ROP GTPases Structure-Function and Signaling Pathways^,

Abstract

Interactions between receptor like kinases and guanyl nucleotide exchange factors together with identification of effector proteins reveal putative ROP GTPases signaling cascades.

Rho of Plants (ROP) proteins, also known as RACs, are the plant-specific subfamily of Rho small GTP-binding proteins, referred to here as small G proteins (Zheng and Yang, 2000a; Brembu et al., 2006; Eliáš and Klimeš, 2012). Like other members of the Ras superfamily of small G proteins, ROPs function as molecular switches due to changes in conformation upon GTP binding and hydrolysis (Berken and Wittinghofer, 2008). The conformational differences between the GTP- and GDP-bound states facilitate transient interactions with effector and regulatory proteins that, in turn, result in periodic activation/inactivation cycles of signaling cascades. Small G protein function is characterized by two central features: (1) due to inefficient GTP hydrolysis, these proteins remain in the GTP-bound active form for extended periods of time; and (2) due to the low dissociation coefficient of GDP, its release is inefficient and depends on enzymatic activity (Bourne et al., 1991; Vetter and Wittinghofer, 2001). Because of these features, the GTP-/GDP-dependent activation/inactivation cycles of small G proteins are regulated in time and space by GDP/GTP Exchange Factors (GEFs) that facilitate the release of GDP and GTPase-Activating Proteins (GAPs) that enhance GTP hydrolysis (Berken and Wittinghofer, 2008).

ROPs relay signaling from distinct plasma membrane domains; ROPs bind through posttranslational lipid modifications and interaction with membrane lipids (Bloch and Yalovsky, 2013). Thus, ROPs function as molecular switches that transduce intracellular and extracellular stimuli in a spatially regulated fashion, resulting in localized regulation of intracellular responses.

A third group of regulatory proteins, Rho GDP Dissociation Inhibitors (RhoGDIs), interact with high and low affinity with GDP- and GTP-bound Rho proteins, respectively, remove them from the membrane, and stabilize them in the cytoplasm until their reactivation by GEFs (Boulter and Garcia-Mata, 2010). The recycling induced by RhoGDIs is thought to maintain Rho proteins in distinct plasma membrane domains and to play an essential role in the establishment and maintenance of cell polarity (Johnson et al., 2011). As will be discussed below, in plants, RhoGDIs are critical for cell polarity maintenance.

Like Rho family proteins from fungi and mammalian cells, ROPs regulate the organization and dynamics of the actin and microtubule (MT) cytoskeleton (Box 1; Bashline et al., 2014; Hamada, 2014; Hashimoto, 2015; Henty-Ridilla et al., 2013; Pick, 2014), endocytosis and exocytosis, and the activation of NADPH oxidase and intracellular kinase cascades. Through effects on actin, MTs, vesicle trafficking, reactive oxygen species (ROS) production, and protein phosphorylation, ROPs regulate cell growth and shape, cytokinesis, subcellular protein localization, and responses to pathogens (Gu et al., 2004; Basu et al., 2008; Yalovsky et al., 2008; Yang, 2008; Nagawa et al., 2010; Bloch and Yalovsky, 2013; Kawano et al., 2014; Oda and Fukuda, 2014; Rivero et al., 2017). ROPs also are implicated in the regulation of abscisic acid (ABA) and auxin signaling and transport (Wu et al., 2011; Liao et al., 2017) and in the regulation of mRNA transcription (Zhang et al., 2016) and protein translation (Schepetilnikov et al., 2017) (Fig. 1; Table I).

Figure 1.

ROP signaling pathways. A, ROP activation/deactivation cycles. ROPs cycle between a GTP-bound active state and a GDP-bound inactive state. Activation is regulated by ROP-specific GEFs, inactivation is enhanced by ROP-specific GAPs, and recycling of type I ROPs between the cytosol and the plasma membrane is facilitated by RhoGDIs. Some GEFs are activated by peptide-activated, plasma membrane-associated RLKs. ROPs also are up-regulated by auxin by an as yet unknown pathway (dashed arrow) and inactivated by ABA, which enhances the degradation of ROPGEFs. B, The Dock family GEF homolog SPK1 is associated with the WAVE complex. Upon activation, ROP-GTP interacts with the WAVE SRA1 subunit and possibly other subunits, resulting in WAVE activation. Activated WAVE activates actin nucleation/branching induced by Arp2/3. C, ROP downstream signaling. ROP regulated pathways and known effectors. The dashed arrow indicates uncertainty regarding the cellular targets of ROP. CPL1 is regulated by ROP signaling but does not interact physically with ROPs. TOR interacts weakly with GTP-bound ROP2 and more strongly with the GDP-bound form.

Table I. Regulators and effectors of ROP signaling.

At, Arabidopsis thaliana; Hv, Hordeum vulgare (barley); Lj, Lotus japonicus; Mt, Medicago trancatula; Nt, Nicotiana tabacum (tobacco); Os, Oryza sativa (rice); Sl, Solanum lycopersicum (tomato); Zm, Zea mays (maize).

Name	Process	Cellular Function	Species	Reference
ROPs
AtROP1	Pollen tube growth, root growth, response to endophytic fungus		At	Fu et al. (2001); Gu et al. (2005); Lee et al. (2008); Venus and Oelmüller (2013)
AtROP2	Pavement cell growth, root hair initiation and growth, stomata opening, phytochrome-mediated light responses, ABA responses, transcriptional and translational regulation		At	Li et al. (2001); Fu et al. (2002, 2005); Jones et al. (2002); Fischer et al. (2006); Jeon et al. (2008); Ikeda et al. (2009); Hwang et al. (2011); Stanislas et al. (2015); Zhao et al. (2015); Hong et al. (2016); Zhang et al. (2016); Schepetilnikov et al. (2017); Wang et al. (2017)
AtROP3	Pollen growth, embryo and root development, auxin distribution		At	Expression data; Huang et al. (2014)
AtROP4	Pavement cell growth, root hair initiation and growth, pathogen response		At	Molendijk et al. (2001, 2008); Fu et al. (2005)
AtROP5	Pollen tube growth		At	Feng et al. (2016)
AtROP6	Pavement cell growth, root hair growth, ROS production, inhibition of endocytosis, root and lateral root development, salicylic acid-regulated pathogen response		At	Molendijk et al. (2001); Fu et al. (2005, 2009); Sorek et al. (2010); Poraty-Gavra et al. (2013); Venus and Oelmüller (2013); Wang et al. (2015)
AtROP7	Vascular development		At	Brembu et al. (2005); Wang et al. (2017)
AtROP8	Pollen nuclei migration in the central cell		At	Kawashima et al. (2014)
AtROP9	ABA and auxin responses, migration of pollen nuclei in pollen tubes		At	Li et al. (2013); Nibau et al. (2013); Choi et al. (2014)
AtROP10	ABA responses		At	Zheng et al. (2002); Xin et al. (2005); Choi et al. (2014)
AtROP11	ABA signaling, secondary cell wall patterning, root hair growth, inhibition of endocytosis		At	Bloch et al. (2005, 2011a, 2011b); Li et al. (2012b); Li and Liu (2012); Oda and Fukuda (2012); Yu et al. (2012)
ZmROP2	Regulation of asymmetric cell division during stomata development		Zm	Humphries et al. (2011)
ZmROP9	Regulation of asymmetric cell division during stomata development		Zm	Humphries et al. (2011)
OsRAC1	Pathogen response, lignin biosynthesis, activation of RBOH		Os	Kawasaki et al. (2006); Wong et al. (2007); Akamatsu et al. (2013); Nagano et al. (2016)
NtRAC5	Pollen tube growth		Nt	Kost et al. (1999); Klahre et al. (2006); Klahre and Kost (2006); Sun et al. (2015)
HvRACB	Susceptibility to powdery mildew, regulation of cell growth		Hv	Schultheiss et al. (2002, 2005); Pathuri et al. (2008)
HVRAC1	Susceptibility to powdery mildew, regulation of cell growth, ROS production		Hv	Pathuri et al. (2008)
HvRAC2	Susceptibility to powdery mildew, regulation of cell growth		Hv	Pathuri et al. (2008)
MtROP10	Regulation of symbiotic interaction with rhizobium		Mt	Lei et al. (2015)
LjROP6	Regulation of symbiotic interaction with rhizobium		Lj	Ke et al. (2012)
Cytoskeleton-associated ROP effectors
RIC1	Pollen tube growth, pavement cell growth, cell wall sensing	Actin severing and capping, cortical MT severing by KTN1	At	Fu et al. (2005, 2009); Lin et al. (2013a); Zhou et al. (2015); Higaki et al. (2017)
RIC3	Pollen tube growth	Actin disassembly	At	Gu et al. (2005); Lee et al. (2008)
RIC4	Pollen tube growth, pavement cell growth, root growth, response to endophytic fungus	Actin assembly and bundling	At	Fu et al. (2005, 2009); Gu et al. (2005); Lee et al. (2008); Venus and Oelmüller (2013)
RISAP	Pollen tube growth	Actin-dependent vesicle transport	Nt	Stephan et al. (2014)
NtADF1	Pollen tube growth	Actin depolymerization	Nt	Chen et al. (2002, 2003)
SCAR1-4	Pavement cell growth, trichome growth, stomata development and asymmetric division	Actin branching	At, Zm	Uhrig et al. (2007); Basu et al. (2008); Facette et al. (2015)
SPK1	Pavement cell and trichome growth	ROPGEF	At	Basu et al. (2008)
SRA1	Pavement cell and trichome growth	Actin branching	At	Uhrig et al. (2007); Basu et al. (2008)
ICR1	Vesicle trafficking, cell polarity	MT organization (putative)	At	Lavy et al. (2007); Hazak et al., (2010, 2014)
ICR2	Unknown	MT organization (putative)	At	Lavy et al. (2007)
MIDD1	Secondary cell wall deposition in metaxylem cells	Localized MT depolymerization through AtKinesin-13A	At	Mucha et al. (2010); Oda et al. (2010); Oda and Fukuda (2012)
HvRBK1	Basal resistance to pathogenic fungi	MT stability	Hv	Huesmann et al. (2012)
Non-cytoskeleton-associated ROP effectors
RIC7	Stomata movement	Vesicle trafficking	At	Hong et al. (2016)
HvRIC171	Pathogen response	Fungal penetration susceptibility	Hv	Schultheiss et al. (2008)
RBK1	Pathogen response	Receptor-like cytoplasmic kinase	At	Molendijk et al. (2008)
RBK2	Pathogen response	Receptor-like cytoplasmic kinase	At	Molendijk et al. (2008)
NCRK	Pathogen response	Receptor-like cytoplasmic kinase	At	Molendijk et al. (2008)
AtRLCK VI_A3	Pathogen response	Receptor-like cytoplasmic kinase	At	Reiner et al. (2015)
ABI1	ABA signaling	PP2C phosphatases	At	Li et al. (2012a)
ABI2	ABA signaling	PP2C phosphatases	At	Yu et al. (2012)
CPL1	Transcriptional regulation	Pol II CTD phosphatase	At	Zhang et al. (2016)
TOR	Auxin response	Translation control	At	Schepetilnikov et al. (2017)
RBOHB/D	Pathogen response	NADPH oxidase	Os	Wong et al. (2007); Kosami et al. (2014); Nagano et al. (2016)
ROPGAPs
MAGAP	Response to pathogenic fungi	Polar organization of cortical MT	Hv	Hoefle et al. (2011)
PHGAP1	Cell division plane selection	PH domain ROPGAP: positioning of the cortical division zone regulator kinesin-12 POK1	At	Stöckle et al. (2016)
PHGAP2	Cell division plane selection	PH domain ROPGAP: positioning of the cortical division zone regulator kinesin-12 POK1	At	Stöckle et al. (2016)
ROPGAP3	Secondary wall pits in metaxylem	Local MT destabilization	At	Oda and Fukuda (2012)
RhoGAP1	Pollen tube polar growth	Regulation of ROP activity	Nt	Klahre and Kost (2006); Sun et al. (2015)
REN1	Pollen tube polar growth	Regulation of ROP activity	At	Hwang et al. (2008)
ROPGAP1	Pollen tube polar growth	Actin organization	At	Fu et al. (2001); Hwang et al. (2010)
ROPGAP4	Response to oxygen deprivation	Regulation of ROS levels	At	Baxter-Burrell et al. (2002)
ROPGEFs
SPK1	Pavement cells and trichome growth	Dock-like ROPGEF	At	Basu et al. (2008)
ROPGEFs1/8/9/12/14	Pollen tube growth	Mediate PRK6 signaling and AtPRK2 signaling	At	Zhang and McCormick (2007); Chang et al. (2013)
ROPGEF4	Secondary wall pits in metaxylem, ABA responses, cell wall sensing	MT destabilization	At	Li and Liu (2012); Oda and Fukuda (2012); Huang et al. (2013)
ROPGEF7	Regulation of embryo and root development	Regulation of PLETHORA2 expression	At	Chen et al. (2011)
ROPGEF1	Root hair and embryo development, ABA responses	Mediate FER signaling	At	Duan et al. (2010); Li and Liu (2012); Liu et al. (2017)
ROPGEF10	ABA responses	Mediate FER signaling	At	Huang et al. (2013)
OsSWAP70	Responses to pathogens	Putative Dbl-like ROPGEF	Os	Yamaguchi et al. (2012)
RhoGDIs
AtRhoGDI1/SCN1	Regulation of ROP2 and RBOHC distribution at root hair initiation sites, pollen tube growth		At	Carol et al. (2005); Hwang et al. (2010); Feng et al. (2016)
AtRhoGDI2a	Pollen tube growth		At	Hwang et al. (2010); Feng et al. (2016)
AtRhoGDI2b	Pollen tube growth		At	Hwang et al. (2010); Feng et al. (2016)
RLKs
FERONIA	Root hair growth, pollen tube rupture, defense responses	Cell wall sensor and a signaling scaffold, signals through ROPGEFs	At	Duan et al. (2010, 2014); Yu et al. (2012); Huang et al. (2013); Stegmann et al. (2017)
THESEUS	Defense responses	Cell wall sensor signaling through ROPGEFs	At	Höfte (2015); Qu et al. (2017a)
PRK6	Pollen tube growth	LURE receptor signaling through ROPGEFs	At	Takeuchi and Higashiyama (2016)
AtPRK2	Pollen tube growth	Plasma membrane LRK that signals through ROPGEFs	At	Chang et al. (2013)
SlPRK1	Pollen tube germination and growth	Plasma membrane LRK signaling through ROGEFS	Sl	Kaothien et al. (2005)
SlPRK2	Pollen tube germination and growth	Plasma membrane LRK signaling through ROGEFS	Sl	Kaothien et al. (2005)
OsCEBiP/OsCERK1	Pathogen response	Plasma membrane LRK that senses chitin and signals through ROPGEFs	Os	Akamatsu et al. (2013)
MtNFP	Symbiotic interaction with rhizobium	Interacts with GTP-MtROP10	Mt	Lei et al. (2015)
LjNFR5	Symbiotic interaction with rhizobium	Interacts with GTP-LjROP6	Lj	Ke et al. (2012)

In recent years, there have been a plethora of new studies on ROPs. Studies on pollen tube and root hair development and plant immune responses have indicated that ROPGEFs are activated by plasma membrane-bound Receptor-Like Kinases (RLKs). The next steps will be elucidation of ROP signaling pathways from identification of the upstream receptors to mechanisms of ROPGEF and ROP activation to identification of downstream effectors and their functions (Fig. 1). In this Update, we aim to summarize our current understanding of ROP structure/function and signaling with an emphasis on new findings that have not been subject to previous reviews. Because of space limitations and focus, we are unable to cover all published literature in the field, including the proposed functions of ROPs in regulating transcription and translation. Thus, we apologize to colleagues whose works are not cited.

ROP EVOLUTION, STRUCTURE, AND FUNCTION

Evolution

ROPs have been identified in red and green algae and are ubiquitous in embryophytes, where they form multimember protein families (Brembu et al., 2006; Eliáš and Klimeš, 2012). The number of ROP proteins varies: there are two in the lycophyte Selaginella moellendorffii, four in the moss Physcomitrella patens, four in the gymnosperm Pinus taeda (loblolly pine), seven in rice (Oryza sativa), nine in maize (Zea mays), nine in tomato (Solanum lycopersicum), 11 in Arabidopsis (Arabidopsis thaliana), and 13 in poplar (Populus spp.; Fowler, 2010). Phylogenetic analyses indicate that the expansion of the ROP family resulted from multiple gene duplication events (Winge et al., 2000; Fowler, 2010). Interestingly, ROPs and ROPGEFs or other Rho GTPases have not been identified in some green algae, including Chlamydomonas reinhardtii and Volvox carteri, indicating a loss of the ROP/Rho signaling module in these organisms. Phylogenetic analyses indicated that ROPs emerged prior to the diversification of the RAC and Cdc42 subfamilies and represent a unique subfamily of Rho GTPases (Zheng and Yang, 2000a; Vernoud et al., 2003; Brembu et al., 2006; Boureux et al., 2007; Rojas et al., 2012).

Analysis of Rho GTPases from divergent eukaryotes revealed that early phylogenetic analyses of the Rho GTPases were biased by a focus on the fungal and animal members of the family. For example, Cdc42, which plays a central role in the regulation of cell polarity (Johnson and Pringle, 1990; Woods and Lew, 2017), emerged with the opisthokonts (fungi and metazoa) but is absent in other eukaryotes. The amoeba Dictyostelium discoideum has proteins that have been historically called Racs but are as divergent from each other as are Rac, Cdc42, and Rho (Eliáš and Klimeš, 2012). The analysis by those authors further confirmed that ROPs are unique to the plant kingdom (Eliáš and Klimeš, 2012). Collectively, the phylogenetic analyses carried out by several groups justify the name Rho of Plants (Zheng and Yang, 2000a). As will be discussed in this Update, ROPs combine functions of Rac, Cdc42, and possibly other Rho proteins, as could be expected from their early divergence during evolution. Plants also possess a unique family of ROPGEFs (Berken et al., 2005; Gu et al., 2006; Elias, 2008) and forms of the Cdc42 Rac Interacting Binding (CRIB) domain containing ROPGAPs (Wu et al., 2000). Furthermore, most of the ROP effectors identified to date are unique to plants. Hence, it appears that many components of the ROP signaling module are unique to plants.

Structure/Function

ROPs have molecular masses of 21 to 24 kD. Like other members of the Ras superfamily, ROPs are composed of an N-terminal catalytic G-domain where nucleotide and effector binding take place and a C-terminal hypervariable domain (HVR), which is responsible for subcellular targeting (Fig. 2A). The G-domain contains five conserved sequence motifs known as the G-box motifs (G1–G5), where nucleotide binding, GTP hydrolysis, and Mg²⁺ binding take place. The G2 and G3 motifs also are known as the switch I and switch II loops; these regions have different conformations in GDP- and GTP-bound states and are essential for the transient interaction with effector and regulatory proteins. Within the G domain, ROPs also contain a helical domain unique to the Rho family known as the insert region or αi. The structural conservation of the G-domain has been confirmed by three-dimensional crystal structures of the GDP-bound Arabidopsis ROP9 (Sørmo et al., 2006), ROP4 in complex with the catalytic PRONE (for Plant-Specific ROP Nucleotide Exchanger) domain of ROPGEF3 (Thomas et al., 2007), ROP5 (Thomas and Berken, 2010; Protein Data Bank code 3BWD), the nucleotide-free ROP7 in complex with the PRONE domain of ROPGEF8 (Thomas et al., 2009), and GTP-bound rice RAC1 (Kosami et al., 2014; Fig. 2, B–D).

Figure 2.

Structures of ROPs. A, Schematic model highlighting the conserved G-box motifs (G1–G5), switch I and II domains, the insert region, and the HVR. The positions of the conserved residues mutated in the constitutively active and dominant-negative mutations are highlighted by green and magenta arrowheads, respectively. C21 and C156 are conserved G domain Cys residues that undergo activation-dependent transient S-acylation (black arrows). The HVR terminates with a canonical CaaX box geranylgeranylation motif and a PBR in type I ROPs or with a GC-CG box and a proximal PBR in type II ROPs. B, Three-dimensional structure of GTP-bound OsRAC1 [OsRAC1 (GMPPNP)]. C, Three-dimensional structure of the GDP-bound AtROP9 [AtROP9 (GDP)]. In B and C, the switch I, switch II, and insert regions are colored green. GMPPNP and GDP are shown as stick models (red, oxygen; blue, nitrogen; orange, phosphorus). The Mg²⁺ ions are shown as yellow spheres. D, Superimposed structures of OsRAC1 (GMPPNP) and AtROP9 (GDP). The main chains of OsRAC1 (GMPPNP) and AtROP9 (GDP) (chain B; Protein Data Bank code 2J0V) were superimposed using PyMOL. Regions of the OsRAC1 and AtROP9 proteins are colored blue and pink, respectively. The side chains of four key switch I residues in OsRAC1 (Val-43, Phe-44, Asp-45, and Tyr-39) and equivalent residues in AtROP9 (Val-39, Phe-40, Asp-41, and Tyr-35) are shown in stick representation. B to D were prepared and contributed by Izuru Ohki and Chojiro Kojima and are based on their article describing the structure of GTP-bound OsRAC1 (Kosami et al., 2014).

Based on structural conservation and known mutations in Ras (Feig, 1999), it was predicted and subsequently shown that point mutations in a G1 Gly (G15 in Arabidopsis ROP1–ROP7 and ROP9; G17 in ROP10 and ROP11; and G27 in ROP8) or G3 Gln (Q64 in Arabidopsis ROP1–ROP7 and ROP9; Q66 in ROP10 and ROP11; and Q76 in ROP8) would prevent GTP hydrolysis, resulting in GTP-locked, constitutively active (CA) mutants. Conversely, mutations in a G1 Thr (T20 in Arabidopsis ROP1–ROP6, ROP7, and ROP9; T22 in ROP10 and ROP11; and T32 in ROP8) or a G4 Asp (D121 in Arabidopsis ROP1–ROP6, ROP7, and ROP9; D123 in ROP10 and ROP11; and D133 in ROP8) are dominant negative (DN), reducing affinity for nucleotides and stabilizing the interaction with GEFs, thereby preventing ROP activation (Berken and Wittinghofer, 2008). In addition to the structural conservation of ROPs, the functional conservation of the constitutively active and dominant-negative mutations was confirmed by biochemical assays (Berken et al., 2005; Sorek et al., 2010; Kosami et al., 2014). The constitutively active and dominant-negative ROP mutants have been used extensively to study ROP function.

A comparison between the structure of the nonhydrolyzable GTP analog guanosine 5′-[β,γ-imido]triphosphate [Gpp(NH)p]-bound rice OsRAC1 and the GDP-bound AtROP9 revealed the conformational differences between the GDP- and GTP-bound states of ROPs (Fig. 2, B–D; Kosami et al., 2014). The structural analysis showed conformational differences in the switch I and switch II domains and the insert region (Fig. 2, B–D). However, the conformational changes in the switch I domain are the most critical for function. It was shown that GTP binding by OsRAC1 results in surface exposure of switch I Glu (D45) and a Tyr (Y39); both are required for its interaction with rice Respiratory Burst Oxidase Homolog (OsRBOHB) NADPH oxidase (Kosami et al., 2014). D45 and Y39 are highly conserved, and it will be interesting to examine whether they play central roles in the interaction of ROPs with other effectors. The structure of ROPs solved so far reveal differences in the structures of the switch II and insert regions (Berken and Wittinghofer, 2008; Thomas and Berken, 2010; Kosami et al., 2014). It is not known yet whether these differences also imply different substrate and regulatory specificities. Collectively, the sequence and structural and functional conservation of ROPs indicate that, regardless of their early divergence, they function in a similar fashion to the well-characterized Rho GTPases from fungi and metazoa.

The C-terminal HVRs of ROPs are composed of Cys-containing sequence motifs, which undergo posttranslational lipid modifications, and proximal Arg-Lys-rich polybasic regions (PBRs). Type I ROPs (in Arabidopsis ROP1–ROP8) terminate with a canonical CaaL box motif (where C is Cys, a₁ and a₂ are aliphatic residues, and L is Leu) in which the Cys residue is modified by the C20 isoprenyl lipid geranylgeranyl. Type II ROPs terminate with GC-CG boxes (where G is Gly and C is Cys) in which the two Cys residues are separated by five or six aliphatic residues and undergo S-acylation by the C16 palmitate or C18 stearate fatty acids. Type II ROPs are likely not regulated by RhoGDIs, since geranylgeranylation is required for the interaction between Rho protein and RhoGDI. The PBRs in animal Rhos, RACs, and Cdc42 enhance membrane attachment by interaction with phosphatidylphosphoinositide (4,5) diphosphate (PIP2) and phosphatidylphosphoinositide (3,4,5) triphosphate (PIP3; Heo et al., 2006). The PBR is required for the plasma membrane attachment of type II ROPs (Lavy and Yalovsky, 2006), possibly by facilitating the interaction with PIPs. In addition, ROPs undergo activation-dependent transient S-acylation on conserved G-domain Cys residues (C21 and C156 in AtROP6). The transient S-acylation stabilizes the interaction of ROPs with the plasma membrane, leads to their accumulation in lipid rafts, and is required for their function (Sorek et al., 2010, 2011, 2017).

ROP REGULATORS: GEFS, GAPS, AND RHOGDI

GEFs

Plants have two major types of ROP GEFs: (1) a plant-unique family known as ROPGEFs that contain the PRONE catalytic domain (Berken et al., 2005; Gu et al., 2006); and (2) distantly related homologs of the animal Ced5, Dock180, Myoblast city (CDM) Zizimin Homology (CZH) RhoGEFs. The CZH RhoGEFs share three tandem Doc Homology Regions (DHRs), of which DHR2 has GEF activity. A single CZH RhoGEF homolog called SPIKE1 (SPK1) was identified in Arabidopsis (Qiu et al., 2002), and it was shown that either the full-length protein or its DHR2 domain can catalyze ROP nucleotide exchange (Basu et al., 2008). The rice OsSWAP70A and OsSWAP70B bear homology to Diffuse B cell lymphoma (Dbl)-type RhoGEFs that are prevalent in yeast and mammalian cells. OsSWAP70A and OsSWAP70B have GEF activity and were suggested to function as GEFs for ROPs (Yamaguchi et al., 2012).

ROPGEFs

ROPGEFs catalyze nucleotide exchange on ROPs but not on nonplant Rho GTPases (Berken et al., 2005). The specificity for ROPs is likely associated with the presence of a small Gly residue in the insert regions of ROPs at a position that, in nonplant ROPs, is occupied by a large Arg residue (Thomas et al., 2007; Berken and Wittinghofer, 2008). Structural studies revealed that ROPGEFs function as dimers and interact with two ROP molecules. Interestingly, it was found that the structure of ROPGEFs is different from that of the Dbl RhoGEFs, but the mechanism of guanyl nucleotide exchange is similar (Thomas et al., 2007). Furthermore, based on the cocrystal structure of ROP4 and PRONE8, it was predicted that the interaction between ROPs and ROPGEFs could be regulated by phosphorylation (Thomas et al., 2007; Berken and Wittinghofer, 2008; Thomas and Berken, 2010). The catalytic PRONE domain of ROPGEFs is constitutively active, and its function is regulated by the ROPGEF C-terminal domain (Berken et al., 2005; Gu et al., 2006). Studies in pollen tubes indicated that ROPGEFs interact with membrane-associated RLKs, which phosphorylate the C-terminal domain and relieve the autoinhibition of the PRONE domain, leading to ROPGEF activation (Kaothien et al., 2005; Zhang and McCormick, 2007; Chang et al., 2013). Similarly, studies in root hairs have demonstrated that ROPGEFs function downstream of the RLK FERONIA (FER; Duan et al., 2010; Yu et al., 2012; Huang et al., 2013). Thus, the activation of at least some ROPGEFs depends on intercellular signaling that involves phosphorylation by membrane-bound RLKs (Fig. 3).

Figure 3.

ROP signaling pathways. A schematic overview of identified ROP signaling pathways is shown. Components of the signaling pathways mentioned in the text are noted, and their relationships to each other are indicated by arrows. The question mark notes an area of insufficient data. From top to bottom in each section are listed the upstream signal, the receptors that relay the signal into the cell, the regulation of ROP activity, and the downstream effectors that modulate the cellular response.

ROPGEFs have only been identified in red algae (Rhodophyta) and green plants (Chloroplastida) and appear to be a family of proteins unique to the plant kingdom (Archeaplastida/Plantae) that are not of a cyanobacterial origin (Elias, 2008). They form protein families with numbers ranging from two in S. moellendorffii, to six in P. patens, to 14 in Arabidopsis (Berken et al., 2005; Gu et al., 2006; Eklund et al., 2010). Arabidopsis ROPGEF mutants often display mild phenotypes (Li and Liu, 2012; Chang et al., 2013; Huang et al., 2013; Zhao et al., 2015; Wang et al., 2017), indicating that they likely function redundantly. Two reports indicated that ROPGEF1 and ROPGEF7 have specific functions during embryo development. ROPGEF7 RNA interference (RNAi) plants displayed compromised stem cell maintenance in embryo and roots (Chen et al., 2011). The ropgef1 mutants are characterized by defective embryo development and embryo auxin distribution and a mild gravitropic response phenotype (Liu et al., 2017), suggesting that the ROPGEF1 function in embryos is unique. However, ROPGEF1 might function in conjunction with other ROPGEFs in root gravitropic responses.

ROPGEFs are soluble proteins that are, however, observed in the plasma membrane, suggesting that they interact with components at the plasma membranes that facilitate their attachment to the membrane. Deletion of the AtROPGEF12 C-terminal domain abolished the membrane association following transient expression in tobacco (Nicotiana tabacum) pollen tubes. Upon transient expression in tobacco pollen tubes, several ROPGEF-GFP fusion proteins were detected at the pollen tube tip (Gu et al., 2006). In embryos, the AtROPGEF1-GFP fusion protein, driven by the ROPGEF1 prompter, was localized in a polarized manner (Liu et al., 2017). Taken together, these data indicate that ROPGEFs likely activate ROPs in the plasma membrane and that their spatial distribution influences the site or sites of ROP activation.

SPK1

SPK1 was discovered in a forward genetic screen in Arabidopsis for mutants with altered trichome development. spk1 mutants are seedling lethal and display a pleotropic phenotype that includes stunted plant size and abnormal leaf pavement cells and trichome growth, cotyledon shape, and cytoskeleton organization (Qiu et al., 2002). In in vitro pull-down assays, SPK1 interacts weakly with GDP-bound and nucleotide-free type I and type II ROPs but not with human RAC1 (Basu et al., 2008), and similar results were obtained in yeast two-hybrid assays (Uhrig et al., 2007). Coimmunoprecipitation and protein interaction assays indicated that SPK1 is associated with the Suppressor of cAMP Receptor (SCAR)/Wiskot-Aldrich syndrome protein-family Verprolin homology protein (WAVE) complex, which activates actin nucleation/branching by the Actin-Related Protein2/3 (Arp2/3) complex (Uhrig et al., 2007; Basu et al., 2008). GTP-bound type I ROPs interact with the SCAR/WAVE complex subunit SRA1 (also known as PIROGI/KLUNKER; Basu et al., 2004, 2008) or with other subunits of the complex (Uhrig et al., 2007). Importantly, analysis of SPK1, SRA1, and Arp2/3 mutants demonstrated that they function in the same pathway (Basu et al., 2008). Taken together, the genetic and biochemical analyses demonstrate that SPK1, type I ROPs (possibly ROP2 and ROP4), SCAR/WAVE, and Arp2/3 complexes regulate actin nucleation (Figs. 4 and 5).

Figure 4.

Mechanisms of spatial ROP activity regulation. A, Local activation of ROPs in the pollen tube. ROPs are activated at the tip by the activation of ROPGEFs by associated RLKs. CRIB domain ROPGAPs inactivate ROPs at the shank to maintain polar growth. PH domain GAPs are localized in subapical cytoplasmic vesicles, restricting ROP active domains. RhoGDIs recycle ROPs from the membrane. B, Formation of active ROP domains in differentiating metaxylem. During metaxylem differentiation, ROP11 is coexpressed with ROPGEF4 and ROPGAP3, resulting in the formation of active ROP11 domains and global inactivation. The size of the ROP11 domains is restricted by MTs and the MT- and plasma membrane-associated protein IQD13. C, Activation of ROPs by SPK1 and the formation of active actin nucleation/branching domains. SPK1 is associated with the WAVE complex. Activation of ROPs by SPK1 causes their association with the SRA1 subunit of the WAVE complex and its activation. Active WAVE can then activate actin nucleation/branching by Arp2/3. ER, Endoplasmic reticulum; PM, plasma membrane.

Figure 5.

ROP regulation of cytoskeleton organization. A, Cortical MT reordering by ROP6, RIC1, and KTN1. RIC1 interacts physically with and promotes the MT severing by KTN1, leading to MT reordering. B, Localized cortical MT depolymerization. In developing metaxylem, MT depolarization takes place by the recruitment of MIDD1 and Kinesin13A to active ROP11 domains. C, Actin dynamics in tip-growing pollen tubes. ROP regulates actin assembly by two counteracting pathways. RIC4 promotes assembly, whereas RIC3 causes disassembly by stimulating Ca²⁺ influx into the cytoplasm. D, Actin nucleation/branching in trichomes and pavement cells. ROPs and the ROPGEF SPK1 interact with the SCAR/WAVE complex and promote actin filament nucleation/branching.

In developing trichomes, nucleation of actin at the tip depends on the WAVE-dependent function of Arp2/3 (Yanagisawa et al., 2015), which is likely locally regulated by SPK1. Thus, the combinatorial regulation of SPK1, ROPs, and WAVE subcellular distribution is very likely responsible for localized actin nucleation. It is not known how SPK1 function is regulated; like ROPGEFs, it may be activated by upstream signaling. It is also not known whether SPK1 regulates additional pathways. In P. patens, there are six SPK1 homologs, whereas in S. moellendorffii and Arabidopsis, SPK1 is encoded by a single gene (Eklund et al., 2010). Sequence analysis of SPK1 homologs from additional plant species will be required before it can be determined whether the reduction in SPK1 homologs followed an evolutionary trend. Genetic analysis in P. patens should reveal whether SPK1 function in the regulation of actin nucleation is evolutionarily conserved and whether it had additional functions that were lost during evolution.

OsSWAP70

The rice proteins OsSWAP70A and OsSWAP70B contain Pleckstrin Homology (PH) and Diffuse B cell Homology domains similar to many animal RhoGEFs and display similarity to human SWAP70 RhoGEF (Shinohara et al., 2002; Yamaguchi et al., 2012). OsSWAP70A displays GEF activity toward OsRAC1 and enhances OsRAC1-dependent ROS production. Surprisingly, however, OsSWAP70A interacts with both constitutively active and dominant-negative OsRAC1 and with Osrac7^CA mutants but not with the dominant-negative form Osrac7^DN. Furthermore, OsSWAP70B does not interact with any rice ROP (Yamaguchi et al., 2012). SWAP70 homologs have been identified in other plants (Yamaguchi et al., 2012). However, given that OsSWAP70A interacts with GTP-bound RAC1, which contradicts a long-held dogma regarding RhoGEF activity (Berken and Wittinghofer, 2008; Thomas and Berken, 2010), and in the absence of loss-of-function mutants to support the biochemical analysis, more work is required to determine whether the plant SWAP70 homologs are, indeed, ROPGEFs.

ROP GAPs

Fungal and metazoan RhoGAPs share a conserved GAP domain that contains an invariable Arg residue required for the catalytic activity. In addition to the GAP domain, RhoGAPs usually contain additional domains that are required for activity and specificity (Lamarche and Hall, 1994; Rittinger et al., 1997; Graham et al., 1999). In plants, there are two types of GAPs for ROPs known as ROPGAPs and RENGAPs (ROP1 Enhancer GAPs; Wu et al., 2000; Hwang et al., 2008; Eklund et al., 2010).

ROPGAPs

ROPGAPs are characterized by a CRIB domain in addition to the GAP domain. The CRIB domain is found in Rho effectors such as WASP in animal cells and ROP Interacting CRIB-containing proteins (RICs) in plants and not in fungal and metazoan RhoGAPs. In ROPGAPs, the CRIB domain is required for activity, as it enhances the binding between ROPs and ROPGAPs (Wu et al., 2000; Schaefer et al., 2011). The CRIB domain also is required for the subcellular localization of ROPGAPs (Wu et al., 2000; Klahre and Kost, 2006). ROPGAPs function as dimers in a 2:2 stoichiometry with ROPs (Schaefer et al., 2011). Arabidopsis AtROPGAP2 has differential binding affinities for various ROPs (Schaefer et al., 2011), and phosphorylation regulates the interaction of tobacco RhoGAP1 and an interacting 14-3-3 protein (Klahre and Kost, 2006), suggesting how ROPGAP function might be regulated.

Following the transient expression of NtRhoGAP1 in tobacco pollen tubes, the protein was localized at the plasma membrane below the tip (Klahre and Kost, 2006; Sun et al., 2015). In agreement with this finding, the ectopic expression of Arabidopsis AtROPGAP1 in tobacco pollen tubes restricts the active ROP domain (Hwang et al., 2010; Fig. 4). ROPGAPs also are important rheostats of ROP signaling. In Arabidopsis, oxygen deprivation induces the ROP-regulated production of ROS. In turn, ROS induce the expression of ROPGAP4, which down-regulates ROP activity and contributes to plant tolerance under anoxic conditions (Baxter-Burrell et al., 2002). Gene expression studies in apple (Malus domestica) showed that ethylene coordinately down-regulates ROPs, ROPGAPs, and ROPGEFs, leading to altered apoplastic hydrogen peroxide levels, suggesting that the regulation of ROP activation status has an important function in the regulation of different abiotic stresses and physiological responses (Zermiani et al., 2015).

RENGAPs

RENGAPs were initially identified by a mutant screen in pollen overexpressing ROP1. RENGAPs contain a PIP-binding PH domain in addition to the GAP domain (Hwang et al., 2008). In pollen tubes, REN1 (also known as RENGAP1) is localized in subapical cytoplasmic vesicles, and its PH domain is not required for this localization. ren1 mutant pollen tubes are swollen, indicating that REN1 functions to restrict the active ROP domain to the pollen tube tips (Hwang et al., 2008; Fig. 4).

RENGAP2 and RENGAP3 interact with the mitosis-specific kinesin-12 POK1. During interphase, RENGAP2 and RENGAP3 are cytoplasmic, but during mitosis, they are localized to the cortical division zone (CDZ). The rengap2 rengap3 double mutant displays mild cell division reorientation, indicating that rapid inactivation of ROPs at the CDZ is required for the correct orientation of cell divisions (Stöckle et al., 2016). The division of labor between ROPGAPs and RENGAPs is currently unknown, and in some cases, such as during mitosis, they might function redundantly (Stöckle et al., 2016).

Formation of Active ROP Domains by the Combinatorial Function of ROPGEFs and ROPGAPs

During metaxylem development, the simultaneous expression of ROP11, ROPGEF4, and ROPGAP3 results in the formation of active ROP domains. The formation of the active ROP11 domain was recapitulated by transient coexpression of ROP11, the PRONE domain of ROPGEF4 (ROPGEF4PRONE), and ROPGAP3. Within the domain, ROP11 and ROPGEF4 are localized in the center and ROPGAP3 in the circumference (Oda and Fukuda, 2012). These experiments demonstrated that ROP-ROPGEF-ROPGAP modules have self-organization properties, suggesting that coordinated expression and localized activation of ROPGEFs together can result in cellular patterning (Figs. 3 and 4).

RhoGDIs

Plant RhoGDIs are structurally conserved (Bischoff et al., 2000). Structural and biochemical analyses have established that the interaction between Rho proteins and RhoGDI under physiological conditions depends on prenylation by geranylgeranyl of the GTPases and involves a hydrophobic pocket in the RhoGDI (Hoffman et al., 2000; DerMardirossian and Bokoch, 2005). In plants, type I ROPs are geranylgeranylated (Sorek et al., 2011, 2017), but type II ROPs do not contain a conserved geranylgeranylation CaaL box motif and are S-acylated but not prenylated (Lavy et al., 2002; Lavy and Yalovsky, 2006). Therefore, it is likely that type II ROPs are not regulated by RhoGDIs.

Transient expression of NtRhoGDI1 in tobacco pollen tubes indicated that it functions in the recycling of ROPs and in their focusing at the pollen tube tip (Klahre et al., 2006; Sun et al., 2015). Analysis of RhoGDI mutants in Arabidopsis showed their involvement in the maintenance of ROP activation domains and ROP stability (Carol et al., 2005; Feng et al., 2016). There are three RhoGDIs in Arabidopsis called RhoGDI1, RhoGDI2a, and RhoGDI2b. RhoGDI1 mutants, also known as SUPERCENTIPIDE1 (SCN1), develop multiple root hair (RH) initials instead of a single RH in each trichoblast. In scn1 mutants, GFP-ROP2 was observed along the trichoblast membrane and ROS production was detected at multiple sites, in contrast to wild-type plants, where ROP2 accumulation and ROS formation were detected specifically at the RH initiation sites (Carol et al., 2005). Interestingly, the phenotype of scn1 fits a mathematical model predicting that a loss of RhoGDI would result in traveling waves instead of a focal polar domain (Jilkine et al., 2007).

Silencing of RhoGDI2a by RNAi resulted in the development of shorter pollen tubes and expansion of ROP distribution; however, possible redundancy with other RhoGDIs was noted (Hwang et al., 2010). rhogdi1 rhogdi2a rhogdi2b triple mutants develop shorter and wider pollen tubes with higher levels of ROPs in the pollen tube tip plasma membranes and expanded ROP activation domains. In addition, protein immunoblots indicated that the overall ROP levels are lower in the RhoGDI triple mutants (Feng et al., 2016). In mammalian cells, reduction in RhoGDI1 levels by small interfering RNA resulted in higher levels of plasma membrane-associated Rho proteins concomitant with reduction in their levels (Boulter et al., 2010). It appears, therefore, that RhoGDIs have an evolutionarily conserved function in the regulation of homeostasis of Rho GTPases. Notably, although the RhoGDI triple mutant plants are less fertile than wild-type plants, they produce viable seeds and their sporophytes do not seem to be significantly different from wild-type controls (Feng et al., 2016). This indicates that type I ROPs are functional and, possibly, that there is redundancy between type I and type II ROPs.

ROP FUNCTION IN TIP AND DIFFUSE GROWTH

Regulation of Pollen Tube Tip Growth

Pollen tubes are characterized by highly polarized oscillatory tip growth that depends on differential cell wall composition, tip-focused vesicle transport, specialized actin organization, and Ca²⁺, ROS, and pH gradients (Cárdenas et al., 2008; Cheung and Wu, 2008; Chebli et al., 2012; Bloch et al., 2016; Michard et al., 2017). Actin organizes into multiple forms in different regions of pollen tubes, including thick long actin cables that are axially aligned in the shank, a mesh ring or funnel-like F-actin structure known as the actin fringe in the subapical region, and a group of fine and less abundant F-actin in the pollen tube apex. Various forms of F-actin structures participate in cytoplasmic streaming and vesicle trafficking to and from the tip and in the accumulation of exocytotic vesicles at the pollen tube apex (Cárdenas et al., 2008; Cheung and Wu, 2008; Qin and Yang, 2011; Rounds et al., 2014; Fu, 2015; Qu et al., 2017b). ROP signaling regulates pollen growth and was shown to affect actin organization, Ca²⁺ levels, and vesicle trafficking.

LUREs are ovule-derived small peptides that attract the pollen tube toward ovules (Dresselhaus and Franklin-Tong, 2013; Higashiyama and Takeuchi, 2015). Several LURE1 receptors have been identified (Takeuchi and Higashiyama, 2016; Wang et al., 2016). It was shown that Pollen-specific Receptor Kinase6 (PRK6) is a LURE1 receptor that localizes to the pollen tube tip and interacts with the regulatory C-terminal domain of pollen-expressed ROPGEF8/9/12/13 (Takeuchi and Higashiyama, 2016; Fig. 3). In agreement, ROPGEF8/9/12-GFP fusion proteins are localized at the pollen tube tip following transient expression in tobacco pollen tubes, and AtROPGEF9 has high specific activity toward the pollen-expressed AtROP1 (Gu et al., 2006). Tomato pollen PRK1 and PRK2 interact with a ROPGEF, originally called Kinase Protein Partner (Kaothien et al., 2005). Arabidopsis PRK2a interacts with ROPGEF12 in yeast through the C-terminal regulatory domain. Coexpression of PRK2a and ROPGEF12 in the tobacco pollen tube induces isotropic growth, and a phospho-mimicking mutation at the C-terminal domain activates ROPGEF12 (Zhang and McCormick, 2007). Taken together, the results from studies described in this section strongly suggest that, upon LURE perception, PRKs activate pollen-specific ROPGEFs that, in turn, activate pollen-specific ROPs. It should be noted that ROPGEF-dependent ROP activation by LUREs has yet to be demonstrated.

Ectopic expression of constitutively active ROP mutants results in isotropic pollen tube growth, whereas the expression of dominant-negative ROP mutant or ROPGAPs inhibits pollen tube growth and is associated with changes in actin organization (Fu et al., 2001; Klahre et al., 2006; Klahre and Kost, 2006). Several studies addressed the mechanisms by which ROPs regulate actin organization. Coexpression studies in the tobacco pollen tube suggest that the tobacco ROP NtRAC5 can lead to the phosphorylation and suppression of actin depolymerization activity of Actin Depolymerizing Factor (ADF; also known as Cofilin; Chen et al., 2003).

Three RIC proteins were implicated in the regulation of pollen tube growth. RIC3 function is associated with increased intracellular Ca²⁺ levels, leading to actin depolymerization, whereas RIC4 is implicated in actin polymerization. The antagonistic functions of RIC3 and RIC4 are suggested to regulate actin dynamics in pollen tubes (Fig. 5; Gu et al., 2005). The mechanisms of RIC3 and RIC4 function remain to be elucidated. RIC1 regulates pollen growth through its F-actin-severing and -capping activities. RIC1 displays oscillatory localization at the apical plasma membrane of the pollen tube tip, which depends on an intact CRIB domain. The severing activity of plasma membrane-localized RIC1 mediates the release of F-actin from the pollen tube apical plasma membrane into the cytoplasm. RIC1 also contributes to severing F-actin and capping the barbed ends in the cytoplasm (Zhou et al., 2015).

An effector of the tobacco ROP called RAC5 Interacting Subapical Pollen tube protein (RISAP) was identified in a yeast two-hybrid screen. RISAP-XFP fusion proteins localize to the trans-Golgi network and interact with actin and with the globular tail domain of a tobacco myosin XI through a protein called Domain of Unknown Function593. RISAP overexpression interferes with apical membrane trafficking and blocks tip growth. It was suggested that RISAP functions in tip-directed vesicle trafficking in a ROP-dependent fashion (Stephan et al., 2014). It would be of interest to analyze RISAP loss-of-function mutants to further examine its function. An Arabidopsis protein (At1g18990) is 43% identical to RISAP, and homologs also have been identified in other plant species. Thus, it should be possible to examine the conservation of RISAP function.

Using RabA4d as an exocytotic marker in transient expression assays in tobacco pollen tubes, it was shown that its accumulation displays an oscillatory pattern that precedes the growth phase and is regulated by F-actin (Lee et al., 2008). Analysis of the exocyst Sec3a subunit mutant demonstrated that polarized exocytosis is required for both the selection of the pollen tube initiation site and polarized growth (Bloch et al., 2016). It is not known whether ROPs are involved directly in the regulation of polarized exocytosis in pollen. An analysis of multiple pollen-expressed ROP mutants has not yet been described, and it is not known whether they are required for selection of the pollen tube germination sites, similar to the function of Cdc42 in the selection of the bud site in Saccharomyces cerevisiae (Johnson and Pringle, 1990; Woods and Lew, 2017).

Regulation of RH Tip Growth

RH tip growth is characterized by actin/myosin-dependent vesicle trafficking toward the growing tip and cell wall relaxation at the tip (Mendrinna and Persson, 2015). RHs form close to the rootward side of trichoblasts, and their initiation and growth require tip-focused cell wall relaxation that depends on ROS-dependent pH fluctuations (Monshausen et al., 2007) and the activity of cell wall expansins (Cho and Cosgrove, 2002). During RH growth, F-actin forms a fine meshwork at the tip and longitudinally oriented filaments axially along the RH shank. When growth ceases, the actin filaments also are detected at the RH tip (Jones et al., 2002; Bloch et al., 2005, 2011b). ROPs regulate both RH initiation and growth. Prior to RH initiation, ROP2/4/6 accumulate at the site of future RH formation in response to a local auxin gradient (Molendijk et al., 2001; Jones et al., 2002; Fischer et al., 2006; Ikeda et al., 2009; Fig. 4). The accumulation of ROPs at future RH formation sites was found to be altered in the procuste1 cellulose synthase mutant, in clasp and sabre mutants that affect MT organization, and in actin7 and actin interacting protein1-2 mutants, suggesting involvement of the cell wall MTs and F-actin in the regulation of the RH initiation sites (Singh et al., 2008; Pietra et al., 2013; Kiefer et al., 2015).

ROP2 is expressed in RHs, and its overexpression induces ectopic RH initiation, resulting in higher RH density; expression of the dominant-negative ROP2 mutant reduces RH density. Importantly, overexpression of the activated ROP2^CA also induces the formation of additional RHs, but the numbers were significantly lower than observed for wild-type ROP2 overexpressors (Jones et al., 2002). This suggests that ROP2 recycling between active and inactive states is essential for RH initiation. The rop2 null mutant develops shorter RHs, indicating that ROP2 is required for RH growth but that its function is redundant with other ROPs. Interestingly, ectopic expression of ROP6/7/11 did not increase RH initiation, suggesting that ROP2 and possibly ROP4 signaling in RH initiation is specific (Molendijk et al., 2001; Jones et al., 2002; Bloch et al., 2005). Interestingly, RHs developed in pluripetala (plp) mutant plants, which lack protein prenylation, suggesting that type II ROPs possibly also contribute to RH initiation (Running et al., 2004; Sorek et al., 2011; Chai et al., 2016). ROP2 activity also is regulated by Microtubule Associated Protein18, which interacts preferentially with GDP-bound ROP2 and competes with RhoGDI (Kang et al., 2017).

The expression of either type I or type II constitutively active ROP mutants results in RHs swelling, which is associated with disorganized actin filaments at the RH tips, inhibition of endocytosis, and nonpolar ROS distribution (Molendijk et al., 2001; Jones et al., 2002; Bloch et al., 2005; Sorek et al., 2010). ROPs activate RBOH-type NADPH oxidases during immune responses (Wong et al., 2007; Nagano et al., 2016). The SCN1/RhoGDI1-dependent and RHD2/AtRBOHC-dependent ROS accumulation at RH formation sites strongly suggest that ROP2/4 activates RBOHC during RH formation and growth (Carol et al., 2005). Consistent with the link between ROPs, ROS formation, and pH regulation, it was found that the ROP11^CA-induced RH swelling was suppressed by the omission of nitrogen-containing compounds (either NH₄⁺ or NO₃⁻) from the growth medium. The omission or addition of NH₄⁺ and NO₃⁻ affected intracellular and extracellular pH fluctuations and the overall pH at RH tips, implying a close correlation between ROP signaling and the pH at the RH tip (Bloch et al., 2011a, 2011b).

Upon activation, at least some ROPs undergo activation-dependent transient S-acylation on one or two G-domain Cys residues and, consequently, partition into detergent-resistant membranes, which is required for their function (Sorek et al., 2009, 2010, 2011, 2017). Consistently, ROP2 association with the plasma membrane is compromised in Protein S-Acyl Transferase4 (PAT4) mutants. pat4 RHs are shorter, and the multiple RH phenotype of the scn1/rhogdi1 mutant is suppressed in the pat4 background. Furthermore, ROPs activate rice RBOHB/H by recruiting them into lipid microdomains (Nagano et al., 2016). At the future RH initiation sites, ROP2/6 colocalize with PIP 4 phosphate 5 kinase 3, Dynamin Related Protein1A (DRP1A) and DRP2B, and the AGCVIII kinase D6 Protein Kinase (D6PK) in sterol-enriched domains (Stanislas et al., 2015). Auxin has been implicated in ROP activation (Tao et al., 2002; Wu et al., 2011; Miyawaki and Yang, 2014), yet the underlying mechanisms remain to be elucidated. PIN phosphorylation by D6PK regulates auxin transport (Barbosa et al., 2014; Zourelidou et al., 2014). Hence, the association of ROPs and D6PK in the RH initiation domain may provide a link between ROP signaling and auxin (Fig. 4; Box 2; Abo et al., 1991).

Signaling between cell wall extensibility and mechanisms that regulate cell growth is mediated in RHs by the RLK FER. FER is a member of the CrRLK1 (Catharanthus roseus RLK1) family that are thought to sense the cell through extracellular malectin-like domains that are presumed to interact with carbohydrates (Höfte, 2015; Li et al., 2016a; Voxeur and Höfte, 2016). RHs collapse or burst in fer mutant alleles. FER interacts with ROPGEF1 in yeast two-hybrid and bimolecular fluorescence complementation assays, and active ROP2 levels and ROS production are reduced in fer mutant plants (Duan et al., 2010). Hence, FER appears to regulate ROP function during the RH growth phase (Fig. 3).

To summarize, local auxin concentration gradients direct the formation of sterol-enriched ROP domains. The formation of single rather than multiple ROP domains depends of RhoGDI-dependent recycling of ROPs to and from the plasma membrane as well as cycling of ROPs between active and inactive states. During RH growth, ROP function is regulated by FER, which coordinates and adjusts cell wall composition in response to intracellular growth signals. Auxin signaling also is known to regulate RH growth (Knox et al., 2003), but it is unknown yet whether it signals through ROPs. Activated ROPs likely activate RBOHC, leading to ROS production and associated changes in intracellular and extracellular pH and intracellular Ca²⁺ levels (Foreman et al., 2003; Takeda et al., 2008). Through effectors, which have yet to be identified, ROPs regulate MTs and F-actin organization and dynamics (Fig. 3).

ROP Function during Diffuse Growth

With the exception of RHs and pollen tubes, plant cells grow by diffuse growth; this implies that their growth proceeds along the cell and is not limited to a single domain, as in tip growth. Three model systems have been used extensively to study ROP function during diffuse growth: trichomes, leaf epidermal pavement cells, and petal epidermal cells. Trichomes grow faster at the region closer to the tip, where the cell wall is thinner than at the basal side. Dense transverse MT arrays exist at the upper half but are absent from the tip and correspond to higher cell wall pressure and tension. Arp2/3-driven actin nucleation takes place at the MT clear zone at the tip and depends on an active WAVE complex (Yanagisawa et al., 2015). It is likely that ROPs are activated locally by SPK1 to promote WAVE activation. The expression of constitutively active rop2^CA resulted in the development of trichomes with thicker stalks and branches and altered branch positions, while the expression of dominant-negative ROP mutants did not alter trichome growth (Fu et al., 2002). However, expression of the Clostridium difficile toxin B catalytic domain, which glycosylates and inhibits ROP function, alters trichome morphogenesis (Singh et al., 2013). It is expected that genetic analysis of loss-of-function ROP mutants will reveal which ROPs are involved in trichome development.

Pavement cells are interdigitated, characterized by their lobes and indentations. Mutual stresses operating between neighboring cells affect the orientation of the MTs and, in turn, the patterning of cell growth (Sampathkumar et al., 2014). Compromised isotropic pavement cells with reduced interdigitation are observed in plants with gain- and loss-of-function mutants in genes encoding ROP, ROP effectors, SPK1, and PLP and in transgenic plants expressing anti-ROP toxins (Fu et al., 2002, 2005, 2009; Qiu et al., 2002; Bloch et al., 2005; Lavy et al., 2007; Sorek et al., 2010, 2011; Poraty-Gavra et al., 2013; Singh et al., 2013). Based on the analysis of Arabidopsis ROP2, ROP4, and ROP6 loss-of-function and constitutively active mutants, it was proposed that these proteins coregulate pavement cell growth by affecting the organization of both cortical MTs and F-actin via different ROP effectors. ROP6 interacts with RIC1, a ROP effector and MT-binding protein. MT-associated RIC1 interacts with the P60 subunit of the MT-severing protein katanin (KTN1). RIC1 promotes the MT-severing activity of KTN1 and the formation of the transverse cortical MTs (Fig. 5; Fu et al., 2005, 2009; Lin et al., 2013a). Overexpression of RIC1 induced highly ordered transverse MTs that result in isomorphic cell growth. ric1 loss-of-function pavement cells exhibited more randomly oriented cortical MTs and a wider neck/indentation region than the wild type (Fu et al., 2009). ROP2 and possibly ROP4 were shown to promote the formation of actin microfilaments via another ROP effector, RIC4. ROP2 also was shown to recruit RIC1 to the plasma membrane and away from MTs, thereby inhibiting its action on MTs (Fu et al., 2005). As ric1 loss-of-function pavement cells are interdigitated (Fu et al., 2009), RIC1 function is likely redundant during pavement cell growth. Treating cotyledons with cellulase results in the development of isomorphic noninterdigitated pavement cells. Interestingly, the cellulase-induced isomorphic pavement cell growth phenotype was suppressed in a ric1 loss-of-function background (Higaki et al., 2017), suggesting that RIC1 could be part of a pathway that senses cell wall integrity.

The physical and signaling interaction between FER and ROPGEF1 established a connection between cell wall sensing and ROP signaling. It was recently reported that ROPGEF4 interacts with the cell wall-sensing CrRLK THESEUS1 during the defense response to Botrytis cinerea (Qu et al., 2017a). It will be interesting to study the function of RIC1 in these signaling cascades. Further work will be required to elucidate how ROP signaling is integrated with cell wall pressure and strain as well as the signals that activate and inactivate ROPs during pavement cell growth.

The abaxial petal epidermal cells are interdigitated. The interdigitation of the cells is compromised in the spk1 mutant and in the rop2 rop6 rop4RNAi mutant, and cells display transverse MT arrays. Furthermore, the levels of active ROP2 and ROP6 are lower in spk1 than in wild-type plants (Ren et al., 2016). Taken together, these results indicated that SPK1 is responsible for activating ROP2, ROP6, and likely ROP4, which together regulate the growth of petal epidermal cells. The adaxial petal epidermal cells have cone-like structures with sharp-angled tips and transverse MT arrays. In ktn1 mutants, the cone-shaped cells have wider tip angles and less ordered MT arrays. Similarly, spk1 and rop2 rop6 rop4RNAi mutants develop cells with wider tip angles. It was proposed that ROP2/6 and ROP4 regulate MTs via KTN1 (Ren et al., 2017). These new findings also suggest that ROP signaling networks play important and general roles in the regulation of cell expansion and cell shape; however, the precise regulatory mechanisms might be distinct in specific organs.

ROP FUNCTION IN SECONDARY WALL PATTERNING, CYTOKINESIS, AND NUCLEI MIGRATION

Regulation of Secondary Wall Patterning by ROPs

During vascular differentiation, the ROP effector Microtubule Depletion Domain1 (MIDD1), also known as Interactor of Constitutively Active ROP5 (ICR5) or ROP Interacting Partner3, is a regulator of secondary cell wall deposition in metaxylem. MIDD1 is an MT-binding protein that interacts with the MT-destabilizing protein KINESINE13A (KIN13A; Mucha et al., 2010). During metaxylem differentiation, MIDD1 promotes the depolymerization of cortical MTs, leading to the formation of secondary wall pits (Oda et al., 2010). During metaxylem differentiation, coexpression of ROP11, ROPGEF4, and ROPGAP3 results in the formation of activated ROP11 domains. The active ROP11, in turn, recruits MIDD1, which, in turn, recruits KIN13A, leading to local MT breakdown (Oda and Fukuda, 2012). It was demonstrated recently that the size of the active ROP domains in the metaxlyem is limited by a plasma membrane-associated protein designated IQD13, which affects MT orientation and dynamics (Fig. 5; Sugiyama et al., 2017).

The Function of ROPs in the Regulation of Cytokinesis

Mutant and protein localization studies have implicated ROPs in the regulation of cytokinesis. In dividing BY2 cells, GFP-AtROP4 is associated with the cell plate (Molendijk et al., 2001). During stomata development in maize, partial knockout of type I ROPs (rop2^−/− rop9^+/−) or treatment with a RAC inhibitor results in weak cell division polarization defects. ROP2/9 accumulate at the junctions between subsidiary mother cells (SMCs) and guard mother cells (GMCs), and their accumulation at these sites depends on the membrane RLK PANGLOS1 (PAN1). When expressed at high levels, GFP-ZmROP2 is not polarly localized in SMCs, and cell division polarization defects are observed. Furthermore, ROP accumulation in detergent-resistant membranes is lower in the pan1 mutant than in wild-type plants, suggesting that PAN1 is required for ROP activation. The accumulation of ROPs and PAN at the SMC/GMC junctions precedes the formation of actin patches at these locations (Humphries et al., 2011).

The first component known to accumulate at the SMC/GMC junctions is the SCAR/WAVE complex. PAN1 and PAN2 polarization and accumulation at the SMC/GMC junction is compromised in a brk1 SCAR/WAVE subunit mutant. Based on these results and on the interaction between SCAR/WAVE subunits and ROPs and SPK1, it was proposed that the SCAR/WAVE complex recruits both PAN1 and ROP2/9 and that the resulting ROP activation leads to the activation of SCAR/WAVE and, in turn, Arp2/3 activation and formation of the actin patch (Facette et al., 2015). It is not yet known whether SPK1 is associated with the SCAR/WAVE complex in maize, whether the PAN1-dependent ROP activation is associated with the catalytic activity of PAN1 on ROPs or ROPGEFs, or whether it is required for the recruitment of ROPs to the SMC/GMC junctions through a protein-protein interaction.

A role for ROP signaling in the regulation of cytokinesis polarity in the Arabidopsis root was established recently (Stöckle et al., 2016). Two PH domain-containing ROPGAPs designated PHGAP1/2 interact the PHRAGMOPLAST ORIENTING KINESIN1 (POK1). phgap1 phgap2 double mutants display a weak cell wall-positioning phenotype, and during cytokinesis, PHGAP1 and PHGAP2 proteins localize to the CDZ in a POK1-dependent fashion (Stöckle et al., 2016). Taken together, these data suggest that the down-regulation of ROPs at specific sites at the cell division zone is required for proper orientation of the cell division planes.

ROP Signaling and Nuclear Migration: ROP8 and Nuclei Migration in the Ovule Central Cell

The central cell contains actin filaments that grow from the plasma membrane toward the nucleus. Upon entry of the sperm nucleus, it is engulfed by actin filaments that make an astral-like structure that pull it toward the central cell nucleus. AtROP8 is expressed uniquely in the central cell and is localized at the plasma membrane. Expression of rop8^DN disrupted the organization of the astral-like actin filaments and the migration of the sperm nuclei. Interestingly, the expression of rop8^CA did not affect actin organization or pollen nucleus migration, suggesting that ROP8 is maintained in an active state in the central cell (Kawashima et al., 2014). Nuclei migration is a vital process in the differentiation of different cell types in plants, and it would be of interest to examine whether other ROPs have a broad role in other cell types.

The Regulation of Nuclei Migration during RH Formation by ROP Signaling

Prior to RH emergence, trichoblast nuclei are localized close to the inner cell membrane at the center of the long cell axis. Consequently, the nuclei migrate into the growing RHs. It was reported recently that the positioning of the nuclei close to the inner membrane and their migration are regulated by ROP and auxin signaling that synergistically regulate actin organization and dynamics (Nakamura et al., 2018).

ROP FUNCTION IN BIOTIC AND ABIOTIC STRESS RESPONSES

Biotic Responses

The roles of ROP GTPases in plant pathogen and biotic responses have been the subject of recent detailed reviews (Bhagatji et al., 2010; Kawano et al., 2014; Rivero et al., 2017). This section will briefly highlight certain relevant aspects. Class VI Receptor-Like Cytoplasmic Kinases (RLCKs) belong to the large superfamily of RLKs, which are involved in a variety of cellular processes, including plant growth, development, and immune responses (Jurca et al., 2008; Lin et al., 2013b). RLCKs of the VI class appear to be downstream ROP effectors that regulate the pathogen response (Molendijk et al., 2008; Huesmann et al., 2012; Reiner et al., 2015). Barley (Hordeum vulgare) HvRBK1 is a type VI_A RLCK that is activated by HvRACB upon infection by the fungal pathogen Blumeria graminis f. sp. hordei (Bgh; Huesmann et al., 2012). Transient knockdown of HvRBK1 influences the stability of the cortical MTs in barley epidermal cells. HvRACB also regulates a RIC family member, RIC171, in the pathogen defense response. Overexpression of RIC171, similar to overexpression of constitutively activated RACB, renders epidermal cells more susceptible to penetration by Bgh. Additionally, RIC171 accumulates at sites of fungal attack, suggesting an enhanced ROP activity at sites of attempted fungal penetration (Schultheiss et al., 2008). HvRACB signaling is regulated by the barley MICROTUBULE-ASSOCIATED ROPGAP1 (HvMAGAP1), limiting the susceptibility of barley to Bgh (Hoefle et al., 2011).

In rice, OsRAC1 plays a critical role in defense responses induced by various microbe-associated molecular patterns, including chitin. OsRAC1 is activated by chitin perception mediated by OsCERK1, an RLK that forms a complex with OsCEBiP, a chitin-binding protein (Shimizu et al., 2010). In the presence of chitin, OsCERK1 phosphorylates and activates OsRACGEF. In turn, OsRAC1 activation activates lignin biosynthesis by activating Cinnamoyl CoA Reductase and ROS production by the activation of NADPH oxidase (Wong et al., 2007; Akamatsu et al., 2013; Nagano et al., 2016).

In Medicago trancatula and Lotus japonicus ROPs, MtROP10 and LjROP6 interact with the RLKs MtNFP and LjNFR5 in the membrane, respectively, and are required for rhizobium-induced nodule formation and for tip growth (Ke et al., 2012, 2016; Lei et al., 2015; Rivero et al., 2017). It is not known yet whether ROPGEFs colocalize with the respective ROPs and RLKs.

The expression of dominant-negative Atrop6^DN under the regulation of the ROP6 promoter induced a salicylic acid-dependent systemic acquired resistance (SAR) response characterized by constitutive expression of pathogenesis-related genes. The rop6^DN plants are small, have twisted leaves with rectangular pavement cells, and have altered actin and MT organization. rop6^DN plants with mutations in SA biosynthesis and signaling factors demonstrated that the developmental and SAR phenotypes are separable. Interestingly, regardless of the constitutive SAR response, the rop6^DN plants did not display enhanced resistance to Golovinomyces orontii powdery mildew (Poraty-Gavra et al., 2013). Possibly, the rop6^DN-induced changes in cytoskeleton organization and cell wall influence the SAR response. In the future, it will be important to carry out gene expression analyses because, in ROP mutants, as this analysis could potentially reveal, hidden phenotypes that affect the plant would otherwise remain undetectable.

ROPs and Abiotic Signaling Responses

Accumulating evidence indicates that ROPs and ABA form negative feedback loops in which ABA signaling suppresses ROP activation and ROP signaling suppresses ABA responses (Lemichez et al., 2001; Zheng et al., 2002; Xin et al., 2005; Li et al., 2012a, 2012b, 2016; Li and Liu, 2012; Yu et al., 2012). ROP10 and ROP11 inhibit ABA signaling by interacting physically with ABA negative regulators, ABA Insensitive1 (ABI1) and ABI2 PP2C phosphatases. It was postulated that the interactions with ROPs prevent ABI1/2 ABA-dependent inactivation by PYR/PIL ABA receptors (Li et al., 2012b; Yu et al., 2012). ABA inhibits ROP activation by promoting the degradation of several ROPGEFs. ROPGEF1 interacts with ABI1 in yeast and in vitro and is degraded in response to ABA. In the abi1 abi2 hab1 pp2^CA quadruple mutant, it is localized to prevacuolar compartments and undergoes degradation (Li et al., 2016b). Thus, ABI1 and possibly other phosphatases may form regulatory hubs that interact with both ROPGEFs and ROPs, forming a positive feedback loop that suppresses ABA responses. Upon an increase in ABA levels, PP2C is sequestered by PYR1/PYL ABA receptors, leading to ROPGEF degradation and ROP inactivation. Oxygen deprivation induces the expression of ROPGAP4, and ropgap4 mutant plants displayed lower oxygen deprivation tolerance due to ROS-induced overproduction of alcohol dehydrogenase (Baxter-Burrell et al., 2002). Hence, ROP activation/inactivation loops play an important function in abiotic stress tolerance and ABA signaling.

A recent report indicates that ROPGEF2 is activated directly or indirectly by phyB and, in turn, activates ROP2 and ROP7 to suppress red light-induced stomatal opening (Wang et al., 2017). Nucleotide exchange assays presented in this work suggest that ROPGEF2 catalyzes nucleotide activity on Cdc42. Yet, it is well documented that ROPGEFs are specific to plant ROPs and cannot catalyze GDP/GTP exchange on nonplant Rho GTPases (Berken et al., 2005; Thomas et al., 2007; Thomas and Berken, 2010). Hence, some of the data presented in this work might have to be revisited.

ROP signaling is involved in the regulation of abiotic stress via regulation of the cytoskeleton. The biphasic reorganization of MTs (rapid MT disassembly followed by reassembly of new MTs) induced by high salinity is required for plant tolerance to salt stress. The ROP2-RIC1 signaling pathway was found to modulate MT reorganization in response to salt stress in Arabidopsis (Li et al., 2017). During salt stress, activation of ROP2 leads to the removal of RIC1 from MTs, leading to enhanced MT reassembly and, in turn, an increase in plant tolerance to salt stress (Li et al., 2017).