Plants have evolved a multitude of strategies to adjust their growth according to external and internal signals. Interconnected metabolic and phytohormonal signalling networks allow adaption to changing environmental and developmental conditions and ensure the survival of species in fluctuating environments. In agricultural ecosystems, many of these adaptive responses are not required or may even limit crop yield, as they prevent plants from realizing their fullest potential. By lifting source and sink activities to their maximum, massive yield increases can be foreseen, potentially closing the future yield gap resulting from an increasing world population and the transition to a carbon-neutral economy. To do so, a better understanding of the interplay between metabolic and developmental processes is required. In the past, these processes have been tackled independently from each other, but coordinated efforts are required to understand the fine mechanics of source–sink relations and thus optimize crop yield. Here, we describe approaches to design high-yielding crop plants utilizing strategies derived from current metabolic concepts and our understanding of the molecular processes determining sink development.
Subscribe to Journal
Get full journal access for 1 year
only $9.92 per issue
All prices are NET prices.
VAT will be added later in the checkout.
Tax calculation will be finalised during checkout.
Rent or Buy article
Get time limited or full article access on ReadCube.
All prices are NET prices.
South, P. F., Cavanagh, A. P., Liu, H. W. & Ort, D. R. Synthetic glycolate metabolism pathways stimulate crop growth and productivity in the field. Science 363, eaat9077 (2019).
Sonnewald, U. & Fernie, A. R. Next-generation strategies for understanding and influencing source-sink relations in crop plants. Curr. Opin. Plant Biol. 43, 63–70 (2018).
Ort, D. R. et al. Redesigning photosynthesis to sustainably meet global food and bioenergy demand. Proc. Natl Acad. Sci. USA 112, 8529–8536 (2015).
Sweetlove, L. J., Nielsen, J. & Fernie, A. R. Engineering central metabolism - a grand challenge for plant biologists. Plant J. 90, 749–763 (2017).
Korner, C. Paradigm shift in plant growth control. Curr. Opin. Plant Biol. 25, 107–114 (2015).
Burnett, A. C., Rogers, A., Rees, M. & Osborne, C. P. Carbon source-sink limitations differ between two species with contrasting growth strategies. Plant Cell Environ. 39, 2460–2472 (2016).
Jonik, C., Sonnewald, U., Hajirezaei, M. R., Flugge, U. I. & Ludewig, F. Simultaneous boosting of source and sink capacities doubles tuber starch yield of potato plants. Plant Biotechnol. J. 10, 1088–1098 (2012).
Rossi, M., Bermudez, L. & Carrari, F. Crop yield: challenges from a metabolic perspective. Curr. Opin. Plant Biol. 25, 79–89 (2015).
Ainsworth, E. A. & Long, S. P. What have we learned from 15 years of free-air CO2 enrichment (FACE)? A meta-analytic review of the responses of photosynthesis, canopy properties and plant production to rising CO2. N. Phytol. 165, 351–371 (2005).
Xiao, Y. G. et al. Genetic gains in grain yield and physiological traits of winter wheat in Shandong Province, China, from 1969 to 2006. Crop Sci. 52, 44–56 (2012).
Sage, R. F., Sage, T. L. & Kocacinar, F. Photorespiration and the evolution of C4 photosynthesis. Ann. Rev. Plant Biol. 63, 19–47 (2012).
Sage, R. F. A portrait of the C4 photosynthetic family on the 50th anniversary of its discovery: species number, evolutionary lineages, and Hall of Fame. J. Exp. Bot. 68, 4039–4056 (2017).
Sage, R. F., Christin, P. A. & Edwards, E. J. The C(4) plant lineages of planet Earth. J. Exp. Bot. 62, 3155–3169 (2011).
Arrivault, S. et al. Metabolite profiles reveal inter-specific variation in operation of the Calvin-Benson cycle in both C4 and C3 plants. J. Exp. Bot. (in the press).
Zhu, X. G., de Sturler, E. & Long, S. P. Optimizing the distribution of resources between enzymes of carbon metabolism can dramatically increase photosynthetic rate: a numerical simulation using an evolutionary algorithm. Plant Physiol. 145, 513–526 (2007).
Galmes, J. et al. Rubisco catalytic properties optimized for present and future climatic conditions. Plant Sci. 226, 61–70 (2014).
Sharwood, R. E., Ghannoum, O. & Whitney, S. M. Prospects for improving CO2 fixation in C3-crops through understanding C4-Rubisco biogenesis and catalytic diversity. Curr. Opin. Plant Biol. 31, 135–142 (2016).
Borghi, G. L. et al. Relationship between irradiance and levels of Calvin-Benson cycle and other intermediates in the model eudicot Arabidopsis and the model monocot rice. J. Exp. Bot. 70, 5809–5825 (2019).
Long, S. P., Marshall-Colon, A. & Zhu, X. G. Meeting the global food demand of the future by engineering crop photosynthesis and yield potential. Cell 161, 56–66 (2015).
Li, Y., Heckmann, D., Lercher, M. J. & Maurino, V. G. Combining genetic and evolutionary engineering to establish C4 metabolism in C3 plants. J. Exp. Bot. 68, 117–125 (2017).
Andralojc, P. J., Carmo-Silva, E., Degen, G. E. & Parry, M. A. J. Increasing metabolic potential: C-fixation. Essays Biochem. 62, 109–118 (2018).
Foyer, C. H., Ruban, A. V. & Nixon, P. J. Photosynthesis solutions to enhance productivity. Philos. Trans. R. Soc. B 372, 20160374 (2017).
Nuccio, M. L. et al. Strategies and tools to improve crop productivity by targeting photosynthesis. Philos. Trans. R. Soc. B 372, 20160377 (2017).
Jansson, C., Vogel, J., Hazen, S., Brutnell, T. & Mockler, T. Climate-smart crops with enhanced photosynthesis. J. Exp. Bot. 69, 3801–3809 (2018).
Sharwood, R. E., Ghannoum, O., Kapralov, M. V., Gunn, L. H. & Whitney, S. M. Temperature responses of Rubisco from Paniceae grasses provide opportunities for improving C3 photosynthesis. Nat. Plants 2, 16186 (2016).
Sharwood, R. E. & Whitney, S. M. Correlating Rubisco catalytic and sequence diversity within C3 plants with changes in atmospheric CO2 concentrations. Plant Cell Environ. 37, 1981–1984 (2014).
Driever, S. M. et al. Increased SBPase activity improves photosynthesis and grain yield in wheat grown in greenhouse conditions. Philos. Trans. R. Soc. B 372, 20160384 (2017).
Lefebvre, S. et al. Increased sedoheptulose-1,7-bisphosphatase activity in transgenic tobacco plants stimulates photosynthesis and growth from an early stage in development. Plant Physiol. 138, 451–460 (2005).
Simkin, A. J. et al. Simultaneous stimulation of sedoheptulose 1,7-bisphosphatase, fructose 1,6-bisphophate aldolase and the photorespiratory glycine decarboxylase-H protein increases CO2 assimilation, vegetative biomass and seed yield in Arabidopsis. Plant Biotechnol. J. 15, 805–816 (2017).
Simkin, A. J., McAusland, L., Lawson, T. & Raines, C. A. Overexpression of the RieskeFeS protein increases electron transport rates and biomass yield. Plant Physiol. 175, 134–145 (2017).
Rae, B. D. et al. Progress and challenges of engineering a biophysical CO2-concentrating mechanism into higher plants. J. Exp. Bot. 68, 3717–3737 (2017).
Schuler, M. L., Mantegazza, O. & Weber, A. P. Engineering C4 photosynthesis into C3 chassis in the synthetic biology age. Plant J. 87, 51–65 (2016).
Nolke, G., Houdelet, M., Kreuzaler, F., Peterhansel, C. & Schillberg, S. The expression of a recombinant glycolate dehydrogenase polyprotein in potato (Solanum tuberosum) plastids strongly enhances photosynthesis and tuber yield. Plant Biotechnol. J. 12, 734–742 (2014).
Kromdijk, J. et al. Improving photosynthesis and crop productivity by accelerating recovery from photoprotection. Science 354, 857–861 (2016).
Lawson, T. & Blatt, M. R. Stomatal size, speed, and responsiveness impact on photosynthesis and water use efficiency. Plant Physiol. 164, 1556–1570 (2014).
Lawson, T., Kramer, D. M. & Raines, C. A. Improving yield by exploiting mechanisms underlying natural variation of photosynthesis. Curr. Opin. Biotech. 23, 215–220 (2012).
Glowacka, K. et al. Photosystem II Subunit S overexpression increases the efficiency of water use in a field-grown crop. Nat. Commun. 9, 868 (2018).
Papanatsiou, M. et al. Optogenetic manipulation of stomatal kinetics improves carbon assimilation, water use, and growth. Science 363, 1456–1459 (2019).
Caine, R. S. et al. Rice with reduced stomatal density conserves water and has improved drought tolerance under future climate conditions. N. Phytol. 221, 371–384 (2018).
Hughes, J. et al. Reducing stomatal density in barley improves drought tolerance without impacting on yield. Plant Physiol. 174, 776–787 (2017).
van Bezouw, R., Keurentjes, J. J. B., Harbinson, J. & Aarts, M. G. M. Converging phenomics and genomics to study natural variation in plant photosynthetic efficiency. Plant J. 97, 112–133 (2019).
Nuccio, M. L. et al. Expression of trehalose-6-phosphate phosphatase in maize ears improves yield in well-watered and drought conditions. Nat. Biotech. 33, 862–869 (2015).
Oszvald, M. et al. Trehalose 6-phosphate regulates photosynthesis and assimilate partitioning in reproductive tissue. Plant Physiol. 176, 2623–2638 (2018).
Gould, N. et al. AtSUC2 has a role for sucrose retrieval along the phloem pathway: evidence from carbon-11 tracer studies. Plant Sci. 188–189, 97–101 (2012).
Zhang, C. & Turgeon, R. Mechanisms of phloem loading. Curr. Opin. Plant Biol. 43, 71–75 (2018).
De Schepper, V., De Swaef, T., Bauweraerts, I. & Steppe, K. Phloem transport: a review of mechanisms and controls. J. Exp. Bot. 64, 4839–4850 (2013).
Braun, D. M., Wang, L. & Ruan, Y. L. Understanding and manipulating sucrose phloem loading, unloading, metabolism, and signalling to enhance crop yield and food security. J. Exp. Bot. 65, 1713–1735 (2014).
Ma, S. et al. Phloem loading in cucumber: combined symplastic and apoplastic strategies. Plant J. 98, 391–404 (2019).
Chen, L. Q. et al. Sucrose efflux mediated by SWEET proteins as a key step for phloem transport. Science 335, 207–211 (2012).
Truernit, E. & Sauer, N. The promoter of the Arabidopsis thaliana SUC2 sucrose-H+ symporter gene directs expression of beta-glucuronidase to the phloem: evidence for phloem loading and unloading by SUC2. Planta 196, 564–570 (1995).
Wingenter, K. et al. Increased activity of the vacuolar monosaccharide transporter TMT1 alters cellular sugar partitioning, sugar signaling, and seed yield in Arabidopsis. Plant Physiol. 154, 665–677 (2010).
Sosso, D. et al. Seed filling in domesticated maize and rice depends on SWEET-mediated hexose transport. Nat. Genet. 47, 1489–1493 (2015).
Wang, H. et al. A subsidiary cell-localized glucose transporter promotes stomatal conductance and photosynthesis. Plant Cell 31, 1328–1343 (2019).
Kryvoruchko, I. S. et al. MtSWEET11, a nodule-specific sucrose transporter of Medicago truncatula. Plant Physiol. 171, 554–565 (2016).
Yang, J., Luo, D., Yang, B., Frommer, W. B. & Eom, J. S. SWEET11 and 15 as key players in seed filling in rice. N. Phytol. 218, 604–615 (2018).
Srivastava, A. C., Ganesan, S., Ismail, I. O. & Ayre, B. G. Functional characterization of the Arabidopsis AtSUC2 Sucrose/H+ symporter by tissue-specific complementation reveals an essential role in phloem loading but not in long-distance transport. Plant Physiol. 148, 200–211 (2008).
van Dongen, J. T., Schurr, U., Pfister, M. & Geigenberger, P. Phloem metabolism and function have to cope with low internal oxygen. Plant Physiol. 131, 1529–1543 (2003).
Giaquinta, R. Mechanism and control of phloem loading of sucrose. Ber. Deut. Bot. Ges. 93, 197–201 (1980).
Deeken, R. et al. Loss of the AKT2/3 potassium channel affects sugar loading into the phloem of Arabidopsis. Planta 216, 334–344 (2002).
Gajdanowicz, P. et al. Distinct roles of the last transmembrane domain in controlling Arabidopsis K+ channel activity. N. Phytol. 182, 380–391 (2009).
Gajdanowicz, P. et al. Potassium (K+) gradients serve as a mobile energy source in plant vascular tissues. Proc. Natl Acad. Sci. USA 108, 864–869 (2011).
Chen, L. Q. et al. A cascade of sequentially expressed sucrose transporters in the seed coat and endosperm provides nutrition for the Arabidopsis embryo. Plant Cell 27, 607–619 (2015).
Zhang, Z. et al. Suppressing a putative sterol carrier gene reduces plasmodesmal permeability and activates sucrose transporter genes during cotton fiber elongation. Plant Cell 29, 2027–2046 (2017).
Grison, M. S. et al. Specific membrane lipid composition is important for plasmodesmata function in Arabidopsis. Plant Cell 27, 1228–1250 (2015).
Yan, D. et al. Sphingolipid biosynthesis modulates plasmodesmal ultrastructure and phloem unloading. Nat. Plants 5, 604–615 (2019).
Kraner, M. E. et al. Choline transporter-like1 (CHER1) is crucial for plasmodesmata maturation in Arabidopsis thaliana. Plant J. 89, 394–406 (2017).
Bologa, K. L., Fernie, A. R., Leisse, A., Loureiro, M. E. & Geigenberger, P. A bypass of sucrose synthase leads to low internal oxygen and impaired metabolic performance in growing potato tubers. Plant Physiol. 132, 2058–2072 (2003).
Lu, M. Z., Snyder, R., Grant, J. & Tegeder, M. Manipulation of sucrose phloem and embryo loading affects pea leaf metabolism, carbon and nitrogen partitioning to sinks as well as seed storage pools. Plant J. 101, 217–236 (2019).
Amthor, J. S. et al. Actionable engineering strategies to cut respiratory carbon loss and boost crop productivity. Plant Cell Environ. (in the press).
Wan, H., Wu, L., Yang, Y., Zhou, G. & Ruan, Y. L. Evolution of sucrose metabolism: the dichotomy of invertases and beyond. Trends Plant Sci. 23, 163–177 (2018).
Ruan, Y. L. Sucrose metabolism: gateway to diverse carbon use and sugar signaling. Annu. Rev. Plant Biol. 65, 33–67 (2014).
Barratt, D. H. et al. Normal growth of Arabidopsis requires cytosolic invertase but not sucrose synthase. Proc. Natl Acad. Sci. USA 106, 13124–13129 (2009).
Baroja-Fernandez, E. et al. Enhancing sucrose synthase activity in transgenic potato (Solanum tuberosum L.) tubers results in increased levels of starch, ADPglucose and UDPglucose and total yield. Plant Cell Physiol. 50, 1651–1662 (2009).
Zhang, L. et al. Overriding the co-limiting import of carbon and energy into tuber amyloplasts increases the starch content and yield of transgenic potato plants. Plant Biotechnol. J. 6, 453–464 (2008).
Geigenberger, P. et al. Inhibition of de novo pyrimidine synthesis in growing potato tubers leads to a compensatory stimulation of the pyrimidine salvage pathway and a subsequent increase in biosynthetic performance. Plant Cell 17, 2077–2088 (2005).
Regierer, B. et al. Starch content and yield increase as a result of altering adenylate pools in transgenic plants. Nat. Biotechnol. 20, 1256–1260 (2002).
Slewinski, T. L. Non-structural carbohydrate partitioning in grass stems: a target to increase yield stability, stress tolerance, and biofuel production. J. Exp. Bot. 63, 4647–4670 (2012).
Smidansky, E. D. et al. Enhanced ADP-glucose pyrophosphorylase activity in wheat endosperm increases seed yield. Proc. Natl Acad. Sci. USA 99, 1724–1729 (2002).
Fernie, A. R. Extending the cascade: identification of a mitogen-activated protein kinase phosphatase playing a key role in rice yield. Plant J. 95, 935–936 (2018).
Weber, H., Buchner, P., Borisjuk, L. & Wobus, U. Sucrose metabolism during cotyledon development of Vicia faba L is controlled by the concerted action of both sucrose-phosphate synthase and sucrose synthase: expression patterns, metabolic regulation and implications for seed development. Plant J. 9, 841–850 (1996).
Boyer, J. S. & McLaughlin, J. E. Functional reversion to identify controlling genes in multigenic responses: analysis of floral abortion. J. Exp. Bot. 58, 267–277 (2007).
Lauxmann, M. A. et al. Reproductive failure in Arabidopsis thaliana under transient carbohydrate limitation: flowers and very young siliques are jettisoned and the meristem is maintained to allow successful resumption of reproductive growth. Plant Cell Environ. 39, 745–767 (2016).
Liu, Y. H., Offler, C. E. & Ruan, Y. L. Cell wall invertase promotes fruit set under heat stress by suppressing ROS-independent cell death. Plant Physiol. 172, 163–180 (2016).
Seki, M. et al. A mathematical model of phloem sucrose transport as a new tool for designing rice panicle structure for high grain yield. Plant Cell Physiol. 56, 605–619 (2015).
Ruan, Y. L., Patrick, J. W., Bouzayen, M., Osorio, S. & Fernie, A. R. Molecular regulation of seed and fruit set. Trends Plant Sci. 17, 656–665 (2012).
Van Dingenen, J. et al. Limited nitrogen availability has cultivar-dependent effects on potato tuber yield and tuber quality traits. Food Chem. 288, 170–177 (2019).
Olas, J. J. et al. Nitrate acts at the Arabidopsis thaliana shoot apical meristem to regulate flowering time. N. Phytol. 223, 814–827 (2019).
Tegeder, M. & Masclaux-Daubresse, C. Source and sink mechanisms of nitrogen transport and use. N. Phytol. 217, 35–53 (2018).
Fan, X. et al. Plant nitrate transporters: from gene function to application. J. Exp. Bot. 68, 2463–2475 (2017).
O’Brien, J. A. et al. Nitrate transport, sensing, and responses in plants. Mol. Plant 9, 837–856 (2016).
Xuan, W., Beeckman, T. & Xu, G. Plant nitrogen nutrition: sensing and signaling. Curr. Opin. Plant Biol. 39, 57–65 (2017).
Wang, L. & Ruan, Y.-L. Shoot–root carbon allocation, sugar signalling and their coupling with nitrogen uptake and assimilation. Funct. Plant Biol. 43, 105–113 (2016).
Habash, D. Z., Bernard, S., Schondelmaier, J., Weyen, J. & Quarrie, S. A. The genetics of nitrogen use in hexaploid wheat: N utilisation, development and yield. Theor. Appl. Genet. 114, 403–419 (2007).
Martin, A. et al. Two cytosolic glutamine synthetase isoforms of maize are specifically involved in the control of grain production. Plant Cell 18, 3252–3274 (2006).
Cai, H. et al. Overexpressed glutamine synthetase gene modifies nitrogen metabolism and abiotic stress responses in rice. Plant Cell Rep. 28, 527–537 (2009).
Bao, A. et al. Accumulated expression level of cytosolic glutamine synthetase 1 gene (OsGS1;1 or OsGS1;2) alter plant development and the carbon-nitrogen metabolic status in rice. PLoS ONE 9, e95581 (2014).
James, D. et al. Concurrent overexpression of OsGS1;1 and OsGS2 genes in transgenic rice (Oryza sativa L.): impact on tolerance to abiotic stresses. Front. Plant Sci. 9, 786 (2018).
Hu, M. et al. Transgenic expression of plastidic glutamine synthetase increases nitrogen uptake and yield in wheat. Plant Biotechnol. J. 16, 1858–1867 (2018).
Yamaya, T. et al. Genetic manipulation and quantitative-trait loci mapping for nitrogen recycling in rice. J. Exp. Bot. 53, 917–925 (2002).
Tabuchi, M., Abiko, T. & Yamaya, T. Assimilation of ammonium ions and reutilization of nitrogen in rice (Oryza sativa L.). J. Exp. Bot. 58, 2319–2327 (2007).
Ameziane, R., Bernhard, K. & Lightfoot, D. Expression of the bacterial gdhA gene encoding a NADPH glutamate dehydrogenase in tobacco affects plant growth and development. Plant Soil 22, 147–157 (2000).
Abiko, T. et al. Changes in nitrogen assimilation, metabolism, and growth in transgenic rice plants expressing a fungal NADP(H)-dependent glutamate dehydrogenase (gdhA). Planta 232, 299–311 (2010).
Zhou, Y. et al. Over-expression of a fungal NADP (H)-dependent glutamate dehydrogenase PcGDH improves nitrogen assimilation and growth quality in rice. Mol. Breed. 34, 335–349 (2014).
Seiffert, B., Zhou, Z., Wallbraun, M., Lohaus, G. & Möllers, C. Expression of a bacterial asparagine synthetase gene in oilseed rape (Brassica napus) and its effect on traits related to nitrogen efficiency. Physiol. Plant. 121, 656–665 (2004).
Thomsen, H. C., Eriksson, D., Moller, I. S. & Schjoerring, J. K. Cytosolic glutamine synthetase: a target for improvement of crop nitrogen use efficiency? Trends Plant Sci. 19, 656–663 (2014).
Good, A. et al. Engineering nitrogen use efficiency with alanine aminotransferase. Can. J. Bot. 85, 252–262 (2007).
Shrawat, A. K., Carroll, R. T., DePauw, M., Taylor, G. J. & Good, A. G. Genetic engineering of improved nitrogen use efficiency in rice by the tissue-specific expression of alanine aminotransferase. Plant Biotechnol. J. 6, 722–732 (2008).
Beatty, P. H., Carroll, R. T., Shrawat, A. K., Guevara, D. & Good, A. G. Physiological analysis of nitrogen-efficient rice overexpressing alanine aminotransferase under different N regimes. Botany 91, 866–883 (2013).
Pena, P. A. et al. Molecular and phenotypic characterization of transgenic wheat and sorghum events expressing the barley alanine aminotransferase. Planta 246, 1097–1107 (2017).
Snyman, S. J., Hajari, E., Watt, M. P., Lu, Y. & Kridl, J. C. Improved nitrogen use efficiency in transgenic sugarcane: phenotypic assessment in a pot trial under low nitrogen conditions. Plant Cell Rep. 34, 667–669 (2015).
Miyashita, Y., Dolferus, R., Ismond, K. P. & Good, A. G. Alanine aminotransferase catalyses the breakdown of alanine after hypoxia in Arabidopsis thaliana. Plant J. 49, 1108–1121 (2007).
McAllister, C. H. & Good, A. G. Alanine aminotransferase variants conferring diverse NUE phenotypes in Arabidopsis thaliana. PLoS ONE 10, e0121830 (2015).
Santiago, J. P. & Tegeder, M. Connecting source with sink: the role of Arabidopsis AAP8 in phloem loading of amino acids. Plant Physiol. 171, 508–521 (2016).
Zhang, L., Garneau, M. G., Majumdar, R., Grant, J. & Tegeder, M. Improvement of pea biomass and seed productivity by simultaneous increase of phloem and embryo loading with amino acids. Plant J. 81, 134–146 (2015).
Tan, Q., Zhang, L., Grant, J., Cooper, P. & Tegeder, M. Increased phloem transport of S-methylmethionine positively affects sulfur and nitrogen metabolism and seed development in pea plants. Plant Physiol. 154, 1886–1896 (2010).
Carter, A. M. & Tegeder, M. Increasing nitrogen fixation and seed development in soybean requires complex adjustments of nodule nitrogen metabolism and partitioning processes. Curr. Biol. 26, 2044–2051 (2016).
Nunes-Nesi, A., Fernie, A. R. & Stitt, M. Metabolic and signaling aspects underpinning the regulation of plant carbon nitrogen interactions. Mol. Plant 3, 973–996 (2010).
Vincentz, M., Moureaux, T., Leydecker, M. T., Vaucheret, H. & Caboche, M. Regulation of nitrate and nitrite reductase expression in Nicotiana plumbaginifolia leaves by nitrogen and carbon metabolites. Plant J. 3, 315–324 (1993).
Athwal, G. S., Huber, J. L. & Huber, S. C. Phosphorylated nitrate reductase and 14-3-3 proteins. Site of interaction, effects of ions, and evidence for an amp-binding site on 14-3-3 proteins. Plant Physiol. 118, 1041–1048 (1998).
Bachmann, M., Huber, J. L., Liao, P. C., Gage, D. A. & Huber, S. C. The inhibitor protein of phosphorylated nitrate reductase from spinach (Spinacia oleracea) leaves is a 14-3-3 protein. FEBS Lett. 387, 127–131 (1996).
Bachmann, M., McMichael, R. W. Jr., Huber, J. L., Kaiser, W. M. & Huber, S. C. Partial purification and characterization of a calcium-dependent protein kinase and an inhibitor protein required for inactivation of spinach leaf nitrate reductase. Plant Physiol. 108, 1083–1091 (1995).
Bachmann, M. et al. Identification of Ser-543 as the major regulatory phosphorylation site in spinach leaf nitrate reductase. Plant Cell 8, 505–517 (1996).
Kaiser, W. M. & Huber, S. C. Posttranslational regulation of nitrate reductase in higher plants. Plant Physiol. 106, 817–821 (1994).
Figueroa, C. M. et al. Trehalose 6-phosphate coordinates organic and amino acid metabolism with carbon availability. Plant J. 85, 410–423 (2016).
Murchie, E. H., Ferrario-Mery, S., Valadier, M. H. & Foyer, C. H. Short-term nitrogen-induced modulation of phosphoenolpyruvate carboxylase in tobacco and maize leaves. J. Exp. Bot. 51, 1349–1356 (2000).
Le Van, Q. & Champigny, M. L. NO(3) enhances the kinase activity for phosphorylation of phosphoenolpyruvate carboxylase and sucrose phosphate synthase proteins in wheat leaves: evidence from the effects of mannose and okadaic acid. Plant Physiol. 99, 344–347 (1992).
Perchlik, M. & Tegeder, M. Improving plant nitrogen use efficiency through alteration of amino acid transport processes. Plant Physiol. 175, 235–247 (2017).
Perchlik, M. & Tegeder, M. Leaf amino acid supply affects photosynthetic and plant nitrogen use efficiency under nitrogen stress. Plant Physiol. 178, 174–188 (2018).
Stitt, M. & Sonnewald, U. Regulation of metabolism in transgenic plants. Ann. Rev. Plant Physiol. Mol. Biol. 46, 341–336 (1995).
Baena-Gonzalez, E., Rolland, F., Thevelein, J. M. & Sheen, J. A central integrator of transcription networks in plant stress and energy signalling. Nature 448, 938–942 (2007).
Shi, L., Wu, Y. & Sheen, J. TOR signaling in plants: conservation and innovation. Development 145, dev160887 (2018).
Xiong, Y. et al. Glucose-TOR signalling reprograms the transcriptome and activates meristems. Nature 496, 181–186 (2013).
Chen, X. et al. Shoot-to-root mobile transcription factor HY5 coordinates plant carbon and nitrogen acquisition. Curr. Biol. 26, 640–646 (2016).
Kurai, T. et al. Introduction of the ZmDof1 gene into rice enhances carbon and nitrogen assimilation under low-nitrogen conditions. Plant Biotechnol. J. 9, 826–837 (2011).
Yanagisawa, S., Akiyama, A., Kisaka, H., Uchimiya, H. & Miwa, T. Metabolic engineering with Dof1 transcription factor in plants: Improved nitrogen assimilation and growth under low-nitrogen conditions. Proc. Natl Acad. Sci. USA 101, 7833–7838 (2004).
Marchive, C. et al. Nuclear retention of the transcription factor NLP7 orchestrates the early response to nitrate in plants. Nat. Commun. 4, 1713 (2013).
Gaudinier, A. et al. Transcriptional regulation of nitrogen-associated metabolism and growth. Nature 563, 259–264 (2018).
Liu, K. H. et al. Discovery of nitrate-CPK-NLP signalling in central nutrient-growth networks. Nature 545, 311–316 (2017).
Guan, P. et al. Interacting TCP and NLP transcription factors control plant responses to nitrate availability. Proc. Natl Acad. Sci. USA 114, 2419–2424 (2017).
Olas, J. J. & Wahl, V. Tissue-specific NIA1 and NIA2 expression in Arabidopsis thaliana. Plant Signal. Behav. 14, 1656035 (2019).
Navarro, C. et al. Control of flowering and storage organ formation in potato by FLOWERING LOCUS T. Nature 478, 119–122 (2011).
Kloosterman, B. et al. Naturally occurring allele diversity allows potato cultivation in northern latitudes. Nature 495, 246–250 (2013).
Abelenda, J. A., Cruz-Oro, E., Franco-Zorrilla, J. M. & Prat, S. Potato StCONSTANS-like1 suppresses storage organ formation by directly activating the FT-like StSP5G repressor. Curr. Biol. 26, 872–881 (2016).
Sharma, P., Lin, T. & Hannapel, D. J. Targets of the StBEL5 transcription factor include the FT ortholog StSP6A. Plant Physiol. 170, 310–324 (2016).
Kondhare, K. R., Vetal, P. V., Kalsi, H. S. & Banerjee, A. K. BEL1-like protein (StBEL5) regulates CYCLING DOF FACTOR1 (StCDF1) through tandem TGAC core motifs in potato. J. Plant Physiol. 241, 153014 (2019).
Banerjee, A. K., Prat, S. & Hannapel, D. J. Efficient production of transgenic potato (S. tuberosum L. ssp. andigena) plants via Agrobacterium tumefaciens-mediated transformation. Plant Sci. 170, 732–738 (2006).
Gonzalez-Schain, N. D., Diaz-Mendoza, M., Zurczak, M. & Suarez-Lopez, P. Potato CONSTANS is involved in photoperiodic tuberization in a graft-transmissible manner. Plant J. 70, 678–690 (2012).
Hannapel, D. J. & Banerjee, A. K. Multiple mobile mRNA signals regulate tuber development in potato. Plants 6, 8 (2017).
Taoka, K. et al. 14-3-3 proteins act as intracellular receptors for rice Hd3a florigen. Nature 476, 332–335 (2011).
Teo, C. J., Takahashi, K., Shimizu, K., Shimamoto, K. & Taoka, K. I. Potato tuber induction is regulated by interactions between components of a tuberigen complex. Plant Cell Physiol. 58, 365–374 (2017).
Lunn, J. E. et al. Sugar-induced increases in trehalose 6-phosphate are correlated with redox activation of ADPglucose pyrophosphorylase and higher rates of starch synthesis in Arabidopsis thaliana. Biochem. J. 397, 139–148 (2006).
Figueroa, C. M. & Lunn, J. E. A tale of two sugars: trehalose 6-phosphate and sucrose. Plant Physiol. 172, 7–27 (2016).
Eastmond, P. J. et al. Trehalose-6-phosphate synthase 1, which catalyses the first step in trehalose synthesis, is essential for Arabidopsis embryo maturation. Plant J. 29, 225–235 (2002).
Gomez, L. D., Gilday, A., Feil, R., Lunn, J. E. & Graham, I. A. AtTPS1-mediated trehalose 6-phosphate synthesis is essential for embryogenic and vegetative growth and responsiveness to ABA in germinating seeds and stomatal guard cells. Plant J. 64, 1–13 (2010).
van Dijken, A. J., Schluepmann, H. & Smeekens, S. C. Arabidopsis trehalose-6-phosphate synthase 1 is essential for normal vegetative growth and transition to flowering. Plant Physiol. 135, 969–977 (2004).
Wahl, V. et al. Regulation of flowering by trehalose-6-phosphate signaling in Arabidopsis thaliana. Science 339, 704–707 (2013).
Sanz, M. J., Mingo Castel, A., van Lammeren, A. A. M. & Vreugdenhil, D. Changes in the microtubular cytoskeleton precede in vitro tuber formation in potato. Protoplasma 191, 46–54 (1996).
Chincinska, I. A. et al. Sucrose transporter StSUT4 from potato affects flowering, tuberization, and shade avoidance response. Plant Physiol. 146, 515–528 (2008).
Abelenda, J. A. et al. Source-sink regulation is mediated by interaction of an FT homolog with a SWEET protein in potato. Curr. Biol. 29, 1178–1186 (2019).
Viola, R. et al. Tuberization in potato involves a switch from apoplastic to symplastic phloem unloading. Plant Cell 13, 385–398 (2001).
Hancock, R. D. et al. Physiological, biochemical and molecular responses of the potato (Solanum tuberosum L.) plant to moderately elevated temperature. Plant Cell Environ. 37, 439–450 (2014).
Hastilestari, B. R. et al. Deciphering source and sink responses of potato plants (Solanum tuberosum L.) to elevated temperatures. Plant Cell Environ. 41, 2600–2616 (2018).
Lehretz, G. G., Sonnewald, S., Hornyik, C., Corral, J. M. & Sonnewald, U. Post-transcriptional Regulation of FLOWERING LOCUS T modulates heat-dependent source-sink development in potato. Curr. Biol. 29, 1614–1624 (2019).
Fridman, E., Carrari, F., Liu, Y. S., Fernie, A. R. & Zamir, D. Zooming in on a quantitative trait for tomato yield using interspecific introgressions. Science 305, 1786–1789 (2004).
Bermudez, L. et al. Silencing of the tomato sugar partitioning affecting protein (SPA) modifies sink strength through a shift in leaf sugar metabolism. Plant J. 77, 676–687 (2014).
Soyk, S. et al. Bypassing negative epistasis on yield in tomato imposed by a domestication gene. Cell 169, 1142–1155 (2017).
Soyk, S. et al. Variation in the flowering gene SELF PRUNING 5G promotes day-neutrality and early yield in tomato. Nat. Genet. 49, 162–168 (2017).
Park, S. J. et al. Optimization of crop productivity in tomato using induced mutations in the florigen pathway. Nat. Genet. 46, 1337–1342 (2014).
Liu, J. et al. The conserved and unique genetic architecture of kernel zize and weight in maize and rice. Plant Physiol. 175, 774–785 (2017).
Siebers, T., Catarino, B. & Agusti, J. Identification and expression analyses of new potential regulators of xylem development and cambium activity in cassava (Manihot esculenta). Planta 245, 539–548 (2017).
Hannapel, D. J., Sharma, P., Lin, T. & Banerjee, A. K. The multiple signals that control tuber formation. Plant Physiol. 174, 845–856 (2017).
Villordon, A. Q., Ginzberg, I. & Firon, N. Root architecture and root and tuber crop productivity. Trends Plant Sci. 19, 419–425 (2014).
Smetana, O. et al. High levels of auxin signalling define the stem-cell organizer of the vascular cambium. Nature 565, 485–489 (2019).
Shi, D., Lebovka, I., Lopez-Salmeron, V., Sanchez, P. & Greb, T. Bifacial cambium stem cells generate xylem and phloem during radial plant growth. Development 146, dev171355 (2019).
Eviatar-Ribak, T. et al. A cytokinin-activating enzyme promotes tuber formation in tomato. Curr. Biol. 23, 1057–1064 (2013).
Matsumoto-Kitano, M. et al. Cytokinins are central regulators of cambial activity. Proc. Natl Acad. Sci. USA 105, 20027–20031 (2008).
Etchells, J. P. & Turner, S. R. The PXY-CLE41 receptor ligand pair defines a multifunctional pathway that controls the rate and orientation of vascular cell division. Development 137, 767–774 (2010).
Miyashima, S. et al. Mobile PEAR transcription factors integrate positional cues to prime cambial growth. Nature 565, 490–494 (2019).
Gancheva, M. S. et al. Identification, expression, and functional analysis of CLE genes in radish (Raphanus sativus L.) storage root. BMC Plant Biol. 16(Suppl. 1), 7 (2016).
Kaachra, A., Vats, S. K. & Kumar, S. Heterologous expression of key C and N metabolic enzymes improves re-assimilation of photorespired CO2 and NH3, and growth. Plant Physiol. 177, 1396–1409 (2018).
Naqvi, S. et al. Transgenic multivitamin corn through biofortification of endosperm with three vitamins representing three distinct metabolic pathways. Proc. Natl Acad. Sci. USA 106, 7762–7767 (2009).
Bachem, C. W. B., van Eck, H. J. & de Vries, M. E. Understanding genetic load in potato for hybrid diploid breeding. Mol. Plant 12, 896–898 (2019).
Zhang, C. et al. The genetic basis of inbreeding depression in potato. Nat. Genet. 51, 374–378 (2019).
Ghosh, S. et al. Speed breeding in growth chambers and glasshouses for crop breeding and model plant research. Nat. Protoc. 13, 2944–2963 (2018).
Fernie, A. R. & Yan, J. De novo domestication: an alternative route toward new crops for the future. Mol. Plant 12, 615–631 (2019).
Mehdi, R. et al. Symplasmic phloem unloading and radial post-phloem transport via vascular rays in tuberous roots of Manihot esculenta. J. Exp. Bot. 70, 5559–5573 (2019).
Ikematsu, S., Tasaka, M., Torii, K. U. & Uchida, N. ERECTA-family receptor kinase genes redundantly prevent premature progression of secondary growth in the Arabidopsis hypocotyl. N. Phytol. 213, 1697–1709 (2017).
Kubo, M. et al. Transcription switches for protoxylem and metaxylem vessel formation. Genes Dev. 19, 1855–1860 (2005).
Knott, J. M., Romer, P. & Sumper, M. Putative spermine synthases from Thalassiosira pseudonana and Arabidopsis thaliana synthesize thermospermine rather than spermine. FEBS Lett. 581, 3081–3086 (2007).
Ge, C. et al. BUD2, encoding an S-adenosylmethionine decarboxylase, is required for Arabidopsis growth and development. Cell Res. 16, 446–456 (2006).
Research in the authors’ laboratories was supported by the following grants: the cassava source–sink (CASS) project of the Bill and Melinda Gates Foundation (to A.R.F., H.E.N., M.S. and U.S.); the ERA-CAPs project SourSi (to A.R.F. and L.J.S.); the BIO2015-3019-EXP grant from the Spanish Ministry of Economy, Industry and Competitiveness and the PCIN-2017-032 CONCERT-JAPAN project financed by the Ministry of Science, Innovation and Universities (to S.P.); Australian Research Council DP180103834 (to Y.L.R.); the US National Science Foundation (grant no. IOS-1457183); the Agriculture and Food Research Initiative (AFRI; grant no. 2017-67013-26158) from the USDA National Institute of Food and Agriculture (to M.T.); the Finnish Centre of Excellence in Molecular Biology of Primary Producers (Academy of Finland CoE program 2014–2019; grant no. 271832); the Gatsby Foundation (grant no. GAT3395/PR3); the University of Helsinki (grant no. 799992091); the European Research Council Advanced Investigator Grant SYMDEV (grant no. 323052; to Y.H.); the BMBF (grant no. 031B0191); the DFG (SPP1530: WA3639/1-2, 2-1); and the Max-Planck-Society (to V.W.). We additionally thank D. Ko and R. Ruonala for their comments on the manuscript.
The authors declare no competing interests.
Peer review information Nature Plants thanks Christine Foyer and the other, anonymous, reviewers for their contribution to the peer review of this work.
Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
About this article
Cite this article
Fernie, A.R., Bachem, C.W.B., Helariutta, Y. et al. Synchronization of developmental, molecular and metabolic aspects of source–sink interactions. Nat. Plants 6, 55–66 (2020). https://doi.org/10.1038/s41477-020-0590-x
A source-sink model explains the difference in the metabolic mechanism of mechanical damage to young and senescing leaves in Catharanthus roseus
BMC Plant Biology (2021)
Source–sink manipulations differentially affect carbon and nitrogen dynamics, fruit metabolites and yield of Sacha Inchi plants
BMC Plant Biology (2021)
Transcriptome integrated metabolic modeling of carbon assimilation underlying storage root development in cassava
Scientific Reports (2021)
Theoretical and Applied Genetics (2021)
Plant Growth Regulation (2021)