interactions between plant circadian clocks and solute transport

Journal of Experimental Botany, Vol. 62, No. 7, pp. 2333–2348, 2011doi:10.1093/jxb/err040 Advance Access publication 4 March, 2011

REVIEW PAPER

Interactions between plant circadian clocks and solutetransport

Michael J. Haydon, Laura J. Bell and Alex A. R. Webb*

Department of Plant Sciences, University of Cambridge, Downing Street, Cambridge CB2 3EA, UK

* To whom correspondence should be addressed. E-mail: [email protected]

Received 28 September 2010; Revised 27 January 2011; Accepted 28 January 2011

Abstract

The Earth’s rotation and its orbit around the Sun leads to continual changes in the environment. Many organisms,

including plants and animals, have evolved circadian clocks that anticipate these changes in light, temperature, and

seasons in order to optimize growth and physiology. Circadian timing is thought to derive from a molecular

oscillator that is present in every plant cell. A central aspect of the circadian oscillator is the presence of

transcription translation loops (TTLs) that provide negative feedback to generate circadian rhythms. This review

examines the evidence that the 24 h circadian clocks of plants regulate the fluxes of solutes and how changes in

solute concentrations can also provide feedback to modulate the behaviour of the molecular oscillator. It highlights

recent advances that demonstrate interactions between components of TTLs and regulation of solute concentrationand transport. How rhythmic control of water fluxes, ions such as K+, metabolic solutes such as sucrose,

micronutrients, and signalling molecules, including Ca2+, might contribute to optimizing the physiology of the plant

is discussed.

Key words: Calcium, circadian, micronutrients, nitrogen, transport, sugar, starch, water.

The circadian clock

The circadian clock is a 24 h timekeeper that regulates at

least 30% of the Arabidopsis transcriptome (Michael et al.,

2008) and also photosynthesis, metabolism, growth, and

stress signalling (Harmer, 2009). For these reasons there are

rhythmic fluxes of water and solutes, which depend on

regulated transporters to permit movement across mem-

branes throughout the plant. This might occur through

regulation of transcripts, post-translational regulation ofprotein abundance, or regulation of transport activity. For

example, there are circadian oscillations in the abundance

of transcripts encoding auxin transporters (Table 1;

Covington and Harmer, 2007). A role for transport in

circadian behaviour cannot be assumed based on transcrip-

tional regulation of transporters alone (Covington and

Harmer, 2007); it is also necessary to consider whether there

are oscillations in the fluxes and concentrations of thetransported solutes. A solute is any dissolved substance, but

the focus here is on a selection of physiologically important

solutes that fulfil diverse roles in energy storage, nutrition,

signalling, and osmotic potential, and for which there is

evidence of regulation of solute flux by the circadian clock.

These include sucrose (Suc), essential nutrients such as

nitrogen (N) and sulphur (S), potassium ions (K+), micro-

nutrients, and signalling molecules such as calcium (Ca2+).

In C3 and C4 plants the day/night cycle of temperatureand light drives rhythms in solute concentration and flux

due to the rhythmic production of sugars by photosynthesis

and the daily changes in turgor resulting from transpiration

during the day. To cope with these daily rhythms, plants,

along with cyanobacteria, fungi, mammals, and most other

forms of life have developed internal timing mechanisms to

anticipate the light/dark cycle. These timing mechanisms are

called the circadian clock. Circadian clocks are character-ized by generation of rhythms that approximate to 24 h, the

Abbreviations: ATP, adenosine triphosphate; CAM, crassulacean acid metabolism; [Ca2+]cyt, cytosolic free Ca2+; DD, continuous dark; LL, continuous light; TTL,transcription translation loop; ROS, reactive oxygen species.ª The Author [2011]. Published by Oxford University Press [on behalf of the Society for Experimental Biology]. All rights reserved.For Permissions, please e-mail: [email protected]

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Table 1. Circadian regulated transcripts for transporters in A. thaliana

Compiled from published lists of circadian regulated transcripts from Dodd et al. (2007) and Covington and Harmer (2007).

AGI locus Protein description [The ArabidopsisInfomation Resource (TAIR)10]

Gene name

Carbohydrate transport

{EM#}AT1G11260 Hexose/H+ symporter STP1

{EM#}AT5G26340 Hexose/H+ symporter, high-affinity STP13

{EM#}AT1G77210 Monosaccharide transporter STP14

{EM#}AT3G51490 Monosaccharide transporter, vacuole

{EM#}AT5G42420 Nucleotide-sugar transporter family protein




{EM#}AT1G71890 Sucrose transporter SUC5

{EM#}AT1G22710 Sucrose transporter, high-affinity, phloem SUC2

{EM#}AT5G23660 Sugar efflux (SWEET) family, nodulin MtN3 family SWEET12










{EM#}AT4G23010 UDP-galactose transporter

{EM#}AT5G47560 Malate/fumarate transporter, vacuole TDT

{EM#}AT5G64290 Dicarboxylate transporter DIT2.1

{EM#}AT5G46110 Triose phosphate translocator, chloroplast TPT/APE2

{EM#}AT3G01550 Phosphoenolpyruvate/phosphate translocator PPT2

{EM#}AT2G43330 Myo-inositol exporter, vacuole INT1

Nitrogen transport

{EM#}AT1G31820 Amino acid permease family

{EM#}AT2G01170 Amino acid transporter family BAT1

{EM#}AT5G65990 Amino acid transporter family



{EM#}AT3G11900 Aromatic/neutral amino acid transporter ANT1

{EM#}AT5G04770 Cationic amino acid transporter CAT6

{EM#}AT1G58360 Neutral amino acid transporter AAP1

{EM#}AT5G19500 Tryptophan/tyrosine permease

{EM#}AT3G24290 Ammonium transporter AMT1;5

{EM#}AT1G64780 Ammonium transporter, high-affinity AMT1;2

{EM#}AT4G13510 Ammonium transporter, plasma membrane AMT1;1

{EM#}AT1G69870 Nitrate transporter, low-affinity phloem NRT1.7

{EM#}AT3G45650 Nitrate efflux transporter, plasma membrane NAXT1

{EM#}AT1G12110 Nitrate transporter, dual-affinity nitrate uptake NRT1.1/CHL1

{EM#}AT5G14570 Nitrate transporter, vacuolar membrane NRT2.7

{EM#}AT1G79410 Organic cation/carnitine transporter OCT5

{EM#}AT3G15380 Choline transporter family, plasma membrane

{EM#}AT1G26440 Ureide permease UPS5

{EM#}AT4G12030 Bile acid/sodium symporter BAT5

{EM#}AT2G26900 Bile acid/sodium symporter

Phosphate/sulphate transport

{EM#}AT3G26570 Phosphate transporter, low-affinity PHT2;1

{EM#}AT2G29650 Inorganic phosphate transporter, thylakoid membrane. PHT4;1

{EM#}AT3G51895 Sulphate transporter SULTR3;1

Water transport

{EM#}AT4G23400 Plasma membrane intrinsic protein (PIP1 subfamily) PIP1D

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Table 1. Continued


Gene name

{EM#}AT4G00430 Plasma membrane intrinsic protein (PIP1 subfamily) PIP1E

{EM#}AT2G45960 Plasma membrane intrinsic protein (PIP1 subfamily) PIP1B

{EM#}AT3G53420 Plasma membrane intrinsic protein (PIP2 subfamily) PIP2A

{EM#}AT4G01470 Tonoplast intrinsic protein (TIP), water, and urea channel TIP1;3

Ion transport

{EM#}AT3G27170 Anion channel protein family CLC-B

{EM#}AT5G57110 Ca2+-ATPase, plasma membrane ACA8

{EM#}AT2G41560 Ca2+-ATPase, calmodulin-regulated, vacuolar ACA4

{EM#}AT5G54250 Cyclic nucleotide gated channel family CNGC4

{EM#}AT2G46450 Cyclic nucleotide gated channel family CNGC12

{EM#}AT5G15410 Cyclic nucleotide-gated channel family CNGC2/DND1

{EM#}AT1G01790 Potassium efflux antiporter KEA1

{EM#}AT4G00630 Potassium transporter family



{EM#}AT2G35060 Potassium transporter, K uptake KUP11




{EM#}AT4G22200 Shaker family K channel, photosynthate- and light-dependent

inward rectifying potassium channel

AKT2

{EM#}AT5G27150 Sodium/proton antiporter, vacuolar NHX1

{EM#}AT2G29110 Ligand-gated ion channel subunit family GLR2.8

{EM#}AT4G35290 Putative glutamate receptor like-protein, putative ligand-gated

ion channel subunit family

GLUR2

{EM#}AT2G17260 Glutamate receptor GLR2

{EM#}AT3G49920 Voltage-dependent anion channel VDAC5

Micronutrient transport

{EM#}AT1G59870 ATP binding cassette (ABC) transporter, plasma membrane

Cd exporter

PDR8/PEN3

{EM#}AT5G23760 Copper transporter family

{EM#}AT3G46900 Copper transporter family COPT2

{EM#}AT5G59030 Copper transporter family COPT1

{EM#}AT1G51610 Cation efflux family protein



{EM#}AT5G26820 Iron-regulated (IREG) transporter IREG3/MAR1

{EM#}AT2G39450 Metal tolerance protein (MTP) family,

Mn transporter, Golgi

MTP11

{EM#}AT4G37270 Metal transporting P-type ATPase (HMA),

chloroplast Cu transporter

HMA1

{EM#}AT4G33520 Metal transporting P-type ATPase (HMA),

chloroplast Cu transporter

PAA1/HMA6

{EM#}AT3G17650 Metal-nicotianamine transporter YSL5



{EM#}AT2G25680 Molybdate transporter, high-affinity MOT1

{EM#}AT5G67330 NRAMP metal transporter family NRAMP4

{EM#}AT3G25190 Vacuolar iron transporter (VIT) family protein

{EM#}AT3G43630 Vacuolar iron transporter (VIT) family protein

{EM#}AT3G20870 ZIP metal ion transporter family ZTP29

{EM#}AT1G55910 ZIP metal ion transporter family ZIP11

Auxin transport

{EM#}AT3G53480 ATP binding cassette (ABC) transporter, plasma

membrane IBA efflux

PDR9

{EM#}AT2G47000 ABC transporter, putative auxin transporter MDR4/ABCB4

{EM#}AT2G36910 ABC transporter, putative auxin efflux protein ABCB1

Circadian regulation of solute transport | 2335D

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Table 1. Continued


Gene name

{EM#}AT1G76530 Auxin efflux carrier family protein

{EM#}AT1G70940 Auxin efflux protein PIN3

{EM#}AT1G23080 Auxin efflux protein PIN7

Proton transport

{EM#}AT2G16510 Plasma membrane ATPase, F0/V0 complex, subunit C protein

{EM#}AT1G64200 Vacuolar H+-ATPase subunit E isoform 3 VHA-E3

{EM#}AT1G15690 H+-translocating inorganic pyrophosphatase (H+-PPase), vacuole AVP1

Nucleotide transport

{EM#}AT3G10960 Adenine-guanine transporter AZG1

{EM#}AT1G15500 ATP/ADP antiporter NTT2

{EM#}AT2G47490 NAD+ transporter, chloroplast NDT1

{EM#}AT1G57990 Purine transporter PUP18

{EM#}AT1G19770 Purine transporter PUP14

{EM#}AT5G03555 Nucleic acid permease

Other/unknown transport

{EM#}AT1G30400 Glutathione S-conjugate transporting ATPase MRP1

{EM#} A4G16370 Oligopeptide transporter (OPT) family OPT3

{EM#}AT4G10770 Oligopeptide transporter (OPT) family OPT7

{EM#}AT5G55930 Oligopeptide transporter (OPT) family OPT1

{EM#}AT3G55110 ABC-2 type transporter family protein

{EM#}AT5G06530 ABC-2 type transporter family protein

{EM#}AT2G36380 ATP binding cassette (ABC) transporter PDR6

{EM#}AT4G39850 ATP binding cassette (ABC) transporter, peroxisomal

{EM#}AT4G34950 Major facilitator superfamily (MFS) protein






















{EM#}AT2G04090 MATE efflux family protein













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ability to persist in constant light or dark, and a degree of

buffering of period length against temperature changes,a process called temperature compensation.

In plants, the molecular nature of the circadian clock

timing mechanism has been best characterized in Arabidopsis

thaliana. It is believed that every cell in A. thaliana

possesses a circadian oscillator (Thain et al., 2000). It is

possible that there is specialization of circadian clock

function in specific cells (Xu et al., 2007) including root

cells (James et al., 2008) and vascular tissue (Para et al.,2007). Circadian clocks are conceptualized as comprising

a rhythm-generating oscillator(s), light input pathways that

adjust the phase of the rhythm to the local environment and

allow tracking of dawn through the changing seasons, and

output pathways that regulate physiology, metabolism, gene

expression, and development.

The rhythm-generating oscillator is a complex network of

interlocking loops that have been proposed to providerobustness (Troein et al., 2009). Transcription translation

loops (TTLs) that form negative feedbacks are important

components of the rhythm-generating oscillator (Harmer,

2009; Hubbard et al., 2009; Fig. 1). In A. thaliana the TTL

begins at dawn with the accumulation of the myb-like

transcriptional regulators, CIRCADIAN CLOCK ASSO-

CIATED 1 (CCA1) and LATE ELONGATED HYPO-

COTYL (LHY), which activate expression ofPSEUDORESPONSE REGULATORS 7 (PRR7) and

PRR9. PRR7 and PRR9 encode transcriptional repressors

that in turn act on the LHY and CCA1 promoters

(Nakamichi et al., 2010) and complete the so-called

morning loop (Fig. 1). E3 ubiquitin ligase-mediated degra-

dation of CCA1 and LHY allows expression of PRR1 [also

known as TIMING OF CHLOROPHYLL A/B BINDING

PROTEIN1 (TOC1)], which is repressed by CCA1 andLHY. PRR1/TOC1 expression peaks in the evening and is

therefore described as being part of the evening loop of the

clock that probably includes GIGANTEA (GI) with PRR1/

TOC1 being repressive to GI and GI in turn activatingPRR1/TOC1. The expression of GI is also repressed by

LHY and CCA1. The evening-expressed transcription

factors, EARLY FLOWERING3 (ELF3) and LUX

ARRHYTHMO (LUX), directly repress the PRR9 pro-

moter. LUX also represses its own promoter and ELF3

negatively affects expression of PRR7, GI, and TOC1

(Dixon et al., 2011; Helfer et al., 2011). GI and TOC1 are

proteins of unknown function that have both been impli-cated in protein–protein interactions. Steady-state levels of

PRR1/TOC1 are increased by protein–protein interactions

with other PRR proteins. PRR3 competes for binding with

ZEITLUPE (ZTL) preventing degradation of TOC1 (Para

et al., 2007), whereas PRR5, also a target for ZTL-

dependent proteasome degradation (Kiba et al., 2007),

enhances nuclear accumulation of PRR1/TOC1 (Wang

et al., 2010). The TTL is closed by an unknown pathway inwhich PRR1/TOC1 activates CCA1 and LHY expression

just before dawn. It is possible that interactions of PRR1/

TOC1 with CCA1 HIKING EXPEDITION (CHE) permit

activation of CCA1 expression because TOC1 antagonizes the

transcriptional repression of CCA1 by CHE (Pruneda-Paz

et al., 2009; Fig. 1).

Light input to the oscillator is mediated by the crypto-

chromes and phytochromes (Somers et al., 1998; Devlin andKay, 2000) although the precise pathways are not well

characterized. There are several points in the oscillator that

permit light entry and it is thought that this evolved to

allow tracking of dawn in changing photoperiods and to

cope with the environmental noise in the light signal (Troein

et al., 2009). In the TTL, light has a number of effects

including increasing the promoter activity of CCA1 and

LHY, while blue light directly activates ZTL to bind GI.Interaction with GI stabilizes ZTL and at dusk ZTL and GI

disassociate allowing ZTL to target PRR1/TOC1 for

Table 1. Continued


Gene name


{EM#}AT4G25640 MATE family transporter

{EM#}AT3G21390 Mitochondrial substrate carrier family protein





{EM#}AT2G39510 Nodulin MtN21/EamA-like transporter family protein










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degradation (Fig. 1). The light-mediated stabilization of

ZTL through its interaction with GI amplifies the rhythm of

PRR1/TOC1 protein levels (Kim et al., 2007).

There is circadian regulation of transcripts for solute

transporters involved in fluxes of ions, sugars, metals, and

metabolites (Table 1, Covington and Harmer, 2007; Doddet al., 2007). While much focus in recent years has been

placed on the TTLs, an emerging theme suggests that the

changes in solute concentration as a consequence of

alteration in the metabolic status of the cell might also

contribute to oscillator function and possibly rhythm

generation (Blasing et al., 2005; Dodd et al., 2007; James

et al., 2008; Gutierrez et al., 2008; Legnaioli et al., 2009;

Dalchau et al., 2011; Rust et al., 2011). In severalcases, these observations imply an involvement of solute

transport.

Fig. 1. TTLs of the A. thaliana circadian clock. The morning loop (yellow circle) contains the Myb-like transcription factors CCA1 and

LHY, which peak in expression early in the subjective day and promote the expression of PRR7 and PRR9, which reciprocally repress

CCA1 and LHY expression. CCA1 and LHY protein levels are regulated by E3 ubiquitin ligase-mediated degradation. In the central loop

(pink circle), reduction in CCA1 and LHY levels allows expression of TOC1, which peaks in expression in the evening. TOC1

subsequently activates expression of CCA1 and LHY through a hypothetical component ‘X’, possibly involving CHE. CHE binds directly

to the CCA1 promoter inhibiting CCA1 expression, CHE expression is in turn inhibited by CCA1 and TOC1 prevents binding of CHE to

the CCA1 promoter by a direct protein–protein interaction. The evening loop (blue circle) compromises TOC1 and a hypothetical

component ‘Y’, a role which is partly fulfilled by GI. TOC1 represses GI expression, and GI in turn activates TOC1 expression. GI

expression is also repressed by CCA1 and LHY. ELF3 and LUX are also components of the evening loop that directly repress PRR7 to

prevent expression in the night. Clock components are also subject to significant post-translational modifications (green box). TOC1

protein levels are balanced by the combined effects of degradation by the ZTL–SCF–E3 ubiquitin ligase complex. This degradation is

inhibited by GI binding to ZTL, an interaction which is stabilized in blue light, and PRR3, which competes with ZTL for binding to TOC1,

an interaction that is enhanced by phosphorylation. Interaction of TOC1 with PRR5 promotes the accumulation of TOC1 in the nucleus.

There are several targets for light input to the clock, as indicated in the figure (see key). Other input signals to the clock include

temperature cycles and circadian oscillations of cADPR. Physiological outputs include water flux, stomatal aperture, starch

accumulation, and degradation rates, leaf movement and [Ca2+]cyt oscillations. Components that have been demonstrated to act on

clock function but whose position has not been fully elucidated have been omitted for clarity. Diagram modified and updated from

Harmer (2009) and Pruneda-Paz and Kay (2009).


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Carbon assimilation and fluxes associatedwith the circadian clock

There is circadian control of C assimilation from TTLs in

Arabidopsis (Dodd et al., 2004) and daily cycles of flux of C

into sugar and from starch (Stitt et al., 2010). Photosynthe-

sis provides Suc for the rest of the plant during the day and

carbohydrate is stored as starch in plastids. At night, starchis converted to maltose (Mal) in chloroplasts and exported

to the cytosol, where it is converted back into Suc and

utilized in respiration (Stitt et al., 2010). These aspects of

carbohydrate metabolism require coordinated regulation of

transporters for triose-phosphate (triose-P), Mal, and glu-

cose-6-phosphate (Glc-6-P) across chloroplast and plastid

membranes and for Suc across plasma membranes (Fig. 2).

Recent reports have demonstrated the importance ofcircadian TTLs for regulating C assimilation and starch

metabolism for optimizing plant growth (Dodd et al., 2005;

Graf et al., 2010). In addition, it seems that transported

product(s) of photosynthesis, such as sugars, are important

inputs for regulating circadian TTLs (Blasing et al., 2005;

James et al., 2008; Dalchau et al., 2011).

Circadian regulation of sugar and starch metabolism foroptimal growth

Starch accumulates during the day at a generally constant

rate but the rate of breakdown of this stored starch depends

on the length of the night. In light/dark cycles, starch

content peaks at dusk, and is utilized at a rate that exhaustsstarch around dawn (Fig. 2; Gibon et al., 2004; Lu et al.,

2005). Transcripts involved in starch metabolism are

rhythmically expressed in light/dark cycles (Smith et al.,

2004) which, in addition to carbohydrate content, persists in

continuous light (LL) suggesting circadian regulation of

starch metabolism (Harmer et al., 2000; Lu et al., 2005). In

A. thaliana, expression of transcripts for the chloroplast

triose-P translocator (TPT; Flugge et al., 1989; Schneideret al., 2002) and MALTOSE EXCESS1 (MEX1), the

chloroplast Mal exporter (Nittyla et al., 2004), are rhythmi-

cally expressed in light/dark cycles (Smith et al., 2004) and

TPT is also under circadian regulation (Table 1). Several

known and putative hexose transporters are circadian

regulated in LL, peaking late in the subjective light period

(Harmer et al., 2000), including SUGAR TRANSPORTER1

(STP1), which encodes a plasma membrane monosaccha-ride/proton symporter in A. thaliana (Sherson et al., 2000).

Transcripts for several members of a recently defined class

of sugar efflux proteins, including a plasma membrane low-

affinity glucose (Glc) uniporter, SWEET1 (Chen et al.,

2010), are also circadian regulated (Table 1). In continuous

dark (DD), oscillations in sugar concentrations or tran-

scripts for starch degradation were not observed (Lu et al.,

2005). The former is likely due to the requirement ofphotosynthesis for carbohydrate cycling, but the latter

might imply a mechanism for light-dependent transcrip-

tional regulation or that sugars are required to maintain

oscillations in transcript abundance (see following section).

Fig. 2. Interaction of sugars with the circadian clock. Light/dark

cycles result in daily shifts in plant metabolism as plant cells

switch between photosynthesis and respiration. Sugars are

synthesized by photosynthesis during the day and stored as

starch for utilization in respiration during the night giving rise to

diel oscillations of sugar and starch concentrations and depend

on various carbohydrate (CHO) transporters at different stages in

the light/dark cycle. The rate of starch utilization is under

circadian regulation, capable of adapting to changes in light

period to ensure sufficient C supply for the entire night,

irrespective of its duration. The schematic summarizes

oscillations under long day (LD, dashed lines) and short day (SD,

continuous lines) conditions from Gibon et al. (2004), Lu et al.

(2005) and Graf et al. (2010). Sugars have a positive effect on

clock function and this input pathway might involve SFR6 and

GI. The production of ROS and ATP as a consequence of

photosynthesis might also have an effect on clock function. In turn,

the circadian clock and oscillations of sugars themselves

contribute to rhythmic expression of sugar-responsive transcripts

(red line) and transcripts involved in starch metabolism (black line).

The schematic represents data from Smith et al. (2004) and

Blasing et al. (2005). Long-distance movement by intercellular

transport of a product of photosynthesis might couple root and

shoot clocks.


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Furthermore, diel changes in protein abundance of several

key enzymes in starch degradation were not detected,

suggesting that in addition to transcriptional regulation

there likely exist post-translational mechanisms for

circadian regulation of starch metabolism (Lu et al., 2005).

The pattern of starch metabolism is remarkably dynamic

in its ability to adjust to changes in light period (Gibon

et al., 2004; Graf et al., 2010). In entrained plants, extensionof the night led to exhaustion of starch before dawn but

optimal rates of starch metabolism were restored within

a single light/dark cycle (Gibon et al., 2004). Furthermore,

when plants were prematurely transferred to the dark, the

rate of utilization was immediately adjusted to prevent

depletion of starch before dawn of the extended night (Graf

et al., 2010; Fig. 2). Midday dark treatments did not affect

the rate of starch degradation in the night, suggestingregulation by the 24-h period of the circadian clock rather

than light/dark transitions (Graf et al., 2010). Consistent

with this, wild-type plants grown in 28-h T cycles (14L/14D)

grew more slowly than in 24-h T cycles, exhausted starch

before dawn, and up-regulated sugar-starvation response

transcripts with coincident peak expression of LHY (Graf

et al., 2010). The difference was suppressed by supplying

Suc suggesting that this might be a consequence ofperturbed sugar metabolism (Graf et al., 2010). Further-

more, starch metabolism oscillated with a 17-h period in

cca1-11, lhy-21 but was unaffected in toc1-2 or ztl-3

mutants, which affect the evening oscillator (Graf et al.,

2010), suggesting that regulation of starch metabolism by

TTLs might be directed from the morning loop (Fig. 2).

Together, these data indicate that starch metabolism,

including transcripts for sugar transporters, is regulatedfrom TTLs of the circadian clock with important implica-

tions for optimal plant growth.

Effect of sugars on clock gene expression

Awareness of circadian regulation of photosynthesis and

carbohydrate metabolism has existed for some time. More

recently, however, it has become apparent that transported

photosynthates, such as sugars, might also provide feedback

to directly regulate circadian clocks through TTLs, thus

demonstrating that solutes themselves can affect regulation

of the clock (Blasing et al., 2005; James et al., 2008;

Dalchau et al., 2011). Microarray experiments indicatedthat 30–50% of leaf transcripts had significant rhythmic

changes in transcript abundance in light/dark cycles

(Blasing et al., 2005). Comparison of highly Glc-, Suc-, and

CO2-responsive transcripts with rhythmically expressed

transcripts revealed that a high proportion of these

C-responsive transcripts are rhythmically regulated in

a pattern that correlates with the endogenous abundance of

the respective solutes according to the time of day (Blasinget al., 2005). This suggested that these stimuli might

contribute to diel oscillations in transcript abundance.

Consistent with this, starchless mutants of PHOSPHOGLYC-

ERATE/BISPHOSPHOGLYCERATE MUTASE (PGM),

which have an amplified oscillation in endogenous sugar

content, have a higher amplitude of oscillation of Glc-

responsive and circadian-regulated transcripts (Blasing

et al., 2005). Principal component analysis supported the

importance of sugar over light or water stress in daily

oscillations in gene expression and it was concluded that

sugars and the circadian clock are the two most significant

inputs to rhythmic regulation of transcripts with sugars

reinforcing the oscillations of circadian transcripts (Blasinget al., 2005; Fig. 2). This is consistent with loss of rhythmic

expression of starch degradation genes in DD (Lu et al.,

2005), which might also be driven by oscillations in sugar

concentrations.

Although the mechanisms by which sugars feed into the

clock remain unclear, Knight et al. (2008) reported that

addition of Suc increased the amplitude of clock gene

expression and reduced period length of leaf movement ofwild-type Arabidopsis in LL, and that this response was

diminished in sensitive to freezing6 (sfr6) mutants. SFR6 is

involved in post-translational regulation of nuclear targets

(Knight et al., 2009) and might contribute mechanistically

to an input pathway for sugars into TTLs of the circadian

clock (Fig. 2). More recently, a combined mathematical and

experimental study demonstrated that exogenous Suc is

necessary for sustained circadian oscillations in DD in A.

thaliana and that a clock component, GI, is required for the

long-term response of TTLs to Suc (Dalchau et al., 2011).

Therefore, GI might integrate metabolic signals into the

molecular oscillator. The finding that the circadian clock of

A. thaliana distinguishes between short- and long-term

changes in metabolic status (Dalchau et al., 2011) might be

related to the observation that the circadian system of the

crassulacean acid metabolism (CAM) plant Kalanchoe

daigremontiana is insensitive to transient metabolic alter-

ations (Wyka et al., 2004). The fluxes of malate at the

tonoplast in CAM are very large, and it was proposed that

there might be a novel CAM circadian oscillator at the

tonoplast based on metabolic fluxes (Blasius et al., 1999).

However, the insensitivity of CAM circadian rhythms to

transient fluxes of metabolites (Wyka et al., 2004) and the

presence of orthologues of the A. thaliana TTL genes in theCAM plant Mesembryanthemum crystallinum suggest

instead that the genetic basis of circadian oscillations in C3

and CAM plants are conserved (Boxall et al., 2005).

The root clock and the role for a photosynthesis-derivedsignal

A transported photosynthate, possibly Suc, might be re-

quired to couple the root and shoot clocks, providing

further evidence for the role of sugars in regulating TTLs

(James et al., 2008). The majority of circadian clock

experiments in A. thaliana have focused on shoots of plants

grown in the presence of exogenous Suc, which woulddiminish the effect of oscillations in endogenous sugars

produced from photosynthesis in light/dark cycles. James

et al. (2008) investigated circadian clock gene regulation in

roots of hydroponically grown A. thaliana without Suc in

the solution. In LL, LHY, CCA1, and TOC1 were rhythmic


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in shoots. In contrast, expression of LHY and CCA1 in

roots oscillated with a lengthened period and expression of

TOC1, as well as other transcripts that are regulated by the

cis-acting evening element (EE), was high and arrhythmic.

Furthermore, in toc1-10 mutants, the circadian period of

LHY was shortened in shoots, but it was unaffected in roots

(James et al., 2008). In the absence of Suc, 13.7% of shoot

transcripts were rhythmic in LL, but only 3.2% wererhythmic in roots and, similar to LHY and CCA1, the

period of these transcripts was ;2 h longer. In light/dark

cycles, most clock transcripts were regulated in the same

phase in roots and shoots but chemical inhibition of

photosynthesis, or addition of Suc at dusk, uncoupled the

root and shoot clocks (James et al., 2008). Together, these

indicate that in LL and in the absence of exogenous Suc

supply, the morning loop of the circadian clock runs slowlyin roots and the evening loop seems to be non-functional,

implying a dependence of the root circadian clock on the

shoot clock. Furthermore, this relationship appears to be

dependent on photosynthesis and suggests a requirement

for transport of a photosynthate. Since Suc is the major

transported sugar, this seems a likely candidate but the

effect of reduced Suc transport on circadian clock function

is currently unknown. Alternatively, adenosine triphosphate(ATP), another direct product of photosynthesis, which can

drive the core oscillator of cyanobacteria in vitro, might be

important (Rust et al., 2011).

Circadian regulation of calcium signalling

In addition to oscillations in the concentrations of bulk

solutes such as Suc, there are circadian oscillations in

solutes that do not contribute directly to the osmotic status

of the cell. In A. thaliana and Nicotiana plumbaginifolia

there are circadian oscillations in [Ca2+]cyt in LL and in

light/dark cycles (Johnson et al., 1995). Chloroplastic free[Ca2+] oscillates with a circadian period in DD, although

not in LL (Johnson et al., 1995). These data demonstrate

that Ca2+, an essential ion that controls numerous signalling

events (Dodd et al., 2010), undergoes daily rhythmic fluxes

in and out of the cytosol and organelles of plant cells. The

amplitude of the oscillations (;350 nM) is sufficient to

activate signalling pathways regulating both physiology and

gene expression (Love et al., 2004; Dodd et al., 2010). Thepurpose of the circadian oscillations is not known but they

have been proposed to participate in photoperiodism and

stress signalling, and might also regulate TTL function

(Johnson et al., 1995; Love et al., 2004; Dodd et al., 2007).

At the plasma membrane, Ca2+ flux into the cytosol occurs

at the plasma membrane through hyperpolarization-acti-

vated Ca2+ channels and possibly depolarization-activated

Ca2+ channels, glutamate-like receptors, cyclic nucleotide-gated channels, non-specific cation channels, and annexins

(Laohavisit et al., 2009; Dodd et al., 2010), at the tonoplast

through the slow vacuolar two-pore channel 1 (SV/TPC1)

(Peiter et al., 2005), and at the tonoplast and endoplasmic

reticulum membranes through inositol (1,4,5) trisphos-

phate- and cyclic adenosine diphosphate ribose (cADPR)-

gated Ca2+ channels (Dodd et al., 2010). It has been

proposed that cADPR-mediated Ca2+ influx is required for

circadian oscillations of [Ca2+]cyt but because the molecular

identity of neither the cADPR-regulated channel nor

ADPR cyclase that makes cADPR are known in plants, the

mechanisms by which the clock regulates this pathway are

not fully explained (Dodd et al., 2007).The phase of the [Ca2+]cyt rhythm in whole leaves is

responsive to day length, with the peak being close to dusk

in short days (8 h L/16 h D) but in the middle of the day in

long days (16 h L/8 h D) (Love et al., 2004). There are no

oscillations of [Ca2+]cyt in DD (Johnson et al., 1995). The

different dynamics of [Ca2+]cyt in different light conditions

is a consequence of dual regulation by rapid light signalling

pathways of both the TTLs and [Ca2+]cyt (Dalchau et al.,2010). In the morning red light promotes increases in

[Ca2+]cyt, but in the afternoon, blue light promotes

decreases in [Ca2+]cyt (Dalchau et al., 2010). Circadian

oscillations of [Ca2+]cyt are dependent on the presence of

CCA1 and regulated by many of the genetic oscillator

components described above, suggesting that these oscilla-

tions depend on a TTL similar to those previously described

(Xu et al., 2007). However, some of the effects of clockmutations on circadian [Ca2+]cyt cycles suggest that [Ca

2+]cytoscillations are dependent on a cell-specific oscillator re-

stricted to particular cell types (Xu et al., 2007). Most

notably, [Ca2+]cyt rhythms are unaffected by the toc1-1

mutation, which causes a short period of other circadian

outputs, and rhythms are absent or very damped in cca1-1

nulls, while other rhythms persist with a short period (Xu

et al., 2007). In LL, circadian rhythms of [Ca2+]cyt areabsent in A. thaliana seedlings grown on 3% Suc (Johnson

et al., 1995). This might suggest that circadian rhythms of

[Ca2+]cyt are linked to the metabolic status of the cell.

In mammals, it is proposed that metabolic status as

reported by nicotinamide adenine dinucleotide (NAD+)

levels has a significant role in circadian function. It is

proposed that the NAD+ synthesis pathway oscillates in

a circadian manner to regulate the activity of NAD+-dependent protein deacetylases/ADP-ribosyltransferase,

SIRT1 (the mammalian orthologue of SIR2, silent informa-

tion regulator 2, or sirtuins), and in turn regulates circadian

gene expression (Nakahata et al., 2009; Ramsey et al.,

2009). Nicotinamide, an inhibitor of sirtuins and other

pathways involving NAD+ synthesis, lengthens clock period

independently of SIRT1 activity (Asher et al., 2008;

Nakahata et al., 2008) suggesting an additional NAD+-dependent mechanism that regulates circadian period and

this might include the activity of poly(ADP-ribose) poly-

merase 1 (Asher et al., 2010). In plants, it was proposed that

the synthesis of a product of NAD+, cADPR, oscillates in

a circadian manner to regulate Ca2+ fluxes (Dodd et al.,

2007). cADPR-driven oscillations of [Ca2+]cyt might also

link NAD+ metabolism to the clock in mammals (Ikeda

et al., 2003). Daily alterations in solutes such as Suc andCa2+ therefore have the potential to couple energy status to

rhythm generation. The sensitivity of the circadian clock in


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plants and animals to nicotinamide provides evidence of

diverse roles for NAD+ in timekeeping, and NAD+ might

act through cADPR-mediated changes in [Ca2+]cyt in both

systems.

Interactions between light and circadian Ca2+

oscillations

To study the regulation and function of circadian [Ca2+]cytoscillations, Dalchau et al. (2010) developed a mathematical

model that provides insight into the control of Ca2+ fluxes

and exposed underlying control principles of circadianrhythms. This model, which was verified experimentally,

demonstrated that the dynamics of [Ca2+]cyt were best

described by a network that consists of light signalling

pathways that operate over rapid timescales directly regu-

lating [Ca2+]cyt, in addition to regulation by TTLs (Fig. 3).

This basic network structure was also found to apply to

>1000 genes regulated by the circadian clock. The in-

corporation of light signalling into the control of circadianoutputs, including [Ca2+]cyt and gene expression, is associ-

ated with the ability to adjust phase to photoperiods of

differing lengths (Dalchau et al., 2010). This provides

evidence that the phase of circadian rhythms of [Ca2+]cytand gene expression occurs through external coincidence

between circadian signals and the timing of the external

light/dark cycle (Fig. 3). In an external coincidence model,

gene expression, for example, could be regulated bya transcription factor, the expression of which is an

oscillating output of the clock and thus expression of this

transcription factor will be controlled by the phase of the

oscillator. However, the activity of this transcription factor

would be modified by rapid light signalling pathways due to

phosphorylation, degradation, and/or activation/repression

by Ca2+-signalling networks (Fig. 3).

Circadian control of water fluxes

Regulation of water fluxes within the plant is critical for

maintenance of the transpiration stream. This is necessary

to meet changing water demands according to the light/dark

cycle and to facilitate nutrient transport through the xylem.

The control of water fluxes is also critical for regulating

cellular turgor, which is required for circadian stomatalmovements (Lebaudy et al., 2008) and has been implicated

in circadian leaf movement (Moshelian et al., 2002a, b;

Siefritz et al., 2004). Regulation of water fluxes requires

coordinated regulation of aquaporin water channels and

transport of ions (Table 1) to regulate osmotic potentials,

thereby driving the flow of water through the plant.

Maintenance of water balance is dependent on components

of circadian TTLs (Dodd et al., 2004, 2005; Legnaioli et al.,2009), and circadian-regulated aquaporins (Moshelian

et al., 2002a; Siefritz et al., 2004), and K+ channels

(Moshelian et al., 2002b; Lebaudy et al., 2008) have been

implicated in these processes. These transport processes

are driven by proton gradients established by plasma

membrane H+-ATPases (Moran et al., 1996; Kim et al.,

2010), but the present authors are not aware of evidence for

circadian regulation of these proton pumps.

Regulation of stomata

Stomata each comprise two guard cells, which form a pore

that allow transpiration and gas exchange in the aerial parts

of the plant to increase water use efficiency and photosyn-

thesis. In C3 and C4 plants, stomata are closed during the

night and open during the day. Stomata open in the hours

before dawn to permit rapid CO2 fixation during first lightwhile temperatures are low to reduce water loss from

transpiration. Stomatal aperture is controlled by changes in

turgor pressure of the guard cells through regulated activity

of anion and K+ channels in the plasma membrane,

together with the formation of malate from osmotically

inactive starch (Kim et al., 2010). Stomatal aperture is

regulated in a circadian manner, persisting in LL and DD

(Stafelt, 1963). CCA1-ox plants are arrhythmic for stomatalconductance in LL and fail to anticipate dawn and dusk in

light/dark cycles (Dodd et al., 2005) and ztl-1 mutants have

a lengthened circadian period of stomatal conductance in

LL (Dodd et al., 2004). Knockouts or overexpressers of

TOC1 have altered stomatal conductance with impacts on

water balance and drought tolerance, which is dependent on

ABSCISSIC ACID-BINDING PROTEIN (ABAR). Absci-

sic acid (ABA) promotes TOC1 expression and TOC1 bindsdirectly to the ABAR promoter and represses ABAR

expression, demonstrating a possible mechanism for TTL-

mediated regulation of water balance through ABA

(Legnaioli et al., 2010).

Although the transport mechanisms underlying changes

in stomatal aperture have been well characterized and

depend on vacuole and plasma membrane channels and

transporters for K+, Cl-, NO3, and Ca2+ (Kim et al., 2010),it remains largely unknown how these are regulated by

TTLs to pre-empt dawn and dusk. Arabidopsis kincless

mutants, which have undetectable plasma membrane in-

ward K+ channel activity specifically in stomata by domi-

nant-negative suppression of guard cell Shaker K+

channels, fail to increase transpiration rates in anticipation

of dawn in light/dark cycles or in DD (Lebaudy et al.,

2008). kincless mutants also have reduced growth comparedwith wild type when exposed to high light intensity at the

beginning of each day, reflecting the importance of circa-

dian regulation of water fluxes through solute transport for

optimal plant growth.

Changes in concentrations of sugars in guard cells have

also been proposed to be critical for regulating stomatal

aperture (Talbott and Zeiger, 1996) and this is probably

contributed to by hexose transporters (Ritte et al., 1999).STP3, STP5, and circadian-regulated STP1 are expressed

in guard cells and STP1 is localized to guard cell plasma

membranes (Stadler et al., 2003). These sugar transporters

might, therefore, be important for regulating turgor

pressure in guard cells.


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Circadian regulation of leaf movement bywater fluxes

A further role for water fluxes in plants has been described

for the circadian regulation of leaf movement. The best-

characterized example is the daily lifting and falling of

leaves of the rain tree (Samanea saman), which persists for

>3 d in DD (Moshelian et al., 2002a). This movement is

driven by the pulvini (motor) cells, which comprise upper

flexor cells and lower extensor cells that act to lower and

raise the leaf, respectively, by regulated changes in turgor

pressure (Moran et al., 1996). Much like the regulation of

stomatal aperture, this is thought to involve activation of

plasma membrane K+ and Cl– channels (Satter et al., 1974;

Moran et al., 1988) and aquaporins (Moshelian et al.,2002a), which act to coordinately and differentially regulate

turgor pressure in pulvini flexor and extensor cells. One of

two pulvini-specific plasma membrane aquaporins,

SsAQP2, was shown to increase water permeability when

expressed in Xenopus oocytes (Moshelian et al., 2002a).

Addition of Hg or phloretin as transport inhibitors specifi-

cally reduced water permeability in SsAQP2-expressing

oocytes and of extensor cells, and high, pulvinus-specificSsAQP2 expression correlated with high leaf angle in light/

dark and DD cycles (Moshelian et al., 2002a). The water

permeability of flexor and extensor cell protoplasts was

Fig. 3. Circadian outputs are mediated by both a central oscillator and rapid light signalling pathways. Circadian-regulated [Ca2+]cyt and

gene expression oscillations are controlled by external coincidence of clock and external light/dark signals (Dalchau et al., 2010). For

example, the dynamics of circadian oscillations in the expression of an output gene might depend on circadian changes in expression of

an activating transcription factor due to central oscillator activity. The activity of the transcription factor might also depend on light-

dependent degradation, light-dependent [Ca2+]cyt signals, and phosphorylation. Thus the output gene expression will integrate both

circadian control and fast light-dependent signalling to adapt the phase and shape of the oscillation to match the photoperiod.


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higher in the morning and the evening and was reduced by

addition of Hg or phloretin (Moshelian et al., 2002a). This

would facilitate water flux during periods of leaf movement

but is not sufficient to explain changes in turgor. Putative

inward-rectifying (SPICK1/2) and putative outward-

rectifying (SPOCK1 and SPORK1) K+ channels were also

identified in S. saman pulvini that showed high identity to

characterized K+ channels from other species (Moshelianet al., 2002b). All transcripts were rhythmically expressed in

LD cycles, specifically in pulvini, and this persisted in DD

suggesting circadian regulation. SPICK2, SPOCK1, and

SPORK1 were all most highly expressed in both extensor

and flexor cells in the morning, whereas SPICK1 is highly

expressed in extensor cells during the dark when leaf angle

is low and might contribute to lifting of leaves at dawn

(Moshelian et al., 2002b). Although there appears to becircadian regulation of transcripts for K+ channels, it is not

easy to explain how these expression patterns could account

for leaf movement and might suggest additional mecha-

nisms for circadian regulation of these, or alternative,

channels.

In plants that lack motor cells, such as Arabidopsis and

tobacco, leaf movement also follows a circadian rhythm.

A tobacco aquaporin, NtAQP1 is under circadian regula-tion in petioles, being rhythmically expressed in both light/

dark and LL conditions (Siefritz et al., 2004). NtAQP1

expression and protein abundance correlated with petiole

lengthening and epinastic leaf movement, peaking in the

early light phase. In wild type, petiole protoplasts had

higher swelling kinetics at dawn than dusk, and this

difference was lost in protoplasts isolated from AQP1

antisense-silenced transgenic tobacco. Circadian oscillationsin leaf angle were absent in these antisense lines, consistent

with a role for aquaporin in circadian-regulated leaf

movement in plants (Siefritz et al., 2004).

Nutrient acquisition and circadian behaviour

Regulation of micronutrient homeostasis depends on trans-

port proteins for uptake, long-distance transport, tissue

distribution, and subcellular localization, and requires

maintenance of sub-toxic cytosolic concentrations of free

ions. There is circadian regulation of transcripts for micro-

nutrient transporters (Table 1). This might be expected dueto the heavy reliance on metals for the photosynthetic

apparatus, which demands a higher requirement for Mg,

Fe, Mn, and Cu by orders of magnitude compared with

non-photosynthetic organisms (Shcolnick and Keren, 2006),

and dependence on the transpiration stream for long-

distance movement of nutrients (Clemens et al., 2002), both

of which are circadian regulated. Furthermore, recent

reports have associated the circadian clock with regulationof micronutrient homeostasis and flux (Duc et al., 2009;

Andres-Colas et al., 2010).

In Arabidopsis, FERRETIN1 (FER1) encodes an Fe-

storage protein that is highly expressed in Fe-excess

conditions and has been implicated in protection against

Fe-dependent oxidative stress (Ravet et al., 2009). A genetic

screen for de-repressed mutants that express AtFER1 in low

Fe conditions identified novel alleles of tic, a mutant

implicated in circadian clock function (Duc et al., 2009).

Mutants of TIC have a short circadian period with effects

on regulation of the evening oscillator (Ding et al., 2007).

Leaf chlorosis of tic-2 mutants grown in standard con-

ditions was rescued by supplementing with Fe and themutants were sensitive to high Fe (Duc et al., 2009). FER1

expression was rhythmic in light/dark cycles peaking in

early morning, persisting in LL and the oscillation was

absent in lhy-21 and cca1-11 mutants indicating regulation

of FER1 by circadian TTLs. In tic-2, FER1 expression

continues to oscillate in light/dark cycles, but at an elevated

level by a mechanism that might be light dependent

(Duc et al., 2009). Expression of APX1, encoding thereactive oxygen species (ROS) scavenger ascorbate peroxi-

dase, FER3 and FER4 were also higher in tic but the

Fe-dependent regulation of these transcripts was not

affected and de-repression of FER1 in tic occurs indepen-

dently of Fe-dependent regulatory motifs (Duc et al., 2009).

Similarly, the expression of IRON REGULATED TRANS-

PORTER1 (IRT1) and FERRIC REDUCTION OXI-

DASE2 (FRO2) transcripts for Fe uptake in roots was notaffected in tic (Duc et al., 2009) but this could be a reflection

of distinctions between root and shoot clocks (James et al.,

2008). Therefore, TIC might be required for light-dependent

repression of FER1, independently of the circadian clock

and given the possible role for ferritin in oxidative stress,

this might be related to regulation of oxidative stress

responses, rather than Fe homeostasis per se. This seems

reasonable since ROS would be elevated during the day asa consequence of photosynthesis (Fig. 2), are also associated

with fluxes of free metal ions, and might represent an

output of photosynthetic energy production that affects

regulation of the circadian clock.

More recently, it was shown that overexpression of either

of two plasma membrane Cu+ transporters, COPPER

TRANSPORTER1 (COPT1) or COPT3, conferred

increased Cu accumulation and hypersensitivity to highexogenous Cu concentrations (Andres-Colas et al., 2010). In

addition, these transgenics had developmental phenotypes

that the authors suggested were reminiscent of circadian

clock mutants. Subtle differences were observed between

growth of the COPT1/2 overexpressers and wild type in LL.

This could be due to altered circadian clock function or

might reflect elevated oxidative stress under LL, which is

consistent with the elevated anthocyanin levels in thetransgenics (Andres-Colas et al., 2010). LHY and CCA1

transcripts were lower in overexpressers at dawn compared

with wild type. In wild type, addition of Cu to Murashige

and Skoog (MS) medium reduced the magnitude of LHY

expression in light/dark cycles, LL, or DD, and chelation of

Cu increased the amplitude of LHY expression in LL

(Andres-Colas et al., 2010). It has been suggested that MS

is somewhat Cu deficient since activity of Cu-dependentproteins is relatively low in plants grown on this medium

(Abdel-Ghany et al., 2005). Therefore, these data might


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suggest that mild Cu deficiency enhances amplitude of

circadian clock gene expression and that the COPT1/3

overexpressers are less Cu deficient due to increased Cu

uptake, consistent with higher expression of transcripts for

Cu-dependent proteins in these lines. Alternatively, these

observations might be somewhat analogous to those observed

for the TIC-dependent regulation of FER1, which might

implicate oxidative stress responses (Duc et al., 2009).Nevertheless, these studies suggest that interactions between

micronutrient concentrations and circadian TTLs might exist.

A mechanism for how circadian regulation of nutrient

acquisition could derive from TTLs has been reported

(Gutierrez et al., 2008). Assimilation of mineral nutrients

such as N and S depends on uptake of inorganic forms from

soil and subsequent reduction to organic forms for utiliza-

tion by the plant. Transcripts encoding components ofN and S assimilation are circadian regulated including

nitrate, ammonium, and sulphate transporters (Table 1).

Analysis of regulation of N assimilation revealed that

surprisingly few transcripts were regulated by inorganic N,

including several high-affinity nitrate transporters and

nitrite reductase, and indicated that organic/inorganic

N balance is more important for transcriptional regulation

of N reduction, N assimilation, and amino acid metabolismin Arabidopsis (Gutierrez et al., 2008). Network analysis

identified CCA1 as a central regulator of N metabolism and

CCA1 was shown to directly bind promoters of genes for

glutamine synthetase (GLN3.1) and glutamate dehydroge-

nase (GDH) and affect expression of these as well as

downstream transcripts (Gutierrez et al., 2008). Pulses of

organic or inorganic N induced positive and negative phase

shifts in CCA1 expression indicating that N can provideinput to TTLs of the circadian clock and demonstrated

a mechanistic link between a component of circadian TTLs

and regulation of solute transport (Gutierrez et al., 2008).

Conclusions

The tremendous progress in determining the nature of the

Arabidopsis circadian clock in the last 15 years has identified

circadian or daily rhythms of a myriad physiological

functions. Fluxes of solute transport are no exception. The

plant cell is a rhythmic milieu that progresses through

changes in metabolic status in response to, and to copewith, the light and temperature stresses imposed by the

rotation of the Earth and the associated oscillations in

water content. These phenomena imply an important role

for the regulation of solute transport in light/dark cycles

and evidence and mechanisms of how this is dependent on

circadian TTLs are beginning to emerge. In some cases it

has been proposed that these oscillating solutes feed back

into the circadian oscillator to modulate the functioning ofthe circadian clock. A major challenge is to identify those

potential regulators that have major effects on circadian

function, and more specifically to determine the consequen-

ces of this regulation for the physiology of the plant in the

diel cycles in which the organism grows.

Supplementary data

Supplementary data are available at JXB online.

Supplementary Table S1 gives a list of gene nomenclature

used.

Acknowledgements

MJH is supported by BBSRC grant BB/H006826/1 and LJB

is supported by a BBSRC-CASE studentship in partnership

with Bayer Crop Science, both awarded to AARW. Wethank Dr Maria Eriksson (University of Umea) for useful

comments on the figures.

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interactions between plant circadian clocks and solute transport

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