IDPs (intrinsically disordered proteins) are highly abundant in eukaryotic proteomes and important for cellular functions, especially in cell signalling and transcriptional regulation. An IDR (intrinsically disordered region) within an IDP often undergoes disorder-to-order transitions upon binding to various partners, allowing an IDP to recognize and bind different partners at various binding interfaces. Plant-specific GRAS proteins play critical and diverse roles in plant development and signalling, and act as integrators of signals from multiple plant growth regulatory and environmental inputs. Possessing an intrinsically disordered N-terminal domain, the GRAS proteins constitute the first functionally required unfoldome from the plant kingdom. Furthermore, the N-terminal domains of GRAS proteins contain MoRFs (molecular recognition features), short interaction-prone segments that are located within IDRs and are able to recognize their interacting partners by undergoing disorder-to-order transitions upon binding to these specific partners. These MoRFs represent potential protein–protein binding sites and may be acting as molecular bait in recognition events during plant development. Intrinsic disorder provides GRAS proteins with a degree of binding plasticity that may be linked to their functional versatility. As an overview of structure–function relationships for GRAS proteins, the present review covers the main biological functions of the GRAS family, the IDRs within these proteins and their implications for understanding mode-of-action.

INTRODUCTION

The GRAS proteins are a plant-specific protein family named after the first three members: GAI [GA (GIBBERELLIC ACID)-INSENSITIVE], RGA (REPRESSOR OF GAI) and SCR (SCARECROW). They play essential roles in transcriptional regulation and various signal transduction pathways controlling plant development. On the basis of the protein family database (Pfam) [1], the GRAS family of genes (PF03514) has been found in 294 embryophyta species and is represented by 1035 sequences. Several independent phylogenetic analyses of GRAS proteins have revealed similarly structured family trees with the following ten subfamilies that have been named either after one of their members or after a common motif: AtLAS (Arabidopsis LATERAL SUPPRESSOR), AtSCL (Arabidopsis SCR-like) 4/7, HAM (HAIRY MERISTEM), AtSCR (Arabidopsis SCR), DLT (DWARF AND LOW TILLERING), AtSCL3, DELLA, AtPAT1 [Arabidopsis pat (phytochrome A signal transduction) 1-1], AtSHR [Arabidopsis SHR (SHORTROOT)] and LISCL (Lilium longiflorum SCR-like) [26]. Some GRAS proteins are not yet assigned to a subfamily and may represent further distinct subfamilies with their functions remaining unknown (Figure 1A). Most GRAS family members possess a variable N-terminal domain (N-domain) and a widely and highly conserved C-terminal domain (GRAS domain), whereas a small number of GRAS proteins have double GRAS domains or a single GRAS domain followed by another functional domain in the C-terminus. When comparing the sequence identities among and between GRAS subfamilies (Table 1), GRAS proteins can show as little as 10% fundamental sequence identity, which is largely due to the conserved GRAS domains present in the whole GRAS family (see below). Higher sequence identities (up to 98%) can be shown within some subfamilies, suggesting that these proteins share a similar function and/or a common mode-of-action.

GRAS protein family and domain structure

Figure 1
GRAS protein family and domain structure

(A) The ten current plant GRAS subfamilies with nine unclassified rice GRAS proteins and the human HsSRC (Homo sapiens SRC) protein as an outgroup. (B) Domain structure, showing the variable N-terminal domain and C-terminal domain with widely conserved motifs. With the exception of the LRI, LRII, NLS and SH2 motifs, the rest of motifs are named after the most prominent amino acids of each motif. (C) The 14 conserved subfamily-restricted motifs (I–XIV) in the N-domains of GRAS subfamilies. As examples, motifs VIII–X from the DELLA subfamily and motifs XIII and XIV from the LISCL subfamily are shown. The conserved hydrophobic and aromatic residue repeats in each motif are indicated by the black filled circles. The fragments predicted as α-helical MoRFs are underlined. With kind permission from Springer Science+Business Media: Plant Molecular Biology, A functionally required unfoldome from the plant kingdom: intrinsically disordered N-terminal domains of GRAS proteins are involved in molecular recognition during plant development, volume 77, 2011, pages 205–223, X. Sun, B. Xue, W. T. Jones, E. Rikkerink, A. K. Dunker and V. N. Uversky.

Figure 1
GRAS protein family and domain structure

(A) The ten current plant GRAS subfamilies with nine unclassified rice GRAS proteins and the human HsSRC (Homo sapiens SRC) protein as an outgroup. (B) Domain structure, showing the variable N-terminal domain and C-terminal domain with widely conserved motifs. With the exception of the LRI, LRII, NLS and SH2 motifs, the rest of motifs are named after the most prominent amino acids of each motif. (C) The 14 conserved subfamily-restricted motifs (I–XIV) in the N-domains of GRAS subfamilies. As examples, motifs VIII–X from the DELLA subfamily and motifs XIII and XIV from the LISCL subfamily are shown. The conserved hydrophobic and aromatic residue repeats in each motif are indicated by the black filled circles. The fragments predicted as α-helical MoRFs are underlined. With kind permission from Springer Science+Business Media: Plant Molecular Biology, A functionally required unfoldome from the plant kingdom: intrinsically disordered N-terminal domains of GRAS proteins are involved in molecular recognition during plant development, volume 77, 2011, pages 205–223, X. Sun, B. Xue, W. T. Jones, E. Rikkerink, A. K. Dunker and V. N. Uversky.

Table 1
Sequence identity ranges among and between GRAS subfamily proteins

Modified from [2].

Subfamily AtLAS AtSCL4/7 HAM AtSCR DLT AtSCL3 DELLA AtPAT1 AtSHR LISCL 
AtLAS 37–63%          
AtSCL4/7 23–34% 47–70%         
HAM 11–31% 16–27% 14–86%        
AtSCR 21–33% 20–31% 13–31% 31–98%       
DLT 19–31% 19–22% 13–23% 21–29% 38%      
AtSCL3 16–31% 17–31% 9–25% 17–35% 17–29% 24–93%     
DELLA 21–33% 21–28% 13–25% 24–32% 23–28% 16–35% 54–94%    
AtPAT1 16–31% 19–33% 12–26% 17–34% 14–29% 15–29% 17–35% 23–74%   
AtSHR 16–30% 16–24% 9–23% 16–25% 13–23% 12–26% 15–25% 15–27% 21–71%  
LISCL 16–28% 16–25% 8–25% 16–29% 13–20% 11–28% 16–27% 16–34% 14–25% 29–91% 
Subfamily AtLAS AtSCL4/7 HAM AtSCR DLT AtSCL3 DELLA AtPAT1 AtSHR LISCL 
AtLAS 37–63%          
AtSCL4/7 23–34% 47–70%         
HAM 11–31% 16–27% 14–86%        
AtSCR 21–33% 20–31% 13–31% 31–98%       
DLT 19–31% 19–22% 13–23% 21–29% 38%      
AtSCL3 16–31% 17–31% 9–25% 17–35% 17–29% 24–93%     
DELLA 21–33% 21–28% 13–25% 24–32% 23–28% 16–35% 54–94%    
AtPAT1 16–31% 19–33% 12–26% 17–34% 14–29% 15–29% 17–35% 23–74%   
AtSHR 16–30% 16–24% 9–23% 16–25% 13–23% 12–26% 15–25% 15–27% 21–71%  
LISCL 16–28% 16–25% 8–25% 16–29% 13–20% 11–28% 16–27% 16–34% 14–25% 29–91% 

The function and characterization of at least one member of each of the ten GRAS subfamilies has been analysed, using either loss- or gain-of-function phenotype mutants and biochemical assays, such as yeast two-hybrid methods [4,7,8]. This information was hampered by a lack of structural information about GRAS proteins until recently. Murase et al. [9] reported the first crystal structure of an N-terminal fragment of the DELLA protein AtGAI (Arabidopsis GAI). Analysis of sets of proteins that contain IDRs (intrinsically disordered regions) (the so-called unfoldome) and the role of these unfolded regions has become a key new area of focus. Sun et al. [10] investigated the intrinsically disordered features of the N-domains of the DELLA subfamily and their binding-induced folding properties [10]. The disorder analysis was further extended to the whole GRAS family to reveal that the intrinsically disordered N-domains of GRAS proteins constitute a plant-specific unfoldome and may act as molecular bait by initiating the key molecular recognition events in plant development encoded by these proteins [2]. In the animal kingdom, numerous IDPs (intrinsically disordered proteins) have been systematically studied and shown to play essential roles in cellular functions, especially signalling and transcriptional regulation [1113]. In contrast, there have been fewer reports of IDPs from the plant kingdom. As an overview of structure–function relationships for GRAS proteins, the present review covers various biological functions of the GRAS family, with emphasis on the potential role played by intrinsic disorder to understand the mode-of-action of GRAS proteins.

DOMAIN STRUCTURES OF GRAS FAMILY PROTEINS

The sequences widely and highly conserved in the GRAS domain can be subdivided into five distinct motifs with their name derived from the most prominent amino acids: LRI (leucine-rich region I), VHIID, LRII (leucine-rich region II), PFYRE and SAW (Figure 1B). Flanked by the two leucine-rich regions, the VHIID motif is present in all GRAS family members identified with occasional interchanges of the valine, leucine and isoleucine residues. Proline, asparagine, histidine, aspartic acid, glutamine and leucine residues are also highly conserved within the larger region of the VHIID motif. The two leucine-rich regions LRI and LRII are characterized by leucine repeats, but, in most cases, not regular heptad repeats as in leucine zippers. The LRI motif is conserved in all GRAS proteins with putative NLSs (nuclear localization signals) found in the latter part of the motif, conforming to the consensus for either bipartite NLSs (as confirmed for the DELLA subfamily) or the non-typical SV40 (simian virus 40)-like NLS [14], as reported for some other GRAS proteins. The LRII motif also contains an LXXLL (Leu-Xaa-Xaa-Leu-Leu; Xaa is for any amino acid) pattern in the latter part of the motif that is conserved in half of the GRAS proteins. A similar LXXLL pattern has been demonstrated to mediate the binding of steroid receptor co-activator complexes to cognate nuclear receptors in mammals [15]. The LRI-VHIID-LRII pattern has been experimentally confirmed to be involved in binding between GRAS proteins and their nucleic acid or protein partners [1621].

The PFYRE motif contains three distinct parts: proline (P), aromatic phenylalanine and tyrosine (FY), and arginine and glutamic acid (RE) residues. There is also an aspartic acid/glutamic acid residue existing in the FY part which is conserved in nearly all GRAS proteins. Although the PFYRE motif is not as strictly conserved at the sequence level as the VHIID motif, the sequences of the PFYRE motif are largely co-linear and this motif still shows a high degree of similarity in sequence among all members of the GRAS family. The SAW motif contains three sequential units: WX7G (X7 is for any seven amino acids), L-W and SAW. The three tryptophan residues are conserved in nearly all of the GRAS family, with the SAW unit located close to the C-terminus of GRAS proteins. Although the roles of the PFYRE and SAW motifs are currently unknown, the absolute conservation of the residues in these motifs may indicate that these motifs are required either for the function or for the structural integrity of GRAS proteins. In addition, the GRAS domains also contain a consensus sequence similar to the SH2 (Src homology 2) domain of STAT (signal transducer and activator of transcription) proteins. In animals, this domain mediates the effect of extracellular ligands on gene transcription through selective binding of the SH2 domain to a phosphorylated tyrosine residue [22]. Present in most members of the GRAS family, the SH2-like motif overlaps with the latter part of the PFYRE motif. Both the order and identity of conserved residues in this plant SH2-like domain are the same as those in the animal SH2 domain of STATs [23].

Consistent with higher sequence identities within each subfamily than between subfamilies, some conserved subfamily-restricted motifs have also been found within nine of the ten subfamilies (Figure 1C). In contrast with the widely conserved motifs in the GRAS domain discussed above, these subfamily-restricted motifs occur in the intrinsically disordered N-domains of GRAS proteins and they are believed to play important roles in molecular recognition events involving GRAS proteins [2]. All of these subfamily-restricted motifs share a common pattern consisting of either repeated hydrophobic or aromatic residues that form the framework for the conserved motif. For example, the N-domains of the DELLA subfamily contain three subfamily-restricted motifs with repeated hydrophobic or aromatic residues (VIII, IX and X in Figure 1C, corresponding to the DELLA, VHYNP and LR/KXI motifs). It has been experimentally confirmed that most of the repeated hydrophobic or aromatic residues in motifs VIII and IX directly interact with the GA-bound receptor GID1 (GA-INSENSITIVE DWARF 1), and thereby play a critical role in perceiving GA signals that control plant development [9,10]. It has been proposed that the repeated hydrophobic or aromatic residues in the other subfamily-restricted motifs (I–VII and XI–XIV) play a similar role in binding to the interacting partners of GRAS proteins [2].

On the other hand, short leucine-rich segments often act as NESs (nuclear export signals), also known as the leucine-rich NES. Containing approximately 10–20 amino acids, the NES motif is ambiguous in terms of its length, consensus sequence and conformation when bound to its receptor [2426]. Normally, the NES motifs comprise three or four regularly spaced leucine residues, but these leucine residues can be substituted by other hydrophobic residues such as isoleucine, valine, phenyalanine or methionine. The exportin-1 or CRM1 (chromosomal region maintenance 1) protein was identified as the cellular receptor for the NES to mediate the export of proteins containing leucine-rich NESs from the nucleus. As shown in the crystal structures of CRM1–SNUPN (Snurportin-1) complexes, the leucine-rich NES of the SNUPN, consisting of its first 16 residues and protruding away from the rest of SNUPN molecule, binds to a hydrophobic groove of CRM1 [26]. The NES motif of SNUPN forms a short three-turn amphiphatic helix followed by an extended coil in which the hydrophobic side chains are aligned to interact with the hydrophobic groove of CRM1, whereas polar residue side chains are exposed on the surface of the complex [26]. Apart from the helical structure, the NES motifs can also have non-helical or extended conformations [25,26]. The ambiguity of the leucine-rich NES together with the prevalence of the leucine-rich patterns in the proteome make it challenging to predict true NESs and proteins containing the NES motif. However, it is critical that a nuclear-targeting signal be accessible within the transported protein for efficient interactions with its NES receptor, meaning that a structurally disordered region harbouring the NES would be favourable [26]. A new predictor has recently been developed, by using IDR prediction and other derived features, along with more direct primary sequence features, to predict both the correct position of the NES within a specific protein and all potential NES-containing proteins [27]. Simultaneous existence of the NLS and NES within the same protein renders it capable of constant nucleocytoplasmic shuttling. Such signal-directed nuclear import and export are widely used by cells to modulate signal transduction pathways, and both the NLS and NES are required for some transcription factors to possess optimal transcriptional activation [28]. Moreover, the basic import/export machinery is highly conserved between animals, plants and yeast [24]. Although most GRAS proteins are localized in the nucleus, some of them, e.g. AtSCL13 from the AtPAT1 subfamily, AtSCL14 from the LISCL subfamily, and AtSHR from the AtSHR subfamily, have been found in both the nucleus and cytoplasm. The partial cytosolic localization of AtSCL14 is most probably due to exportin-dependent nuclear export [20]. Both nuclear and cytoplasmic localization are required for cell-to-cell transport of AtSHR, and such capacity for intercellular movement of AtSHR may be conserved among other GRAS family proteins [29]. Given that NLS motifs, the widely conserved LRI and LRII regions and the subfamily-restricted hydrophobic or aromatic residue-rich motifs in the N-domains exist in most GRAS proteins, there is potential for the existence of NES motifs and thus nucleocytoplasmic shuttling to occur with other GRAS proteins. Nevertheless, systematic experimental investigations are needed in this area.

BIOLOGICAL FUNCTIONS OF GRAS SUBFAMILIES

Below we focus on the biological functions of the eight subfamilies with most direct relevance to later discussions. For the sake of completeness, a brief summary of the main biological functions of all ten GRAS subfamilies are listed in Table 2.

Table 2
Main biological functions of the ten GRAS protein subfamilies

The proteins listed are some of the experimentally characterized GRAS proteins along with some representative references. AtRGA, Arabidopsis REPRESSOR OF GAI; AtRGL1, Arabidopsis RGA-like 1; MtNSP1, M. truncatula nodulation signalling pathway 1; OsCIGR, O. sativa CIGR; PeSCL7, Populus euphratica SCARECROW-LIKE 7; SLN1, barley SLENDER 1; ZmD8, Z. mays DWARF.

Subfamily Proteins Functions Reference(s) 
DELLA AtGAI, AtRGA, AtRGL1, AtRGL2, AtRGL3, SLR1, StRGA, Rht1, ZmD8, SLN1 Repressors of GA-responsive genes, transcriptional co-activators of PIFs, control fruit patterning, modulate JA signalling, integrators of regulatory and environmental signals. [9,10,16,17,3033,35,36,6771
AtSCR AtSCR, OsSCR1, ZmSCR Root radial patterning and root growth, QC identity, asymmetric cell division. [18,29,38,40,41
AtSHR AtSHR, OsSHR1, OsSHR2, MtNSP1 Root radial patterning and root growth, cell division and endodermis specification, transcription factor for nodule development. [18,19,29,37,39,45
AtSCL3 AtSCL3 Positive regulator of the GA response pathway, integrator of GA/DELLA signalling and the SCR/SHR pathway in root cell elongation. [42,43
LISCL LISCL, AtSCL14, NtGRAS1, CsSCL1, PrSCL1 Transcriptional regulator, transcriptional regulation or activation associated with the plant stress responses, adventitious root formation in response to auxin. [6,20,46,48
AtSCL4/7 PeSCL7 Transcriptional regulator in response to environmental stresses such as salt, osmotic shock and drought. [86
AtPAT1 AtPAT1, AtSCL13, OsCIGR1, OsCIGR2 PhyA-specific signalling, positive regulator of phyB-dependent red light signalling, hypocotyl elongation, transcriptional regulators in the early stages of plant defence signalling. [5053
DLT Rice semi-dwarf mutant with low-tillering (DLT or Os29) BR (brassinosteroid) signalling, negatively regulated by either exogenous or endogenous BRs; modulating BR responses and participating in the control of rice tillering. DLT and OsBZR1 (Oryza sativa brassinazole-resistant 1) regulate each other for the fine-tuning of BR responses. [87,88
AtLAS AtLAS, LeLs, OsMOC1 Axillary shoot formation, initiation of axillary meristems, control of tillering. [5456
HAM HAM, BnSCL1, MtNSP2 Shoot meristem maintenance, transcriptional activator in response to auxin, transcriptional co-activator in nodulation signalling. [19,21,44,57
Subfamily Proteins Functions Reference(s) 
DELLA AtGAI, AtRGA, AtRGL1, AtRGL2, AtRGL3, SLR1, StRGA, Rht1, ZmD8, SLN1 Repressors of GA-responsive genes, transcriptional co-activators of PIFs, control fruit patterning, modulate JA signalling, integrators of regulatory and environmental signals. [9,10,16,17,3033,35,36,6771
AtSCR AtSCR, OsSCR1, ZmSCR Root radial patterning and root growth, QC identity, asymmetric cell division. [18,29,38,40,41
AtSHR AtSHR, OsSHR1, OsSHR2, MtNSP1 Root radial patterning and root growth, cell division and endodermis specification, transcription factor for nodule development. [18,19,29,37,39,45
AtSCL3 AtSCL3 Positive regulator of the GA response pathway, integrator of GA/DELLA signalling and the SCR/SHR pathway in root cell elongation. [42,43
LISCL LISCL, AtSCL14, NtGRAS1, CsSCL1, PrSCL1 Transcriptional regulator, transcriptional regulation or activation associated with the plant stress responses, adventitious root formation in response to auxin. [6,20,46,48
AtSCL4/7 PeSCL7 Transcriptional regulator in response to environmental stresses such as salt, osmotic shock and drought. [86
AtPAT1 AtPAT1, AtSCL13, OsCIGR1, OsCIGR2 PhyA-specific signalling, positive regulator of phyB-dependent red light signalling, hypocotyl elongation, transcriptional regulators in the early stages of plant defence signalling. [5053
DLT Rice semi-dwarf mutant with low-tillering (DLT or Os29) BR (brassinosteroid) signalling, negatively regulated by either exogenous or endogenous BRs; modulating BR responses and participating in the control of rice tillering. DLT and OsBZR1 (Oryza sativa brassinazole-resistant 1) regulate each other for the fine-tuning of BR responses. [87,88
AtLAS AtLAS, LeLs, OsMOC1 Axillary shoot formation, initiation of axillary meristems, control of tillering. [5456
HAM HAM, BnSCL1, MtNSP2 Shoot meristem maintenance, transcriptional activator in response to auxin, transcriptional co-activator in nodulation signalling. [19,21,44,57

The DELLA subfamily: GA, light and JA (jasmonate) signalling and transcriptional co-activation in control of plant growth

DELLA proteins, named after the conserved DELLA motif in their N-domains, are one of the most extensively studied GRAS subfamilies. DELLAs function as repressors of GA-responsive plant growth and are key regulatory targets in the GA signalling pathway. Gain-of-function mutants of the DELLA genes show dwarfism and GA-insensitivity, whereas loss-of-function mutations cause constitutive GA-response phenotypes [30,31]. The GA signal modulates plant growth by causing degradation or deactivation of DELLAs, thereby de-repressing GA-regulated genes. The GA signal is firstly perceived by the GA receptor GID1 in rice, and its three homologous genes in Arabidopsis thaliana (AtGID1a, AtGID1b and AtGID1c). GA-bound GID1 then facilitates the formation of a ternary complex of GA, GID1 and the N-domain of DELLAs through the conserved DELLA and VHYNP motifs [9]. This ternary complex allows recruitment of an F-box protein, referred to as GID2 (GA-INSENSITIVE DWARF 2) in rice and SLY1 (SLEEPY1) in A. thaliana, by binding to the VHIID and LRII motifs of the GRAS domain of DELLAs [32]. This GA-dependent interaction confers specificity on the SCFGID2/SLY1(Skp1/cullin/F-box) E3 ubiquitin ligase complex toward DELLAs by promoting addition of a polyubiquitin chain, and targets DELLAs for their subsequent degradation by the 26S proteasome [7]. The disappearance of DELLAs stimulates GA-responsive processes, such as seed germination, stem and root elongation, and transition to flowering.

DELLAs have also been revealed to act as transcriptional co-activators in interacting with PIFs (PHYTOCHROME-INTERACTING FACTORs), members of group VII of the bHLH (basic helix-loop-helix) transcription factors [16,33]. In the absence of GA, DELLAs interact with the light-responsive PIF3 and PIF4 using its LRI region, preventing PIF3 and PIF4 from binding to their target promoters and leading them to be an inactive complex. The GA-induced degradation of DELLAs release PIF3 and PIF4, allowing them to regulate gene expression and promoting plant growth in response to light signalling. This DELLA–bHLH regulatory module may also be implicated in fruit development in that DELLAs bind another Arabidopsis bHLH factor ALC (ALCATRAZ) that specifies tissues required for fruit dehiscence [34], preventing activation of its targets. Thus the GA-induced degradation of DELLAs may release ALC to modulate the expression of its target genes and direct the differentiation of the SL (separation layer) cells [35]. On the other hand, DELLAs have been shown to prevent the JAZ1 (JA ZIM-domain 1) protein (a key repressor of MYC2 in JA signalling) from binding to MYC2 (a key transcriptional activator of JA responses) through a direct DELLAs–JAZ1 interaction, thereby facilitating the release of MYC2 for activating the JA response. The N-domain and the LRI motif are responsible for the direct interaction between DELLAs and JAZ1 [17]. In addition, DELLAs can serve as integrators that mediate the cross-talk of various regulatory inputs, such as abiotic stresses, auxin and ethylene signals [36], although less is known about the mechanisms of this integration.

The AtSCR and AtSHR subfamilies: root radial patterning and root growth

The proteins SCR and SHR belong to different subfamilies, but together they play an important role in the control of radial patterning for both the root and shoot in Arabidopsis [8,37]. The plant root shows a radial organization in which epidermis and ground tissue layers (cortex and endodermis) surround a central stele consisting of the pericycle and vasculature. At the root tip, stem cells are maintained by a group of slowly dividing cells termed the QC (quiescent centre). AtSCR is required for the asymmetric cell division of the initial cell that is responsible for the generation of ground tissue and is expressed in the QC [38]. Expressed within the central stele, AtSHR has been shown to move outwards into adjacent ground tissue where it enters the nucleus and is necessary for QC and endodermis specification [39].

SCR transcription is limited to the ground tissue and is positively regulated by SHR and a positive-feedback loop. After SHR moves into the root endodermal cell layer, SCR blocks SHR movement by sequestering it into the nucleus through direct interaction and forming an SCR–SHR complex. Sequestered in the nucleus, the complex promotes the production of more SCR to ensure that there is sufficient SCR to trap SHR in the endodermal cell layer [18]. The homologous genes in maize [ZmSCR (Zea mays SCR)] and rice [OsSCR (Oryza sativa SCR)] were shown to have comparable expression patterns with AtSCR in roots [40,41]. This suggests that the mechanism may be evolutionarily conserved, providing an explanation as to why nearly all plants possess only a single layer of endodermis. Direct interactions between SHR and SCR control some aspects of SHR function, such as asymmetric cell division, QC specification and stem cell maintenance, whereas SHR may form complexes with other proteins to fulfil other aspects of SHR function, such as endodermis specification [18]. On the other hand, OsSCR can be expressed in other processes involving asymmetric cell divisions, such as stomatal development and ligule formation. OsSHR1 (O. sativa SHR1) expression is less restricted than in Arabidopsis, suggesting a similar, but not identical, function in monocots [41].

SHR directly interacts with SCR through the LRI-VHIID-LRII pattern in the GRAS domain [18]. Further studies of interactions between SHR and SCR revealed that SHR movement is required for normal root development, and the VHIID and PFYRE motifs in the GRAS domain of SHR are essential for this movement [29]. Similarly, the GRAS domain of SCR alone has the capability to move, although it is not as efficient as the movement of SHR. The N-domain of SHR functions to stabilize its GRAS domain and enhance the movement of SHR, whereas the N-domain of SCR is likely to play a role in inhibiting movement of the intact SCR protein [29]. Both inter- and intra-molecular protein interactions are important events for SHR and SCR in root development.

The AtSCL3 subfamily: integration of GA signalling with the SCR/SHR pathway during root development

AtSCL3 was found to express predominantly in the root endodermis with a pattern similar to that of SCR [8] and its promoter was directly bound by the SCR–SHR complex [18], implying that AtSCL3 may also play a role in endodermal specification. Recently, two independent studies revealed that AtSCL3 is nuclear-localized in root cells, AtSCL3 functions as a positive regulator of GA signalling, and AtSCL3 and DELLAs antagonize each other in modulating both downstream GA responses by direct protein–protein interaction and GA homoeostasis by feedback-regulating upstream GA biosynthetic genes [42,43]. The direct protein interaction between AtSCL3 and DELLAs has been experimentally confirmed using yeast two-hybrid and in vitro pull-down assays [42], whereas the precise locations of the interacting motifs are still to be identified. It has also been demonstrated that AtSCL3 controls the co-ordination of root cell elongation in the elongation-differentiation zone by attenuating DELLAs. Furthermore, AtSCL3 modulates the timing and extent of the formative division for ground tissue maturation in the meristem zone by both maintaining functional GA signalling through interaction with DELLAs and employing the SHR–SCR module [43]. Thus AtSCL3, acting downstream of both GA/DELLA and SHR/SCR pathways and serving as an integrator, forms a regulating network together with DELLAs and SHR–SCR to mediate GA-promoted cell elongation in the root endodermis.

NSP (NODULATION SIGNALLING PATHWAY) 1 and NSP2: GRAS protein–DNA complex for nodulation signalling in legumes

Legume plants develop a symbiotic relationship with rhizobia that reside in nodules developed on the roots or stems of legumes after plant host perception of the Nod factor, a signalling molecule released by rhizobia. Two GRAS proteins, NSP1 and NSP2, have been discovered to be essential for nodule development and function in legumes [44,45]. As for SHR and SCR, NSP1 and NSP2 belong to two different subfamilies (AtSHR and HAM respectively with 20.1% identity). These two proteins perform different but complementary roles in Nod-factor signalling and nodule morphogenesis. A previous study of the model legume Medicago truncatula has revealed that NSP1 and NSP2 form homo- and hetero-polymers, and that NSP1 is a DNA-binding transcription factor that attaches directly to the promoters of Nod factor-inducible early nodulin genes (ENOD11) [19]. The binding of NSP1 to the promoter of ENOD11 is enhanced by Nod factor application and requires the presence of NSP2; therefore it is the complex of NSP1–NSP2 that is directly associated with the promoters of Nod factor-inducible genes.

As well as the interactions between NSP1 and DNA, protein–protein interactions play an essential role in nodulation signalling. The LRI and LRII regions of NSP1 are required for DNA binding, whereas the LRI region of NSP2 is responsible for its association with NSP1. Specific removal or a site-directed mutation in the LRI region of NSP2 results in its loss of, or reduced interaction with, NSP1 and interferes with its ability to function in nodulation signalling [19].

The LISCL subfamily: transcriptional regulation in response to auxin and stress-induced signals

The LISCL subfamily consists of two clades, a rice-specific subgroup and a second subgroup that contains proteins from Arabidopsis and other plants. The LISCL subfamily appears to function in transcriptional regulation. The LISCL protein itself, a representative of the subfamily, was the first reported GRAS protein directly involved in regulation of expression of a specific gene. Expressed in the premeiotic phase within the anthers of L. longiflorum, LISCL plays a role in the transcriptional activation of a meiosis-associated promoter during meiosis and it requires a specific factor(s) for transcriptional activation in PMCs (pollen mother cells) [46]. There are two conserved subfamily-restricted motifs in the N-domains of the LISCL subfamily, motifs XIII and XIV (Figure 1C), that have many acidic amino acids flanking hydrophobic or aromatic residue repeats [2]. Motif XIII and the following neutral region are responsible for the strong transcriptional activation function of LISCL [46], consistent with the finding that hydrophobic residues interspersed between acidic residues are associated with transcriptional activation [47]. Other members of the LISCL subfamily play similar transcriptional activation or co-activation roles, but in response to different signal pathways. Predominantly expressed in roots, PrSCL1 (Pinus radiata SCL1) and CsSCL1 (Castanea sativa SCL1) are induced in response to exogenous auxin in rooting-competent cuttings and play a role at the very early stages of adventitious root formation [6]. These two GRAS proteins from distantly related forest species share similarity in both sequences and auxin-induced expression in roots, suggesting a conserved function for these proteins in auxin signalling associated with adventitious root formation in both species. A nuclear-localized tobacco GRAS protein [NtGRAS1 (Nicotiana tabacum GRAS1)] is strongly up-regulated by a wide array of both abiotic and biotic stress treatments (antimycin A, H2O2, salicylic acid and L-cysteine) and it may play a role in the transcriptional regulation of genes associated with the plant stress response [48]. NtGRAS1 expression is strongly induced by a number of stress treatments in both tobacco suspension cells and tobacco leaf tissue. Another member of the LISCL subfamily involved in the response to xenobiotic stress, AtSCL14, is recruited to target promoters by forming a complex with class II TGA transcription factors, a conserved plant subfamily of basic leucine zipper transcription factors binding to the TGACG motif in certain promoters [49]. This recruitment is essential for the transcriptional activation of the genes involved in detoxifying harmful chemicals. AtSCL14 serves as a transcriptional co-activator of TGA factors [20].

The mechanisms involved in the transcriptional regulation and interactions among the LISCL subfamily protein, interacting partners and promoters remain unclear. However, all members of the LISCL subfamily share both widely conserved GRAS domains (motifs LRI, VHIID, LRII, PFYRE and SAW) and two conserved subfamily-restricted acidic motifs (XIII and XIV in Figure 1C) in their N-domains. It was proposed that a similar function of transcriptional co-activation through these subfamily-restricted acidic motifs in the N-domains may be conserved for the members of the LISCL subfamily [2].

The AtPAT1 subfamily: phytochrome light signal transduction and plant defence

AtPAT1 is a positive regulator involved in the phyA (phytochrome A)-specific signalling pathway and acts at an early stage of the phyA signalling cascade [50]. As it lacks a putative NLS common in other GRAS proteins, AtPAT1 is located only in the cytoplasm, suggesting that important early signalling events involving phyA occur in the cytoplasm. Analysis with a truncated AtPAT1 containing the first 341 amino acids showed that the whole C-terminal domain of AtPAT1 is required for function. In contrast, AtSCL13, another member of the AtPAT1 subfamily, serves as a positive regulator of continuous red light signals downstream of phyB (phytochrome B) [51]. Predominantly expressed in the active elongation zones of hypocotyls, AtSCL13 plays its major role in hypocotyl elongation during de-etiolation. Genetic evidence suggests that AtSCL13 is also needed to modulate phyA signal transduction in a phyB-independent way. Localized in both the cytoplasm and nucleus, AtSCL13 could play a biological role in both of these cellular compartments. Although AtPAT1 and AtSCL13 are both involved in phytochrome signalling, each is specific for a different light condition and regulates a different subset of responses.

Two CIGR (chitin-inducible cibberellin-responsive) rice GRAS proteins from the AtPAT1 subfamily (CIGR1 and CIGR2) have been shown to be rapidly induced in suspension-cultured rice cells upon N-acetyl-chito-oligosaccharide elicitor perception and in co-cultivation with the rice blast fungus [52]. They may play key roles as transcriptional regulators in the early stages of defence signalling following fungal perception and/or pathogenesis. It was further revealed that CIGR1 and CIGR2 are also responsive to exogenous bioactive GA in a dose-dependent manner in suspension-cultured rice cells, showing multiple signalling mechanisms involving CIGR1 and CIGR2 activation [53]. CIGR1 and CIGR2 also appear to be positively regulated by the rice DELLA protein SLR1 in both suspension-cultured cells and in planta, suggesting that SLR1 may act as a transcriptional activator of CIGR1 and CIGR2 before its degradation in response to GA. Unlike AtPAT1, CIGR1 and CIGR2 are nuclear-localized, which is consistent with their functions as transcriptional regulators rather than signal transmitters.

The AtLAS subfamily: axillary meristem development

The AtLAS subfamily is associated with developmental processes of the axillary meristem. The AtLAS protein itself is required for axillary shoot formation during vegetative development, but not for the initiation of primary axillary meristems during the reproductive phase of development. In the latter period, either AtLAS is not needed or it is replaced by other proteins, perhaps a different GRAS family member [54]. LeLs (Lycopersicon esculentum lateral suppressor), a member of the AtLAS subfamily from tomato, is responsible for the initiation of axillary meristems leading to lateral shoot formation during vegetative growth [55], indicating that AtLAS and LeLs are functionally relevant and that the mechanism controlling side-shoot development is conserved between tomato and Arabidopsis. OsMOC1 (O. sativa monoculm 1), a homologue of AtLAS in rice, also has a critical role in the initiation of axillary meristem, and formation and outgrowth of tiller buds [56]. Overexpression of OsMOC1 leads to an increased number of tillers, as well as a reduction in height, indicating that OsMOC1 has distinct functions, such as promotion of tiller bud outgrowth, and a negative effect on plant height. Differences between these AtLAS family proteins may reflect fundamental differences between monocotyledonous tillering and dicotyledonous branching.

Members of the AtLAS subfamily contain all of the widely conserved motifs of the GRAS domain. One common feature of the AtLAS subfamily is that they have a very short N-terminal domain characterized by the presence of homopolymeric stretches of serine and threonine residues that are also common in other GRAS proteins. However, there are no conserved subfamily-restricted motifs currently identified in their N-domains.

The HAM subfamily: shoot meristem maintenance

Plant shoot development depends on the perpetuation of a SAM (shoot apical meristem) at the very summit of the growth axis. Petunia HAM, a GRAS protein from the HAM subfamily, functions in shoot meristem maintenance and mediates a signal from lateral organ primordia and stem provasculature which is essential and specific for maintaining the SAM [57]. HAM is likely to signal cell-fate decisions in the shoot apex, promoting the undifferentiated state as a distinct cellular identity, possibly in a similar way to AtSHR which controls cell fate by intercellular movement during root development [39]. Both HAM and AtSHR act non-cell-autonomously, functioning as relays between the differentiating cells and the non-committed cells. Maintaining the undifferentiating cells in the SAM is achieved, at least in part, by the expression of HAM in the surrounding cells, whereas AtSCR expression in the QC is needed to maintain the undifferentiated state of the initial cells [4,58].

Alternatively, BnSCL1 (Brassica napus SCR-like), a member of the HAM subfamily from B. napus, plays a role in transcriptional activation. Expressed predominantly in roots, BnSCL1 gene expression is regulated by auxin, suggesting that the role of BnSCL1 in root development may be associated with auxin [21]. BnSCL1 has been shown to act as a transcriptional activator or co-activator, and both the N- and C-domain of BnSCL1 are required for the transactivation. BnSCL1 regulates the plant auxin response through interaction with AtHDA19, an Arabidopsis HDAC (histone deacetylase). HDACs are generally found in large complexes constituting transcriptional repressors and co-repressors or transcriptional activators [59]. Under elevated auxin concentrations in cells, BnSCL1 may interact with AtHDA19 using its VHIID motif in the GRAS domain to relieve repression exerted by HDA19 on auxin-responsive genes [21]. As mentioned above, NSP2, another member of the HAM subfamily, has also been shown to interact with NSP1 as a transcriptional co-activator in nodulation signal transduction.

INTRINSICALLY DISORDERED N-DOMAINS CONTRIBUTE TO FUNCTIONAL POLYMORPHISM

The discovery and characterization of functional IDPs is one of the fastest growing areas of protein science in the past decade [13]. IDPs or proteins containing IDRs lack secondary and/or tertiary structures in solution, yet possess crucial biological functions under physiological conditions [11]. Bioinformatics studies have predicted that approximately 25–30% of eukaryotic proteins are IDPs, and more than 50% of eukaryotic proteins have IDRs [60]; furthermore, more than 70% of signalling proteins have IDRs [61], and 82–94% of transcription factors from three transcription factor datasets possess extended disordered regions [62]. In the animal kingdom, numerous IDPs have been characterized by experimental and bioinformatics analyses to be over-represented in the cell nucleus, and overwhelmingly prevalent in cell signalling and transcriptional regulation.

An IDR enables a protein to fold differently in order to recognize and bind different partners at various binding interfaces – binding promiscuity that makes IDPs or IDRs central in signalling and functional regulation of the cell [63]. Studies have revealed that intrinsic disorder is a common feature of hub proteins from four eukaryotic interactomes, and disordered domains confer such hub proteins with the crucial ability to interact with multiple structurally diverse partners in interaction networks [6466]. Such binding promiscuity is true for GRAS proteins as they not only regulate various processes of plant development, but also act as integrators of signals from multiple plant growth regulatory and environmental inputs. The N-domains of the DELLA subfamily were characterized using both theoretical and experimental approaches to be IDPs, whereas the C-terminal GRAS domains of DELLA proteins possess a basically folded structure [10,67]. Furthermore, disorder studies of the whole GRAS family support the hypothesis that that the N-domains of all GRAS proteins are intrinsically disordered, forming the first functionally required unfoldome identified in the plant kingdom [2].

A generalized model for the domain structure of GRAS proteins is shown in Figure 2. The model proposes that GRAS proteins have a variable N-terminal region consisting of IDRs and a structurally folded C-terminal region consisting of a series of widely and highly conserved motifs. We suggest [2,10] that the IDRs in GRAS proteins act as a readily accessible platform to display their binding promiscuity and qualify the GRAS family as hub proteins. The functional polymorphism of GRAS proteins is reflected by the fact that: (i) the GRAS family has evolved into different subfamilies (with ten currently identified) that possess various primary functions targeting specific developmental and growth processes; (ii) different members within the same subfamily, e.g. the LISCL subfamily, play similar roles of transcriptional regulation, but in response to distinct signal pathways; (iii) the GRAS proteins within the same subfamily, e.g. DELLA proteins, can cross-talk with many different signal pathways in plant development by interacting with key proteins in these pathways (e.g. DELLA–GID1, DELLA–GID2/SLY1, DELLA–PIFs, DELLA–ALC, DELLA–JAZ1 and DELLA–AtSCL3); and (iv) in addition to their primary functions, GRAS proteins can act as pairs of interacting partners from different subfamilies (AtSHR–AtSCR, NSP1–NSP2, SLR1–CIGR1/CIGR2, AtSCL3–DELLA and AtSCL3–SCR/SHR) to modulate specific signalling and developmental processes. It has been proposed that the widely and highly conserved GRAS domains throughout the entire GRAS family could be commonly involved in transcriptional regulation. In contrast, an assortment of intrinsically disordered N-domains renders GRAS proteins capable of being involved in diverse aspects of plant development and lends a degree of specificity to the transcriptional regulation imposed by the GRAS domains [2]. In other words, nuclear-localized GRAS proteins are functionally required to accommodate different partners, whereas the intrinsically disordered nature of their N-domains enables GRAS proteins to act as hub proteins and allows them to perform their key role in integrating multiple developmental and environmental signals.

Model of functional polymorphism of GRAS proteins defined by their IDR

Figure 2
Model of functional polymorphism of GRAS proteins defined by their IDR

The wiggly random coils indicate the intrinsically disordered nature of the N-domains of each subfamily. The grey core part indicates the widely and highly conserved GRAS domains found throughout the entire GRAS family and involved in transcriptional regulation. The beads on the random coils indicate the various MoRFs (potential binding sites) located in the disordered N-domains that act as molecular baits by which each GRAS protein recognizes and interacts with its specific partner in specific signalling pathways.

Figure 2
Model of functional polymorphism of GRAS proteins defined by their IDR

The wiggly random coils indicate the intrinsically disordered nature of the N-domains of each subfamily. The grey core part indicates the widely and highly conserved GRAS domains found throughout the entire GRAS family and involved in transcriptional regulation. The beads on the random coils indicate the various MoRFs (potential binding sites) located in the disordered N-domains that act as molecular baits by which each GRAS protein recognizes and interacts with its specific partner in specific signalling pathways.

INTRINSIC DISORDER CONTRIBUTES TO PHOSPHORYLATION/DEPHOSPHORYLATION OF GRAS PROTEINS

Phosphorylation and dephosphorylation play important roles in protein–protein interactions. Phosphorylated sites in proteins can either be directly recognized by interacting partners or introduce allosteric changes that trigger a series of downstream effects. Phosphorylation and dephosphorylation have been observed as essential factors in the interactions of some GRAS proteins. The phosphorylation and dephosphorylation states of DELLA proteins were correlated with their stability, plant growth repressive activity and their GA-induced degradation [6870]. GA signal activates both dephosphorylation of serine/threonine residues and phosphorylation of tyrosine residues prior to triggering the GA-induced DELLA protein degradation [71]. Acting upstream of NSP1 and NSP2, the nuclear-localized CCaMK (calcium- and calmodulin-dependent protein kinase) may modify either NSP1 or NSP2, or both through phosphorylation to facilitate their interaction or association with DNA, given that CCaMK is required for nodulation signalling [72], and that NSP1 and NSP2 are also required for CCaMK-induced gene expression [73]. Reversible phosphorylation is required for the plant-stress-induced response of NtGRAS1 [48]. Rice CIGR1 and CIGR2, induced by the GA signal, depend on both phosphorylation and dephosphorylation events [53]. Phosphorylation can be a major mechanism of intracellular signal transduction in response to both abiotic and biotic stress in plants [74].

Protein phosphorylation occurs with much higher frequency in disordered regions than in ordered regions, indicating a strong preference for phosphorylated sites in the IDRs. This may be because the open structure of IDRs reduces steric hindrance to access by kinases and phosphatases [75]. In a model of dynamic disordered protein complexes, phosphorylation has been shown to be efficiently utilized as a means to fine-tune the electrostatic interactions of disordered protein regions for signal transduction [76]. Most GRAS proteins have homopolymeric stretches of serine, threonine and sometimes tyrosine residues in their intrinsically disordered N-domains, implying that phosphorylation and dephosphorylation would be a high-probability event for the N-domains of GRAS proteins. This is consistent with the phosphorylation site predictions of GRAS proteins by combining position-specific amino acid frequencies and disorder information [2]. Recently, a rice kinase (EL1) has been identified that directly phosphorylates the rice DELLA protein SLR1 [77]. Given the complexity that reversible phosphorylation may be involved in the regulatory mechanisms of GRAS proteins [6871], the studies of phosphorylation and dephosphorylation of GRAS proteins still have a long way to go. Nevertheless, disorder-assisted prediction of phosphorylation sites would provide a new approach to look into the phosphorylated sites of GRAS proteins and their mediating effects on interactions and recognitions with partners in signalling cascades.

INTRINSICALLY DISORDERED N-DOMAINS CONTRIBUTE TO MOLECULAR RECOGNITION

One of the most important functions of IDPs or IDRs is molecular recognition in regulatory and cell-signalling processes via protein–protein interactions [11,66]. In addition to promoting binding promiscuity by interacting with numerous partners, IDRs provide other important functional advantages over structurally ordered proteins for molecular recognition in signalling: IDRs can bind their partners with high specificity and low affinity. To permit specific recognition, IDRs usually undergo binding-induced folding and display binding plasticity by accommodating diverse binding sites of different partners during protein interactions. As a result of the disorder-to-order transition, IDRs adopt a preferred ordered conformation upon binding to their biological partners [63,78,79]. Such binding-induced folding is further characterized by the presence of MoRFs (molecular recognition features), short rigid segments that are located within extended disordered regions and are able to recognize their interacting partners and thus play a key role in initiating disorder-to-order transitions [80]. Existing examples of the preferred conformations upon binding to partners include α-helix (α-MoRFs), β-strand (β-MoRFs) or irregular (ι-MoRFs) structures [81,82]. A large decrease in conformational entropy due to folding of disordered regions in the disorder-to-order transition is able to uncouple specificity from binding strength [83,84]. The resultant high-specificity/low-affinity combination associated with the regulatory interaction between an IDP and its partner is therefore both highly specific and easily dispersed. These properties are essential in signalling networks as activating and terminating a signal are equally important for signalling cascades [85].

Most GRAS proteins utilize their widely conserved LRI-VHIID-LRII pattern in the C-terminal GRAS domains for their interactions with interacting partners, which have been experimentally confirmed for the following pairs of interacting proteins using either a single motif or the entire LRI-VHIID-LRII pattern: DELLA–PIFs, DELLA–JAZ1, DELLA–GID2, AtSHR–AtSCR, NSP1–NSP2 and BnSCL1–AtHDA19. Although not yet confirmed, it is tempting to speculate that a similar mechanism may be employed for the interacting pairs in which DELLAs, AtSCR, NSP2 and BnSCL1 act as transcriptional co-activators by either blocking or enhancing the transcriptional activity of their partners. The highly conserved leucine-rich regions have been revealed to be involved in various types of transcriptional regulations [15]. Therefore it is likely that the widely and highly conserved LRI-VHIID-LRII pattern or the entire GRAS domain is a key component in allowing GRAS proteins to be involved in the transcriptional regulatory machinery. It is even possible that these transcriptional regulation functions could be evolutionarily conserved for the whole GRAS family.

In contrast, fewer interactions utilizing the less widely conserved (subfamily-restricted) motifs in the N-domains of GRAS proteins have been reported, probably due to the dynamic and flexible nature of the interactions occurring between intrinsically disordered N-domains and their interacting partners for molecular recognition or signal transmission. Nevertheless, many MoRFs have been predicted by bioinformatics analyses to overlap with these subfamily-restricted motifs in the N-domains of GRAS proteins [2], representing additional potential protein–protein binding sites for molecular recognition during plant development. The intrinsically disordered N-domains bear one or more MoRFs (shown in Figure 2 as beads on the random coils) as molecular baits to hook one or more interacting partners. Multiple MoRFs within the same protein may provide some GRAS proteins with their functional versatility and/or ability to integrate multiple regulatory and environmental signals.

Such predicted binding sites have been experimentally confirmed in the case of the DELLA subfamily. As shown in Figure 3, the N-domains of the DELLA subfamily were both theoretically and experimentally demonstrated to be IDPs. Both the DELLA and LK/RXI motifs were predicted as α-helix-forming MoRFs, whereas the VHYNP motif was excluded from the prediction of α-helix-forming MoRFs because this motif adopted an irregular loop conformation (ι-MoRFs) rather than an α-helix conformation after binding to the GID1 receptor [10]. Acting as MoRFs, the conserved DELLA and VHYNP motifs in the N-domains of DELLAs undergo disorder-to-order transitions upon binding to the GA receptor GID1, seeding the DELLA–GID1 interactions. The third conserved LK/RXI motif in the N-domains of DELLAs, predicted as an α-helix-forming MoRF, was totally disordered and projected into the solvent region in the crystal structure of the ternary complex (GA/GID1/short AtGAI fragment), indicating that it is not involved in the interactions with the GID1 receptor. This implies that the LK/RXI motif is a potential binding site of DELLAs for an unknown component in signalling pathways. The LK/RXI motif sequence blast search of the PDB database mostly resulted in α-helical structures (Figure 3). We proposed that pull-down experiments utilizing the LK/RXI motif may help to find the potential new interacting partner of DELLAs [10].

Disorder predictions and the protein interactions between the DELLA and GA GID1 receptor

Figure 3
Disorder predictions and the protein interactions between the DELLA and GA GID1 receptor

Scores of intrinsic disorder were predicted using PONDR® VL-XT for two members of the DELLA subfamily, AtRGL2 (orange line) and AtGAI (black line, dotted lines indicate shifting to align the AtGAI sequence with AtRGL2). The residues with scores >0.5 (threshold) are disordered and those <0.5 are ordered. The thick black horizontal line represents the location of the N-domains of DELLA proteins. The N-domains are predicted as disordered, whereas the C-domains are basically ordered. The X-ray crystal structure of GA3-bound receptor AtGID1a from A. thaliana complexed with a fragment of the N-domain of AtGAI (residues 11–113, encompassing the DELLA, VHYNP motifs and most of the LK/RXI motif; PDB code 2ZSH) is displayed on the top to indicate its interactions with the DELLA and VHYNP motifs. The short red and cyan bars indicate the locations of the α-helix-forming MoRFs predicted for the DELLA and LK/RXI motif respectively. The short magenta bar indicates the location of ι-MoRFs (irregular loop) for the VHYNP motif. All of the MoRFs (binding sites) are at or near the downward spikes in the plot of disorder scores, short rigid segments in the extended disorder region. The LK/RXI motif of AtGAI is totally disordered and not defined in the crystal structure, supporting that this motif does not interact with AtGID1a, but its α-helix-forming MoRF is supported by its sequence blast search of the PDB database. A representative of the α-helices searched is shown in cyan, and the unknown partner interacting with DELLA proteins through the LK/RXI motif is shown as a grey oval shape.

Figure 3
Disorder predictions and the protein interactions between the DELLA and GA GID1 receptor

Scores of intrinsic disorder were predicted using PONDR® VL-XT for two members of the DELLA subfamily, AtRGL2 (orange line) and AtGAI (black line, dotted lines indicate shifting to align the AtGAI sequence with AtRGL2). The residues with scores >0.5 (threshold) are disordered and those <0.5 are ordered. The thick black horizontal line represents the location of the N-domains of DELLA proteins. The N-domains are predicted as disordered, whereas the C-domains are basically ordered. The X-ray crystal structure of GA3-bound receptor AtGID1a from A. thaliana complexed with a fragment of the N-domain of AtGAI (residues 11–113, encompassing the DELLA, VHYNP motifs and most of the LK/RXI motif; PDB code 2ZSH) is displayed on the top to indicate its interactions with the DELLA and VHYNP motifs. The short red and cyan bars indicate the locations of the α-helix-forming MoRFs predicted for the DELLA and LK/RXI motif respectively. The short magenta bar indicates the location of ι-MoRFs (irregular loop) for the VHYNP motif. All of the MoRFs (binding sites) are at or near the downward spikes in the plot of disorder scores, short rigid segments in the extended disorder region. The LK/RXI motif of AtGAI is totally disordered and not defined in the crystal structure, supporting that this motif does not interact with AtGID1a, but its α-helix-forming MoRF is supported by its sequence blast search of the PDB database. A representative of the α-helices searched is shown in cyan, and the unknown partner interacting with DELLA proteins through the LK/RXI motif is shown as a grey oval shape.

On the basis of the above example of the DELLA subfamily, the intrinsically disordered N-domains of the GRAS family are most likely to be involved in various molecular recognitions, since all of the predicted MoRFs of GRAS proteins fall exclusively in the N-domains. Structural information from intrinsic disorder-based MoRFs provides a conceptual framework that can guide future experiments to understand the mechanism of signalling and regulation for the entire GRAS family [2].

CONCLUSIONS AND PERSPECTIVES

Plant-specific GRAS proteins play critical and diverse roles in plant growth and development, and often act as an integrator of signals from multiple plant growth regulatory and environmental inputs. IDPs are highly abundant in eukaryotic proteomes and are important for a wide range of biological processes, especially in cell signalling and transcriptional regulation. As the first identified unfoldome from the plant kingdom, GRAS proteins may well utilize their intrinsically disordered N-terminal domains in molecular recognition during plant development and signalling, whereas their structurally folded C-terminal domains are involved in transcriptional regulation. The intrinsic disorder of GRAS proteins renders them capable of functional polymorphism. Having been confirmed in the case of the DELLA subfamily, all MoRFs of GRAS proteins appear to be concentrated into the disordered N-terminal domains and thus may undergo disorder-to-order transitions upon binding to their various interacting partners.

We postulate that functional analysis of the subfamily-restricted motifs of repeated hydrophobic or aromatic residues identified in the N-domains will give clues to possible binding sites of those unexamined GRAS proteins in the interactions with their partners. For example, motif III in BnSCL1 (HAM subfamily) could be important in initiating protein–protein interactions with other members of the large HDAC transcriptional repressor complex because both the N- and C-domain of BnSCL1 are required for transactivation [21]; motif X (DELLA subfamily) is likely to be a potential binding site of DELLAs for components in cross-talk pathways other than GA signalling [10]; and motifs XIII and XIV in AtSCL14 (LISCL subfamily) may be important for interacting with co-activating TGA transcription factors since motif XIII in LISCL has been shown to be associated with a strong transcriptional activation function [46].

In addition to the LRI and LRII regions in the GRAS domain, the repeated hydrophobic or aromatic residues, forming the framework for the subfamily-restricted motifs in the N-domains, also provide extra sites to be tested for potential NES activity. For example, motif XII in the AtSHR (AtSHR subfamily) may be important in potentiating AtSHR movement by stabilizing its GRAS domain and providing nuclear export activity. In contrast, the N-domain of AtSCR (AtSCR subfamily) plays a role in inhibiting movement of the intact AtSCR protein [29]. It is possible that the NES motif in AtSCR could be masked by motifs IV or V of AtSCR, and that this action may be enhanced by the flexibility resulting from the intrinsic disorder of the N-domain. As these subfamily-restricted motifs often coincide with the MoRFs of GRAS proteins, they allow the development of a conceptual framework for the role of specific motifs within the N-domains and provide guidance for rational experimental approaches, such as targeting sequences for site-directed mutation analysis in functional studies or pull-down experiments, as well as for finding new interacting proteins. These approaches should help to elucidate the roles of these motifs in the modulatory functions of GRAS proteins.

Abbreviations

     
  • ALC

    ALCATRAZ

  •  
  • AtLAS

    Arabidopsis LATERAL SUPPRESSOR

  •  
  • AtSCL

    Arabidopsis SCARECROW-like

  •  
  • AtSCR

    Arabidopsis SCARECROW

  •  
  • AtSHR

    Arabidopsis SHORTROOT

  •  
  • bHLH

    basic helix-loop-helix

  •  
  • BnSCL1

    Brassica napus SCARECROW-like

  •  
  • CCaMK

    calcium- and calmodulin-dependent protein kinase

  •  
  • CIGR

    chitin-inducible cibberellin-responsive

  •  
  • CRM1

    chromosomal region maintenance 1

  •  
  • DLT

    DWARF AND LOW TILLERING

  •  
  • GA

    GIBBERELLIC ACID

  •  
  • GAI

    GA-INSENSITIVE

  •  
  • AtGAI

    Arabidopsis GAI

  •  
  • GID1

    GA-INSENSITIVE DWARF 1

  •  
  • GID2

    GA-INSENSITIVE DWARF 2

  •  
  • HAM

    HAIRY MERISTEM

  •  
  • HDAC

    histone deacetylase

  •  
  • IDP

    intrinsically disordered protein

  •  
  • IDR

    intrinsically disordered region

  •  
  • JA

    jasmonate

  •  
  • JAZ1

    jasmonate ZIM-domain 1

  •  
  • LeLs

    Lycopersicon esculentum lateral suppressor

  •  
  • LISCL

    Lilium longiflorum SCR-like

  •  
  • LRI

    leucine-rich region I

  •  
  • LRII

    leucine-rich region II

  •  
  • MoRF

    molecular recognition feature

  •  
  • NES

    nuclear export signal

  •  
  • NLS

    nuclear localization signal

  •  
  • NSP

    NODULATION SIGNALLING PATHWAY

  •  
  • NtGRAS1

    Nicotiana tabacum GRAS1

  •  
  • OsMOC1

    Oryza sativa monoculm 1

  •  
  • OsSCR

    Oryza sativa SCARECROW

  •  
  • OsSHR1

    Oryza sativa SHORTROOT 1

  •  
  • phyA

    phytochrome A

  •  
  • phyB

    phytochrome B

  •  
  • PIF

    PHYTOCHROME-INTERACTING FACTOR

  •  
  • QC

    quiescent centre

  •  
  • SAM

    shoot apical meristem

  •  
  • SCR

    SCARECROW

  •  
  • SH2

    Src homology 2

  •  
  • SHR

    SHORTROOT

  •  
  • SLY1

    SLEEPY1

  •  
  • SNUPN

    Snurportin-1

  •  
  • STAT

    signal transducer and activator of transcription

  •  
  • ZmSCR

    Zea mays SCR

We thank Dr Kimberley Snowden from The New Zealand Institute for Plant and Food Research for helpful comments prior to submission. We also thank the reviewers for giving detailed comments and suggestions that have been helpful in improving the paper.

FUNDING

Work of the authors is supported by the New Zealand Foundation for Research, Science and Technology [FRST contract C06X0812], and The New Zealand Institute for Plant & Food Research capability funding.

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