Focus Article
Nutritional regulation of root development León Francisco Ruiz Herrera,1 Michael W. Shane2 and José López-Bucio1∗ Mineral nutrients such as nitrogen (N), phosphorus (P), and iron (Fe) are essential for plant growth, development, and reproduction. Adequate provision of nutrients via the root system impacts greatly on shoot biomass and plant productivity and is therefore of crucial importance for agriculture. Nutrients are taken up at the root surface in ionic form, which is mediated by specific transport proteins. Noteworthy, root tips are able to sense the local and internal concentrations of nutrients to adjust growth and developmental processes, and ultimately, to increase or decrease the exploratory capacity of the root system. Recently, important progress has been achieved in identifying the mechanisms of nutrient sensing in wild- and cultivated species, including Arabidopsis, bean, maize, rice, lupin as well as in members of the Proteaceae and Cyperaceae families, which develop highly sophisticated root clusters as adaptations to survive in soils with very low fertility. Major findings include identification of transporter proteins and transcription factors regulating nutrient sensing, miRNAs as mobile signals and peptides as repressors of lateral root development under heterogeneous nutrient supply. Understanding the roles played by N, P, and Fe in gene expression and biochemical characterization of proteins involved in root developmental responses to homogeneous or heterogeneous N and P sources has gained additional interest due to its potential for improving fertilizer acquisition efficiency in crops. © 2015 Wiley Periodicals, Inc. How to cite this article:
WIREs Dev Biol 2015. doi: 10.1002/wdev.183
INTRODUCTION
C
onsiderable information has been gathered in the last decade on plant responses regulated by nutrient signaling, including: (1) elucidation of hormonal pathways with auxin as a central player mediating changes in root architecture by nutrient availability,1–3 (2) identification of transporter proteins for nutrient uptake, and transcription factors as master regulators of root architecture,4–6 (3) the involvement of ubiquitin system components in nutrient signaling,7,8 (4) identification of miRNAs as mobile signals,9–13 and ∗ Correspondence
to: [email protected]
1 Instituto
de Investigaciones Químico-Biológicas, Universidad Michoacana de San Nicolás de Hidalgo, Edificio A-1’, Ciudad Universitaria Morelia, Michoacán, México
2 School
of Plant Biology, Faculty of Science, University of Western Australia, Crawley, Australia Conflict of interest: The authors have declared no conflicts of interest for this article.
(5) characterization of peptides as repressors of lateral root development under heterogeneous nutrient supply.14 Several of these responses are systemically controlled by leaf nutrient status, whereas others are locally regulated by nutrient availability in the root environment.15–17 This focus article highlights signaling role(s) of N, P, and Fe in controlling root development. Nitrate (NO−3 ) and ammonium (NH+4 ) are the major sources of N taken up by roots, and regulate gene expression, energy transfer, protein activation, as well as metabolic and physiological processes.17 P is taken up by roots as fully oxidized soluble phosphate (Pi), which is a component of many key biomolecules such as nucleic acids and phospholipids and thus participates in various enzymatic reactions and metabolic pathways.18 Fe is an essential micronutrient for virtually all organisms because it functions as a component of many enzymes and proteins and its deficiency can
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block the synthesis of chlorophyll precursors, inducing leaf yellowing and reducing photosynthesis.19 As each of these elements is of vital importance for plant growth and development, plant root systems evolved the ability to acclimate to nutrient stress by eliciting a complex array of morphological and biochemical adaptations to optimize N, P, and Fe uptake capacity, in which localized cell division, elongation and differentiation function coordinately to maximize soil exploration and root nutrient foraging.20,21
NITRATE AND AMMONIUM DIFFERENTIALLY MODULATE LATERAL ROOT DEVELOPMENT Arabidopsis and crop seedlings can modify their root architectures to acquire sources of N that are uniformly or unevenly distributed in soil. One important response involves increased growth of lateral roots (LR) within patches enriched in NO−3 or greater LR formation to uniformly supplied NH+4 ions (Figure 1(a)).22,23 The stimulation of LR elongation in NO−3 -rich patches is the result of a signaling effect of the external NO−3 ion itself rather than its downstream metabolites glutamine or glutamate. An important breakthrough in understanding the role of NO−3 in stimulating LR growth was the identification of the ARABIDOPSIS NITRATE-REGULATED1 (ANR1) gene, a member of the plant MADS-box family of transcription factors, which is a vital component in this regulatory signaling pathway (Figure 1(a)).22 Transgenic plants in which ANR1 is repressed show altered suppression of LR growth in NO−3 -rich patches.22 LR primordia initiation is strongly repressed in Arabidopsis by the combination of high external sucrose and low external NO−3 , while sucrose alone appears to act at the level of meristem activation.23 Local supply of NH+4 to Arabidopsis plants increases LR formation. This effect is independent from cumulative uptake of (15)N-labeled NH+4 , suggesting that this process is unlikely due to a nutritional effect but most likely occurs via a sensing event.24 In contrast to wild-type plants, NH+4 -induced LR formation was almost absent in a quadruple AMMONIUM TRANSPORTER mutant qko (amt1;1 amt1;2 amt1;3 amt2;1) and significantly lower in the amt1;3 single mutant.24 To define long-distance signals that affect root growth in response to homogenous or heterogenous N supply, a split-root design has been applied.25,26 On the root experiencing N-limited conditions, both the rate of LR initiation and further LR growth is decreased, while in the N-rich
LRs proliferate (Figure 1(b)). These effects correlated with altered cytokinin biosynthesis and transcriptome reprograming.25 The transcription factor TEOSINTE BRANCHED1/CYCLOIDEA/ PROLIFERATING CELL FACTOR20 (TCP20) was implicated in NO−3 signaling by its ability to bind the promoters in more than 100 NO−3 -regulated genes including NIA1, NRT2.1, and NRT1.1.26 Analysis of insertion mutants of TCP20 showed that they had normal primary and LR growth under uniform supply of NO−3 , but were impaired in preferential LR growth supplied with high NO−3 concentration. Inhibition of LR growth in the high NO−3 was still evident in the mutants even when NH+4 was supplied in the opposite halve, indicating that the TCP20 response was specific to NO−3 . Thus, homogeneous or heterogeneous N sources interfere with LR development apparently acting in different ways.
AUXIN, MICRO RNAs AND PEPTIDE SIGNALING ARE CRITICAL FOR NITROGEN REGULATED ROOT GROWTH The auxin receptor AFB3 and downstream components are essential to regulate LR responses to local supply of NO−3 , likely via miR393 and miR167 action.2,27–29 AFB3 is induced by NO−3 and repressed by miR393 in roots. In contrast, miR393 is not induced in a nitrate reductase (NR)-null mutant of Arabidopsis, suggesting that N metabolites produced by NO−3 reduction and assimilation acts in a negative feedback loop regulating AFB3 levels over time according to N availability. AFB3 acts specifically in the context of the NO−3 response by regulating a connected gene network under control of the NAC4 transcription factor.2 Lateral root responses to N have also been shown to depend on miR167 and its target, the auxin response factor ARF8 mRNA.28 This module acts specifically in pericycle cells in a positive manner to control LR initiation and later in development by decreasing the final length of LRs. Downstream of auxin, nitric oxide (NO) homeostasis appears to be a key regulator of early NO−3 perception and root elongation.30 Accordingly, NO production by NR increased with NO−3 supply in the elongation zone of maize roots and correlated with increased rates of growth. Decreased levels of NO in roots affected their elongation under high NO−3 conditions, whereas treatment of NO−3 -deprived roots with sodium nitroprusside (SNP), a NO-releasing compound, stimulated root growth. An increased formation of LRs in response to homogenous low NO−3 availability depends on
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FIGURE 1 | Regulatory elements involved in LR growth in response to homogeneous or heterogeneous N sources. Homogenous supply of NH+4
triggers lateral root initiation, while NO−3 -rich patches stimulates lateral root elongation. (a) In Arabidopsis, NRT1.1 is a dual-affinity NO−3 transporter involved in both high- and low-affinity NO−3 uptake. The switch between the two affinities is controlled by phosphorylation at the T101, and this phosphorylation is regulated by the calcineurin B-like-interacting protein kinase CIPK23. NRT1.1 promotes LR growth in NO−3 -rich patches likely through the transcription factor ANR1. NO−3 levels affect root development through modulation of auxin synthesis and signaling via TAR2, AFB3 and ARF8. Solid arrows indicate positive effects, blunt solid arrows indicate negative effects, and dashed lines indicate possible interactions. (b) Root foraging for NO−3 involves both local and systemic signaling. Roots respond to low local levels of NO−3 by producing CEP1 and CLE3 peptides, which are repressors of lateral root growth. CEP1 moves upward to the shoot and is received by the LRR-RKs proteins CEPR1 and CEPR2. Long-distance signaling to the root halve grown under high NO−3 involves cytokinin (CK) biosynthesis and requires the transcription factor TCP20, which activates the expression of genes involved in N uptake and is necessary for the induction of development.
the function of the auxin biosynthesis gene TAR2 (tryptophan aminotransferase-related 2).3 TAR2 is expressed in the pericycle of the primary root and is induced under low N conditions. In wild-type
plants, low N stimulated auxin accumulation in LR primordia at early developmental stages. Conversely, auxin accumulation and LR growth were impaired in the tar2 null mutant. Overexpression of TAR2
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leads to increased LR numbers under both high and low N conditions.3 All together, these reports show that AFB3 and downstream signaling proteins may increase auxin accumulation and/or auxin sensitivity in LR primordia depending on NO−3 or NH+4 availability and thus activate root branching. Several homologs of the CLE (CLAVATA3/ ESR-related) gene family are upregulated by N deficiency. CLAVATA3 (CLV3) is known as a peptide that binds to the CLAVATA1 (CLV1) receptor and regulates shoot-cell proliferation and differentiation.31 Interestingly, the signaling pathway composed of N-responsive CLE peptides and the CLAVATA1 (CLV1) leucine-rich repeat receptor-like kinase is expressed in the Arabidopsis root vasculature and pericycle. Overexpression of CLE1, −3, −4, and −7 repressed the development of LR primordia. In contrast, clv1 mutants showed progressive outgrowth of primordia from the parent root under N-deficient conditions.31 These data implicate the N-responsive CLE-CLV1 signaling module as an essential mechanism restrictively controlling root branching in N-deficient environments. N-deprived roots also secrete small peptides belonging to the Arabidopsis C-terminally encoded peptide (CEP) family, which share short, conserved domains near the C terminus, a feature that is common to several modified small peptides from plants.14 CEP1 was used to screen a library of cultured plant cells expressing all known plant leucine-rich repeat receptor kinase (LRR-RK)—encoding genes and identified two homologous LRR-RKs (CEPR1 and CEPR2) that efficiently bind CEP1.14 Exposure of roots to low N increased CEP1 levels in the xylem sap of the shoot, indicating its long-distance transport to the shoot (Figure 1(b)). Arabidopsis plants deficient in this signaling pathway show decreased growth accompanied with N-deficiency symptoms, thus indicating that signaling from the root to the shoot via a peptide—LRR-RK relay helps the plant adapt to fluctuations in local N availability.
ROLE OF NITRATE AND AMMONIUM TRANSPORTERS IN NITROGEN SENSING BY ROOTS The NRT1.1 nitrate transporter plays a key role in NO−3 sensing. NRT1.1 loss of function causes a decreased root proliferation in NO−3 patches, resulting from reduced LR elongation (Figure 1(a)). This response is not due to lower specific NO−3 uptake activity in the nrt1 mutants, but is associated with dramatically decreased ANR1 expression in
LR primordia and in the tips of the LRs.4 Intriguingly, NRT1.1 not only transports NO−3 but also facilitates uptake of auxin, while NO−3 inhibits NRT1.1-dependent auxin uptake.32 These results show that NRT1.1 promotes localized root growth either as a NO−3 sensor, as a facilitator of NO−3 influx into cells responsible of local N perception, or via local cell-specific auxin accumulation. However, mutations of a second NO−3 transporter NRT2.1 have been shown to repress or stimulate LR initiation depending on NO−3 and sucrose supply,23,33 thus implying NO3 transport in the regulation of LR formation via photosynthate supply to roots. Cereals contain a large PTR/NRT1 gene family (PTR, di/tripeptide transporter; NRT1, low-affinity nitrate transporter) for the uptake and translocation of NO−3 , NH+4 , or small peptides. Rice plants have at least 84 PTR/NRT1 genes but, OsPTR9 is one of the few that has been functionally characterized as having a role in root development.34,35 Over-expression of OsPTR9 in transgenic rice plants resulted in increased NH+4 uptake, and LR formation. Conversely, downregulation of OsPTR9 in a T-DNA insertion line (osptr9) and in OsPTR9-RNAi rice plants had the opposite effect. These results indicate that OsPTR9 might influence root surface area for nutrient absorption by affecting LR density.35
PHOSPHORUS DEFICIENCY INHIBITS PRIMARY ROOT GROWTH AND INCREASES ROOT-ABSORPTIVE CAPACITY Plants can modify their root architecture to improve Pi acquisition in P-deficient acid or alkaline soils in which Pi is precipitated out of the soil solution due to combination with cations. Early studies in Arabidopsis and crops have demonstrated that Pi deficiency affects root development by reducing growth of primary roots while increasing the number and length of LRs and root hairs (Figure 2(a)–(d)).36,37 Modifications in root growth angle may also enhance root foraging and exploration in upper soil layers.38,39 This response is associated with litter decomposition, so P accumulates as mostly organic Pi, but with some inorganic Pi as well.39 The first visible event upon low Pi stress in the primary root is a strong reduction of cell elongation, which occurs within hours after plants are transferred from a relatively high to low Pi-containing medium.16 This is followed by a reduction of cell division as revealed by the cell-cycle marker CYCB1;1 in parallel with loss of quiescent center (QC) identity as reported by the QC46 marker. Eventually, differentiation
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FIGURE 2 | Arabidopsis root architectural responses to low Pi stress. Representative root architectures of seedlings grown for 12 days on medium supplied with (a) 1000 μM (high Pi) or (b) 1 μM (low Pi). Note the highly branched root systems in low Pi conditions due to proliferation of LR. Expression of Pi transporter AtPT2 in primary root tips of seedlings grown under high Pi (c) or low Pi (d) supply coincides with high root-hair proliferation capacity. (e) Phenotypes of Arabidopsis WT (Col-0) and low phosphorus insensitive 1 mutant seedlings grown under low P stress. Note the long primary roots developed by the mutants indicative of altered perception of low Pi availability.
events such as root-hair formation increase dramatically further increasing the root absorptive capacity (Figure 2(d)).40 Thus, the primary roots of plants grown under low Pi stress initiate a developmental program that blocks primary root elongation and favors LR formation and root hair proliferation. Genes and proteins that control primary root growth and LR development under low Pi stress have been identified through screening primary root growth in low P medium. Characterization of alleles of BIG, a protein required for normal auxin transport in Arabidopsis showed that this protein is required for pericycle cell activation to form LR in both high (1 mM) and low (1 μM) P conditions, but not for the low P-induced alterations in primary root growth and root hair elongation.41 In contrast, the low Pi-insensitive (LPI) mutants show the typical increase in LR and root hair number under low P conditions but the primary root growth is unaffected (Figure 2(e)), thus implying LPI genes in sensing Pi deficiency (Figure 3).15 Loss-of-function mutations in LOW PHOSPHATE ROOT1 (LPR1) and its close paralog LPR2 strongly reduce the primary root growth inhibition by low Pi stress similarly to the phenotype shown by lpi mutants.16 Interestingly, LPR1 is expressed in the root cap, the possible region sensing Pi deficiency. LPR1 and LPR2 encode multicopper oxidases (MCOs), enzymes that oxidize their substrates with the concomitant reduction of dioxygen to two water molecules.16 Human MCOs possess both
ferroxidase and Fe2+ ferroportin activities and play a role in Fe metabolism.42 The phosphate deficiency response 2 (pdr2) mutant seedlings display hypersensitive responses to Pi deficiency evidenced by conditional short-root phenotype caused by selective inhibition of root cell division followed by meristem death under low Pi conditions.43 Higher density of LRs is observed in pdr2 mutants under low Pi stress, indicating that PDR2 is necessary both for maintenance of root meristem function and pericycle cell activation when Pi availability is limited. PDR2 encodes the single P(5)-type ATPase of Arabidopsis thaliana.44 PDR2 functions in the endoplasmic reticulum (ER) and is required for proper expression of SCARECROW (SCR), a key regulator of root patterning, and for stem-cell maintenance in Pi-deprived roots. Indeed, the MCO encoded by LPR1 is targeted to the ER and interacts with PDR2.44 Because the expression domains of both genes overlap in the stem-cell niche and root meristem, PDR2 and LPR1 may function together in an ER-resident pathway that adjusts root meristem activity to external Pi sensing (Figure 3).
AUXIN MEDIATES ROOT ARCHITECTURE REMODELING BY PHOSPHORUS DEPRIVATION Accumulating evidence suggests that auxins modulate the responses of root architecture to low Pi stress. This was first supported by a study showing that
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Anthocyanin accumulation Nucleus
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Low P sensing IAA14 QC identity and meristem division (CYCB1;1)
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FIGURE 3 | Phosphorus starvation signaling pathways regulate primary and LR development. Pi is sensed in root tips by an unknown P starvation sensor, which triggers the signaling pathways that modulate root architecture. Low Pi promotes LR proliferation through increasing auxin transport and BIG function, and auxin signaling that involves TIR1, IAA14, ARF7, and ARF19. SLR1/IAA14 is a member of Aux/IAA family that represses the activity of ARF7 and ARF19 in lateral root initiation. The effects of low Pi stress on cell division and differentiation is regulated through the transcription factor SCR and LPR and PDR proteins. Other transcription factors such as WRKY75 regulate LR proliferation and anthocyanin accumulation.
exogenous application of auxins to P-adequate roots resulted in localized alterations in root architecture in Arabidopsis that mimicked those observed under low P growth conditions.37 Another study showed that LR proliferation in roots of plants under low Pi stress required normal auxin transport mediated by BIG.41 The auxin signalling pathway also contributes to the Pi deficiency-induced cluster-root formation in white lupin (Lupinus albus L).45,46 Compelling evidence indicates that the pattern of LR formation and emergence in response to low Pi stress is mediated by changes in auxin sensitivity in roots.1 This study showed that low Pi signaling altered the expression of auxin-responsive genes and stimulated pericycle cells to divide. Modulation of auxin sensitivity by Pi was found to depend on the auxin receptor TRANSPORT INHIBITOR RESPONSE 1 (TIR1) and the transcription factor AUXIN RESPONSE FACTOR 19 (ARF19) (Figure 3). Low Pi stress increases the expression of TIR1 and causes AUXIN/INDOLE-3-ACETIC ACID (AUX/IAA) auxin response repressors to be degraded.1 It was proposed that auxin sensitivity is enhanced in Pi-deprived plants by an increased expression of TIR1, which accelerates the degradation of AUX/IAA proteins, thereby unshackling ARF transcription factors that activate/repress genes involved in LR initiation and growth.
TRANSCRIPTION FACTORS MEDIATE ROOT ARCHITECTURAL RESPONSES TO PHOSPHORUS AND IRON AVAILABILITY Several transcription factors act downstream or in concert with auxin signaling to mediate root responses to low Pi stress. RNAi knockdown studies showed that a reduction in WRKY75 mRNA levels caused early anthocyanin accumulation and reduced Pi uptake during Pi starvation.5 Silencing of WRKY75 also caused an increase in LR and root-hair growth that was independent of Pi status, suggesting WRKY75 is a modulator of root development. Microarray data from Pi-deficient rice identified an R2R3 MYB transcription factor, OsMYB2P-1.47 Primary roots of OsMYB2P-1-overexpressing plants were shorter than those in wild-type plants under Pi-sufficient conditions, while primary roots and adventitious roots of OsMYB2P-1-overexpressing plants were longer than those of wild-type plants under low Pi conditions. In contrast to the accumulated information about the regulatory networks controlled by N or P availability that shape root form and function, less is known about root responses to Fe. Noteworthy, plants exposed to Fe deprivation display increased primary root waving and LR initiation, with the
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FIGURE 4 | Iron signaling regulates root architecture. Fe is taken up into the symplast by IRT1 transporters at the root surface. Reduction of Fe by FRO2, and FRO5 reductases, contribute to increased Fe acquisition. Fe can diffuse through the symplastic space to the pericycle and then to the vasculature. BTS and PYE, a putative E3 ligase and a bHLH transcription factor, respectively, are specifically induced in root pericycle cells under Fe deficiency. PYE and other bHLHs transcription factors likely regulate the genes responsive to activate pericycle cells for LR initiation under Fe deprivation. Fe translocation inside the plant is associated with suitable chelating molecules such as organic acids and loading from vasculature to mesophyll tissue occurs via YSL transporters.
pericycle acting as a regulatory center for sensing and responding to local Fe availability (Figure 4).6 Whole root time–course microarray analyses showed that numerous transcription factors are temporally coexpressed under Fe deficiency. A screen of mutants of these transcription factors looking for alterations in metabolic and developmental responses to Fe deficiency identified a bHLH transcription factor named POPEYE (PYE), and a second gene tightly coregulated with PYE called BRUTUS (BTS).6 PYE appears to play an important role in root responses to Fe-deficient conditions, as pye1 mutant seedlings display decreased tolerance to Fe deprivation in primary root growth and are more sensitive to develop leaf chlorosis.
ROLE OF NUTRIENTS IN ROOT CLUSTER FORMATION: SIMILARITY IN FUNCTION, DIVERSITY IN STRUCTURE While basic knowledge about the pathways involved in root development by nutrient availability has been obtained from Arabidopsis, root clusters are particularly remarkable for their diversity of morphological structure and roles in plant adaptation to soils with very low nutrient availability (Figure 5). These highly
specialised ephemeral root structures evolved to perform critical functions in nutrient acquisition. They occur on the root systems of a range of perennial dicotyledonous (e.g., ‘proteoid roots’ in Proteaceae; Figure 5(a) and (d) and monocotyledonous (‘dauciform roots’ in Cyperaceae; Figures 5(c) and 6) native plants, but are also formed on several agriculturally significant crops (e.g., ‘cluster roots’ in Fabaceae; Figure 5(b)).48 Proteoid root clusters are characterised by incredibly dense clusters of numerous short, determinate rootlets, covered by root hairs at maturity. Superficially, proteoid roots exhibit a huge range of morphologies but most common are ‘tree-like’ compound clusters (e.g., Banksia species; Figure 5(a)) or ‘bottle-brush-like’ simple proteoid-roots (e.g., Lupinus albus, Figure 5(b)). Dauciform root clusters also exhibit a vast array of structural diversity, but are characterized by short, carrot shaped LRs with remarkably dense and long root hairs (Figure 5(c)). The formation of root clusters is most highly represented amongst nonmycorrhizal plants that are particularly well-adapted to soils with low Pi availability.49,50 Detailed developmental studies have shown that root clusters can be physiologically active for as little as 10 days or up to 4 weeks.50–52,48 For example, after initiation, dauciform roots in Schoenus unispiculatus reach maturity within ∼ 7 days
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FIGURE 5 | Root-cluster morphologies in Proteaceae, Cyperaceae, and Fabaceae. All species are well adapted to soils of extremely low P
concentrations. Plants were grown in nutrient solutions containing ≤1 μM phosphate. (a) Proteoid roots (compound type) of Banksia prionotes (Proteaceae). (b) Cluster roots (simple type) of Lupinus albus L. (Fabaceae). (c) Dauciform roots of Schoenus unispiculatus (Cyperaceae). (d) Proteoid roots of Hakea prostrata (simple type, Proteaceae). (e) Stages of dauciform root development in S. unispiculatus. Dauciform roots were initially white, but often turned bright yellow as they matured. Remarkably large numbers of long root hairs with a similar final length were fully developed after ca. 7–9 days. The axes of very young dauciform roots (1–3 days old) illustrate the ‘carrot shape’ of the dauciform root axes, and a slender stalk connected the carrot-shaped axis to the main root. After 10–12 days, the dauciform roots senesced. (f) Stages of proteoid-root development in H . prostrata from rootlet initiation to senescence (∼20 days old, far right). Scale bars in mm (a) 10; (b) 4; (c) 3; (d) 11; (e) 5; and (f) 9.
(Figure 5(e)), whereas proteoid-roots of Hakea prostrata (Proteaceae) require approximately 14 days (Figure 5(f)). The ephemeral nature of root clusters necessitates their continual replacement by sequential formation along axes of growing LRs (Figures 5(d) and 6), or as a singular structure upon short LRs as illustrated in the time–course of dauciform root development in an Australian sedge (Figure 5(e)).53
Phosphorus has been repeatedly shown to be a key nutrient influencing initiation (low Pi supply) or suppression (higher Pi supply) of root clusters (Figure 6), but their formation is also influenced by N and Fe.54 Split-root or foliar feeding experiments suggest that the extent of root-cluster formation can be determined by internal (leaf) Pi status or Pi availability in the external root environment in a species-specific manner. With little exception, the root architecture
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FIGURE 6 | Influence of phosphate supply on dauciform-root formation in the south-western Australian native Cyperaceae Carex fascicularis. The formation of highly specialised dauciform roots increases dramatically on plants as the Pi level is decreased in nutrient solution (from left to right). This particular species of Carex develops relatively small dauciform roots in series (arrows) along the axes of growing LR (cf . Figure 5(c) and (e)). Scale bar; 13 mm.
of field-grown root cluster-forming species is almost always patterned toward greater abundance of root clusters, and hence, root-foraging in upper soil horizons where levels of nutrients are often enriched.55 Formation of dense clusters of rootlets and/or root hairs in a small soil volume markedly increases root surface area for nutrient foraging;20,48 however, those ‘zones’ from which rootlets or root hairs acquire Pi and micronutrients greatly overlap. This morphology is likely to have been customized during evolution to allow the build-up of exuded compounds rather than to enhance the rate of nutrient uptake. A universal response of root clusters to Pi deprivation is accumulation and exudation of copious amounts of carboxylates (i.e., anions of organic acids) during specific stages in their lifecycle (Figure 5). Citrate plays a central role in nutrient acquisition in virtually all root clusters studied to date. In Lupinus albus, specific transcript expression of ALMT and MATE transporter genes correlated with the exudation of citrate and flavonoids.56 In the rhizosphere, the role of citrate is to mobilize inorganic and organic Pi that is sorbed onto soil particles replacing Pi via ligand exchange, thereby releasing Pi, which thus enters into the soil solution. Both inorganic and organic Pi compounds are solubilized by citrate, the latter then becoming available for hydrolysis by phosphatases.51,56–58 Field and glasshouse studies demonstrate that root-cluster biomass can comprise 5–60% of whole plant mass (Figure 6), indicating a wide variation in carbon costs to the whole plant. In addition, the amount of carbon
exuded by root clusters as carboxylates can be enormous, up to 25% of the total plant dry weight.51 Enhanced synthesis and exudation of citrate by root clusters has been correlated with upregulation of genes encoding enzymes pivotal to enhanced P acquisition and P-use efficiency in many species that form root clusters.56–58 Phosphoenolpyruvate carboxylase (PEPC) is a tightly controlled cytosolic enzyme located at the core of plant carbon metabolism that catalyzes the irreversible 𝛽-carboxylation of the glycolytic intermediate PEP to form the tricarboxylic acid intermediate oxaloacetate and inorganic Pi.58 Post-translational mechanisms that activate or inhibit PEPC activity are crucial to regulate metabolism associated with citrate synthesis. Recent studies indicated that the post-translational activation of PEPC by deubiquitination and phosphorylation, coupled with an approximately 200% increase in the in vivo concentration of PEPC’s allosteric activator glucose-6-phosphate, will facilitate rapid anaplerotic PEP carboxylation in support of substantial organic anion synthesis and exudation by mature proteoid roots of harsh hakea.59 Identification and biochemical and functional characterization of metabolic flexibility in native species adapted to nutrient impoverished habitats have shown novel traits that can, with model plants like Arabidopsis help move toward generating P-use-efficient crops.
CONCLUSIONS Plants require N, P, and Fe for most metabolic and cellular functions. While N is the most abundant gas in the atmosphere, roots acquire it as NO−3 or NH+4 to grow, and these are essential components of N for fertilizers. Whilst our global P reserves are being depleted, Pi levels in many agricultural soils are building up, because 80–90% of Pi applied as fertilizer is precipitated by cations or soil particles, rendering it unavailable for plants that lack-specific adaptations to access sorbed Pi. Fe deficiency may cause leaf yellowing to crops particularly in alkaline soils and may affect plant growth by interfering with Pi nutrition.60 Plants are able to sense N, P, and Fe deficiencies and alter root architecture via LR initiation and growth, a process that occurs at the pericycle or root tip cells and involves auxin signaling. NO−3 transporters are required for NO−3 modulated root development, while transcription factors such as WRKY75 and PYE are important for P and Fe regulated root responses. Although auxin can be considered a general modulator of root morphogenesis under contrasting N and P availabilities, considerable crosstalk occurs with other plant growth regulators
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such as cytokinins,25 gibberellins,61 strigolactones,62 and brassinosteroids.63 Cytokinins are strong antagonists of auxin-mediated cluster-root formation. Addition of cytokinins to nutrient-solution strongly suppresses cluster-root formation in hydroponically grown white lupin.64 In cluster roots of P-deficient white lupin, genes related to proteins involved in the synthesis of cytokinins are up-regulated. Moreover, upregulation of genes involved in sensing cytokinins corresponds with young, juvenile cluster roots that have relatively higher tissue concentration of cytokinins whereas, mature cluster roots have been shown to up regulate expression of genes that produce proteins involved in cytokinin degradation.64 Interestingly, a recent study showed the central role of ethylene during cluster root maturation in white lupin, which was associated with the upregulation of the Fe-deficiency regulated network that mediates ethylene-induced expression of Fe-deficiency responses in Arabidopsis.56,6 Several transcription factors such as PLT, SCR, PHB, PHV, or AUX/IAA with a known role in the control of meristem activity and developmental processes in
Arabidopsis show an increased expression in the tips of the cluster.65 Genes involved in hormonal responses (PIN, LAX, and YUC) and cell-cycle control (CYCA/B and CDK) are also differentially expressed, indicating some commonalities in root developmental programs between different plant species. Desciphering the contribution of phytohormone signaling and their downstream effectors on cellular programs that determine the changes in root architecture to single or combined nutrient deficiencies is an ongoing avenue for research. The study of root clusters in native plants has provided great insight into the degree of root-architecture remodelling that is possible in plants naturally adapted to extremely infertile soils. However, an even deeper understanding (as in Arabidopsis) of nutrient-sensing in root-cluster initiation and development is required, especially if the goal is to apply the knowledge gained for agriculture. Understanding how gene expression, protein activity, and developmental processes change in crops and native plants adapted to soils of low nutrient availability is basic toward understanding plant functioning but also for improving fertilizer-use efficiency and crop yield.
ACKNOWLEDGMENTS We thank Dr. Scott Poethig for the kind invitation to write this review. This work was supported by grants from the Consejo Nacional de Ciencia y Tecnología (CONACYT, México), the Consejo de la Investigación Científica (UMSNH, México, Grant No. CIC 2.26), and the Marcos Moshinsky Foundation. This work was also supported by the DP1092856 from the Australian Research Council (ARC) to MS, an ARC Australian Research Fellow.
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9. Bari R, Datt E, Pant B, Stitt M, Scheible WR. PHO2, microRNA399, and PHR1 define a phosphate-signaling pathway in plants. Plant Physiol 2006, 141:988–999. 10. Pant BD, Buhtz A, Kehr J, Scheible WR. MicroRNA399 is a long-distance signal for the regulation of plant phosphate homeostasis. Plant J 2008, 53:731–738. 11. Kant S, Peng M, Rothstein SJ. Genetic regulation by NLA and microRNA82 for maintaining nitratedependent phosphate homeostasis in Arabidopsis. PLoS Genet 2011, 7:e1002021. 12. Lin WY, Huang TK, Chiou TJ. NITROGEN LIMITATION ADAPTATION, a target of microRNA827, mediates degradation of plasma membrane-localized phosphate transporters to maintain phosphate homeostasis in Arabidopsis. Plant Cell 2013, 25:4061–4074.
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30. Manoli A, Begheldo M, Genre A, Lanfranco L, Trevisan S, Quaggiotti S. NO homeostasis is a key regulator of early nitrate perception and root elongation in maize. J Exp Bot 2014, 65:185–200. 31. Araya T, Miyamoto M, Wibowo J, Suzuki A, Kojima S, Tsuchiya Y, Sawad S, Fukuda H, von Wirén N, Takahashi H. CLE-CLAVATA1 peptide-receptor signaling module regulates the expansion of plant root systems in a nitrogen-dependent manner. Proc Natl Acad Sci USA 2014, 111:2029–2034. 32. Krouk G, Lacombe B, Bielach A, Perrine-Walker F, Malinska K, Mounier E, Hoyerova K, Tillard P, Leon S, Ljung K, et al. Nitrate-regulated auxin transport by NRT1.1 defines a mechanism for nutrient sensing in plants. Dev Cell 2010, 18:927–937. 33. Remans T, Nacry P, Pervent M, Girin T, Tillard P, Lepetit M, Gojon A. A central role for the nitrate transporter NRT2.1 in the integrated morphological and physiological responses of the root system to nitrogen limitation in Arabidopsis. Plant Physiol 2006, 140: 909–921. 34. Léran S, Varala K, Boyer JC, et al. A unified nomenclature of NITRATE TRANSPORTER 1/PEPTIDE TRANSPORTER family members in plants. Trends Plant Sci 2014, 19:5–9. 35. Fang Z, Xia K, Yang Z, et al. Altered expression of the PTR/NRT1 homologue OsPTR9 affects nitrogen
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37. López-Bucio J, Hernández-Abreu E, Sánchez-Calderón L, Nieto-Jacobo MF, Simpson J, Herrera-Estrella L. Phosphate availability alters architecture and causes changes in hormone sensitivity in the Arabidopsis root system. Plant Physiol 2002, 129:244–256. 38. Bonser AM, Lynch J, Snapp S. Effect of phosphorus deficiency on growth angle of basal roots in Phaseolus vulgaris. New Phytol 1996, 132:281–288. 39. Lynch JP. Root phenes for enhanced soil exploration and phosphorus acquisition: Tools for future crops. Plant Physiol 2011, 156:1041–1049. 40. Sánchez-Calderón L, López-Bucio J, Chacón-López A, et al. Phosphate starvation induces a determinate developmental program in the roots of Arabidopsis thaliana. Plant Cell Physiol 2005, 46:174–184. 41. López-Bucio J, Hernández-Abreu E, Sánchez-Calderón L, et al. An auxin transport independent pathway is involved in phosphate stress-induced root architectural alterations in Arabidopsis: identification of BIG as a mediator of auxin in pericycle cell activation. Plant Physiol 2005, 137:681–691. 42. Vashchenko G, MacGillivray RTA. Multi-copper oxidases and human iron metabolism. Nutrients 2013, 5:2289–2313. 43. Ticconi CA, Delatorre CA, Lahner B, Salt DE, Abel S. Arabidopsis pdr2 reveals a phosphate-sensitive checkpoint in root development. Plant J 2004, 37:801–814. 44. Ticconi CA, Lucero RD, Sakhonwasee S, et al. ER-resident proteins PDR2 and LPR1 mediate the developmental response of root meristems to phosphate availability. Proc Natl Acad Sci USA 2009, 106: 14174–14179. 45. Gilbert G, Knight JD, Vance CP, Allan DL. Proteoid root development of phosphorus deficient lupin is mimicked by auxin and phosphonate. Ann Bot 2000, 85: 921–928. 46. Meng ZB, You XD, Suo D, Chen YL, Tang C, Yang JL, Zheng SJ. Root-derived auxin contributes to the phosphorus-deficiency-induced cluster-root formation in white lupin (Lupinus albus). Physiol Plant 2013, 148: 481–489. 47. Dai X, Wang Y, Yang A, Zhang WH. OsMYB2P-1, an R2R3 MYB transcription factor, is involved in the regulation of phosphate-starvation responses and root architecture in rice. Plant Physiol 2012, 159:169–183. 48. Shane MW, Lambers H. Cluster roots: a curiosity in context. Plant Soil 2005, 274:101–125. 49. Vance CP, Uhde-Stone C, Allan DL. Phosphorus acquisition and use: critical adaptations by plants for
51. Shane MW, Cramer MD, Funayama-Noguchi S, Cawthray GR, Millar AH, Day DA, Lambers H. Developmental physiology of cluster-root carboxylate synthesis and exudation in harsh hakea. Expression of phosphoenolpyruvate carboxylase and the alternative oxidase. Plant Physiol 2004, 135:549–560. 52. Delgado M, Zúñiga-Feest A, Alvear M, Borie F. The effect of phosphorus on cluster-root formation and functioning of Embothrium coccineum (R. et J. Forst.). Plant Soil 2013, 373:765–773. 53. Shane MW, Cawthray GR, Cramer MD, Kuo J, Lambers H. Specialized ‘dauciform’ roots of Cyperaceae are structurally distinct, but functionally analogous with ‘cluster’ roots. Plant Cell Environ 2006, 29: 1989–1999. 54. Lambers H, Shane MW, Cramer MD, Pearse SJ, Veneklaas EJ. Root structure and functioning for efficient acquisition of phosphorus: matching morphological and physiological traits. Ann Bot 2006, 98:693–713. 55. Lambers H, Clode P, Hawkins HJ, Laliberté E, Oliveira RS, Reddell P, Shane MW, Stitt M, Weston P. Metabolic adaptations of the non-mycotrophic Proteaceae to soils with low phosphorus availability. In: Plaxton WC, Lambers H, eds. Phosphorus Metabolism in Plants. Annual Plant Reviews. Oxford: Wiley-Blackwell; 2015, 48:289–336. doi:10.1002/9781118958841.ch11. 56. Wang Z, Straub D, Yanga H, Kania A, Shen J, Ludewig U, Neumann G. The regulatory network of cluster-root function and development in phosphate-deficient white lupin (Lupinus albus) identified by transcriptome sequencing. Physiol Plant 2014, 151:323–338. 57. Shane MW, Stigter K, Fedosejevs E, Plaxton WC. Senescence-inducible cell wall and intracellular purple acid phosphatases: implications for phosphorus remobilization in Hakea prostrata (Proteaceae) and Arabidopsis thaliana (Brassicaceae). J Exp Bot. In press. 58. Plaxton WC, Shane MW. The role of post-translational enzyme modifications in the metabolic adaptations of phosphorus-deprived plants. In: Plaxton WC, Lambers H, eds. Phosphorus Metabolism in Plants. Annual Plant Reviews. Oxford: Wiley-Blackwell; 2015, 48:99–124. doi:10.1002/9781118958841.ch4. 59. Shane MW, Fedosejevs ET, Plaxton WC. Reciprocal control of anaplerotic phosphoenolpyruvate carboxylase by in vivo monoubiquitination and phosphorylation in developing proteoid roots of phosphate-deficient harsh hakea. Plant Physiol 2013, 161:1634–1644. 60. Ward JT, Lahner B, Yakubova E, Salt DE, Raghothama KG. The effect of iron on the primary root elongation of Arabidopsis during phosphate deficiency. Plant Physiol 2008, 147:1181–1191.
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Nutritional regulation of root development León Francisco Ruiz Herrera,1 Michael W. Shane2 and José López-Bucio1∗ Mineral nutrients such as nitrogen (N), phosphorus (P), and iron (Fe) are essential for plant growth, development, and reproduction. Adequate provision of nutrients via the root system impacts greatly on shoot biomass and plant productivity and is therefore of crucial importance for agriculture. Nutrients are taken up at the root surface in ionic form, which is mediated by specific transport proteins. Noteworthy, root tips are able to sense the local and internal concentrations of nutrients to adjust growth and developmental processes, and ultimately, to increase or decrease the exploratory capacity of the root system. Recently, important progress has been achieved in identifying the mechanisms of nutrient sensing in wild- and cultivated species, including Arabidopsis, bean, maize, rice, lupin as well as in members of the Proteaceae and Cyperaceae families, which develop highly sophisticated root clusters as adaptations to survive in soils with very low fertility. Major findings include identification of transporter proteins and transcription factors regulating nutrient sensing, miRNAs as mobile signals and peptides as repressors of lateral root development under heterogeneous nutrient supply. Understanding the roles played by N, P, and Fe in gene expression and biochemical characterization of proteins involved in root developmental responses to homogeneous or heterogeneous N and P sources has gained additional interest due to its potential for improving fertilizer acquisition efficiency in crops. © 2015 Wiley Periodicals, Inc. How to cite this article:
WIREs Dev Biol 2015. doi: 10.1002/wdev.183
INTRODUCTION
C
onsiderable information has been gathered in the last decade on plant responses regulated by nutrient signaling, including: (1) elucidation of hormonal pathways with auxin as a central player mediating changes in root architecture by nutrient availability,1–3 (2) identification of transporter proteins for nutrient uptake, and transcription factors as master regulators of root architecture,4–6 (3) the involvement of ubiquitin system components in nutrient signaling,7,8 (4) identification of miRNAs as mobile signals,9–13 and ∗ Correspondence
to: [email protected]
1 Instituto
de Investigaciones Químico-Biológicas, Universidad Michoacana de San Nicolás de Hidalgo, Edificio A-1’, Ciudad Universitaria Morelia, Michoacán, México
2 School
of Plant Biology, Faculty of Science, University of Western Australia, Crawley, Australia Conflict of interest: The authors have declared no conflicts of interest for this article.
(5) characterization of peptides as repressors of lateral root development under heterogeneous nutrient supply.14 Several of these responses are systemically controlled by leaf nutrient status, whereas others are locally regulated by nutrient availability in the root environment.15–17 This focus article highlights signaling role(s) of N, P, and Fe in controlling root development. Nitrate (NO−3 ) and ammonium (NH+4 ) are the major sources of N taken up by roots, and regulate gene expression, energy transfer, protein activation, as well as metabolic and physiological processes.17 P is taken up by roots as fully oxidized soluble phosphate (Pi), which is a component of many key biomolecules such as nucleic acids and phospholipids and thus participates in various enzymatic reactions and metabolic pathways.18 Fe is an essential micronutrient for virtually all organisms because it functions as a component of many enzymes and proteins and its deficiency can
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block the synthesis of chlorophyll precursors, inducing leaf yellowing and reducing photosynthesis.19 As each of these elements is of vital importance for plant growth and development, plant root systems evolved the ability to acclimate to nutrient stress by eliciting a complex array of morphological and biochemical adaptations to optimize N, P, and Fe uptake capacity, in which localized cell division, elongation and differentiation function coordinately to maximize soil exploration and root nutrient foraging.20,21
NITRATE AND AMMONIUM DIFFERENTIALLY MODULATE LATERAL ROOT DEVELOPMENT Arabidopsis and crop seedlings can modify their root architectures to acquire sources of N that are uniformly or unevenly distributed in soil. One important response involves increased growth of lateral roots (LR) within patches enriched in NO−3 or greater LR formation to uniformly supplied NH+4 ions (Figure 1(a)).22,23 The stimulation of LR elongation in NO−3 -rich patches is the result of a signaling effect of the external NO−3 ion itself rather than its downstream metabolites glutamine or glutamate. An important breakthrough in understanding the role of NO−3 in stimulating LR growth was the identification of the ARABIDOPSIS NITRATE-REGULATED1 (ANR1) gene, a member of the plant MADS-box family of transcription factors, which is a vital component in this regulatory signaling pathway (Figure 1(a)).22 Transgenic plants in which ANR1 is repressed show altered suppression of LR growth in NO−3 -rich patches.22 LR primordia initiation is strongly repressed in Arabidopsis by the combination of high external sucrose and low external NO−3 , while sucrose alone appears to act at the level of meristem activation.23 Local supply of NH+4 to Arabidopsis plants increases LR formation. This effect is independent from cumulative uptake of (15)N-labeled NH+4 , suggesting that this process is unlikely due to a nutritional effect but most likely occurs via a sensing event.24 In contrast to wild-type plants, NH+4 -induced LR formation was almost absent in a quadruple AMMONIUM TRANSPORTER mutant qko (amt1;1 amt1;2 amt1;3 amt2;1) and significantly lower in the amt1;3 single mutant.24 To define long-distance signals that affect root growth in response to homogenous or heterogenous N supply, a split-root design has been applied.25,26 On the root experiencing N-limited conditions, both the rate of LR initiation and further LR growth is decreased, while in the N-rich
LRs proliferate (Figure 1(b)). These effects correlated with altered cytokinin biosynthesis and transcriptome reprograming.25 The transcription factor TEOSINTE BRANCHED1/CYCLOIDEA/ PROLIFERATING CELL FACTOR20 (TCP20) was implicated in NO−3 signaling by its ability to bind the promoters in more than 100 NO−3 -regulated genes including NIA1, NRT2.1, and NRT1.1.26 Analysis of insertion mutants of TCP20 showed that they had normal primary and LR growth under uniform supply of NO−3 , but were impaired in preferential LR growth supplied with high NO−3 concentration. Inhibition of LR growth in the high NO−3 was still evident in the mutants even when NH+4 was supplied in the opposite halve, indicating that the TCP20 response was specific to NO−3 . Thus, homogeneous or heterogeneous N sources interfere with LR development apparently acting in different ways.
AUXIN, MICRO RNAs AND PEPTIDE SIGNALING ARE CRITICAL FOR NITROGEN REGULATED ROOT GROWTH The auxin receptor AFB3 and downstream components are essential to regulate LR responses to local supply of NO−3 , likely via miR393 and miR167 action.2,27–29 AFB3 is induced by NO−3 and repressed by miR393 in roots. In contrast, miR393 is not induced in a nitrate reductase (NR)-null mutant of Arabidopsis, suggesting that N metabolites produced by NO−3 reduction and assimilation acts in a negative feedback loop regulating AFB3 levels over time according to N availability. AFB3 acts specifically in the context of the NO−3 response by regulating a connected gene network under control of the NAC4 transcription factor.2 Lateral root responses to N have also been shown to depend on miR167 and its target, the auxin response factor ARF8 mRNA.28 This module acts specifically in pericycle cells in a positive manner to control LR initiation and later in development by decreasing the final length of LRs. Downstream of auxin, nitric oxide (NO) homeostasis appears to be a key regulator of early NO−3 perception and root elongation.30 Accordingly, NO production by NR increased with NO−3 supply in the elongation zone of maize roots and correlated with increased rates of growth. Decreased levels of NO in roots affected their elongation under high NO−3 conditions, whereas treatment of NO−3 -deprived roots with sodium nitroprusside (SNP), a NO-releasing compound, stimulated root growth. An increased formation of LRs in response to homogenous low NO−3 availability depends on
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Control
(a)
NH4+ NRT1.1 NO3–
CIPK23
N
Cytosol T101 miR167
P
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Auxin biosynthesis
Nucleus ANR1
ARF8
TAR2
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(b) CEPR1,2 CK
Pericycle
Nucleus
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TCP20 Xylem
Companion cell CLV1
CEP1 NO3–
N response genes NRT2.1, NRT3.1, NRT1.1 Lateral root development
NO3–
Downstream elements Lateral root development
FIGURE 1 | Regulatory elements involved in LR growth in response to homogeneous or heterogeneous N sources. Homogenous supply of NH+4
triggers lateral root initiation, while NO−3 -rich patches stimulates lateral root elongation. (a) In Arabidopsis, NRT1.1 is a dual-affinity NO−3 transporter involved in both high- and low-affinity NO−3 uptake. The switch between the two affinities is controlled by phosphorylation at the T101, and this phosphorylation is regulated by the calcineurin B-like-interacting protein kinase CIPK23. NRT1.1 promotes LR growth in NO−3 -rich patches likely through the transcription factor ANR1. NO−3 levels affect root development through modulation of auxin synthesis and signaling via TAR2, AFB3 and ARF8. Solid arrows indicate positive effects, blunt solid arrows indicate negative effects, and dashed lines indicate possible interactions. (b) Root foraging for NO−3 involves both local and systemic signaling. Roots respond to low local levels of NO−3 by producing CEP1 and CLE3 peptides, which are repressors of lateral root growth. CEP1 moves upward to the shoot and is received by the LRR-RKs proteins CEPR1 and CEPR2. Long-distance signaling to the root halve grown under high NO−3 involves cytokinin (CK) biosynthesis and requires the transcription factor TCP20, which activates the expression of genes involved in N uptake and is necessary for the induction of development.
the function of the auxin biosynthesis gene TAR2 (tryptophan aminotransferase-related 2).3 TAR2 is expressed in the pericycle of the primary root and is induced under low N conditions. In wild-type
plants, low N stimulated auxin accumulation in LR primordia at early developmental stages. Conversely, auxin accumulation and LR growth were impaired in the tar2 null mutant. Overexpression of TAR2
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leads to increased LR numbers under both high and low N conditions.3 All together, these reports show that AFB3 and downstream signaling proteins may increase auxin accumulation and/or auxin sensitivity in LR primordia depending on NO−3 or NH+4 availability and thus activate root branching. Several homologs of the CLE (CLAVATA3/ ESR-related) gene family are upregulated by N deficiency. CLAVATA3 (CLV3) is known as a peptide that binds to the CLAVATA1 (CLV1) receptor and regulates shoot-cell proliferation and differentiation.31 Interestingly, the signaling pathway composed of N-responsive CLE peptides and the CLAVATA1 (CLV1) leucine-rich repeat receptor-like kinase is expressed in the Arabidopsis root vasculature and pericycle. Overexpression of CLE1, −3, −4, and −7 repressed the development of LR primordia. In contrast, clv1 mutants showed progressive outgrowth of primordia from the parent root under N-deficient conditions.31 These data implicate the N-responsive CLE-CLV1 signaling module as an essential mechanism restrictively controlling root branching in N-deficient environments. N-deprived roots also secrete small peptides belonging to the Arabidopsis C-terminally encoded peptide (CEP) family, which share short, conserved domains near the C terminus, a feature that is common to several modified small peptides from plants.14 CEP1 was used to screen a library of cultured plant cells expressing all known plant leucine-rich repeat receptor kinase (LRR-RK)—encoding genes and identified two homologous LRR-RKs (CEPR1 and CEPR2) that efficiently bind CEP1.14 Exposure of roots to low N increased CEP1 levels in the xylem sap of the shoot, indicating its long-distance transport to the shoot (Figure 1(b)). Arabidopsis plants deficient in this signaling pathway show decreased growth accompanied with N-deficiency symptoms, thus indicating that signaling from the root to the shoot via a peptide—LRR-RK relay helps the plant adapt to fluctuations in local N availability.
ROLE OF NITRATE AND AMMONIUM TRANSPORTERS IN NITROGEN SENSING BY ROOTS The NRT1.1 nitrate transporter plays a key role in NO−3 sensing. NRT1.1 loss of function causes a decreased root proliferation in NO−3 patches, resulting from reduced LR elongation (Figure 1(a)). This response is not due to lower specific NO−3 uptake activity in the nrt1 mutants, but is associated with dramatically decreased ANR1 expression in
LR primordia and in the tips of the LRs.4 Intriguingly, NRT1.1 not only transports NO−3 but also facilitates uptake of auxin, while NO−3 inhibits NRT1.1-dependent auxin uptake.32 These results show that NRT1.1 promotes localized root growth either as a NO−3 sensor, as a facilitator of NO−3 influx into cells responsible of local N perception, or via local cell-specific auxin accumulation. However, mutations of a second NO−3 transporter NRT2.1 have been shown to repress or stimulate LR initiation depending on NO−3 and sucrose supply,23,33 thus implying NO3 transport in the regulation of LR formation via photosynthate supply to roots. Cereals contain a large PTR/NRT1 gene family (PTR, di/tripeptide transporter; NRT1, low-affinity nitrate transporter) for the uptake and translocation of NO−3 , NH+4 , or small peptides. Rice plants have at least 84 PTR/NRT1 genes but, OsPTR9 is one of the few that has been functionally characterized as having a role in root development.34,35 Over-expression of OsPTR9 in transgenic rice plants resulted in increased NH+4 uptake, and LR formation. Conversely, downregulation of OsPTR9 in a T-DNA insertion line (osptr9) and in OsPTR9-RNAi rice plants had the opposite effect. These results indicate that OsPTR9 might influence root surface area for nutrient absorption by affecting LR density.35
PHOSPHORUS DEFICIENCY INHIBITS PRIMARY ROOT GROWTH AND INCREASES ROOT-ABSORPTIVE CAPACITY Plants can modify their root architecture to improve Pi acquisition in P-deficient acid or alkaline soils in which Pi is precipitated out of the soil solution due to combination with cations. Early studies in Arabidopsis and crops have demonstrated that Pi deficiency affects root development by reducing growth of primary roots while increasing the number and length of LRs and root hairs (Figure 2(a)–(d)).36,37 Modifications in root growth angle may also enhance root foraging and exploration in upper soil layers.38,39 This response is associated with litter decomposition, so P accumulates as mostly organic Pi, but with some inorganic Pi as well.39 The first visible event upon low Pi stress in the primary root is a strong reduction of cell elongation, which occurs within hours after plants are transferred from a relatively high to low Pi-containing medium.16 This is followed by a reduction of cell division as revealed by the cell-cycle marker CYCB1;1 in parallel with loss of quiescent center (QC) identity as reported by the QC46 marker. Eventually, differentiation
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FIGURE 2 | Arabidopsis root architectural responses to low Pi stress. Representative root architectures of seedlings grown for 12 days on medium supplied with (a) 1000 μM (high Pi) or (b) 1 μM (low Pi). Note the highly branched root systems in low Pi conditions due to proliferation of LR. Expression of Pi transporter AtPT2 in primary root tips of seedlings grown under high Pi (c) or low Pi (d) supply coincides with high root-hair proliferation capacity. (e) Phenotypes of Arabidopsis WT (Col-0) and low phosphorus insensitive 1 mutant seedlings grown under low P stress. Note the long primary roots developed by the mutants indicative of altered perception of low Pi availability.
events such as root-hair formation increase dramatically further increasing the root absorptive capacity (Figure 2(d)).40 Thus, the primary roots of plants grown under low Pi stress initiate a developmental program that blocks primary root elongation and favors LR formation and root hair proliferation. Genes and proteins that control primary root growth and LR development under low Pi stress have been identified through screening primary root growth in low P medium. Characterization of alleles of BIG, a protein required for normal auxin transport in Arabidopsis showed that this protein is required for pericycle cell activation to form LR in both high (1 mM) and low (1 μM) P conditions, but not for the low P-induced alterations in primary root growth and root hair elongation.41 In contrast, the low Pi-insensitive (LPI) mutants show the typical increase in LR and root hair number under low P conditions but the primary root growth is unaffected (Figure 2(e)), thus implying LPI genes in sensing Pi deficiency (Figure 3).15 Loss-of-function mutations in LOW PHOSPHATE ROOT1 (LPR1) and its close paralog LPR2 strongly reduce the primary root growth inhibition by low Pi stress similarly to the phenotype shown by lpi mutants.16 Interestingly, LPR1 is expressed in the root cap, the possible region sensing Pi deficiency. LPR1 and LPR2 encode multicopper oxidases (MCOs), enzymes that oxidize their substrates with the concomitant reduction of dioxygen to two water molecules.16 Human MCOs possess both
ferroxidase and Fe2+ ferroportin activities and play a role in Fe metabolism.42 The phosphate deficiency response 2 (pdr2) mutant seedlings display hypersensitive responses to Pi deficiency evidenced by conditional short-root phenotype caused by selective inhibition of root cell division followed by meristem death under low Pi conditions.43 Higher density of LRs is observed in pdr2 mutants under low Pi stress, indicating that PDR2 is necessary both for maintenance of root meristem function and pericycle cell activation when Pi availability is limited. PDR2 encodes the single P(5)-type ATPase of Arabidopsis thaliana.44 PDR2 functions in the endoplasmic reticulum (ER) and is required for proper expression of SCARECROW (SCR), a key regulator of root patterning, and for stem-cell maintenance in Pi-deprived roots. Indeed, the MCO encoded by LPR1 is targeted to the ER and interacts with PDR2.44 Because the expression domains of both genes overlap in the stem-cell niche and root meristem, PDR2 and LPR1 may function together in an ER-resident pathway that adjusts root meristem activity to external Pi sensing (Figure 3).
AUXIN MEDIATES ROOT ARCHITECTURE REMODELING BY PHOSPHORUS DEPRIVATION Accumulating evidence suggests that auxins modulate the responses of root architecture to low Pi stress. This was first supported by a study showing that
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FIGURE 3 | Phosphorus starvation signaling pathways regulate primary and LR development. Pi is sensed in root tips by an unknown P starvation sensor, which triggers the signaling pathways that modulate root architecture. Low Pi promotes LR proliferation through increasing auxin transport and BIG function, and auxin signaling that involves TIR1, IAA14, ARF7, and ARF19. SLR1/IAA14 is a member of Aux/IAA family that represses the activity of ARF7 and ARF19 in lateral root initiation. The effects of low Pi stress on cell division and differentiation is regulated through the transcription factor SCR and LPR and PDR proteins. Other transcription factors such as WRKY75 regulate LR proliferation and anthocyanin accumulation.
exogenous application of auxins to P-adequate roots resulted in localized alterations in root architecture in Arabidopsis that mimicked those observed under low P growth conditions.37 Another study showed that LR proliferation in roots of plants under low Pi stress required normal auxin transport mediated by BIG.41 The auxin signalling pathway also contributes to the Pi deficiency-induced cluster-root formation in white lupin (Lupinus albus L).45,46 Compelling evidence indicates that the pattern of LR formation and emergence in response to low Pi stress is mediated by changes in auxin sensitivity in roots.1 This study showed that low Pi signaling altered the expression of auxin-responsive genes and stimulated pericycle cells to divide. Modulation of auxin sensitivity by Pi was found to depend on the auxin receptor TRANSPORT INHIBITOR RESPONSE 1 (TIR1) and the transcription factor AUXIN RESPONSE FACTOR 19 (ARF19) (Figure 3). Low Pi stress increases the expression of TIR1 and causes AUXIN/INDOLE-3-ACETIC ACID (AUX/IAA) auxin response repressors to be degraded.1 It was proposed that auxin sensitivity is enhanced in Pi-deprived plants by an increased expression of TIR1, which accelerates the degradation of AUX/IAA proteins, thereby unshackling ARF transcription factors that activate/repress genes involved in LR initiation and growth.
TRANSCRIPTION FACTORS MEDIATE ROOT ARCHITECTURAL RESPONSES TO PHOSPHORUS AND IRON AVAILABILITY Several transcription factors act downstream or in concert with auxin signaling to mediate root responses to low Pi stress. RNAi knockdown studies showed that a reduction in WRKY75 mRNA levels caused early anthocyanin accumulation and reduced Pi uptake during Pi starvation.5 Silencing of WRKY75 also caused an increase in LR and root-hair growth that was independent of Pi status, suggesting WRKY75 is a modulator of root development. Microarray data from Pi-deficient rice identified an R2R3 MYB transcription factor, OsMYB2P-1.47 Primary roots of OsMYB2P-1-overexpressing plants were shorter than those in wild-type plants under Pi-sufficient conditions, while primary roots and adventitious roots of OsMYB2P-1-overexpressing plants were longer than those of wild-type plants under low Pi conditions. In contrast to the accumulated information about the regulatory networks controlled by N or P availability that shape root form and function, less is known about root responses to Fe. Noteworthy, plants exposed to Fe deprivation display increased primary root waving and LR initiation, with the
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FIGURE 4 | Iron signaling regulates root architecture. Fe is taken up into the symplast by IRT1 transporters at the root surface. Reduction of Fe by FRO2, and FRO5 reductases, contribute to increased Fe acquisition. Fe can diffuse through the symplastic space to the pericycle and then to the vasculature. BTS and PYE, a putative E3 ligase and a bHLH transcription factor, respectively, are specifically induced in root pericycle cells under Fe deficiency. PYE and other bHLHs transcription factors likely regulate the genes responsive to activate pericycle cells for LR initiation under Fe deprivation. Fe translocation inside the plant is associated with suitable chelating molecules such as organic acids and loading from vasculature to mesophyll tissue occurs via YSL transporters.
pericycle acting as a regulatory center for sensing and responding to local Fe availability (Figure 4).6 Whole root time–course microarray analyses showed that numerous transcription factors are temporally coexpressed under Fe deficiency. A screen of mutants of these transcription factors looking for alterations in metabolic and developmental responses to Fe deficiency identified a bHLH transcription factor named POPEYE (PYE), and a second gene tightly coregulated with PYE called BRUTUS (BTS).6 PYE appears to play an important role in root responses to Fe-deficient conditions, as pye1 mutant seedlings display decreased tolerance to Fe deprivation in primary root growth and are more sensitive to develop leaf chlorosis.
ROLE OF NUTRIENTS IN ROOT CLUSTER FORMATION: SIMILARITY IN FUNCTION, DIVERSITY IN STRUCTURE While basic knowledge about the pathways involved in root development by nutrient availability has been obtained from Arabidopsis, root clusters are particularly remarkable for their diversity of morphological structure and roles in plant adaptation to soils with very low nutrient availability (Figure 5). These highly
specialised ephemeral root structures evolved to perform critical functions in nutrient acquisition. They occur on the root systems of a range of perennial dicotyledonous (e.g., ‘proteoid roots’ in Proteaceae; Figure 5(a) and (d) and monocotyledonous (‘dauciform roots’ in Cyperaceae; Figures 5(c) and 6) native plants, but are also formed on several agriculturally significant crops (e.g., ‘cluster roots’ in Fabaceae; Figure 5(b)).48 Proteoid root clusters are characterised by incredibly dense clusters of numerous short, determinate rootlets, covered by root hairs at maturity. Superficially, proteoid roots exhibit a huge range of morphologies but most common are ‘tree-like’ compound clusters (e.g., Banksia species; Figure 5(a)) or ‘bottle-brush-like’ simple proteoid-roots (e.g., Lupinus albus, Figure 5(b)). Dauciform root clusters also exhibit a vast array of structural diversity, but are characterized by short, carrot shaped LRs with remarkably dense and long root hairs (Figure 5(c)). The formation of root clusters is most highly represented amongst nonmycorrhizal plants that are particularly well-adapted to soils with low Pi availability.49,50 Detailed developmental studies have shown that root clusters can be physiologically active for as little as 10 days or up to 4 weeks.50–52,48 For example, after initiation, dauciform roots in Schoenus unispiculatus reach maturity within ∼ 7 days
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FIGURE 5 | Root-cluster morphologies in Proteaceae, Cyperaceae, and Fabaceae. All species are well adapted to soils of extremely low P
concentrations. Plants were grown in nutrient solutions containing ≤1 μM phosphate. (a) Proteoid roots (compound type) of Banksia prionotes (Proteaceae). (b) Cluster roots (simple type) of Lupinus albus L. (Fabaceae). (c) Dauciform roots of Schoenus unispiculatus (Cyperaceae). (d) Proteoid roots of Hakea prostrata (simple type, Proteaceae). (e) Stages of dauciform root development in S. unispiculatus. Dauciform roots were initially white, but often turned bright yellow as they matured. Remarkably large numbers of long root hairs with a similar final length were fully developed after ca. 7–9 days. The axes of very young dauciform roots (1–3 days old) illustrate the ‘carrot shape’ of the dauciform root axes, and a slender stalk connected the carrot-shaped axis to the main root. After 10–12 days, the dauciform roots senesced. (f) Stages of proteoid-root development in H . prostrata from rootlet initiation to senescence (∼20 days old, far right). Scale bars in mm (a) 10; (b) 4; (c) 3; (d) 11; (e) 5; and (f) 9.
(Figure 5(e)), whereas proteoid-roots of Hakea prostrata (Proteaceae) require approximately 14 days (Figure 5(f)). The ephemeral nature of root clusters necessitates their continual replacement by sequential formation along axes of growing LRs (Figures 5(d) and 6), or as a singular structure upon short LRs as illustrated in the time–course of dauciform root development in an Australian sedge (Figure 5(e)).53
Phosphorus has been repeatedly shown to be a key nutrient influencing initiation (low Pi supply) or suppression (higher Pi supply) of root clusters (Figure 6), but their formation is also influenced by N and Fe.54 Split-root or foliar feeding experiments suggest that the extent of root-cluster formation can be determined by internal (leaf) Pi status or Pi availability in the external root environment in a species-specific manner. With little exception, the root architecture
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FIGURE 6 | Influence of phosphate supply on dauciform-root formation in the south-western Australian native Cyperaceae Carex fascicularis. The formation of highly specialised dauciform roots increases dramatically on plants as the Pi level is decreased in nutrient solution (from left to right). This particular species of Carex develops relatively small dauciform roots in series (arrows) along the axes of growing LR (cf . Figure 5(c) and (e)). Scale bar; 13 mm.
of field-grown root cluster-forming species is almost always patterned toward greater abundance of root clusters, and hence, root-foraging in upper soil horizons where levels of nutrients are often enriched.55 Formation of dense clusters of rootlets and/or root hairs in a small soil volume markedly increases root surface area for nutrient foraging;20,48 however, those ‘zones’ from which rootlets or root hairs acquire Pi and micronutrients greatly overlap. This morphology is likely to have been customized during evolution to allow the build-up of exuded compounds rather than to enhance the rate of nutrient uptake. A universal response of root clusters to Pi deprivation is accumulation and exudation of copious amounts of carboxylates (i.e., anions of organic acids) during specific stages in their lifecycle (Figure 5). Citrate plays a central role in nutrient acquisition in virtually all root clusters studied to date. In Lupinus albus, specific transcript expression of ALMT and MATE transporter genes correlated with the exudation of citrate and flavonoids.56 In the rhizosphere, the role of citrate is to mobilize inorganic and organic Pi that is sorbed onto soil particles replacing Pi via ligand exchange, thereby releasing Pi, which thus enters into the soil solution. Both inorganic and organic Pi compounds are solubilized by citrate, the latter then becoming available for hydrolysis by phosphatases.51,56–58 Field and glasshouse studies demonstrate that root-cluster biomass can comprise 5–60% of whole plant mass (Figure 6), indicating a wide variation in carbon costs to the whole plant. In addition, the amount of carbon
exuded by root clusters as carboxylates can be enormous, up to 25% of the total plant dry weight.51 Enhanced synthesis and exudation of citrate by root clusters has been correlated with upregulation of genes encoding enzymes pivotal to enhanced P acquisition and P-use efficiency in many species that form root clusters.56–58 Phosphoenolpyruvate carboxylase (PEPC) is a tightly controlled cytosolic enzyme located at the core of plant carbon metabolism that catalyzes the irreversible 𝛽-carboxylation of the glycolytic intermediate PEP to form the tricarboxylic acid intermediate oxaloacetate and inorganic Pi.58 Post-translational mechanisms that activate or inhibit PEPC activity are crucial to regulate metabolism associated with citrate synthesis. Recent studies indicated that the post-translational activation of PEPC by deubiquitination and phosphorylation, coupled with an approximately 200% increase in the in vivo concentration of PEPC’s allosteric activator glucose-6-phosphate, will facilitate rapid anaplerotic PEP carboxylation in support of substantial organic anion synthesis and exudation by mature proteoid roots of harsh hakea.59 Identification and biochemical and functional characterization of metabolic flexibility in native species adapted to nutrient impoverished habitats have shown novel traits that can, with model plants like Arabidopsis help move toward generating P-use-efficient crops.
CONCLUSIONS Plants require N, P, and Fe for most metabolic and cellular functions. While N is the most abundant gas in the atmosphere, roots acquire it as NO−3 or NH+4 to grow, and these are essential components of N for fertilizers. Whilst our global P reserves are being depleted, Pi levels in many agricultural soils are building up, because 80–90% of Pi applied as fertilizer is precipitated by cations or soil particles, rendering it unavailable for plants that lack-specific adaptations to access sorbed Pi. Fe deficiency may cause leaf yellowing to crops particularly in alkaline soils and may affect plant growth by interfering with Pi nutrition.60 Plants are able to sense N, P, and Fe deficiencies and alter root architecture via LR initiation and growth, a process that occurs at the pericycle or root tip cells and involves auxin signaling. NO−3 transporters are required for NO−3 modulated root development, while transcription factors such as WRKY75 and PYE are important for P and Fe regulated root responses. Although auxin can be considered a general modulator of root morphogenesis under contrasting N and P availabilities, considerable crosstalk occurs with other plant growth regulators
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such as cytokinins,25 gibberellins,61 strigolactones,62 and brassinosteroids.63 Cytokinins are strong antagonists of auxin-mediated cluster-root formation. Addition of cytokinins to nutrient-solution strongly suppresses cluster-root formation in hydroponically grown white lupin.64 In cluster roots of P-deficient white lupin, genes related to proteins involved in the synthesis of cytokinins are up-regulated. Moreover, upregulation of genes involved in sensing cytokinins corresponds with young, juvenile cluster roots that have relatively higher tissue concentration of cytokinins whereas, mature cluster roots have been shown to up regulate expression of genes that produce proteins involved in cytokinin degradation.64 Interestingly, a recent study showed the central role of ethylene during cluster root maturation in white lupin, which was associated with the upregulation of the Fe-deficiency regulated network that mediates ethylene-induced expression of Fe-deficiency responses in Arabidopsis.56,6 Several transcription factors such as PLT, SCR, PHB, PHV, or AUX/IAA with a known role in the control of meristem activity and developmental processes in
Arabidopsis show an increased expression in the tips of the cluster.65 Genes involved in hormonal responses (PIN, LAX, and YUC) and cell-cycle control (CYCA/B and CDK) are also differentially expressed, indicating some commonalities in root developmental programs between different plant species. Desciphering the contribution of phytohormone signaling and their downstream effectors on cellular programs that determine the changes in root architecture to single or combined nutrient deficiencies is an ongoing avenue for research. The study of root clusters in native plants has provided great insight into the degree of root-architecture remodelling that is possible in plants naturally adapted to extremely infertile soils. However, an even deeper understanding (as in Arabidopsis) of nutrient-sensing in root-cluster initiation and development is required, especially if the goal is to apply the knowledge gained for agriculture. Understanding how gene expression, protein activity, and developmental processes change in crops and native plants adapted to soils of low nutrient availability is basic toward understanding plant functioning but also for improving fertilizer-use efficiency and crop yield.
ACKNOWLEDGMENTS We thank Dr. Scott Poethig for the kind invitation to write this review. This work was supported by grants from the Consejo Nacional de Ciencia y Tecnología (CONACYT, México), the Consejo de la Investigación Científica (UMSNH, México, Grant No. CIC 2.26), and the Marcos Moshinsky Foundation. This work was also supported by the DP1092856 from the Australian Research Council (ARC) to MS, an ARC Australian Research Fellow.
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