as
a repressor of cellular quiescence (Tsai et al., 2014). Here I show the global transcript profiles of
cht7 and PL in two additional conditions of N availability: 6 and 12 h of N resupply following 48
h of N deprivation (designated NR6 and NR12, respectively; Figure 3.2). These time points were
chosen to reflect the metabolic and physiological contexts. For instance, both the regeneration of
chlorophyll and turnover of TAGs restarted at NR6 (see below), and at NR12 cell division
resumed. Importantly, the NR6 and NR12 RNAs were harvested, prepared, and sequenced in
parallel with the samples for the N-replete and N-deprived data sets and, thus, can be directly
compared. In each of the experiments, three replicates were taken independently to allow
statistically sound analyses.

Figure 3.2. Experimental Design of RNA-Seq Experiments. Parental line (dw15) and cht7
cells grown to mid-log phase in N-replete medium were sampled (N-replete) before being
deprived of N (-N); cells were sampled after 48 h following N deprivation (N-deprived) and
resupplied with N (+N) and sampled at 6 (NR6) and 12 h (NR12).

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One key objective of this investigation was to understand the molecular changes that
facilitate cells to exit and regrow from the N deprivation-induced quiescence. Towards this goal,
transcripts of the NR6 PL or NR12 PL were compared with those of the N-deprived PL (for
complete data set see Online Supplemental Data Set 4). Previously we showed that when PL
cells transited from N-replete to N-deprived condition, 2647 genes were up-regulated and 3346
down-regulated (Tsai et al., 2014). We found that in response to N resupply these genes reversed
their expression in a time-dependent manner and can be categorized according to three different
stages, i.e. early, mid, and late (Figure 3.3A). Taking the genes for which expression was upregulated during N deprivation as an example, 1405 of them reversed at least two-fold (log2 ≤ -1
with a p value of < 0.05) at NR6 and were defined as early-reverse (1309 of the 1405 retained at
least two-fold reversion at NR12); another 519 genes reversed not until NR12 and were defined
as mid-reverse; 723 genes had not reversed by NR12 and were assumed to reverse at later time
and were defined as late-reverse. Likewise, 1208 (1136 retained at least two-fold reversion at
NR12), 991 and 1147 genes of those down-regulated during N deprivation are defined as early,
mid and late-reverse, respectively (Online Supplemental Data Set 5). Selected metabolic
pathways are discussed in detail in the later result sections.

Figure 3.3. Summary Scheme of Transcriptomic Analyses in the Parental Line. (A) Early,
Mid, and Late-reverse gene groups in the parental line. Numbers represent transcript changes by
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Figure 3.3 (cont’d)
the N status using a 2-fold cutoff (log2 = 1) and a p value of < 0.05. +N, N-replete; -N, Ndeprived; NR6 and NR12, 6 and 12 h of N resupply. Dashed line means the numbers shown are
indirectly based on extrapolation of s , the RNA-Seq data. (B) NR-specific gene groups in the
parental line. Numbers represent the transcripts whose abundance did not vary in the –N over +N
RNA-Seq comparison (-1 < log2 < 1), but changed over 2-fold with a p value of < 0.05) in the
NR over +N and NR over –N comparison.

The regrowth delay of cht7 suggests the existence of a specific exit program that governs
the quiescence exit. I therefore asked whether there are transcriptional responses specific to N
resupply (NR-specific). To be classified as a NR-specific gene, its relative RNA abundance must
not vary when shifting from N-replete to N-deprived condition (less than two-fold), but is either
up or down-regulated in both the NR versus N-replete and the NR versus N-deprived
transcriptomes (using a 2-fold cutoff and a p value of < 0.05, Figure 3.3B). It should be noted
that NR-specific regulation means the transcripts are differentially regulated only when exposed
to N resupply, but this does not necessarily mean they are only expressed during the period of N
resupply. Given these criteria, 852 (NR6-specific) and 347 (NR12-specific) genes were
specifically up-regulated after 6 and 12 h of N resupply, respectively, among which 197 genes
overlapped. For genes specifically down-regulated during N resupply, 547 (NR6-specific) and
411 (NR12-specific), 224 overlapped (Online Supplemental Data Set 6). The events of NRspecific regulation were more frequent soon after N was refed (NR6). The overlaps (197 and 224)
were judged to be the most robust set of NR-specific genes. Selected metabolic pathways are
discussed in detail in the later result sections. It is important to note that the NR-reverse and NRspecific gene groups are mutually exclusive, enabling me to better dissect the responses in the
phase of quiescence exit.

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Comparative Transcriptomics of cht7 and the Parental Line following N-resupply
CHT7 encodes a putative transcription factor and without it, quiescent cells show delayed
regrowth. It would not be unexpected to see differences in transcript abundance between cells
that undergo orderly progression and cells that fail to do so. Therefore, to distill meaningful
dysregulation in cht7 when PL cells would be exiting quiescence, I adopted the strategy
described previously for similar purposes (Gonzalez-Ballester et al., 2010; Castruita et al., 2011).
Transcripts of the PL after 6 h and 12 h of N resupply were compared with those of N-deprived
PL (Figure 3.4 A and B, blue circles); transcripts of the PL after 6 h and 12 h of N resupply were
compared with those from cht7 (Figure 3.4 A and B, yellow circles). It is useful to know that the
blue circles in Figure 3.4A contain all the early-reverse and NR6-specific genes mentioned above
and the blue circles in Figure 3.4B contain all the mid-reverse and NR12-specific genes. While
most transcripts responsive to N resupply in the PL were readjusted normally in cht7 (Figure 3.4
A and B, blue circles outside the overlap), a specific group did not respond in cht7 and remained
at expression levels that were similar to those of N-deprived PL cells (Figure 3.4 A and B,
overlap between blue and yellow circles; See Online Supplemental Data Set 7 for complete
information). I hypothesize that these overlapping genes constitute the core of the CHT7 regulon
during quiescence exit that is required for resuming cell growth, whereas the changes in
expression of other genes (yellow circles outside the overlap) are secondary or compensatory
effects due to the impaired growth resulting from the loss of CHT7. Moreover, 446 of the 1293
NR6 overlapping genes (594 plus 699) and 164 of the 1176 NR12 overlapping genes (639 plus
537) are NR-specific (Online Supplemental Data Set 7), arguing that CHT7 impacts quiescence
exit in ways that are distinct from ordinary cell proliferation.

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Figure 3.4. Comparative Transcriptomics during Quiescence Exit. (A, B) Global gene
expression analysis of the parental line (dw15) and cht7 following N resupply. Large blue circles:
total number of genes changed in expression in a comparison of dw15 after 48 h of N deprivation
followed by 6 h (NR6) or 12 h (NR12) of N resupply and dw15 N-deprived for 48 h. Small
circles: total number of genes changed in expression in a comparison of dw15 NR6 and cht7
NR6 and dw15 NR12 and cht7 NR12.

Closer examination of the full data sets provided striking examples in support of this
hypothesis, namely genes encoding enzymes involved in the tetrapyrrole pathway (chlorophyll
synthesis, in the NR6 and NR12 overlaps) and peroxisomal redox homeostasis (mainly in the
NR12 overlaps). At 6 and 12 h following N resupply, nearly every gene of tetrapyrrole
biosynthesis tended to show lower transcript levels in cht7 compared to PL following N resupply
(Figure 3.5A, included here were genes that fall below the 2-fold threshold; Online Supplemental
Data Set 8). These genes did not greatly differ in expression between cht7 and the PL in Nreplete condition; hence, I conclude that they are specifically misregulated in cht7 in response to
the signals that should cause the cells to exit quiescence. The RNA-Seq data were confirmed for
representative genes by quantitative PCR (qPCR) including cht7 complemented lines in this
analysis (Figure 3.6A). These changes in transcript abundance were corroborated at the
metabolite level. Shortly after 6 h following N resupply, the cellular chlorophyll content of the
PL and complementation lines increased; it decreased after 12 h, likely because cells were
dividing (Figure 3.5C). In contrast, the chlorophyll cell content of cht7 did not increase during
the entire observation period. Genes encoding enzymes maintaining redox homeostasis during
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high demand of fatty acid β-oxidation were down-regulated in cht7 following N resupply
(Figures 3.5B and 3.6B; Online Supplemental Data Set 8). Among these were catalase (CAT1
and CAT2), ascorbate peroxidase (APX1 and APX2) and monodehydroascorbate reductase
(MDAR1) which are responsible for detoxifying ROS generated within peroxisomes as a
byproduct of β-oxidation (Eastmond, 2007). Consistently, thiobarbituric acid reactive substances
(TBARS), the cellular metabolites reflecting damage caused by ROS, accumulated in cht7
following N resupply (Figure 3.5D). Intriguingly, Arabidopsis homologues of HPR1, MAS1,
MDH2, MDH4, MDAR1 and PXN1 (all misregulated in NR12 cht7) have defects in seed oil
breakdown and seedling establishment, reminiscent of the cht7 mutant phenotypes (Graham,
2008; Theodoulou and Eastmond, 2012).

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Figure 3.5. Changes in Gene Expression and Metabolite of Tetrapyrrole Pathway and
Peroxisomal Redox Homeostasis. (A, B) Overview of expression of genes involved in the
tetrapyrrole pathway (A) and in peroxisomal redox homeostasis (B). RNA-Seq comparisons of
dw15 at different N status and the comparisons between dw15 and cht7 at each N status are
shown in the heatmap. +N, N-replete; -N, N-deprived; NR6 and NR12, 6 and 12 h of N resupply.
Early, Mid, Late, NR6 or NR12 is indicated on the left of the heatmap for the gene whose
response to N resupply has been classified. (C) Left panel: total cellular chlorophyll content. C1C4 individual complemented lines. Right panel: cellular TBARS content. For all quantitative
data, averages (n=3) and standard deviation are indicated.

85

Figure 3.6. Confirmation of Transcript Changes at Quiescence Exit. (A, B, C) qPCR analysis
of selected transcripts of genes encoding enzymes of the tetrapyrrole pathway at 6 h of N
resupply (A), of genes involved in peroxisomal redox homeostasis at 12 h of N resupply (B), and
of genes involved in fatty acid synthesis at 12 h of N resupply (C) (dw15, parental line; cht7
mutant; C1-4, complemented cht7 mutant lines).

Metabolite Analyses Confirm the Effect of Transcriptional Changes on Lipid Metabolism
Upon N deprivation, changes in TAG metabolism are reflected in the way genes in lipid
metabolic pathways are regulated (Miller et al., 2010; Blaby et al., 2013; Schmollinger et al.,
2014). Genes encoding 3-ketoacyl-ACP synthase (KAS1), stearoyl-ACP-Δ9-desaturase
(FAB2) – the key enzyme for the signature TAG fatty acid oleate (18:1Δ9), acyl-ACPthioesterase (FAT1) and long-chain acyl-CoA synthetase (LACS1) were upregulated, implying

86

elevated export of de novo synthesized fatty acids from chloroplast to where TAGs are made.
Phosphatidate phosphatase (PAP), which increases the DAG pool through the Kennedy pathway,
and the putative DAG acyltransferase (e.g. DGTT1) also showed increased transcript levels. All
of the above genes reversed their expression in response to N resupply (Online Supplemental
Data Set 9). Interestingly, the two long-chain acyl-CoA synthetase isoforms (LACS1 and LACS2)
behaved just in the opposite. LACS2 transcripts decreased when N-deprived and increased when
N was resupplied, which paralleled the need of β-oxidation, and thus likely to catalyze the
activation of fatty acids for resulting from TAG breakdown for β-oxidation. In contrast, some
modifications only occur upon N resupply, that is, NR-specific. Betaine lipid synthase (BTA1)
synthesizes the diacylglyceryl-trimethylhomoserine (DGTS), a major lipid component presumed
to take the role of PC in extraplastidic membranes in Chlamydomonas (Riekhof et al., 2005).
Transcript levels of BTA1 remained constant when entering N deprivation, but increased
approximately 4 fold after 6 h and 12 h of N resupply, a typical NR-specific gene. DGTT4 is one
of the five putative DAG acyltransferases; however its association with TAG production has
been unclear. I found a near 4-fold increase of DGTT4 RNAs after 6 h and 12 h of N resupply –
the time when TAGs were being hydrolyzed – which could suggest a need to detoxify excessive
free fatty acids by redirecting some back to TAGs, or that DGTT4 was to transfer acyl groups
from TAG to other lipids (e.g. MGDG).
N status also has an impact on membrane lipids. The relative fraction of the thylakoid
lipid MGDG to total membrane lipids in PL was reduced following N deprivation, and restored
following N resupply (Figure 3.7C). Digalactosyldiacylglycerol (DGDG) on the other hand, was
more abundant when N is absent (Figure 3.7C) in accord with the upregulation of DGDG
synthase (DGD1). The increase of DGTS following N deprivation was small but statistically

87

significant; however unlike DGDG which gradually returned to its initial level when the
condition was reversed, the elevated amount of DGTS was retained throughout the observation
period (Figure 3.7C), consistent with the NR-specific upregulation of BTA1 mRNA levels. The
significance of the increase in DGDG and DGTS when TAGs are built is perhaps related to LD
formation as discussed later. No statistically significant changes were observed for
phosphatidylethanolamine (PE), phosphatidylglycerol (PG), phosphatidylinositol (PI) and
sulfoquinovosyldiacylglycerol (SQDG) (Figure 3.8A).

Figure 3.7. Changes in Gene Expression and Metabolite Levels Related to MGDG and
Fatty Acid Syntheses. (A, B) Overview of expression of genes expression involved in MGDG
(A) and in fatty acid (B) syntheses. RNA-Seq comparisons of dw15 at different N status and the
comparisons between dw15 and cht7 at each N status are shown in the heatmap. Arrows indicate
the sequence of reactants and products of the catalyzed reactions for each enzyme encoded by
the gene as indicated. +N, N-replete; -N, N-deprived; NR6 and NR12, 6 and 12 h of N resupply.
18:1/16:0 indicates the fatty acid (FA) at the sn-1/sn-2 position of DAG (diacylglycerol) or
MGDG (monogalactosyldiacylglycerol). (C) Polar lipid content (depicted as the ratio of each
polar lipid FAs over total polar lipid FAs) in the presence (+N, N-replete) or absence (-N, Ndeprived) of N or following N resupply at times indicated (h). DGDG, digalactosyldiacylglycerol;
DGTS, diacylglyceryl-trimethylhomoserine. Averages (n=4) and standard deviation are indicated.
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Figure 3.7 (cont’d)
(D) Per cell FA content for total lipid and TAG following N resupply in dw15. Averages (n=3)
and standard deviation are indicated.

Figure 3.8. Detailed Lipid Analysis at Quiescence Exit. (A) Quantification of polar lipids in
dw15 and cht7 grown at different N status (+N, N-replete; -N, N-deprived; NR, N-resupplied) for
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Figure 3.8 (cont’d)
the times (h) indicated, including phosphatidylethanolamine (PE), phosphatidylglycerol (PG),
sulfoquinovosyldiacylglycerol (SQDG) and phosphatidylinositol (PI). The y-axis is depicted as
ratio of each polar lipid fatty acids (FAs) over total polar FAs. (B) Relative fatty acid
composition of dw15 and cht7 following N resupply (NR) at times indicated (h). (C, D) Relative
fatty acid composition of dw15 of total lipids (C) and of TAGs (D) following N resupply at times
indicated (h).
Fatty acid profiles of total lipids remained almost unchanged in cht7 after N resupply,
while in the PL, 16:4 and 18:3ω3 fatty acids increased over time in parallel to increases in
MGDG content (Figure 3.8B). 16:4 and 18:3ω3 are the major fatty acids of the MGDG (Giroud
et al., 1988). Not surprisingly, among all the membrane lipids cht7 was not able to readjust
MGDG in response to N resupply (Figures 3.7C and 3.8A). I found that MGD1 encoding the
MGDG synthase, three genes for MGDG palmitate-delta7-desaturase (Cre09.g397250, g4555
and g4570), and virtually every gene involved in fatty acid synthesis were misregulated in cht7
(Figures 3.7 A-B and 3.6C). Nevertheless, these genes might not be under the direct control of
CHT7 since they were not present in the NR6 or NR12 overlap groups (Figure 3.4 A and B;
Online Supplemental Data Sets 7 and 9), except for HAD1, KAR1 and MCT1 but only in NR12.
TAG degradation started soon after 6 h of N resupply in almost a linear fashion; however, the
total lipid content per cell did not vary noticeably by 12 h of N resupply (Figure 3.7D),
implicating that the fatty acids removed from TAGs were not sent to peroxisomes for β-oxidation,
but probably to the chloroplast for rebuilding of MGDG, the predominant lipid of photosynthetic
membranes. Indeed, 16:4 and 18:3ω3 fatty acids were not being degraded after N is resupplied
and had been stored in TAGs during N deprivation (Figure 3.8 C and D). Therefore we
hypothesize that the transcription of MGDG and fatty acid synthesis is subjected to a feedback
control by metabolite levels. In cht7 following N resupply, the low 16:4 and 18:3ω3 levels (as
free fatty acids or in DAGs) and the high overall fatty acid levels (in TAGs) created a negative
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feedback that does not allow upregulation of genes for MGDG and fatty acid syntheses,
respectively. Moreover, MGDG desaturation was also part of the NR-specific responses, as the
genes encoding the enzymes that catalyze the second and third double bonds on the sn-1 and sn-2
position of MGDG were especially up-regulated following N resupply (Cre06.g288650 and
Cre01.g038600, Figure 3.7A; Online Supplemental Data Set 9), suggesting that the demand for
MGDG in exiting quiescence was higher than that for ordinary growth.

Two Classes of Lipases Affect TAG Accumulation in opposite ways
Lipases are a subclass of acyl hydrolases that deesterify carboxylic esters. Traditionally, they are
referred to as class 3 lipases when involved in the hydrolysis of TAG, with a serine protease triad
forming the catalytic center (Brady et al., 1990; Winkler et al., 1990). Since it has become clear
that during N deprivation membrane lipid turnover is an alternative route for fatty acids
channeled into TAGs (Li et al., 2012b), lipases can actually affect TAG accumulation in opposite
ways. However, it is a challenge to distinguish them. There are 131 proteins predicted to be
lipases, phospholipases, or patatin-like proteins based on the GXSXG motif common to
hydrolases (Online Supplemental Data Set 8), and biochemical validation often comes with
surprises. PGD1, the galactoglyceride lipase responsible for half of the TAG production, was
purported to be a TAG lipase based on in silico annotation (Li et al., 2012b). Although LIP1 is a
class 3 lipase, it can act on polar lipids in addition to its involvement in TAG turnover (Li et al.,
2012a). Through integrated pairwise comparison, I was able to filter and classify lipases
according to the gene expression patterns. As TAGs increase during N deprivation and decrease
following N resupply, I expected genes encoding lipases involved in TAG turnover (e.g. LIP1) to
be down-regulated during N deprivation (Figure 3.9, -N/+N), and up-regulated following N

91

resupply (Figure 3.9, NR6/-N or NR12/-N), and genes encoding lipases assisting TAG
production (e.g. PGD1) to behave in the opposite way. Thus I narrowed down the number of
potential functional lipases to 9 (TAG turnover, Figure 3.9 upper panel) and 23 (TAG production,
Figure 3.9 middle panel), including LIP1 and PGD1 in their expected class. We also included 2
(TAG turnover) and 6 (TAG production) NR-specific lipase candidates whose abundance might
be critical for TAGs when N is again available. Transcript profiling of candidate genes for
peroxisomal β-oxidation (e.g. 3-oxoacyl-CoA thiolase (ATO1) and acyl-CoA oxidase) is in tune
with TAG turnover under these conditions, with the exception of enoyl-CoA oxidase/isomerase
(ECH1) which is particularly important for degradation of unsaturated fatty acids (Goepfert et al.,
2008). A primary phenotype of cht7 is the compromised hydrolysis of TAGs, after which the
mutant is named. After 6 h of N resupply, among the genes of two classes of lipases,
Cre17.g707300, Cre06.g265850, Cre03.g195200 and Cre03.g152800 (possibly TAG-reducing)
and PGD1, g9707 and Cre03.g174900 (possibly TAG-enhancing) were misregulated in cht7 and
behaved in a way that would hinder the degradation and promote the accumulation of TAGs,
making them promising candidates for reverse genetic studies (Figure 3.9 and Online
Supplemental Data Set 8). These seven genes are also among the NR6 overlaps mentioned above.

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Figure 3.9. Changes in Gene Expression of Putative Lipases and β-oxidation. Transcript
profiles of genes encoding lipases potentially involved in the degradation of TAGs (upper panel)
or in the production TAGs (middle panel), and of the genes involved in β-oxidation (lower panel).
RNA-Seq comparisons of dw15 between at N status and the comparisons between dw15 and cht7
at each N status are shown in the heatmap. +N, N-replete; -N, N-deprived; NR6 and NR12, 6 and
12 h of N resupply. Early, Mid, Late, NR6 or NR12 is indicated on the left of the heatmap for the
gene whose response to N resupply has been classified.

The cAMP-Dependent Protein Kinase A Pathway Is Involved in Quiescence Exit
I also explored signaling cascades that may impact TAG metabolism. In adipocytes, adenylyl
cyclase and protein kinase A (PKA) act as transducers between hormone signals and lipolytic
responses (Guo et al., 2009). Upon activation, adenylyl cyclase converts ATP to cyclic AMP
93

(cAMP) and binding of cAMP to the regulatory domains of PKA activates the enzyme and leads
to the phosphorylation of the respective targets, e.g. hormone sensitive lipase and perilipin.
Adenylyl and guanylyl cyclases form one of the largest enzyme families in the Chlamydomonas
genome (Merchant et al., 2007), and discrimination based solely on primary sequences is
difficult. At a glance, many of the adenylyl cyclase candidates were differentially regulated by N
status and in the cht7 background (Online Supplemental Data Set 10). A cAMP specific ELISA
assay showed that concentrations of cAMP positively correlate with the activation of TAG
breakdown (Figure 3.10A). I therefore intended to clarify the involvement of PKA in the
lipolysis in Chlamydomonas. A 20-amino acid fragment of the naturally occurring protein kinase
inhibitor (PKI) has been shown to bind to the catalytic domain of PKA (Knighton et al., 1991). A
derivative of this fragment has been used in the study of protein kinases in flagellar assembly in
Chlamydomonas (Howard et al., 1994). When applied simultaneously with N refeeding, PKI
blocked lipolysis in the PL in a dosage-dependent manner (Figure 3.10B). 10 μM of PKI caused
chlorosis indicative of cell death, thus none of the TAGs in the PL or cht7 being degraded even
at 24 h of N resupply. Fatty acid profiles of cells treated with PKI resembled those of cht7
(Figure 3.11). Importantly, within the nontoxic range the TAG contents of cht7 were not affected
by PKI. PKI treatment also mimicked the regrowth defect of cht7 in the PL (Figure 3.10C).
Phosphodiesterases are the enzyme counteracting adenylyl cyclase by breaking cAMP into AMP.
Stimulating the activity of phosphodiesterase attenuates cAMP-mediated lipolysis (Botion and
Green, 1999). We curated potential adenylyl cyclases and phosphodiesterases whose expression
profile matched the fluctuation of cAMP (Figure 3.10D; Online Supplemental Data Set 10), with
the intent to provide possible candidates for reverse genetic studies, such as Cre13.g607150,
Cre05.g237800, Cre02.g080550 and g11603 considering their regulation in cht7. The remainder

94

of the enzymes (in Online Supplemental Data Set 10 but not listed in Figure 3.10A) might be
responsible for cyclic GMP signaling.

Figure 3.10. Possible Role o cAMP-PKA Signaling in Quiescence Exit and Transcript
Profiles of Related Genes. (A) ELISA assay to quantify the cellular content of cAMP in dw15.
+N, N-replete; -N, N-deprived; NR, N resupply at times indicated (h). Asterisks indicate a
statistically significant difference (unpaired t-test, p < 0.05). (B) Efficacy of TAG degradation in
the presence of the inhibitor PKI . TAG contents (depicted as ratio of TAG fatty acids (FA) over
total FAs) right before N resupply (NR) are shown under NR0 (equivalent to –N, N-deprived for
48 h). TAG contents after 24 h of N resupply with and without PKI treatment are shown under
NR24. (C) Regrowth during PKI treatment is shown as fold increase by comparing the cell
number counted at 24 h of N resupply to that counted right before N resupply with and without
PKI treatment. (D) Transcript profiles of genes encoding putative adenylyl cyclase and
phosphodiesterase. RNA-Seq comparisons of dw15 at different N status and the comparisons
95

Figure 3.10 (cont’d)
between dw15 and cht7 at each N status are shown in the heatmap. +N, N-replete; -N, Ndeprived; NR6 and NR12, 6 and 12 h of N resupply. Early, Mid, Late, NR6 or NR12 is indicated
on the left of the heatmap for the gene whose response to N resupply has been classified. For all
quantitative data, averages (n=3) and standard deviation are indicated.

Figure 3.11. Fatty Acid Profiles of PKI-Treated Samples. Relative fatty acid composition of
total lipids (upper panel) and of TAGs (lower panel) in dw15. After 48 h of N deprivation, cells
were sampled right before N resupply (NR0) and at 24 h of N resupply with and without PKI
treatment (μM).

96

DISCUSSION
N deprivation is as an effective approach to induce TAG accumulation in microalgae, attracting
interest from public and private sectors due to the potential in creating a renewable energy source
(Hu et al., 2008). From a biology standpoint, the reversible cessation of growth provides a facile
experimental system to study cellular quiescence. Chlamydomonas offers perspectives beyond
yeast in studying quiescence because, apart from the unique features related to photosynthesis,
the underlying regulatory processes in this green alga utilize a Retinoblastoma (RB) tumor
suppressor homologue and CXC domain proteins such as CHT7 and thus more closely resemble
regulation in mammalian cells (Armbrust et al., 1995; Burkhart and Sage, 2008; Sadasivam and
DeCaprio, 2013). Yeast cells possess neither RB nor CXC domain proteins. Cellular responses to
N deprivation have been studied on multiple omic levels, including transcripts, proteins, and
metabolites, drawing an integrated picture of N sparing mechanisms (Miller et al., 2010; Blaby et
al., 2013; Schmollinger et al., 2014; Wase et al., 2014). In contrast, the research on quiescence
exit or the means that triggers it (i.e. N resupply) has not yet been studied; in part due to the
misconception that quiescence exit might simply be the reverse of entry. In this work, global
transcriptomic analysis uncovered a subset of genes whose expression only fluctuated during N
resupply (e.g. BTA1 and DGTT4), as a first entry point to explore the unique aspects of
quiescence exit. Metabolite levels are usually lower in quiescent cells than newly divided ones
because of a higher catabolic and lower anabolic rate, for example, chlorophylls (Gray et al.,
2004; Schmollinger et al., 2014). It is natural to imagine the regrowth out of quiescence demands
more than ordinary cell proliferation does. NR-specific genes were dispersed in diverse pathways
with genes that reverted expression pattern in response to N resupply. I discussed many

97

examples, such as BTA1 in DGTS synthesis and MGDG desaturation, to show that NR-specific
responses are important to boost the rapid resumption of growth.

The CHT7 regulon and possible influence of cAMP-PKA signaling
NR overlaps (Figure 3.4 A and B) highlight programs of genes most likely responsible for—not
only responsive to—quiescence exit and at the same time under the control of CHT7. I
hypothesize that when cells receive signals to exit quiescence, CHT7 governs these programs
that affect metabolic pathways necessary for the resumption of growth. The discussed examples
of misregulated pathways, i.e., tetrapyrrole pathway and peroxisomal redox homeostasis,
represent just a fraction of genes affected in their expression by CHT7. Particularly important is
that these metabolic pathways misregulated in cht7 during N resupply were normal during Nreplete growth. As such CHT7 may play distinct roles in repressing transcriptional programs
prior to quiescence entry and in adjusting transcriptional networks during the exit from
quiescence. Whether the control of CHT7 on these genes is direct or indirect remains to be
answered through further analysis, e.g. chromatin immunoprecipitation followed by sequencing
of fragments bound to CH7 or its complex.
My finding of cAMP-PKA signaling being potentially involved in lipolysis and
quiescence exit in Chlamydomonas was a logical prediction based on the current knowledge in
adipocytes and in yeast quiescence cells. However, none of the two classic examples of PKA
phosphorylation in lipolysis, hormone sensitive lipase and perilipin, has apparent homologues
encoded in the C. reinhardtii genome (Yeaman, 1990; Marcinkiewicz et al., 2006). Even so it
does not rule out the existence of unknown PKA targets that can modulate lipolysis. Another
possibility is that the cAMP-PKA signaling pathway controls upstream aspects of quiescence,

98

and the inhibition of TAG turnover by PKI treatment is a secondary effect. It drew my attention
when PKI treatment did not exacerbate the TAG phenotype in cht7, which strongly suggests that
the CHT7 and cAMP-PKA pathways converge at some point, at least in the regulation of TAG
turnover. In yeast, cAMP-PKA singling, like CHT7 in Chlamydomonas, prevents the entry into
quiescence. Mutants of adenylyl cyclase fail to proliferate and display phenotypes superficially
similar to quiescence (van Aelst et al., 1991). On the contrary, constitutive activation of PKA
causes premature cell death when entering quiescence, indicative of the necessity to deactivate
the negative regulators of quiescence to ensure a proper entry (Uno et al., 1984; Shin et al., 1987).
Downregulation of CHT7 activity in quiescent cells might as well be necessary in
Chlamydomonas. In addition, reactivation of the cAMP-PKA pathway is critical for successful
exit from quiescence in yeast. Yeast mutants unable to raise a transient cAMP level when fresh
nutrients are again available show extended delay in resuming growth (Jiang et al., 1998).
Pharmacological and biochemical approaches have uncovered the existence of PKA catalytic
subunits in Chlamydomonas even though respective genes are still missing (Howard et al., 1994).
Future studies of a possible CHT7-cAMP-PKA interplay may shed light on the complex network
underlying quiescence control. At present, whether the Chlamydomonas cAMP-PKA pathway
directly controls lipolysis, which in turn affects quiescence exit or vise versa remains unclear, but
either way it points to unique opportunities to unravel questions concerning LD biology and
cellular quiescence.

99

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CHAPTER 4
The Role of Lipid Droplet in Coordinating Lipid Dynamics to Ensure the Reversibility of
Cellular Quiescence in Chlamydomonas1

_____________________________
1

I contributed to the data shown in all of the figures. Krzysztof Zienkiewicz also made significant
contribution to Figures 4.4C, 4,7 D, E, and F.
ABSTRACT

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ABSTRACT
Lipid droplets (LDs) are unique among organelles for their hydrophobic core which allows the
storage of neutral lipids. They are also known as hubs for lipid metabolism and temporary
shelters for protein under stress conditions. The fate of LD is closely tied to the state of
quiescence in unicellular organisms. However, whether or how LDs influence quiescence is
largely unknown. Here, I provide evidence that LDs coordinate critical responses for quiescence.
These comprise a possible deacylation/reacylation cycle that exchanges the polyunsaturated acyl
groups of LD surface lipids with de novo synthesized saturated ones; providing temporary shelter
for refugee proteins; and being in close proximity to the chloroplast outer envelope to allow
protein and lipid trafficking between the two organelles during quiescence exit. The mature LD
requires stabilization by a proteinaceous coating. Failure to maintain an optimized surface areato-volume ratio of LD compromises the efficacy of triacylglycerol turnover, and disturbs the
timely progression out of quiescence.

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INTRODUCTION
TAGs synthesized during quiescence are stored in lipid droplets (LDs), the dynamic organelles
found ubiquitously across species and cell types. Long perceived as inert, lipid droplets are now
known to have many cellular functions (Martin and Parton, 2006; Walther and Farese, 2012).
Such recognition comes from LD proteomes and biochemical evidence. The localization of a
long chain acyl-CoA synthetase (ACSL3), two phosphocholine cytidylyltransferase isoforms
(CCT1 and CCT2), and a diacylglycerol acyltransferase (DGAT2) to LDs in animals suggests
local synthesis of phosphotidylcholine (PC) and TAG needed for LD growth (Fujimoto et al.,
2007; Guo et al., 2008; Kuerschner et al., 2008). Activities of ACSL3 and CCT1 associated with
LDs are experimentally verified (Fujimoto et al., 2007; Krahmer et al., 2011). Proteomic studies
in Chlamydomonas also portray LDs as a hub for lipid synthesis, acyl exchange, lipid signaling
and trafficking (Moellering and Benning, 2010; Nguyen et al., 2011). Under stress conditions
(e.g. heat shock or endoplasmic reticulum (ER) stress) or certain developmental stages (e.g.
Drosophila embryos or viral infection), LDs recruit seemingly unrelated proteins from other
cellular compartments, hypothetically for temporary sheltering and to proteins sequestered when
not needed, or as a way to transport proteins (Welte, 2007). It is a common belief that LDs in
eukaryotes emerge from the ER. However, recent studies in Chlamydomonas proposed that the
ER and chloroplast are both involved in LD biogenesis. Evidence shows that LDs are invariably
cytosolic but snuggled into the folds of the chloroplast envelope, forming numerous punctate
associations with the outer envelope, and that nearly all the diacylglycerols (DAGs) acylated into
TAGs in LDs are derived from the chloroplast (Fan et al., 2011; Goodson et al., 2011).
The mutant phenotype of cht7 shows that LD formation as well as its degradation is
tightly linked to the state of quiescence (Tsai et al., 2014). However, whether or how LDs

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contribute to the entry, maintenance, and exit of quiescence has not yet been addressed. In this
study, I conducted functional analysis of LDs using biochemical, genetic, microscopic and
proteomic approaches. The detailed characterization of LDs may transform our thinking about
mechanisms responsible for the reversibility of quiescence.

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MATERIALS AND METHODS
Strains and Growth Conditions
As described in Chapter 2.

Fatty Acid and Lipid Analysis
As described in Chapter 2.

Lipid Droplet Isolation, HDN-PAGE, and Co-immunoprecipitation
Cells grown in 400 mL of N-deprived or 800 mL of N-resupplied cultures were spun down and
resuspended in 10 mL of isolation buffer (50 mM HEPES pH 7.5, 5 mM KCl, 5 mM MgCl2, 0.5
mM sucrose, 1 mM phenylmethylsulfonylfluoride (PMSF), 1 mM 2,2’-dipyridyl, and 1X
protease inhibitor cocktail Roche), and lysed by sonication on ice with 10 s on/off cycle for 2
min. The homogenate was overlaid with 25 mL of floating buffer 1 (isolation buffer with only
0.1 M sucrose) and centrifuged in swing bucket at 23,000 g for 30 min. Lipid droplets floating on
top of the overlay buffer were collected using a Potter S homogenizer. Lipid droplets were then
solubilized in 200 μL of floating buffer 2 (isolation buffer without sucrose), and mixed 1:1 with
2X HDN sample buffer (200 mM Tris-HCl pH 8.0, 1 M 6-aminocapronic acid, 20 % glycerol, 10
mM dithiothreitol, 2 µM PMSF) supplemented with 0.5 or 1% n-Dodecyl-β-D-maltopyranoside
(DDM). The mixture was incubated on ice for 20 min followed by centrifugation at 20,000g for
10 min at 4°C. The aqueous phase was carefully transferred to a new tube to avoid the
contamination from the lipid layer on top and the precipitates, and then run on 4-14% histidine
deoxycholate native gels as described (Kikuchi et al., 2006; Ladig et al., 2011). Electrophoresis
was done at 50 V for an hour, and then switched to 30 V overnight at 4°C. The gels were

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denatured and either subjected directly to immunobloting, or run in a second dimension on
denaturing SDS-PAGE before immunobloting. In all cases of the study, the RC DC Protein
Assay Kit (Bio-Rad) was adopted for protein quantification.
For immunoblot analysis SDS-PAGE gels were blotted onto polyvinylidene fluoride
(PVDF) membranes in transfer buffer (25 mM Tris, 192 mM Gly, and 10% methanol) for 60 min
at 100V. Membranes were blocked for 60 min in TBST (50 mM Tris, 150 mM NaCl, 0.05 %
(v/v) Tween 20, pH 7.6) with 5% nonfat dry milk, and probed with primary antibodies overnight
at 4°C. The primary antibodies were used at 1:1000 dilutions, except for mouse anti-αtubulin
(1:10,000). Goat anti-mouse or anti-rabbit secondary antibodies coupled to horseradish
peroxidase were used at 1:10000 dilution and incubated with blots at room temperature for 30
min. Blots were then washed six times with TBST at room temperature for 10 min each. Antigen
was detected by chemiluminescence (Bio-Rad Clarity Western ECL substrate) using a chargecoupled device (CCD) imaging system. Anti-MLDP was generated as described (Tsai et al.,
2014). Anti-NAB1 was a gift from Krishna K. Niyogi, Department of Plant and Microbial
Biology, Berkeley, CA. To generate antibodies against Chlamydomonas TGD2, the full length
coding sequence of TGD2 was amplified using total cDNA as the template, and inserted into the
expression vector pET28B(+) (Novagen). Construct was introduced into E. coli BL21 (DE3).
Recombinant proteins (6xHis-TGD2) were purified using Ni-NTA agarose (Qiagen) and
separated by SDS-PAGE to examine the purity. Few other proteins were co-purified with 6xHis
MLDP. Roughly 2 mg of each protein was sent for antibody production in rabbits by Cocalico
Biologicals, Inc.
For Co-immunoprecipitation (CoIP) experiments, 1.2 L N-deprived or 2.4 L N-resupplied
cultures were centrifuged. Cell pellets were washed twice in phosphate-buffered saline pH 7.2

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(PBS), resuspended in PBS with protease inhibitor (P9599, Sigma-Aldrich) and phosphatase
inhibitor (Thermo Scientific) at a concentration of 109 cells/mL, and then crosslinked on ice with
1 mM freshly prepared DSP (Dithiobissuccinimidyl propionate, Pierce). To quench the crosslinking, 1 M Tris-HCL pH 7.5 was added to a final concentration of 100 mM and incubated on
ice for 15 min. Cells were then snap frozen in liquid nitrogen and stored at -80°C until use. Lipid
droplets were isolated as described above, but instead of dissolving in the floating buffer 2, they
were washed in 500 μL of washing buffer (same as isolation buffer but with 150 mM KCl). The
suspension was overlaid with 1 mL of floating buffer 1 and centrifuged at 20,000 g for 15 min at
4°C. The washing was repeated three times until no green pellet was seen after centrifugation.
LDs were then solubilized in 400 μL of floating buffer added with 1% DDM. After incubating on
ice for 20 min, insoluble materials and the lipid layer were separated by centrifugation at 20,000
g for 10 min at 4°C. The supernatant was divided into two portions, one incubated with 100 μL
Dynabeads Protein A (Life Technologies) coupled with 10 μg of purified MLDP IgG, and the
other with Dynabeads coupled with MLDP prebleed as negative control. Incubation was carried
out overnight at 4°C with gentle rotation. Dynabeads were collected magnetically and washed
with PBS supplemented with 100 mM NaCl and 1 mM PMSF. The washing was repeated three
times and finished by an additional wash using PBS only. Proteins were eluted with nonreducing sodium dodecyl sulfate (SDS) sample buffer by incubating at room temperature for 15
min. Before being processed for mass spectrometry, DSP was cleaved by boiling in the presence
of 5% β-mercaptoethanol. CoIP elutes were separated by SDS-PAGE but shortly after the
samples had entered the gel, the gel bands were excised. Gel bands were digested in-gel
according to (Shevchenko et al., 1996) with modifications. Briefly, gel bands were dehydrated
using 100% acetonitrile and incubated with 10 mM dithiothreitol in 100 mM ammonium

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bicarbonate, pH~8, at 56°C for 45min, dehydrated again and incubated in the dark with 50 mM
iodoacetamide in 100 mM ammonium bicarbonate for 20min. Gel bands were then washed with
ammonium bicarbonate and dehydrated again. Sequencing grade modified Trypsin was prepared
to 0.01ug/μL in 50mM ammonium bicarbonate and ~50 μL of this was added to each gel band so
that the gel was completely submerged. Bands were then incubated at 37°C overnight. Peptides
were extracted from the gel by water bath sonication in a solution of 60% ACN/1% TCA and
vacuum dried to ~2 μL. Peptides were then re-suspended in 2% acetonitrile/0.1% TFA to 25 μL.
From this, 5 μL were automatically injected by a Thermo (www.thermo.com) EASYnLC 1000
onto a Thermo Acclaim PepMap RSLC 0.075mm x 150mm C18 column and eluted over 120min
with a gradient of 2% B to 30% B in 109 min, ramping to 100% B at 110 min and held at 100%
B for the duration of the run (Buffer A = 99.9% Water/0.1% Formic Acid, Buffer B = 99.9%
Acetonitrile/0.1% Formic Acid). Sample injection order was randomized to avoid bias. Eluted
peptides were sprayed into a ThermoFisher Q-Exactive mass spectrometer (www.thermo.com)
using a FlexSpray spray ion source. Survey scans were taken in the Orbi trap (35000 resolution,
determined at m/z 200) and the top ten ions in each survey scan are then subjected to automatic
higher energy collision induced dissociation (HCD) with fragment spectra acquired at 17,500
resolution. The resulting MS/MS spectra are converted to peak lists using Mascot Distiller,
v2.4.3.3 (www.matrixscience.com) and searched against all protein entries in the C. reinhardtii
v5.3.1, protein database (downloaded from www.phytozome.net) and appended with common
laboratory contaminants (downloaded from www.thegpm.org, cRAP project) using the Mascot
searching algorithm, v 2.4. The Mascot output was then analyzed using Scaffold Q+S, v4.3.0
(www.proteomesoftware.com) to probabilistically validate protein identifications. Assignments
validated using the Scaffold 1% FDR filter (protein threshold) and 95% confidence filter

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(peptide threshold) are considered true. Variable modifications of Oxidation of Methionine and
of DSP cleavable x-linker were adjusted accordingly.

Immunofluorescence, Confocal, and Transmission Electronic Microscopy
For immunofluorescence, cells were fixed in a mixture of 4% (w/v) paraformaldehyde and 0.15
M sucrose in PBS (pH 7.4) for 1 h at room temperature. After two washes in phosphate-buffered
saline (PBS) buffer, cells were permeabilized by incubation in 0.01% Triton-X100 in PBS for 5
min. at room temperature and washed twice in PBS. Samples of 200 μl of volume were
transferred into sterile Eppendorrf tubes and blocked with a solution containing 1% (w/v) BSA in

PBS (pH 7.2) for 1 h. Then, samples were incubated with primary antibodies: 1) mouse antiαtubulin (T6074, Sigma-Aldrich) diluted 1:50 in PBS buffer, pH 7.2, containing 1% (w/v) BSA
or 2) rabbit anti-MLDP diluted 1:50 in PBS buffer, pH 7.2, containing 1% (w/v) BSA or 3)
mixture of both antibodies, overnight at 4 °C on rotary shaker. Anti-mouse IgG conjugated with
Alexa Fluor 405 (Molecular Probes) or anti-rabbit IgG conjugated with Alexa Fluor 488
(Molecular Probes) or a mixture of both diluted 1:200 in PBS buffer (pH 7.2) were used as the
secondary antibodies. In order to visualize LDs, just before examination, aliquots (100 μl) of cell
suspension were mixed with 10 μl of a solution of 0.1 mg ml−1 Nile Red (Sigma-Aldrich) in
acetone or with 1 μl of 1 mM solution of Bodipy 665/676 in DMSO. Samples were then washed
with PBS, re-suspended in ProLong Gold anti-fade reagent (Invitrogen) and observed with a
Spectral-based Olympus FluoView 1000 confocal laser scanning microscope (Olympus, Japan)
using a combination of: 1) a diode 405 nm laser for Alexa Fluor 405 signal detection, 2) an argon
(488 nm) laser for Alexa Fluor 488 signal or Nile Red staining visualization, or 3) solid state
(633 nm) laser for Bodipy 665/676 detection. Chloroplast autofluorescence was excited by using

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solid state (556 nm) laser. Z-series images were collected and processed with the Olympus
FluoView FV1000 confocal microscope software (Olympus, Japan). Negative controls were
treated as above, but the primary antibody was omitted. For the observation other than
immunofluorescence such as counting LD number and size in the MLDP RNAi lines, live cells
were used without fixation. For electron microscopy, centrifuged cell pellets were processed as
previously described (Harris, 1989), and transmission electron micrographs were captured using
a JEOL100 CXII instrument (Japan Electron Optics Laboratories, Tokyo, Japan).

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RESULTS
Transmission Electronic Microscopy Reveals Two Stages of LD Mobilization
N deprivation led to drastic changes in cellular ultra-structures in Chlamydomonas, such as
vacuolization, replacing of stacked thylakoids by starch granules, and the accumulation of LDs
(Figures 4.1A and 4.2 A-C). On the other hand, ultra-structures of Chlamydomonas during
quiescence exit have not been explored in detail. To fill this gap, transmission electron
micrographs were taken over a time course of N resupply and showed that, initially, while LDs
decreased in size they remained connected with the chloroplast outer envelope (Figures 4.1 B, D
and 4.2 D-F). Later on, two types of LDs were visible with usually the larger ones still making
contact with the chloroplast and the smaller ones moving toward the vacuoles (Figures 4.2 G-I
and 4.3 A-I), in accordance with the finding that interaction of oil bodies with vacuoles
contributes to the degradation of storage lipids in seeds (Poxleitner et al., 2006). Completion of
LD degradation is not necessary for the cell cycle to proceed as LDs were found in newly
produced daughter cells (Figures 4.1C and 4.2 J-L).

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Figure 4.1. Ultrastructure of Cells during Quiescence Exit. (A-C) Transmission electron
micrographs of a representative 21gr cell grown in N deprivation (-N) for 48 h (A) and
representative cells resupplied with N (NR) and grown for r 9 (B) and 12 h (C). (D) Close
contact of lipid droplet (LD) with chloroplast the outer envelope during early stage of LD
mobilization is shown. Scale bars are indicated. ES, eyespot; P, pyranoid; S, starch granules.

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Figure 4.2. Ultrastructure of Cells during Quiescence Exit. (A-L) Transmission electron
micrographs of a representative 21gr cell grown in N deprivation (-N) for 48 h (A-C) and
representative cells N-resupplied (NR) for 9 (D-F), 12 h (G-I) and 18 h (J-L). Scale bars are
indicated.

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Figure 4.3. Ultrastructure of Cells during Quiescence Exit. (A-C) Transmission electron
micrographs of a representative 21gr cell grown following N deprivation for 48 h followed by 9
h of N resupply (A), at higher magnification in (B, C). LD, lipid droplet; ES, eyespot; P,
pyranoid; S, starch granules. (D-F) Electron micrographs of LD-chloroplast contacts in cells
fixed at 12 h of N resupply. (G-I) Electron micrographs showing smaller LDs detaching from the
chloroplast and moving towards the vacuole. Scale bars are indicated.

MLDP Recruits Different Sets of Proteins to LDs
Previously we showed that the reduction of MLDP – the major lipid droplet protein in
Chlamydomonas – increases LD size without affecting TAG content (Moellering and Benning,
2010). In addition, its abundance is linked to the status of quiescence (Tsai et al., 2014). This
protein could provide a unique entry for a more in-depth study of the role of LDs in cellular
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quiescence. MLDP abundance decreased following N resupply as shown before (Figure 4.4A)
(Tsai et al., 2014). It was highly enriched in the fraction containing of LDs, and much less in the
ER fractions (Figure 4.4B). BIP was used as an ER marker. I used indirect immuno-fluorescence
to probe the localization of MLDP in fixed N-deprived cells. Signals of MLDP predominantly
formed a ring-like structure around LDs (Figure 4.4C), which is typical of proteins located on the
LD surface (Welte, 2007). They could also be detected in the cytosol albeit to a lesser extent, and
seemingly in a reticulate pattern. Notably, the distribution of MLDP was not evenly around LDs
but often polarized on one side or on multiple spots, and the larger the LDs the more
concentrated the MLDP. We think that in Chlamydomonas the coating with scaffold proteins – at
least in the case of MLDP – is a separate process carried out after LDs have been formed.
Many of the enzymes identified in LD proteomes possess neither amphipathic helices nor
hydrophobic spans (Walther and Farese, 2012). I hypothesized that in Chlamydomonas these
proteins associate with LDs by protein-protein interactions, e.g. with LD-bound protein, such as
MLDP. LDs isolated from cells deprived of N for 48h were solubilized and complexes were
fractionated by histidine and deoxycholate-based native PAGE (HDN-PAGE) (Ladig et al.,
2011). Immunoblots recognized MLDP in five distinct complexes of between 66 to 480 kDa
with no detection of the MLDP monomer (~28 kDa) (Figure 4.4D). A gel slice from the first
dimension was excised and the proteins therein were separated by SDS-PAGE in the second
dimension. The 2D SDS-PAGE confirmed the identity of these immunoreactive proteins to be
MLDP (Figure 4.4E). While the change in MLDP abundance was modest at 9 h of N resupply
(Figure 4.4A), the difference in complex formation was great – only the complex of the lowest
molecular mass was detected (Figure 4.4D). To characterize the protein components in these
MLDP complexes, I performed co-immunoprecipitation (Co-IP) in combination with LC-

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MS/MS. Dynabeads coupled with MLDP antibodies were incubated with LDs isolated from cells
exposed to 48 h of N deprivation (-N LDs) or 48 h of N deprivation followed by 9 h of N
resupply (NR9 LDs). As a negative control, Dynabeads coupled with MLDP prebleed were used.
Although in total 817 (-N LDs) and 349 (NR9 LDs) proteins were found, quantitative
information was only obtained for 188 (-N LDs) and 25 (NR9 LDs) distinct protein sets by
requiring that at least two peptides of each protein were repeatedly quantified in all three
replicates and that the ratio of total spectrum count between the experiments and the negative
control using MLDP prebleed was less than that for MLDP (Online Supplemental Data Set 11).
In theory, this approach allowed us to identify proteins that localized to LDs through the
interaction with MLDP. Compared with two recently reported Chlamydomonas LD proteomes
(Moellering and Benning, 2010; Nguyen et al., 2011), our experiments uncovered 97 newly
identified LD proteins (Figure 4.4F). Examples are MGD1 and LIP1. It is likely that MGD1is
sheltered by association with LDs from proteolysis and, thus is readily available without
transcription and translation as N is resupplied, to resynthesize the MGDG degraded during N
deprivation without delay. LIP1 is a TAG-reducing lipase. Being constitutively on LDs shows
that LIP1-mediated lipolysis is not controlled by a translocation event that moves the enzyme
from the cytosol to LDs, but requires activation of activity, as has been shown for the yeast TAG
lipase Tgl4 (Kurat et al., 2009). 28 proteins were found in all three data sets (Figure 4.4F),
including those involved in lipid metabolism such as BTA1, LCS1, a phosphoinositide
phosphatase (TEF21), a phospholipase D (Cre13.g591900), a glycosyl hydrolase with similarity
to the galactolipid:galactolipid galactosyltransferase SFR2 of Arabidopsis, although this activity
has not been shown to exist in Chlamydomonas) and a cyclopropane-fatty-acyl-phospholipid
synthase (CFA2). On the contrary, only 25 proteins were identified by MLDP CoIP with the LDs

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isolated following N resupply (Figure 4.4G; Online Supplemental Data Set 11), in accord with
the finding that only one type of MLDP complexes was detected by HDN-PAGE under this
condition. 11 of the 25 proteins were identified in both CoIP experiments, some of which may
constitute the core of MLDP complexes, such as BTA1 and β-tubulin (TUB1).

Figure 4.4. Lipid Droplet Localization and MLDP Complex Formation. (A) Immunoblot of
MLDP in dw15 following N resupply (NR) at times indicated (h). Coomassie Brilliant Blue
(CBB) staining shows equal protein loading. (B) Relative abundance of MLDP in different cell
fractions is shown by immunobloting. WCL, whole cell lysate; LD, lipid droplet; ER,
endoplasmic reticulum. BIP is used as an ER marker. Equal amounts of protein were loaded. (C)
Immunofluorescence detection of MLDP during N deprivation. BF, bright field. Scale bars in
each panel represent 2 μm. (D) LDs isolated at 48 h of N deprivation (-N) and 9 h of N resupply
(NR) were fractionated by HDN-PAGE and detected by MLDP immunoblot. Arrows indicate
each MLDP complex identified. (E) Proteins of –N LDs were separated by HDN-PAGE in the
first dimension (on the top) and then SDS-PAGE in the second dimension (on the left), and
detected by MLDP immunoblot. (F) Overlapping proteins between the data sets of Moellering et
al., of Nguyen et al., and of the LD MLDP-CoIP conducted with the LDs harvested at 48 h of N
deprivation. (G) Overlapping proteins between the data sets of the LD MLDP-CoIP conducted
with the LDs harvested at 48 h of N deprivation (-N) and at 9 h of N resupply (NR).
MLDP Is Indispensible for α-tubulin Association with LDs
Interaction with MLDP might be important for the stability or the sequestration of its associated
protein partners. Given the limited number of tools developed for Chlamydomonas, I decided to

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focus on two representative examples of the CoIP results. These are α-tubulin (TUA1), which
like β-tubulin is the major component of microtubules, and the presumed ortholog of TGD2, a
protein involved in ER-chloroplast lipid trafficking (Awai et al., 2006). In fact, α-tubulin is one
of the 28 proteins found in all three data sets, whereas TGD2 is present in two of the data sets but
not in the one generated by Moellering et al. I first verified the association of α-tubulin and
TGD2 with lipid droplets by subcellular fractionation, with respective negative controls to judge
the purity of lipid droplets. α-tubulin is universally known as cytosolic; therefore the
Chlamydomonas cytosolic marker NAB1 (for putative nucleic acid binding protein) was used as
a negative control (Mussgnug et al., 2005). Although not more enriched than whole cell lysates,
α-tubulin was seen in the LD fractions while NAB1 was not (Figure 4.5A). As in Arabidopsis
(Awai et al., 2006), Chlamydomonas TGD2 was presumed to be on the inner envelope of
chloroplasts at least during regular growth. However, upon N deprivation it was strongly
enriched in the LD fraction, but not another chloroplast inner envelope protein, Tic40. Early on
we suppressed MLDP by RNA interference (RNAi) and showed the effect on LD size in
multiple independent lines (Moellering and Benning, 2010). Here, when MLDP is reduced, αtubulin and TGD2 were not affected on the whole cell level (Figure 4.5B). Strikingly, α-tubulin
virtually disappeared from the LDs isolated from MLDP RNAi lines (Figure 4.5C). It appears
that MLDP is required for α-tubulin association with LDs, and perhaps also for mediating the
physical interaction between microtubules and LDs. TGD2, on the other hand, persisted on LDs
regardless of MLDP. Perhaps MLDP was helpful but not required for the relocation of TGD2, or
the level of MLDP was not low enough to have an impact. I also examined the protein levels of
MLDP, α-tubulin, and TGD2 in the LDs fraction when cells were exiting quiescence. Recall that
within the first 9 h of N resupply, MLDP only slightly altered its abundance on whole cell level

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(Figure 4.4A). It was similar for MLDP in the LDs fraction (Figure 4.5D). Protein abundance of
α-tubulin (modestly) and TGD2 (strongly) decreased in LDs in response to N resupply. TGD2
may have been relocated back to chloroplast, perhaps due to the close contact during the early
stage of LD degradation (Figure 4.1D).

Figure 4.5. Lipid Droplet Protein Quantification in the MLDP RNAi Lines. (A) Immunoblot
detection of cytosolic proteins α-tubulin (α-tub) and NAB1, and putative chloroplast inner
envelope proteins TGD2 and Tic40. WCL, whole cell lysate; LD, lipid droplet. Equal amounts of
protein were loaded. (B) MLDP, α-tubulin, and TGD2 abundance in the whole cell lysates of
dw15 and MLDP RNAi lines (mldp). Equal amounts of protein were loaded. (C) Protein
abundance in the LD fractions. 1X, 1/2X and 1/4X represent the serial dilutions of protein
loading, and are equal in dw15 and MLDP RNAi lines. (D) Comparison of protein abundance in
the –N (48 h of N deprivation) and NR9 (9h of N resupply) LDs. All data shown here are
representative of three or four independent biological replicates.

The Surface of Lipid Droplet Has Unique Lipid and Fatty Acid Composition
By and large, LDs consist of a neutral lipid core surrounded by a phospholipid monolayer
(Tauchi-Sato et al., 2002). While it is true for yeast and animals, it might be different in
Chlamydomonas since the commonly most abundant phospholipid, PC, is missing. I was
therefore interested in characterizing the polar lipid fractions of LDs. Lipid species from total
cells or LDs were separated by thin layer chromatography (TLC) plate (Figure 4.6A and B).
DGDG and DGTS appeared to be most abundant among polar lipids associated with LDs,
confirmed by quantitative analysis using gas chromatography (Online Supplemental Table 1).

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These data explained the increase of DGTS and especially DGDG during N deprivation (Figure
5C), as LDs grow in size and number so does the demand for surface lipids. Under closer
examination, phospholipids all together (i.e. PE, PG and PI) only accounted for less than 25% of
LD surface lipids, contrasting to ~25% of DGTS, and more than 50% of galactolipids which
could only originate from the chloroplast. Surprisingly, the fatty acid spectrum of LD surface
lipids was very different from that of whole cells, generally more enriched in saturated fatty
acids (~70% for lipid droplets versus ~40% for whole cells). This finding strongly suggested that
these polar lipids were bona fide LD lipids. The abundance of DGDG dropped in half (~32% to
~16%) after 9 h of N resupply, implying a regulated lipid trafficking happened on the surface of
LD or a targeted degradation of DGDG.
To my surprise, two additional lipid species were seen chromatographingly close to
DGTS and DGDG in fractions from the LDs of the RNAi lines (Figure 4.6 B). Since they were
not detected in the total cell fractions, this effect was LD-specific. We deduced that these lipids
could be different molecular species of DGTS and DGDG, according to their responses to
Dragendorff reagent which reacts with tertiary amines of DGTS, and to α-naphthol which binds
to sugar group such as galactose of DGDG (Figure 4.6C) (Benning et al., 1995). Also, the
combined amount of the original and putative DGTS in the RNAi lines (~26%) was comparable
to the DGTS in the PL (~24%), so was for DGDG (~35% in RNAi lines versus ~32% in the PL).
The difference in migration could result from the difference in fatty acid composition. Lipids
with more saturated fatty acids are slightly less hydrophobic and thus migrate slower on TLC
plate compared to the same species of lipid carrying more unsaturated fatty acids. Indeed, in the
RNAi lines, saturated fatty acids were ~90% in the putative DGTS and ~92% in the putative
DGDG, whereas the original DGTS and DGDG contained ~78% and ~69% of saturated fatty

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acids, respectively. Overall, the surface of LDs isolated from RNAi lines was even more
enriched in saturated fatty acids (~88%). Collectively, these results suggest that MLDP affects
the lipid composition at the surface of lipid droplets.

Figure 4.6. Lipid Profiles of Lipid Droplet Surface Membrane. (A) Whole cell lipids of dw15
and MLDP RNAi lines (mldp) were separated by thin layer chromatography (TLC). MGDG,
monogalactosyldiacylglycerol;
DGTS,
diacylglyceryl-trimethylhomoserine;
PE,
phosphatidylethanolamine; PG, phosphatidylglycerol; DGDG, digalactosyldiacylglycerol;
SQDG, sulfoquinovosyldiacylglycerol; PI, phosphatidylinositol. (B) Lipids of lipid droplets were
separated by TLC. Arrow and asterisk indicate additional lipids from the mldp lipid droplets. (C)
Lipids separated by TLC were stained by α-naphthol (left panel) Dragendorff reagent (right
panel). (A), (B) and (C) are not from the same TLC plate, except left panel of (C) is the same
plate as in (B).

Proper Surface Area of Lipid Droplet Ensures Orderly Progression out of Quiescence
Fortuitously I observed that MLDP RNAi lines did not grow in fresh medium if inoculated from
plates over three weeks old, whereas regular strains can be stored for months under the same
conditions, implying a defect in quiescence exit. To follow up, we transferred cells in the midlog phase to liquid medium without N, and waited for various times before adding N back. Cell
biomass based on optical density and cell concentration were monitored at 0 and 24 h of N
resupply. The regrowth of PL gradually slowed down over the longer period of N deprivation,
and RNAi lines began to lag further behind after being incubated in N-free medium for 6 days
(Figure 4.7A and B). TAGs continued to accumulate in the PL and RNAi lines and finally
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saturated at day 6 of N deprivation. The delayed turnover of TAGs in the PL after prolonged N
deprivation correlated with the slower regrowth, and starting from the samples being deprived
for 4 days, RNAi lines exhibited a lipolysis defect even though it did not yet affect regrowth
(Figure 4.7C). Knowing that the reduction of MLDP causes morphological changes of LDs, we
examined the dynamics of LD formation in the PL and RNAi lines. As reported before, after 2
days of N deprivation RNAi lines had fewer LDs but larger in size (Figure 4.7D and E). At day 4,
LDs in the PL grew to near the size of LD in RNAi lines at day 2 but decreased in number,
indicating LD fusions. A critical observation was made on day 6. In the PL there was no obvious
increase of large LDs (1 μm up in diameter), but numerous small ones (1 μm down in diameter)
were formed, positing that large LDs had grown to their maximal capacity and new LDs were
needed to accommodate the incoming TAGs. On the contrary, LDs in the RNAi lines continued
to enlarge. By day 8, small LDs were no longer seen in the PL, indicating that another round of
fusion events had occurred. LDs in RNAi lines never ceased to fuse even though after day 6
there was no increase in TAGs, and in some cells the entire cytosol was occupied by a giant
droplet (Figure 4.7F). In summary, our data demonstrated that MLDP balanced the surface areaper-volume ratio which is important for metabolizing TAGs, and ensured orderly progression out
of quiescence.

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Figure 4.7. Reduction of MLDP Affects Quiescence Exit, TAG Turnover, and Lipid
Droplet Homeostasis. (A) Cultures of dw15 and MLDP RNAi lines (mldp) at 24 h of N resupply
(NR) following N deprivation (-N) as times indicated (d, day). (B) Growth of dw15 and mldp
during the course of N deprivation followed by N resupply measured at the times indicted. (C)
TAG contents (depicted as ratio of TAG fatty acids (FA) over total FAs) at different days of N
deprivation (-N) and at 24 h of N resupply (NR) following N deprivation as times indicated. (D,
E) Lipid droplets (LD) of 15 cells for each sample were counted (D) over days of N deprivation
as indicated, and their diameter (E) was quantified with imaging software and shown in data
point-box plot. (F) BODIPY staining of LDs in representative cells. Scale bars represent 2 μm.

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DISCUSSION
The role of lipid droplet in cellular quiescence
Biochemical assays validated some of the inferences based on transcriptional analysis described
in Chapter 3. In fact, many key enzymes in lipid metabolic pathways were identified by MLDP
CoIP from LDs, the most prominent of which is BTA1. Normal growth of LD requires
increasing amount of DGTS in the surface area. It is reasonable to assume that BTA1 is deployed
on LDs so that DGTS can be synthesized locally. BTA1 was only upregulated during N resupply,
the time when membrane rebuilding is heavily in demand. It is also possible that BTA1 is
protected on LDs from active proteolysis, so that during transit out of quiescence it can
immediately participate in the restoration of organelles. These hypotheses are not mutually
exclusive. LD localization of LACS1 has been observed repeatedly including the study described
here. Its possible function can be discussed in view of two conceptual phases. First, fatty acids
are synthesized by 3-ketoacyl-ACP synthase in the form of acyl-ACP (acyl carrier protein). To
be exported out of chloroplast the ACP moiety must be removed by the activity of acyl-ACP
thioesterase, and almost instantly long-chain acyl-CoA synthetase activates the resulting free
fatty acids so they can be incorporated into glycolipids. Previously it was thought that fatty acids
after being converted to acyl-CoA were transported to the ER where TAG was assembled (Guixe
and Babul, 1985). Here I propose a new possibility, namely that activation of fatty acids occurs
at the contact sites between chloroplast and LDs after ACP is removed. Second, several putative
lipases and acyltransferases are found on LDs including examples from the current work, a lysophosphatidate:acyl-ACP acyltransferase (g9888) and a phospholipid/glycerol acyltransferase
(Cre17.g738350), indicative of active deacylation and reacylation. LACS1 may intervene in the
processes by turning fatty acids released by lipase into acyl-CoA, which can then be utilized by

129

the acyltransferase. Theoretically, the proteins isolated by MLDP CoIP from LDs are LDlocalized by interacting with MLDP. However, an inevitable pitfall of CoIP experiments is
protein contamination through the hydrophobic interaction with bait protein, a fact that needs to
be taken into consideration when interpreting the data.
The monolayer surface of LD contained all the polar lipids that can be found in
Chlamydomonas. At first, it may seem like contamination, especially with more than 50% of
galactolipids whose biosynthesis and distribution are thought to be exclusively with the
chloroplast. However, the fatty acid profile differed in every polar lipid when compared between
LDs and whole cells. Except for DGTS, there is no indication of possible local synthesis.
Therefore, one can only assume that these lipids were transported to LDs from the chloroplast
(i.e. MGDG, DGDG, SQDG and PG) and the ER (i.e. PE, PI, PG and perhaps DGTS) where
syntheses occur. Despite coming from different sources, these lipids share a common feature
which is a higher saturation level than their counterparts in whole cell extracts. This has led me
to hypothesize a LD-specialized acyl exchange mechanism that utilizes the de novo-synthesized
acyl groups palmitate (16:0) and oleate (18:1Δ9) to replace the mature, mostly unsaturated fatty
acids from polar lipids, which are then stored in lipid droplets (Figure 4.8). Oleate seemed to be
specific for the LD DGDG (Online Supplemental Table 1). It is reminiscent of the
deacylation/acylation cycle catalyzed in part by PGD1 during N deprivation, which prefers
18:1Δ9/16:0 MGDG but not its mature 18:3/16:4 molecular species (Li et al., 2012). The
proposed mechanism can be potentially executed by any of the numerous LD-bound lipases and
acyltransferases that have been reported in Chlamydomonas (Moellering and Benning, 2010;
Nguyen et al., 2011). The fact that LD surface lipids in MLDP RNAi lines had an even higher
saturation rate, to an extent that distinct DGTS and DGDG species were possible separated by

130

TLC, further confirms the acyl exchange took place on the LDs. This might result from the lower
total LD surface area in the RNAi lines, in other words, lower substrate-to-enzyme ratio, and
give rise to a high degree of acyl exchange.

Figure 4.8. A model for lipid droplet-specialized deacylation/reacylation cycle. De novo
synthesized fatty acids (FA) (16:0, 18:0, and 18:1Δ9) in the form of acyl-ACP are exported from
chloroplast either right after fatty acid synthesis (FAS) or after the deacylation/reacylation cycle
mediated by PGD1. Acyl-ACP is transformed into acyl-CoA by lipid droplet (LD)-bound
LACS1 before it can be incorporated into polar lipids or triacylglycerols (TAGs) on the LD. The
ER-originated and chloroplast (CP)-originated polar lipids after transported to LD are subjected
to deacylation, following by reacylation using the de novo synthesized FA stored in the LD. The
deacylation/reacylation cycle may repeat until both acyl groups have been replaced. The
polyunsaturated FA removed during the deacylation is then incorporated into TAG and stored in
the LD.

A new model of lipid droplet formation
LD assembly in Chlamydomonas can be divided into two distinct phases (Figure 4.9). TAGs
were packed into a hydrophobic compartment in proximity to the chloroplast. Whether this is a
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direct budding from chloroplast is not clear at this point, but the close proximity and punctate
fusions ought to have physiological relevance, for instance, to allow chloroplast lipids or proteins
to diffuse naturally to LDs. When LDs reached a certain size (approximately 0.5 μm in diameter),
MLDP proteins would begin to gather around them. This proteinaceous coating is apparently
important in determining LD size, because LDs would no longer fuse or enlarge once they have
been stabilized by MLDP (approximately 2 μm in diameter). Supposedly, as more TAGs were
continuously being made, new rounds of LD formation and stabilization have to occur. How is
MLDP transported to LDs? Unique peptides such as sequences that direct proteins to the ER or
chloroplast have not been found for LDs in any cell type, despite that hundreds of LD proteins
have been identified. Nevertheless we presume microtubule-mediated transport is likely involved
(Hirokawa et al., 2009). Microtubules are a component of cytoskeleton, commonly comprised of
α and β-tubulin, and they are known for LD fusion as well as LD trafficking by cytoplasmic
motor proteins in animal cells (Gross et al., 2000; Bostrom et al., 2005; Shubeita et al., 2008).
Both α and β-tubulin were shown to interact with MLDP (Online Supplemental Data Set 11) and
have been found in Chlamydomonas LD proteomes (Moellering and Benning, 2010; Nguyen et
al., 2011). In my hypothesis, minute droplets or vesicles carrying MLDP are secreted from the
ER, and then the droplets are bound to microtubules through MLDP, and transported to the
uncoated LDs that are attached to the chloroplast (Figure 4.9). Afterwards fusions occur and
release the cargo. Microtubule units remained bound to the mature LDs through MLDP as
demonstrated by the example of α-tubulin. In view of this, MLDP functionally resembles a
perilipin homolog LSD2 which coordinates LD motion in Drosophila embryos by clustering with
another LD protein Klar and the motor protein dynein that move along microtubules towards
minus-end (Welte et al., 2005). This model suggests that in Chlamydomonas the main role of the

132

ER in LD formation is to provide decorated proteins and ER-originated polar lipids, and less in
TAGs, consistent with the finding that the reduction of MLDP did not affect the total TAG
content stored in the mature LDs.

Figure 4.9. A model for lipid droplet formation and deformation. Small lipid droplet evolves
from the chloroplast (CP) and thus allows natural diffusion of CP proteins to LD, such as TGD2.
As the LD grows, MLDP complexes are transported to LD through microtubule to stabilize the
mature LD. The complexity of MLDP complex increases when interacting with proteins
preoccupied the LD. Upon N resupply, some proteins dissociate from MLDP complexes and
move away from the LD. Small LD detaches from the CP and moves toward vacuole for final
turnover.

Is MLDP part of a LD targeting mechanism? MLDP forms multi-order protein complexes
on the mature LDs. These complexes or key components of the complexes could have been
assembled on the ER, and then moved along microtubules to LDs through the mediation of
MLDP. The proteins preexisted on the developing LDs, such as those from the chloroplast, then

133

interacted with the MLDP complexes after being transported, thus contributing to the complexity
of the complexes (Figure 4.9). This might explain why TGD2 did not require MLDP to locate on
LD in spite of a possible interaction. Thus far underlying mechanisms for LD protein targeting
are poorly understood, with some insights derived from the Arf1/COPI machinery that secures
the ER-LD connections for protein trafficking (Wilfling et al., 2014). Future investigation of the
transport of MLDP will provide unique insights in photosynthetic cells.
The dynamics of LDs has direct impact on the reversibility of quiescence. TAGs stored in
LDs are the main source of energy and membrane building blocks to fuel the regrowth.
Therefore, when droplets fail to optimize the surface area-to-volume ratio it not only affects the
efficacy of TAG turnover, which is most likely due to the limited accessibility to lipases, but also
disturbs the timeliness of quiescence exit, and this situation could become exacerbated if the time
in quiescence is extended. Close LD-to-chloroplast proximity was maintained during the first
stage of LD mobilization, which might create a conduit for lipids (e.g. DGDG) and proteins (e.g.
TGD2) to move back to chloroplast. CHT7 and MLDP appear to represent different classes of
effectors for cellular quiescence, according to their respective mutant phenotypes. Once the cells
of cht7 enter quiescence (approximately 12 to 24 h of N deprivation), they are destined to
experience severe challenges at the exit (Tsai et al., 2014). On the other hand, a defect in
regrowth of the MLDP RNAi lines only became obvious in prolonged quiescence. In a sense, the
depth of quiescence is manifested by the developmental stage of LDs.

134

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CHAPTER 5
Conclusions and Perspectives

139

Nature is the integration of environmental factors and biological complexity. In response to the
dynamic environment all living cells need to make a resolute choice between division and
quiescence to ensure the continuity of life; the decision made is then executed through cell cycle
determinants and metabolic fluxes. Plants, due to their sessile nature, are inevitably exposed to
more biotic and abiotic stresses that drive them to redirect resources from growth and cell
division to respective defence mechanisms. However, how plant cells recover from the
conditional, temporary cessation of growth/division has never been explored in detail. In this
dissertation, I present progress towards the comprehensive understanding of cellular quiescence
in photosynthetic eukaryotes, using Chlamydomonas reinhardtii as a reference model. Among
the principle findings are the discovery of CHT7 as a repressor of cellular quiescence (Chapter 2),
the CHT7 regulon and the transcriptional programs unique to N resupply (Chapter 3), and the
aspects in which lipid droplets (LD) integrate with cellular metabolism to ensure the orderly
progression from quiescence (Chapter 4). In this final chapter, I will discuss the unanswered
questions and related future issues.

Regulatory processes by which CHT7 maintains the reversibility of quiescence
Current data point toward a role of CHT7 as a repressor of transcriptional programs associated
with quiescence, based on which, I hypothesized that CHT7 secures the orderly and rapid exit
from quiescence by resuppressing the induced responses. Nevertheless, comparative
transcriptomics of N-resupplied conditions uncover a subset of the CHT7 regulon that differs
from the gene regulation CHT7 imposes under normal N-replete growth, e.g. the tetrapyrrole
pathway and peroxisomal redox homeostasis. As such, CHT7 may have additional, more direct
impacts on quiescence exit. These speculations can only be addressed through the identification

140

of the in vivo chromatin binding sites for CHT7 or CHT7-containing complexes under different
growth conditions. Integrative analysis of the transcriptome and chromatin binding data will help
distinguish the genes that are primary or secondary targets of CHT7, as well as clarify the
potential feedback regulations by metabolite levels.

Potential modification of CHT7 activity during the entry into quiescence
In yeast, the pathways that prevent the entry into quiescence (e.g. cAMP-PKA) must be
deactivated during the transition; otherwise cells will not be able to survive (Uno et al., 1984;
Shin et al., 1987). Since CHT7 also represses quiescence when cell cycle progresses regularly, it
is reasonable to speculate that the activity of CHT7 is curtailed during quiescence. As a matter of
fact, CHT7 seems dispensable during quiescence for its mutant remains viable even under
extended period of N deprivation. Overexpression of CHT7 during quiescence using promoters
responsive to N deprivation, such as that of MLDP or NIT1 (nitrate reductase 1), will help
address this issue. The overexpression lines are expected to lose viability during quiescence.
Provided that CHT7 is indeed deactivated during quiescence, the question arises: By
what means can this be done? The persistence of CHT7 and CHT7 complexes before, during,
and after quiescence suggests that fluctuation of abundance or change of interacting partners
likely is not part of the regulatory mechanism to regulate CHT7. Nuclear-cytoplasmic shuttling is
neither the case as CHT7 remains in the nucleus throughout the quiescence cycle. A recently
published global phosphoproteome of C. reinhardtii reports two phosphorylation sites of CHT7
during regular growth condition (Wang et al., 2014). Posttranslational modifications of CHT7
depending on the nutritional status of the cell may modulate its possible DNA binding

141

preferences. But again, the experimental evidence of posttranslational modification and whether
it causes functional impact remain to be addressed.

The regulatory logic underlying the CHT7 and Rb complexes to enable transitions between
quiescence and cell division
In Chlamydomonas, a protein complex containing orthologs of DP1, E2F1 and Rb/MAT3 is
present in the nucleus throughout the cell cycle (Olson et al., 2010). Two lines of evidence let me
hypothesize that CHT7 may be a part of larger regulatory complex that transiently associates
with the Rb/MAT3 protein of Chlamydomonas. First, CXC domain proteins have been found in
large regulatory protein complexes with Rb in animal cells (Sadasivam and DeCaprio, 2013; van
den Heuvel and Dyson, 2008), and second, the mat3 mutant of Chlamydomonas not only shows
a cell size defect, but like the cht7 mutant is also slow to respond when resupplied with N
following N-deprivation (Armbrust et al., 1995). Precedence is set by the dynamic nature of
CXC protein-containing complexes such as the human DREAM/LINC complex. Five proteins,
including the CXC domain protein LIN54, form the complex core and remain together
throughout the cell cycle (Litovchick et al., 2007). During quiescence, the core binds the Rb-like
pocket protein p130, E2F4, and DP to repress cell cycle-dependent gene expression. This
association is reversed upon phosphorylation of the core subunit LIN52, hence allowing the core
to recruit the transcription factors BMYB and then FOXM1 to promoters with highest activity
during the G2/M phases (Sadasivam and DeCaprio, 2013). Thus, multiple CHT7 containing
complexes of varying abundance depending on the cellular conditions seem possible and will
have to be explored in future to gain a more complete understanding of the function of CHT7.

142

The Rb/MAT3 protein of Chlamydomonas clearly has functions in addition to CHT7
such as in the control of cell size (Armbrust et al., 1995; Umen and Goodenough, 2001). At this
time, no data describing global changes in transcriptional profiles for the mat3 mutant under
different N supply conditions are available that would allow me to compare possible regulatory
networks governed by CHT7, Rb/MAT3 or both.

Functional interplay between CHT7 and other quiescence regulators
Recall that contrasting to yeast, rapamycin functions only to slow down, not to
completely abolish the growth and division of Chlamydomonas. The cht7 mutant is more
sensitive to rapamycin treatment, judged by the even slower growth in the medium with
rapamycin (Tsai et al., 2014), which means that both CHT7 and TOR pathways are negative
regulators of quiescence and are not redundant. On the other hand, the cAMP-PKA signaling
pathway, to the most part, might be redundant with CHT7 in regulating the exit from quiescence.
It is reasonable to hypothesize functional interplay among pathways of CHT7, TOR, cAMP-PKA,
and even Rb/MAT3, as illustrated in Figure 5.1. Hints about the role of CHT7 in the regulatory
networks of quiescence can be obtained by characterizing the protein components of CHT7
complex. This comparative proteomic analysis should provide direct evidence, whether and to
what extent the CHT7 complex shares common components with TOR, cAMP-PKA, and Rb
complexes. Arguably, current knowledge does not even begin to accurately portray the multiple
flavors of the N deprivation-induced quiescence or quiescence in general, and other key
regulators doubtless remain to be uncovered.

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Figure 5.1. Summary of pathways thought to control transitions between growth/division
and quiescence in Chlamydomonas reinhardtii. The CHT7 and TOR pathways are thought to
repress aspects of quiescence. CHT7, Rb, and PKA signaling are required for orderly transition
out of quiescence. CHT7 and PKA pathways may share a certain level of redundancy.

Lipid droplets are important for the survival during quiescence and for the quiescence exit
The three constituents of LD—a TAG core, polar lipid monolayer, and associated proteins—give
hints to the multiple roles of LD in cellular quiescence. In response to quiescence,
Chlamydomonas cells deposit TAGs into LDs. On the one hand, it likely takes away the acetylCoA from TCA cycle so prevent it from producing chemical energy (i.e. ATP and NADH), as a
way to restrict cell growth and division. On the other hand, it consumes the reducing power
generated from the photosynthetic electron transport chain, which otherwise will cause lethal
oxidative damage to the cell (Li et al., 2012). To be used as membrane building blocks in most
organelles, polar lipids must undergo several steps of desaturation on their acyl chains. During
quiescence, although large portion of polar lipids, thylakoid MGDG in particular, are degraded
by autophagy, their polyunsaturated acyl chains are preserved safely. The proposed LDspecialized deacylation/reacylation cycle provides one such mechanism, which removes
polyunsaturated fatty acids from the polar lipids imported to the LD and stores them in the TAG
core. Supposedly, this process allows rapid reconstruction of organelle lipids without remaking
the polyunsaturated fatty acids when conditions reverse. Autophagy also degrades proteins. It has
been hypothesized and observed that LDs serve as temporary shelters for proteins under stress
144

conditions (Welte, 2007). In Chlamydomonas, LDs may function in a similar way, and MLDP
may establish LD targeting through protein-protein interaction for proteins unable to anchor
themselves on the lipid monolayer. Again, preservation of these proteins might be crucial for the
rapid restoration of cellular status during the transit out of quiescence.

Biotechnological applications of cellular quiescence and lipid droplet biology
Microalgae accumulate valuable compounds under conditions adverse to growth. For example,
nutrient starvation causes accumulation of TAGs but also induces cellular quiescence,
characterized by the reversible cessation of growth. Among other factors, this inverse
relationship between biomass productivity and TAG accumulation has long hampered efforts
toward the efficient generation of biofuel feedstocks from microalgae. The discovery of a mutant
and the corresponding protein CHT7 affecting the orderly transition of algal cells from
quiescence to normal growth provides mechanistic insights to address this problem. For example,
CHT7 acts as a repressor of cellular quiescence. It might be possible to use CHT7 to lower the
expressivity of quiescence in the oil-producing conditions, and thus to engineer a high-biomasshigh-oil algal strain. In addition, overexpression of CHT7 in the regular growth condition might
delay the entry into stationary phase, and enhance the biomass capacity per growth unit.
LDs can also be readily isolated, thus facilitating harvesting of the desired compound.
Knowledge of how complex enzyme arrays can be assembled on LDs allows the design of a
synthetic intracellular platform that can produce both known secondary metabolites and novel
compounds in unicellular photosynthetic organisms, as well as having the benefit that lipophilic
toxic compounds can be sequestered into them and thus rendered harmless to the cell.

145

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