THE INTEGRATION of LIGHT and PLASTID SIGNALS by Michael E

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THE INTEGRATION OF LIGHT AND PLASTID SIGNALS

Michael E. Ruckle

A DISSERTATION

Submitted to Michigan State University in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

Biochemistry and Molecular Biology

2010

ABSTRACT

THE INTEGRATION OF LIGHT AND PLASTID SIGNALS

Michael E. Ruckle

Proper chloroplast biogenesis and function are essential for agriculture and life on earth because photosynthesis drives plant growth, development, and reproduction.

Photosynthesis-related gene expression was previously reported to be induced by light signaling and repressed by plastid signaling. Although light signaling and plastid signaling were previously thought to independently regulate the expression of these genes, data indicating that the regulation of photosynthesis-related gene expression by light and plastid signals depends on common promoter elements led me to hypothesize that light signaling and plastid signaling might be interactive processes and that these interactions might be significant. I first tested this hypothesis by screening a group of

Arabidopsis mutants with defects in plastid signaling for light signaling phenotypes.

Based on results from these experiments, I conclude that the blue light receptor cryptochrome1 (cry1) contributes to both the light and the plastid signaling that regulates the expression of genes encoding the light-harvesting chlorophyll a/b-binding protein (Lhcb) of photosystem II. I provide evidence that plastid signaling broadly

³UHZLUHV´OLJKWVLJQDOLQJDQGWKDWLQWKHFDVHRIWKHFU\VLJQDOLQJWKDWUHJXODWHVLhcb

H[SUHVVLRQWKLV³UHZLULQJ´LVODUJHO\FDXVHGE\ the conversion of long hypocotyl 5 (HY5) from a positive regulator to a negative regulator of Lhcb expression. HY5 is a bZIP-type

transcription factor that acts downstream of cry1 and other photoreceptors. I found that cry1-dependent plastid signals are genetically distinct from GENOMES UNCOUPLED 1

(GUN1)-dependent plastid signals and that the interactions between light and plastid signals appear critical for proper chloroplast biogenesis. Addtionally, I found that plastid signals can broadly affect light-regulated development of Arabidopsis seedlings.

Results from these developmental assays are consistent with cry1 and GUN1 helping integrate chloroplast function with light regulated development. Based on these findings, I hypothesized that the interactions between light and plastid signaling promote chloroplast biogenesis by optimizing the expression of chloroplast-related genes for particular light environments and that plastid signaling broadly regulates light signaling by affecting possibly numerous signaling factors that act downstream of photoreceptors.

We tested these ideas with time-resolved- expression profiling. Results from the expression profiling are consistent with interactions between light and plastid signaling optimizing not only chloroplast biogenesis but also coordinating plant growth and development with chloroplast function. Results from the reverse genetic analyses of

Arabidopsis mutants yielded mutant alleles that cause abnormal chloroplast biogenesis.

These alleles have defects in eighteen genes that encode transcription factors, signaling factors, and proteins of no known function. These findings provide evidence that light and plastid signaling are interactive processes that not only promote chloroplast biogenesis and function but also affect diverse processes related to plant growth and development.

ACKNOWLEDGEMENTS

First and foremost I would like to thank Rob Larkin, for always having his door open for helpful discussions, for pushing me as hard as he could to help me reach my potential, for always providing me with support for my ideas, and for being an excellent mentor. I owe much of my success to him.

I would also like to thank my committee members Drs. Beronda Montgomery,

Robert Last, Sheng Yang He, and Christoph Benning, for their support and guidance.

I would like to thank my family, especially my mom, for providing me with the strong support that I needed along the way.

I would like to my friends at MSU, for making graduate school enjoyable, and for always offering a helping hand when I needed it.

I would like to thank Neil Adhikari for putting up with me as his office partner and lab-mate.

Much of the work presented here came at the aid of several amazing students. I would like to thank Andrea Stavoe, Chris Sinkler, Lauren Lawrence, and Stephanie

Demarco for all of their help.

TABLE OF CONTENTS

LIST OF TABLES««««.«««««««««««««««««««««««««Yii

LIST OF ),*85(6««««««««««««««««««««««««««««YLLL

CHAPTER 1: Introduction ,QWURGXFWLRQ««««««««««...... 2 $GDSWLQJ WR WKH OLJKW«««««...... 14 Light-UHJXODWHG VLJQDO WUDQVGXFWLRQ QHWZRUNV«««««...... 12 Chloroplast Development...... 22 3ODVWLG 6LJQDOLQJ««««...... 34 Integration of light and plastLG VLJQDOV«««««««««««««««42 )LJXUHV««««««««««««««««««««««««««««44 5HIHUHQFHV««««««««««««««««««««««««««1

CHAPTER 2: Plastid signals remodel light signaling networks and are essential for efficient chloroplast biogenesis in Arabidopsis Abstract...... 71 Introduction...... 72 Results...... 77 Discussion...... 95 Materials and methods...... 105 $FNQRZOHGJHPHQWV«««««««««««««««««««««««10 )LJXUHV««««««««««««««««««««««««««««11 5HIHUHQFHV«««««««««««««««««««««««««««8

CHAPTER 3: Plastid signals that affect photomorphogenesis in Arabidopsis thaliana are dependent on GENOMES UNCOUPLED 1 and cryptochrome 1 Abstract...... 144 Introduction...... 145 Results...... 148 Discussion...... 157 Materials and methods...... 164 AcknowledgementV««««««««««««««««««««««8 )LJXUHV«««««««««««««««««««««««««««9 References««««««««««««««««««««««««««2

CHAPTER 4: Characterization of transcriptomes and signaling factors that contribute to chloroplast biogenesis in Arabidopsis Abstract...... 199 Introduction...... 200 Results...... 206 Discussion...... 231 Materials and methods...... 248 v

$FNQRZOHGJPHQWV««««««««««««««««««««««««««3 )LJXUHV««««««««««««««««««««««««««««4 ReferencHV«««««««««««««««««««««««««««5

LIST OF TABLES -2 -1 Table 2.1 Segregation of the long hypocotyl phenotype in 25 ȝmol m s blue light inF seedlings 6 3 ««««««««««««««««««««««««««««««

Table 4.1 Genes that exhibit enhanced light-induced expression in lincomycin-treated seedlings and their publicly available T-DNA alleles««««««««««««««3

Table 4.2 Genes that exhibit enhanced light-induced exprssion in lincomycin-treated seedlings and that do not have publicly available T-'1$DOOHOHV«««««««««7

Table 4.3 Alleles of genes that exhibit similar light-induced expression in lincomycin- treated and untreated seedlings that cause end phenotypes««««««««««8

Table 4.4 The diverse regulators of genes that exhibit enhanced light-induced expression in lincomycin-treated seedlings««««««««««««««««««2

vii

LIST OF FIGURES

Figure 1.1 The affect of light quality and quantity on plant growth and development««««««««««««««««««««««««««««««««4

Figure 1.2 3ODQWSKRWRUHFHSWRUV«««««««««««««««««««««««6

Figure 1.3 A simplified model of the known components of light signaling during phRWRPRUSKRJHQHVLV«««««««««««««««««««««««««47

Figure 1.4 The known light signaling network«««««««««««««««««8

Figure 1.5 Agriculturally important plastid types««««««««««««««««9

Figure 1.6 Examples the chloroplast adapWLQJWRWKHOLJKW««««««««««««1

Figure 1.7 Examples of mutations in chloroplast development««««««««««3

Figure 1.8 The balance of light and plastid signals during chloroplast development...54

Figure 1.9 Plastid signaling during chloroplast development«««««««««««6

Figure 1.10 Strategy IRUREWDLQLQJPXWDQWVWKDWUHJXODWHFKORURSODVWGHYHORSPHQW«8

Figure 2.1 Allelism of new gun mutants and cry1 mutants«««««««««««1

Figure 2.2 Expression of Lhcb and Rbcs in gun1 and cry mutants after chloroplast biogenesis was blocked««««««««««««««««««««««««««3

Figure 2.3 Expression of Lhcb and Rbcs in gun1 and cry1 after chloroplast biogenesis was blocked with various inhibitors of chloroplast biogenesis««««««««««4

Figure 2.4 Expression of Lhcb and Rbcs in cop1-4 and hy5 mutants after chloroplast biogenesis was blocked««««««««««««««««««««««««««5

Figure 2.5 Expression of Lhcb and Rbcs in the dark or in various qualities of light...117

Figure 2.6 Expression of Lhcb and Rbcs in gun1, phyA, and phyB mutants after chloroplast biogenesis was blocked«««««««««««««««««««««9

Figure 2.7 The effects of plastid development on the fluence rate response of Lhcb and Rbcs..««««««««««««««««««««««««««««««««««20

Figure 2.8 Chlorophyll-deficient cotyledons in gun1 and gun1 cry mutants««««1

viii

Figure 2.9 HL sensitivity of gun1 and light signaling mutants««««««««««3

Figure 2.10 Model for PhANG regulation by a network of plastid and light signaling pathways«««««««««««««««««««««««««««««««««5

Figure 2.11 The gun mutant screen procedure««««««««««««««««6

Figure 2.12 Analysis of T-DNA alleles««««««««««««««««««««7

Figure 2.13 Expression of Lhcb in cry1 after chloroplast biogenesis was blocked in red light«««««««««««««««««««««««««««««««««««9

Figure 2.14 Lhcb and Rbcs expression in gun1-1 and various photoreceptor mutants after chloroplast biogenesis was blocked in blue light««...... 130

Figure 2.15 Lhcb and Rbcs mRNA levels in wild type and gun1 mutants grown in darkness and various light qualities without inhibitors of chloroplast biogenesis««1

Figure 2.16 Analysis of det/cop/fus phenotypes in gun1-1 and cop1-4««««««2

Figure 2.17 Chlorophyll-deficient leaves in gun1 and gun1cry1 double mutants««3

Figure 2.18 Phenotypes of gun1 and light signaling mutants following HL incubations«««««««««««««««««««««««««««««««.134

Figure 2.19 Cosegregation analysis of long hypocotyl in blue light and gun phenotypes««««««««««««««««««««««««««««««««5

Figure 3.1 Cotyledon opening in gun1-1 and wild-type seedlings in different fluence rates of blue light«««««««««««««««««««««««««««««9

Figure 3.2 Flattened cotyledon areas in gun1-1 and wild-type seedlings«««...... 170

Figure 3.3 Cotyledon areas in gun1, cry1, and hy5 mutants«««««««««««71

Figure 3.4 Analysis of cotyledon expansion in gun1-1 and phyB««««««««172

Figure 3.5 Analysis of epidermal cells in the cotyledons of gun1 and cry1 mutants«««««««««««««««««««««««««««««««««3

Figure 3.6 Analysis of anthocyanin levels in lincomycin-treated seedlings««««5

Figure 3.7 Analysis of anathocyanin levels during de-etiolation in bright white light «««««««««««««««««««««««««««««««««««6

Figure 3.8 Analysis of hypocotyl lengths in gun1, cry1, and hy5 mutants«««««8

Figure 3.9 Models for the regulation of photomorphogenesis by plastid signals««80

Figure 3.10 Representative seedlings and cotyledons from the fluence response experiments«««««««««««««««««««««««««««««««2

Figure 3.11 Analysis of cotyledon opening in seedlings grown on media containing lincomycin or erythromycin«««««««««««««««««««««««««4

Figure 3.12 Cotyledon expansion in treated and untreated gun1-1 and wild-type seedlings«««««««««««««««««««««««««««««««««5

Figure 3.13 Analysis of cotyledon expansion in gun1 mutants grown in blue light«7

Figure 3.14 Quantitative analysis of SEMs«««««««...... 188

Figure 3.15 Representative seedlings from Figure 3.8«««««««««««««90

Figure 3.16 Light microscopy of wild type, gun1, cry1, and gun1-1 cry1 cotyledons««««««««««««««««««««««««««««««««.191

Figure 4.1 Kinetic analysis of Lhcb1 and RbcS mRNA expression following a fluence- rate shift«««««««««««««««««««««««««««««««««54

Figure 4.2 The light-regulated transcriptome in lincomycin-treated and untreated seedlings«««««««««««««««««««««««««««««««««55

Figure 4.3 Summary of biological process and cellular component GO terms enriched in particular expression patterns«««««««««««««««««««««««8

Figure 4.4 Summary of biological process and biological response to stimulus GO terms enriched in particular expression patterns««««««««««««««««2

Figure 4.6 Analysis of chlorophyll phenotypes caused by T-DNA insertion alleles following de-etiolation«««««««««««««««««««««««««««8

Figure 4.7 De-etiolation efficiencies of end mutants in various fluence rates«««90

Figure 4.8 Expression of Lhcb1 and RbcS following a fluence-rate shift«««««5

Figure 4.9 Agglomerative clustering of early time points (i.e., 0.5 h and 1 h) and late time points (i.e., 4 h and 24 h)«««««««««««««««««««««««7

Figure 4.10 Distribution of user-defined expression patterns among light- and plastid- regulated genes««««««««««««««««««««««««««««««3

Figure 4.11 Agglomerative hierarchical clustering of significantly enriched GO terms with expression clusters««««««««««««««««««««««««««.306

Figure 4.12 Clustering of GO terms enriched with user-defined expression patterns«««««««««««««««««««««««««««««««««9

Figure 4.13 Analysis of chlorophyll phenotypes caused by mutant alleles of genes that encode light-signaling factors««««««««««««««««««««««««2

CHAPTER 1

INTRODUCTION

The ever increasing demand for low cost food and the need for alternative sources of energy to maintain economic growth throughout the world requires society to continually ask the same question to an ancient problem; can we continue to convert the same amount of solar energy into more useable biomass? From the selection of the first cereal crops from wild grasses to modern industrial farming practices, plant biologist have provided solutions that have lead to an exponential increase in crop productivity and food production. It is predicted that to maintain economic growth and

WKHFXUUHQWTXDOLW\RIOLIHIRUWKHZRUOG¶VSRSXODWLRQ, food production rates will need to increase by at least 50% by 2050 (Murchie et al., 2009). Because the increased productivity associated with the combination of modern breeding and industrial farming is no longer sufficient to meet the future demands for increases in productivity, science has turned to the potential of genetic engineering to answer the question; can the fundamental process of photosynthesis be further optimized to harvest more light energy and more importantly covert that energy into usable biomass? There are several lines of evidence that argue that the potential of photosynthetic output in crop plants has not reached it theoretical limits (Murchie et al., 2009). Many of the metabolic processes that are predicted to be targets for optimization are carried out in the chloroplast. Here we analyze the chloroplast, but not solely for its role as the site of photosynthesis, but as the source of quantitative information that enables the plant to couple photosynthetic output with its environment, and its growth and development.

Ultimately the goal is to move toward a better understanding of the biology of the

chloroplast and expand what is currently known about the mechanisms that plants use to optimize chloroplast biology to maximize light capture and photosynthetic output.

ADAPTING TO THE LIGHT

Light is arguably the most important environmental cue to shape plant development. To maximize photon capture, growth and development, and reproduction, plants must be able to continually adapt to a complex and ever changing light environment. The unobstructed spectral distribution of sunlight falls into three general domains: Ultra violet light (<400nm), visible light (400 to 700nm), and far-red light

(>700nm). To respond to the broad spectrum of light, plants are thought to possess distinct photoreceptors for sensing UV-B, UV-A, blue, green, red, and far-red light

(Kendrick and Kronenberg, 1994). The ability to sense various spectral qualities allows plants to optimize photon capture to different light environments, such as under a canopy or to different times of the day. The light spectrum under a canopy is depleted of most of the UV and visible light and enriched in far-red light. Additionally, plants can sense and respond to the intensity, duration and direction of the light using some of the same photoreceptors that sense light quality. Light perception also plays an important role in regulating germination, flowering, and senescence, which are essential for plant survival, but the focus presented here is directed toward understanding the role that light plays in regulating photosynthesis, chloroplast development, and photosynthetic tissue development.

Skotomorphogenesis to Photomorphogenesis

The transition from dark growth or skotomorophogenesis to light-dependent growth or photomorphogenesis is characterized by the most dramatic and evident

morphological changes caused by light, although plants constantly adapt their morphology to maximize photosynthesis. Skotomorphogenic growth is characterized by an elongated hypocotyl, and small, unexpanded cotyledons that contain etioplasts rather than chloroplasts, and appear yellow in color due to the lack of chlorophyll

(Figure 1.1A). During skotomorphogenesis plastids in the cotyledon are in the form of etioplasts, which contain a crystalline structure called the prolaminar body (PLB). The

PLB mostly consists of thylakoid lipids and the chlorophyll precursor protochlorophyllide, which remains bound by protochlorophyllide oxidoreductase (POR) until it is exposed to light. In the dark, genes that encode photosynthesis-related proteins are expressed at very low levels. When a seedling is exposed to light, the photomorphogenic growth pattern rapidly establishes the seedling as a photoautotrophic organism and the seHGOLQJ¶VHQHUJ\LVGLUHFWHGWRFRW\OHGRQDQGOHDIGHYHORSPHQWDQGQRWK\SRFRW\O elongation. Light induces the expression of genes that encode photosynthetic proteins and etioplasts develop into functional chloroplasts (Chen et al., 2004).

Plant responses to light intensity are generally divided into three categories: very

-2 low fluence (VFL) responses, which require as little as a single pulse of 100 pmol m ,

-2 low fluence (LF) responses, which require multiple pulses adding up to 1mmol m , and high irradiance (HI) responses, which requires prolonged exposure to light in excess of

-2 10 mmol mol . VLF responses include the initiation of transcription of certain photosynthetic genes and the initiation of germination (Fankhauser and Chory, 1997,

Botto et al., 1996). Low fluence responses include the initiation of de-etiolation and greening of the cotyledons (Fankhauser and Casal, 2004). High irradiance responses

result in a continuous developmental gradient of responses depending on fluence quality and quantity, and are generally the most commonly measured responses that lead to morphological changes such as the inhibition of hypocotyl elongation, cotyledon opening, cotyledon expansion and anthocyanin accumulation (Fankhouser and Casal,

2004). As fluence rates of photosynthetically active light increase, more of the

VHHGOLQJ¶VHQHUJ\LV distributed into cotyledon expansion and diverted away from hypocotyl elongation (Figure 1.1A). At high fluence rates the photosynthetic capacity of the seedling is saturated and excess photon capture leads to photoinhibition, and the seedling invests its energy resources into the accumulation of sunscreens like anthocyanin, carotenoids, isoprenoids, and flavonoids (McNellis and Deng, 1995,

Pogson et al., 2008) (Figure 1.1A).

Optimizing growth to maximize photon capture

Although the initial morphological changes evident in young seedlings are perhaps the most dramatic and the most well studied, plants continually adapt to light throughout their life cycle. Light environments can vary in the amount of photosynthetically active light that is available. To compete for sunlight plants use stems, trunks and vines to position leaves out of the canopy of other plants. Under a dense canopy most of the photosynthetically active light is absorbed by chlorophyll in leaves at the top of the canopy, leaving only far-red light to the leaves at the lower levels (McNellis and Deng, 1995). To optimize electron capture plants adjust their leaf anatomy to their light environment. When grown in unobstructed sunlight plants have

relatively thick leaves, that contain multiple layers of palisade cells, which are dense in chlorophyll (Weston et al 2000). The light that passes through the thick palisade cell layer is then scattered and collected by the spongy mesophyll cell layer (Figure 1.1B).

In shade-grown plants, the palisade layer is reduced and thus the leaf is thinner allowing for the same amount of light-harvesting tissue to be extended over a larger area (Figure 1.1B). Shade-grown plants also invest more resources in light harvesting antenna complexes, where sun grown plants have more reaction centers and higher levels of Rubisco (Anderson et al., 1995). The molecular mechanism that controls the morphological changes and optimizes leaf structure to its light environment is poorly understood. There is evidence that blue light plays a major role in this process, however, mutants with defects in blue light photoreceptors cry1, cry2, and phot1 did not have altered leaf structure (Weston et al., 2000). One possible explanation is that signals are derived from the chloroplast indirectly measure light intensity by the over reduction of the plastoquinone pool; these signals have been shown to regulate Lhcb

(Light harvesting complex B) expression, but these signals have not been connected to the control of leaf anatomy (Escoubas et al., 1995).

In C3 plants, leaf photosynthesis is saturated at about 25% of the maximum full

-2 -1 sunlight (1000-2000 µmol m s ), and thus any photosynthetically active light that is intercepted above is level is wasted (Long et al., 2006). Therefore plant architecture becomes an important trait to maximize light capture. Although plants that maintain a large horizontal canopy may have a competitive advantage by preventing other plants from becoming established, they are also limiting their own productivity by shading their

lower leaves (Long et al., 2006). To maximize photosynthesis, a leaf architecture that positions upper leaves at a more vertical angle allows for over half of the sunlight to reach lower leaves (Figure 1.1C). By distributing the solar energy across multiple leaves a plant would achieve over double the efficiency of light energy use when compared to a horizontal canopy plant (Long et al., 2006). This aspect of optimizing leaf architecture is especially important in monoculture crop species where competition from non-crop plants is controlled by modern industrial farming practices.

Phototropisms

Although plants are sessile organisms, they have the ability to reorient their leaves and chloroplasts to grow toward photo synthetic ally active light, and away from shade to optimize solar capture. When under a partial canopy plants have the ability to grow toward the pockets of photosynthetically active unidirectional light. Phototropisms were initially described by Charles Darwin (Holland et al., 2009). Phototropism enables plants to bend toward incident light to optimize light absorbtion. Phototropisms are mediated by a family of blue light photorecetors called phototropins (Holland et al.,

2009). After perception of unidirectional light the phototropins initiate a signal transduction cascade that ends with the redistribution of auxin that promotes cell elongation on the side of the plant that faces away from the light (Holland et al., 2009).

Another phototropism that is mediated by phototropins is solar tracking. Solar tracking allows the leaf to move perpendicular to the incident light to maintain the highest level of light absorption and thus maximize photosynthesis (Figure 1.1D). Solar-tracking allows

plants to follow the movement of the sun across the sky during the day to maximize solar capture (Koller, 2000).

Phototropins are also required for chloroplast and stomatal movements. In low light or a shaded canopy, chloroplasts distribute evenly throughout the cells in the leaf to maximize the capture of incident light, but in full sunlight the chloroplasts move to the periphery of the cell, where they shade each other and thus minimize excess light absorption (Figure 1.1E) (Christie, 2007). During the day, stomata open to allow for the gas exchange required for photosynthesis and carbon fixation, but during the night if the stomata remained open they would allow for excess water loss. Phototropins in the guard cell coordinate stomatal opening with photosyntheticlly active light (Christie,

2007).

The affect of light quality on development

The photosynthetically active light spectrum is mostly dependent on the absorption spectrum of chlorophyll, although the carotenoids expand the wavelengths that are collected for photosynthesis. Chlorophylls primarily absorb in the blue and red regions of the visible light spectrum. To couple photosynthetic light with photosynthetic tissue development, plants have developed multiple photoreceptors that initiate strong morphogenic changes in the blue and red regions of the visible spectrum. Perception of red and far-red light is mediated by the phytochrome family of photoreceptors. In

Arabidopsis, five distinct genes designated PHYA, PHYB, PHYC, PHYD, and PHYE encode the apoproteins (Quail et al 1995). phyA is light labile, and is expressed at high

levels in the dark. Upon exposure to light phyA levels drop 100 fold (Quail et al., 1995). phyA is solely responsible for the perception of far-red light, and phyB is the primary photoreceptor in continuous red light (McNellis and Deng., 1995). Although phytochromes perceive blue light, the cryptochromes are the primary blue light photoreceptors that drive blue-light-mediated photomorphogenesis. In Arabidopsis the cryptochromes are encoded by two genes, CRY1 and CRY2. During photomorphogenesis cry1 mediates high fluence rate responses and cry2 mediates low fluence rate responses (Lin et al., 1998). Unlike phytochromes, cryptochromes do not obey the law of reciprocity, meaning they only sense fluence rate and not total amount of light perceived (Neff et al., 2000).

Continuous red light is the most effective wavelength for inducing cotyledon expansion and chloroplast biogenesis in developing seedlings, but it is not as effective as blue or far-red light at inhibiting hypocotyl elongation (Figure 1.1F) (McNellis and

Deng, 1995). Continuous far-red light is effective at inducing cotyledon opening and expansion and at higher fluence rates it can strongly inhibit hypocotyl elongation, but the far-red wavelength is too long for the light-dependent conversion of protochlorophyllide to chlorophyllide and therefore the seedlings are unable to green and carryout photoautotrophic growth (Figure 1.1G) (McNellis and Deng, 1995).

Because continuous blue light stimulates both phytochromes and cryptochromes it is a very effective inhibitor of hypocotyl elongation and inducer of cotyledon expansion

(Figure 1.1H) (McNellis and Deng, 1995).

The ability of plants to perceive multiple wavelengths of light, allows plants to have a better sense of time and place. At dusk sunlight is enriched in longer

wavelengths of light. Therefore the perception of the higher ratio of far-red to red light allows plants to perceive the end of the day and adjust and maximize their metabolism and development. The understory of a canopy also has a high ratio of far-red to red light, and plants alter their development to avoid the shade as well as to maximize the capture of small amount of light that is available. End-of-the-day and shade-avoidance responses lead to early flowering, long stems, and longer and wider leaves (Figure 1I and 1J) (Neff et al., 2000).

LIGHT-REGULATED SIGNAL TRANSDUCTION NETWORKS

The regulation of plant growth has evolved to maximize photon capture and photosynthetic output. Because chlorophyll is the light-absorbing pigment that captures photons for photosynthesis, the photosynthetic output of a plant is highly dependent on the absorption spectra of chlorophyll, and peak photosynthetic output is seen in the blue and red regions of the light spectrum (Figure 1.2A).

Photoreceptors

As mentioned above three classes of photoreceptors are responsible for most of the developmental changes caused by light. Phytochromes and cryptochromes are the primary photoreceptors that mediate hypocotyl inhibition in developing seedlings. The combined action spectrum of these two classes of photoreceptors on the inhibition of hypocotyl length closely follows the action spectrum of photosynthesis (Figure 1.2B).

By perceiving the same light qualities that are active in photosynthesis, cryptochromes and phytochromes can more quickly and accurately adapt various aspects of metabolism and development to changing light conditions, than if light perception was simply controlled by the feedback from photosynthesis.

The absorption spectrum of phytochrome is similar to chlorophyll, because the chromophore, phytochromobilin 3Ɏ% LVGHULYHGIURPWKHsame tetrapyrrole biosynthetic pathway that synthesizes chlorophyll. Unlike chlorophyll, which is a tetrapyrrole ring, phytochromobilin is a linear tetrapyrrole that isomerizes, like a switch, between Pr and Pfr forms. The absorption spectrum of the Pfr form is shifted to the far-

red light spectrum when compared to the Pr form (Figure 1.2D). Upon absorption of far- red light the Pfr form isomerizes to the Pr form, which subsequently can convert back to

Pfr form upon absorption of red light (Neff et al., 2000). The Pr form is only active in type I phytochromes (PHYA in Arabidopsis), whereas in type II phytochromes the Pfr form is active (PHYB-PHYE in Arabidopsis). The conversion from the Pr to Pfr form of phytochromobilin leads to conformational changes in phytochrome protein. This conformational change is important for signal transduction with the proteins that interact and phosphorylate the phytochromes. Phosphorylation of phyA controls its subcellular localization, its stability and its affinity toward downstream signal transducers (Jiao et al.,

2007). Additionally, phytochromes have intrinsic kinase activity at the C-terminal kinase-related domain (Yeh and Lagarias, 1998), but when this domain is truncated from PHYB, the holoprotein is still functional in signaling (Matushita, 2003).

Cryptochromes absorb blue light with with dual chromophores made up of a flavin adenine dinucleotide (FAD) and a pterin. The two chromophores are bound by a domain that has a similar structure to blue/UV light activated DNA photolyases (PHR domain) (Figure 1.2E). Like DNA photolyases cryptochromes absorb light in the blue and UV spectrum (Figure 1.2E). Photoactivation of the N terminal domain causes a conformational change that drives the autophosphorylation of the C-terminal CCT/DAS domain (Sang et al. 2005) and dimerization of cry1. Dimerization affects its interaction with downstream signal transduction components and shifts the distribution between nucleus to the cytoplasm (Jaio et al., 2007).

Because phototropins are the photoreceptors that mediate phototropisms in plants, the action spectrum of phototrophic movement closely follows the absorption

spectrum of the two flavin mononucleotide chromophores (FMN) bound by the phototropin proteins (Figure 1.2C and 1.2F). The two chromophores are bound by two

LOV (light-oxygen-voltage) domains, LOV1 and LOV2. The first LOV domain is thought to be important for dimerization and attenuation of the signal, and the second LOV domain is important for inhibiting the kinase activity in the dark (Kimura and Kagawa,

2006) (Figure 1.2F). C-terminal Ser/Thr kinase domain is important for the autophosphorylation of the phototropins, which is essential for signal transduction, but the phosphorylation of downstream components has not been discovered (Demarsy and Fankhauser et al. 2009).

There is one more class of photoreceptors that perceives blue light. The

Zeitlupe/FKF1/LKP2 family of photoreceptors have FMN chromophores that bind LOV1 domains at the N-terminal end of the protein similar to the phototropins. The C-terminus contains an F-box domain and 6 Kelch repeats. The F-Box domain is important for the ubiquitination and the subsequent degradation of targets of the ZTL1/FKF1/LKP2 family

(Demarsy and Fankhauser et al., 2009). The interaction between FKF1 and GI mediate the light-dependent degradation of CYCLING DOF FACTOR1 (CDF1), which represses

CONSTANS expression, a positive regulator of flowering (Imaizumi, 2010). The

ZTL1/LPK2 interation with GI leads to the degradation of TOC1, which is a central component of the circadian clock (Imaizumi, 2010).

Regulation of transcription by light

In Arabidopsis and rice, at least 20% of the transcriptome is regulated by light.

Most of the major metabolic pathways in the plant cell are coordinately regulated by the light (Ma et al., 2001, Jiao et al., 2005 $OPRVWDOORIWKHOLJKW¶VDIIHFWRQWKH transcriptome is downstream of the photoactivation of the phytochromes and cryptochromes. The phototropins only affect the transcriptional regulation of a limited number of genes (Jiao et al., 2007).

The most widely studied light responsive gene is the gene that encodes the Light harvesting complex B protein (Lhcb), because it is robustly induced by the light. Lhcb is essential for chloroplast development and photosynthesis, it is coordinately regulated with most genes that encode proteins involved in photosynthesis, and it is one of the most abundant proteins in the plant cell. The light responsive genes are regulated by promoter elements known as light responsive elements (LRE). The most well understood and one of the most abundant LREs is the G-box and its variants, although no single element has been found in all the light-regulated promoters (Jiao et al., 2007).

The G-box is generally described as having the consensus sequence of CACGTG, which has been pared down since it was initially described by Giuliano et al., 1988. The

G-box has been shown to be bound by several different transcription factors in vitro and in vivo (Jiao et al., 2007). Two G-box binding factors which have dramatic affects on photomorphogenesis are the bZIP transcription factors HY5 (LONG HYPOCOTYL5) and its homolog HYH (HY5 HOMOLOG) (Holm et al. 2002), which bind and regulate a number of genes in vivo (Lee et al., 2007), and the PIF (PHYTOCHROME

INTERACTING FACTOR) family of bHLH transcription factors that act as repressors of photomorphogenesis (Shin et al., 2009). HY5 is a positive regulator of

photomorphogenesis under a broad spectrum of light, suggesting that it acts downstream of both the phytochromes and cryptochromes (Chory, 1992), and HY5 has been shown to regulate 20% of all light-regulated genes (Ma et al., 2002). Additionally

60% of the genes regulated by phytochromes after 1h of light exposure are bound by

HY5 (Lee et al., 2007). HY5 is activated transcriptionally and post-translationally by light (Jiao et al., 2007), which further propagates the HY5-dependent induction of photomorphogenesis. The best understood mechanism for the regulation of HY5 is that in the dark HY5 is degraded by COP1 (CONSTITUTIVE PHOTOMORPHGENIC1).

Photoactivated cry1 inhibits COP1 activity in the nucleus and initiates the translocation of COP1 to the cytoplasm away from positive regulators of photomorphogenesis like

HY5 (Jiao et al., 2007) (Figure 1.3). HY5 is considered to be positioned relatively high in the hierarchy of transcription factors that regulate photomorphogenesis (Chen et al.,

2004). The mechanism by which phytochrome signaling acts through HY5, is not well understood. The phytochrome regulation of photomorphogenesis through the PIFs is better understood. In the dark, PIFs bind the G-box and prevent positive regulators like

HY5 from binding (Shin et al. 2009) (Figure 1.3). Upon exposure to light, phyB transolocates from the cytoplasm to the nucleus where it colocalizes to nuclear speckles with HEMERA (HMR) (Chen et al., 2010). HEMERA is structurally similar to multiubiquitin-binding protein RAD23 in yeast, and can partially rescue yeast RAD23 mutants. Additionally, HEMERA mutants accumulate PHYA, PIF1 and PIF3 in the light, in wild-type PIF1 and PIF3 are phosphorylated and degraded in the presence of photoactivated phytochromes, and PHYA is light labile (Chen et al., 2010) (Figure 1.3).

Thus HEMERA appears to mediate phytochrome-dependent proteolysis. Another

interesting aspect of HEMERA is that unlike all other light signaling mutants HEMERA is albino and is unable to develop chloroplasts. In addition to the nucleus, HEMERA also localizes to the chloroplast and is known to associate with the plastid transcription machinery (Pfalz et al., 2006). Both the nuclear and chloroplast fractions of HEMERA are required for chloroplast biogenesis (Chen et al., 2010). The nuclear fraction of

HEMERA appears to promote chloroplast biogenesis by degrading the PIFs. PIFs are known to strongly repress the expression of HEMA1 and GUN5, which are required for chlorophyll biosynthesis and LHCA1 and psaE1, which encode subunits of PSI (Shin et al., 2009). The pif1 pif3 pif4 pif5 quadruple mutant is constitutively photomorphogenic in both the light and the dark, which indicates that this family of transcription factor plays a major role in repressing photomorphogenic growth and chloroplast biogenesis, and that there is significant activity of positive regulators in the dark even when COP1 is active

(Shin et al., 2009). Because HY5 and the PIFs essentially bind the same promoter element, the removal of the PIFs by photoactivated phytochrome would indirectly activate HY5 and thus explain the mechanism that phytochromes regulate HY5 transcription.

The role of protein degradation in regulating photomorphogenesis

As discussed above, turnover of HY5, PIF1 and PIF3 plays a critical role in regulating light-dependent transcription. Our understanding of the turnover process was initially revealed by a class of mutants that carried out photomorphogenic growth in the dark. The cop/det/fus are a class of mutants that show open and expanded cotyledons,

inhibition of hypocotyl elongation, but cannot develop chloroplasts because of the light requirement of protochlorophyllide oxidoreductase (POR), although the etioplasts do develop thylakoid structures (Chory and Peto, 1990). At the genetic level the

COP/DET/FUS mutants were described as making up D³FHQWUDOSURFHVVRU´of the light- dependent reprogramming that leads to photomorphogenic growth (Figure 1.4)

(McNellis and Deng, 1995). Later it was discovered that COP1, COP9, COP10, and the

FUS (FUSCA) proteins are important for the ubiquitin mediated degradation of proteins.

Although first discovered in plants the COP9 signalosome (CNS) or proteosome is found in all prokaryotes and eukaryotes, and is made up of a complex of proteins that were initially described based on studies of fus mutants, and is the eventual site of degradation of all E3 ubiquitin ligase targets (Wei et al., 2008). Based on its strong mutant phenotype COP1 has always been considered the major E3 ubiquitin ligase regulating photomorphogenic growth, but the more recent discovery of HEMERA suggests that COP1 may only be part of the story. Additionally, the F-box protein, EID1

(for Empfindlicher Im Dunkelroten Licht)KDVEHHQVKRZQWR³ILQHWXQH´WKHOLJKWVLJQDO, but unlike cop1 mutants eid1 mutants require light before they have a visible phenotype, which is hypersentivity to far-red and blue light (Dieterle et al., 2000). Because eid1 mutants accumulate the same level of photoactivated PHYA as wild type, it is thought that EID1 acts on downstream signaling components of PHYA signaling cascade

(Buche et al., 2000) (Figure 1.4). Like the eid1 mutant, spa1 mutant (SUPRESSOR OF

PHYTOCHROME A-105) is also hypersensitive to light (Hoecker et al., 2005). SPA1 is a member of a four gene family in Arabidopsis. The quadruple spa1 spa2 spa3 spa4 mutant is constitutively photomorphogenic. Additionally, SPA1 interacts with COP1 in

vivo. Members of the SPA gene family have distinct but overlapping functions. Taken together it appears that the SPA proteins give specificity to the central regulator COP1

(Hoecker et al., 2005) (Figure 1.3).

Chromatin remodeling

Another important aspect of the molecular regulation of photomorphogenesis is chromatin remodeling. Chromatin remodeling does not appear to be affected by light quality, but acts as a general mechanism to ensure that photomorphogenic genes are

³RII´LQWKHGDUN -LDRHWDO ,WZDVLQLWLDOO\GHVFULEHGWKDWKLVWRQHDFHW\ODWLRQ allowed for increased nuclease accessibility of the promoter of the pea plastocyanin gene (Chua et al., 2001). Additionally, repression of photomorphogenesis is observed in two histone acetylatransferase mutants haf2 and gcn5, which are hypoacetylated. In the histone deacetylase hd1 mutant, which is hyperacetylated, photomorphogenesis is promoted (Bertrand et al., 2005 and Benhamed et al., 2006). The additional importance of histone modification is suggested by the constitutive photomorphogenic mutation in

DE-ETIOLATED1 (DET1). In tomato DET1 interacts with the non-acetylated tail of histone H2B (Benvenuto et al., 2002). In vivo DET1 forms a complex with the E2 ubiquitin ligase, COP10, and DAMAGED DNA BINDING PROTEIN 1 (DBB1). DBB1 is able to bind DNA and is thought to aid in the recruitment of histone acetyltransferases.

The COP1 functions together with the DET1/COP10/DDB1 complex to promote the degradation of proteins like HY5 (Yanagawa et al., 2004) (Figure 1.3). Taken together it appears that histone acetylation has two functions; first chromatin acetylations opens

the chromatin to allow transcriptional activators and repressors access to the promoters of light-responsive genes, secondly through the DET1/COP10/DDB complex chromatin modification recruits COP1 to the chromatin to enhance the degradation of factors like

HY5 in the dark.

The light signaling network

Based on phenotypic analysis of mutant alleles, factors like HY5, the PIFs, COP1,

HEMERA, and DET1 appear to be central components of the light signaling network that regulates photomorphogenesis. But several other factors have been discovered that provide the plant with more specific information from the light signaling network, such as light intensity, quality, direction, time, and place. Transcription factors such as

HFR1, and LAF1 are positive regulators of gene transcription like HY5 and are also targets of COP1 for degradation, but unlike HY5, which is active in all fluence qualities, they are primarily active in far-red light (Figure 1.4) (Jiao et al., 2007). Other signaling components like FHY3 and FAR1 are also specific to far-red light responses. Other signaling components appear to modulate blue light responses. SUB1, PP7, and GBF1 are positive regulators of blue light responses (Guo et al., 2001, Moller et al., 2003,

Schindler et al., 1992) (Figure 1.4). HRB1 contributes to the fluence quality response to attenuate the light response to blue and red light (Kang et al., 2005). The transcription factor MYC2 is a repressor of blue light signaling but also appears to play a role in jasmonic acid and abscisic acid signaling, potentially participating in cross talk between light and stress hormone signals (Yadav et al., 2005). As mentioned above, plants

perceive light direction through the phototropins. Downstream components of the phototropins such as NPH3 and RPT2 appear to connect the photoreceptors with transcription regulation (Kimura and Kagawa, 2006). Because phototropism leads to the redistribution of auxin in the plant, several components of the auxin signaling network, such as ARF7, TIR1, and IAA19, have been found to be aphototrophic in the corresponding mutants (Kimura and Kagawa, 2006). To accurately match growth and metabolism with the time of day, plants maintain a circadian clock, which consists of 4 primary cyclic regulatory loops; the CCA1/LHY-PRR7/9 loop, the CCA1-TOC1/CHE loop, CCA1/LHY-TOC1 loop and the TOC1 loop (Imaizumi et al., 2010). These loops

PDLQWDLQWKHFORFN¶VWLPLQJEXWWKLVWLPLQJLVFRXSOHGWROLJKWVLJQDOLQJWKURXJKLWV regulation by COP1. Interestingly, CRY2 appears to have more affect on circadian control than CRY1 and the phytochromes. As previously mentioned the circadian clock also couples light perceived by the ZTL1/LKP2/FKF2 class of photoreceptors to the time of day (Imaizumi et al., 2010).

CHLOROPLAST DEVELOPMENT

The transition from autotrophic growth to photoautotrophic growth is arguably one of the most important steps in a SODQW¶VOLIHF\FOHIn a mature seed, chloroplasts are not present, only proplastids, and therefore germinating seedlings do not start with a source of chloroplasts that are simply divided and partitioned into newly developed cells.

,QVWHDGFKORURSODVWVGHYHORSIURPSURSODVWLGVDQGHWLRSODVWV7KHWHUP³SURSODVWLGV´ generally refers to plastids that are small, undifferentiated, and are simple in structure, but this term is not well defined in the literature and likely encompasses plastid types with significantly different functions (Wise and Hoober, 2007). Proplastids that have the ability to differentiate into chloroplasts are sometimes referred to as eoplasts, or germinal proplastids to differentiate them from other nonspecific proplastids types

(Wise., 2007) Chloroplasts only develop from germinal proplastids in the developing embryo and at the shoot apical meristem. (Figure 1.5B). The extent to which photosynthetic tissue development is coupled to chloroplast development in the shoot apical meristem is not well understood. When the light releases the shoot apical meristem from growth arrest there is a rapid upregulation the cytokinin and gibberellin responses prior to leaf emergence (Lopez-Juez et al., 2008). Interestingly, ent-kaurene synthase (KS) activity, which is required for gibberellin biosynthesis, only colocalizes with developing tissues that contain proplastids and not tissues that contain chloroplasts

(Aach et al., 1997), and localization of KS by organeller fractionation found that it was enriched in the plastid stroma (Aach et al., 1995). Additionally, mutants that are unable to develop chloroplasts, or seedlings treated with inhibitors of chloroplast biogenesis,

are only able to develop leaves that contain undifferentiated cells and they do not have the characteristic ultrastructure of a photosynthetically active leaf (Yu et al., 2007).

When photosynthetic tissues are grown in the dark germinal proplastids develop into etioplasts. The defining structure of the etioplast is the prolamellar body (PLB), which is a tetrahedrally branched matrix of membranes that contain carotenoids, protochlorophyllide, galactolipids, and proteins. Upon exposure to light the PLB is quickly converted into thylakoid membranes, and the developed chloroplast is formed

(Figure 1.5A). Although the transition from germinal proplastids to chloroplasts maybe more common in nature, etioplast have been used extensively to study chloroplast development, because the transition from etioplast to chloroplast is easily controlled by the presence of light, the plastid development state is uniform in all photosynthetic tissues, it occurs rapidly, and it occurs in a measurable step-wise process (Wise and

Hoober, 2007).

Other Plastid types

Although chloroplasts receive the most attention, plastids can differentiate into multiple different plastid types. Most of the plastid types that have been characterized are differentiated based on their ultrastructure and metabolic function. Leucoplast is a general term that describes a non-pigmented plastid. Leucoplasts are diverse in function and agriculturally important. For example, leucoplast in the basal and stalk cells of the trichome are the site of the early steps of monoterpene biosynthesis in mint

(Figure 1.5C) (Turner et al., 1999). Another agriculturally important leucoplast is the

amyloplast. This plastid type is the site of the complete starch biosynthetic pathway (Yu et al., 1998), and the site of starch storage. Although diurnal starch synthesis is important for maintaining metabolism in photoautotrophic tissues, starch synthesis in non-photosynthetic tissues like fruits, seeds, tubers, and stems is important for the long term storage of starch. Amyloplasts in these storage tissues contain several large starch grains with minimal internal membranes (Figure 1.5D). Amyloplasts also play a role in gravity perception. Because starch grains are more dense than water, amyloplasts sediment in the direction of the gravity, which is perceived by the root tip cells of the plant (Figure 1.5D) (Wise and Hoober, 2007). Leucoplasts can also accumulate oil. These oil dense plastids are called elaioplasts. Although the plastid is the site of fatty acid synthesis in plant cells, elaioplasts are not the site of oil storage in mature oilseeds, which is stored in oleosomes. Elaioplasts are important for pollen maturation, where they temporally store the oil for the production of sterol lipids that coat the pollen grain (Figure 1.5E) (Hernandez-Pinzon et al., 1999).

Unlike leucoplasts, chromoplasts are brightly colored, and are found in various fruits, flowers, leaves and roots, and are used in part to attract pollinators and fruit- dispering animals. Chromoplast often develop from chloroplasts in fruits and leaves, but they can also develop from proplastids in other tissues like tubers (Figure 1.5F)

(Wise and Hoober, 2007). Chromoplasts accumulate large amounts of carotenoids in lipid droplets and fibrils (Figure 1.5F) (Deruere., et al 1994). Chromoplasts are

DJULFXOWXUDOO\LPSRUWDQWEHFDXVHWKH\DUHULFKLQȕ-carotene (e.i. carrots), and lycopene

(e.i. tomato), which make up a significant source of antioxidants in the human diet

(DellaPenna and Pogson, 2006). Because chloroplasts contain approximately 75% of

the total leaf protein, plants dismantle the chloroplast during senescence to regain some of the resources tied up in the organelle during photoautotrophic growth. These senescing chloroplasts have been named gerontoplasts. Gerontoplasts are characterized by the unstacking of the grana, loss of thylakoid membranes, and an accumulation of plastoglobuli, which are thought to contain the lipid-protein remnants of the thylakoids (Figure 1.5G) (Wise and Hoober, 2007). Another agriculturally important plastid type has the appearance of a proplastid, but is found in the nodules of plants that through the symbiotic relationship with bacteria, fix nitrogen. The main metabolic function of the nodule proplastids is to carry out the GS-GOGAT cycle, which produces amino acids from the glutamine that is derived from the ammonia assimilated by the symbiotic bacteria (Figure 1.5H) (Wise and Hoober, 2007).

High and low light chloroplast development

As mentioned earlier plants experience a wide verity of light environments depending on their location in the canopy, time of day, or cloud cover. Chloroplasts adjust their developmental stage to optimize light capture and carbon fixation. In low light or shade conditions, light becomes the limiting factor for carbon fixation and in high light or full sun conditions CO2 concentrations become the limiting factor for carbon fixation (Figure 1.6A). To maximize light capture in low light a greater proportion of the chloroplasts¶ resources are placed into light-capture, and they have more grana, more thylakoids per grana, and more light-harvesting antenna (Figure 1.6B). In high light more of the chloroplasts¶ resources are placed into carbon fixation and Rubisco, and to

avoid the over reduction of the photosynthetic electron transport (PET) chain the light- harvesting antenna is down regulated and there are fewer grana and fewer thylakoids per grana (Figure 1. 6C) (Weston et al., 2000). The control of this morphology shift is thought to be controlled by the light and retrograde signals derived from the chloroplast

(Pfannschmidt et al., 2009). The regulation of the shift from low light chloroplast morphology to high light chloroplast morphology would be consistent with its regulation by the golden-like transcription factors (GLK). The Arabidopsis glk1 glk2 double mutant is pale green and the chloroplasts contain few grana stacks (Fitter et al., 2002). When

GLK genes are overexpressed in the double mutant background, the resulting plants have more grana stacks and more chlorophyll than wild-type (Waters et al., 2008,

2009a). Interestingly, in these plants it was found that the */.¶s specifically upregulate the genes that encode the proteins required for chlorophyll biosynthesis and the light harvesting complex, but they do not affect the regulation of genes required for the

Calvin cycle (Waters et al., 2009a 7KH*/.¶s are induced by the light and are regulated by plastid development signals (Waters et al., 2009a). Taken together the data would be consistent with a model LQZKLFKWKH*/.¶VDUHWKHSULPDU\UHJXODWRUVRI chloroplast morphology in response to high and low light. In low light GLKs are induced by the light and upregulate chlorophyll synthesis and the light harvesting complex leading to more grana. In high light, plastid retrograde singles down regulate GLK activity and thus there is less chlorophyll synthesis and light harvesting complex, but the

Calvin cycle is continually induced by the light (Figure 1.6 B-D) (Waters et al., 2009b).

Dimorphic chloroplasts are required for C4 photosynthesis

In full sun the concentration of CO2 availability becomes the limiting factor for photosynthetic output. To increase the efficiency of rubisco, CO2 is initially fixed in the mesophyll cells by phosphoenolpyruvate carboxylase to a C4 acid and then transported to the bundle sheath cells where it is decarboxylated back into CO2, which is then assimilated into the Calvin cycle by rubisco. In bright sun, the difference in photosynthetic output of a C4 plant compared to a C3 plant is about 30-50% greater depending on the temperature (Henning and Brown, 1986) (Figure 1.6D). The metabolism of the bundle sheath and mesophyll cells are distinct. In C4 plants, mesophyll cells accumulate PSII and therefore have many stacked grana. Bundle sheath cells only accumulate PSI and therefore lack stacked grana. Bundle sheath cells also accumulate more starch granules. Additionally the chloroplasts of C4 bundle sheath cells contain a peripherial reticulum (PR), which is a system of tubules and saccules that are frequently attached to the inner envelope. The function of the PR is not well understood, but they have been proposed to be important for metabolite transport (Wise and Hoober, 2007). Because of the morphological differences of the chloroplast between the two cell types, these chloroplasts are called dimorphic chloroplasts (Figure 1.6E). Although the mechanism that regulates chloroplasts

GLPRUSKLVPLVQRWZHOOXQGHUVWRRGWKH*/.¶VDSSHDUWREHLPSRUWDQWIRUWKHUHJXODWLRQ

of the photosynthetic development differences that lead to dimorphic chloroplasts

(Langdale and Kidner, 1994, Rossini et al., 2001).

Mutants in chloroplast development

Our understanding of the mechanisms that are required for chloroplast assembly has mostly come from mutants that are impaired in chloroplast development and from the use of inhibitors and herbicides that specifically block plastidic processes that are required for chloroplast development. Defects in chloroplast biogenesis, whether caused by a mutation or induced by an inhibitor, lead to several different classes of phenotypes. Each phenotype provides information about the role that each defective process has in chloroplast biogenesis.

Variegated mutants produce leaves with white and green sectors. In the white sectors the cells are deficient of developed chloroplasts, and in the green sectors the chloroplast develop normally (Figure 1.7). The variegated phenotype can be caused by mutations in nuclear, chloroplast and mitochondrial genomes (Yu et al., 2007). The mutations that lead to the variegated phenotype have provided information about the molecular mechanisms that are important for the transition from proplastid to chloroplast in shoot apical meristem. In the variegated mutant immutans (im), white sector formation is proportional to the amount of light that is received by the shoot apical meristem. IM encodes a plastid terminal oxidase (PTOX) that shares similarity with the mitochondrial alternative oxidase (AOX). IM prevents the over reduction of the plastoquinone (PQ) pool by PSII and phytoene desaturase (PDS) (Peltier and Cournac,

2002). This function appears to be more important during chloroplast biogenesis, because downstream electron transport components are not present to the same extent as in green tissue, and carotenoids, synthesized by PDS, are essential for the assembly of the photosynthetic complexes in the thylakoid membrane (Yu et al., 2007). Like IM,

VAR3 might be important for the regulation of carotenoid synthesis, and in the var3 mutant, the threshold of cartenoid accumulation is not reached during chloroplast

ELRJHQHVLV 1पVWHGHWDO Two other variegated mutants var1 and var2 have been shown to encode FtsH metalloproteases (Chen et al., 2000, Sakamoto et al.,

2002). VAR1 and VAR2 appear to be important for the turnover of damaged D1 subunit of PSII, in the var1 and var2 mutants damaged D1 accumulates and the chloroplast become photo-oxidized (Lindahl et al., 2000). Like IM it appears that VAR1 and VAR2 are important for maintaining a photo-oxidative-damage threshold during chloroplast development.

Virescent mutants appear to carry out chloroplast biogenesis at a slower rate than wild-type and thus have a range of chloroplast development states in developing leaf tissue. Virescent mutants are characterized by the lack of chloroplast development at the leaf primordia, with the leaves slowly becoming greener as the tissue becomes older (Figure 1.7). Three mutant alleles that cause virescence have been cloned, clpR1, dg1 and ys1 have defects in plastid gene expression or plastid translation (Koussevitzky et al., 2007, Chi et al., 2008, Zhou et al., 2008). DG1 and YS1 are pentatricopeptide repeat proteins (PPR). This class of protein is typically associated with editing transcripts of plastid-localized genes (Chi et al., 2008, Zhou et al., 2009). YS1 is required for the proper editing of RpoB, a subunit of the plastid encoded RNA

polymerase (PEP). In the ys1 mutants plastid transcription is slower than in wild type and therefore chloroplast development is limited by PEP transcription. When grown in the dark and then allowed to de-etiolate ys1 mutants green at the same rate as wild type, suggesting that this mutation does not affect the transition from etioplast to chloroplast (Zhou et al., 2009). Similarly the dg1 mutant has reduced transcript levels of the genes that encode for the PEP subunits, but normal transcript levels of the nuclear- encoded polymerase (NEP) (Chi et al., 2008).

Albino mutants are unable to produce functional chloroplasts at anytime during development (Figure 1.7). Albino mutants are able to maintain a functional proplastids, but are unable to develop chloroplasts (Figure 1.7). Albino mutants provide information on the processes that are required for chloroplast development. Unlike the ys1 and dg1 virescent mutants in which the rate of PEP transcription is slowed, if plastid transcription or plastid translation is completely blocked, chloroplast development is unable to take place. To study chloroplast development scientists commonly exploit this fact and disrupt chloroplast translation with inhibitors. For example, lincomycin specifically inhibits the plastid translation machinery and at high enough concentrations completely blocks chloroplast biogenesis (Mulo et al., 2003). Chloroplast development also requires the import of nuclear-encoded photosynthetic proteins. The protein import mutants ppi1 (Toc33) and ppi2 (Toc159) are specifically unable to import photosynthetic proteins (Kessler and Schnell, 2009). ppi1 and ppi2 are viable as seedlings, but mutants in the core protein import machinery like Tic32 are embryo lethal, because they are unable to maintain a functional proplastid (Hormann et al., 2004). Another mechanism that leads to albinism is the loss of carotenoid synthesis by the use of

inhibitors, like norflurazon an inhibitor of PDS, or carotenoid-deficient mutants (Nott et al., 2006). Additionally, mutants in the mevalonate 2-C-methyl-D-erythritol-4-P (MEP) pathway, like cla1 are unable to carry out chloroplast biogenesis but do have functional proplastids (Mandel et al., 1996). The MEP pathway is required for the synthesis of isoprenes, carotenoids, chlorophyll, and quinones (Estevez et al., 2000). Additionally, mutants that are unable to synthesize thylakoid membrane lipids are albino (Kobyashi et al., 2007). Although most of the mutant alleles that cause albinism have defects in genes that encode plastid-localized proteins, there are exceptions. HEMERA is localized in the nucleus and in the plastid. Both the nuclear and the plastidic forms of

HEMERA are required for chloroplast development (Chen et al., 2010). HEMERA mutants are albino because the transcription of genes required for photosynthesis are strongly repressed in hmr (Chen et al., 2010).

As mentioned before chloroplast biogenesis only occurs in two different tissue types, the shoot apical meristem/leaf primordia and cotyledons. The photosynthetic tissue of the embryos develops into cotyledons and tissues derived from the SAM develop into the true leaves. If chloroplast biogenesis is regulated differently in these two tissues then researchers should be able to isolate mutant alleles that specifically cause chloroplast biogenesis defects in each tissue. Indeed, Arabidopsis mutants that have albino cotyledons and green true leaves, and mutants that have albino true leaves with green cotyledons have been isolated (Figure 1.7). These mutants suggest that the mechanisms by which chloroplast develop in these two tissue types is at least partially distinct. SCO1 (SNOWY COTYLEDON1) encodes elongation factor G, and mutants have reduced plastid translation (Albrecht et al., 2006). SIG6 is a plastid-localized

transcription factor that is expressed during the early stages of development and is important for PEP-dependent gene expression (Chi et al., 2010). SCO2/CYO1 has chaperone like activity and is important for protein disulfide isomerization, and it localizes to the thylakoids (Shimada et al., 2007). The mechanism that leads to cotyledon-specific albinism is not clear, but one explanation could come from genetic redundancy. This is likely the case for SIG6, which is expressed early in development, where other sigma factors that are functionally redundant with SIG6 are expressed later in development (Loschelder et al., 2006). Mutations in ATD2 have the opposite phenotype, where the cotyledons are green and the true leaves are white (Yu et al.

2007). ATD2 encodes ATase2 (amidophosphoribosyl transferase), which is the first committed step in purine synthesis, like sig6 the true leaf specific albanisim could come from tissue-specific expression and functional redundancy with ATase1 (Yu et al., 2007).

One of the most common phenotypes that is caused by a disruption of chloroplast function is a pale phenotype. A few examples are mentioned here.

Mutations that lead to less chlorophyll synthesis are generally pale, depending on the severity of the mutation, chlorophyll mutants can appear almost as green as wild type , such as gun5-1 or albino such as a chlm null mutant (Mochizuki et al., 2001, Pontier et al., 2007). Inefficient assembly of either photosystem I or photosystem II can lead to a pale phenotype. For example the CNFU is a scaffold for iron sulfur clusters and subsequently PSI assembly (Yabe et al., 2004). LPA2 is required for efficient PSII assembly (Ma et al., 2007). HY1 and HY2 (HYPOCOTYL1/2) are required for the synthesis of phytochromobilin, and thus hy1 and hy2 mutants are not able to perceive red light, and subsequently induce chlorophyll and photosynthesis-related genes (Parks

et al., 1991). The dpe1 mex1 double mutant is pale, because it accumulates high levels of maltose, which subsequently induces autophagy of the chloroplast (Stettler et al.,

2009). Although pale mutants are important because unlike albino or variegated mutants these mutations are at least partially tolerated by the chloroplast and do not completely knock out the function of the chloroplast.

The reticulate mutant phenotype is typically characterized by the presence of green vasculature tissue and pale green/yellow lamina (Figure 1.7). This patterning reflects the differences in chloroplast development and dysfunction in the mesophyll versus the bundle sheath cells (Yu et al., 2007). Although several mutant alleles exist that cause reticulate phenotypes, scabrous3 (sca3) and cab underexpressed1 (cue1) are the only two to be extensively characterized. The SCABROUS3 gene encodes

RpoTp, which is a nuclear-encoded plastid RNA polymerase (Hricova et al., 2006). The phenotype caused by the sca3 mutation, indicates that RpoTp-mediated transcription is required for chloroplast development. The reticulate pattern could be due to the redundant activity of RpoTmp (Yu et al., 2007). The CUE1 encodes the plastid phosphoenolpyruvate/phosphate translocator (PPT) (Steatfield et al., 1999). The pale green sectors are a result of the loss of flux into the shikimate pathway. Like sca3 the reticulate pattern in cue1 is likely due to functional redundancy of AtPP1 and AtPP2 which are distinctly expressed in the vasculature and interveinal regions respectively

(Knappe et al., 2003).

PLASTID SIGNALING

Although more is known about the regulation of chloroplast development by extraplastidic signals like the light, biotic and abiotic stress, less is known about the feedback regulation from the plastid. The regulation of nuclear genes that encode chloroplast proteins by extraplastidic signals and subsequent import into the chloroplast is often referred to as the anterograde flow of information. The feedback control of nuclear gene expression by the plastid is often referred to as retrograde control or flow of information. Plastid retrograde signals coordinate nuclear gene expression with the functional and developmental state of the plastid, and coordinate the expression of the plastid and nuclear genomes.

Anterograde and retrograde control of chloroplast development

After years of research and the isolation of multiple mutants in chloroplast development, light signaling, and plastid signaling, the picture of chloroplast development as a continual flow of information is slowing starting to become clearer. In developing embryos, and in the shoot apical meristem, proplastid metabolism and function are maintained in part by tissue-specific programming. Very little is known about the regulation of proplastid function, in part because proplastids are essential, and mutants that do not have functional proplastids are embryo lethal. In Arabidopsis, of 339 non-redundant genes that are required for embryogenesis, about one third are predicted to be localized in the plastid (Hsu et al., 2010). Plastid metabolism is required throughout embryogenesis, whereas plastid gene expression only appears to be

important after the globular stage (Hsu et al., 2010). Proteins required for proplastid function are encoded by the plastid and nuclear genomes. The proteins that are encoded by the nuclear genome are imported through the class I import machinery

(Figure 1.8). The class I import pathway imports proteins that are required for housekeeping functions such as plastid gene expression and non-photosynthetic plastid metabolism (Kessler and Schnell, 2009).

Proplastids are thought to be maintained in part by repressive plastid-to-nucleus signals. These repressive signals couple the functional state of the plastid with nuclear gene expression, and thus strongly repress photosynthesis-associated nuclear genes

(PhANGs). Although it is not clear how these plastid signals are perceived or how they are transmitted to the nucleus, it is known that disruptions in plastid gene expression, plastid translation, photosynthetic protein import, and carotenoid synthesis lead to the production of these signals (Figure 1.9) (Woodson and Chory, 2008, Kakizaki et al.,

2009). The mechanism by which the repressive plastid signal is attenuated to allow for chloroplast development to proceed is not well understood, but all of the conditions mentioned above that lead to the production of plastid signals have one thing in common, they prevent the formation of photosynthetic membranes or lead to dysfunction of the photosynthetic membranes (Figure 1.9). Therefore the production of photosynthetic membranes or thylakoids likely leads to the attenuation of repressive plastid signals.

The proteins required for photosynthesis and subsequently thylakoid biogenesis are encoded by both the plastid and nuclear genomes. As mentioned above, light is required for chloroplast development and is a potent regulator of PhANG expression.

Light up-regulates photosynthetic gene expression by attenuating repressive transcription factors like the PIFs, and inducing the activity of positive trans-acting factors like the GLKs and HY5. Light also induces the expression of nuclear encoded sigma factors, which in turn translocate to the plastid and activate the expression of the core proteins of photosystem I and II, which are encoded by the plastid genome (Figure

1.8) (Waters et al., 2009b). The photosynthetic proteins that are encoded by the nucleus are then imported through the class II protein import machinery, which specifically imports photosynthetic proteins (Figure 1.8). The selectivity of the class II protein import machinery allows for photosynthetic protein import to be regulated independently of plastid housekeeping functions, which are important for maintaining homeostasis during chloroplast development (Kessler and Schnell, 2009). After being imported into the plastid, the photosynthetic proteins begin to assemble with thylakoid membrane lipids to form photosynthetic membranes, but if the assembly is not properly coordinated with the plastid genome, such as in seedlings treated with lincomycin or plastid gene expression mutants, repressive plastid signals are maintained until proper coordination occurs. If seedlings are removed from lincomycin treatment chloroplasts develop properly and the seedlings green (Larkin and Ruckle, 2008). If the assembly of the photosynthetic membranes leads to excessive photooxidative damage, such as in the case of seedlings treated with norflurazon, or in mutants like im, var1, var2 or var3, plastid signals are maintained until the damage is repaired. If thylakoid biogenesis does not occur at least to a minimal extent before the plastids move out of the leaf primordia and into the newly developed leaf, the prolastids never develop into functional chloroplasts, which suggest that chloroplast development requires the cellular

environment provided by the leaf primordia (Yu et al. 2007). In wild type or plants grown in the absence of inhibitors of chloroplast development, thylakoid biogenesis is a relatively efficient process, and as the photosynthetic membranes develop, repressive plastid signals are attenuated (Larkin and Ruckle, 2008). The mechanism by which the plastid signals are attenuated is not well understood, especially because plastid signals repress the genes that encode the thylakoid proteins that appear to be required to attenuate the plastid signal.

After the chloroplasts develop, their functional and developmental state remains coupled to nuclear gene expression. The functional state of the chloroplast is highly influenced by the light environment that it experiences. When plants are exposed to intense light there are three different sources of chloroplast-derived information that couple the functional state of chloroplasts with nuclear gene expression: (1) reactive oxygen species (ROS), such as singlet oxygen, super-oxide, and hydrogen peroxide

(Pogson et al., 2008), (2) the redox state of the plastoquinone pool (Pfannschmidtet al.,

2009), and (3) the functional and developmental state of the thylakoid membranes similar to those in seedlings treated with chloroplast development inhibitors.

Developmental signals likely occur in high light due to the down regulation of photosynthetic proteins and the general thylakoid dysfunction induced by high light stress (Figure 8 and 9). The extent of overlap and functional redundancy between these three processes is not well understood. The role of these signals is to reduce the stress on the chloroplast caused by high light. To relieve stress, these three chloroplast signaling pathways downregulate photosynthetic genes, upregulate sunscreens such as anthocyanins and other flavoniods, upregulate proteins involved in cyclic electron flow

and upregulate ROS scavenging mechanisms (Pogson et al., 2008) (Figure 1.8 and 1.9)

The presence of first two signaling pathways allows the plants to be responsive to the environment, and the presence of the third signaling pathway allows the plants to maintain homeostasis in high light without continually operating with a highly reduced

PQ pool, or continually generating toxic reactive oxygen species.

Plastid signal transduction

One approach investigators use to study plastid signals is to grow the plants in the presence of inhibitors that block chloroplast biogenesis, such as lincomycin or norfluazon. In response to this chloroplast dysfunction, plastid signals potently repress

PhANG transcription (Nott et al., 2006). The genome uncoupled (gun) mutants have enhanced our understanding of plastid-to-nucleus signaling. In gun mutants PhANGs are partially derepressed when chloroplast biogenesis is blocked. Koussevitzky et al.

(2007a) reported that GUN1, encodes a chloroplastic pentatricopeptide repeat (PPR) protein, which acts downstream of plastid-derived signals such as Mg-protoporphyrin IX, high light signals, sugar signals, plastid import signals, and signals related to the expression of the plastid genome (Koussevitzky et al., 2007a, Kakizaki et al., 2009)

(Figure 1.9). All of the signals that are thought to be integrated by GUN1 induces a severe reduction in thylakoid development or induce thylakoid dysfunction. The only exception is Mg-Proto IX, which appears to be a specific for the transmission of the signal that couples defective chloroplast biogenesis caused by the loss of carotenoid synthesis with nuclear gene expression, because GUN5 mutants do not have a gun

phenotype when treated with lincomycin (Figure 1.9) (Grey et al. 2003). Although the

GUN1 pathway has been reported to be epistatic to GUN5, and thus they define the same signaling pathway (Koussevitzky et al., 2007a), the data is inconsistent.

Mochizuki et al. 2001 reported a synergistic derepression of Lhcb in the gun1-1 gun5 double mutant, which resulted in 4 to 5 fold more Lhcb transcript accumulation than in the gun1-1 single mutant. This result was measured by two independent methods and has been replicated by other groups (Ruckle et al., unpublished). The 4 to 5 fold increase in Lhcb transcript accumulation seen in the gun1-1 gun5 double mutant as compared to the gun1-1 single mutant is much more than the 50 to 100 percent increase in Lhcb transcript accumulation that has been reported for null alleles of GUN1 as compared to the gun1-1 allele (Koussevitzky et al., 2007a, Ruckle et al., 2007).

Whether GUN1 and GUN5 define the same signaling pathway is not clear. GUN1 is a component of a major plastid-to-nucleus signaling pathway, but additional pathways likely exist because gun1 null mutants treated with inhibitors of chloroplast biogenesis do not accumulate PhANG transcripts at the same levels as untreated wild-type seedlings (Koussevitzky et al., 2007a). The plastid development signal that is synthesized or transduced by GUN1 (factor W in Figure 1.9) activates the transcription factor ABI4, which binds the CACC element of the G-box, preventing transcriptional activators, like HY5 and other G-Box binding factors, from binding the G-box and activating PhANG expression (Figure 1.9) (Koussevitzky et al., 2007a).

The GUN1 and the GUN5 pathways couple the functional state of the chloroplast with nuclear gene expression, these pathways appear to be more active in proplastids and during chloroplast development. But the GUN1 pathway is active in developed

chloroplasts that are under high light stress (Koussevitzky et al., 2007a). In developed chloroplasts there are signaling pathways that couple the redox status of the plastoquinone (PQ) pool, and the production of ROS by photosynthesis with nuclear gene expression (Figure 1.9) (Pfannschmidt et al., 2008). Although redox signals can be triggered by high light, high light is non-specific and more specific experiments can be carried out with the use of inhibitors of photosynthetic electon transport like DBMIB or DCMU. Although plastid signals mediated by the developmental state of the chloroplast regulate several thousand genes (Strand et al., 2003 and Koussevitzky et al.,

2007a), only 54 genes were strictly regulated by the PQ redox status (Fey et al., 2005).

Although the mechanism by which redox signals are perceived and transmitted to the nucleus is not well understood, two different pathways appear to be involved. The first is a putative phosphorylation cascade that involves the action of the thylakoid membrane kinase STN7 and its paralog STN8 (Figure 1.9) (Bonardi et al., 2005,

Pfannschmidt et al., 2009). The second is through the modulation of thioredoxins (TRX), by ferredoxin. Thioredoxins modify the activity of many chloroplast enzymes and processes depending on their redox status, although it is not clear if these modification leave the chloroplast (Pogson et al., 2008). There are many conditions that stimulate the production of singlet oxygen and ROS in the chloroplast. Singlet oxygen can be derived from PSII when the PQ pool is over reduced or from photosensitizing chlorophyll precursors (Pogson et al., 2008). If singlet oxygen has been shown to induce programmed cell death, this programmed cell death response is mediated at least in part by EXECUTER1 and EXECUTER2 (Figure 1.9) (Wagner et al., 2004, Lee et al., 2007). Additionally, plastid-derived superoxide and hydrogen peroxide

specifically regulate gene expression in the nucleus, but the mechanism by which ROS signals are perceived and transuded are not well understood, but the transduction might involve the activation of a MAPK cascade (Figure 1.9) (Pfannschmidt et al., 2008).

INTEGRATION OF LIGHT AND PLASTID SIGNALS

Light and plastid signals are the major regulators of chloroplast development and the expression of photosynthetic genes in the nucleus. But the mechanism and the extent to which these signaling networks are integrated remains relatively unstudied.

Additionally, the significance of this integration is only hypothetical, but it does make sense that these two signals are integrated to in order to couple the light environment with the functional state of the chloroplast. Prior to the work presented here, plastid signals were thought to regulate PhANG expression independently of light signals

(Koussevitzky et al., 2007a).

Strategy to identify common components of the plastid and light signaling networks

The hypothesis tested here, is that plastid signals and light signals are not completely independent and that PhANG expression would be more effectively coordinated if these signaling networks are integrated through common trans acting factors (Factor X in

Figure 1.10A). To identify these common components we took a forward genetic approach. First we identified plastid signaling mutants (gun mutants) that were unable

+ to couple the expression the Lhcb::Luc reporter gene with the functional state of the chloroplast when grown on norflurazon (Figure 1.10B). Because of the nature of this mutant screen, which was based on the sensitive nature of the luciferase-based reporter gene, we were able to identify mutants with relatively subtle phenotypes.

These new gun mutants were then grown under various light qualities to identify

mutants that also had defects in light signaling or perception (Figure 1.10B). Because of the subtle gun phenotype of these mutants the light signaling phenotype was much more advantageous for the positional cloning of the mutant alleles. From this screen we identified multiple mutations in CRY1. The initial characterization of the integration of light and plastid signals mediated by CRY1 and other photoreceptor mutants is presented in Chapter 2. The role that plastid signals mediated by cry1 and GUN1 pathway plays in development is presented in Chapter 3.

A second approach was taken to address two questions about the integration of plastid and light signals. (1) To what extent do plastid and light signals coordinate the expression of the nuclear and plastid genomes? (2) Can we identify factors that are coordinately regulated by plastid and light signals and that potentially mediate the signal transduction of these signaling pathways? We hypothesize that genes whose expression is induced early in the response to light and plastid signals are important for the subsequent regulation of PhANGs by the light and plastid signals (Figure 1.10C-D).

Mutants in these early-induced factors are expected to provide useful tools to test ideas on the significance of the integration of plastid and light signals in nature (Figure 1.10D).

The analysis of the light and plastid-regulated transcriptomes revealed that these signals are important for regulation of a wide variety of plant functions, but these interactions appear the most important for regulating photosynthesis and chloroplast function. Characterization of the mutants in genes that are coordinately regulated by light and plastid signals, indicate that the integration of light and plastid signals is important for the repression of chloroplast biogenesis. These results are presented in

Chapter 4.

Figure 1.1 The affect of light quality and quantity on plant growth and development. A) Representation of the affect of increasing light intensity on seedling development B) The affect of light intensity on leaf cell structure. Adapted from Weston et al 2000. C) The optimization of photon capture by adjusting leaf architecture to allow more unshaded light to leaves lower in the canopy. Adapted from Long et al 2006. D) Illustration of how plants can adjust leaf position perpendicular to the direction of the light source. E) Illustration of chloroplast movement in high and low intensity light. F) Representative seedling growth in continuous-red light. G) Representative seedling growth in continuous-far-red light. H) Representative seedling growth in continuous-blue light. I) Representative seedling growth in white light with a high far-red to red light ratio (shade grown). J) Representative seedling growth in white light with a high red to far-red light ratio.

For interpretation of the references to color in and all other figures, the reader is referred to the electronic version of this dissertation.

Figure 1.2 Plant photoreceptors A) The action spectrum of photosynthetic output for a unit of incident light in Pisum sativum. Adapted from Inada 1976 Plant and Cell Physiol. B) The affect of fluence quality on the inhibition of hypocotyl length in white mustard (Sinapis alba). Adapted from Beggs et al. 1980. C) The affect of fluence quality on the curvature toward light of oat coleoptiles. Adapted from Thimann and Curry 1960) D-F) Domain structure with chromophore binding sites of phytochrome, cryptochrome, and phototropin Jiao et al 2007. Absorption spectrum of purified phytochrome in either the Pr and Pfr conformation was adapted from Vierstra and Quail 1983. Absorbtion spectra of purified chryptochrome and phototropin were adapted from Ahmad et al. 2002 Plant Physiology.

Figure 1.3 A simplified model of the known components of light signaling during photomorphogenesis. A) In dark grown, etiolated seedlings the transcriptonal repressor PIF3 binds the promoter of light responsive genes that encode proteins required for chloroplast biogenesis and photomorphogenesis at the G-Box. The trasciptional activator HY5 is phosphorylated, which attenuates its activity, and it is targeted for degradation by the CSN by the COP1/SPA1 complex. The DET1/DDB1/COP10 complex further activates COP1 activity and localizes it to transcriptionally active sites. B) When exposed to light PHYB (and other phytochromes) translocate to the nucleus, where it forms a complex with HMR. The formation of the PHYB/HMR complex then leads to the degradation of PIF1 and PIF3. Concurrently, blue light activation of CRY1 leads to an inactivation of COP1 and the relocalization of COP1 to the cytoplasm. HY5 becomes unphosphroylated and accumulates because COP1 is inactive. HY5 then binds to the G-Box and recruits the PIC (pre-initiation complex), and chloroplast biogenesis and photomorphogenesis genes are transcribed. For abbreviations see the text.

Figure 1.4 The known light signaling network. Four know classes of photoreceptors initiate the signal transduction cascade. Each factor is color coded to represent its relative importance to the respective photoreceptor signaling cascade. COP/DET/FUS and the transciption factors known to be regulated by the COP/DET/FUS signalosome are coded in yellow. The flow of information and the nature of regulation between signaling components is indicated with arrows and t-bars. The relative thickness of the line represents an approximate estimation of the contribution of a given factor in the regulation of a downstream process, based on the relative phenotype of the mutant in regulating photomorphogenic growth.

Figure 1.5 Agriculturally important plastid types. A) The transition from etioplast to chloroplast in dark grown seedlings. B) The transition from proplastids to chloroplast only occurs in developing embryos and at the leaf primordia. C) Representation of a mint leucoplast in a glandular trichrome. Adapted from Turner et al., 1999. D) Representation of an amyloplast in the root tip as a gravity sensor and in a potato storage tuber. Adapted from Wise et al., 2007 E) Representation of an elaioplast in mature pollen grain. Adapted from Wise et al., 2007 F) A chromoplast developing in a ripening tomato fruit G) A gerontoplast disassembling from a chloroplast during senescence. H) A nodule proplastid in a nitrogen fixing root nodule.

Figure 1.6 Examples the chloroplast adapting to the light. A) Photosynthetic carbon fixation rate as a function of light intensity in two different plants. The first plant was grown in full sun conditions before being measured at various fluence rates (Red), the second plant was grown in the shade before being measured at various fluence rates (Blue). Adapted from Harvey et al 1979. B) The potential regulation of low light adapted chloroplast development by the GLK transcription factors. In low light GLKs are activated by the light and upregulate chlorophyll synthesis and Lhcb, which leads to more stacked grana, and more thylakoids. C) In high light, the GLKs are repressed by plastid derived signals and there is less chlorophyll synthesis and Lhcb, resulting in less stacked grana and fewer thylakoids. D) Photosynthetic carbon fixation rate as a function of light intensity in two different plant species. Panicum prionitis, a C4 plant (Red) and Panicum laxum a C3 plant (Blue). Adapted from Henning et al., 1986. E) One of the characteristic traits of C4 metabolisim is dimorphic chloroplasts. In mesophyll cells there are stacked grana, but in bundle sheath cells there is only unstacked thylakoids as well as the presence of the peripherial reticulum (PR).

Figure 1.7 Examples of mutations in chloroplast development.

Figure 1.8 The balance of light and plastid signals during chloroplast development. Red lines, transcripts ,and proteins indicate the flow of information from the nucleus to the plastid and from the plastid to the nucleus to maintain the proplastid development state. In part through the initiation by light perceived by the photoreceptors (PR) PhANGs are expressed and transported to the plastid. Blue lines, transcripts and proteins indicate the flow of information that is required to develop an maintain chloroplast function. If a developed chloroplast becomes stressed redox, ROS, and developmental signals activate the expression of ROS scavenging enzymes, and sunscreens, while simultaneously repressing PhANGs. The orange lines, transcripts and proteins indicate the flow of information that is required to adapt to stress. Once the adaptation is complete the chloroplast returns to homoeostasis.

Figure 1.9 Plastid signaling during chloroplast development. When Arabidopsis seedlings are treated with inhibitors of chloroplast development

(LIN and NFL), sugars, or high light, thylakoid biogenesis is blocked or thylakoid dysfunction is induced. Ether the general dysfunction of the thylakoids or the individual treatments trigger the production of a second messenger W that requires GUN1 for its synthesis or transduction. W induces ABI4 dependent repression of PhANG expression by binding to the CCAC consensus sequence of the G-box, and WKXVGLVSODFLQJ*%)¶V,QIXQFWLRQDOFKORURSODVWVWKHSKRWRV\QWKHWLFHOHFWURQ transport chain generates singlet oxygen, superoxide, and hydrogen peroxide. EX1 and EX2 are required for the perception or the transduction of singlet oxygen signal in the chloroplast then transduced to the nucleus by X. Superoxide and hydrogen peroxide signals transduced by Z leads to the expression of ROS scavenging enzymes. Redox signals that couple the redox state of the PQ pool with nuclear gene expression is perceived or transduced in part by STN7 and STN8 in the chloroplast , but requires Y to affect nuclear gene expression.

Figure 1.10 Strategy for obtaining mutants that regulate chloroplast development. A) Although it has been established that plastid and light signals regulate PhANG expression by distinct signaling pathways (factors W and Y) through the same promoter elements (Koussevitzki et al., 2007) we propose that this regulation may occur through the integration common trans acting factor(s) X. B) Schematic of the forward genetic screen designed to identify common factors to light and plastid signaling networks. C) The hypothesis that the coordinated regulation by the light and plastid signals of factor(s) Z are at least in part responsible for the downstream coordinated regulation by the light and plastid signals. D) Schematic of the reverse genetic screen designed to identify common factors to light and plastid signaling networks. Seedlings grown in the presence or absence of lincomycin are shifted from low to high fluence rate red and blue light. Tissue is collected at early and late time points and the changes in transcription are analyzed with microarrays. Candidate genes are selected and mutants are isolated and characterized.

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CHAPTER 2

PLASTID SIGNALS REMODEL LIGHT SIGNALING NETWORKS AND ARE ESSENTIAL FOR EFFICIENT CHLOROPLAST BIOGENESIS IN ARABIDOSIS

This research was originally published in The Plant Cell. Michael E. Ruckle, Stephanie M. DeMarco, and Robert M. Larkin. Plastid signals remodel light signaling networks and are essential for efficient chloroplast biogenesis in Arabidopsis. The Plant Cell. 2007; Vol. 19: pp. 3944-60 © 2007 The American Society of Plant Biologist.

PLASTID SIGNALS REMODEL LIGHT SIGNALING NETWORKS AND ARE ESSENTIAL FOR EFFICIENT CHLOROPLAST BIOGENESIS IN ARABIDOSIS

ABSTRACT

Plastid signals are among the most potent regulators of genes that encode proteins active in photosynthesis. Plastid signals help coordinate the expression of the nuclear and chloroplast genomes and the expression of genes with the functional state of the chloroplast. Here, we report the isolation of new cryptochrome 1 (cry1) alleles from a screen for Arabidopsis genomes uncoupled (gun) mutants, which have defects in plastid-to-nucleus signaling. We also report genetic experiments showing that previously unidentified plastid signal converts multiple light signaling pathways that perceive distinct qualities of light from positive to negative regulators of some but not all photosynthesis-associated nuclear genes (PhANGs) and change the fluence rate response of PhANGs. At least part of this remodeling of light signaling networks involves converting HY5, a positive regulator of PhANGs, into a negative regulator of

PhANGs. We also observed that mutants with defects in both plastid-to-nucleus and cry1 signaling exhibited severe chlorophyll deficiencies. These data show that the remodeling of light signaling networks by plastid signals is a mechanism that plants use to integrate signals describing the functional and developmental state of plastids with signals describing particular light environments when regulating PhANG expression and performing chloroplast biogenesis.

INTRODUCTION

Proteins that perform functions related to photosynthesis are encoded by both nuclear and chloroplast genomes. The chloroplast contains a small genome that encodes less than 100 proteins; some 2100 nuclear genes are predicted to encode chloroplast proteins in Arabidopsis thaliana and as many as 4800 in Oryza sativa

(Richly and Leister, 2004). Thus, the majority of proteins active in photosynthesis are encoded by nuclear genes. Coordinating PhANG expression with the expression of photosynthesis-associated plastidic genes is central to the establishment and maintenance of the photoautotrophic lifestyle of plants. The regulation of PhANGs has been studied for decades, but a number of significant gaps remain in our knowledge of their regulation (Nott et al., 2006; Rook et al., 2006; Jiao et al., 2007).

Regulation of PhANGs is complex, involving signals perceived by multiple signaling pathways, such as those triggered by light, the circadian clock, tissue-specific signals, carbohydrates, hormones, and plastids (Nott et al., 2006; Rook et al., 2006;

Jiao et al., 2007). Plastid signals affect photosystem stoichiometry, stress responses, and are thought to regulate PhANGs as proplastids develop into chloroplasts, a process that is coordinated with the development of leaf cells from the leaf primordia and with the transition of cotyledons from heterotrophic to photoautotrophic organs after germination (Mullet, 1988, 1993; Nott et al., 2006). To study plastid signals, laboratories often use inhibitors or mutations to block chloroplast development or perturb chloroplast function. PhANG expression is most potently repressed by plastid signals when chloroplast biogenesis is blocked (Nott et al., 2006). These repressive plastid signals are stronger than inductive signals, such as the robust inductive signals

from extraplastidic photoreceptors, and can repress PhANG expression to lower levels than are observed in the dark (Sullivan and Gray, 1999). The plastid signals emitted when chloroplast biogenesis is blocked are thought to contribute to proper gene expression during the early stages of chloroplast biogenesis and to help coordinate the expression of genes that encode functions related to photosynthesis that reside in both nuclear and chloroplast genomes. Proper coordination of nuclear and chloroplast genome expression is thought to be critical for proper chloroplast biogenesis because much of the photosynthetic machinery is composed of large multisubunit protein complexes composed of both plastid and nuclear gene products (Nott et al., 2006).

Retrograde signaling pathways analogous to these plastid-to-nucleus signaling pathways have been reported for the mitochondria and endoplasmic reticulum. These other retrograde signaling pathways, which are understood in much more detail, inform the nucleus on the status of their respective organelles, causing an adjustment in the anterograde flow of information from the nucleus (Liu and Butow, 2006; Ron and Walter,

2007). During chloroplast biogenesis, in an analogous fashion, retrograde plastid-to- nucleus signaling pathways are thought to regulate the anterograde flow of information from the nucleus to the plastid, which is driven by extraplastidic signaling pathways that sense endogenous and environmental cues.

Although the molecular nature of plastid-to-nucleus signaling pathways active during chloroplast biogenesis remains poorly understood, our understanding of the regulation of PhANG expression by plastid-to-nucleus signaling pathways has been enhanced by the genomes uncoupled (gun) mutants. gun mutants uncouple the expression of genes that encode proteins active in photosynthesis with chloroplast

function. When chloroplast biogenesis is blocked in wild-type seedlings, genes that encode proteins active in photosynthesis are repressed, but when chloroplast biogenesis is blocked in gun mutants, PhANGs are partially derepressed. The derepression of PhANG expression in gun mutants is thought to be due to at least partial inactivation of a plastid-to-nucleus signaling pathway that represses nuclear gene expression in response to particular plastid signals (Nott et al., 2006; Koussevitzky et al.,

2007). The expression of the nuclear and chloroplastic genomes has also been reported to be uncoupled in gun1 mutants when chloroplast biogenesis is blocked

(Susek et al., 1993).

Previous work with gun mutants indicates that accumulation of Mg- protoporphyrin IX, inhibiting the expression of the chloroplast genome, high levels of glucose, and exposure to high-intensity light all produce a second messenger that triggers a plastid-to-nucleus signaling pathway that represses PhANG expression.

GUN1, a chloroplastic pentatricopeptide repeat (PPR) protein is required for the biosynthesis or the transduction of this second messenger. ABI4, an Apetala 2-type transcription factor, functions downstream of GUN1 by binding promoter elements found in PhANGs (Nott et al., 2006; Koussevitzky et al., 2007). The discoveries that GUN1 and ABI4 act downstream of multiple plastid signals is consistent with a master switch integrating diverse plastid signals proposed by Richly et al. (2003). Although these recent advances are very exciting, significant gaps in our understanding of this form of interorganellar communication remain. For example, the identity of plastid signals, the mechanism by which signals exit the plastid, the mechanism by which the plastid interacts with other cellular compartments, and the impact of plastid signals on growth

and development remain open questions. Additionally, because strong gun1 alleles express lower levels of PhANGs, when treated with inhibitors of chloroplast biogenesis compared to wild-type seedlings that are not treated with inhibitors of chloroplast biogenesis (Koussevitzky et al., 2007), it is likely that one or more additional plastid-to- nucleus signaling pathways that do not utilize GUN1 help coordinate PhANG expression with chloroplast biogenesis and function.

Much more is known about the regulation of PhANG expression by light than by plastid signals. In Arabidopsis and rice, at least 20% of the transcriptome is regulated by light. A number of photoreceptors and downstream signaling components have been shown to function in light-regulated transcriptional networks, and multiple light- responsive promoter elements have been identified in PhANGs (Jiao et al., 2007).

Well-studied PhANGs, such as the genes that encode the light-harvesting chlorophyll a/b-binding protein of photosystem II (Lhcb or CAB, hereafter referred to as Lhcb) and the Rubisco small subunit (Rbcs), are light induced via the phytochrome and cryptochrome signaling pathways (Gao and Kaufman, 1994; Reed et al., 1994; Folta and Kaufman, 1999; Mazzella et al., 2001; Martinez-Hernandez et al., 2002). These pathways transduce far-red, red, and blue light signals (Jiao et al., 2007). Regulation of transcription by light is complex, involving the regulation of activity, subcellular localization, and concentrations of particular photoreceptors and downstream signaling components (Jiao et al., 2007).

Although light and plastid signals trigger distinct signaling pathways (Sullivan and

Gray, 1999), it is known that plastid signals and light signals can regulate PhANG expression using common or adjacent promoter elements (Nott et al., 2006;

Koussevitzky et al., 2007). To test whether plastid-to-nucleus signaling pathways and light signaling pathways might interact at some point upstream of these common promoter elements, we isolated a group of new gun mutants and screened them for light signaling phenotypes. We discovered that four of these mutants were allelic to cry1 and that cry1 alleles isolated by other laboratories were also gun mutants. We found that along with plastid signals and cry1, phyB and likely another phytochrome can also contribute to the repression of Lhcb when chloroplast biogenesis is blocked. Moreover, we found that the mechanism by which cry1 represses Lhcb expression when chloroplast biogenesis is blocked involves the conversion of HY5, a well-studied bZIP transcription factor that acts downstream of cry1 and other photoreceptors (Jiao et al.,

2007), from a positive regulator to a negative regulator of Lhcb and Rbcs. This remodeling of light signaling pathways was independent of the plastid-to-nucleus signaling pathway defined by previously isolated gun mutants and affected the response of PhANGs to both light quality and quantity. We also observed that gun1-101, a null allele, exhibited chlorophyll deficiencies that increased strikingly in either cry1 or hy5 null mutant backgrounds, which indicates that GUN1 and cry1 signaling pathways contribute to efficient chloroplast biogenesis in a redundant manner.

RESULTS

Isolation of cry1 mutants from a gun mutant screen

We performed a screen to isolate new gun mutants that might contain defects in one or more of the steps in plastid-to-nucleus signaling that are poorly understood

(Figure 2.11). In this screen, following EMS mutagenesis, new gun mutants were identified as M2 seedlings that exhibited derepression of an Lhcb1*1:luciferase + (luc+) reporter gene, as judged by bioluminescence of seedlings grown on media containing norflurazon. Norflurazon blocks carotenoid biosynthesis by inhibiting phytoene desaturase. Without carotenoids, plastids experience severe photooxidative stress in bright light, chloroplast biogenesis is arrested at an early stage that resembles the proplastid, and PhANGs are severely repressed (Oelmüller, 1989). We expected that the Lhcb:luc+ reporter gene would provide a more sensitive screen than the original gun screen, which utilized an Lhcb-driven reporter gene that conferred hygromycin resistance upon gun mutants (Susek et al., 1993). With greater sensitivity, we expected to identify mutants with subtle phenotypes that were missed in the original screen.

Progeny that inherited the Lhcb:luc+ reporter gene-based gun phenotype were grown on media that contained norflurazon and screened for derepression of the endogenous

Lhcb genes. Mutants that exhibited derepression of endogenous Lhcb genes when grown on media containing norflurazon were then grown on media containing lincomycin and tested for derepression of endogenous Lhcb genes. Like norflurazon, lincomycin blocks chloroplast biogenesis and causes severe repression of PhANGs.

Lincomycin inhibits chloroplast biogenesis by specifically inhibiting plastid translation, which is an entirely different mechanism than norflurazon (Mulo et al., 2003). Thus, the

mutants obtained at the end of this process cannot be resistant to a particular inhibitor and likely have defects in signaling pathways that regulate endogenous Lhcb genes.

Because light and plastid-to-nucleus signaling pathways regulate Lhcb and a number of other PhANGs through common promoter elements, we tested whether some of our new gun mutants might also have light signaling defects. We found that four of our mutants exhibited long hypocotyls when grown in high-fluence-rate blue light

(Figure 2.1A), which is consistent with these mutants having defects in cry1 signaling

(Ahmad and Cashmore, 1993; Ahmad et al., 1995; Shalitin et al., 2003; Ohgishi et al.,

2004). The gun phenotypes of these mutants were subtle compared to mutants isolated from the first gun screen such as gun1-1 (Susek et al., 1993; Mochizuki et al., 2001). In fact, Lhcb mRNA accumulated to only two- to threefold above wild-type levels in most of these new gun mutants. In contrast, we repeatedly observed that gun1-1 accumulated approximately eight- to ten-fold more Lhcb mRNA than wild type when chloroplast biogenesis was blocked (Figure 2.1B). Like other gun mutants, Lhcb mRNA accumulated to similar levels as in the wild type when these mutants were not treated with inhibitors of chloroplast biogenesis (Figure 2.1B).

To test whether the gun phenotypes and the long-hypocotyl phenotypes might be linked, one of these new mutants was crossed to the parental line, Columbia-0 (Col-0) containing the Lhcb::luc+ reporter gene. In the F2 progeny, the long-hypocotyl phenotype segregated like a semi-dominant allele when seedlings were grown in high- fluence-rate blue light (S.M. DeMarco, M.E. Ruckle, and R.M. Larkin, unpublished data), as has previously been reported for cry1 mutants (Koornneef et al., 1980; Ahmad and

Cashmore, 1993). Four F2 seedlings that exhibited long hypocotyls in blue light were propagated and F3 progeny were found to be homozygous for the long hypocotyl in blue light and gun phenotypes (Table 2.1 and Figure 2.19), which indicates that these two phenotypes are linked. Because the wild-type hypocotyl phenotype could be scored unambiguously, we were able to determine that this phenotype mapped to a 1.5-

Mb interval on chromosome 4 that contained CRY1 (Figure 2.1C). From these data, we hypothesized that this new mutant and possibly all four of our new gun mutants that exhibited long hypocotyls in blue light might be allelic to cry1. To test this idea, we sequenced CRY1 in all four mutants. We found G to A transitions in each mutant that caused substitutions in the derived amino acid sequence (Figure 2.1D). To further test the possibility that cry1 mutants were also gun mutants, we obtained a cry1 allele in which a T-DNA is inserted into the third exon of CRY1 (Salk_069292; Alonso et al.,

2003). Because the last published set of cry1 alleles was numbered in the three hundreds (Shalitin et al., 2003), we refer to this T-DNA allele as cry1-400. We determined that cry1-400 does not accumulate CRY1 mRNA (Figure 2.12C) and therefore must be a null allele. We observed that cry1-400 is a gun mutant and exhibits a similar gun phenotype compared to the aforementioned missense alleles. hy4-1, a cry1 mutant in the Landsberg erecta (Ler) ecotype (Ahmad and Cashmore, 1993), accumulated similar amounts of Lhcb mRNA compared to the other mutants, but because Lhcb is more severely repressed in Ler compared to Col-0, the gun phenotype of hy4-1 is actually more robust than a typical cry1 mutant in the Col-0 ecotype (Figure

2.1B). From these data we concluded that cry1 contributes to the repression of Lhcb genes when chloroplast biogenesis is blocked and that our four new gun mutants are

allelic to cry1. We named these new missense alleles cry1-401, cry1-402, cry1-403, and cry1-404. We have not yet determined whether the twofold stronger gun phenotype observed in cry1-401 is caused by an unlinked mutation or by the amino acid substitution caused by the cry1-401 allele.

Double mutant studies with cry1 and gun1 mutants

To learn more about the mechanism of PhANG regulation by GUN1 and cry1 when chloroplast biogenesis is blocked, we analyzed gun1 cry1 double mutants.

Double mutants were constructed using cry1-400 and either gun1-1, a leaky missense allele (Susek et al., 1993; Koussevitzky et al., 2007), or a publicly available gun1 T-DNA allele (SAIL_33_D01; Sessions et al., 2002) that we refer to as gun1-101. gun1-101 expresses a partial GUN1 mRNA that encodes a truncated PPR domain but not the small mutS-related (SMR) domain that is thought to be required for DNA binding activity (Koussevitzky et al., 2007) ( Figure 2.12A and 2.12B online). According to the current model of GUN1 activity (Koussevitzky et al., 2007), gun1-101 should at least be a severe loss-of-function allele and possibly a null allele. Because cry1-400 was used in all subsequent experiments, for simplicity, we hereafter refer to cry1-400 as cry1. gun1-101 accumulated more Lhcb mRNA than gun1-1 when chloroplast biogenesis was blocked. This difference was always greater in blue than white light. In nine independent experiments with different inhibitors of chloroplast biogenesis and in either blue or white light, we observed 1.5- to 2.7-fold more Lhcb mRNA when chloroplast biogenesis was blocked in cry1 gun1-1 or cry1 gun1-101 double mutants than would be expected for additive increases caused by two pathways acting independently (Figures

2.2, 2.3, and 2.4). These data indicate that GUN1 and cry1 are partially redundant in their repression of Lhcb gene expression when chloroplast biogenesis is blocked in these light conditions.

In blue light, Lhcb mRNA accumulated to the same level in wild-type seedlings that were not treated with inhibitors of chloroplast biogenesis (i.e., green seedlings) and lincomycin-treated gun1-101 cry1 double mutants (Figures 2.2 and 2.4). These data suggest that when chloroplast biogenesis is blocked in blue light, most, if not all, of the repression of Lhcb is mediated by both GUN1 and cry1 and that gun1-101 is likely a null.

In white light, the gun1-101 cry1 double mutant accumulated 70% of Lhcb mRNA found in untreated controls, which suggests that perception of at least one additional light quality besides blue light or that higher fluence rates might be important for maximal repression of Lhcb under these conditions.

We observed that although Lhcb is derepressed in cry1 when chloroplast biogenesis is blocked in either blue light or white light, Lhcb mRNA accumulated to similar levels in cry1 and wild type when chloroplast biogenesis was blocked in red light

(i.e., cry1 is not a gun mutant in red light) (Figure 2.13). We conclude that when chloroplast biogenesis is blocked, maximum repression of Lhcb is dependent on photoactivated cry1. The long hypocotyl in blue light phenotypes of all of the cry1 alleles described in this report (Figure 2.1A) further supports our conclusion that photoactivated cry1 represses Lhcb when chloroplast biogenesis is blocked.

Because redundancies have been observed between cry1 and cry2 in the regulation of some blue-light-responsive processes (Casal, 2006), we tested whether a

cry2 mutant might also be a gun mutant. We observed that cry2-1, a null allele (Guo et al., 1998), is not a gun mutant. However, the cry1 cry2-1 double knockout repeatedly exhibited a slightly stronger gun phenotype than cry1, but only in blue light (Figure 2.2).

These results indicate that cry2 can partially compensate for cry1 in the cry1 background under these conditions. We also tested whether other blue-light photoreceptor T-DNA insertion mutants (Figure 2.12) might also be gun mutants. We found that phot1, phot2, phyA, phyB, nph3, and cry3 T-DNA insertion mutants are not gun mutants in blue light (Figure 2.14).

Like Lhcb, Rbcs is repressed in wild type and derepressed in gun1 mutants when chloroplast biogenesis is blocked (Figure 2.2; Susek et al., 1993). In contrast to Lhcb, however, Rbcs is not derepressed in cry1 or gun1 cry1 double mutants when chloroplast biogenesis is blocked. In fact, Rbcs mRNA usually accumulated to slightly lower levels in cry1 than in wild type and always accumulated to lower levels in gun1 cry1 double mutants than in gun1 mutants (Figures 2.2, 2.3, and 2.4). These results indicate that cry1 induces Rbcs gene expression under these conditions. Because in gun1 mutants Rbcs mRNA accumulated to only approximately 10-20% of untreated wild-type controls (Figure 2.2), we conclude that at least one additional pathway besides the GUN1 pathway likely contributes to the repression of Rbcs under these conditions.

The more severe repression of Rbcs in white light than blue light (Figure 2.2) suggests that, as with Lhcb, perception of multiple qualities of light or higher fluence rates might be required for the maximal repression of Rbcs when chloroplast biogenesis is blocked.

We observed the same patterns of Lhcb and Rbcs expression in gun1, cry1, and gun1-1 cry1 double mutants regardless of whether chloroplast biogenesis was blocked

by treatments using lincomycin, erythromycin, or norflurazon (Figure 2.3). Both lincomycin and erythromycin specifically inhibit plastid translation, but utilize different mechanisms (Mulo et al., 2003), and, as mentioned above, norflurazon is a phytoene desaturase inhibitor. Each of these inhibitors has previously been shown to arrest chloroplast biogenesis at a stage resembling a proplastid and to severely repress

PhANGs in Arabidopsis (Nott et al., 2006). Therefore, we conclude that the repression of Lhcb and Rbcs expression are likely caused by a reduction in the activities of particular signaling pathways and cannot be caused by resistance to particular inhibitors.

Moreover, when seedlings were not treated with inhibitors of chloroplast biogenesis (i.e., in green seedlings), Lhcb mRNA accumulated to similar levels in gun1-1, cry1, gun1-1 cry1, and wild type (Figure 2.3). When seedlings were not treated with inhibitors of chloroplast biogenesis, Rbcs mRNA accumulated to essentially the same levels in gun1-1 and wild type, but Rbcs mRNA accumulated to lower levels in cry1 backgrounds regardless of whether chloroplast biogenesis was blocked (Figures 2.2, 2.3, and 2.4).

These data indicate that cry1 does not play an essential role in regulating Lhcb in green seedlings under these light conditions and that cry1 induces Rbcs regardless of the developmental state of the plastid.

Genetic analyses of downstream signaling components

cry1 has been shown to promote photomorphogenesis by inhibiting COP1, an E3 ubiquitin ligase that targets positive regulators of photomorphogenesis for degradation via the proteasome (Yi and Deng, 2005; Jiao et al., 2007). We tested whether cry1

utilized a COP1-dependent mechanism to repress Lhcb and simultaneously induce

Rbcs expression when chloroplast biogenesis was blocked and whether GUN1 utilizes a COP1-dependent mechanism to repress PhANGs. When we analyzed Lhcb expression, we observed that cop1-4, a weak allele (Deng and Quail, 1992; McNellis et al., 1994), was not a gun mutant and that cop1-4 was epistatic to cry1 but had a minor impact on gun1-1 when chloroplast biogenesis was blocked (Figure 2.4A). These data indicate that cry1 functions through a COP1-dependent mechanism to repress Lhcb and suggest that GUN1 does not likely utilize a COP1-dependent mechanism to repress

Lhcb under these conditions.

A different pattern of regulation was observed when we monitored Rbcs expression in cop1-4 single and double mutants. Because cry1 induces Rbcs when chloroplast biogenesis is blocked, we expected that lincomycin-treated cop1-4 would express higher levels of Rbcs than wild type. cop1-4 and cry1 cop1-4 repeatedly accumulated slightly higher levels of Rbcs than wild type under these conditions.

Moreover, we did observe an enhanced derepression of Rbcs in cop1-4 gun1-1 compared to the cop1-4 and gun1-1 single mutants (Figure 2.4A). These data indicate that cop1-4 is epistatic to cry1 but not gun1-1, suggesting that cry1 and COP1 function in the same pathway and that GUN1 and cry1 function in different pathways.

Because cry1 contributes to the repression of Lhcb when chloroplast biogenesis is blocked, we speculated that other positive regulators of PhANG expression might also inhibit Lhcb expression when chloroplast biogenesis is blocked. We chose to test whether HY5, a positive regulator of PhANG expression in vivo (Lee et al., 2007) that acts downstream of multiple photoreceptors (Jiao et al., 2007), might contribute to the

repression of PhANGs when chloroplast biogenesis is blocked. Indeed, we observed that a T-DNA insertion in HY5 (SALK_096651; Alonso et al., 2003) that we determined to be a null allele (Figure 2.12) and refer to as hy5 was a subtle gun mutant with an

Lhcb expression phenotype similar to cry1 (Figure 2.4B). We also observed that Lhcb was expressed at similar levels in the cry1 hy5 double mutant as the cry1 and hy5 single mutants. Additionally, the gun1-101 hy5 double mutant resembled the gun1 cry1 double mutants in that it exhibited enhanced Lhcb expression compared to the gun1-

101 and hy5 single mutants. These data indicate that HY5 functions in the same pathway as cry1 and is responsible for much of the cry1-mediated repression of Lhcb when chloroplast biogenesis is blocked. These data also indicate that HY5 functions in a pathway that is distinct from GUN1 that represses Lhcb when chloroplast biogenesis is blocked. When we monitored Rbcs expression in the same mutants, we observed that hy5 and wild type contained similar levels of Rbcs mRNA and that more Rbcs mRNA accumulated in the gun1-101 hy5 double mutant than in gun1-101 (Figure 2.4B).

These data indicate that HY5 does not contribute to the cry1-mediated induction of

Rbcs under these conditions. HY5 may contribute to the repression of Rbcs as it does for Lhcb, but these repressive effects can be observed only when GUN1 is not active.

Analysis of gun phenotypes in different light qualities and in phy mutants

Because we repeatedly observed that regardless of genetic background, Lhcb and Rbcs mRNAs accumulated to higher levels when chloroplast biogenesis was blocked in blue light compared to white light, we hypothesized that the crosstalk

between plastid-to-nucleus signaling pathways and light signaling pathways may be more complex than the interactions between the GUN1 and cry1 pathways described above. To test this idea, we compared PhANG expression in lincomycin-treated and untreated wild-type and gun1 seedlings in darkness, white light, or in a fluence rate of blue, red, and far-red light that was equivalent to the fluence rate of each of these light

-2 -1 qualities in our 125 Pmol m s white light. In these experiments, we blocked etioplast and chloroplast biogenesis by treating seedlings with lincomycin, which prevents plastid development beyond a proplastid stage and severely represses PhANG expression in either dark- or light-grown seedlings (Sullivan and Gray, 1999). We observed that in lincomycin-treated wild type, gun1-1, and gun1-101, Lhcb mRNA accumulated to the lowest levels in darkness, but accumulated to only slightly higher levels in white light within a genetic background (Figure 2.5A). In contrast, Lhcb was expressed at approximately threefold or higher levels in blue, red, or far-red light than in white light and darkness within a genetic background (Figure 2.5A).

These data indicate that perception of multiple qualities of light might be necessary for maximum repression of Lhcb or that maximum repression of Lhcb might require high fluence rates of light when chloroplast biogenesis is blocked. Moreover, these data show that different light qualities do not have a major impact on GUN1 activity, and that light stimulates Lhcb expression compared to darkness under these conditions. When Lhcb expression levels were analyzed as a percent of untreated wild-type seedlings in the same light conditions, we observed that Lhcb mRNA accumulated to lower levels in treated gun1 mutants compared to untreated wild type in all light conditions except far-red. In far-red light, Lhcb mRNA accumulated to similar

levels in treated gun1 mutants and untreated wild type (Figure 2.5A). These data indicate that in addition to GUN1, at least one other pathway is required to repress Lhcb expression in the dark, blue light, and red light when etioplast or chloroplast biogenesis is blocked and indicate that only the GUN1 pathway is necessary to repress Lhcb in far- red light.

To test whether the simultaneous perception of blue and red light might be sufficient to produce the strong repression of Lhcb observed in white light, we compared

Lhcb mRNA levels in lincomycin-treated and untreated wild type and gun1 seedlings irradiated with white light to seedlings irradiated with a combination of blue and red light that were equivalent to the fluence rates of each of these light qualities in our 125 Pmol

-2 -1 m s white light. Although seedlings exposed to white light received a larger quantity of light than seedlings exposed to a combination of blue and red light, we observed that seedlings accumulated lower levels of Lhcb mRNA when chloroplast biogenesis is blocked in a combination of blue and red light than in white light (Figure 2.5B). From these data we conclude that most if not all of the Lhcb repression observed when chloroplast biogenesis is blocked in white light is caused by GUN1 and a combination of blue and red light, and a component of white light that is distinct from blue and red may have a slight stimulatory effect on Lhcb when chloroplast biogenesis is blocked.

We obtained different results from a similar analysis of Rbcs expression. Like

Lhcb, Rbcs was expressed at higher levels in blue light than white light when chloroplast biogenesis was blocked (Figure 2.5A), as previously observed (Figure 2.2).

In contrast to Lhcb, however, Rbcs was expressed at very low levels in red light

compared to white light. In contrast to untreated seedlings (Figure 2.15), treated seedlings accumulated similar levels of Rbcs mRNA when chloroplast biogenesis was blocked in either red light or darkness. GUN1 had a minor effect on Rbcs expression in red light compared to white and blue light. An additional difference between Lhcb and

Rbcs was that Rbcs mRNA did not accumulate to the same level as untreated controls in far-red light (Figure 2.5A). These data suggest that GUN1 and at least one other pathway are required to repress Rbcs when chloroplast biogenesis is blocked in far-red light. From these and previous results, it would appear that cryptochromes and phytochromes regulate Rbcs and Lhcb very differently when chloroplast biogenesis is blocked, which is in contrast to the similar regulation that we observed in these same light conditions when seedlings were not treated with inhibitors of chloroplast biogenesis

(Figure 2.15) and the similar regulation that was previously reported for both Lhcb and

Rbcs by cryptochromes and phytochromes when seedlings were not treated with inhibitors of chloroplast biogenesis (Mazzella et al., 2001; Martinez-Hernandez et al.,

2002).

We observed that the gun1 mutants accumulate more Lhcb and Rbcs mRNA than wild type when etioplast biogenesis is blocked in the dark and that in gun1 mutants neither Lhcb nor Rbcs mRNA accumulate to the levels observed in untreated wild-type seedlings grown in the dark (Figure 2.5A). These data are consistent with a previous report showing that lincomycin treatments can block etioplast biogenesis and that plastid-to-nucleus signaling does not depend on light (Sullivan and Gray, 1999). These data also show that GUN1 and at least one additional light-independent pathway repress these PhANGs when etioplast biogenesis is blocked. In the dark, as in all of the

different light conditions that we tested, untreated gun1 mutants and untreated wild type accumulated similar levels of Lhcb and Rbcs mRNA ( Figures 2.15 and 1.16).

Therefore, gun1 is distinct from the det/cop/fus mutants, which can express higher levels of PhANGs than wild type, regardless of whether they are treated with inhibitors of etioplast biogenesis (Sullivan and Gray, 1999; Figure 2.16).

Because our data indicate that perception of red light is required for maximal repression of PhANGs when chloroplast biogenesis is blocked, we tested whether phyA, phyB, phyA gun1-1, and phyB gun1-1 exhibit gun phenotypes. In these experiments, we used T-DNA insertion alleles of phyA and phyB (Salk_ 014575 and Salk_ 022035;

Alonso et al., 2003), which we determined to be null alleles (Figure 2.12). When Lhcb was monitored, we observed that neither phyA nor phyB were gun mutants. We observed that phyA gun1-1 accumulated less and that phyB gun1-1 accumulated more

Lhcb mRNA than gun1-1 (Figure 2.6). These data indicate that, like GUN1 and cry1, phyB can contribute to the repression of Lhcb when chloroplast biogenesis is blocked, but that phyB is only a gun mutant when GUN1 is not active. In contrast, phyA only induces Lhcb when GUN1 is inactive. Rbcs mRNA accumulated to similar levels in phyA and phyB mutants, regardless of whether these alleles were in a wild-type or gun1-1 background (Figure 2.6). These data indicate that, by themselves, neither phyA nor phyB is critical for either inducing or repressing Rbcs in wild-type seedlings when chloroplast biogenesis is blocked.

Analysis of gun phenotypes in different fluence rates of white light

In addition to light quality, fluence rate can have a major impact on PhANG expression (Terzaghi and Cashmore, 1995). Because our data indicate that plastid signals can change the nature of PhANG regulation in response to light quality, we tested whether plastid signals might also influence the response of Lhcb and Rbcs to different quantities of light. The Lhcb and Rbcs mRNA levels were measured in wild- type and gun1 seedlings that were grown in increasing fluence rates of white light and either treated or not treated with lincomycin. In untreated seedlings, we found that the fluence rate response of Lhcb and Rbcs expression differed: Lhcb expression was inhibited at the highest fluence rate, but Rbcs was stimulated only by increasing the fluence rate (Figure 2.7). In contrast, when either wild type or gun1-1 was treated with lincomycin, the peak of Lhcb expression was shifted to a lower fluence rate relative to the untreated control and Rbcs expression was inhibited by higher fluence rates, which was not observed in untreated seedlings. When seedlings were treated with lincomycin, the expression of each gene was most strongly inhibited by fluence rates above 1 Pmol

-2 -1 m s relative to the untreated control grown in the same fluence rate (Figure 2.7).

These data indicate that the inhibition of Lhcb and Rbcs expression by increasing fluence rates of white light is enhanced when chloroplast biogenesis is blocked and that

GUN1 does not affect this response to fluence rate. These data also show that under these conditions, GUN1 plays a major role in repressing Lhcb in low fluence rates but plays a less important role in higher fluence rates.

Although Lhcb genes have been reported to be strongly light-responsive

(Terzaghi and Cashmore 1995), we observed only a three to fourfold increase in Lhcb mRNA levels in white light-grown compared to dark-grown seedlings (Figure 2.7; Figure

2.15). Differences in experimental conditions likely account for these conflicting results.

For example, most reports on the light regulation of Lhcb genes analyze the transient induction of Lhcb in etiolated seedlings grown on media that lacks sucrose (Terzaghi and Cashmore, 1995). In contrast, we compared the steady-state levels of Lhcb after several days of growth in either the dark or the light, and we included sucrose in the growth media, which has been shown to inhibit Lhcb expression (Rook et al., 2006).

-2 -1 Additionally, we found that the 125 Pmol m s used in these experiments can have an inhibitory effect on Lhcb expression (Figure 2.7).

Analysis of chlorophyll deficiencies in gun1 and light signaling mutants

If plastid-to-nucleus signaling contributes to efficient chloroplast biogenesis, we would expect that chloroplast biogenesis would be less efficient in gun mutants than wild type and even more inefficient in gun1 cry1 double mutants compared to wild type.

Indeed, we observed that under growth conditions in which wild-type seedlings greened normally, a small percentage of gun1 mutants developed chlorophyll-deficient cotyledons (Figure 2.8). Consistent with our earlier experiments indicating that gun1-

101 was a stronger allele than gun1-1 (Figures 2.2, 2.4, and 2.5), chlorophyll-deficient cotyledons were observed more frequently in gun1-101 than in gun1-1 (Figure 2.8). In contrast, neither cry1 nor cry1 cry2 developed chlorophyll-deficient cotyledons under these same conditions (Figure 2.8). Consistent with the idea that GUN1 and cry1 are at least partially redundant in the regulation of genes required for proper chloroplast biogenesis, the frequency of seedlings with chlorophyll-deficient cotyledons increased

approximately threefold in the gun1-101 cry1 double mutant compared to gun1-101. A similar increase was observed when gun1-1 cry1 cry2 was compared to gun1-1 cry1, which indicates that cry2 also contributes to efficient chloroplast biogenesis (Figure 2.8).

Chlorophyll-deficient cotyledons varied from partially green organs that contained chlorophyll deficient areas to uniformly albino cotyledons in both single and higher order mutants. Regardless of genetic background, seedlings with chlorophyll-deficient cotyledons produced mostly green primary leaves but a minority produced primary leaves that only partially greened and in some cases appeared variegated (Figure 2.17).

If chloroplast biogenesis were inefficient in these mutants, we would expect them to be more susceptible than wild type to photooxidative stress and albinism when chloroplast biogenesis proceeds in continuous high-intensity white light (HL). To test these mutants for HL sensitivity during chloroplast biogenesis, we allowed these seedlings to germinate in the dark for 23 h and then transferred them to various fluence rates of continuous white light. Smaller seedlings that contained less chlorophyll were judged to be more sensitive to HL. With the exception of gun1-101 cry1, cop1-4 , and all cop1-4 double mutants, all mutants contained essentially the same level of

-2 -1 chlorophyll as wild type when seedlings were grown in 100 Pmol m s white light. At the higher fluence rates of white light, chlorophyll deficiency was uniformly distributed throughout a population of seedlings and not restricted to the cotyledons. We found that as the fluence rate of continuous white light was increased, the severity of chlorophyll deficiency increased more in the gun1, cry1, and hy5 single and double mutants compared to wild type, and that chlorophyll deficiency increased more in the double mutants made from combinations of gun1, cry1, and hy5 compared to the

-2 -1 corresponding single mutants. In 500 Pmol m s , chlorophyll levels were reduced synergistically in gun1 cry1 and gun1 hy5 double mutants compared to the corresponding single mutants. At higher fluence rates, the differences between these single and double mutants were less pronounced (Figure 2.9). We observed that gun1 mutants and cry1 mutants were similarly sensitive to HL under most fluence rates, but

-2 -1 that cry1 was more sensitive to HL than gun1 mutants at 1500 Pmol m s . hy5 appeared to be more sensitive to HL than either gun1 or cry1 mutants. Accordingly, gun1-101 hy5 was more sensitive than gun1-101 cry1. When seedling size was considered, gun1-101 hy5 appeared to be more sensitive to HL than cry1 hy5 (Figure

2.9). These data are consistent with previous studies indicating that HY5 is a downstream component of multiple signaling pathways (Jiao et al., 2007) and with our gene expression results indicating that cry1 and GUN1 trigger distinct pathways.

We found that cop1-4 contained significantly less chlorophyll than wild type at

-2 -1 100 and 500 Pmol m s . Similar results have been reported previously (Deng and

Quail, 1992). However, we observed that cop1-4, gun1-1 cop1-4, cry1 cop1-4, and wild

-2 -1 type contained similar levels of chlorophyll at 1000 and 1500 Pmol m s . These data indicate that inhibition of COP1 is a major component of HL stress protection and are consistent with cry1 using a COP1-dependent mechanism to protect plants from HL.

Because our gene expression studies indicate that GUN1 probably does not utilize a

COP1-dependent mechanism to regulate PhANG expression, we suggest that the suppression of HL sensitivity in gun1-1 cop1-4 is likely caused by indirect effects.

To test whether mature chloroplasts in these single and double mutants are simply more sensitive to HL than wild type, we grew wild type and all of these mutants

-2 -1 in 125 Pmol m s for 7 d and then transferred green seedlings to HL for 3 d. All of these mutants were green before they were transferred to continuous HL. We did not observe any consistent and striking differences in the pigmentation of mutants and wild type except that the hy5 single and double mutants were always noticeably paler than wild type at the highest fluence rate, but only in the youngest leaves (Figure 2.18).

Similar results were obtained in an experiment with three-week-old plants grown in soil

(M.E. Ruckle and R.M. Larkin, unpublished data). Altogether, our analysis of greening in these mutants indicates that the photoprotective functions provided by GUN1, cry1, and

HY5 are more important during chloroplast biogenesis than in seedlings that contain mature chloroplasts.

DISCUSSION

New cry1 alleles isolated from a gun mutant screen

The cryptochromes are composed of amino-terminal DNA photolyase-related

(PHR) domains and carboxy termini that are not related to DNA photolyases. The amino termini bind the flavin adenine dinucleotide and methenyltetrahydrofolate chromophores and are necessary for dimerization. Both of these activities are necessary for light-dependent activation of the carboxy-terminal domains (Lin and

Shalitin 2003; Sang et al., 2005). The blue light signal perceived by the PHR domain is transduced by stimulating the carboxy-terminal domains, which inhibit COP1 (Yi and

Deng, 2005; Jiao et al., 2007). A large number of missense alleles that cause amino acid substitutions throughout cry1 were isolated previously. Like the new cry1 mutants described here, all of the previously isolated missense alleles exhibit long hypocotyls in blue light and all are loss-of-function alleles (Ahmad and Cashmore, 1993; Ahmad et al.,

1995; Shalitin et al., 2003). cry1-401, cry1-402, and cry1-404 cause amino acid substitutions D21N, S286N, and G340E, respectively, and likely render cry1 defective in one or more of the activities attributed to the amino terminus. A missense allele that, like cry1-404, causes a G340E substitution in a photolyase signature sequence was isolated previously (Ahmad and Cashmore, 1993; Ahmad et al., 1995), but missense alleles that cause D21N and S286N substitutions have not been reported previously.

The S286N substitution in cry1-402 is interesting because the mechanism by which cry1 transduces the blue light signal involves phosphorylation of serine residue(s) (Bouly et al., 2003; Shalitin et al., 2003) and because missense alleles that cause substitutions at serine residues have not been reported previously. Although it is possible that the

S286N substitution in cry1-402 removes an important phosphorylation site, it is also possible that replacing a serine with an asparagine residue at position 286 is simply disruptive to folding.

cry1-403 was the only allele we isolated that caused an amino acid substitution in the carboxy terminus of cry1. Carboxy termini are poorly conserved among cryptochromes, but the carboxy termini of most cryptochromes contain three well- conserved motifs referred to as DAS. The three motifs that make up the DAS motif are

DQXVP (D), an acidic region (A), and STAES followed by GGXVP (S) (Lin and Shalitin,

2003). Other missense alleles have previously been reported to cause amino acid substitutions in and around the D and A motifs (Ahmad et al., 1995). cry1-403 is the only missense allele reported to alter the S motif, changing the highly conserved STAES motif to STAKS.

Plastid signals change the nature of Lhcb regulation by HY5

The cryptochromes have been shown to regulate gene expression in a blue light- dependent manner by binding and inhibiting COP1, an E3 ubiquitin ligase that targets photoreceptors and transcription factors that positively regulate photomorphogenesis

(e.g., HY5) for degradation in the dark via the proteasome. Consistent with this mechanism, the short-hypocotyl phenotype of cop1 has been reported to be epistatic to the long-hypocotyl phenotype of a cry1 (Ang and Deng, 1994). The results from our analysis of PhANG expression in double mutants resembles the analysis of hypocotyl length reported by Ang and Deng (1994) and are consistent (1) with cry1 utilizing a

COP1-based mechanism to regulate PhANG expression regardless of whether cry1 is functioning as a positive or a negative regulator of PhANG expression and (2) with

GUN1 not using a COP1-dependent mechanism to repress PhANGs when both etioplast and chloroplast biogenesis is blocked. From our double mutant studies we also conclude that plastid signals convert cry1 signaling pathways from positive to negative regulators of Lhcb by converting HY5 from a positive to a negative regulator of

Lhcb. Previously, HY5 has been reported to function only as a positive regulator of

PhANGs like Lhcb and Rbcs in vivo, but HY5 has been reported to negatively regulate a number of other genes (Lee et al., 2007). The mechanism by which HY5 is converted from a positive regulator to a negative regulator of Lhcb expression is an open question whose answer may include posttranslational modifications, heterodimerization with distinct transcription factors, changes in the concentrations of coactivators and corepressors, or some combination of these mechanisms.

Koussevitzky et al. (2007) showed that GUN1 prevents light signaling pathways from inducing PhANGs by promoting the binding of ABI4 adjacent to promoter elements that contain G-boxes, which are important for light induction of PhANGs (Koussevitzky et al., 2007). Our finding that HY5 only functions as a negative regulator of Rbcs expression when the GUN1 pathway was inactivated is consistent with the GUN1 pathway preventing G-box-binding factors such as HY5 (Lee et al., 2007) from regulating Rbcs expression as proposed by Koussevitzky et al. (2007). In contrast, we found that full repression of Lhcb genes requires not only GUN1 but also cry1 and HY5.

Thus, our findings indicate that Lhcb and Rbcs are repressed by distinct mechanism when chloroplast biogenesis is blocked.

Crosstalk between plastids and light signaling networks

Cryptochromes have previously been shown to regulate blue-light-inducible genes, especially genes that encode proteins with functions related to photosynthesis

(Ohgishi et al., 2004). cry1 induces Rbcs expression (Martinez-Hernandez et al., 2002). cry1 can induce Lhcb expression but has been suggested to repress Lhcb during high light stress (Mazzella et al., 2001). Consistent with these reports, we observed that

Lhcb mRNA accumulated to lower levels in green seedlings when white light fluence

-2 -1 rates were increased above 50 Pmol m s . Little is known about mechanisms plants use to sense different quantities of light (Jiao et al., 2007). Our finding that Lhcb was repressed at much lower fluence rates when chloroplast biogenesis was blocked than in the untreated green seedlings indicates that the functional and developmental state of chloroplasts has a major impact on the response of PhANGs to fluence rates of light.

In the light, cry1 induces Lhcb, but it is not possible to observe these inductive effects in a cry1 mutant because of redundant induction by phyA and phyB (Mazzella et al., 2001). Moreover, phyA, phyB, and cry1 are all important for the light induction of

Rbcs (Martinez-Hernandez et al., 2002). Interactions between cry1 and phytochromes that impact photomorphogenesis have also been reported previously (Casal, 2006).

Therefore, we expected that the crosstalk between plastid and light signaling pathways that impact PhANG expression when chloroplast biogenesis is blocked might also involve one or more phytochromes in addition to GUN1 and cry1. Indeed, our results showing (1) that GUN1 and cry1 are necessary and sufficient for most if not all

repression of Lhcb in high-fluence-rate blue light but not in white light and (2) that perception of both blue light and red light is essential for maximal repression of Lhcb and Rbcs when chloroplast biogenesis is blocked suggests that phy activity is probably also required for the repression of Lhcb under these conditions.

Both phyA and phyB have been shown to induce Lhcb and Rbcs expression in seedlings that are not treated with inhibitors of chloroplast biogenesis (Reed et al., 1994;

Martinez-Hernandez et al., 2002). However, we found that phyA and phyB regulate

Lhcb differently when chloroplast biogenesis is blocked. phyA remains a positive regulator of Lhcb in seedlings treated with inhibitors of chloroplast biogenesis, which is apparent when GUN1 is inactive. In contrast to phyA, and like cry1, phyB acts as a negative regulator of Lhcb when chloroplast biogenesis is blocked but only in the gun1-

1 background. Although these data indicate that phyB contributes to the repression of

Lhcb in white light when chloroplast biogenesis is blocked, these data also suggest that the repression of Lhcb by phytochromes is likely complex. Our analysis of Rbcs expression in different light conditions also suggests that perception of both blue and red light is critical for maximum repression when chloroplast biogenesis is blocked and that this repression likely requires multiple phytochromes. Consistent with this idea,

Arabidopsis contains five phytochromes (phyA-phyE), and functional redundancies have been reported among some of these phytochromes (Casal, 2006). A comprehensive analysis of phy mutants will likely be required to understand the interactions between plastids and red light that impacts PhANG expression.

Interactions between light and plastid signaling have already been suggested, because light and plastid signals utilize common or adjacent promoter elements (Nott et

al., 2006; Koussevitzky et al., 2007) and phytochrome regulation of PhANGs was reported to be impaired in mutants with defective chloroplasts (Vinti et al., 2005). One interpretation of these data would be that PhANG expression is controlled by a balance between inductive light signaling pathways and repressive plastid signaling pathways, acting independently. In this model, the gun phenotypes of cry1, hy5, and phyB could be explained if cry1, HY5, and phyB enhance plastid stress (e.g., photooxidative stress), thereby enhancing the activity of inhibitors that block chloroplast biogenesis. Although this model is difficult to rule out completely, it is inconsistent with our data and other published results. First, some of these light signaling proteins have been reported to protect chloroplasts from stress. For example, cry1 was previously reported to protect plants from chloroplast stress induced by high-intensity light (Kleine et al., 2007). We observed that cry1, HY5, and GUN1 also protect plants from albinism induced by HL

-2 -1 (Figure 2.9) and that cry1 and GUN1 protect plants from albinism in 125 Pmol m s white light (Figure 2.8). Second, if cry1, HY5, and phyB promoted plastid stress in lincomycin-treated seedlings, plastids in dark-grown lincomycin-treated seedlings would be less stressed than those in light-grown lincomycin-treated seedlings. Such differences in plastid stress might affect plastid size and ultrastructure. However, plastid development has been reported to be similar in light-grown and dark-grown lincomycin- treated seedlings in both wild-type and COP1-deficient backgrounds (Sullivan and Gray,

1999; 2000). Analysis of plastid ultrastructure indicates that GUN1 also does not enhance plastid stress in norflurazon-treated seedlings (Susek et al., 1993).

Nonetheless, analysis of plastid ultrastructure in wild type and gun1 cry1 double mutants would help test this model.

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Our analysis of PhANG expression in seedlings treated with inhibitors of chloroplast biogenesis also suggests that these light signaling proteins most likely do not induce plastid stress. For example, the GUN1 pathway appears to function as a master switch that integrates multiple plastid signals (Koussevitzky et al., 2007). If cry1,

HY5, and phyB promote stress that triggers the GUN1 pathway, a seedling would not have a more robust gun phenotype than observed in gun1-101. In other words, gun1-

101 would be epistatic to cry1, hy5, and phyB; but gun1 mutants are not epistatic to any of these mutants. If these light signaling proteins induce plastid stress that triggers a

GUN1-independent plastid-to-nucleus signaling pathway, we would expect enhanced derepression of PhANG expression in cry1, hy5, and phyB mutants treated with inhibitors of chloroplast biogenesis. However, our analysis of Lhcb expression is consistent with both cry1 and HY5 inducing plastid stress in lincomycin-treated seedlings, but our analysis of Rbcs expression is consistent with HY5 inducing plastid stress and cry protecting plastids from stress under these conditions (Figure 2.4). It is difficult to imagine a simple mechanism in which HY5 could promote plastid stress that simultaneously represses Lhcb and Rbcs, while cry1 concurrently promotes plastid stress that represses Lhcb and in parallel reduces plastid stress that represses Rbcs.

Moreover, because our analysis of Lhcb and Rbcs expression indicates that these

PhANGs are similarly repressed when chloroplast biogenesis is blocked, these data are also not likely explained by Rbcs being more sensitive than Lhcb to chloroplast stress.

Although a mechanism in which cry1, HY5, and phyB promote plastid stress when seedlings are treated with inhibitors of chloroplast biogenesis would need to be complex

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to be consistent with available data, more work will be required to completely rule out this possibility.

A model in which the functional and developmental state of the plastid controls the nature of PhANG regulation by light signaling pathways is consistent with all available data (Figure 2.10). In this model, PhANG expression (1) is repressed by the

GUN1-dependent plastid-to-nucleus signaling pathway and (2) may (e.g., Lhcb) or may not (e.g., Rbcs) be repressed by a plastid signal that converts HY5 from a positive to a negative regulator of PhANGs and is distinct from the plastid signal that is either produced or transduced by a GUN1-dependent pathway. In this model, a plastid signal determines whether cry1 is a positive or a negative regulator of Lhcb and the fluence rate of blue light determines the amount of pathway activity. The conversion of light signaling pathways from positive to negative regulators of Lhcb allows plants to repress

Lhcb more severely than if light signaling pathways remained inductive and simply competed with repressive plastid-to-nucleus signaling pathways. The additional flexibility afforded by integrating light and plastid signals in this manner might facilitate chloroplast biogenesis and repair in diverse light environments.

Because cry1 remains a positive regulator of Rbcs regardless of whether seedlings are treated with inhibitors of chloroplast biogenesis, we might expect that

Rbcs would be expressed at higher levels than Lhcb when chloroplast biogenesis is blocked. However, we found that Lhcb and Rbcs are similarly repressed when chloroplast biogenesis is blocked in white light. These data, our analysis of Lhcb and

Rbcs expression in darkness and in particular light qualities, and our analysis of phy mutants argue that the plastid signals have a broad impact on the nature of PhANG

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regulation by light signaling pathways and that multiple pathways repress PhANGs in response to chloroplast function and developmental state. Thus, the mechanisms by which plastid signals inhibit PhANG expression appears to be more complex than suggested by Koussevitzky et al. (2007).

Plastid signals are required for efficient chloroplast biogenesis

gun1 mutants were shown to have much greater difficulty greening than wild type after prolonged periods of growth in the dark, which is consistent with GUN1 performing an important function during chloroplast biogenesis (Mochizuki et al., 1996). Aside from this phenotype, gun1 mutants have not been reported to have other morphological or pigmentation defects (Nott et al., 2006; Koussevitzky et al., 2007). However, the triggering of the GUN1 pathway by HL implicates this pathway in HL resistance

(Koussevitzky et al., 2007). Indeed we observed that gun1mutants are more sensitive to HL than wild type. Moreover, the striking increase in chlorophyll deficiencies that we

-2 -1 observed in gun1 cry mutants in 125 Pmol m s white light and the synergistic decrease in chlorophyll levels that we observed in gun1 cry and gun1 hy5 double

-2 -1 mutants in 500 Pmol m s white light lead us to several conclusions: (1) these two pathways are required for efficient chloroplast biogenesis; (2) efficient chloroplast biogenesis likely requires two plastid signals, one transduced by the GUN1-dependent pathway and the other by the GUN1-independent pathway that is partially transduced by cry1, phyB, and HY5; and (3) the full impact of plastid-to-nucleus signaling on

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chloroplast biogenesis has been underappreciated because of redundancies between these two plastid-to-nucleus signaling pathways.

-2 -1 In 125 Pmol m s white light, we observed that GUN1 and cryptochromes are more important for chloroplast biogenesis in cotyledons than in primary leaves, which is consistent with previous reports indicating that chloroplast biogenesis in cotyledons and true leaves requires different genes (Yamamoto et al., 2000; Albrecht et al., 2006).

However, the impact of plastid-to-nucleus signaling on chloroplast biogenesis in both cotyledons and primary leaves may be far greater than suggested by these data.

Because we found that the plastid-to-nucleus signaling pathway that controls the nature of PhANG regulation impacts both blue and red light signaling and because we found that GUN1 and at least one other GUN1-independent pathways repress PhANGs when etioplast biogenesis is blocked in the dark, we conclude that plastid-to-nucleus signaling is likely complex. To determining the full impact of plastid-to-nucleus signaling on chloroplast biogenesis it will be necessary to analyze chloroplast biogenesis in mutants in which all plastid-to-nucleus signaling pathways are inactivated.

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MATERIALS AND METHODS

Plant material and growth conditions

hy4-1 was in the Ler ecotype (Ahmad and Cashmore, 1993). All other mutants were in the Columbia-0 (Col-0) ecotype. cry2-1, hy4-1, and all of the T-DNA alleles were obtained from the Arabidopsis Biological Resource Center (Ohio State University).

Seeds were surface sterilized by mixing them in 70% ethanol, 0.5% Triton X-100 solution for 10 min on a tube mixer, then incubating them in 95% ethanol for 10 min on a tube mixer, followed by air drying on filter paper soaked in 95% ethanol in a laminar flow hood. Seeds were plated on Linsmaier and Skoog media containing 2.0% sucrose and 0.5% phytoblend (Caisson Laboratories, Inc). Five PM norflurazon, 0.5 mM lincomycin, or 0.5 mM erythromycin was included in the growth media to block chloroplast biogenesis (Nott et al., 2006); all were purchased from Sigma. Seeds were

-2 -1 stratified for 4 d at 4° C, irradiated with 125 Pmol m s red light for 1 h at 21° C, incubated in the dark for 23 h at 21° C as recommended by Fankhauser and Casal

(2004), and grown for 6 d at 21° C in the specified light conditions in environmentally controlled chambers (Percival Scientific). For experiments in white light, other than HL,

-2 -1 light was provided by broad-spectrum fluorescent tube lamps at 125 Pmol m s . To measure the spectral quality of our white light, we used a StellarNet EPP2000 spectroradiometer (Apogee Instruments). For HL experiments, a combination of high- pressure sodium and metal halide lamps was used. For single light quality experiments and experiments with combinations of blue and red light, seedlings were grown in

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controlled-environment chambers containing light-emitting diodes (Percival Scientific).

In these chambers, the blue light peak was at 470 nm with a spectral bandwidth of 25 nm, the red light peak was at 669 nm with a spectral bandwidth of 25 nm, and the far- red light peak was at 739 nm with a spectral bandwidth of 31 nm. Far-red light was passed through one filter (number 116, Lee Filters) to remove wavelengths that were less than 700 nm. During fluence rate response experiments, white light was filtered through neutral density filters (Roscolux #397, Rosco Laboratories). For far-red light, fluence rates were measured with a StellarNet EPP2000 spectroradiometer (Apogee

Instruments). All other fluence rates were measured with an LI-250A photometer using a PAR sensor (LI-COR Biosciences).

Genetic methods

A Col-0 line harboring an Lhcb1*1:luciferase+ (Lhcb:luc+) reporter gene was mutagenized using EMS as recommended by Weigel and Glazebrook (2002). Pools of

M2 seeds representing approximately 20 to 30 M2 families were surface sterilized and plated in media that contained 5 PM norflurazon as described above. Seeds were

-2 stratified as described above and then incubated for 8 to 9 d in constant 125 Pmol m

-1 s white light at 21° C. Photobleached seedlings were screened for derepression of

Lhcb:luc+ by imaging bioluminescence as recommended by Chinnusamy et al. (2002) using a low-light imaging camera from EG & G Berthold. Bioluminescence of putative mutants was compared to that of the Col-0 Lhcb:luc+ parental line and gun1-1 and gun5

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lines (Mochizuki et al., 2001) in which the Lhcb:luc+ reporter gene was introduced from the Col-0 Lhcb:luc+ parental line by crossing.

For mapping, gun mutants were crossed to a Ler line in which the Lhcb:luc+ reporter gene was introgressed by 12 crosses to Ler. F2 progeny that exhibited a wild-

-2 -1 type hypocotyl phenotype in 25 Pmol m s blue light were used to map cry1-401 with

SSLP markers (Bell and Ecker, 1994) using the Cereon Genomics Indel database

(Jander et al., 2002) and procedures described by Weigel and Glazebrook et al. (2002).

To sequence CRY1 in the mutants isolated from the gun mutant screen, the CRY1

® coding sequence was amplified by means of Platinum Pfx DNA polymerase

(Invitrogen) using CRY1-specific oligonucleotides in at least 10 aliquots that were subsequently pooled, purified from agarose gels using the QIAquick Gel Extraction Kit

(Qiagen), and sequenced with gene-specific oligonucleotides by the Research

Technology Support Facility (Michigan State University).

Oligonucleotides for identifying T-DNA alleles were designed using the recommendations of the Salk Institute Genomic Analysis Laboratory

(http://signal.salk.edu/). Double mutants were identified among progeny of appropriate crosses using SSLP, CAPS (Konieczny and Ausubel, 1993; Bell and Ecker, 1994) or dCAPS markers (Neff et al., 1998).

Hypocotyl measurements

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Seedlings were grown as described above but on media lacking sucrose.

Hypocotyl measurements were performed as recommended by Fankhauser and Casal

(2004).

Analysis of RNA

For RT-PCR, RNA was isolated from 7-d-old seedlings using the RNeasy Plant

Miniprep Kit (Qiagen), including the on-column DNase treatment. First strand cDNA was synthesized from 2 Pg RNA using the Omniscript RT Kit (Qiagen). PCR reactions were programmed with Taq polymerase (Invitrogen) and gene-specific oligonucleotides.

PCR products were analyzed after 20, 25, and 30 cycles. UBQ10 expression was analyzed to test whether the same amounts of cDNA were used for each PCR, as recommended by Weigel and Glazebrook (2002).

For northern blotting, RNA was extracted as described for RT-PCR without the on-column DNase treatment. Northern blotting was performed as recommended by

Chory et al. (1991) with the indicated quantities of RNA. The Lhcb probe was prepared by amplifying the entire open reading frame (ORF) from cDNA clone U13603, which encodes Lhcb1*1 (At1g29920), using CCGGAATTCATGGCCTCAACAAT,

TCCCCGCGGTCACTTTCCGGGAACAA, and Taq DNA polymerase (Invitrogen). The

Rbcs probe was prepared by amplifying part of the Rbcs ORF from cDNA clone U15710, which encodes Rbcs-1A (At1g67090), essentially as described for Lhcb but using

TATGGTCGCTCCTTTCAACG and TGATGCACTGGACTTGACGG. Both cDNA clones were obtained from the ABRC. To prepare each probe, PCR products were purified by

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agarose gel electrophoresis and extracted from gel slices using the QIAquick Gel

Extraction Kit (Qiagen). Purified PCR products were labeled using the Random Primers

DNA Labeling System (Invitrogen). Hybridized RNA blots were imaged using Imaging

Screen K (Bio-Rad) and analyzed using the Molecular Imager FX (Bio-Rad). Lhcb and

Rbcs mRNAs were quantitated using the Quantity One 1-D Analysis Software (Bio-Rad) and normalized to methylene blue-stained 18S rRNA that was quantititated using the same software. Normalization by this method was found to be in the linear range of

2 detection from 2 to 6 Pg of total RNA (R = 0.97). The same relative levels of Lhcb and

Rbcs mRNAs accumulation among genetic backgrounds were observed repeatedly and were consistent within a particular light condition.

Chlorophyll measurements

Chlorophyll was extracted and quantitated as recommended by Porra et al.

(1989).

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ACKNOWLEDGMENTS

We are grateful to Steve Kay (The Scripps Research Institute) for providing the

Arabidopsis line containing the Lhcb::luc+ reporter gene, Joanne Chory (The Salk

Institute for Biological Studies) for providing gun1-1 and gun5 seeds, Xing Wang Deng

(Yale University) for providing cop1-4 seeds, and Neil Adhikari (Michigan State

University) for providing the SSLP marker nda22. We thank Abby Lott, Todd Lydic,

Stephanie Buck, and Jackson Gehan (Michigan State University) for providing helpful assistance during the EMS mutant screen, and Beronda Montgomery-Kaguri (Michigan

State University) for many helpful discussions during the course of this work and helpful comments on this manuscript. We thank Gregg Howe (Michigan State University) for helpful comments on this manuscript. This work was supported by DOE grant no. DE±

FG02±91ER20021 and NSF grant no. IOB 0517841 to RML.

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Figure 2.1 Allelism of new gun mutants and cry1 mutants. A) Similar long hypocotyl phenotypes of new gun mutants and cry1 mutants. -2 -1 Seedlings were grown in 25 µmol m s blue light. Representative seedlings (above) and hypocotyl measurements (below) are shown. Hypocotyls were measured in blue light and in the dark for each line shown. Error bars indicate 95% confidence intervals, n>36. B) Similar gun phenotypes of new gun mutants and cry1 mutants. Seedlings were -2 -1 grown in 125 µmol m s white light on media that either contained (+Lin) or lacked (-Lin) lincomycin. RNA was extracted and Lhcb mRNA levels were determined by northern blotting using 3.0 µg of RNA. The levels of Lhcb transcripts were normalized to total RNA stained with methylene blue. Numbers below each lane indicate the amount of hybridized RNA as a percent of hybridized RNA in untreated wild type grown in the same light condition. C) Rough mapping of the long hypocotyl in blue light phenotype. The wild-type hypocotyl phenotype was rough mapped based on an analysis of 118 chromosomes from F progeny that were obtained from a cry1-401 (Col-0) × Ler cross and 2 -2 -1 exhibited wild-type hypocotyl lengths in 25 µmol m s blue light. Chromosomes were analyzed using two SSLP markers, nda22 and smd1 (Table 2.1). Bacterial artificial chromosome (BAC) clones that contain the SSLP marker sequences for nda22 and smd1 are indicated. The number of centromere proximal recombinants (top) and centromere distal recombinants (bottom) identified with each marker is indicated. D) CRY1 nucleotide and derived amino acid substitutions found in the new gun mutants. The altered codons and the resulting amino acid substitutions found in new cry1 mutants are indicated. Lines and boxes indicate introns and exons, respectively. The photolyase-related (PHR) domain and the DAS motifs have been reviewed by Lin and Shalitin (2003).

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Figure 2.2 Expression of Lhcb and Rbcs in gun1 and cry mutants after chloroplast biogenesis was blocked. -2 -1 -2 -1 Seedlings were grown in either 50 µmol m s blue or 125 µmol m s white light on media that either contained (+Lin) or lacked (-Lin) lincomycin. The levels of Lhcb and Rbcs mRNA were quantitated as described in Figure 2.1B, except that 4.0 µg of RNA was used.

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Figure 2.3 Expression of Lhcb and Rbcs in gun1 and cry1 after chloroplast biogenesis was blocked with various inhibitors of chloroplast biogenesis. -2 -1 Seedlings were grown in 50 µmol m s blue light on media that lacked any inhibitor of chloroplast biogenesis (-Lin) or contained lincomycin (+Lin), erythromycin (+Ert), or norflurazon (+Nfl). Lhcb and Rbcs mRNA levels were quantitated as described in Figure 2.1B, except that 4.0 µg of RNA was used.

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Figure 2.4 Expression of Lhcb and Rbcs in cop1-4 and hy5 mutants after chloroplast biogenesis was blocked. -2 -1 A) gun phenotypes of cop1-4 . Seedlings were grown in 50 µmol m s blue light on media that contained (+Lin) or lacked (-Lin) lincomycin. Lhcb and Rbcs mRNA levels were quantitated as described in Figure 2.1B. B) gun phenotypes of hy5. The growth of seedlings and the quantitation of the levels of Lhcb and Rbcs mRNA were as described in (A).

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Figure 2.5 Expression of Lhcb and Rbcs in the dark or in various qualities of light. A) Lhcb and Rbcs expression levels in darkness, white light, blue light, red light, and far-red light. Seedlings were grown on media containing lincomycin in the -2 -1 -2 -1 -2 -1 dark or in 125 µmol m s white light, 25 µmol m s blue light, 35 µmol m s -2 -1 red light, or 2 µmol m s far-red light. The levels of Lhcb and Rbcs mRNA were quantitated as described in Figure 2.1B, except that 2.5 µg of RNA was used. B) Expression of Lhcb and Rbcs in white light and in a combination of blue and red light. Seedlings were grown on media containing (+Lin) or lacking (-Lin) lincomycin in either white (W) or a combination of blue and red (B+R) light. Fluence rates were as described in (A). The levels of Lhcb and Rbcs mRNA were quantitated as described in (A). The number below the B+R (-Lin) lane indicates the amount of hybridized RNA as a percentage of mRNA in the untreated control grown in W (-Lin).

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Figure 2.6 Expression of Lhcb and Rbcs in gun1, phyA, and phyB mutants after chloroplast biogenesis was blocked. -2 -1 Seedlings were grown in 125 µmol m s white light on media that lacked (- Lin) or contained (+Lin) lincomycin. The levels of Lhcb and Rbcs mRNA were quantitated as described in Figure 2.1B

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Figure 2.7 The effects of plastid development on the fluence rate response of Lhcb and Rbcs. - Seedlings were grown in white light fluence rates of 0, 1.0, 10, 50, or 125 µmol m 2 -1 s in either the presence (+Lin) or absence (-Lin) of lincomycin. The levels of Lhcb and Rbcs mRNA were quantitated as described in Figure 2.1B. For untreated wild type, the number below each lane indicates the amount of hybridized RNA as a percent of hybridized RNA in untreated wild type grown in -2 -1 125 µmol m s white light.

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Figure 2.8 Chlorophyll-deficient cotyledons in gun1 and gun1 cry mutants. A) Percentage of seedlings exhibiting chlorophyll-deficient phenotypes. Seedlings were grown on media without an inhibitor of chloroplast biogenesis. The total number of seedlings and the seedlings that were visibly chlorophyll deficient were -2 -1 counted after 6 d of growth in 125 µmol m s white light. Four independent experiments were performed and each experiment contained a total of approximately fifty seedlings. Error bars represent 95% confidence intervals between independent experiments. B) Chlorophyll-deficient seedlings. Representative wild type (Col-0) and representative chlorophyll-deficient mutant seedlings are shown after 6 (Col-0, gun1-101, gun1-1 cry1, gun1-1 cry1 cry2) or 7 (gun1-1 and gun1-1 cry1) d of growth in white light. Arrows indicate chlorophyll-deficient areas. Bars = 2 mm.

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Figure 2.9 HL sensitivity of gun1 and light signaling mutants. A) gun1 and light signaling mutant seedlings grown in the indicated fluence rates of continuous white light. One-d-old etiolated seedlings were irradiated with the indicated fluence rates of continuous white light for 7 d. Representative seedlings are shown. Bars = 2 mm. B) Comparisons of total chlorophyll levels in gun1 and light signaling mutants in various fluence rates of continuous white light. Seedlings were grown as described in (A). Chlorophyll was extracted from at least three samples for each line in each condition. Error bars represent 95% confidence intervals.

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Figure 2.10 Model for PhANG regulation by a network of plastid and light signaling pathways. The current model for the GUN1-dependent plastid-to-nucleus signaling pathway was adapted from Koussevitzky et al. (2007). In this model a second messenger (indicated with Y) that requires GUN1 for either its production or transduction (dotted arrows) triggers a plastid-to-nucleus signaling pathway that represses PhANGs. A plastid signal(s) that is independent of GUN1 (indicated with X), represses both Lhcb and Rbcs in the dark (not shown) and also converts cry1 and one or more photoreceptors that perceive red light into negative regulators of Lhcb and Rbcs. cry1 becomes a negative regulator of Lhcb when X converts HY5 from a positive to a negative regulator of Lhcb. Under these same conditions, Rbcs is induced by cry1 and simultaneous repressed by a combination of blue and red light.

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Figure 2.11 The gun mutant screen procedure. Ethyl methanesulfonate (EMS); other abbreviations are defined in the text.

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Figure 2.12 Analysis of T-DNA alleles. A) Map of the GUN1 gene. The T-DNA flanking sequence is represented as a green arrow. RT-PCR primer locations are represented by black arrows. B) Truncation of GUN1 mRNA in gun1-101. RT-PCR analysis was performed with gene-specific primers that either target the gene product upstream of the predicted T- DNA insertion site (Rp+Lp1) or span the insertion site (Rp+Lp2). The UBQ10 transcript was used as a control. C) The effects of T-DNA insertions on the expression of CRY1, HY5, PHYA, PHYB, PHOT1, PHOT2, NPH3 and CRY3. RT-PCR analysis was performed using primers that span the T-DNA insertion site in their respective genes. The UBQ10 transcript was used as a control. PPR, pentatricopeptide repeat; SMR, small mutS-related domain

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Figure 2.13 Expression of Lhcb in cry1 after chloroplast biogenesis was blocked in red light. -2 -1 Seedlings were grown in 35 µmol m s red light in the presence of lincomycin (+ Lin) and the absence of Lincomycin (-Lin). RNA was analyzed as described in Figure 2.1B.

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Figure 2.14 Lhcb and Rbcs expression in gun1-1 and various photoreceptor mutants after chloroplast biogenesis was blocked in blue light. Seedlings were grown on media that contained lincomycin (+Lin) or lacked -2 -1 lincomycin (-Lin) under 50 µmol m s blue light. RNA was analyzed as described in Figure 2.1B.

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Figure 2.15 Lhcb and Rbcs mRNA levels in wild type and gun1 mutants grown in darkness and various light qualities without inhibitors of chloroplast biogenesis. Seedlings were grown in the indicated light conditions on media that did not contain an inhibitor of chloroplast biogenesis. Fluence rates for each light quality were the same as in Figure 2.5 in the text. RNA was analyzed as described in Figure 2.5 in the text.

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Figure 2.16 Analysis of det/cop/fus phenotypes in gun1-1 and cop1-4. Seedlings were grown in the dark on media that contained lincomycin (+Lin) or lacked lincomycin (-Lin). RNA was analyzed as described in Figure 2.1B.

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Figure 2.17 Chlorophyll-deficient leaves in gun1 and gun1cry1 double mutants. -2 -1 Seedlings were grown in 125 µmol m s white light. Two-week-old seedlings with both green and partially chlorophyll-deficient true leaves are shown. Arrows indicate some of these chlorophyll deficiencies. Bars = 2 mm.

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Figure 2.18 Phenotypes of gun1 and light signaling mutants following HL incubations. Green plants were transferred to HL as described in the text. gun1-101 is indicated as gun1 for clarity. Bars = 2 mm.

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Figure 2.19 Cosegregation analysis of long hypocotyl in blue light and gun

phenotypes. cry1-403 was backcrossed and four F2 seedlings that displayed long hypocotyls in blue light were propagated for segregation analysis. F seedlings were grown 3 -2 -1 in 50 ȝmol m s blue light on media that contained lincomycin (+Lin). Lhcb expression was analyzed in four F3 lines in which the long hypocotyl phenotype did not segregate. Quantitation of Lhcb mRNA levels was as described in Figure 2.1B except that numbers below each lane indicate fold increase relative to wild type.

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-2 -1 Table 2.1 Segregation of the long hypocotyl phenotype in 25 ȝmol m s blue light in F seedlings. 3 1. F mutant line, 2. Number of seedlings with long hypocotyls, 3. Total number 3 of seedlings, 4. Percentage of seedlings with long hypocotyls. Less than 100% long hypocotyls in some lines may be caused by unlinked mutations.

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CHAPTER 3

PLASTID SIGNALS THAT AFFECT PHOTOMORPHOGENESIS IN ARABIDOPSIS THALIANA ARE DEPENDENT ON GENOMES UNCOUPLED 1 AND CRYPTOCHROME 1

This research was originally published in New Phytologist. Michael E. Ruckle, Robert M. Larkin. Plastid signals that affect photomorphogenesis in Arabidopsis thaliana are dependent on GENOMES UNCOUPLED 1 and cryptochrome 1. New Phytologist 2009; Vol. 182: pp. 367-79 © 2009 New Phytologist Trust.

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PLASTID SIGNALS THAT AFFECT PHOTOMORPHOGENESIS IN ARABIDOPSIS THALIANA ARE DEPENDENT ON GENOMES UNCOUPLED 1 AND CRYPTOCHROME 1

ABSTRACT When plastids experience dysfunction they emit signals that help coordinate nuclear gene expression with their functional state. One of these signals can remodel a light signaling network that regulates the expression of nuclear genes that encode particular antenna proteins of photosystem II. These findings led us to test whether plastid signals might impact other light regulated processes. We monitored photomorphogenesis in genomes uncoupled 1 (gun1), cryptochrome 1 (cry1), and long hypocotyl 5 (hy5), which have defects in light and plastid signaling, by growing seedlings under various light conditions and either treating or not treating them with antibiotics that induce chloroplast dysfunction and trigger plastid signaling. We found that plastid signals that depend on GUN1 can affect cotyledon opening and expansion, anthocyanin biosynthesis, and hypocotyl elongation. We also found that plastid signals that depend on CRY1 can regulate cotyledon expansion and development. Our findings suggest that plastid signals triggered by plastid dysfunction can broadly affect photomorphogenesis and that plastid and light signaling can promote or antagonize each other, depending on the responses studied. These data suggest that GUN1 and cry 1 help to integrate chloroplast function with photomorphogenesis.

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INTRODUCTION

Light is one of the most important environmental signals perceived by plants. It is required for chloroplast biogenesis and photosynthesis, regulates metabolism, entrains the circadian clock, and has striking effects on organ growth and development

(Jiao et al., 2007). Many photoreceptors and downstream signaling components that transduce these light signals have been identified. Mechanisms for light signaling networks have been partially defined and are characterized by extensive interactions and redundancies (Casal, 2006; Jiao et al., 2007).

Driving chloroplast biogenesis and function is perhaps one of the most important functions of light-regulated signaling networks because photosynthesis provides the energy required for growth, development, and reproduction. Light regulates chloroplast biogenesis and function by controlling the expression of genes that encode proteins with chloroplast-related functions (Tyagi & Gaur, 2003; Monte et al., 2004; Ohgishi et al.,

2004; Toyoshima et al., 2005; Marín-Navarro et al., 2007) and activating a light- dependent enzyme in the chlorophyll biosynthetic pathway (Tanaka & Tanaka, 2007).

However, the plastid is not completely subordinate to light conditions. Plastids emit signals that help coordinate nuclear gene expression with the functional state of the plastid. Plastid signals have been reported to coordinate the expression of photosynthesis-associated nuclear genes (PhANGs) and stress-related nuclear genes with plastid function (Strand et al., 2003; Pesaresi et al., 2007; Kim et al., 2008; Dietzel et al., 2008). Plastid signals also contribute to efficient chloroplast biogenesis

(Mochizuki et al., 1996; Ruckle et al., 2007), optimize the stoichiometry of photosystems

I and II (Dietzel et al., 2008), and trigger stress and programmed cell death responses

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(Nott et al., 2006; Kim et al., 2008). At least some of these plastid signals can be transduced in the dark (Sullivan et al., 1999; Ruckle et al., 2007).

Plastid signals have been reported to affect light responses. Because a subset of mutants with defective plastid-localized proteins exhibits aberrant leaf anatomy, plastid signals are thought to contribute to proper leaf morphogenesis (Rodermel 2001;

Yu et al., 2007). Additionally, plastid signals are thought to be required for mesophyll cell division and to promote mesophyll cell elongation when leaves acclimate to high- intensity light (Tan et al., 2008). Also, an analysis of the chlorophyll-deficient chlorophyll a/b-binding (CAB) protein-underexpressed (cue) mutants gives evidence that plastid signals can have mild inhibitory effects on cotyledon opening and hypocotyl elongation in continuous red and far-red light (Vinti et al., 2005). The long after far-red 6 (laf6) mutant, which has a defect in a plastid-localized ATP-binding-cassette protein, exhibits a striking increase in hypocotyl elongation in far-red light (Møller et al., 2001). The photomorphogenic phenotype of laf6 was proposed to result from accumulation of protoporphyrin IX that might act as a signaling factor (Møller et al., 2001) or defects in iron homeostasis (Xu et al., 2005). Alternatively, Cornah et al. (2003) suggested abnormal tetrapyrrole metabolism in laf6 may lead to reduced levels of phytochromobilin, the tetrapyrrole chromophore of phytochrome, and therefore lower levels of active phytochromes in laf6 relative to wild type.

In seedlings treated with inhibitors of chloroplast biogenesis, two genetically distinct plastid signals repress genes that encode the light harvesting chlorophyll a/b- binding proteins of photosystem II (Lhcb) when plastids experience dysfunction. One of these plastid signals is dependent on GENOMES UNCOUPLED 1 (GUN1), which

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encodes a plastid-localized pentatricopeptide repeat protein that colocalizes with nucleoids (Koussevitzky et al., 2007). The other plastid signal affects a light signaling network that depends in part on the basic leucine zipper transcription factor long hypocotyl 5 (HY5) acting downstream of the blue light photoreceptor cryptochrome 1

(cry1) (Ruckle et al., 2007). The finding that knocking out both of these signals causes a synergistic up-regulation of Lhcb expression implies that the full impact of plastid signals on various processes may have been obscured by genetic complexity during previous analyses of plastid signaling mutants. To test whether the plastid signals that are dependent on GUN1 and light signaling networks contribute to photomorphogenesis, we quantitated a variety of commonly studied photomorphogenic processes

(Fankhauser & Casal, 2004) in gun1 and light signaling mutants of Arabidopsis thaliana treated with inhibitors of chloroplast biogenesis in a variety of light qualities and quantities. Our results indicate that plastid signals that are dependent on GUN1 and cry1 can broadly affect photomorphogenesis.

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RESULTS

Analysis of cotyledon opening and expansion in gun1 and light signaling mutants

We expected that GUN1 might have a greater impact on photomorphogenesis of the cotyledons in low-fluence rates than in high-fluence rates because (1) the light signaling network that represses Lhcb is more important to the repression of Lhcb when fluence rates are increased and (2) GUN1-dependent plastid signals contribute to the repression of Lhcb regardless of light conditions (Ruckle et al., 2007). To test this hypothesis, we analyzed the fluence-rate responses of cotyledon opening and expansion under conditions that either induced or did not induce plastid signaling. We tested these fluence rate responses in blue light because the effects of plastid signals on blue light signaling are better understood than the effects of plastids on the components of the light signaling network that perceive other light qualities (Ruckle et al., 2007). To trigger plastid-to-nucleus signaling, we treated seedlings with lincomycin and erythromycin, which inhibit plastid translation (Mulo et al,. 2003). These inhibitors have been shown to block both etioplast and chloroplast biogenesis (Sullivan & Gray,

1999) and to repress PhANG expression in both light and dark (Sullivan & Gray, 1999;

Ruckle et al., 2007). Seedlings treated with these inhibitors have nonphotosynthetic plastids rather than chloroplasts, are viable when provided sucrose, and become green upon removal of the inhibitor (Sulliavn & Gray, 1999; Ruckle et al., 2007). GUN1 has been localized to chloroplasts and is thought to also reside in the nonphotosynthetic plastids found in the aerial parts of lincomycin- and erythromycin-treated seedlings where it contributes to the biosynthesis or transduction of plastid signals (Koussevitzky et al., 2007). We observed that cotyledon opening requires higher fluence rates of blue

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light when wild-type seedlings were treated with lincomycin compared to untreated wild- type seedlings (Figure 3.1 and 3.10). In contrast, cotyledon opening exhibited a similar fluence rate response in treated gun1-1, untreated gun1-1, and untreated wild type

(Figure 3.1 and 3.10). gun1-1 is a leaky allele (Koussevitzky et al., 2007). Similar results were obtained with lincomycin and erythromycin (Figure 3.11). Erythromycin and lincomycin both inhibit plastid translation, but by different mechanisms (Mulo et al.,

2003). These data indicate that (1) the observed differences in cotyledon opening are likely caused by reduced GUN1 activity in the mutant and not by resistance to particular inhibitors and (2) that GUN1-dependent plastid signals can repress cotyledon opening in low-fluence-rate blue light.

We observed that cotyledon expansion increased similarly in gun1-1 and wild- type seedlings that were not treated with an inhibitor of chloroplast biogenesis as the fluence rate of blue light increased (Figure 3.10 and 3.12A). This relationship between cotyledon expansion and fluence rate of blue light has been reported previously for untreated wild-type seedlings (Jackson & Jenkins, 1995; Ohgishi et al., 2004). In contrast, when seedlings were treated with lincomycin, cotyledon expansion was significantly greater in gun1-1 compared to wild type, but these differences diminished

-2 -1 -2 above 5 µmol m s blue light and there were no significant differences in 50 µmol m

-1 s blue light (Fig. 3.11 and 3.12A). These data indicate that GUN1-dependent plastid signals can inhibit cotyledon expansion in lincomycin-treated seedlings and that high- fluence-rate blue light can also repress cotyledon expansion when chloroplast biogenesis is blocked regardless of whether GUN1 is active.

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In high fluence rates, the cotyledons of wild-type seedlings curled downward

(Figure 3.10 and 3.15A). To account for this effect, we repeated this fluence response experiment but fixed and flattened cotyledons before measuring cotyledon area (Figure

3.2). When wild type seedlings were treated with inhibitors of chloroplast biogenesis in

-2 -1 50 µmol m s blue light, the flattened cotyledons of lincomycin-treated wild type were

50% larger than cotyledons that were not processed in this manner (Figure 3.12B).

These data indicate that accounting for the curling of the cotyledons is most important

-2 -1 when measuring cotyledon areas in treated seedlings grown in 50 µmol m s blue light. Cotyledon expansion was greater in gun1-1 than in wild type, regardless of whether chloroplast biogenesis was blocked with lincomycin or erythromycin (Figure

3.13A, B and C). Similar results were obtained with gun1-101 (Figure 3.13A, B and C), a T-DNA allele that appears to be a null (Ruckle et al., 2007).

Because plastid signaling triggered by inhibitors of chloroplast biogenesis had been shown to affect the nature of Lhcb gene regulation by cry1 and HY5 (Ruckle et al.,

2007) and because cry1 had been shown to affect the photomorphogenesis of the cotyledons in high-fluence-rate blue light (Jackson & Jenkins, 1995; Neff & Chory, 1998;

Ohgishi et al., 2004), we tested whether cry1 and HY5 contribute to the repression of cotyledon opening and expansion in high-fluence-rate blue light. We grew gun1-1, cry1, and hy5 mutants in the presence and absence of inhibitors of chloroplast biogenesis in

-2 -1 50 µmol m s blue light. cry1 and hy5 are T-DNA insertion alleles that are known to

-2 -1 be nulls (Ruckle et al., 2007). In 50 µmol m s blue light, cotyledons were completely unfolded in gun1-1 cry1, the single mutants, and wild type, regardless of whether

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seedlings were treated with inhibitors of chloroplast biogenesis (Figure 3.1, 3.3B, and

3.10). When plants were not treated with inhibitors of chloroplast biogenesis in this same fluence rate of blue light, no difference was observed in cotyledon areas, except for cotyledons of cry1 hy5, which were two-thirds the size of wild type (Figure 3.3A and

B).

When seedlings were treated with inhibitors of chloroplast biogenesis in 50 µmol

-2 -1 m s blue light, the areas of the cotyledons in gun1 or cry1 were indistinguishable from wild type (Figure 3.3A and B). In contrast, the areas of the cotyledons increased in the gun1-1 cry1 double mutant by approximately 50%, but were still approximately half the size of untreated seedlings (Figure 3.3A and B). Because the gun1 cry1 double mutant has larger cotyledons than wild type or the single mutants, we conclude that either the cry1- or the GUN1-dependent plastid signals can compensate for the loss of the other and that these signals are functionally redundant for the repression of cotyledon expansion when chloroplast biogenesis is blocked. Additionally, because knocking out the CRY1 gene ameliorates the repressive effect of high-fluence-rate blue light on cotyledon expansion, we conclude that this repression likely depends on a functional cry1 photoreceptor. Cotyledon areas were indistinguishable among hy5, gun1 hy5, and cry1 hy5 double mutants that were treated with inhibitors of chloroplast

-2 -1 biogenesis (Figure 3.3A and B). In contrast to 50 µmol m s blue light, we found that gun1-1 had larger cotyledons than wild type when chloroplasts biogenesis was blocked

-2 -1 in 100 µmol m s red light (Figure 3.4A, B, and C). Moreover, we found that phyB

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remained a positive regulator of cotyledon expansion regardless of whether seedlings are treated with inhibitors of chloroplast biogenesis (Figure 4A, B, and C).

Analysis of epidermal cells in gun1 and cry1 mutants

The light-driven expansion of the cotyledons is largely a consequence of cell expansion rather than cell division (Neff & Van Volkenburgh, 1994; Stoynova-Bakalova et al., 2004). To test whether plastid signals might affect cell expansion in the cotyledons, we examined cotyledon surfaces using scanning electron microscopy

(SEM). The epidermal cells of the cotyledons from untreated wild-type and mutant seedlings all exhibited a similar pattern of interdigitating lobes that are typical of epidermal pavement cells (Figure 3.5A). In fact, these pavement cells were comparable in area, shape, and stomatal density (Figure 3.14A, B, and C). In contrast, when wild- type seedlings were treated with inhibitors of chloroplast biogenesis, the pavement cells lacked the interdigitated lobes of untreated seedlings but had the same surface area and stomatal density as untreated wild type (Fig. 3.5B, 3.14A, B and C). The pavement cells of treated gun1-1 and cry1 were half the size of treated wild type and the density of stomata was strikingly increased in treated gun1-1 relative to the wild type (Figure 3.5B,

3.14A and C). In contrast to the treated single mutants and wild type, pavement cell area, shape, and density of stomata were very similar in treated gun1-1 cry1 and untreated seedlings (Figure 3.5B, 3.14A, B and C). These data indicate that when chloroplast biogenesis is blocked, the development of abnormal epidermal cells is dependent on GUN1 and cry1.

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Analysis of anthocyanin levels in gun1 and cry1 mutants

Anthocyanins are induced by photoreceptors and stress. These pigments function in part to protect the chloroplast from high-fluence-rate light-induced photooxidative stress and photoinhibition (Gould et al., 2004). Because high-fluence- rate light induces both ROS-dependent (Nott et al., 2006; Kim et al., 2008) and GUN1- dependent plastid signals (Koussevitzky et al., 2007), we tested whether GUN1- dependent plastid signals also contribute to anthocyanin biosynthesis. We measured

-2 -1 anthocyanin levels in seedlings grown in 50 µmol m s blue light to learn whether plastid signals affect the important role of cry1 and HY5 in the induction of anthocyanin biosynthesis (Ahmad et al., 1995; Jackson & Jenkins, 1995; Shin et al., 2007;

Vandenbussche et al., 2007). We found that gun1-1 and gun1-101 contained fourfold less anthocyanin than wild type, but only when chloroplast biogenesis was blocked, and that cry1 and hy5 contained 1.5- to 2.0-fold less anthocyanin than wild type, regardless of whether chloroplast biogenesis was blocked (Figure 3.6A and B). Additionally, anthocyanin levels were 1.5- and 3.0-fold lower in the gun1-1 cry1 and gun1-101 hy5 double mutants, respectively, than in the single mutants, but only when chloroplast biogenesis was blocked (Figure 3.6A and B). These data indicate that GUN1- dependent plastid signals do not induce anthocyanin biosynthesis by affecting cry1 or

HY5 activity and that GUN1-dependent plastid signals have a greater effect on the induction of anthocyanin biosynthesis than cry1 and HY5 when chloroplast biogenesis is blocked in blue light.

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We also tested whether plastid signals induce anthocyanins in high-intensity white light because, as stated above, high-intensity light triggers plastid signaling (Nott et al., 2006; Koussevitzky et al., 2007; Kim et al., 2008), plastid signals contribute to successful greening in high-fluence-rate light (Ruckle et al., 2007), and anthocyanin- deficient plants exhibit an enhanced sensitivity to photooxidative stress (Gould, 2004).

We allowed seedlings to germinate in the dark for 23 h and then transferred them to

-2 -1 1000 µmol m s continuous white light for six days. Under these conditions, gun1, cry1, hy5 and the corresponding double mutants are chlorophyll deficient relative to the wild type (Ruckle et al., 2007). We found that anthocyanin levels were indistinguishable in gun1 mutants and wild type, but anthocyanins accumulated to 3- and 50-fold lower in cry1 and hy5, respectively, compared to wild type. Similarly, anthocyanins accumulate to 30- and 500-fold lower levels in the gun1 cry1 and gun1 hy5 double mutants, respectively, than in wild type and anthocyanin levels were fivefold lower in the gun1-

101 cry1 double mutant than in gun1-1 cry1 (Figure 3.7A and B). These data indicate that although cry1 and HY5 are major regulators of anthocyanin biosynthesis in 1000

-2 -1 µmol m s white light, GUN1 can contribute to anthocyanin biosynthesis when seedlings de-etiolate under these conditions and either cry1 or HY5 is absent.

Analysis of hypocotyl elongation in gun1 mutants

To test whether the effects of plastid signals on photomorphogenesis are restricted to photosynthetic organs or whether they can also affect the growth of nonphotosynthetic organs, we tested the effect of plastid signals on light-regulated

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elongation of the hypocotyl. cry1 and HY5 contribute to the light-dependent inhibition of hypocotyl elongation in untreated seedlings; for cry1, these effects are more striking in blue light (Koornneef et al., 1980; Chory, 1992; Ahmad & Cashmore, 1993; Ang & Deng,

1994). In blue light, gun1 mutants and wild type have similar hypocotyl lengths.

However, when seedlings are treated with lincomycin, gun1-101 did have 1.3- to 1.4- fold shorter hypocotyls than wild type in blue light (Figure 3.3B and 3.8A). cry1 and hy5 have 2- to 4-fold longer hypocotyls than gun1 and wild type, regardless of whether chloroplast biogenesis is blocked (Figure 3.3B and 3.8A). In white light, the hypocotyls of untreated cry1, gun1, and wild type are essentially the same length, and the hypocotyls of untreated hy5 are 1.7-fold longer than cry1, gun1, and wild type. However, when treated with inhibitors of chloroplast biogenesis, cry1 and hy5 have 1.6- to 4.3-fold longer hypocotyls than treated wild type (Figure 3.8B and 3.15A). These data indicate that cry1 and HY5 are negative regulators of hypocotyl elongation, regardless of whether seedlings are treated with inhibitors of chloroplast biogenesis, and that cry1 and HY5 have more important roles in regulating hypocotyl elongation when seedlings are treated with inhibitors of chloroplast biogenesis than in untreated seedlings in white light. In the dark, all of the hypocotyl lengths were the same, whether or not seedlings were treated with inhibitors of chloroplast biogenesis (M.E. Ruckle & R.M. Larkin, unpublished data). We found that hypocotyls in untreated cry1 hy5 are 1.5 and 4 times as long as hypocotyls in corresponding untreated single mutants in blue and white light, respectively (Figure 3.3B, 3.8A, B, and 3.15A). These findings indicate that cry1 and

HY5 can only partially compensate for each other in the inhibition of hypocotyl elongation under these conditions.

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The enhanced hypocotyl lengths of treated cry1 and hy5 seedlings in white light lead us to suggest that plastid signals can stimulate hypocotyl elongation. To test this possibility, we inhibited plastid-to-nucleus signaling in cry1 and hy5 backgrounds by preparing gun1-101 cry1 and gun1-101 hy5 double mutants. We observed that hypocotyls were 2- to 3-fold shorter in gun1-101 hy5 relative to hy5 in both light conditions and 1.4-fold shorter in gun1-101cry1 relative to cry1 in blue light (Figure 3.3B,

3.8A, B, and 3.15A). To test whether GUN1 can stimulate hypocotyl elongation under other conditions that trigger GUN1-dependent plastid signaling, we measured the hypocotyl lengths of seedlings grown in continuous high-fluence-rate white light. We observed that high-fluence-rate white light stimulated hypocotyl elongation fourfold in hy5 relative to wild type and gun1-101 and that the longer hypocotyls of hy5 were suppressed in the hy5 gun1-101 double mutant (Figure 3.8C and 3.15B).

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DISCUSSION

Plastid signals repress the photomorphogenesis of the cotyledons

The cryptochromes are primarily responsible for photomorphogenesis in blue light (Lin & Shalitin 2003; Ohgishi et al., 2004). Because we found that GUN1-

-2 -1 dependent plastid signals inhibit cotyledon opening in 5 µmol m s blue light and in lower fluence rates of blue light, we suggest that plastid signals likely act downstream of the light signaling network that inhibits cotyledon opening in these fluence rates.

Cotyledon opening was independent of plastid signals when the fluence rate of blue light was increased. Increased photoreceptor activity in high-fluence-rate blue light (Lin et al., 1998; Neff & Chory, 1998; Ohgishi et al., 2004) may explain these data.

In untreated seedlings, cry1 is a positive regulator of cotyledon expansion in blue light (Jackson & Jenkins, 1995; Neff & Chory, 1998; Ohgishi et al., 2004), but the contribution of cry1 to cotyledon expansion is lower when the fluence rate of blue light is increased (Jackson & Jenkins, 1995). Our analysis of cotyledon areas in cry1, hy5, and

- cry1 hy5 indicates that cry1 is a positive regulatoURIFRW\OHGRQH[SDQVLRQLQımol m

2 -1 s blue light and that HY5 can compensate for a loss of cry1 activity in this fluence rate of blue light. Consistent with this interpretation, other photoreceptors have been reported to contribute to cotyledon expansion in high-fluence-rate blue light (Neff &

Chory, 1998; Ohgishi et al., 2004) and HY5 appears to act downstream of multiple photoreceptors (Jiao et al., 2007). A similar model likely explains our finding that cry1 hy5 has longer hypocotyls than the corresponding single mutants.

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We also found that when chloroplast biogenesis is blocked, cry1 contributes to the repression rather than the promotion of cotyledon expansion (Figure 3.9A). These data indicate that the conversion of cry1 from a positive to a negative regulator by plastid signals likely impacts the expression of nuclear genes that affect cotyledon expansion and not only of Lhcb genes, as has been previously shown (Ruckle et al.,

2007). Plastid signals convert cry1 from a positive to a negative regulator of Lhcb genes by a mechanism that involves converting HY5, which acts downstream of cry1, from a positive to negative regulator of Lhcb genes (Ruckle et al., 2007). Because in contrast to Lhcb, the conversion of cry1 from a positive to a negative regulator of cotyledon expansion does not appear to involve HY5, plastid signals must regulate cotyledon expansion by a mechanism that does not require HY5 or at least also utilizes other factors that can compensate for a loss of HY5 activity. In animal systems, nuclear receptors and Myc can function as positive or negative regulators of transcription depending on which coregulators or transcription factors they bind (Wanzel et al., 2003;

Feige & Auwerx 2007; Kassel et al. 2007).

Hormones were previously reported to cause abnormal curling of leaves and cotyledons (Keller & Van Volkenburgh,1997; Hamant et al., 2002; Kakiuchi et al., 2007).

Because we found that the cotyledons of wild type but none of the mutants curled

-2 -1 downward when chloroplast was blocked in 50 µmol m s blue light but not in lower fluence rates of blue light, we suggest that plastid signals might affect hormone

-2 -1 biosynthesis or responses in 50 µmol m s blue light. Additionally, because we found

-2 -1 that the cotyledons of gun1-1 were larger than wild type in 100 µmol m s red light but

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-2 -1 not 50 µmol m s blue light, we conclude that unlike blue light, red light probably does not repress cotyledon expansion when chloroplast biogenesis is blocked. We cannot rule out the possibility that red light might contribute to the inhibition of cotyledon expansion when combined with a distinct light quality, such as blue light. Consistent with this idea, interactions between light signaling pathways that are triggered by a combination of blue and red light have been reported previously (Casal, 2006).

We found that the pavement cells in the epidermis of cotyledons from wild-type seedlings treated with inhibitors of chloroplast biogenesis did not exhibit a jigsaw-puzzle appearance that is typical of untreated seedlings (Smith, 2003; Panteris & Galatis,

2005). The finding that in contrast to wild type and the single mutants, the epidermis of treated gun1cry1 bears striking resemblance to the epidermis of untreated seedlings suggests that at least two plastid signals, one dependent on GUN1 and the other transduced by cry1, are part of a signaling network that can regulate cell differentiation in the epidermis (Figure 3.9A). A role for plastids in the development of the epidermis has been noted previously. For example, similar abnormal epidermal cells have been observed in the pale cress (pac) mutant of Arabidopsis and the defective chloroplasts and leaves-mutable (dcl-m) mutant of tomato, which are both deficient in chloroplast- localized proteins that participate in RNA processing (Keddie et al., 1996; Meurer et al.,

1998; Reiter et al., 1994; Tirlapur et al., 1999; Bellaoui et al., 2003).

We expected that differences in pavement cell area (Figure 3.14A) would explain gun1 cry1 having larger cotyledons than the single mutants (Figure 3.3). However, the observation that the cotyledons of treated wild type have pavement cells of

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approximately the same size as gun1 cry1 (Figure 3.14A) but smaller cotyledons than treated gun1 cry1 (Figure 3.3) would seem paradoxical. One possible explanation for these data might lie in the rough appearance of the epidermis in treated wild type

(Figure 3.5B). Indeed, from a light microscopy analysis of cotyledon sections, we found that the epidermis of treated wild type is extensively invaginated relative to the treated mutants (Figure 3.16A and B). These data indicate that measurements of cotyledon area provide a less accurate estimate of cotyledon surface area in treated wild type than in these treated mutants.

One or more GUN1-dependent plastid signals affect anthocyanin biosynthesis and elongation of the hypocotyl

Both cry1 and HY5 had been reported to be important positive regulators of anthocyanin biosynthesis, but whether plastid signals affect anthocyanin biosynthesis has not been previously reported. We induced GUN1-dependent plastid signaling by treating seedlings with either lincomycin or high-fluence-rate white light. Lincomycin provides a complete and uniform block to greening among wild type and all of the mutants tested. In contrast, high-fluence-rate white light inhibits greening most in the double mutants tested, less in the single mutants tested, and least in the wild type

(Ruckle et al., 2007). We observed that accumulation of anthocyanins is dependent on

GUN1 only when chloroplast biogenesis is severely impaired by lincomycin treatments or by high-fluence-rate light treatments of gun1 cry1 and gun1 hy5 double mutants

(Figure 3.9B). These findings are consistent with GUN1-dependent plastid signals

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inducing anthocyanin biosynthesis when plastids experience dysfunction to protect plastids from excess light and to scavenge reactive oxygen species. An alternative interpretation, that gun1 mutants contain fewer anthocyanins than wild type because they experience less plastid dysfunction than wild type during these treatments, is inconsistent with previous reports, which indicate that gun1 mutants experience greater or equal plastid dysfunction compared to wild type (Susek et al., 1993; Mochizuki et al.,

1996; Koussevitzky et al., 2007; Ruckle et al., 2007).

Our findings also suggest that GUN1-dependent plastid signals can contribute to the elongation of hypocotyls and that cry1 and HY5 contribute to the repression of hypocotyl elongation regardless of the functional status of the plastid. The finding that plastid signals remodel cry1 signaling that affects Lhcb expression (Ruckle et al., 2007) and cotyledon expansion but not hypocotyl elongation is consistent with (1) the distinct cotyledon and hypocotyl transcriptome responses to light (Ma et al., 2005) and (2) the composition of downstream components of light signaling pathways varying among distinct organs (Jiao et al., 2007). The plastid signals that affect hypocotyl elongation may originate from plastids in the hypocotyls or in another organ, such as the cotyledon.

Lending support to this idea, Tanaka et al. (2002) reported that photoreceptors in cotyledons affect processes in hypocotyls.

The suppression of the long hypocotyl phenotype of hy5 in gun1 hy5 is consistent with two models. (1) Hypocotyls are driven to elongate and hypocotyl elongation is inhibited by a light signaling network regardless of the functional state of the plastid. In this model, GUN1-dependent plastid signaling, which is triggered by both inhibitors of chloroplast biogenesis and high-fluence-rate white light, stimulate hypocotyl

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elongation not only in hy5 but also in wild type. In wild type, HY5 would dominate the system, thereby obscuring the stimulatory effect of GUN1 and resulting in an overall reduced rate of hypocotyl elongation. (2) During inhibitor and high light treatments, gun1 hy5 mutants may experience greater plastid dysfunction than hy5, to the extent of being unable to support hypocotyl elongation, for example because of abnormal metabolism. Although the second model is difficult to completely rule out, the first model is more consistent with the available data. For instance, we observed that cry1, phyB, and hy5 exhibit long hypocotyl phenotypes regardless of whether they are treated with inhibitors of chloroplast biogenesis that induce severe plastid dysfunction. In fact, the hypocotyls of cry1 and hy5 treated with lincomycin and erythromycin were longer than untreated controls in white light. Also, alleles of hy1 and hy2, which are deficient in phytochrome signaling, also exhibit elongated hypocotyls when seedlings are photobleached with norflurazon (Mochizuki et al., 2001). Therefore, hypocotyls are not necessarily less driven to elongate when plastids experience dysfunction. Our finding that in the dark, the hypocotyl lengths are the same in wild type and all mutants regardless of whether seedlings are treated with lincomycin indicates that these effects of GUN1-dependent plastid signals on hypocotyl length depend on light.

Connections between plastid function and hypocotyl elongation have been previously reported. For example, phytochrome interacting factor 3 (PIF3) promotes both chloroplast biogenesis (Monte et al., 2004) and hypocotyl elongation in continuous red light (Bauer et al., 2003; Kim et al., 2003; Monte et al., 2004). Additionally, hypocotyl lengths are reduced in the chlorophyll-deficient cue mutants (Vinti et al., 2005) but enhanced in laf6 (Møller et al., 2001). The suppression of the long hypocotyl

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phenotype of hy5 in gun1 hy5 and these other reports are consistent with chloroplast biogenesis and hypocotyl elongation interacting by a complex mechanism. Although a mechanism is not clear at present, we suggest that the current data are consistent with

GUN1-dependent plastid signals stimulating hypocotyl elongation by (1) inhibiting a factor (Y) that represses hypocotyl elongation and acts downstream of cry1 and in a separate pathway from HY5 (Figure 3.9C) or by (2) acting downstream of HY5 and stimulating a factor (Z) that induces hypocotyl elongation and is repressed by HY5

(Figure 3.9D).

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MATERIALS AND METHODS

Plant material and growth conditions

Plant material and all growth conditions (i.e., growth media, temperature, and light sources) were as described in Ruckle et al. (2007). Briefly, seedlings were grown on Linsmeier and Skoog media containing 2% sucrose and either 0.5 mM lincomycin,

0.5 mM erythromycin, or no inhibitor of chloroplast biogenesis, as described by Ruckle et al. (2007). For each experiment, seeds were stratified, germination was promoted by irradiating seeds with red light, seedlings were grown in the dark for 23 h as described by Ruckle et al. (2007), and seedlings were then grown for six days in the indicated conditions.

Analysis of cotyledon opening and expansion

To quantitate cotyledon opening, we placed seedlings in a horizontal position on moist sheets of BioDesignGelWrap (BioDesign Inc. of New York, Carmel NY) alongside a ruler and imaged them using a flatbed scanner at a resolution of 300 points per inch.

Angles between the cotyledons were quantitated using ImageJ (National Institutes of

Health) as recommended by Fankhauser & Casal (2004). For these measurements and all other measurements, statistical significance (P<0.05) was tested by either one or two-way ANOVA as indicated. For anthocyanin measurements, log transformation of the data was necessary to meet the assumptions of the statistical test.

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To quantitate cotyledon area, we excised cotyledons from each seedling and placed them on moist sheets of BioDesignGelWrap, imaged them, and quantitated them as described for cotyledon opening. Fixed and flattened cotyledons were prepared to account for the downward curling of the cotyledons observed in lincomycin-treated wild

-2 -1 type in 50 µmol m s blue light. Separate groups of seedlings were fixed in an ethanol:acetic acid (3:1) solution overnight as recommended by Kakiuchi et al. (2007) that also contained 0.01% Coomassie Brilliant Blue R-250 to facilitate the subsequent imaging of the cotyledons. The fixed cotyledons were subsequently washed once with ethanol:acetic acid (3:1) to remove excess stain. The fixed and stained cotyledons were flattened as recommended by Neff & Chory (1998). Briefly, the cotyledons were placed on the sticky side of transparent tape and flattened with forceps. A dry sheet of

BioDesignGelWrap was placed on top of these cotyledons, which were then scanned as described for cotyledon opening. We quantitated cotyledon areas using ImageJ, as recommended by Fankhauser & Casal (2004). We calculated the amount of curling by dividing the flattened cotyledon areas by the unflattened areas.

Scanning electron microscopy (SEM) and cross sections

For SEM, samples were fixed in 2.5% glutaraldehyde/2.5% paraformaldehyde with a 0.1 M cacodylate buffer, pH 7.4, and dehydrated. Critical-point-dried samples were osmium coated and examined using the 6400 JEOL scanning electron microscope with accelerating voltage of 10 kV. For cross sections, tissue was fixed as described for

SEM, postfixed in 1% osmium tetroxide in 0.1 M cacodylate buffer, dehydrated in a

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graded acetone series, and infiltrated and embedded in Poly/Bed 812 resin. Sections of

1 µm were cut with a Power Tome XL ultramicrotome (RMC, Boeckeler Instruments,

Tucson AZ) and stained with Epoxy Tissue Stain (Electron Microscope Sciences,

Hatfield, PA). Cross sections were visualized with an Axio Imager M1 microscope (Carl

Zeiss Inc.).

Analysis of SEMs and cross sections

We measured cell areas from the SEM images using the free-hand tool of

ImageJ, as recommended by Djakovic et al. (2006). The interdigitating lobes of the epidermal pavement cells were quantitated as recommended by Djakovic et al. (2006).

For this calculation, we measured cell perimeter and area using ImageJ and, with these values, calculated a form factor, as recommended by Russ (2002). The form factor is

2 HTXDOWR ʌ FHOODUHD FHOOSHULPHWHU ; it describes the amount of convolution at the cell periphery. For example, a circle has a form factor of 1 and the form factor becomes smaller than 1 as convolution, such as interdigitating lobes, increases. We calculated the density of stomata by counting the number of stomata in a micrograph and dividing

2 by the area of each micrograph (120,000 µm ), as recommended by Berger & Altmann

(2000). To quantitate the wrinkling of the cotyledon surface, we calculated a cotyledon surface factor, which is the length of the abaxial cotyledon surface divided by the diameter of the cotyledon. These values were determined using ImageJ.

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Analysis of anthocyanin levels and hypocotyl lengths

Anthocyanin levels were quantitated exactly as recommended by Fankhauser &

Casal (2004). We measured hypocotyl lengths by placing seedlings onto moist

BioDesignGelWrap. These seedlings were imaged and hypocotyl lengths were quantitated as described for cotyledon opening.

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ACKNOWLEDGEMENTS

We thank Andrea Stavoe, Chris Sinkler, and Stephanie DeMarco for helpful assistance. We thank Ewa Danielewicz and Alicia Pastor at the Center for Advanced

Microscopy (Michigan State University) for technical assistance with SEM and light microscopy experiments, Joanne Chory for providing gun1-1 seeds, and Beronda

Montgomery-Kaguri and Lyle Burgoon for helpful discussions. This work was supported by DOE grant no. DE±FG02±91ER20021 and NSF grant no. IOB 0517841 to RML.

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Figure 3.1 Cotyledon opening in gun1-1 and wild-type seedlings in different fluence rates of blue light. Col-0 (squares) and gun1-1 mutants (circles) were grown in the presence (filled shapes) or absence (open shapes) of lincomycin in the indicated fluence rates of blue light. For each data point, n > 35. Error bars represent 95% confidence intervals. Statistical significance (P<0.05) was tested with a two-way ANOVA. * indicates a significant difference between lincomycin-treated gun1-1 and lincomycin-treated wild type. ** indicates a significant difference between lincomycin-treated and untreated wild type.

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Figure 3.2 Flattened cotyledon areas in gun1-1 and wild-type seedlings. Seedlings were grown as described in Figure 3.1. Cotyledons were fixed and flattened as described in Experimental Procedures. For each data point, n > 29. Error bars and statistical analyses are as described in Figure.3.1. * and ** are as defined in Figure 3.1.

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Figure 3.3 Cotyledon areas in gun1, cry1, and hy5 mutants. A) Quantitation of cotyledon area measurements in gun1, cry1, and hy5 mutants. The indicated lines were grown on media containing lincomycin (+Lin) or -2 -1 erythromycin (+Ert), or containing no inhibitor (No Inhibitor) in 50 µmol m s blue light. Cotyledons were fixed and flattened, and cotyledon areas were measured as described in Experimental Procedures. For each line in each condition, n > 33. Error bars represent 95% confidence intervals. Statistical significance (P<0.05) was tested with a one-way ANOVA. * indicates a significant difference between single mutants and wild type or a significant difference between double mutants and single mutants grown in the same conditions. B) Representative seedlings and cotyledons were grown as described in (A). Bar = 2 mm.

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Figure 3.4 Analysis of cotyledon expansion in gun1-1 and phyB. A) Cotyledon area measurements in gun1-1 and phyB mutants. Seedlings were grown on media containing lincomycin (+Lin) or no inhibitor of chloroplast biogenesis -2 -1 (No Inhibitor) in 100 µmol m s red light. For each line in each condition, n = 36. Error bars represent 95% confidence intervals. Statistical analyses were as in Figure 3.3. * is defined as in Figure 3.3. B-C) Representative seedlings and cotyledons were grown as described in (A) on media containing lincomycin (B) or no inhibitor of chloroplast biogenesis (C). Bar = 2 mm.

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Figure 3.5 Analysis of epidermal cells in the cotyledons of gun1 and cry1 mutants. A) Analysis of the epidermis from the untreated cotyledons of wild type (Col-0), gun1, and cry1 mutants. The indicated lines were grown on media that did not -2 -1 contain an inhibitor of chloroplast biogenesis in 50 µmol m s blue light. The epidermis of the cotyledons was analyzed by SEM. Micrographs were obtained from the center of 10-15 cotyledons for each line. Representative images are shown. B) Analysis of the epidermis from the cotyledons of lincomycin-treated wild type (Col-0), gun1, and cry1 mutants. The indicated lines were grown on media that contained lincomycin in the same light conditions described in (A). The epidermis was analyzed as in (A). Representative micrographs are shown. Bar = 100 mm.

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Figure 3.6 Analysis of anthocyanin levels in lincomycin-treated seedlings. A) Analysis of anthocyanin levels in lincomycin-treated and untreated gun1 and cry1 -2 -1 mutants in blue light. Seedlings were grown in 50 µmol m s blue light in the presence of lincomycin (+Lin) or the absence of an inhibitor of chloroplast biogenesis (No Inhibitor). For each line in each condition, n> 4 and there were twenty seedlings per replicate. Error bars represent 95% confidence intervals. B) Analysis of anthocyanin levels in lincomycin-treated and untreated gun1 and hy5 mutants in blue light. Seedlings were grown and analyzed as in (A). Error bars are as described in (A). Statistical analyses were as described in Figure 3.3. * is defined as in Figure 3.3.

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Figure 3.7 Analysis of anathocyanin levels during de-etiolation in bright white light. A) Analysis of anthocyanin levels in gun1 and cry1 mutants during de-etiolation in -2 -1 bright white light. Seedlings were grown in continuous 1000 µmol m s white light with no inhibitor of chloroplast biogenesis. For each line in each condition, n> 4 and there were twenty seedlings per replicate. Error bars represent 95% confidence intervals. B) Analysis of anthocyanin levels in gun1 and hy5 mutants during de-etiolation in continuous bright white light. Seedlings were grown at a different time than the seedlings in (A) and were analyzed as described in (A). Error bars are as described in (A). Statistical analyses were as described in Figure 3.3. * is defined as in Figure 3.3.

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Figure 3.8 Analysis of hypocotyl lengths in gun1, cry1, and hy5 mutants. A) Analysis of hypocotyl length in gun1, cry1, and hy5 mutants grown in blue light. -2 -1 For each line, seedlings were grown in the dark or in 50 µmol m s blue light in the presence of lincomycin (+Lin) or erythromycin (+Ert), or in the absence of an inhibitor of chloroplast biogenesis (No Inhibitor). For each line in each condition, n > 26. For all lines, hypocotyl lengths in a particular line in a particular condition are presented as a percent of the average hypocotyl length of that line grown on the same media in the dark. Error bars represent 95% confidence intervals. B) Analysis of hypocotyl length in gun1, cry1, and hy5 mutants grown in white light. -2 -1 For each line, seedlings were grown in the dark or in 125 µmol m s white light in the presence of lincomycin (+Lin) or erythromycin (+Ert), or in the absence of an inhibitor of chloroplast biogenesis (No Inhibitor). For each line in each condition, n > 27. Analysis of hypocotyl lengths and error bars are as described in (A). C) Analysis of hypocotyl length in gun1and hy5 mutants grown in high-fluence-rate -2 -1 white light. For each line, seedlings were grown in the dark or in 1000 µmol m s white light with no inhibitor of chloroplast biogenesis. For each line, n > 22. Analysis of hypocotyl lengths and error bars are as described in (A). Statistical analyses were as in Figure 3.3. * is defined as in Figure 3.3.

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Figure 3.9 Models for the regulation of photomorphogenesis by plastid signals. A) Model for the regulation of cotyledon expansion by plastid signals. A plastid signal that requires GUN1 for its biosynthesis or transduction (X) (dotted arrows) exits the plastid and triggers an extraplastidic signaling pathway that ultimately represses PhANG expression (Koussevitzky et al., 2007). GUN1-dependent plastid signals also repress photomorphogenesis of the cotyledons. A plastid signal that is genetically distinct from the GUN1-dependent plastid signals (W) converts cry1 from a positive to a negative regulator of cotyledon expansion in high-fluence-rate blue light, as discussed in the text. The same signal may be involved in blocking pavement cell differentiation in the cotyledons. B) Model for the regulation of anthocyanin biosynthesis by plastid signals. GUN1 and cry1 protect plastids from stress induced by high-intensity light (HL) during de- etiolation (Ruckle et al., 2007). Photoactivated cry1 induces anthocyanin biosynthesis (Ahmad et al., 1995). GUN1, cry1, and HY5 induce anthocyanin biosynthesis, but the GUN1 pathway only appears to be an important regulator of anthocyanin biosynthesis when chloroplasts are stressed or damaged by HL or inhibitors of chloroplast biogenesis (Inhibitors), as discussed in the text. HY5 is not shown but probably acts downstream of cry1 (Jiao et al., 2007). C-D) Models for the regulation of hypocotyl elongation by GUN1-dependent plastid signals during chloroplast dysfunction. (C) Repression of an inhibitor of hypocoytyl elongation by GUN1-dependent plastid signals. In this model, Y is a factor that inhibits hypocotyl elongation by a mechanism that does not involve HY5. When chloroplasts experience dysfunction, GUN1-dependent plastid signals inhibit Y, and cry1 induces Y. (D) Stimulation of an inducer of hypocotyl elongation by GUN1- dependent plastid signals. In this model, Z is a factor that induces hypocotyl elongation and acts downstream of HY5. When chloroplasts experience dysfunction, GUN1-dependent plastid signals stimulate factor Z.

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Figure 3.10 Representative seedlings and cotyledons from the fluence response experiments. Seedlings were grown as described for Figure 3.1 and 3.2 at the indicated fluence rates of blue light. Bar = 2 mm.

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Figure 3.11 Analysis of cotyledon opening in seedlings grown on media containing lincomycin or erythromycin. Seedlings were grown in the presence of erythromycin (+Ert) or lincomycin (+Lin), or -2 - in the absence of an inhibitor of chloroplast biogenesis (No inhibitor) in 1 µmol m s 1 blue light. For each line in each condition, n > 16. Error bars represent 95% confidence intervals. Statistical significance (P<0.05) was tested as in Figure 3.3. * is defined as in Figure 3.3.

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Figure 3.12 Cotyledon expansion in treated and untreated gun1-1 and wild-type seedlings. A) Cotyledon areas in gun1-1 and wild-type seedlings in various fluence rates of blue light. Seedlings were grown as described in Figure 3.1. For each data point, n > 38. Error bars represent 95% confidence intervals and are smaller than the squares or circles for some data points. B) Quantitation of cotyledon curling in various fluence rates of blue light. The amount of curling in the cotyledons was calculated by dividing the flattened cotyledon areas from Figure 3.2 by the unflattened cotyledon areas from (A). Ratios that are greater than 1 indicate curling. Statistical significance (P<0.05) was tested as in Figure 3.1. * and ** is defined as in Figure 3.1.

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Figure 3.13 Analysis of cotyledon expansion in gun1 mutants grown in blue light. -2 -1 A) Cotyledon area measurements in 1 µmol m s blue light. The indicated lines were grown on media containing erythromycin (+Ert), lincomycin (+Lin), or no inhibitor of chloroplast biogenesis (No Inhibitor). Cotyledons were fixed and flattened as described in Experimental Procedures. For each line in each condition, n > 34. Error bars represent 95% confidence intervals. -2 -1 B) Cotyledon area measurements in 2.5 µmol m s blue light. The indicated lines were grown and analyzed as in (A). Error bars are as in (A). For each line in each condition, n> 32. Statistical significance (P<0.05) was tested as in Figure 3.3. * is defined as in Figure 3.3.

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Figure 3.14 Quantitative analysis of SEMs. A) Quantitation of epidermal pavement cell area. B) Quantitation of epidermal pavement cell shape. C) Density of stomata. Statistical significance (P<0.05) was tested as in Figure 3.3. * is defined as in Figure 3.3.

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Figure 3.15 Representative seedlings from Figure 3.8. A) Representative seedlings grown on media containing lincomycin (+Lin), erythromycin (+Ert), or no inhibitor of chloroplast biogenesis (No Inhibitor) in white light. Growth conditions were as described in Figure 3.8B. Representative seedlings are shown. B) Representative seedlings grown in high-fluence-rate white light. Growth conditions were as described in Figure 3.8C. Representative seedlings are shown. Bar = 2 mm.

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Figure 3.16 Light microscopy of wild type, gun1, cry1, and gun1-1 cry1 cotyledons. A) Representative sections of gun1, cry1, and wild-type (Col-0) cotyledons. Seedlings were grown on media containing lincomycin as described in Figure 3.5. Representative sections of cotyledons are shown. Bar = 100 mm.

B) Quantitative analysis of cotyledon surface area. The cotyledon surface factor was calculated as described in Experimental Procedures. For each line, n = 4. Statistical significance (P<0.05) was tested as in Figure 3.3. * is defined as in Figure 3.3.

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Vandenbussche F, Habricot Y, Condiff AS, Maldiney R, Van der Straeten D, Ahmad M. (2007). HY5 is a point of convergence between cryptochrome and cytokinin signalling pathways in Arabidopsis thaliana. Plant J. 49: 428-441. Vinti G, Fourrier N, Bowyer JR, López-Juez E. (2005). Arabidopsis cue mutants with defective plastids are impaired primarily in the photocontrol of expression of photosynthesis-associated nuclear genes. Plant Mol. Biol. 57: 343-357. Wanzel M, Herold S, Eilers M. (2003). Transcriptional repression by Myc. Trends Cell Biol. 13:146-150 Xu XM, Adams S, Chua NH, Møller SG. (2005). AtNAP1 represents an atypical SufB protein in Arabidopsis plastids. J Biol Chem. 280: 6648-6654. Yu F, Fu A, Aluru M, Park S, Xu Y, Liu H, Liu X, Foudree A, Nambogga M, Rodermel S. (2007). Variegation mutants and mechanisms of chloroplast biogenesis. Plant Cell Environ. 30: 350-365.

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CHAPTER 4

PLASTIDS ARE MAJOR REGULATORS OF LIGHT SIGNALING IN ARABIDOPSIS

Michael E. Ruckle, Lyle D. Burgoon, Lauren Lawerence, Christopher Sinkler, Robert M. Larkin. (2010) Characterization of transcriptomes and signaling factors that contribute to chloroplast biogenesis in Arabidopsis.Submitted

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PLASTIDS ARE MAJOR REGULATORS OF LIGHT SIGNALING IN ARABIDOPSIS

ABSTRACT

Plastids emit signals that broadly affect cellular processes. Based on previous genetic analyses, we proposed that plastid signaling regulates the downstream components of a light signaling network and that this signal integration coordinates chloroplast biogenesis with both the light environment and development by regulating gene expression. We tested these ideas by analyzing light-regulated and plastid- regulated transcriptomes and by performing a reverse genetic analysis of particular light- and plastid-regulated genes. We found that the chloroplast dysfunction is a major regulator of light signaling, attenuating the expression of more than half of all light- regulated genes in our dataset and changing the nature of light regulation for a smaller fraction of these light-regulated genes. Our transcriptome analyses are consistent with these interactions promoting the biogenesis and function of chloroplasts and helping coordinate diverse light-regulated processes such as growth, circadian clock activity, and stress responses with chloroplast function. Consistent with our transcriptome analyses, our reverse genetic screen yielded eighteen genes that attenuate chloroplast biogenesis during de-etiolation and contribute to signaling, transcription, or no known function. Our findings provide evidence that light and plastid signaling are interactive processes that broadly influence diverse cellular processes by regulating a complex network of signaling factors.

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INTRODUCTION

In plants, chloroplasts are derived from nonphotosynthetic proplastids during the development of photosynthetic organs such as cotyledons and leaves (Thomson and

Whately, 1980; Saito et al., 1989; Mullet, 1993; Asokanthan et al., 1997; Vothknecht and Westhoff, 2001). Regulated gene expression plays a major role in chloroplast biogenesis and function and is complex on at least two levels: (1) chloroplast biogenesis requires the coordinated expression of both the nuclear and chloroplast genomes and

(2) the gene expression that drives chloroplast biogenesis is regulated by a number of cues²both environmental and endogenous. Light is a major driver of chloroplast biogenesis, not only because light is a major regulator of chloroplast-related gene expression but also because a light-dependent enzyme is required for chlorophyll biosynthesis (Masuda and Fujita, 2008; Waters and Langdale, 2009). In addition to light, endogenous cues such as the circadian clock, hormones, and carbohydrates are important regulators of photosynthesis-related gene expression (Rook et al., 2006; Jiao et al., 2007). These extraplastidic cues constitute the anterograde control of chloroplast biogenesis. Anterograde control is not the sole regulator of chloroplast biogenesis and function; the chloroplast emits signals that can have a major effect on the expression of nuclear genes. This retrograde plastid-to-nucleus signaling helps coordinate nuclear gene expression with the functional state of the chloroplast. This bidirectional exchange of information between the nucleus and the plastid (i.e., anterograde control and retrograde signaling) is thought to help coordinate the expression of the nuclear and chloroplast genomes and promote chloroplast biogenesis and function (Woodson and

Chory, 2008). Such bidirectional communication that promotes homeostasis in various

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conditions is well established between the mitochondria and the nucleus and between the endoplasmic reticulum (ER) and the nucleus (Liu and Butow, 2006; Ron and Walter,

2007).

Light and plastid signals are major regulators of chloroplast biogenesis. Light regulates ca. 20% of the transcriptome in Arabidopsis and rice. A number of photoreceptors and downstream signaling components are known to drive light- regulated transcriptional networks. In photosynthesis-related nuclear genes, light- regulated transcription depends on a combination of at least two distinct promoter elements (Tyagi and Gaur, 2003; Jiao et al., 2007). Light induces the expression of well-studied photosynthesis-related genes, such as the genes that encode the light- harvesting chlorophyll a/b-binding protein of photosystem II (Lhcb or CAB, hereafter referred to as Lhcb) and the Rubisco small subunit (RbcS), by regulating the phytochrome and cryptochrome photoreceptors (Gao and Kaufman, 1994; Reed et al.,

1994; Folta and Kaufman, 1999; Mazzella et al., 2001; Martinez-Hernandez et al., 2002).

These photoreceptors transduce far-red, red, and blue light signals (Jiao et al., 2007).

Light signaling mechanisms include the regulation of activity, subcellular localization, and concentration of particular photoreceptors and downstream signaling components

(Jiao et al., 2007; Chory, 2010).

Plastid signals that regulate nuclear gene expression help coordinate the expression of photosynthesis-related genes and stress-related nuclear genes with plastid function (Larkin and Ruckle, 2008; Woodson and Chory, 2008; Galvez-

Valdivieso and Mullineaux, 2010; Lemeille and Rochaix, 2010), help coordinate the expression of nuclear and plastid genomes (Susek et al., 1993), contribute to efficient

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chloroplast biogenesis (Mochizuki et al., 1996; Ruckle et al., 2007), and affect development (Ruckle and Larkin, 2009; Cottage et al., 2010). Thus, plastid signals that regulate nuclear gene expression are thought to contribute significantly to the development of photosynthetic organs. Additionally, because the various biotic and abiotic stresses that plants experience in nature can cause chloroplast dysfunction and the down-regulation of photosynthesis-related gene expression (Takahashi and Murata,

2008; Saibo et al., 2009; Bilgin et al., 2010), plastid signaling might contribute to biotic and abiotic stress responses. Consistent with this idea, the plastid-to-nucleus signaling mutant genomes uncoupled 1 (gun1) exhibits reduced sensitivity to intense-light stress

(Koussevitzky et al., 2007), many type III effectors of Pseudomonas syringae are predicted to reside in chloroplasts (Guttman et al., 2002), and chloroplast-localized proteins contribute to biotic stress responses (Seo et al., 2000; Slaymaker et al., 2002;

Jelenska et al., 2007; Caplan et al., 2008; Wangdi et al., 2010). Plastid signaling is connected to tetrapyrrole metabolism, plastid genome expression, reactive oxygen species production, and photosynthetic electron transport, but the molecular nature of signals derived from these processes and many of the proteins that are required for plastid signal biosynthesis and transduction remain unknown. Also, the full impact of plastid-to-nucleus signaling is not clear because we cannot completely knock out plastid signaling at this time (Larkin and Ruckle, 2008; Woodson and Chory, 2008; Galvez-

Valdivieso and Mullineaux, 2010; Lemeille and Rochaix, 2010).

Some of the first proteins that contribute to broadly significant mitochondrial-to- nucleus and ER-to-nucleus signaling pathways were discovered using reporter-gene- based forward-genetic mutant screens in yeasts that were experiencing severe

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mitochondrial or ER dysfunction (Liu and Butow, 2006; Ron and Walter, 2007). These screens were performed with ı° yeast that lack mitochondrial DNA and therefore do not respire (Liao and Butow, 1993; Jia et al., 1997), or with yeast having ER dysfunction because of a mutation or an inhibitor treatment that blocks the normal glycosylation of proteins that occurs in the ER (Cox et al., 1993; Mori et al., 1993). A rationale for these approaches is that when organelles malfunction, retrograde signaling is a major regulator of genes that are regulated by retrograde signaling. Therefore, retrograde signaling phenotypes are more robust and are more easily scored with reporter genes when mitochondria and the ER show severe dysfunction. A similar approach was developed to study plastid-to-nucleus signaling in Arabidopsis. Reporter-gene-based forward genetic screens were developed using Arabidopsis seedlings treated with inhibitors of chloroplast biogenesis (Susek et al., 1993; Koussevitzky et al., 2007;

Ruckle et al., 2007; Cottage et al., 2008). Plastid-to-nucleus signaling screens were also developed using the Arabidopsis flu allele that causes severe chloroplastic reactive oxygen species stress (ROS) derived from an overaccumulation of protochlorophyllide when grown in photoperiodic light. This ROS stress promotes photobleaching in flu seedlings and both stunts growth and promotes lesion formation in leaves of mature flu plants (Meskauskiene et al., 2001; op den Camp et al., 2003). Mutants with defects in plastid-to-nucleus signaling triggered by the redox environment of the plastid were also reported (Heiber et al., 2007). Based on analyses of the mutants yielded by these screens, a complex network of processes would appear to participate in plastid-to- nucleus signaling (Mochizuki et al., 2001; Larkin et al., 2003; Wagner et al., 2004;

Koussevitzky et al., 2007; Lee et al., 2007; Ruckle et al., 2007; Przybyla et al., 2008;

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Baruah et al., 2009; Coll et al., 2009; Kim et al., 2009; Meskauskiene et al., 2009).

Although these findings have provided insight into plastid-to-nucleus signaling, major gaps remain in our understanding of this process.

Results from experiments that rely on inhibitors of chloroplast biogenesis and the flu mutant indicate that plastid signals can remodel light signaling (Danon et al., 2006;

Ruckle et al., 2007). These findings are intriguing because light signals were previously thought to drive only the anterograde control of chloroplast biogenesis and to function independently from the plastid signals that attenuate chloroplast biogenesis and

IXQFWLRQ7KHSODVWLGFDQ³UHZLUH´WKHOLJKWVLJQDOLQJWKDWUHJXODWHVLhcb1 expression largely by converting at least one downstream signaling component, the bZIP transcription factor long hypocotyl 5 (HY5) that acts downstream of cryptochrome 1

(cry1) and other photoreceptors, from a positive regulator of Lhcb1 expression in seedlings that contain well-functioning chloroplasts to a negative regulator of Lhcb1 expression in seedlings that contain dysfunctional chloroplasts. These interactions appear important for efficient chloroplast biogenesis (Ruckle et al., 2007). Based on these findings, interactions between light and plastid signaling were proposed to help balance the many processes required for optimal chloroplast biogenesis (Ruckle et al.,

2007; Larkin and Ruckle, 2008). Chloroplast dysfunction cDQUDSLGO\³UHZLUH´OLJKW signaling. Experiments with the conditional flu mutant indicate that chloroplastic ROS stress rapidly converts cry1 signaling from a process that promotes chloroplast biogenesis and function to a process that promotes albinism and programmed cell death (Danon et al., 2006). Additionally, Ruckle and Larkin (2009) provided evidence that light and plastid signals can help coordinate chloroplast biogenesis and light-

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regulated development. Thus, interactions between light signaling and the plastid signaling triggered by inhibitors of chloroplast biogenesis appear significant and complex. To test predictions from these previous reports, we analyzed light-regulated transcriptomes and plastid-regulated transcriptomes. We also performed reverse genetic analyses of particular genes that were defined by these transcriptome analyses and that are annotated as encoding signaling factors, transcription factors, or proteins of no known function. Our findings indicate that plastid signaling is a major regulator of light signaling and provide evidence that the regulation of downstream signaling factors contributes to these interactions.

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RESULTS

Although light signaling induces the expression of photosynthesis-related nuclear genes such as the Lhcb1 and RbcS genes in seedlings that contain well-functioning chloroplasts (Tyagi and Gaur, 2003; Jiao et al., 2007), this same light signaling network contributes to the repression of these genes when seedlings are treated with inhibitors of chloroplast biogenesis (Ruckle et al., 2007). Increasing fluence rates of white light induces the expression of Lhcb1 and RbcS in seedlings containing well-functioning chloroplasts, but these same fluence rates of white light inhibit the expression of Lhcb1 and attenuate the light-induced expression of RbcS in seedlings treated with lincomycin

(Ruckle et al., 2007). Lincomycin is an antibiotic that functions as a light-independent inhibitor of chloroplast biogenesis by inhibiting the activity of plastid ribosomes (Sullivan and Gray, 1999; Mulo et al., 2003; Ruckle et al., 2007). Blue and red light appear mostly if not entirely responsible for this repressive effect of white light (Ruckle et al.,

2007).

To further study these interactions between light and plastid signaling, we grew

Arabidopsis thaliana ecotype Colombia-0 (Col-0) seedlings in 40% blue and 60% red

(BR) light in either the presence or the absence of lincomycin as previously described

-2 -1 (Ruckle et al., 2007). After 6 d of growth in 0.5 µmol m s of BR light, we transferred

-2 -1 these seedlings to 60 µmol m s BR light. We observed a twofold increase in the levels of mRNA transcribed from Lhcb1 and an eleven-fold increase in the levels of mRNA transcribed from RbcS in untreated seedlings at 24 h after this fluence-rate shift

(Figure 4.1A and B, respectively), which is consistent with previous reports (Gao and

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Kaufman, 1994; Reed et al., 1994; Terzaghi and Cashmore, 1995; Mazzella et al., 2001;

Martinez-Hernandez et al., 2002). We also observed a twofold and a sixfold decrease in Lhcb1 mRNA levels at 4 h and at 24 h after this fluence-rate shift in lincomycin- treated seedlings (Figure 4.1A). In contrast, RbcS expression is induced following this fluence-rate shift in lincomycin-treated seedlings, even though RbcS expression is attenuated in lincomycin-treated relative to untreated seedlings (Figure 4.1B). These data are consistent with chloroplast dysfunction down-regulating the expression of photosynthesis-related genes and are consistent with previous work (Ruckle et al.,

2007). These data also indicate that the inhibition of Lhcb1 expression by light in lincomycin-treated seedlings is a rapid response. RNA blot hybridization analysis indicates that these RNA preparations are of high integrity (Figure 4.8A and B).

Based on this analysis of Lhcb1 and RbcS expression, we conclude that this 0.5

-2 -1 to 60 µmol m s BR-fluence-rate-shift procedure is useful for studying the regulation of transcriptome responses by light and plastid signals that affect photosynthesis- related gene expression. Previous analyses of light-regulated transcriptomes did not test for effects of the plastid on light signaling (Ma et al., 2003; Tepperman et al., 2004;

Tepperman et al., 2006; Jiao et al., 2007; Leivar et al., 2009; Shin et al., 2009).

Previous analyses of plastid-regulated transcriptomes did not test for the effects of light signaling (Strand et al., 2003; Koussevitzky et al., 2007; Aluru et al., 2009). Previous analyses of transcriptomes regulated by the plastidic ROS that rewires cry1 signaling to induce programmed cell death focused on rapid responses that did not include photosynthesis-related genes (op den Camp et al., 2003; Danon et al., 2006).

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Analysis of light- and plastid-regulated transcriptomes

To test the extent to which plastid signals remodel the light-regulated transcriptome, we analyzed transcriptomes in lincomycin-treated and untreated

-2 -1 seedlings before (0 h) and 0.5 h, 1 h, 4 h, and 24 h after a 0.5 to 60 µmol m s BR- fluence-rate shift, as described above, using the Affymetrix GeneChip ATH1. We found that the fluence-rate shift significantly changes the expression of 6424 genes by twofold or more relative to the 0 h control (Figure 4.2A). By comparing transcriptomes in lincomycin-treated and untreated seedlings we found that nearly half of these light- regulated genes were also significantly regulated by the plastid as judged by a lincomycin treatment (Figure 4.2A). Only 680 genes were significantly regulated by only the lincomycin treatment and were not significantly regulated by the fluence-rate shift

(Figure 4.2A).

Consistent with previous reports (Jiao et al., 2007), we found that the number of light-regulated genes only increases with time after the BR-fluence-rate shift (Figure

4.2B). Light regulates fewer genes in lincomycin-treated seedlings compared to untreated seedlings at each time point following the fluence-rate shift. This reduction in genes significantly regulated by light in lincomycin-treated seedlings was apparent for both genes that are significantly regulated only by the fluence-rate shift (Figure 4.2B, orange bars) and genes that are significantly regulated by both the fluence-rate shift and the lincomycin treatment (Figure 4.2B, purple bars). Two- to threefold more of the genes that are significantly regulated only by the fluence-rate shift are regulated by the

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fluence-rate shift in untreated seedlings than in lincomycin-treated seedlings, and two- to fourfold more genes that are significantly regulated by both the fluence-rate shift and the lincomycin treatment were significantly regulated by the fluence-rate shift in untreated seedlings compared to lincomycin-treated seedlings (Figure 4.2B). Thus, lincomycin treatment abolishes significant light regulation for approximately half of all light-regulated genes in this dataset. Consistent with plastid signaling affecting the composition of the light-regulated transcriptome, trajectory analysis indicates that the transcriptomes of lincomycin-treated seedlings and untreated seedlings are most similar at 0.5 h after the BR-fluence-rate shift and become more different as time increases following the BR-fluence-rate shift (Figure 4.2C). Based on a comparison of the transcriptomes of lincomycin-treated and untreated seedlings, the most striking effect is the reduction in the size of the light-regulated transcriptome in lincomycin-treated seedlings relative to untreated seedlings. This effect is illustrated by lower variance in the first principal component of the variance (PC1) in the treated samples (Figure 4.2C).

Additionally, the trajectory plot shown in Figure 4.2C indicates a large divergence between the PC1 and the second principal component of the variance (PC2) of lincomycin-treated and untreated samples at the later time points. These data provide evidence that the expression differences in lincomycin-treated and untreated seedlings are not transient differences and that the plastid changes a component of the light response.

The interpretations of our trajectory plot (Figure 4.2C) and our analysis of significantly regulated genes (Figure 4.2B) were corroborated by agglomerative hierarchical clustering. Light-regulated expression is more robust in untreated seedlings

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than in lincomycin-treated seedlings for genes in clusters B, E, G, H and I (Figure 4.2E;

Figure 4.9C). These clusters represent 63% of significantly regulated genes. This

HIIHFWRIFKORURSODVWG\VIXQFWLRQ³WXUQLQJRIIWKHOLJKWV´LVPRUHREYLRXVZKHQ transcriptomes were analyzed at 4 h and 24 h than at 0.5 h and 1 h (Figure 4.9A and B).

Agglomerative hierarchical clustering of transcriptomes at 4 h and 24 h following the fluence-rate shift reveals that the expression of genes in clusters Y, CC, DD, EE, and

GG are more regulated by light in untreated seedlings than in lincomycin-treated seedlings (Figure 4.9B and F). Genes in these clusters account for 66% of significantly regulated genes (Figure 4.9B and F). At these same time points, the expression of genes that are significantly regulated by the light in both lincomycin-treated and untreated seedlings are found in clusters Z, AA, and FF (Figure 4.9B and F). These genes account for 33% of significantly regulated genes. By contrast, agglomerative hierarchical clustering of transcriptomes at 0.5 h and 1 h after the BR-fluence-rate shift indicates that only the expression of genes in clusters R, U, and X are regulated by light in both lincomycin-treated and untreated seedlings (Figure 4.9A and E). These genes represent only 8.3% of significantly regulated genes.

A relatively small number of genes in several clusters resemble the Lhcb1 genes in that the nature of their light-regulated expression flips (i.e., light is converted from a positive to negative regulator or vice versa) depending on whether the chloroplast is functional or dysfunctional. The expression of genes in clusters G (Figure 4.2E; Figure

4.9C) and DD (Figure 4.9B and F) is like the expression of the Lhcb1 genes in that their expression is mostly repressed by the light in lincomycin-treated and induced by light in untreated seedlings. The expression of genes in cluster D (Figure 4.2E; Figure 4.9C)

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and BB (Figure 4.9B and F) are mostly induced by light in lincomycin-treated but repressed by light in untreated seedlings.

In contrast to the genes significantly regulated by light (i.e., genes expressed at significantly different levels at 0.5, 1, 4, or 24 h following the fluence-rate shift relative to

0 h), the number of genes that are significantly regulated by the plastid (i.e., genes expressed at significantly different levels in lincomycin-treated relative to untreated seedlings at a particular time point) does not only increase as time increases. Among genes whose expression is significantly regulated by both lincomycin and the BR- fluence-rate shift, the expression of two- to threefold more genes is significantly regulated by the lincomycin treatment at 0 h and 24 h than at 4 h following the BR- fluence-rate shift (Figure 4.2B, blue bars). Among genes whose expression is significantly regulated only by lincomycin and not by the BR-fluence-rate shift, the expression of twofold more of these genes is regulated by the lincomycin treatment at 0 h and at 24 h than at 4 h (Figure 4.2B, green bars). One interpretation of these data is that light has a major influence on the composition of the plastid-regulated transcriptome. Indeed, trajectory analysis and agglomerative hierarchical clustering are consistent with (1) light remodeling the plastid-regulated transcriptome at 0 h to a radically different transcriptome at 24 h, and (2) the 4 h time point representing an intermediate state between the 0 h and 24 h transcriptomes (Figure 4.2D and F). The relatively large distance between the three time points on a trajectory plot (Figure 4.2D) indicates that light has a significant impact on the composition of the plastid-regulated transcriptome.

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Analysis of these same data by agglomerative hierarchical clustering provides additional evidence that light signaling has a major influence on the composition of the plastid-regulated transcriptome. Light can convert the plastid from a positive to a negative regulator of the transcriptome or vice versa. Such a conversion is apparent for the genes in cluster K, L, M, N, and O (Figure 4.2F; Figure 4.9D). Other categories of plastid regulation that support this interpretation are exemplified by cluster Q (Figure

4.2F; Figure 4.9D). Genes in this category are repressed in lincomycin-treated

-2 -1 seedlings grown in 0.5 µmol m s BR light and are more severely repressed in

-2 -1 lincomycin-treated seedlings after they are transferred to 60 µmol m s BR light.

Genes in cluster Q represent only 5.2% of significantly regulated genes. Cluster P contains genes that are significantly regulated by the plastid but not significantly regulated by light. These genes are repressed in lincomycin-treated seedlings

-2 -1 regardless of whether seedlings are grown in 0.5 µmol m s BR light or transferred to

-2 -1 60 µmol m s BR light, cluster P genes represent only 2.7% of significantly regulated genes (Figure 4.2F; Figure 4.9D).

We also manually categorized expression patterns to focus on particular patterns of interest, such as those in which the nature of light-regulated gene expression is different in lincomycin-treated and untreated seedlings. Among user-defined expression patterns that exhibit regulation by both the BR-fluence-rate shift and lincomycin, we found that the most common expression pattern is the absence of significant regulation by the BR-fluence-rate shift in lincomycin-treated seedlings (i.e., patterns 32 and 33;

Figure 4.10A and B), which is consistent with the results from agglomerative

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hierarchical clustering. We found that a relatively small group of greater than 100 genes are like the Lhcb1 genes in that they are differently regulated by light in lincomycin- treated and untreated seedlings (i.e., patterns 26 and 27; Figure 4.10A and B). This manual clustering further indicates that light can convert the plastid from a positive to a negative regulator or vice versa for approximately 200 genes (i.e., patterns 28 and 29;

Figure 4.10A), which is also consistent with the results from agglomerative hierarchical clustering.

Enrichment of Gene ontology terms in light- and plastid-regulated transcriptomes

To gain insight into the possible biological significance of the crosstalk between light and plastid signaling, we tested for enrichment of gene ontology (GO) terms in the genes from the expression clusters obtained from agglomerative hierarchical clustering and in the user-defined expression patterns (Figures 4.11 and 4.12). We also performed hierarchical clustering of individual genes from each significantly enriched

GO term. Summaries of this GO analysis and the hierarchical clustering of individual genes associated with these GO terms are presented in Figure 4.3 and 4.4. The expression patterns of individual genes are presented in Figure 4.5A, B, and C. We used this method rather than the commonly used MapMan (Thimm et al., 2004) because MapMan does not allow for the presentation of complex gene expression patterns of a large number of individual genes. Although GO terms representing a number of biological functions were significantly enriched in these expression clusters and user-defined expression patterns (Figure 4.3 and 4.4), genes annotated as

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contributing to photosynthesis exhibit the most significant enrichment in the entire

-29 dataset (p = 7.59 × 10 ) (Figure 4.3). Fourteen additional plastid-related terms

-3 exhibited significant enrichments that range from p = 2.81 × 10 (plastid membrane) to

-20 p = 8.71 × 10 (thylakoid membrane) (Figure 4.3). In general, the expression of the genes associated with these GO terms is induced following the BR-fluence-rate shift, and the lincomycin treatment attenuates this induced expression. The expression of these genes is repressed by the lincomycin treatment and this repression becomes more severe following the BR-fluence-rate shift (Figure 4.3; Figure 4.5A and B).

Translation-related GO terms are also significantly enriched in the entire dataset and are most enriched in expression pattern 32 and expression cluster E (Figure 4.3).

These expression patterns are exemplified by attenuated light-induced expression in lincomycin-treated seedlings relative to untreated seedlings (Figure 4.3 and Figure

4.5C).

We found distinctive expression patterns among particular groups of chloroplast- related genes. (1) The expression of most genes that are annotated as encoding components of the thylakoid lumen is induced following the BR-fluence-rate shift to a greater degree in untreated than lincomycin-treated seedlings and is repressed in lincomycin-treated seedlings at all time points (Ruckle and Larkin unpublished data). (2)

In contrast, the expression of many genes annotated as encoding components of the thylakoid membrane, photosystem I, and photosystem II is only slightly regulated by the

BR-fluence-rate shift, but strongly repressed by the lincomycin treatment (Figure 4.3).

(3) Although the expression of genes annotated as contributing to the plastid stroma,

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plastid organization, and carotenoid metabolic processes is generally induced by the

BR-fluence-rate shift, these genes do not show the same degree of negative regulation by the lincomycin treatment as genes annotated as contributing to the thylakoids (Figure

4.3). (4) Forty nine chloroplast genes were significantly regulated by light or plastid signals. The lincomycin treatment regulates the expression of these chloroplast genes to a greater degree than the fluence-rate shift. In many instances, the lincomycin treatment induced the expression of chloroplast genes that are annotated as contributing to the plastid ribosome but repressed expression of chloroplast genes that are annotated as contributing to photosynthesis (Figure 4.5C).

The expression profile of particular plastid-related genes deviates significantly from the expression profile of the majority of plastid-related genes in that their light- induced expression is greater in lincomycin-treated seedlings relative to untreated seedlings. (1) As discussed above, the expression of a number of genes that are annotated as encoding components of the plastid ribosome is generally induced in lincomycin-treated seedlings relative to untreated seedlings (Figure 4.5C). (2) Some of the genes annotated as contributing to plastid organization exhibit higher expression following the BR-fluence-rate shift in lincomycin-treated seedlings (Ruckle and Larkin unpublished data). (3) The expression of several genes that contribute to photosynthesis, tetrapyrrole metabolism, and thylakoid function are induced more following the BR-fluence-rate shift in lincomycin-treated seedlings. These genes include

VAR2 (FtsH2), FtsH1, FtsH8, and FtsH11 (Figure 4.5A and B; Ruckle and Larkin unpublished data); psbO2 (Figure 4.5A); SEP2 and OHP2 (Ruckle and Larkin unpublished data); HCF107 and HCF136 (Figure 4.5A; Ruckle and Larkin unpublished

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data); GUN4 (Figure 4.5B); CLA1 (Figure 4.5B); and EXECUTER2 (Ruckle and Larkin unpublished data); PIL5/PIF1 (Figure 4.5B); and STN7 (Figure 4.5A).

Previously, light was reported to induce the expression of Lhcb1 when chloroplasts are functional and to repress the expression of Lhcb1 when chloroplasts are dysfunctional (Ruckle et al., 2007). This sort of expression pattern is observed among genes annotated as contributing to diverse functions (Ruckle and Larkin unpublished data). Several groups of these genes are discussed here. (1) We did observe this expression pattern among several other photosynthesis-related genes besides Lhcb1, such as Lhcb6, psbA, ndhA, psaB, ndhI, Lhcb5, and psbI; and tetrapyrrole metabolic cluster-related genes such as GLK1, GLK2, and PORB (Figure

4.5A and B). Although subtle (i.e., less than a twofold difference in expression level) for some of these genes, the finding that this trend is observed for a number of genes that contribute to thylakoid function provides evidence that this form of regulation is meaningful. The highly similar members of the Lhcb1 gene family do not appear in this dataset because probe sets that were assigned to more than one gene were removed from the dataset when the raw data were processed. (2) The lincomycin treatment also changes the nature of light regulation for genes that contribute to circadian rhythms.

The expression of LHYR (Ruckle and Larkin unpublished data), CCA1, FIO1, PIF5/PIL6,

PRR5, PRR7, ZTL, and other genes that are annotated as contributing to the circadian clock (Ruckle and Larkin unpublished data) is repressed at 4 h and/or 24 h following the

BR-fluence-rate shift in untreated seedlings and is induced following the BR-fluence- rate shift in lincomycin-treated seedlings. The lincomycin treatment also changes the nature of the light-regulated expression of RVE1 (Supplemental Ruckle and Larkin

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unpublished data). (3) Although not enriched in the entire dataset, genes annotated as contributing growth- and development-related processes such as the cell cycle and

-7 DNA replication are significantly enriched in expression cluster EE (p = 5.01 × 10 ;

-6 Figure 4.3) and expression pattern 27 (p = 3.80 × 10 ; Figure 4.3), respectively. The expression of both groups of genes is induced following the BR-fluence-rate shift in untreated seedlings and repressed by this BR-fluence-rate shift in lincomycin-treated seedlings (Figure 4.3; Ruckle and Larkin unpublished data). Several expansion genes exhibit a similar expression pattern (Ruckle and Larkin unpublished data). (4) The expression of several genes that are required for the biosynthesis and transduction of the jasmonic acid (JA) signal (i.e., ACS6, AOS, AS1, LOX2, JAZ5, and JAZ6), is induced following the BR-fluence-rate shift in untreated seedlings but repressed following the BR-fluence-rate shift in lincomycin-treated seedlings (Ruckle and Larkin unpublished data).

The production of chloroplastic ROS can contribute to plastid-to-nucleus signaling (Woodson and Chory, 2008; Galvez-Valdivieso and Mullineaux, 2010).

However, based on analyses of plastid signaling mutants, others concluded that ROS is not a major driver of plastid-to-nucleus signaling in seedlings treated with inhibitors of chloroplast biogenesis (Strand et al., 2003; Voigt et al., 2009). These transcriptome data provide a distinct approach for testing whether chloroplastic ROS might contribute to the remodeling of light signaling in lincomycin-treated seedlings as it does in the flu mutants (Danon et al., 2006). Genes annotated as contributing to oxidative stress are

-10 enriched in this dataset (p = 5.75 × 10 ; Figure 3.4). However, the expression of

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these genes was similar following the BR-fluence-rate shift, regardless of whether

-4 seedlings were treated with lincomycin (i.e., positive correlation, p = 3.16 × 10 ; Figure

3.4). Further, the predominant response was down-regulated expression in response to the BR-fluence-rate shift (Ruckle and Larkin unpublished data). Other classes of genes that contribute to oxidative stress tolerance and are significantly enriched in this dataset

-5 include those related to phenylpropanoid metabolic processes (p = 3.89 × 10 ),

-4 carotenoid metabolic processes (p = 7.41 × 10 ), and glycoside metabolic processes (p

-4 = 2.51 × 10 ). In general, the light-induced expression of phenylpropanoid- and carotenoid-related genes is similar in lincomycin-treated and untreated seedlings. The light-induced expression of glycoside-related genes is attenuated in lincomycin-treated seedlings (Figure 4.3 and 4.4; Ruckle and Larkin unpublished data). Further, although

-3 cell-death-related genes are enriched in the entire dataset (p-value = 1.86 × 10 ;

Figure 4.4), the expression patterns of these genes are diverse and are therefore not consistent with interactions between light and plastid signals driving increases in cell death (Ruckle and Larkin unpublished data). Thus, based on these transcriptome analyses, lincomycin-treated seedlings and untreated seedlings would appear to contain similar levels of oxidative stress following the BR-fluence-rate shift.

To further test whether plastidic ROS might drive the remodeling of light signaling observed in lincomycin-treated seedlings, we tested whether light and plastid signals regulate the expression of genes that are induced at least fivefold by diverse ROS from various cellular compartments (Gadjev et al., 2006). We found that the majority of these

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genes are either (1) not significantly enriched in our dataset (i.e., not significantly regulated by light and plastid signals) or (2) significantly enriched but their expression is repressed by the BR-fluence-rate shift regardless of whether seedlings are treated with lincomycin. Genes whose expression is induced by singlet-oxygen or by superoxide

(Gadjev et al., 2006) are enriched in the overall dataset (Ruckle and Larkin unpublished data). The majority of these genes were similarly expressed following the BR-fluence- rate shift regardless of whether seedlings are treated with lincomycin. Based on these analyses of ROS-inducible genes (Gadjev et al., 2006) and our GO analyses, we conclude that light and plastid signals can likely regulate these ROS-responsive genes using signaling mechanisms that do not depend on ROS and that ROS is likely not

HVVHQWLDOIRUWKH³UHZLULQJ´RIOLJKWVLJQDOLQJE\SODVWLGVLJQDOV

Abiotic stress can cause chloroplast dysfunction by attenuating the biosynthesis of proteins that are essential for the function of photosystem II (Takahashi and Murata,

2008). Therefore, treating plants with an inhibitor of chloroplast translation such as lincomycin may to some degree simulate the various chloroplast stresses that plants experience in nature. If this idea is correct, then plants experiencing abiotic stress may convert light signaling from a process that promotes chloroplast biogenesis and function to one that attenuates these processes by a mechanism that at least in part depends on plastid-to-nucleus signaling. Consistent with this idea, abiotic stress response-related genes are significantly enriched in our dataset. These genes include those associated

-13 - with responses to cold stress (p = 1.07 × 10 ), water-deprivation stress (p = 1.62 ×10

13 -12 -8 - ), salt stress (p = 5.75 ×10 ), heat stress (p = 3.72 ×10 ), metal ions (p = 8.51 ×10

219

6 -12 ), and abscisic acid (ABA; p = 3.55 ×10 ) (Figure 4.4). The clustering of genes annotated as contributing to cold, ABA, and osmotic stress responses (i.e., water deprivation and salt stress) with particular user-defined expression patterns (Figure

4.12A) is consistent with considerable overlap in genes annotated as contributing to these GO terms or in considerable overlap in the expression patterns of genes that contribute to these processes. For most of these genes, expression is significantly regulated at 4 h and at 24 h following the BR-fluence-rate shift in untreated seedlings and is attenuated in lincomycin-treated seedlings. In many instances, the lincomycin treatment attenuated the down-regulation of stress-related genes that followed the BR- fluence rate shift. This response is also observable as fluence-rate shift converting the lincomycin treatment from a negative to a positive regulator of stress-related genes

(Figure 4.4). Genes that contribute to particular abiotic stress responses especially osmotic stress responses such as AtMYB60, AZF2, ASN1/DIN6, ERD10, DREB2A,

DREB2B, DREB2D, ERD1, HOS1, KIN10, LEA14, RD22, SIZ1, SnRK2.2, SnRK2.3,

STZ/ZAT10, ZAT12 (Fujita et al., 2009; Saibo et al., 2009) exhibited distinct expression patterns following the fluence-rate shift in lincomycin-treated and untreated seedlings

(Ruckle and Larkin unpublished data). In contrast, heat-inducible genes are only transiently induced following the BR-fluence-rate shift and are similarly regulated in lincomycin-treated and untreated seedlings (Figure 4.4; Ruckle and Larkin unpublished data).

Genes annotated as contributing to the responses to various light intensities and various light qualities are enriched in the overall dataset (Figure 4.4). In general, an effect of the lincomycin treatment on the light-regulated expression of these genes was

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most striking for those genes that are also annotated as contributing to plastid function

(Ruckle and Larkin unpublished data). Additionally, the lincomycin treatment strikingly affects the expression of genes annotated as contributing to the circadian clock. This treatment converts the fluence-rate shift from a negative to a positive regulator of numerous circadian-related genes as described above. This treatment also attenuates the negative regulation that follows the fluence-rate shift in untreated seedlings for several additional genes that contribute to circadian rhythms such as EFL3, GI, LKP2,

TOC1, TIC, and other genes that are annotated as contributing to the circadian clock

(Ruckle and Larkin unpublished data).

The major hormone-related genes enriched in this dataset are those genes

-12 annotated as contributing to ABA responses (p = 3.55x10 ) as discussed above.

-3 Auxin, gibberellin (GA), and JA are enriched to a lesser degree (p = 3.02 × 10 , 2.95 ×

-8 -8 10 , 1.26 × 10 , respectively) (Figure 4.4). The expression of genes annotated as contributing to auxin stimulus and GA stimulus is repressed by the BR-fluence-rate shift and the lincomycin treatment attenuates this regulated expression (Figure 4.4; Ruckle and Larkin unpublished data). In general, the expression of genes annotated as contributing to JA stimulus is induced following the fluence-rate shift, and this expression is attenuated by the lincomycin treatment (Figure 4.4; Ruckle and Larkin unpublished data).

The BR-fluence-rate shift and the lincomycin treatment had diverse effects on the expression of genes annotated as contributing to the regulation of gene expression for several classes of metabolic processes (Figure 4.3). Based on cluster analysis of

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individual genes (Ruckle and Larkin unpublished data), the enrichment of genes

-4 annotated as contributing to carbon utilization in the entire dataset (p = 3.31 × 10 ;

Figure 4.3) appears to result mostly from their contribution to photosynthesis. The expression of these genes is induced by the BR-fluence-rate shift, but attenuated by the lincomycin treatment (Figure 4.3). The expression of genes annotated as contributing to starch metabolism is induced by the BR-fluence-rate shift and attenuated by the lincomycin treatment. In contrast, genes annotated as contributing to monosaccharide metabolic processes are more highly expressed than those annotated as contributing to starch metabolism following the fluence-rate shift in lincomycin-treated seedlings

(Figure 4.3). Genes annotated as contributing to amino acid metabolism are enriched in the overall dataset (p = 3.31 × 10-7; Figure 4.3). In general, the light-regulated expression of these genes is attenuated by lincomycin treatment (Figure 4.3; Ruckle and Larkin unpublished data). The expression of genes annotated as contributing to components of the mitochondria is induced following the BR-fluence-rate shift. In general, this induced expression is attenuated in lincomycin-treated seedlings (Figure

4.3; Ruckle and Larkin unpublished data).

We found that the expression of genes annotated as contributing to transcription regulation dominated the transcriptomes at 0.5 h and 1 h following the fluence-rate shift; the expression of genes annotated as contributing to metabolism, translation, growth, development, and stress and oxidative stress was induced later (4 h to 24 h). These findings are consistent with previous findings for untreated seedlings (Tepperman et al.,

2001; Jiao et al., 2003; Tepperman et al., 2004; Tepperman et al., 2006). As discussed

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above, the lincomycin treatment had a significant effect on the light-regulated expression of a number of these genes (Figure 4.3 and 4.4).

Reverse genetic analysis of particular light- and plastid-regulated genes

The integration of light and plastid signaling is proposed to depend on light and plastid signaling inducing the activity of downstream signaling proteins that contribute to both light and plastid signaling (Ruckle et al., 2007; Larkin and Ruckle, 2008). If this model is correct and if the activities of these proteins and the expression of the genes that encode these proteins are similarly regulated, then at least some of the genes whose expression is more highly induced by light in lincomycin-treated seedlings than in

XQWUHDWHGVHHGOLQJVVKRXOGFRQWULEXWHWRWKLV³UHZLULQJ´RIOLght signaling by plastid signals. To test this idea, we identified genes that (1) exhibit increases in expression 1 h following the BR-fluence-rate shift, (2) are expressed at levels at least 1.5-fold higher in lincomycin-treated seedlings than in untreated seedlings, and (3) are annotated as encoding proteins that contribute to transcription, signaling, and unknown functions. We identified 38 genes that meet these criteria (Table 4.1 and Table 4.2). T-DNA insertion mutants were publicly available for 25 of these genes, and two T-DNA alleles were available for 7 of these genes (Table 4.1). T-DNA alleles were not publicly available for the remaining 13 genes (Figure 4.2). We propagated these mutants and obtained homozygous lines for 32 of these T-DNA insertion mutants. Most of these T-DNA alleles are nulls or severe loss-of-function alleles based on an RT-PCR analysis (Ruckle and Larkin unpublished data). For a control group of mutants that are not expected to

223

DIIHFWWKLV³UHZLULQJ´RIOLJKWVLJQDOLQJEy plastid signals, we identified genes that (1) exhibit increases in expression 1 h after the BR-fluence-rate shift, (2) exhibit similar levels of expression in lincomycin-treated and untreated seedlings, and (3) are also annotated as encoding proteins with functions related to transcription, signaling, or unknown functions. We obtained 28 publicly available mutants, of which 22 had T-DNA insertions. We propagated these mutants and obtained lines that are homozygous for each T-DNA insertion. Most of these T-DNA alleles are nulls or severe loss-of-function alleles based on RT-PCR analysis (Ruckle and Larkin unpublished data).

To determine whether these T-DNA insertion mutants have defects in chloroplast biogenesis, we tested the efficiencies of the etioplasts-to-chloroplast conversion in these mutants and wild type. Dark-grown seedlings contain etioplasts rather than chloroplasts and do not contain chlorophyll. When dark-grown seedlings are transferred to the light, etioplasts are converted into chloroplasts. Chloroplast biogenesis from etioplasts is marked by the accumulation of chlorophyll (Waters and Langdale, 2009).

Thus, quantifying chlorophyll levels in mutants and wild-type seedlings during de- etiolation provides an efficient assay for monitoring the conversion of etioplasts to chloroplasts. To perform this chloroplast biogenesis assay, we first grew wild-type

Arabidopsis and each of these mutants for 4 d in the dark, transferred them to 125 µmol

-2 -1 m s broad spectrum white light for 24 h, and then quantified chlorophyll levels in each mutant. For a control, we used gun1-101; gun1-101 and other gun1 alleles cause inefficient greening, especially when fluence rates are increased (Mochizuki et al., 1996;

Ruckle et al., 2007). Of the 32 T-DNA insertion alleles derived from genes that are more highly expressed in lincomycin-treated than in untreated seedlings following the BR-

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fluence-rate shift, 20 (63%) caused enhanced de-etiolation (end) phenotypes. These 20 mutants accumulate at least twofold more chlorophyll per mg fresh weight than wild type during de-etiolation (Figure 4.6A). This group of mutants defines sixteen genes. Two independently isolated alleles caused an end phenotype for 6 of these genes, but only single alleles were publicly available for the remaining 10 genes (Figure 4.6A).

Nonetheless, the overrepresentation of the end phenotype in this group of mutants provides evidence that for the majority of these 10 alleles, the end phenotype is probably not caused by unlinked alleles. Only 4 mutants from this group (i.e., 13-34,

14-30, 20-26, and 35-83) accumulated essentially the same amount of chlorophyll as wild type (Figure 4.6A). Only one mutant (i.e., 7-85) accumulated significantly less chlorophyll than wild type in this de-etiolation experiment (Figure 4.6A). The remaining

7 mutants from this group accumulated significantly more chlorophyll than wild type during de-etiolation but did not accumulate a mean quantity of chlorophyll that was at least twofold more than wild type (Figure 4.6A). Although we did not classify the mutants with these more subtle phenotypes as end mutants, the genes defined by these mutant alleles may contribute to chloroplast biogenesis.

In contrast to the high frequency of end phenotypes caused by T-DNA insertion alleles of genes that are more highly expressed in lincomycin-treated than untreated seedlings following the BR-fluence-rate shift, T-DNA insertion alleles of genes whose expression is similarly induced in lincomycin-treated and untreated seedlings following the BR-fluence-rate shift do not yield a high frequency of end phenotypes. Indeed, only

3 mutants that define only 2 of the 22 genes (9%) from this group accumulate at least twofold more chlorophyll per mg fresh weight than wild type (Figure 4.6B; Table 4.2).

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Two independently isolated T-DNA alleles were available for only one of these genes

(Figure 4.6B; Table 4.2). Eleven of these alleles caused significantly more chlorophyll to accumulate than in wild type but not at least twofold more chlorophyll than wild type as observed among end mutants. Only one T-DNA allele was available for each of these 11 genes (Figure 4.6B). In addition to the low frequency of end phenotypes, this group of mutants is further distinguished from the previous group in that there was no significant difference in the amount of chlorophyll accumulation relative to wild type for

50% of these mutants (Figure 4.6B). In contrast, only 9% of the mutants from the previous group accumulated levels of chlorophyll that are not significantly different from wild type during this de-etiolation experiment (Figure 4.6A). The results reported in

Figure 4.6A and B indicate that expression profiling of lincomycin-treated and untreated seedlings can provide and efficient screen for genes that help attenuate chloroplast biogenesis.

To further test whether particular groups of mutants might exhibit a high frequency of end phenotypes, we examined end phenotypes in a large group of light signaling mutants. We expect that a group of light signaling mutants will exhibit at least a low frequency of end phenotypes because light signaling is a major regulator of chloroplast biogenesis. Only 25% of mutant alleles from this group of alleles that defines 40 genes cause more chlorophyll to accumulate during de-etiolation than wild type (Figures 4.13).

The end phenotypes of cry1-92, phyA-75, and phyB-35 (Figure 4.13) are consistent with cry1, phyA, and phyB inducing the expression of the END genes. These end phenotypes are also consistent with cry1-92, phyA-75, and phyB-35 experiencing less photooxidatives stress than wild type during de-etiolation and these photoreceptors not

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inducing the expression of the END genes. This overaccumulation of chlorophyll in cry1-92, phyA-75, and phyB-35 would appear to conflict with previously published data showing that loss-of-function alleles of CRY1, PHYA, and PHYB cause chlorophyll deficiencies (Neff and Chory, 1998). The end phenotypes of cry1-92, phyA-75, and phyB-35 are likely conditional. Consistent with this interpretation, specific parameters of de-etiolation experiments were previously reported to cause particular light signaling mutants to either overaccumulate or underaccumulate chlorophyll (Stephenson et al.,

2009). Analysis of chlorophyll accumulation phenotypes of other light signaling mutants provides evidence that, during de-etiolation, loss-of-function alleles of Atmyc2-05, hfr1-

27, gbf1-12, hrb1-68, and spa1-40 can promote the accumulation of chlorophyll and that loss-of-function alleles of det1-1, cop1-4, pif1-72, fhy3-11, and pif3-27 can attenuate the accumulation of chlorophyll during de-etiolation (Figure 4.13). Consistent with these findings, a loss-of-function allele of AtMYC2 was previously reported to accumulate more chlorophyll than wild type (Yadav et al., 2005), det1-1 was previously reported to cause chlorophyll deficiencies (Chory et al., 1989), and cop1-4 and loss-of-function alleles of PIF1 and PIF3 were previously reported to attenuate greening in de-etiolation experiments (Ang and Deng, 1994; Stephenson et al., 2009). Based on these data, we conclude that our reverse genetic screen yields a high frequency of end phenotypes.

The T-DNA alleles that cause end phenotypes may up-regulate thylakoid biogenesis or down-regulate chloroplast stress such as the sort of photooxidative stress that can attenuate the accumulation of chlorophyll. Testing whether increasing fluence rates affect chlorophyll accumulation during de-etiolation can provide evidence of photooxidative stress. Thus, we performed de-etiolation experiments with these end

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-2 mutants in three fluence rates of broad spectrum white light: 15, 100, and 300 µmol m

-1 s . In general, end phenotypes are more striking when de-etiolation is performed in

-2 -1 100 µmol m s and less striking when de-etiolation is performed in either 15 or 300

-2 -1 µmol m s (Figure 4.7A). This trend was also observed in wild-type plants, even

-2 -1 though the differences between 15 and 100 µmol m s were not significant for wild

-2 -1 type (Figure 4.7A). We suggest that 15 µmol m s provides insufficient light for

-2 -1 optimal de-etiolation, 300 µmol m s provides excess light that yields photooxidative

-2 -1 stress during de-etiolation, and that 100 µmol m s provides sufficient light for de- etiolation without causing excessive photooxidative stress. end mutants that exhibit enhanced thylakoid biogenesis relative to wild type and that experience similar levels of photooxidative stress as wild type are expected to accumulate more chlorophyll than

-2 -1 wild type when de-etiolation is performed in 15 and in 100 µmol m s . The end mutants in this class are not expected to accumulate significantly more chlorophyll than

-2 -1 wild type when de-etiolation is performed at 300 µmol m s . Several end alleles that define six genes cause such phenotypes (i.e., 1-83, 3-F12, 5-74, 15-H05, 15-F11, 17-12,

17-43, 23-D06, 23-F03; Figure 4.7A). end mutants that perform similar rates of thylakoid biogenesis as wild type and that experience less photooxidative stress than wild type are expected to accumulate more chlorophyll than wild type when de-etiolation

-2 -1 is performed in both 100 and in 300 µmol m s and accumulate similar levels of

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-2 -1 chlorophyll as wild type when de-etiolation is performed at 15 µmol m s . Although several end alleles that define seven genes (i.e., 2-31, 6-48, 8-49, 8-D01, 19-29, 21-35,

21-76, 25-G09, and 29-18) cause significantly more chlorophyll to accumulate than in

-2 -1 the wild type when de-etiolation is performed in both 100 and 300 µmol m s , these alleles also cause more chlorophyll to accumulate when de-etiolation is performed in 15

-2 -1 µmol m s (Figure 4.7A). Thus, end alleles attenuating photooxidative stress is not a satisfactory explanation for these end phenotypes. Possible mechanisms underlying these end phenotypes are explored below. Two of the end alleles (i.e., 22-B11 and 38-

44) cause more chlorophyll to accumulate than in wild type only when de-etiolation is

-2 -1 performed at 100 µmol m s (Figure 4.7A). The end phenotypes of 22-B11 and 38-44 are more subtle than those exhibited by other end mutants. Allele 7-85 was the only one yielded by this screen that causes less chlorophyll to accumulate relative to wild type during de-etiolation. Allele 7-85 resembles gun1-101 in that it de-etiolates less efficiently as fluence rate increases (Figure 4.7A).

The bulk of the end mutants yielded from this screen either (1) accumulate more

-2 chlorophyll than wild type when de-etiolation is performed at 15, 100, and 300 µmol m

-1 s or (2) accumulate more chlorophyll than wild type when de-etiolation is performed at

-2 -1 15, and 100 µmol m s , but accumulate essentially the same levels of chlorophyll as

-2 -1 wild type when de-etiolation is performed at 300 µmol m s (Figure 4.7A). One possible interpretation of these data is that all of these end alleles enhance chloroplast

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biogenesis relative to wild type and experience similar degrees of photooxidative stress as wild type but that the up-regulation of chloroplast biogenesis caused by several of

-2 - these end alleles cannot withstand the photooxidative stress yielded by 300 µmol m s

1 white light (i.e., 1-83, 3-F12, 5-74, 15-H05, 15-F11, 17-12, 17-43, 23-D06, 23-F03). If this model is correct, all of these end mutants should accumulate chlorophyll more rapidly than wild type and alleles that cause more photooxidative stress during de- etiolation (e.g., gun1-101 and 7-85) should accumulate chlorophyll at the same rate as wild type when de-etiolation is performed under conditions that essentially abolish photooxidative stress. To test this idea, we performed a de-etiolation experiment in BR

-2 -1 light that was 1 µmol m s and extracted and quantified chlorophyll at four intervals

-2 -1 from 0 h to 24 h. Consistent with 1 µmol m s BR light not promoting photooxidative stress, chlorophyll accumulated at similar rates in wild type and in mutants that appear to experience abnormal levels of photooxidative stress (i.e., gun1-101 and 7-85; Figure

4.7B). All of the end mutants tested accumulated chlorophyll more rapidly than wild

-2 -1 type when de-etiolation was performed in 1 µmol m s BR. More chlorophyll per mg fresh weight is apparent at 12 h and at 24 h for 15 of the end mutants (Figure 4.7B).

The remaining 5 end mutants tested accumulated more chlorophyll than wild type at only one time point (Figure 4.7B).

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DISCUSSION

Here, we report that light-regulated transcriptomes are radically different in

Arabidopsis seedlings that contain dysfunctional chloroplasts than in those that contain functional chloroplasts. Further, we report that light signaling can induce major changes in plastid-regulated transcriptomes. We found that the interactions between light and plastid signaling can have major effects on the expression of genes that are annotated as contributing to chloroplast-related functions such as photosynthesis, but also to a broad spectrum of functions including metabolism, stress responses, and both growth and development. Using these transcriptome data, we also developed a reverse genetic strategy that aimed to identify plastid-regulated factors that contribute to the

"rewiring" of light signaling in cells that contain dysfunctional chloroplasts. This screen yielded alleles that define 18 genes encoding proteins with signaling, transcription, or no known function that cause more chlorophyll to accumulate relative to wild type during de-etiolation.

Plastid signals are major regulators of light-regulated gene expression

We found that a major effect of plastid signaling triggered by lincomycin treatments is to reduce the size of the light-regulated transcriptome by approximately twofold. Therefore, we conclude that plastid signals are major regulators of light signaling in cells that contain dysfunctional chloroplasts. Thus, like hormones and organ-specific signals (Jiao et al., 2007; Jaillais and Chory, 2010), plastid signals can serve as major endogenous regulators of light signaling.

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The integration of light and plastid signaling promotes chloroplast function

We found that the BR-fluence-rate shift induced the expression of genes that contribute to a broad spectrum of biological functions that includes photosynthesis and other chloroplast functions in untreated seedlings that contain well-functioning chloroplasts. This finding is consistent with a number of reports that indicate a major role for light signaling in promoting chloroplast biogenesis and function by regulating gene expression (Ma et al., 2001; Tepperman et al., 2001; Ma et al., 2003; Monte et al.,

2004; Ohgishi et al., 2004; Tepperman et al., 2006; Leivar et al., 2009; Shin et al., 2009).

In contrast, we found that a major consequence of the integration of light signaling and the plastid signaling that is triggered by lincomycin treatments is down-regulated expression of genes that contribute to chloroplast functions. This down-regulated expression is especially striking for genes that contribute to the light reactions of photosynthesis. In seedlings treated with lincomycin, the expression of a large number of genes, including many that contribute to the light reactions of photosynthesis, is not induced by light. Additionally, the expression of a smaller number of genes, including several that contribute to the light reactions of photosynthesis and the transcription factor GLK1²a major driver of thylakoid biogenesis (Waters et al., 2009), is negatively regulated by light in lincomycin-treated seedlings.

Although the lincomycin treatment used here attenuates the light-induced expression for most of the genes annotated as contributing to photosynthesis, we found that the lincomycin treatment enhances the light-induced expression of particular

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photosynthesis-related genes. In general, this group of genes contributes to the protection or repair of distressed chloroplasts. These genes include: (1) FtsH1, VAR2

(FTSH2), FTSH8, and FtsH11, which encode ATP-dependent zinc metalloproteases.

FtsH1 and VAR2 help repair damaged photosystem II (Lindahl et al., 2000; Kato et al.,

2009). FtsH1, VAR2, and FtsH8 promote chloroplast biogenesis (Chen et al., 2000;

Takechi et al., 2000; Sakamoto et al., 2002; Zaltsman et al., 2005) and FtsH11 contributes to chloroplast function during heat stress (Chen et al., 2006). (2) PIL5/PIF1 encodes a nuclear transcription factor that attenuates photooxidative stress by helping regulate chlorophyll biosynthesis (Huq et al., 2004; Moon et al., 2008). (3) EXECUTER2 helps protect the chloroplast from singlet oxygen stress (Lee et al., 2007). (4) psbO2 promotes photosystem II activity during high-intensity-light stress (Allahverdiyeva et al.,

2009). (5) STN7 encodes a thylakoid-associated kinase that promotes optimal photosynthesis and recovery from photoinhibition (Goral et al., 2010; Lemeille and

Rochaix, 2010). (6) SEP2 and OHP2, which are related to the Lhc genes, are implicated in high-intensity-light stress tolerance (Heddad and Adamska, 2000; Andersson et al.,

2003). (7) HCF107 and HCF136 promote the assembly of photosystem II (Meurer et al.,

1998; Felder et al., 2001). (8) GUN4 binds chlorophyll precursors, induces chlorophyll metabolism (Larkin et al., 2003; Davison et al., 2005; Verdecia et al., 2005), and contributes to photoprotection (Larkin et al., 2003; Peter and Grimm, 2009). (9) CLA1 encodes 1-deoxy-D-xylose 5-phosphate synthase, a rate-limiting enzyme for the biosynthesis of the photoprotective plastidic isoprenoids (Estévez et al., 2001).

We propose that the integration of light and plastid signaling reported here promotes chloroplast biogenesis and function by tailoring nuclear gene expression to

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both the particular degree of chloroplast function and the particular light environment.

The main effects of this signal integration are the attenuation of chloroplast function and the simultaneous promotion of thylakoid repair and stress tolerance when chloroplasts experience dysfunction. We expect that this signal integration is significant because intense light can cause severe chloroplast stress and dysfunction. By integrating light and plastid signaling, we propose that plants can not only react to chloroplast dysfunction but also to the potential for light-induced chloroplast dysfunction.

Consistent with this idea, both light and plastid signaling affect the gene expression response to increasing light intensity (Ma et al., 2003; Ruckle et al., 2007), and both light and plastid signaling appear essential for efficient chloroplast biogenesis when seedlings are irradiated with intense light (Ruckle et al., 2007).

The idea that these transformative effects on light signaling are a primary effect of plastid-to-nucleus signaling rather than an indirect effect of chloroplast dysfunction is supported by previous kinetic experiments. A burst of chloroplastic singlet oxygen production in the Arabidopsis flu mutant converts cry1 signaling from a process that promotes chloroplast function to a process that inhibits chloroplast function and promotes photobleaching and cell death in as little as 4 h (Danon et al., 2006).

Alternative models in which more indirect mechanisms lead to this conversion, such as light signaling promoting plastid stress by regulating nuclear gene expression (i.e., an anterograde control-based mechanism), require extremely complex mechanisms to fit with previously published analyses of light and plastid signaling mutants (Ruckle et al.,

2007). Further, such models are counterintuitive. Why would a major signaling network such as the light signaling network make stress tolerance more difficult? The

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transcriptome analyses reported here provide an independent approach for testing this anterograde control model. These transcriptome analyses are consistent with light signaling driving no more oxidative stress or chloroplastic ROS production in lincomycin-treated seedlings than in untreated seedlings. This finding is consistent with inhibitors of chloroplast biogenesis depleting chloroplasts of their major sources of photooxidative stress such as photosynthetic electron transport and both chlorophylls and chlorophyll precursors (Mochizuki et al., 2008; Moulin et al., 2008). Indeed, experiments with ROS-responsive genes and dyes that detect ROS are consistent with inhibitors of chloroplast biogenesis causing ROS-deficiencies relative to untreated seedlings (Voigt et al., 2009). Thus, the plastid signals triggered by lincomycin treatments likely affects light signaling by a mechanism that does not necessarily depend on the ROS-based signals and photosynthesis-related signals that are also triggered by changes in the light environment in plants that contain well-functioning chloroplasts (Danon et al., 2006; Bräutigam et al., 2009; Galvez-Valdivieso and

Mullineaux, 2010) because lincomycin-treated seedlings are albinos that do not perform photosynthetic electron transport. Also, we conclude that crosstalk between light an plastid signaling does not necessarily cause cell death as it does in flu mutants (Danon et al., 2006) because inhibitors of chloroplast biogenesis do not cause cell death if seedlings are provided a carbon source (Susek et al., 1993; Ruckle et al., 2007), and our transcriptome analysis are not indicative the integration of light and plastid signaling driving cell death. The simplest interpretation of the available data is that the chloroplast dysfunction caused by inhibitors of chloroplast biogenesis triggers retrograde plastid-to-nucleus signaling that does not necessarily depend on ROS or

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photosynthesis. This chloroplast dysfunction rapidly remodels light signaling, converting it into a process that attenuates chloroplast function and promotes chloroplast repair and protection in cells that contain dysfunctional chloroplasts thereby promoting chloroplast biogenesis and function. These conclusions are consistent with previous analyses of chloroplast biogenesis in light and plastid signaling mutants (Ruckle et al.,

2007; Larkin and Ruckle, 2008). However, in response to extreme stress, the integration of light and plastid signaling can trigger cell death (Danon et al., 2006).

For chloroplast genes, the lincomycin treatment was a more significant regulator of gene expression than the BR-fluence-rate shift. These findings are consistent with the chloroplast genome producing nearly a full complement of mRNAs during chloroplast biogenesis (Toyoshima et al., 2005) and the lincomycin treatment disrupting the biosynthesis of mRNA that occurs during chloroplast biogenesis. We found that chloroplast dysfunction triggered by the lincomycin treatment used here is a major regulator of photosynthesis-related genes that reside in both nuclear and chloroplast genomes and that photosynthesis-related genes residing in both nuclear and chloroplast genomes are coexpressed (Ruckle and Larkin unpublished data). These findings provide evidence that the plastid signals triggered by chloroplast dysfunction help coordinate the expression of photosynthesis-related genes localized in both nuclear and chloroplast genomes. Other evidence that plastid-to-nucleus signaling triggered by chloroplast dysfunction helps coordinate the expression of nuclear and chloroplast genes comes from an analysis of gun1 mutants, which are defective in plastid-to-nucleus signaling. gun1 alleles were found to disrupt the plastid-regulated

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expression of the Lhcb1 and RbcS, two nuclear genes, but not to affect the expression of psaAB, a chloroplast gene (Susek et al., 1993).

The integration of light and plastid signaling appears to affect growth and stress responses

Our transcriptome analyses are consistent with the interactions between light and plastid signaling affecting growth. Consistent with our analyses of light-regulated transcriptomes in untreated seedlings, light was previously reported to promote the expression of genes that contribute to cytosolic ribosomes, the cell cycle, and DNA replication (López-Juez et al., 2008). In contrast, light-induced expression of genes that are annotated as contributing to cytosolic ribosomes is attenuated and the expression of genes annotated as contributing to the cell cycle and DNA replication is repressed by light in lincomycin-treated seedlings. Further, the light-regulated expression of genes annotated as contributing to mitochondrial functions is attenuated by the lincomycin treatment, a finding that is consistent with the considerable interaction between chloroplasts and mitochondria (Noctor et al., 2007; Noguchi and Yoshida, 2008;

Woodson and Chory, 2008; Aluru et al., 2009). These findings are consistent with plastid-to-nucleus signaling converting light signaling from a process that promotes growth to one that inhibits growth and with the integration of light and plastid signaling coordinating growth with chloroplast biogenesis and function. Based on our finding that chloroplast dysfunction affects the light-regulated expression of genes that are annotated as contributing to hormone function, we suggest that integration of hormone,

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light, and plastid signaling may at least partially explain these effects on growth-related gene expression. Previous reports of crosstalk among hormone, light and plastid signaling are consistent with this interpretation (Acevedo-Hernández et al., 2005; Kim et al., 2009; Cottage et al., 2010; Jaillais and Chory, 2010). The idea that the conversion of light signaling from a process that promotes growth to one that inhibits growth is a primary effect of plastid-to-nucleus signaling rather than an indirect effect of chloroplast dysfunction is supported by previous kinetic experiments. A burst of chloroplastic singlet oxygen production in the Arabidopsis flu mutant affects cry1 signaling (Danon et al., 2006) as described above and causes a rapid inhibition of growth (op den Camp et al., 2003). Other data lend further support to the idea that the integration of light and plastid signaling coordinate growth with chloroplast biogenesis and function. (1)

Seedlings treated with inhibitors of chloroplast biogenesis are smaller than untreated even, when both treated and untreated are provided exogenous sucrose and especially as fluence rates are increased (Ruckle and Larkin, 2009). (2) Cells in white sectors of variegated mutants are smaller than those in adjacent green sectors (Yu et al., 2007).

The integration of light and plastid signaling does not absolutely attenuate the expression of growth-related genes. This crosstalk induces the expression of genes that encode components of the plastid ribosome and has little effect on the expression of many genes that encode components of plastid chromosomes. These data are consistent with a significant role for the plastid genome in plastid-to-nucleus signaling

(Koussevitzky et al., 2007) and the elevated expression of particular chloroplast genes that contribute to chloroplast genome expression in both seedlings treated with

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inhibitors of chloroplast biogenesis (Gray et al., 2003) and albino mutants (Cho et al.,

2009).

Biotic and abiotic stresses resemble the lincomycin treatment used here. (1)

Abiotic stress attenuates the expression of the photosynthesis-related chloroplast genes thereby inducing chloroplast dysfunction (Takahashi and Murata, 2008). (2) Both biotic and abiotic stresses down-regulate photosynthesis-related gene expression, attenuate growth, and induce stress responses. This diversion of resources from growth to stress tolerance is a major component of stress responses that remains poorly understood

(Herms and Mattson, 1992; Ballaré, 2009; Saibo et al., 2009; Bilgin et al., 2010).

Because of the similarities among transcriptome responses to biotic and abiotic stress and the lincomycin treatment used here, we propose that the integration of light and plastid signaling may contribute to biotic and abiotic stress tolerance. Consistent with this idea, we found that in general, light up-regulates the expression of photosynthesis- and growth-related genes but down-regulates the expression of most stress-related genes in untreated seedlings. Thus, light signaling appears to help plants invest in growth rather than stress tolerance when seedlings contain well-functioning chloroplasts.

In contrast, light signaling generally down-regulates the expression of photosynthesis- and growth-related genes in lincomycin-treated seedlings. Although stress-related gene expression is not light-induced in lincomycin treated seedlings, the down-regulated expression of stress-related genes that follows the fluence-rate shift in untreated seedlings is attenuated in lincomycin-treated seedlings. In general, stress-related genes are expressed at higher levels in lincomycin-treated seedlings than untreated seedlings following the fluence rate shift. Thus, crosstalk between light and plastid

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signaling would appear to promote stress tolerance in general by helping plants divert resources from growth to stress tolerance when seedlings contain dysfunctional chloroplasts. In addition to promoting this general stress response, the integration of light and plastid signaling appears to promote specific types of stress tolerance by regulating the expression of numerous genes that contribute to particular abiotic stress responses, especially osmotic stress responses.

In addition to abiotic stress responses, we suggest that crosstalk between light and plastid signaling might specifically affect JA signaling. Crosstalk between light and

JA signaling was demonstrated previously (Zhai et al., 2007; Moreno et al., 2009;

Robson et al., 2010). Our finding that chloroplast dysfunction (1) generally attenuates the light-regulated expression of genes that are annotated as contributing to the JA response and (2) converts light from a positive to a negative regulator of several genes that contribute to JA biosynthesis and signaling provides evidence that chloroplast dysfunction can affect the integration of light and JA signaling. These findings also provide evidence that the integration of light and plastid signaling might attenuate JA signaling when chloroplasts are dysfunctional. This interpretation is consistent with light signaling promoting JA signaling in green aerial tissues and sucrose attenuating this JA signaling (Robson et al., 2010); carbohydrates can negatively regulate chloroplast function (To et al., 2003; Stettler et al., 2009).

The integration of light and plastid signaling appears to affect the circadian clock

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We also found that crosstalk between light and plastid signals can have striking effects on the expression of genes that contribute to the circadian clock. Plastid signals can convert the light from a negative to a positive regulator of genes that encode core clock components such as CCA1, LHY, PRR5, PRR7, and ZTL (de Montaigu et al.,

2010; Pruneda-Paz and Kay, 2010). Plastid signals can similarly influence the light- regulated expression of genes that contribute to circadian regulated gene expression such as FIO1 (Kim et al., 2008), PIF5/PIL6 (Nozue et al., 2007), RVE1 (Laubinger et al.,

2006; Pruneda-Paz and Kay, 2010), and other genes that annotated as contributing to the circadian clock. In addition to these striking effects, we found that plastid signals can attenuate the light-mediated repression of several genes that contribute to circadian rhythms. These genes include those that contribute to the core circadian clock such as

GI and TOC1, other genes that contribute to circadian rhythms such as ELF3, LKP2, and TIC (de Montaigu et al., 2010; Pruneda-Paz and Kay, 2010), and other genes that are annotated as contributing to the circadian clock. Although genetic interactions between the circadian clock and both transcription factors that regulate chloroplast function (Stephenson et al., 2009) and chloroplast-localized RNA-binding proteins

(Hassidim et al., 2007) were reported previously, this report is the first to document such a large number of circadian-regulated genes exhibiting abnormal light-regulated expression in response to chloroplast dysfunction. Based on these findings, we suggest that interaction between light signaling and the plastid-to-nucleus signaling triggered by the lincomycin treatment used here and possibly triggered by biotic or abiotic stresses in nature might affect the activity of the circadian clock. We further propose that these interactions may help coordinate circadian clock activity with the functional and

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developmental state of the chloroplast. Consistent with this proposal, both lincomycin treatments and low temperatures induce chloroplast dysfunction by inhibiting the expression of the chloroplast genome (Takahashi and Murata, 2008) and both low temperatures (Bieniawska et al., 2008) and the lincomycin treatment used here affect the expression of genes that encode core components of the circadian clock.

The END genes down-regulate chloroplast biogenesis

Appropriate mutant screens should yield alleles that disrupt the integration of light and plastid signaling. Indeed, hy5 alleles disrupt these interactions (Ruckle et al.,

2007). However, an analysis of light signaling mutants by Ruckle et al. (2007) provides evidence that the light and plastid signals that regulate photosynthesis-related gene expression affect a complex network because single mutants can exhibit subtle gene expression phenotypes whereas double mutants can exhibit synergistically enhanced gene expression phenotypes. For example, the light signaling factors cry1 and HY5 are responsible for approximately 5% of the plastid regulation of Lhcb1 when seedlings are treated with inhibitors of chloroplast biogenesis, and GUN1 is responsible for 20 to 50% of the plastid regulation of Lhcb1 depending on the light conditions. However, cry1 and

GUN1 are apparently responsible for all of the plastid regulation of Lhcb1 in blue light

(i.e., the plastid does not appear to regulate Lhcb1 expression in cry1 gun1 double mutants in blue light) (Ruckle et al., 2007). Consistent with these findings, previous work indicates that both partial and complete redundancies among light signaling factors are common (Jiao et al., 2007; Leivar et al., 2008; Leivar et al., 2009; Sellaro et al., 2009; Shin et al., 2009; Chory, 2010) and that the synergistic regulation of

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transcription by distinct signals is also common (Smale, 2010). Thus, screens that yield mutant alleles that cause subtle phenotypes would appear essential for filling the remaining gaps in our knowledge of this signaling.

Here, we describe a reverse genetic screen based on expression profiling that yielded alleles that cause defects in chloroplast biogenesis. Based on previous analyses of the light and plastid regulation of Lhcb1, we hypothesized that proteins contributing to the integration of light and plastid signaling are most active when seedlings are treated with inhibitors of chloroplast biogenesis and when fluence rates are increased (Ruckle et al., 2007; Larkin and Ruckle, 2008). Because the expression profiles of genes can mirror the activities of their encoded proteins, we expected that genes encoding at least some of these proteins are more highly expressed following the

BR-fluence-rate shift in lincomycin-treated than in untreated seedlings. There is precedence for signaling-related genes exhibiting such expression patterns; the expression of genes that encode particular transcription factors that participate in auxin and JA signaling is induced during auxin and JA signaling, respectively (Reed, 2001;

Thines et al., 2007). Indeed, T-DNA insertion alleles that disrupt 64% of the genes identified using this expression profiling approach cause chlorophyll to accumulate at significantly higher rates during de-etiolation and cause chlorophyll to accumulate to significantly higher levels during de-etiolation than in wild type. Control groups of mutants that were defined using distinct expression patterns or using previously identified light signaling mutants exhibit a much lower frequency of end phenotypes. In total, we defined 18 END genes using expression profiling and 9 END genes by analyzing light signaling mutants. If inhibitors of chloroplast biogenesis like lincomycin

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were not useful for studying the signaling that contributes to chloroplast biogenesis because they induce too many secondary effects, we would expect to randomly isolate mutants with defects in chloroplast biogenesis from this screen at a low frequency. The high frequency of end phenotypes yielded by this screen demonstrates that inhibitors of chloroplast biogenesis are extremely useful tools for studying the signaling that contributes to chloroplast biogenesis. The finding that genes exhibiting enhanced light- induced expression in lincomycin seedlings are more likely to encode proteins that down-regulated chloroplast function supports one of the main conclusions from our transcriptome analyses; a major effect of the crosstalk between light and plastid signaling is to attenuate chloroplast function when chloroplasts are dysfunctional.

Few mutants overaccumulate chlorophyll. The Arabidopsis coi1 and ged1 mutants accumulate more chlorophyll than wild type during de-etiolation (Choy et al.,

2008; Robson et al., 2010). In Chlamydomonas reinhardtii, nab1 mutants also exhibit elevated levels of chlorophyll (Mussgnug et al., 2005). The ged1 allele has not been cloned. None of the end mutants are allelic to coi1. None of the proteins encoded by the END genes exhibit a striking sequence similarity to the NAB1. The end phenotype of the JA signaling mutant coi1 (Robson et al., 2010) further supports an interpretation of our transcriptome data; specifically, crosstalk among JA, light, and plastid signaling might affect chloroplast biogenesis.

For four of the END genes reported here, expression was reported to rapidly increase in response to light, and T-DNA insertion alleles of these END genes were reported to cause developmental defects in hypocotyls and cotyledons during de- etiolation (Khanna et al., 2006). Khanna et al. (2006) did not report whether these

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alleles cause chloroplast phenotypes. The genes that were also identified by Khanna et al. (2006) are SIG5 (At5g24120), a gene that encodes a putative protein kinase

(At2g30040), COL2 (At3g02380), and SPA1 (At2g46340). Other phenotypes were previously reported for sig5 and spa1, besides the enhanced chloroplast biogenesis phenotypes reported here and the developmental phenotypes reported by Khanna et al.

(2006). sig5 mutants are impaired in their ability to tolerate salt stress, osmotic stress, and high-intensity-light stress, but exhibit no striking phenotypes when grown in optimal conditions (Nagashima et al., 2004). Although spa1 mutants exhibit defects in light- regulated development (Hoecker et al., 1998; Khanna et al., 2006), photosynthesis- related gene expression (Hoecker and Quail, 2001; McCormac and Terry, 2002; Zhou et al., 2002), and chloroplast biogenesis following hourly pulses of far-red light

(Baumgardt et al., 2002), this report is the first to show that spa1 mutants can accumulate chlorophyll at higher rates and to higher levels of chlorophyll than wild type during de-etiolation. Ectopic expression of COL2 was reported to cause no striking phenotype (Ledger et al., 2001).

Among the remaining 14 END genes that were not identified by Khanna et al.

(2006), few have experimentally verified functions. Three of these END genes either encode or may encode signaling-related proteins: HAI1 (At5g59220), an F-box family protein (At2g16365), and a WD-40 repeat family protein (At5g23730). HAI1 encodes a protein phosphatase 2C that contributes to ABA signaling (Fujita et al., 2009). Three of these END genes encode transcription factors: SMZ (At3g54900), CDF1 (At5g62430), and RAP2.6 (Related to AP2 6; At1g43160). SMZ encodes an AP2-domain-containing protein that potently represses flowering (Schmid et al., 2003; Mathieu et al., 2009).

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CDF1 encodes a Dof transcription factor that affects flowering time (Imaizumi et al.,

2005). If, as we suggest based on our transcriptome analyses, plastid signaling

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END genes should contribute to stress responses. Indeed, RAP2.6 expression is light- induced in lincomycin-treated seedlings but light-repressed in untreated seedlings

(Ruckle and Larkin unpublished data) and is also affected by both biotic and abiotic stress. Further, RAP2.6 contributes to ABA and abiotic stress responses (He et al.,

2004; Zhu et al., 2010). Although none of these signaling or transcription factors were previously implicated in regulating chloroplast biogenesis or function, relatives of

RAP2.6 promote chloroplast function. ABI4 is related to RAP2.6 (Zhu et al., 2010) and contributes to the plastid- and sugar-regulated expression of photosynthesis-related genes (Acevedo-Hernández et al., 2005; Koussevitzky et al., 2007). Also, RAP2.2 contributes to carotenogenesis (Welsch et al., 2007). Further, RAP2.4 contributes to the production of chloroplastic antioxidants and to chloroplast function in fluctuating light environments (Shaikhali et al., 2008). The remaining eight END genes either have no known function or have functions that are inferred only from sequence similarities or gene expression patterns. Nonetheless, consistent with the end phenotypes reported here, three of these END proteins (At4g28740, At5g08050, and At5g13770) are predicted to reside in the chloroplast.

The expression of the END genes is regulated by a variety of signals besides light and plastid signals (Table 4.4). These diverse expression responses are consistent with the END genes contributing to a signaling network rather than to a linear pathway and with a variety of signals down-regulating chloroplast biogenesis and

246

function. The expression of At2g24540 and CDF (At5g62430) is regulated by the circadian clock (Table 4.4). These data are consistent with the integration of light and plastid signaling affecting circadian-regulated gene expression at least in part by regulating the expression of these genes and those defined by our transcriptome analyses. The expression of At2g24540, At5g35970, At2g41660, COL2 (At3g02380),

At1g43160, and At5g52250 is regulated by wounding and diverse abiotic stresses

(Supplemental Table 3). These data are consistent with the integration of light and plastid signaling affecting stress responses at least in part by regulating the expression of these genes and those defined by our transcriptome analyses.

247

MATERIALS AND METHODS

Plant Materials and Growth Conditions

Seedlings were grown in controlled-environment chambers containing light- emitting diodes or broad-spectrum fluorescent tube lamps (Percival Scientific, Perry, IA) at the indicated fluence rates in the presence and absence of lincomycin as described by Ruckle et al. (2007). Light was filtered though one or more layers of neutral-density filters to obtain different fluence rates of light as previously described by Ruckle et al.

(2007). Isolation and analysis of RNA by northern blotting was as described by Ruckle et al. (2007). T-DNA insertion mutants (Sessions et al., 2002; Alonso et al., 2003) obtained from the Arabidopsis Resource Center (ABRC) at the Ohio State University

(Columbus, OH) were propagated and homozygous lines were isolated. Homozygous lines were found to breed true in at least one subsequent generation using PCR-based genotyping as recommended by the Salk Institute Genomic Analysis Laboratory

(http://signal.salk.edu/). We determined the gene expression phenotype caused by each T-DNA insertion allele by comparing the levels of mRNA transcribed from each T-

DNA allele to the corresponding wild-type gene using RT-PCR as described by Ruckle et al. (2007).

Transcriptome analyses

To establish conditions that are useful for studying the impact of plastid signaling on the light-regulated transcriptome, total RNA was extracted from seedlings that were grown in various qualities and quantities of light, and the levels of Lhcb1 and RbcS

248

mRNA were quantified using RNA blot hybridizations as described by Ruckle et al.

(2007). For transcriptome analyses, biological replicates were grown separately under the same conditions. All experiments were performed with four biological replicates, with one exception. Only three biological replicates were used for the lincomycin- treated seedlings collected one hour after the fluence-rate shift. Each biological replicate contained 50 to 100 seedlings. To minimize variability among independent preparations of RNA, three independent RNA extractions were performed for each biological replicate using the RNeasy Plant Miniprep Kit (Qiagen, Valencia, CA) with the on-column DNase treatment. The three independent RNA extractions from each biological replicate were subsequently combined to generate one RNA preparation for each biological replicate. RNA samples from biological replicates were independently extracted, processed, and analyzed. Biotinylated target RNA was prepared from 5 µg total RNA for each sample using the GeneChip® One-Cycle Target Labeling (Affymetrix,

Santa Clara, CA). For each saPSOHȝJRIODEHOHGWDUJHWF51$ZDVSXULILHG fragmented, and hybridized to the GeneChip® Arabidopsis ATH1 Genome Array

(Affymetrix) as recommended by the manufacturer. The GeneChip arrays were then washed and stained using a GeneChip® Fluidics Station 450 (Affymetrix) and then analyzed using the GeneChip® Scanner 3000 7G (Affymetrix).

All data used in this study passed previously described quality assurance protocols (Burgoon et al., 2005). Microarray data were normalized using GCRMA in

R/Bioconductor (Wu and Irizarry, 2004). Posterior probabilities were calculated using an empirical Bayes analysis on a per gene, per time point, and per treatment basis. Both the unprocessed and normalized microarray data discussed in this

249

publication were deposited in NCBI's Gene Expression Omnibus (Edgar et al., 2002) and are accessible through GEO Series accession number GSE24517

(http://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE24517). Significant

GLIIHUHQFHVLQJHQHH[SUHVVLRQDUHGHILQHGDVWKRVHFDXVLQJDWZRIROGFKDQJH S

0.01). Trajectory analysis of differentially expressed genes was performed in R using singular value decomposition.

Agglomerative hierarchical clustering (Euclidean distance) of differentially expressed genes was performed in R. To create gene lists for each cluster, the

KHDWPDSVZHUHUHFRQVWUXFWHGLQ0LFURVRIW([FHOXVLQJWKH³URZ,QG´IXQFWLRQRI5 and the conditional formatting function from Microsoft Excel 2007. In addition to agglomerative hierarchical clustering, manual filtering of expression patterns was performed to identify genes that fit user-defined terms such as "genes induced early,"

"genes induced late," or "genes repressed by the chloroplast dysfunction." Microsoft

Excel 2007 was used to sort the genes that fit each model of expression to create gene lists for each model of expression.

Gene ontology (GO) terms were tested for enrichment in the expression clusters and user-defined expression patterns using the GO terms from the GO Consortium

(Ashburner et al., 2000) and Ontoligizer 2.0 (Bauer et al., 2008). The gene association file was downloaded from the Gene Ontology website (http://www.geneontology.org/)

CVS version 1.1260, GOC validation from July 21, 2009. The gene association file and the gene lists for both expression clusters and user-defined expression patterns were uploaded into Ontologizer 2.0 to identify GO terms that are enriched in each expression cluster or pattern as previously described (Grossmann et al., 2007; Bauer et al., 2008).

250

*2WHUPVZHUHFRQVLGHUHGVLJQLILFDQWO\HQULFKHGE\2QWRORJL]HULISDIWHUD

Westfall-Young correction for multiple testing as previously recommended (Grossmann et al., 2007). Ontoligizer 2.0 was not used to correct for redundancies between parent-

GO terms and the child-GO terms, which contain a subset of genes found in a particular parent-GO term. Only a term-for-term enrichment that separately tests for the enrichment of parent-GO and child-GO terms was used to test for the enrichment of GO terms. Results from this analysis yielded a list of GO terms that were enriched for at least one expression cluster or pattern. In this list, GO terms are represented by the negative log10 of uncorrected p-values. Next, a matrix that compares the negative log10 of uncorrected p-values and enriched expression patterns was created in

Microsoft Excel 2007. Agglomerative hierarchical clustering was performed on this matrix as described above for differentially expressed genes. When parent- and child-

GO terms clustered together, the parent term was used as previously recommended

(Grossmann et al., 2007). Parent- and child-GO terms that did not cluster together were considered separate terms. The child term was used as the GO-enriched term only in these instances. From this analysis, fifty-five GO terms were classified as distinct and enriched terms. We found the genes that made up these 55 enriched terms at the GO website using the advanced search function in AmiGO (Carbon et al., 2009) version 1.7, release date October 7, 2009. To remove potential artifacts of computational annotation of the GO, only genes that had the following evidence codes were considered: inferred from direct assay (IDA), inferred from experiment (EXP), inferred from expression pattern (IEP), inferred from genetic interaction (IGI), inferred from mutant phenotype

(IMP), inferred from physical interaction (IPI), traceable author statement (TAS), and

251

nontraceable author statement (NAS) (http://www.geneontology.org/)(Rhee et al., 2008).

Agglomerative hierarchical clustering was performed on significantly regulated genes from the enriched GO terms as described above for differentially expressed genes.

Genes that that are significantly regulated by particular types of ROS in particular subcellular locations were previously described by Gadjev et al. (2006). These ROS- regulated genes were tested for significant regulation by the BR-fluence-rate shift and the lincomycin treatment as described for GO terms. Significant regulation was established by calculating a hypergeometric distribution as described for estimating term-for-term overrepresentation of a GO term (Grossmann et al., 2007).

Analysis of chlorophyll levels in Arabidopsis seedlings

Chlorophyll was extracted from Arabidopsis seedlings that were grown in the indicated conditions and quantified as previously described (Porra et al., 1989), except that we homogenized 10 to 20 mg of seedlings in 1.5-ml microfuge tubes that contained a single 3-mm very high-density zirconium oxide bead (Glen Mills Inc., Clifton NJ) using a Retsch TissueLyser (Qiagen).

252

ACKNOWLEDGEMENTS

We thank Stephanie DeMarco for helpful assistance. This work was supported by NSF grants no. IOB-0517841 and IOS-1021755 and DOE grant no. DE±FG02± 91ER20021 to RML.

253

Figure 4.1 Kinetic analysis of Lhcb1 and RbcS mRNA expression following a fluence-rate shift. A) Relative expression of Lhcb1 following a fluence-rate shift. After stratification, -2 -1 the seeds were exposed to 100 mmol m s red light for 1 h and then placed in the dark for 23 h to promote uniform germination. Seedlings were grown for 6 d in -2 -1 -2 -1 0.5 ȝmol m s BR light and then transferred to 60 ȝmol m s BR light. Seedlings were collected and RNA was extracted at 0, 0.5, 1, 4, and 24 h following the fluence-rate shift. The levels of Lhcb1 mRNA relative to Lhcb1 mRNA levels at 24 h were determined from four biological replicates and quantified from RNA blots as described by Ruckle et al. (2007). B) Relative expression of RbcS following a fluence-rate shift. RNA was extracted and quantified as described in (A).

254

255

Log2 fold change

256

Figure 4.2 The light-regulated transcriptome in lincomycin-treated and untreated seedlings. A) Venn diagram of light and plastid-regulated genes. Light-regulated genes are GHILQHGDVWKRVHWKDWDUHH[SUHVVHGWZRIROGKLJKHURUORZHU S DWKK 4 h, or 24 h after the fluence-rate shift than before the shift (0 h). Plastid-regulated genes are those that meet the same fold change and significance criteria used to classify a gene as light regulated when the expression level of a particular gene in lincomycin-treated (+Lin) seedlings is normalized to the expression level in untreated (-Lin) seedlings at 0 h, 4 h, or 24 h. B) Numbers of genes regulated by light and lincomycin treatment after a BR- fluence-rate shift. Numbers of genes that exhibited a significantly different expression level in +Lin or -Lin seedlings at 0.5 h, 1 h, 4 h, and 24 h after a BR fluence-rate shift are indicated. The 3335 genes that are significantly regulated only by light are indicated with orange and light orange. The 3089 genes that are regulated by both light and lincomycin treatment are indicated with purple and light purple. Plastid regulation is presented for the 3089 genes that are regulated by light and plastid signals in blue and light blue. The plastid regulation for the 680 genes regulated only by the plastid is presented in green and light green. C) Principal component analysis of lincomycin treatment affecting the composition of the light-regulated transcriptome. Trajectory plots show the first principal component (PC1) and the second principal component (PC2), which are two orthogonal factors that describe 61% and 18%, respectively, of the variance caused by the BR-fluence-rate shift. D) Principal component analysis of the BR-fluence-rate shift on the lincomycin- regulated transcriptome. These trajectory plots show PC1 and PC2, which account for 79% and 21%, respectively, of the variance in the dataset. E) Agglomerative hierarchical clustering of the 7104 significantly regulated genes based on their regulation by the BR-fluence-rate shift. Nine basic expression patterns were identified (A-I). F) Agglomerative hierarchical clustering of the 7104 significantly regulated genes based on their regulation by lincomycin treatment. Eight basic expression patterns were identified (J-Q).

257

Figure 4.3

A Expressed Enrich- genes / ment in Total all genes in GO Term GO ID genes GO term

-log(p)

Plastid

1 P Photosyntheisis GO:0015979 28.12 74/90 2 C Thylakoid lumen GO:0031977 16.34 45/48 3 C Photosystem II GO:0009523 13.08 14/15 Thylakoid 4 C GO:0042651 19.06 48/64 membrane 5 C Plastid stroma GO:0009532 15.97 80/117 6 C Plastoglobule GO:0010287 14.57 49/56 7 C Photosystem I GO:0009522 4.85 8/9 8 C Stromule GO:0010319 4.32 22/33 NAD(P)H 9 C dehydro-genase GO:0010598 4.12 9/9 complex Tetrapyrrole 10 P GO:0033013 5.25 38/61 metabolic process Plastid 11 P GO:0009657 5.27 49/84 organization Carotenoid 12 P GO:0016116 3.13 16/24 metabolic process 13 C Plastid ribosome GO:0009547 3.45 16/24 Plastid 14 C GO:0009508 3.71 13/16 chromosome

15 C Plastid membrane GO:0042170 2.55 57/104

258

Figure 4.3 (Continued)

GO Term Light pattern of expression with the highest enrichment

Pattern +LIN -LIN -log(p) number 0.5 0.5 24 24 1 Plastid 4 1 4

1 Photosyntheisis 12 14.39 2 Thylakoid lumen 12 16.78 3 Photosystem II 12 4.11 Thylakoid 4 12 16.12 membrane 5 Plastid stroma 34 Positive correlation 10.66 6 Plastoglobule 12 7.19 7 Photosystem I 32 3.92 8 Stromule 12 8.95 NAD(P)H 9 dehydro-genase 12 6.70 complex Tetrapyrrole 10 metabolic 12 5.30 process Plastid 11 30 6.70 organization Carotenoid 12 metabolic N/A N/A N/A process 13 Plastid ribosome N/A N/A N/A Plastid 14 10 5.96 chromosome Plastid 15 N/A N/A N/A membrane

259

Figure 4.3 (Continued)

C Light cluster of expression with the highest GO Term enrichment

Cluster +LIN -LIN -log(p) letter 0.5 0.5 24 24 4 Plastid 1 1 4

1 Photosyntheisis B 20.37 2 Thylakoid lumen B 27.02 3 Photosystem II B 5.97 Thylakoid 4 B 19.39 membrane 5 Plastid stroma B 20.27 6 Plastoglobule B 12.43 7 Photosystem I B 4.35 8 Stromule B 12.80 NAD(P)H 9 dehydro-genase Y 8.90 complex Tetrapyrrole 10 metabolic B 9.77 process Plastid 11 Z 9.16 organization Carotenoid 12 metabolic B 6.84 process 13 Plastid ribosome EE 6.65 Plastid 14 Y 4.96 chromosome Plastid 15 Z 5.76 membrane

260

Figure 4.3 (Continued)

D Plastid pattern of expression Plastid cluster of expression GO Term with the highest enrichment with the highest enrichment

Pattern Cluster +LIN/ -log +LIN/-LIN -log(p) number letter -LIN (p) 24 24 0 4 0 4 Plastid

1 Photosyntheisis 25 33.60 Q 34.74

2 Thylakoid lumen 23 32.38 Q 22.61

3 Photosystem II 25 26.96 Q 22.60 Thylakoid 4 23 25.58 Q 25.59 membrane 5 Plastid stroma 23 16.44 N 12.16

6 Plastoglobule 21 12.89 Q 19.29

7 Photosystem I 22 13.16 Q 14.40

8 Stromule 21 8.11 Q 5.76 NAD(P)H dehydro- 9 22 9.88 Q 10.68 genase complex 1 Tetrapyrrole 23 5.93 P 5.56 0 metabolic process 1 Plastid 21 3.78 N 8.13 1 organization 1 Carotenoid N/A N/A N/A N 4.05 2 metabolic process 1 Plastid ribosome N/A N/A N/A J 4.45 3 1 Plastid 25 1.90 N 4.90 4 chromosome 1 Plastid membrane N/A N/A N/A P 5.61 5

261

Figure 4.3 (Continued) E Average expression of the major cluster

-LIN +LIN /-LIN 0.5 24 24 24 Plastid 1 4 0 4

1 Photosyntheisis 2 Thylakoid lumen 3 Photosystem II Thylakoid 4 membrane 5 Plastid stroma 6 Plastoglobule 7 Photosystem I 8 Stromule NAD(P)H 9 dehydro-genase complex Tetrapyrrole 10 metabolic process Plastid 11 organization Carotenoid 12 metabolic process 13 Plastid ribosome

Plastid 14 chromosome Plastid 15 membrane

262

Figure 4.3 (Continued)

Expressed Enrich- genes / ment in Total all genes in GO Term GO ID genes GO term

-log(p)

Translation

1 P Translation GO:0006412 5.81 32/59

Cytosolic 2 C GO:0022626 0.72 70/181 ribosome

Ribosome 3 P GO:0042254 4.71 8/10 biogenesis

Growth and development 1 P Cell cycle GO:0007049 0.13 39/123 Embryonic 2 P GO:0009790 0.79 123/326 development 3 C Cell wall GO:0005618 0.98 64/123 Cell wall 4 P GO:0042545 0.91 27/61 modification 5 P DNA replication GO:0006260 1.20 21/43 Regulation of 6 P post-embryonic GO:0048580 1.80 57/135 development

263

Figure 4.3 (Continued) G

GO Term Light pattern of expression with the highest enrichment

Pattern +LIN -LIN -log(p) number 0.5 0.5 24 24 Translation 1 4 1 4

1 Translation 32 23.28

Cytosolic 2 32 20.79 ribosome

Ribosome 3 32 11.97 biogenesis

Growth and development 1 Cell cycle N/A N/A N/A Embryonic 2 N/A N/A N/A development 3 Cell wall 31 3.59 Cell wall 4 N/A N/A N/A modification 5 DNA replication 27 5.42 Regulation of 6 post-embryonic N/A N/A N/A development

264

Figure 4.3 (Continued)

GO Term Light cluster of expression with the highest enrichment

Cluster +LIN -LIN -log(p) letter 0.5 0.5 24 24 Translation 1 4 1 4

1 Translation E 18.84

Cytosolic 2 E 17.25 ribosome

Ribosome 3 E 7.09 biogenesis

Growth and development 1 Cell cycle EE 6.03 Embryonic 2 Z 5.98 development 3 Cell wall N/A N/A N/A Cell wall 4 CC 2.78 modification 5 DNA replication DD 4.05 Regulation of 6 post-embryonic GG 4.96 development

265

Figure 4.3 (Continued) I Plastid pattern of expression Plastid cluster of expression GO Term with the highest enrichment with the highest enrichment

Pattern Cluster +LIN/-LIN -log(p) +LIN/-LIN -log(p) number letter 24 24 0 4 0 4 Translation

1 Translation 23 2.15 N 22.07

Cytosolic 2 24 0.34 N 11.60 ribosome

Ribosome 3 23 3.43 N 7.68 biogenesis

Growth and development 1 Cell cycle N/A N/A N/A K 2.32 Embryonic 2 N/A N/A N/A N 0.54 development 3 Cell wall 18 5.81 N/A N/A N/A Cell wall 4 N/A N/A N/A K 5.13 modification 5 DNA replication 28 6.13 K 5.35 Regulation of 6 post-embryonic N/A N/A N/A M 3.48 development

266

Figure 4.3 (Continued) J

Average expression of the major cluster

-LIN +LIN /-LIN

0.5 24 24 24 1 4 Translation 0 4

1 Translation

Cytosolic 2 ribosome

Ribosome 3 biogenesis

Growth and development 1 Cell cycle Embryonic 2 development

3 Cell wall

Cell wall 4 modification DNA 5 replication Regulation of 6 post-embryonic development

267

Figure 4.3 (Continued)

K Expressed Enrich- genes / ment in Total all genes in GO Term GO ID genes GO term

-log(p)

Regulation of gene expression

Regulation of 1 P GO:0045449 0.03 206/558 transcription Cellular protein 2 P GO:0044257 0.52 56/134 catabolic process Ubiquitin ligase 2 C GO:0000151 1.78 19/42 complex 3 P RNA processing GO:0006396 5.16 47/96

Metabolism 1 P Carbon utilization GO:0015976 3.48 11/13 Cellular amino 3 P GO:0006520 6.48 59/113 acid metabolism 4 P Lipid transport GO:0006869 3.32 9/16 Starch metabolic 5 P GO:005982 2.91 21/33 process Monosaccharide 6 P GO:0005996 3.18 22/45 metabolic process

Other cellular components Endoplasmic 1 C GO:0005783 0.70 103/267 reticulum 2 C Mitochondrion GO:0005739 4.09 232/551

268

Figure 4.3 (Continued)

GO Term Light pattern of expression with the highest enrichment

Pattern +LIN -LIN -log(p) number 0.5 0.5 24 24 4 1 4 Regulation of gene 1 expression

Regulation of 1 4 6.05 transcription Cellular protein 2 33 6.38 catabolic process Ubiquitin ligase 2 16 4.66 complex

3 RNA processing 6 5.13

Metabolism 1 Carbon utilization 12 2.98 Cellular amino 3 34 Positive correlation 5.86 acid metabolism 4 Lipid transport N/A N/A N/A Starch metabolic 5 32 3.61 process Monosaccharide 6 metabolic 30 5.37 process

Other cellular components Endoplasmic 1 N/A N/A N/A reticulum 2 Mitochondrion 32 8.69

269

Figure 4.3 (Continued)

GO Term Light cluster of expression with the highest enrichment

Cluster +LIN -LIN -log(p) letter 0.5 0.5 24 24

Regulation of gene 1 4 1 4 expression

Regulation of 1 R 9.43 transcription Cellular protein 2 GG 6.60 catabolic process Ubiquitin ligase 2 GG 1.95 complex 3 RNA processing GG 6.26

Metabolism 1 Carbon utilization Y 3.20 Cellular amino 3 E 5.53 acid metabolism 4 Lipid transport F 6.47 Starch metabolic 5 N/A N/A N/A process Monosaccharide 6 metabolic B 5.11 process

Other cellular components Endoplasmic 1 H 5.00 reticulum 2 Mitochondrion Z 15.19

270

Figure 4.3 (Continued)

Plastid pattern of expression Plastid cluster of expression GO Term with the highest enrichment with the highest enrichment

Pattern +LIN / Cluster +LIN / -log(p) -log(p) number -LIN letter -LIN 24 24 0 4 Regulation of gene 0 4

expression

Regulation of 1 21 0.5 M 4.65 transcription Cellular protein 2 20 3.1 L 3.73 catabolic process Ubiquitin ligase 2 20 7.00 L 5.56 complex

3 RNA processing 21 0.28 O 4.49

Metabolism 1 Carbon utilization 21 7.08 Q 5.56 Cellular amino 3 22 2.66 N 4.44 acid metabolism 4 Lipid transport N/A N/A N/A J 6.06 Starch metabolic 5 23 5.52 N/A N/A N/A process Monosaccharide 6 metabolic 21 3.43 P 2.74 process

Other cellular components Endoplasmic 1 N/A N/A N/A J 1.54 reticulum 2 Mitochondrion 29 1.32 J 4.71

271

Figure 4.3 Summary of biological process and cellular component GO terms enriched in particular expression patterns. (A-N) User defined expression patterns obtained as described in Figure 4.10 and clusters of expression were obtained as described in Figures 4.2 and Figure 4.11. Significant enrichment of 19 GO terms defined as biological processes (P) and 16 GO terms defined as cellular components (C) was determined as described in Figures 4.11 and 4.12. The negative log10 of p-values is used to quantify the GO term enrichment in the user-defined expression patterns, clusters of expression, or in the entire dataset of 7104 genes. To determine the average expression of the major cluster, we performed hierarchical cluster analysis for genes annotated to each GO term to identify the cluster that contains the largest number of genes. Next, the light- and plastid-regulated expression values were averaged at each time point in each condition within the major cluster. Up-regulated (red) and down-regulated (blue) expression is indicated at 0.5 h, 1 h, 4 h, and 24 h following the BR-fluence-rate shift in lincomycin-treated (+LIN) and untreated seedlings (-LIN). Plastid-regulated expression is similarly indicated at 0 h, 4 h, and 24 h relative to the fluence-rate shift. Color intensity is proportional to degree of regulation. Positive correlation describes a similar response to the BR-fluence-rate shift regardless of whether seedlings are treated with lincomycin. For genes that exhibit positive correlation, the correlation coefficient between the expression patterns in lincomycin-treated and untreated seedlings is greater than 0.95. The major clusters and patterns are defined in Figures 4.2E, 4.2F, 4.9A-F, and 4.10. Genes that constitute the major clusters are indicated in Figure 4.5.

272

Figure 4.4 Expressed A Enrich- genes / ment in Total all genes in GO Term GO ID genes GO term

-log(p)

Oxidative Stress and ROS scavenging

1 R Oxidative stress GO:0006979 9.24 102/173 Phenyl propanoid 2 P GO:0009698 4.41 43/82 Metabolic process Glycoside 3 P GO:0016137 3.60 30/48 metabolic process 4 P Cell death GO:0008219 2.73 44/80

Biotic and Abiotic stress 1 R Heat GO:0009408 7.43 57/99 2 R Cold GO:0009409 12.97 128/216 3 R Water deprivation GO:0009414 12.79 97/145 4 R Salt stress GO:0009651 11.24 117/202 5 R Wounding GO:0009611 6.99 67/114 Carbohydrate 6 R GO:0009743 6.37 93/174 stimulus 7 R Metal ion GO:0010038 5.07 44/70

273

Figure 4.4 (Continued)

GO Term Light pattern of expression with the highest enrichment

Pattern +LIN -LIN -log(p) number 0.5 0.5 24 24 1 1 4 Oxidative Stress and 4 ROS scavenging

1 Oxidative stress 34 Positive correlation 3.50 Phenyl propanoid 2 Metabolic N/A N/A N/A process Glycoside 3 metabolic 16 5.93 process 4 Cell death N/A N/A N/A

Biotic and Abiotic stress 1 Heat 6 5.89 2 Cold 34 Positive correlation 7.01 3 Water deprivation 34 Positive correlation 8.20

4 Salt stress 33 5.30 5 Wounding 15 4.23 Carbohydrate 6 15 5.73 stimulus 7 Metal ion 11 2.65

274

Figure 4.4 (Continued)

Light cluster of expression with the highest GO Term enrichment

Cluster +LIN -LIN -log(p) letter 0.5 0.5 24 24

Oxidative Stress and 1 4 1 4 ROS scavenging 1 Oxidative stress I 4.83 Phenyl propanoid 2 Metabolic AA 8.98 process Glycoside 3 metabolic AA 5.99 process 4 Cell death FF 6.38 Biotic and Abiotic stress 1 Heat C 24.10 2 Cold S 4.40 3 Water deprivation H 7.62 4 Salt stress H 7.55 5 Wounding CC 2.49 Carbohydrate 6 D 4.83 stimulus 7 Metal ion D 4.20

275

Figure 4.4 (Continued)

D Plastid pattern of expression Plastid cluster of expression GO Term with the highest enrichment with the highest enrichment

Clust Pattern +LIN / -log(p) er +LIN/-LIN -log(p) number -LIN letter 24 24

Oxidative Stress and 0 4 0 4

ROS scavenging

1 Oxidative stress 21 5.09 O 3.29 Phenyl propanoid 2 Metabolic N/A N/A N/A J 5.61 process Glycoside 3 metabolic 18 6.92 K 5.54 process 4 Cell death N/A N/A N/A M 1.68

Biotic and Abiotic stress 1 Heat 2.50 M 2.74 20 2 Cold 4.28 J 5.73 22 3 Water deprivation 9.21 J 7.80 20 4 Salt stress 4.11 M 6.70 20 5 Wounding 2.57 J 3.17 22 Carbohydrate 6 1.52 M 5.15 stimulus 20 7 Metal ion 1.67 M 2.90 25

276

Figure 4.4 (Continued)

E Average expression of the major cluster

-LIN +LIN /-LIN

Oxidative Stress 0.5 24 24 24 0 4 and ROS 1 4 scavenging Oxidative 1 stress Phenyl propanoid 2 Metabolic process Glycoside 3 metabolic process 4 Cell death Biotic and Abiotic stress 1 Heat 2 Cold Water 3 deprivation 4 Salt stress 5 Wounding Carbohydrat 6 e stimulus 7 Metal ion

277

Figure 4.4 (Continued) Expressed F Enrich- genes / ment in Total all genes in GO Term GO ID genes GO term

-log(p)

Light

1 R UV light GO:0009411 6.33 38/55

2 R High light intensity GO:0009644 9.48 34/41

Red and far-red 3 R GO:0009639 5.21 77/129 light 4 R Blue light GO:0009637 5.88 30/44 5 P Circadian rhythm GO:0007623 5.19 30/43

Hormones Abscisic acid 1 R GO:0009737 11.45 137/245 stimulus

2 R Auxin stimulus GO:0009733 2.52 76/169

Jasmonic acid 3 R GO:0009753 7.90 80/140 stimulus Gibberellin 5 R GO:0009739 7.53 59/99 stimulus

278

Figure 4.4 (Continued)

GO Term Light pattern of expression with the highest enrichment

Pattern +LIN -LIN -log(p) number 0.5 0.5 24 24 1 4 Light 1 4

1 UV light 4 5.76 High light 2 4 4.12 intensity Red and far-red 3 4 5.78 light

4 Blue light 4 3.06

5 Circadian rhythm 33 3.78

Hormones Abscisic acid 1 17 5.61 stimulus

2 Auxin stimulus 3 5.63

Jasmonic acid 3 2 3.96 stimulus Gibberellin 5 11 5.05 stimulus

279

Figure 4.4 (Continued)

Light cluster of expression with the highest GO Term enrichment

Cluster +LIN -LIN -log(p) letter 0.5 0.5 24 24 Light 1 4 1 4

1 UV light AA 15.40 High light 2 C 8.92 intensity Red and far-red 3 R 6.73 light

4 Blue light U 2.86

5 Circadian rhythm BB 3.82

Hormones Abscisic acid 1 H 8.07 stimulus

2 Auxin stimulus X 10.18 Jasmonic acid 3 T 4.97 stimulus Gibberellin 5 H 6.39 stimulus

280

Figure 4.4 (Continued)

I Plastid cluster of Plastid pattern of expression expression with the highest GO Term with the highest enrichment enrichment

Pattern Cluster +LIN / +LIN/ -log(p) -log(p) number letter -LIN -LIN 24 24 0 4 0 4 Light

1 UV light 20 4.45 K 1.52 High light 2 5.05 P 7.02 intensity 21 Red and far- 3 21 7.52 M 5.04 red light

4 Blue light 21 3.40 Q 4.11 Circadian 5 21 4.08 L 3.36 rhythm

Hormones Abscisic acid 1 20 7.21 J 8.04 stimulus

2 Auxin stimulus 21 2.71 M 2.68

Jasmonic acid 3 23 2.73 J 3.82 stimulus Gibberellin 5 20 2.63 L 2.70 stimulus

281

Figure 4.4 (Continued)

Average expression of the major cluster

-LIN +LIN /-LIN

0.5 24 24 24 Light 1 4 0 4

1 UV light High light 2 intensity Red and 3 far-red light 4 Blue light Circadian 5 rhythm

Hormones Abscisic 1 acid stimulus Auxin 2 stimulus Jasmonic 3 acid stimulus Gibberellin 5 stimulus

282

Figure 4.4 Summary of biological process and biological response to stimulus GO terms enriched in particular expression patterns. (A-J) User defined expression patterns obtained as described in Figure 4.10 and clusters of expression were obtained as described in Figures 4.2 and Figure 4.11. Significant enrichment of 4 GO terms defined as biological processes (P) and 16 biological responses to stimulus (R) was determined as described in Figures 4.11 and 4.12. The negative log10 of p-values is used to quantify the GO term enrichment in the user-defined expression patterns, clusters of expression, or in the entire dataset of 7104 genes. To determine the average expression of the major cluster, we performed hierarchical cluster analysis for genes annotated to each GO term to identify the cluster that contains the largest number of genes. Next, the light- and plastid-regulated expression values were averaged at each time point in each condition within the major cluster. Up-regulated (red) and down-regulated (blue) expression is indicated at 0.5 h, 1 h, 4 h, and 24 h following the BR-fluence-rate shift in lincomycin-treated (+LIN) and untreated seedlings (-LIN). Plastid-regulated expression is similarly indicated at 0 h, 4 h, and 24 h relative to the fluence-rate shift. Color intensity is proportional to degree of regulation. Positive correlation is as described in Figure 4.3. The major clusters and patterns are defined in Figures 4.2E,4.2F, 4.9A-F, and 4.10. Genes that constitute the major clusters are from Ruckle and Larkin unpublished data.

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Figure 4.5 Agglomerative hierarchical clustering of significantly regulated genes annotated as contributing to photosynthesis, tetrapyrrole metabolic process, plastid ribosomes, and cytosolic ribosomes. A) Agglomerative hierarchical clustering of significantly regulated genes annotated as contributing photosynthesis and their expression patterns. The criteria for significant regulation are described in Figure 4.2. The photosynthesis-GO term was identified using the criteria described in Figure 4.12. The number of significantly regulated genes from the photosynthesis-GO term/total number of genes in the photosynthesis- GO term is indicated in parentheses. As indicated by the key below each column, squares from the left to the right indicate light-regulated expression at 0.5 h, 1 h, 4 h, and 24 h following the BR-fluence-rate shift in lincomycin-treated seedlings; light- regulated expression at 0.5 h, 1 h, 4 h, and 24 h following the BR-fluence-rate shift in untreated seedlings; and plastid-regulated expression at 0 h, 4 h and 24 h following the BR-fluence-rate shift. Up-regulated expression is indicated with red and down- regulated expression is indicated with blue. Color intensity is proportional to the degree of regulation. B) Agglomerative hierarchical clustering of significantly regulated genes annotated as contributing to tetrapyrrole metabolism with their expression patterns. Genes were clustered and expression is labeled as described in (A). C) Agglomerative hierarchical clustering of significantly regulated genes annotated as contributing to plastid or cytosolic ribosomes with their expression patterns. Genes were clustered and expression is labeled as described in (A).

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Figure 4.6 Analysis of chlorophyll phenotypes caused by T-DNA insertion alleles following de-etiolation. A) Analysis of chlorophyll phenotypes caused by T-DNA insertion alleles of genes that are more highly expressed in lincomycin-treated seedlings than untreated seedlings following a BR-fluence-rate shift. Mutants containing T-DNA insertion alleles of genes that are expressed at least 1.5-fold higher levels in lincomycin- treated relative to untreated seedlings at 1 h following a BR-fluence-rate shift and control lines were grown for 4 d in the dark and then transferred to continuous, broad- -2 -1 spectrum white light that was 125 µmol m s for 24 h. Chlorophyll was extracted, quantified from four biological replicates for each line, and normalized to wild type. Mean chlorophyll levels that were at least twofold greater than the chlorophyll levels of wild type are indicated with a red-dashed line and red bars. Mean chlorophyll levels that were at least twofold less than in wild type are indicated with a blue- dashed line and blue bars. Error bars represent 95% confidence intervals. T-DNA alleles were named using the arbitrary number assigned to each gene (Ruckle and Larkin unpublished data) and the last two numbers of Salk accession code or the last 3 digits of the SAIL accession code. For example, the T-DNA alleles of gene number 1 (At5g24120) are SAIL_1232_H11 and Salk_141383. B) Analysis of chlorophyll phenotypes caused by T-DNA insertion alleles of genes that are similarly expressed in lincomycin-treated and untreated seedlings following a BR-fluence-rate shift. The de-etiolation of mutants and the extraction and quantification of chlorophyll were performed as described in (A).

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Figure 4.7 De-etiolation efficiencies of end mutants in various fluence rates. A) De-etiolation efficiencies of end mutants in three different fluence rates. The end mutants, gun1-101, and wild type (Col-0) were grown in the dark for 4 d and then -2 irradiated with broad-spectrum white light at fluence rates of 15, 100, or 300 µmol m -1 s for 24 h. Chlorophyll was extracted from four biological replicates for each line in each condition. Chlorophyll levels of wild type (Col-0) are indicated with blue bars. Chlorophyll levels of mutants are indicated with red bars and green bars. Error bars represent 95% confidence intervals. -2 -1 B) De-etiolation rates of end mutants in 1 µmol m s BR light. The end mutants, gun1-101, and wild type (Col-0) were grown in the dark for 4 d and then transferred -2 -1 to 1µmol m s BR light. Chlorophyll was extracted from four biological replicates for each line at 0, 6, 12, and 24 h from wild type (Col-0; blue curves) and the end mutants (red and green curves indicate distinct alleles). A Col-0 control was grown on the same plate for each end mutant. Error bars represent 95% confidence intervals.

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Figure 4.8 Expression of Lhcb1 and RbcS following a fluence-rate shift. A) Expression of Lhcb1 and RbcS in lincomycin-treated seedlings after a fluence-rate shift. Seedlings were grown in the presence of lincomycin (+Lin) or in the absence of lincomycin (-Lin) as described in Figure 4.1. RNA was extracted and both Lhcb1 and RbcS mRNA levels were quantified with RNA blot hybridizations that utilized 2.5 µg of total RNA. Four biological replicates from each time point were quantified and normalized to total RNA stained with methylene blue. Numbers below each lane indicate the amount of hybridized RNA as a percentage of hybridized RNA in -2 -1 untreated seedlings after 24 h in 60 µmol m s BR light. B) Expression of Lhcb1 and RbcS in seedlings not treated with lincomycin following a fluence-rate shift. Seedlings were grown in the absence of lincomycin (-Lin) and the levels of Lhcb1mRNA were quantified as in (A).

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Figure 4.9 Agglomerative clustering of early time points (i.e., 0.5 h and 1 h) and late time points (i.e., 4 h and 24 h). A) Agglomerative hierarchical clustering of the 7104 active gene sets based on their regulation by light at 0.5 h and 1 h in lincomycin-treated and untreated seedlings. Up-regulated expression is indicated with red and down-regulated expression is indicated with blue. Color intensity is proportional to the degree of regulation. B) Agglomerative hierarchical clustering of the 7104 active gene sets based on their regulation by the light at 4 h and 24 h in lincomycin-treated and untreated seedlings. Data are colored as in (A). C-F) The average gene expression level (log2) and number of genes represented by each of the clusters from Figure 4.2E (C), Figure 4.2F (D), Figure 4.9A (E), and Figure 4.2B (F).

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Figure 4.10 Distribution of user-defined expression patterns among light- and plastid-regulated genes. A) Expression patterns of light- and plastid-regulated genes. We defined 33 models of expression that arise from the interactions between light and plastid signaling. Gene expression was described in four possible ways: induced (red); repressed (blue); induced or repressed (dark gray); no significant change (light gray); data were not considered (white). Genes were considered induced (red) or repressed (blue) if the change in H[SUHVVLRQZDVJUHDWHUWKDQWZRIROG S 7KHH[SUHVVLRQ patterns were: pattern 1, genes are induced or repressed; patterns 2-5, early-light- responsive genes (i.e., genes are regulated by light only at 0.5 h and 1 h); patterns 6- 9, transiently responsive genes (i.e., genes that are regulated by light only at 4 h); patterns 10-13, light induction or repression at 4 h is maintained at 24 h in lincomycin-treated and/or untreated seedlings; patterns 14-17, late-responsive genes (i.e., genes that are regulated by light only at 24 h); patterns 18-25, plastid- responsive genes (i.e., genes that are regulated in lincomycin-treated seedlings at 0 h, 4 h, or at 24 h); patterns 26-27, genes that are oppositely regulated by light at the same time point in lincomycin-treated and untreated seedlings; patterns 28-29, genes that are oppositely regulated in lincomycin-treated and untreated seedlings 24 h after the BR-fluence-rate shift; patterns 30-31, genes that are either induced or repressed by light in lincomycin-treated but not in untreated seedlings; patterns 32-33 genes that are induced or repressed by light in untreated seedlings but not in lincomycin- treated seedlings. B) Number of genes in each of the expression models described in (A). The 33 expression patterns from (A) and two additional patterns, 34 and 35, are shown. Pattern 34 (a.k.a., positive correlation) describes genes that exhibit a similar response to the BR-fluence-rate shift regardless of whether seedlings are treated with lincomycin. For these genes, the correlation coefficient between the expression patterns in lincomycin-treated and untreated seedlings is greater than 0.95. Pattern 35 (a.k.a., negative correlation) describes genes that have an opposite response to the BR-fluence-rate shift in lincomycin-treated and untreated seedlings. For these genes, the correlation coefficient between the expression pattern in lincomycin- treated and untreated seedlings is less than -0.95.

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Figure 4.11 Agglomerative hierarchical clustering of significantly enriched GO terms with expression clusters.

A) Agglomerative hierarchical clustering of GO terms defined as a biological process with expression clusters. We analyzed 205 GO terms and judged them to be enriched in at least one of the expression cluVWHUVLISDIWHUD:HVWIDOO-Young correction. GO terms were included if they were enriched in at least one expression SDWWHUQDQGS7KHQHJDWLYHORJ10 of the uncorrected p-value for each pattern was clustered. Clusters of GO terms are shown as a dendrogram at the left of the cluster matrix. Summary GO terms are shown at the right of the cluster matrix. These summary terms contain a representative group of genes from multiple GO categories. For example, tetrapyrrole metabolism is a summary term that contains a representative group of genes from the following GO categories: tetrapyrrole metabolic process, 0033013; porphyrin metabolic process, 0006778; chlorophyll metabolic process, 0015994; cofactor metabolic process, 0051186; tetrapyrrole biosynthetic process, 0033014; porphyrin biosynthetic process, 0006779; cofactor biosynthetic process, 0051188; chlorophyll biosynthetic process, 0015995. Expression clusters from Figure 4.2 and Figure 4.9 are shown as a dendrogram at the top of the cluster matrix. Names of expression clusters from Figure 4.2 and Figure 4.9 that correspond to each branch of the dendrogram are indicated at the bottom of the cluster matrix. The letter P next to the cluster letter indicates regulation by plastid dysfunction. No letter P next to the cluster letter indicates regulation by the BR fluence-rate-shift. The branch of the dendogram labeled 1 contains all of the significantly regulated genes in the dataset. The intensity of red color indicates degree of enrichment. Co-regulated GO terms are indicated with dashed boxes. B) Agglomerative hierarchical clustering of GO terms defined as cellular components with expression clusters. Eighty-eight GO terms defined as cellular components were enriched as described in (A). The labeling of this heatmap is as described for (A).

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Figure 4.12 Clustering of GO terms enriched with user-defined expression patterns. A) Agglomerative hierarchical clustering of GO terms defined as biological processes with expression patterns. One hundred ninety-eight active GO terms assigned as a ELRORJLFDOSURFHVVZHUHFRQVLGHUHGHQULFKHGLISDIWHUD:HVWIDOO-Young correction of at least one of the expression clusters. GO terms were included if they were enriched in at least one exSUHVVLRQSDWWHUQDQGS The negative log10 of the uncorrected p-values for each pattern was clustered. These one hundred ninety-eight GO clusters are shown as a dendrogram at the left of the cluster matrix. Summary terms that represent a cluster of GO terms are shown at the right of the cluster matrix and are as described in Figure 4.10A. User-defined expression patterns from Figure 4.10 are shown as a dendrogram at the top of the cluster matrix. User-defined expression pattern numbers from Figure 4.10 that correspond to each branch of the dendrogram are indicated at the bottom of the cluster matrix. The letter P next to the cluster number indicates regulation by the lincomycin treatment. No letter P next to the cluster letter indicates regulation by the BR fluence-rate-shift. The intensity of red color indicates degree of enrichment. Clusters of patterns are indicated with dashed boxes. B) Agglomerative hierarchical clustering of GO terms defined as cellular components with expression patterns. Eighty-seven GO terms assigned as cellular components were enriched as described in (A). Labeling of this heatmap is as described in (A).

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Figure 4.13 Analysis of chlorophyll phenotypes caused by mutant alleles of genes that encode light-signaling factors. De-etiolation was performed and relative chlorophyll levels were quantified for the indicated light-signaling mutants as described in Figure 4.6. These alleles were previously generated using either EMS or T-DNA insertional mutagenesis. Names for T-DNA insertion alleles were formed from the gene name and the last two numbers of the Salk accession code or the last three digits of the SAIL accession code. For example, the T-DNA insertion allele of AtPP7 that is derived from Salk_089764 is named Atpp7-64. cry1-92, phyA-75, phyB-35, phot1-58, phot2-75, nph3-39, cry3-30, and hy5-51 were previously characterized (Ruckle et al., 2007). The remaining alleles derived from T-DNA insertional mutagenesis are described by Ruckle and Larkin unpublished data. The alleles derived from EMS mutagenesis were cop1-4 (Deng et al., 1992; Deng and Quail, 1992), det1-1 (Chory et al., 1989), and phot1-5, which was formerly known as nph1-5 (Liscum and Briggs, 1995; Huala et al., 1997). Error bars represent 95% confidence intervals.

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Table4.1 Light Plastid Insertion effect on No. AGI Code Induction T-DNA Line(s) regulation transrcipt Ratio SAIL_1232_H11 Null 1 At5g24120 3.92 -7.69 SALK_141383 Null 2 At3g56290 3.16 -4.35 SALK_053531 Strong Knockdown 3 At2g30040 2.79 -2.70 SAIL_1175_F12 Null 5 At5g08050 2.35 -3.22 SALK_048774 Null 6 At5g24660 2.35 2.48 SALK_031648 Strong Knockdown 7 At3g17040 2.19 -3.56 SALK_079285 Knockdown SALK_084849 Knockdown 8 At1g44000 2.06 -2.69 SAIL_682_D01 Knockdown 10 At2g24540 2.00 -1.67 SAIL_897_A11 Weak Knowckdown

11 At5g35970 1.99 -4.63 SALK_149757 Strong Knockdown 13 At4g11360 1.97 -1.98 SALK_094834 Null SAIL_210_E05 Null 14 At5g14970 1.94 -2.32 SALK_036830 Null SAIL_1256_F11 Null 15 At5g58650 1.93 -2.47 SAIL_129_H05 Null

16 At2g41660 1.92 -1.21 SALK_076560 Null

SALK_011143 Strong Knockdown 17 At5g13770 1.88 -4.38 SALK_051012 Strong Knockdown 19 At2g16365 1.86 -1.43 SALK_024229 Null

20 At5g52780 1.84 -1.85 SALK_143426 Null SALK_108235 Null 21 At3g54990 1.84 1.11 SALK_135576 Null 22 At5g62430 1.78 -2.33 SAIL_381_B11 Null

SAIL_70_F03 Knockdown 23 At3g02380 1.78 -9.32 SAIL_265_D06 Knockdown 25 At1g43160 1.75 -7.30 SAIL_1225_G09 Null 28 At5g52250 1.72 5.80 SALK_060638 Null 29 At1g04770 1.71 1.68 SALK_091618 Null 35 At2g33250 1.57 -1.19 SALK_033583 Null 36 At2g46340 1.56 -1.27 SALK_023840 Strong Knockdown 38 At4g28740 1.52 -1.92 SALK_133844 Null

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Table4.1 (Continued)

No. AGI Code Name/Description

a 1 At5g24120 SIGE, AtSIG5, SIGMA FACTOR 5 2 At3g56290 Expressed Protein MAPKKK14, MITOGEN-ACTIVATED PROTEIN KINASE KINASE 3 At2g30040 KINASE 14 5 At5g08050 Expressed Protein 6 At5g24660 LSU2, RESPONSE TO LOW SULFUR2 c 7 At3g17040 HCF107, HIGH CHLOROPHYLL FLUORESCENT 107 , tetratricopeptide repeat-containing protein e 8 At1g44000 SGR-Like, STAY GREEN LIKE, Subfamily protein AFR, ATTENUATED FAR-RED RESPONSE, Kelch repeat- 10 At2g24540 g containing F-box family protein 11 At5g35970 Putitive DNA binding protein, DEAD-like helicase domain 13 At4g11360 RHA1b, RING-H2 FINGER A1B 14 At5g14970 Expressed Protein i 15 At5g58650 PSY1, PLANT PEPTIDE CONTAINING SULFATED TYROSINE1 j 16 At2g41660 MIZ1, MIZU-KUSSEI 1 17 At5g13770 Pentatricopeptide (PPR) repeat-containing protein 19 At2g16365 F-box family protein 20 At5g52780 Expressed Protein l 21 At3g54990 SMZ, SCHLAFMUTZE, AP2 domain transcription factor CDF1, CYCLING DOF FACTOR 1, Dof-type zinc finger domain- 22 At5g62430 containing protein COL2, CONSTANS-LIKE 2, zinc-finger protein, CCT Domain, B- 23 At3g02380 box Domain, Transcription factor 25 At1g43160 RAP2.6, RELATED TO AP2 6, AP2 domain transcription factor

28 At5g52250 Transducin family protein, WD-40 repeat family protein, COP1-like 29 At1g04770 Male sterility MS5 family protein, Tetratricopeptide TPR domain 35 At2g33250 Expressed Protein SPA1, SUPPRESSOR OF PHYA, serine/ threonine kinase-like 36 At2g46340 q motif, WD-repeat domain s 38 At4g28740 Similar to LPA1, LOW PSII ACCUMULATION1

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Table4.1 (Continued)

No. AGI Code Biological function, process and location

ISS,TAS b 1 At5g24120 Transcription factor activity , Chloroplast 2 At3g56290 Unknown ISS 3 At2g30040 Kinase activity b 5 At5g08050 Unknown, Thylakoid membrane 6 At5g24660 Unknown c c RNA processing , Regulation of translation , Chloroplast 7 At3g17040 d membrane 8 At1g44000 Unknown f 10 At2g24540 Far-red light phototransduction ISS g 11 At5g35970 DNA binding / Chloroplast ISS h 13 At4g11360 Protein binding , E3 ligase activity 14 At5g14970 Unknown i 15 At5g58650 Cell proliferation and expansion j 16 At2g41660 Hydrotropism IEA 17 At5g13770 Chloroplast 19 At2g16365 Unknown k 20 At5g52780 Chloroplast thylakoid membrane ISS l IC 21 At3g54990 Transcription factor activity , Floral repression , Nucleus ISS m Transcription factor activity , DNA binding , Protein 22 At5g62430 m m m binding , Regulation of flowering time , Nucleus ISS 23 At3g02380 Transcription factor activity ISS,TAS ISS 25 At1g43160 Transcription factor activity , Nucleus ISS Heterotrimeric G-protein complex , CUL4 RING ubiquitin 28 At5g52250 ISS ligase complex 29 At1g04770 Unknown IEA 35 At2g33250 Chloroplast q ISS r 36 At2g46340 Photomorphogenesis , Signal transducer activity , Nucleus s,t 38 At4g28740 Unknown, Chloroplast

315

Table 4.1 Genes that exhibit enhanced light-induced expression in lincomycin- treated seedlings and their publicly available T-DNA alleles. Twenty five genes are ranked by their light induction. Light induction is defined as the ratio of light-induced expression in lincomycin-treated seedlings to light-induced expression in untreated seedlings at 1 h following the BR-fluence-rate shift. Plastid regulation is represented as the ratio of induced or repressed (-) expression in lincomycin-treated seedlings to expression in untreated seedlings at 0 h relative to the BR-fluence-rate shift. Gene names and descriptions are based on available literature or on TIGR gene annotation records. Biological function, process, and locations are based on the current literature or the gene ontology with the following evidence codes: ISS, Inferred from sequence or structural similarity; TAS, Traceable author statement; IEA, Inferred from electronic annotation; IC, Inferred by curator. The publicly available T-DNA insertion alleles used in this study are listed. For each homozygous line, the RNA phenotype caused by the particular T-DNA insertion allele was determined by RT-PCR (Ruckle and Larkin Unpublished datat). References are designated as follows: a (Kanamaru and Tanaka, 2004); b (Friso et al., 2004); c (Felder et al., 2001); d (Sane et al., 2005); e (Barry et al., 2008); f (Harmon and Kay, 2003); g (Kleffmann et al., 2004); h (Stone et al., 2005); i (Amano et al., 2007); j (Kobayashi et al., 2007); k (Peltier et al., 2004); l (Schmid et al., 2003); m (Imaizumi et al., 2005); n (He et al., 2004); o (Zhu et al., 2010); p (Hoecker et al., 1998); q (Fankhauser et al., 1999); r (Lariguet et al., 2006); s (Zybailov et al., 2008); t (Peng et al., 2006).

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Table4.2

Arabidopsis Genome Light Initiative (AGI) Induction Plastid No. Identifier Ratio regulation

4 At1g14345 2.45 -5.92

9 At1g11380 2.06 -1.80 12 At3g49580 1.98 -1.67

18 At2g30520 1.87 -1.21

24 At5g47610 1.76 -3.56

26 At2g42540 1.73 5.80

27 At1g12250 1.73 -2.48 30 At1g16720 1.68 -2.19

31 At1g42550 1.65 -3.08

32 At1g79270 1.65 -1.68

33 At1g32080 1.64 -6.56

34 At5g49330 1.57 1.38

37 At2g02950 1.52 1.17

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Table4.2 (Continued) Arabidopsis Genome Initiative (AGI) Biological function, process No. Identifier Name/Description and location

IEA 4 At1g14345 Aldo/keto reductase Oxidation reduction , domain Chloroplast thylakoid a membrane 9 At1g11380 Cys-rich domain Unknown 12 At3g49580 LSU1, RESPONSE TO Unknown LOW SULFUR 1 b ISS 18 At2g30520 RPT2, ROOT Phototropism , Nucleus b PHOTOTROPISM 2 ISS 24 At5g47610 Zinc finger (C3HC4-type Protein Binding RING finger) c 26 At2g42540 COR15A, COLD- Cold acclimation , REGULATED 15A d Chloroplast a 27 At1g12250 Expressed Protein Chloroplast thylakoid e 30 At1g16720 HCF173, HIGH Translational initiation , CHLOROPHYLL e e Photosystem II assembly , FLUORESCENCE 173 f Chloroplast g 31 At1g42550 PMI1, PLASTID Chloroplast relocation , MOVEMENT h g Plasma membrane IMPAIRED1 32 At1g79270 ECT8, Unknown EVOLUTIONARILY CONSERVED C- TERMINAL REGION 8 i 33 At1g32080 LrgB-like domain Chloroplast inner membrane protein 34 At5g49330 ATMYB111, MYB Transcription factor ISS DOMAIN PROTEIN 111 activity j 37 At2g02950 PKS1, Phototropism , Red/far red PHYTOCHROME k k light signaling , Ctytoplasm , KINASE SUBSTRATE 1 j Plasma membrane

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Table 4.2 Genes that exhibit enhanced light-induced exprssion in lincomycin-treated seedlings and that do not have publicly available T-DNA alleles.

Thriteen genes are ranked by their light induction. Light induction is defined as the ratio of light-induced expression in lincomycin-treated seedlings to light-induced expression in untreated seedlings at 1 h folling the BR-fluence ±rate shift. Plasted regulation is represented as a ratio of induced or repressed (-) expression in lincomycin-treated seedling to the expression in untreated seedlings at 0 h relative to the BR-fluence-rate shift. Gene names and descriptions are based on available literature or on TIGR gene annotation records Biological function, process, and location are based on current literature or the Gene Ontology. References : a (Peltier et al., 2004); b (Sakai et al., 2000); c (Lin et al., 1992); d (Kleffmann et al., 2004); e (Schult et al., 2007); f (Zybailov et al., 2008); g (DeBlasio et al., 2005); h (Nühse et al., 2003); i (Ferro et al., 2003); j (Lariguet et al., 2006); k (Fankhauser et al., 1999).

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Table 4.3

Biological function, Light Plastid Name / process, and No. AGI code Induction regulation Description location

67 At5g59220 1.20 -0.10 HAI1, IEA HIGHLY Chloroplast , ABA- protein serine / INDUCED threonine PP2C phosphatase ISS GENE 1 activity , a ABA signaling

70 At5g23730 0.83 -0.64 WD-40 CUL4 RING repeat ubiquitin ligase family ISS protein complex

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Table 4.3 (Continued)

Transcript phenotype in the No. AGI code T-DNA line(s) homozygote

67 At5g59220 SALK_142672 Null SAIL_520_H12 Upregulated

70 At5g23730 SALK_015765 Knockdown

Table 4.3 Alleles of genes that exhibit similar light-induced expression in lincomycin- treated and untreated seedlings that cause end phenotypes. The light induction and the plastid regulation were calculated as described in Table 4.1. Gene names and descriptions are based on available literature or on TIGR gene annotation records. Biological function, process, and location are based on the current literature or the gene ontology with evidence codes as described in Table 4.1. Reference: a (Fujita et al., 2009).

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Table 4.4

No. AGI Code Regulators of gene expression

E, 1 E, 1 M, 2 SS, 3 1 At5g24120 Far-red light , Red light , UV-B , Plastid signals , E, 4 Blue light 2 At3g56290 UV-BM ,2 E, 5 E, 1 E, 1 E, 6 3 At2g30040 Nitrogen , Far-red light , Red light , Brassinosteroid , E, 6 E, 7 Auxin , Cold 5 At5g08050 NA M, L, 8 6 At5g24660 Sulfur E, 9 7 At3g17040 Red light 8 At1g44000 NA M, 10 M, 10 M, 10 11 E, 12 10 At2g24540 Salt , Osmotic , Cold , Circadian clock , UV-B M,13 E,12 11 At5g35970 High light , UV-B E, 14 L, 15, 13 At4g11360 Phenylglycosides , Sucrose starvation Immune 16 E, 17 E, 18 response to flg22E , Sucrose addition , Chitin 14 At5g14970 NA 15 At5g58650 NA E, 12 16 At2g41660 UV-B 17 At5g13770 NA 19 At2g16365 NA 20 At5g52780 NA 19 E, 5 21 At3g54990 Photoperiod , Nitrogen 20 22 At5g62430 Circadian clock SS, 21 E, 1 E, 1 23 At3g02380 Reactive oxygen , Far-red light , Red light E, 22 E, 23 E, 6 E, 6 25 At1g43160 Wounding , Intense light , Brassinosteroid , Auxin , L, 24 E, 25 M, 26 E, 27 Drought , Cytokinin , Cold , Blue light , Far-red E, 28 E, 28 light , Red light E, 1 E, 1 E, 29 M, 13 28 At5g52250 Far-red light , Red light , UV-B , High light M, L, 8 29 At1g04770 Sulfur 35 At2g33250 NA E, 1 E, 1 E, 30 E, 31 36 At2g46340 Far-red light , Red light , Blue light , Green light 38 At4g28740 NA

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Table 4.4 The diverse regulators of genes that exhibit enhanced light-induced expression in lincomycin-treated seedlings. Only signals that both regulate the expression of these genes and are indicated in the text (including embedded tables) of the indicated references were considered. Large datasets provided in the supplemental material online were not considered. The kinetics of regulated expression is also provided: E, Early, 0-2 h; M, Mid, 2-8 h; L, Late, 8-24 h; SS, Steady State. References: 1 (Khanna et al., 2006); 2 (Brown et al., 2005); 3 (Ankele et al., 2007); 4 (Onda et al., 2008); 5 (Scheible et al., 2004); 6 (Goda et al., 2004); 7 (Lee et al., 2005); 8 (Maruyama-Nakashita et al., 2005); 9 (Monte et al., 2004); 10 (Kreps et al., 2002); 11 (Harmon and Kay, 2003); 12 (Oravecz et al., 2006); 13 (Kleine et al., 2007); 14 (Guan and Nothnagel, 2004); 15 (Contento et al., 2004); 16 (Navarro et al., 2004); 17 (Osuna et al., 2007); 18 (Libault et al., 2007); 19 (Schmid et al., 2003); 20 (Imaizumi et al., 2005); 21 (Charron et al., 2008); 22 (Yan et al., 2007); 23 (Rossel et al., 2007); 24 (Catala et al., 2007); 25 (Rashotte et al., 2003); 26 (Fowler and Thomashow, 2002); 27 (Folta et al., 2003); 28 (Tepperman et al., 2004); 29 (Ulm et al., 2004); 30 (Fittinghoff et al., 2006); 31 (Dhingra et al., 2006).

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