TY - JOUR
AU - Webb, Alex A. R.
AB - 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 concentration and 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. 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 membranes throughout the plant. This might occur through regulation of transcripts, post-translational regulation of protein 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 transcriptional regulation of transporters alone (Covington and Harmer, 2007); it is also necessary to consider whether there are oscillations in the fluxes and concentrations of the transported 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+), micronutrients, and signalling molecules such as calcium (Ca2+). Table 1. Circadian regulated transcripts for transporters in A. thaliana AGI locus  Protein description [The Arabidopsis Infomation 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#}AT5G65000  Nucleotide-sugar transporter family protein    {EM#}AT4G03950  Nucleotide-sugar transporter family protein    {EM#}AT5G17630  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#}AT5G40260  Sugar efflux (SWEET) family, nodulin MtN3 family  SWEET8  {EM#}AT5G50790  Sugar efflux (SWEET) family, nodulin MtN3 family  SWEET10  {EM#}AT3G14770  Sugar efflux (SWEET) family, nodulin MtN3 family  SWEET2  {EM#}AT3G48740  Sugar efflux (SWEET) family, nodulin MtN3 family  SWEET11  {EM#}AT4G15920  Sugar efflux (SWEET) family, nodulin MtN3 family  SWEET17  {EM#}AT1G21460  Sugar efflux (SWEET) family, nodulin MtN3 family  SWEET1  {EM#}AT3G28007  Sugar efflux (SWEET) family, nodulin MtN3 family  SWEET4  {EM#}AT4G25010  Sugar efflux (SWEET) family, nodulin MtN3 family  SWEET14  {EM#}AT5G13170  Sugar efflux (SWEET) family, nodulin MtN3 family  SWEET15  {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#}AT2G41190  Amino acid transporter family    {EM#}AT3G56200  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  {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#}AT4G04850  Potassium transporter family    {EM#}AT5G14880  Potassium transporter family    {EM#}AT2G35060  Potassium transporter, K uptake  KUP11  {EM#}AT4G33530  Potassium transporter, K uptake  KUP5  {EM#}AT1G70300  Potassium transporter, K uptake  KUP6  {EM#}AT3G02050  Potassium transporter, K uptake  KUP3  {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#}AT1G79520  Cation efflux family protein    {EM#}AT1G16310  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#}AT5G24380  Metal-nicotianamine transporter  YSL2  {EM#}AT4G24120  Metal-nicotianamine transporter  YSL1  {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  {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#}AT1G08890  Major facilitator superfamily (MFS) protein    {EM#}AT4G36670  Major facilitator superfamily (MFS) protein    {EM#}AT4G04750  Major facilitator superfamily (MFS) protein    {EM#}AT1G67300  Major facilitator superfamily (MFS) protein    {EM#}AT2G48020  Major facilitator superfamily (MFS) protein    {EM#}AT5G17010  Major facilitator superfamily (MFS) protein    {EM#}AT1G19450  Major facilitator superfamily (MFS) protein    {EM#}AT4G19450  Major facilitator superfamily (MFS) protein    {EM#}AT1G30560  Major facilitator superfamily (MFS) protein    {EM#}AT2G40460  Major facilitator superfamily (MFS) protein    {EM#}AT1G72120  Major facilitator superfamily (MFS) protein    {EM#}AT1G22570  Major facilitator superfamily (MFS) protein    {EM#}AT3G21670  Major facilitator superfamily (MFS) protein    {EM#}AT5G13400  Major facilitator superfamily (MFS) protein    {EM#}AT1G68570  Major facilitator superfamily (MFS) protein    {EM#}AT3G45710  Major facilitator superfamily (MFS) protein    {EM#}AT3G16180  Major facilitator superfamily (MFS) protein    {EM#}AT3G53960  Major facilitator superfamily (MFS) protein    {EM#}AT3G47960  Major facilitator superfamily (MFS) protein    {EM#}AT2G16660  Major facilitator superfamily (MFS) protein    {EM#}AT4G22990  Major facilitator superfamily (MFS) protein    {EM#}AT2G04090  MATE efflux family protein    {EM#}AT2G38510  MATE efflux family protein    {EM#}AT5G17700  MATE efflux family protein    {EM#}AT1G33110  MATE efflux family protein    {EM#}AT1G73700  MATE efflux family protein    {EM#}AT2G34360  MATE efflux family protein    {EM#}AT3G21690  MATE efflux family protein    {EM#}AT1G61890  MATE efflux family protein    {EM#}AT5G38030  MATE efflux family protein    {EM#}AT5G52450  MATE efflux family protein    {EM#}AT1G66760  MATE efflux family protein    {EM#}AT1G51340  MATE efflux family protein    {EM#}AT5G52050  MATE efflux family protein    {EM#}AT4G25640  MATE family transporter    {EM#}AT3G21390  Mitochondrial substrate carrier family protein    {EM#}AT5G56450  Mitochondrial substrate carrier family protein    {EM#}AT2G46320  Mitochondrial substrate carrier family protein    {EM#}AT4G11440  Mitochondrial substrate carrier family protein    {EM#}AT5G01340  Mitochondrial substrate carrier family protein    {EM#}AT2G39510  Nodulin MtN21/EamA-like transporter family protein    {EM#}AT3G18200  Nodulin MtN21/EamA-like transporter family protein    {EM#}AT1G44800  Nodulin MtN21/EamA-like transporter family protein    {EM#}AT3G02690  Nodulin MtN21/EamA-like transporter family protein    {EM#}AT1G11450  Nodulin MtN21/EamA-like transporter family protein    {EM#}AT4G01430  Nodulin MtN21/EamA-like transporter family protein    {EM#}AT2G40900  Nodulin MtN21/EamA-like transporter family protein    {EM#}AT3G53210  Nodulin MtN21/EamA-like transporter family protein    {EM#}AT5G47470  Nodulin MtN21/EamA-like transporter family protein    AGI locus  Protein description [The Arabidopsis Infomation 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#}AT5G65000  Nucleotide-sugar transporter family protein    {EM#}AT4G03950  Nucleotide-sugar transporter family protein    {EM#}AT5G17630  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#}AT5G40260  Sugar efflux (SWEET) family, nodulin MtN3 family  SWEET8  {EM#}AT5G50790  Sugar efflux (SWEET) family, nodulin MtN3 family  SWEET10  {EM#}AT3G14770  Sugar efflux (SWEET) family, nodulin MtN3 family  SWEET2  {EM#}AT3G48740  Sugar efflux (SWEET) family, nodulin MtN3 family  SWEET11  {EM#}AT4G15920  Sugar efflux (SWEET) family, nodulin MtN3 family  SWEET17  {EM#}AT1G21460  Sugar efflux (SWEET) family, nodulin MtN3 family  SWEET1  {EM#}AT3G28007  Sugar efflux (SWEET) family, nodulin MtN3 family  SWEET4  {EM#}AT4G25010  Sugar efflux (SWEET) family, nodulin MtN3 family  SWEET14  {EM#}AT5G13170  Sugar efflux (SWEET) family, nodulin MtN3 family  SWEET15  {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#}AT2G41190  Amino acid transporter family    {EM#}AT3G56200  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  {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#}AT4G04850  Potassium transporter family    {EM#}AT5G14880  Potassium transporter family    {EM#}AT2G35060  Potassium transporter, K uptake  KUP11  {EM#}AT4G33530  Potassium transporter, K uptake  KUP5  {EM#}AT1G70300  Potassium transporter, K uptake  KUP6  {EM#}AT3G02050  Potassium transporter, K uptake  KUP3  {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#}AT1G79520  Cation efflux family protein    {EM#}AT1G16310  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#}AT5G24380  Metal-nicotianamine transporter  YSL2  {EM#}AT4G24120  Metal-nicotianamine transporter  YSL1  {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  {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#}AT1G08890  Major facilitator superfamily (MFS) protein    {EM#}AT4G36670  Major facilitator superfamily (MFS) protein    {EM#}AT4G04750  Major facilitator superfamily (MFS) protein    {EM#}AT1G67300  Major facilitator superfamily (MFS) protein    {EM#}AT2G48020  Major facilitator superfamily (MFS) protein    {EM#}AT5G17010  Major facilitator superfamily (MFS) protein    {EM#}AT1G19450  Major facilitator superfamily (MFS) protein    {EM#}AT4G19450  Major facilitator superfamily (MFS) protein    {EM#}AT1G30560  Major facilitator superfamily (MFS) protein    {EM#}AT2G40460  Major facilitator superfamily (MFS) protein    {EM#}AT1G72120  Major facilitator superfamily (MFS) protein    {EM#}AT1G22570  Major facilitator superfamily (MFS) protein    {EM#}AT3G21670  Major facilitator superfamily (MFS) protein    {EM#}AT5G13400  Major facilitator superfamily (MFS) protein    {EM#}AT1G68570  Major facilitator superfamily (MFS) protein    {EM#}AT3G45710  Major facilitator superfamily (MFS) protein    {EM#}AT3G16180  Major facilitator superfamily (MFS) protein    {EM#}AT3G53960  Major facilitator superfamily (MFS) protein    {EM#}AT3G47960  Major facilitator superfamily (MFS) protein    {EM#}AT2G16660  Major facilitator superfamily (MFS) protein    {EM#}AT4G22990  Major facilitator superfamily (MFS) protein    {EM#}AT2G04090  MATE efflux family protein    {EM#}AT2G38510  MATE efflux family protein    {EM#}AT5G17700  MATE efflux family protein    {EM#}AT1G33110  MATE efflux family protein    {EM#}AT1G73700  MATE efflux family protein    {EM#}AT2G34360  MATE efflux family protein    {EM#}AT3G21690  MATE efflux family protein    {EM#}AT1G61890  MATE efflux family protein    {EM#}AT5G38030  MATE efflux family protein    {EM#}AT5G52450  MATE efflux family protein    {EM#}AT1G66760  MATE efflux family protein    {EM#}AT1G51340  MATE efflux family protein    {EM#}AT5G52050  MATE efflux family protein    {EM#}AT4G25640  MATE family transporter    {EM#}AT3G21390  Mitochondrial substrate carrier family protein    {EM#}AT5G56450  Mitochondrial substrate carrier family protein    {EM#}AT2G46320  Mitochondrial substrate carrier family protein    {EM#}AT4G11440  Mitochondrial substrate carrier family protein    {EM#}AT5G01340  Mitochondrial substrate carrier family protein    {EM#}AT2G39510  Nodulin MtN21/EamA-like transporter family protein    {EM#}AT3G18200  Nodulin MtN21/EamA-like transporter family protein    {EM#}AT1G44800  Nodulin MtN21/EamA-like transporter family protein    {EM#}AT3G02690  Nodulin MtN21/EamA-like transporter family protein    {EM#}AT1G11450  Nodulin MtN21/EamA-like transporter family protein    {EM#}AT4G01430  Nodulin MtN21/EamA-like transporter family protein    {EM#}AT2G40900  Nodulin MtN21/EamA-like transporter family protein    {EM#}AT3G53210  Nodulin MtN21/EamA-like transporter family protein    {EM#}AT5G47470  Nodulin MtN21/EamA-like transporter family protein    Compiled from published lists of circadian regulated transcripts from Dodd et al. (2007) and Covington and Harmer (2007). View Large In C3 and C4 plants the day/night cycle of temperature and 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 characterized by generation of rhythms that approximate to 24 h, the 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 provide robustness (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 ASSOCIATED 1 (CCA1) and LATE ELONGATED HYPOCOTYL (LHY), which activate expression of PSEUDORESPONSE 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 degradation of CCA1 and LHY allows expression of PRR1 [also known as TIMING OF CHLOROPHYLL A/B BINDING PROTEIN1 (TOC1)], which is repressed by CCA1 and LHY. 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 activating PRR1/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 promoter. 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 implicated 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 in which 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). Fig. 1. View largeDownload slide 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). Fig. 1. View largeDownload slide 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). Light input to the oscillator is mediated by the cryptochromes and phytochromes (Somers et al., 1998; Devlin and Kay, 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 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; Dodd et 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 (Bläsing et al., 2005; Dodd et al., 2007; James et al., 2008; Gutiérrez et al., 2008; Legnaioli et al., 2009; Dalchau et al., 2011; Rust et al., 2011). In several cases, these observations imply an involvement of solute transport. Carbon assimilation and fluxes associated with 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). Photosynthesis provides Suc for the rest of the plant during the day and carbohydrate is stored as starch in plastids. At night, starch is 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 glucose-6-phosphate (Glc-6-P) across chloroplast and plastid membranes and for Suc across plasma membranes (Fig. 2). Recent reports have demonstrated the importance of circadian 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 (Bläsing et al., 2005; James et al., 2008; Dalchau et al., 2011). Fig. 2. View largeDownload slide 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 Bläsing et al. (2005). Long-distance movement by intercellular transport of a product of photosynthesis might couple root and shoot clocks. Fig. 2. View largeDownload slide 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 Bläsing et al. (2005). Long-distance movement by intercellular transport of a product of photosynthesis might couple root and shoot clocks. Circadian regulation of sugar and starch metabolism for optimal 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 exhausts starch 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; Flügge et al., 1989; Schneider et al., 2002) and MALTOSE EXCESS1 (MEX1), the chloroplast Mal exporter (Nittylä et al., 2004), are rhythmically 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 monosaccharide/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 transcripts for starch degradation were not observed (Lu et al., 2005). The former is likely due to the requirement of photosynthesis for carbohydrate cycling, but the latter might imply a mechanism for light-dependent transcriptional regulation or that sugars are required to maintain oscillations in transcript abundance (see following section). 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, extension of 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, suggesting regulation 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 of perturbed sugar metabolism (Graf et al., 2010). Furthermore, 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 regulated from TTLs of the circadian clock with important implications 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 (Bläsing et al., 2005; James et al., 2008; Dalchau et al., 2011). Microarray experiments indicated that 30–50% of leaf transcripts had significant rhythmic changes in transcript abundance in light/dark cycles (Bläsing 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 (Bläsing et al., 2005). This suggested that these stimuli might contribute to diel oscillations in transcript abundance. Consistent with this, starchless mutants of PHOSPHOGLYCERATE/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 (Bläsing 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 (Bläsing et 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 of wild-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 Kalanchoë daigremontiana is insensitive to transient metabolic alterations (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 the CAM 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-derived signal A transported photosynthate, possibly Suc, might be required 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 would diminish 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 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% were rhythmic 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 slowly in 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). The purpose 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-activated 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) trisphosphate- 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 oscillations depend on a TTL similar to those previously described (Xu et al., 2007). However, some of the effects of clock mutations on circadian [Ca2+]cyt cycles suggest that [Ca2+]cyt oscillations are dependent on a cell-specific oscillator restricted 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 are absent 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 information 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) polymerase 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 and Ca2+ therefore have the potential to couple energy status to rhythm generation. The sensitivity of the circadian clock in 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+]cyt oscillations, Dalchau et al. (2010) developed a mathematical model that provides insight into the control of Ca2+ fluxes and exposed underlying control principles of circadian rhythms. 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 regulating [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 incorporation of light signalling into the control of circadian outputs, including [Ca2+]cyt and gene expression, is associated 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+]cyt and 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 by a 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). Fig. 3. View largeDownload slide 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. Fig. 3. View largeDownload slide 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. 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 stomatal movements (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 photosynthesis. 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 light while 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 (Ståfelt, 1963). CCA1-ox plants are arrhythmic for stomatal conductance 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). Abscisic acid (ABA) promotes TOC1 expression and TOC1 binds directly 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 inward K+ channel activity specifically in stomata by dominant-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 compared with wild type when exposed to high light intensity at the beginning of each day, reflecting the importance of circadian 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. Circadian regulation of leaf movement by water 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 specifically reduced water permeability in SsAQP2-expressing oocytes and of extensor cells, and high, pulvinus-specific SsAQP2 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 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 (Moshelian et 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 be circadian 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 mechanisms 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 regulation 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 oscillations in 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 transport 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 micronutrient transporters (Table 1). This might be expected due to 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 regulation of 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 conditions was rescued by supplementing with Fe and the mutants 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 the reactive oxygen species (ROS) scavenger ascorbate peroxidase, 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 independently of Fe-dependent regulatory motifs (Duc et al., 2009). Similarly, the expression of IRON REGULATED TRANSPORTER1 (IRT1) and FERRIC REDUCTION OXIDASE2 (FRO2) transcripts for Fe uptake in roots was not affected 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 as a 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 high exogenous 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 the transgenics (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-dependent proteins is relatively low in plants grown on this medium (Abdel-Ghany et al., 2005). Therefore, these data might 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 (Gutiérrez 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 utilization by the plant. Transcripts encoding components of N 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 metabolism in Arabidopsis (Gutiérrez 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 dehydrogenase (GDH) and affect expression of these as well as downstream transcripts (Gutiérrez et al., 2008). Pulses of organic or inorganic N induced positive and negative phase shifts in CCA1 expression indicating that N can provide input to TTLs of the circadian clock and demonstrated a mechanistic link between a component of circadian TTLs and regulation of solute transport (Gutiérrez 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 cope with, 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 of the circadian clock. A major challenge is to identify those potential regulators that have major effects on circadian function, and more specifically to determine the consequences of this regulation for the physiology of the plant in the diel cycles in which the organism grows. Abbreviations 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 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. We thank Dr Maria Eriksson (University of Umeå) for useful comments on the figures. References Abdel-Ghany SE,  Müller-Moulé P,  Niyogi KK,  Pilon M,  Shikanai T.  Two P-type ATPases are required for copper delivery in Arabidopsis thaliana chloroplasts,  The Plant Cell ,  2005, vol.  17 (pg.  1233- 1251) Google Scholar CrossRef Search ADS PubMed  Andrés-Colás N,  Perea-García A,  Puig S,  Peñarrubia L.  Deregulated copper transport affects Arabidopsis development especially in the absence of environmental cycles,  Plant Physiology ,  2010, vol.  153 (pg.  170- 184) Google Scholar CrossRef Search ADS PubMed  Asher G,  Gatfield D,  Stratmann M,  Reinke H,  Dibner C,  Kreppel F,  Mostoslavsky R,  Alt FW,  Schibler U.  SIRT1 regulates circadian clock gene expression through PER2 deacetylation,  Cell ,  2008, vol.  134 (pg.  317- 328) Google Scholar CrossRef Search ADS PubMed  Asher G,  Reinke H,  Altmeyer M,  Gutierrez-Arcelus M,  Hottiger MO,  Schibler U.  Poly(ADP-ribose) polymerase 1 participates in the phase entrainment of circadian clocks to feeding,  Cell ,  2010, vol.  142 (pg.  943- 953) Google Scholar CrossRef Search ADS PubMed  Bläsing OE,  Gibon Y,  Günther M,  Höhne M,  Morcuende R,  Osuna D,  Thimm O,  Usadel B,  Scheible WR,  Stitt M.  Sugars and circadian regulation make major contributions to the global regulation of diurnal gene expression in Arabidopsis,  The Plant Cell ,  2005, vol.  17 (pg.  57- 81) Google Scholar CrossRef Search ADS   Blasius B,  Neff R,  Beck F,  Lüttge U.  Oscillatory model of crassulacean acid metabolism with a dynamic hysteresis switch,  Proceedings of the Royal Society London Series B ,  1999, vol.  266 (pg.  93- 101) Google Scholar CrossRef Search ADS   Boxall SF,  Foster JM,  Bohnert HJ,  Cushman JC,  Nimmo HG,  Hartwell J.  Conservation and divergence of circadian clock operation in a stress-inducible crassulacean acid metabolism species reveals clock compensation against stress,  Plant Physiology ,  2005, vol.  137 (pg.  969- 982) Google Scholar CrossRef Search ADS PubMed  Chen LQ,  Hou BH,  Lalonde S, et al.  Sugar transporters for intercellular exchange and nutrition of pathogens,  Nature ,  2010, vol.  468 (pg.  527- 532) Google Scholar CrossRef Search ADS PubMed  Clemens S,  Palmgren MG,  Krämer U.  A long way ahead: understanding and engineering plant metal accumulation,  Trends in Plant Science ,  2002, vol.  7 (pg.  309- 315) Google Scholar CrossRef Search ADS PubMed  Covington MF,  Harmer SL.  The circadian clock regulates auxin signaling and responses in Arabidopsis,  PLoS Biology ,  2007, vol.  5 pg.  e222  Google Scholar CrossRef Search ADS PubMed  Dalchau N,  Hubbard KE,  Robertson FC,  Hotta CT,  Briggs HM,  Stan GB,  Gonçalves JM,  Webb AAR.  Correct biological timing in Arabidopsis requires multiple light-signaling pathways. Proceedings of the National Academy of Sciences,  USA ,  2010, vol.  107 (pg.  13171- 13176) Google Scholar CrossRef Search ADS   Dalchau N,  Baek SJ,  Briggs HM, et al.  2011. GIGANTEA mediates a long-term response of the Arabidopsis thaliana circadian clock to sucrose,  Proceedings of the National Academy of Sciences, USA,   doi: 10.1073/pnas.1015452108 Devlin PF,  Kay SA.  Cryptochromes are required for phytochrome signaling to the circadian clock but not for rhythmicity,  The Plant Cell ,  2000, vol.  12 (pg.  2499- 2510) Google Scholar CrossRef Search ADS PubMed  Ding Z,  Millar AJ,  Davis AM,  Davis SJ.  TIME FOR COFFEE encodes a nuclear regulator in the Arabidopsis thaliana circadian clock,  The Plant Cell ,  2007, vol.  19 (pg.  1522- 1536) Google Scholar CrossRef Search ADS PubMed  Dixon LE,  Knox K,  Kozma-Bognar L,  Southern MM,  Pokhilko A,  Millar AJ.  Temporal repression of core circadian genes is mediated through EARLY FLOWERING 3 in Arabidopsis,  Current Biology ,  2011, vol.  22 (pg.  120- 125) Google Scholar CrossRef Search ADS   Dodd A,  Parkinson K,  Webb AAR.  Independent circadian regulation of assimilation and stomatal conductance in the ztl-1 mutant of Arabidopsis,  New Phytologist ,  2004, vol.  162 (pg.  956- 966) Google Scholar CrossRef Search ADS   Dodd AN,  Gardner MJ,  Hotta CT, et al.  A cADPR-based feedback loop modulates the Arabidopsis circadian clock,  Science ,  2007, vol.  318 (pg.  1789- 1792) Google Scholar CrossRef Search ADS PubMed  Dodd AN,  Kudla J,  Sanders D.  The language of calcium signaling,  Annual Review of Plant Biology ,  2010, vol.  61 (pg.  593- 620) Google Scholar CrossRef Search ADS PubMed  Dodd AN,  Salathia N,  Hall A,  Kevei E,  Toth R,  Nagy F,  Hibberd JM,  Millar AJ,  Webb AAR.  Plant circadian clocks improve growth, competitive advantage and survival,  Science ,  2005, vol.  309 (pg.  630- 633) Google Scholar CrossRef Search ADS PubMed  Duc C,  Cellier F,  Lobréaux S,  Briat JF,  Gaymard F.  Regulation of iron homeostasis in Arabidopsis thaliana by the clock regulator time for coffee,  Journal of Biological Chemistry ,  2009, vol.  284 (pg.  36271- 36281) Google Scholar CrossRef Search ADS PubMed  Flügge UI,  Fischer K,  Gross A,  Sebald W,  Lottspeich F,  Eckerskorn C.  The triose phosphate-3-phosphoglycerate-phosphate translocator from spinach chloroplasts: nucleotide sequence of a full-length cDNA clone and import of the in vitro synthesized precursor protein into chloroplasts,  EMBO Journal ,  1989, vol.  8 (pg.  39- 46) Google Scholar PubMed  Gibon Y,  Bläsing OE,  Palacios-Rojas N,  Pankovic D,  Hendriks JH,  Fisahn J,  Höhne M,  Günther M,  Stitt M.  Adjustment of diurnal starch turnover to short days: depletion of sugar during the night leads to a temporary inhibition of carbohydrate utilization, accumulation of sugars and post-translational activation of ADP-glucose pyrophosphorylase in the following light period,  The Plant Journal ,  2004, vol.  39 (pg.  847- 862) Google Scholar CrossRef Search ADS PubMed  Graf A,  Schlereth A,  Stitt M,  Smith AM.  Circadian control of carbohydrate availability for growth in Arabidopsis plants at night. Proceedings of the National Academy of Sciences,  USA ,  2010, vol.  107 (pg.  9458- 9463) Google Scholar CrossRef Search ADS   Gutiérrez RA,  Stokes TL,  Thum K, et al.  Systems approach identifies an organic nitrogen-responsive gene network that is regulated by the master clock control gene CCA1. Proceedings of the National Academy of Sciences,  USA ,  2008, vol.  105 (pg.  4939- 4944) Google Scholar CrossRef Search ADS   Harmer SL.  The circadian system in higher plants,  Annual Review of Plant Biology ,  2009, vol.  60 (pg.  357- 377) Google Scholar CrossRef Search ADS PubMed  Harmer SL,  Hogenesch JB,  Straume M,  Chang HS,  Han B,  Zhu T,  Wang X,  Kreps JA,  Kay SA.  Orchestrated transcription of key pathways in Arabidopsis by the circadian clock,  Science ,  2000, vol.  290 (pg.  2110- 2113) Google Scholar CrossRef Search ADS PubMed  Helfer A,  Nusinow DA Chow BY,  Gehrke AR,  Bulyk ML Kay SA.  LUX ARRHYTHMO encodes a nighttime repressor of circadian gene repression in the Arabidopsis core clock,  Current Biology ,  2011, vol.  22 (pg.  126- 133) Google Scholar CrossRef Search ADS   Hubbard KE,  Robertson FC,  Dalchau N,  Webb AAR.  Systems analyses of circadian networks,  Molecular Biosystems ,  2009, vol.  5 (pg.  1502- 1511) Google Scholar CrossRef Search ADS PubMed  Ikeda M,  Sugiyama T,  Wallace CS,  Gompf HS,  Yoshioka T,  Miyawaki A,  Allen CN.  Circadian dynamics of cytosolic and nuclear Ca2+ in single suprachiasmatic nucleus neurons,  Neuron ,  2003, vol.  38 (pg.  253- 263) Google Scholar CrossRef Search ADS PubMed  James AB,  Monreal JA,  Nimmo GA,  Kelly CL,  Herzyk P,  Jenkins GI,  Nimmo HG.  The circadian clock in Arabidopsis roots is a simplified slave version of the clock in shoots,  Science ,  2008, vol.  322 (pg.  1832- 1835) Google Scholar CrossRef Search ADS PubMed  Johnson CH,  Knight MR,  Kondo T,  Masson P,  Sedbrook J,  Haley A,  Trewavas A.  Circadian oscillations of cytosolic and chloroplastic free calcium in plants,  Science ,  1995, vol.  269 (pg.  1863- 1865) Google Scholar CrossRef Search ADS PubMed  Kiba T,  Henriques R,  Sakakibara H,  Chua NH.  Targeted degradation of PSEUDO-RESPONSE REGULATOR5 by an SCFZTL complex regulates clock function and photomorphogenesis in Arabidopsis thaliana,  The Plant Cell ,  2007, vol.  19 (pg.  2516- 2530) Google Scholar CrossRef Search ADS PubMed  Kim WY,  Fujiwara S,  Suh SS,  Kim J,  Kim Y,  Han L,  David K,  Putterill J,  Nam HG,  Somers DE.  ZEITLUPE is a circadian photoreceptor stabilized by GIGANTEA in blue light,  Nature ,  2007, vol.  449 (pg.  356- 360) Google Scholar CrossRef Search ADS PubMed  Kim TH,  Böhmer M,  Hu H,  Nishimura N,  Schroeder JI.  Guard cell signal transduction network: advances in understanding abscisic acid, CO2, and Ca2+ signaling,  Annual Review of Plant Biology ,  2010, vol.  61 (pg.  561- 591) Google Scholar CrossRef Search ADS PubMed  Knight H,  Thomson AJ,  McWatters HG.  Sensitive to freezing6 integrates cellular and environmental inputs to the plant circadian clock,  Plant Physiology ,  2008, vol.  148 (pg.  293- 303) Google Scholar CrossRef Search ADS PubMed  Knight H,  Mugford SG,  Ulker B,  Gao D,  Thorlby G,  Knight MR.  Identification of SFR6, a key component in cold acclimation acting post-translationally on CBF function,  The Plant Journal ,  2009, vol.  58 (pg.  97- 108) Google Scholar CrossRef Search ADS PubMed  Laohavisit A,  Demidchik V,  Mortimer JC, et al.  Zea mays annexins modulate cytosolic free Ca2+, form a Ca2+-permeable conductance and have peroxidase activity,  The Plant Cell ,  2009, vol.  21 (pg.  479- 493) Google Scholar CrossRef Search ADS PubMed  Lebaudy A,  Vavasseur A,  Hosy E,  Dreyer I,  Leonhardt N,  Thibaud JB,  Véry AA,  Simonneau T,  Sentenac H.  Plant adaptation to fluctuating environment and biomass production are strongly dependent on guard cell potassium channels. Proceedings of the National Academy of Sciences,  USA ,  2008, vol.  105 (pg.  5271- 5276) Google Scholar CrossRef Search ADS   Legnaioli T,  Cuevas J,  Mas P.  TOC1 functions as a molecular switch connecting the circadian clock with plant responses to drought,  EMBO Journal ,  2009, vol.  28 (pg.  3745- 3757) Google Scholar CrossRef Search ADS PubMed  Love J,  Dodd AN,  Webb AAR.  Circadian and diurnal calcium oscillations encode photoperiodic information in Arabidopsis,  The Plant Cell ,  2004, vol.  16 (pg.  956- 966) Google Scholar CrossRef Search ADS PubMed  Lu Y,  Gehan JP,  Sharkey TD.  Daylength and circadian effects on starch degradation and maltose metabolism,  Plant Physiology ,  2005, vol.  138 (pg.  2280- 2291) Google Scholar CrossRef Search ADS PubMed  Michael TP,  Mockler TC,  Breton G, et al. ,  Network discovery pipeline elucidates conserved time-of-day-specific cis-regulatory modules. PLoS Genetics 4, e14 ,  2008 Moran N,  Ehrenstein G,  Iwasa K,  Mischke C,  Bare C,  Satter RL.  Potassium channels in motor cells of Samanea saman: a patch-clamp study,  Plant Physiology ,  1988, vol.  88 (pg.  643- 648) Google Scholar CrossRef Search ADS PubMed  Moran N,  Yueh YG,  Crain RC.  Signal transduction and cell volume regulation in plant leaflet movements,  News in Physiological Sciences ,  1996, vol.  11 (pg.  108- 114) Moshelion M,  Becker D,  Biela A,  Uehlein N,  Hedrich R,  Otto B,  Levi H,  Moran N,  Kaldenhoff R.  Plasma membrane aquaporins in the motor cells of Samanea saman: diurnal and circadian regulation,  The Plant Cell ,  2002, vol.  14 (pg.  727- 739) Google Scholar CrossRef Search ADS PubMed  Moshelion M,  Becker D,  Czempinski K,  Mueller-Roeber B,  Attali B,  Hedrich R,  Moran N.  Diurnal and circadian regulation of putative potassium channels in a leaf moving organ,  Plant Physiology ,  2002, vol.  128 (pg.  634- 642) Google Scholar CrossRef Search ADS PubMed  Nakahata Y,  Kaluzova M,  Grimaldi B,  Sahar S,  Hirayama J,  Chen D,  Guarente LP,  Sassone-Corsi P.  The NAD+-dependent deacetylase SIRT1 modulates CLOCK-mediated chromatin remodeling and circadian control,  Cell ,  2008, vol.  134 (pg.  329- 340) Google Scholar CrossRef Search ADS PubMed  Nakahata Y,  Sahar S,  Astarita G,  Kaluzova M,  Sassone-Corsi P.  Circadian control of the NAD+ salvage pathway by CLOCK-SIRT1,  Science ,  2009, vol.  324 (pg.  654- 657) Google Scholar CrossRef Search ADS PubMed  Nakamichi N,  Kiba T,  Henriques H,  Mizuno T,  Chua N-H,  Sakakibara H.  PSEUDO-RESPONSE REGULATORS 9, 7, and 5 are transcriptional repressors in the Arabidopsis circadian clock,  The Plant Cell ,  2010, vol.  22 (pg.  594- 605) Google Scholar CrossRef Search ADS PubMed  Niittylä T,  Messerli G,  Trevisan M,  Chen J,  Smith AM,  Zeeman SC.  A previously unknown maltose transporter essential for starch degradation in leaves,  Science ,  2004, vol.  303 (pg.  87- 89) Google Scholar CrossRef Search ADS PubMed  Para A,  Farré EM,  Imaizumi T,  Pruneda-Paz TL,  Harmon FG,  Kay SA.  PRR3 is a vascular regulator of TOC1 stability in the Arabidopsis circadian clock,  The Plant Cell ,  2007, vol.  19 (pg.  3462- 3473) Google Scholar CrossRef Search ADS PubMed  Peiter E,  Maathuis FJM,  Mills LN,  Knight H,  Pelloux J,  Hetherington AM,  Sanders D.  The vacuolar Ca2+-activated channel TPC1 regulates germination and stomatal movement,  Nature ,  2005, vol.  434 (pg.  404- 408) Google Scholar CrossRef Search ADS PubMed  Pruneda-Paz JL,  Breton G,  Para A,  Kay SA.  A functional genomics approach reveals CHE as a component of the Arabidopsis circadian clock,  Science ,  2009, vol.  323 (pg.  1481- 1485) Google Scholar CrossRef Search ADS PubMed  Ramsey KM,  Yoshino J,  Brace CS, et al.  Circadian clock feedback cycle through NAMPT-mediated NAD+ biosynthesis,  Science ,  2009, vol.  324 (pg.  651- 654) Google Scholar CrossRef Search ADS PubMed  Ravet K,  Touraine B,  Boucherez J,  Briat JF,  Gaymard F,  Cellier F.  Ferritins control interaction between iron homeostasis and oxidative stress in Arabidopsis,  The Plant Journal ,  2009, vol.  57 (pg.  400- 412) Google Scholar CrossRef Search ADS PubMed  Ritte G,  Rosenfeld J,  Rohrig K,  Raschke K.  Rates of sugar uptake by guard cell protoplasts of Pisum sativum L. related to the solute requirement for stomatal opening,  Plant Physiology ,  1999, vol.  121 (pg.  647- 656) Google Scholar CrossRef Search ADS PubMed  Rust MJ,  Golden SS,  O'Shea EK.  Light-driven changes in energy metabolism directly entrain the cyanobacterial circadian oscillator,  Science ,  2011, vol.  331 (pg.  220- 223) Google Scholar CrossRef Search ADS PubMed  Satter RL,  Geballe GT,  Applewhite PB,  Galston AW.  Potassium flux and leaf movement in Samanea saman. I. Rhythmic movement,  Journal of General Physiology ,  1974, vol.  64 (pg.  413- 430) Google Scholar CrossRef Search ADS PubMed  Schneider A,  Häusler RE,  Kolukisaoglu U,  Kunze R,  van der Graaff E,  Schwacke R,  Catoni E,  Desimone M,  Flügge UI.  An Arabidopsis thaliana knock-out mutant of the chloroplast triose phosphate/phosphate translocator is severely compromised only when starch synthesis, but not starch mobilisation is abolished,  The Plant Journal ,  2002, vol.  32 (pg.  685- 699) Google Scholar CrossRef Search ADS PubMed  Shcolnick S,  Keren N.  Metal homeostasis in cyanobacteria and chloroplasts. Balancing benefits and risks to the photosynthetic apparatus,  Plant Physiology ,  2006, vol.  141 (pg.  805- 810) Google Scholar CrossRef Search ADS PubMed  Sherson SM,  Hemmann G,  Wallace G,  Forbes S,  Germain V,  Stadler R,  Bechtold N,  Sauer N,  Smith SM.  Monosaccharide/proton symporter AtSTP1 plays a major role in uptake and response of Arabidopsis seeds and seedlings to sugars,  The Plant Journal ,  2000, vol.  24 (pg.  849- 857) Google Scholar CrossRef Search ADS PubMed  Siefritz F,  Otto B,  Bienert GP,  van der Krol A,  Kaldenhoff R.  The plasma membrane aquaporin NtAQP1 is a key component of the leaf unfolding mechanism in tobacco,  The Plant Journal ,  2004, vol.  37 (pg.  147- 155) Google Scholar CrossRef Search ADS PubMed  Smith SM,  Fulton DC,  Chia T,  Thorneycroft D,  Chapple A,  Dunstan H,  Hylton C,  Zeeman SC,  Smith AM.  Diurnal changes in the transcriptome encoding enzymes of starch metabolism provide evidence for both transcriptional and posttranscriptional regulation of starch metabolism in Arabidopsis leaves,  Plant Physiology ,  2004, vol.  136 (pg.  2687- 2699) Google Scholar CrossRef Search ADS PubMed  Somers DE,  Devlin PF,  Kay SA.  Phytochromes nad cryptochromes in the entrainment of the Arabidopsis circadian clock,  Science ,  1998, vol.  282 (pg.  1488- 1490) Google Scholar CrossRef Search ADS PubMed  Stadler R,  Büttner M,  Ache P,  Hedrich R,  Ivashikina N,  Melzer M,  Shearson SM,  Smith SM,  Sauer N.  Diurnal and light-regulated repression of AtSTP1 in guard cells of Arabidopsis,  Plant Physiology ,  2003, vol.  133 (pg.  528- 537) Google Scholar CrossRef Search ADS PubMed  Ståfelt MG.  Diurnal dark reactions in the stomatal movements,  Physiologia Plantarum ,  1963, vol.  16 (pg.  756- 766) Google Scholar CrossRef Search ADS   Stitt M,  Lunn J,  Usadel B.  Arabidopsis and primary photosynthetic metabolism - more than the icing on the cake,  The Plant Journal ,  2010, vol.  61 (pg.  1067- 1091) Google Scholar CrossRef Search ADS PubMed  Talbott TD,  Zeiger E.  Central roles for potassium and sucrose in guard-cell osmoregulation,  Plant Physiology ,  1996, vol.  111 (pg.  1051- 1057) Google Scholar CrossRef Search ADS PubMed  Thain SC,  Hall A,  Millar AJ.  Functional independence of circadian clocks that regulate plant gene expression,  Current Biology ,  2000, vol.  10 (pg.  951- 956) Google Scholar CrossRef Search ADS PubMed  Troein C,  Locke JCW,  Turner MS,  Millar AJ.  Weather and seasons together demand complex biological clocks,  Current Biology ,  2009, vol.  19 (pg.  1961- 1964) Google Scholar CrossRef Search ADS PubMed  Wang L,  Fujiwara S,  Somers DE.  PRR5 regulates phosphorylation, nuclear import and subnuclear localization of TOC1 in the Arabidopsis circadian clock,  EMBO Journal ,  2010, vol.  29 (pg.  1903- 1915) Google Scholar CrossRef Search ADS PubMed  Wyka TP,  Bohn A,  Duarte HM,  Kaiser F,  Lüttge UE.  Perturbations of malate accumulation and the endogenous rhythms of gas exchange in the Crassulacean acid metabolism plant Kalanchoë daigremontiana testing the tonoplast-as-oscillator model,  Planta ,  2004, vol.  219 (pg.  705- 713) Google Scholar CrossRef Search ADS PubMed  Xu X,  Hotta CT,  Dodd AN,  Love J,  Sharrock R,  Lee YW,  Xie Q,  Johnson CH,  Webb AAR.  Distinct light and clock modulation of cytosolic free Ca2+ oscillations and rhythmic CHLOROPHYLL A/B BINDING PROTEIN 2 promoter activity in Arabidopsis,  The Plant Cell ,  2007, vol.  19 (pg.  3474- 3490) Google Scholar CrossRef Search ADS PubMed  © The Author [2011]. 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TI - Interactions between plant circadian clocks and solute transport
JF - Journal of Experimental Botany
DO - 10.1093/jxb/err040
DA - 2011-03-04
UR - https://www.deepdyve.com/lp/oxford-university-press/interactions-between-plant-circadian-clocks-and-solute-transport-shxAlrx6vS
SP - 2333
EP - 2348
VL - 62
IS - 7
DP - DeepDyve
ER -