WASH has a critical role in NK cell cytotoxicity through Lck-mediated phosphorylation

Natural killer (NK) cells are important effector cells of the innate immune system to kill certain virus-infected and transformed cells. Wiskott–Aldrich Syndrome protein (WASP) and SCAR homolog (WASH) has been identified as a member of WASP family proteins implicated in regulating the cytoskeletal reorganization, yet little is known about its function in lymphocytes. Here we demonstrate that WASH is crucial for NK cell cytotoxicity. WASH was found to colocalize with lytic granules upon NK cell activation. Knockdown of WASH expression substantially inhibited polarization and release of lytic granules to the immune synapse, resulting in the impairment of NK cell cytotoxicity. More importantly, our data also define a previously unappreciated mechanism for WASH function, in which Src family kinase Lck can interact with WASH and induce WASH phosphorylation. Mutation of tyrosine residue Y141, identified here as the major site of WASH phosphorylation, partially blocked WASH tyrosine phosphorylation and NK cell cytotoxicity. Taken together, these observations suggest that WASH has a pivotal role for regulation of NK cell cytotoxicity through Lck-mediated Y141 tyrosine phosphorylation.Natural killer (NK) cells are the first defense line against viral infections and tumors.¹ NK cell-mediated lysis of target cells requires the formation of immunological synapse between NK cells and target cells and subsequent delivery of lytic granules containing perforin and granzymes.^{2, 3} The importance of the actin cytoskeleton in this process has been well documented.⁴ However, the precise mechanism of actin reorganization in NK cells remains to be elucidated.Wiskott–Aldrich syndrome protein (WASP) is the first identified member of an actin regulator family.⁵ WASP family proteins contain a C-terminal domain that binds to and activates the Arp2/3 complex for cytoskeleton remodeling.⁶ In the absence of WASP, cytotoxic activity of NK cells is defective owing to impaired immune synapse formation and perforin localization.⁷ It has also been shown that WASP may be important for integration of NK cell signaling, particularly for nuclear translocation of NFAT2 and NF-κB during the activating receptor NKp46-dependent activation.⁸WASP and SCAR homolog (WASH) has been discovered as a new WASP family member.⁹ Subsequent studies show that WASH interacts with multiple proteins, including FAM21, to form a large core complex and regulate actin dynamics.¹⁰ WASH localizes to sorting and recycling endosomes, where WASH complex activates Arp2/3-mediated actin polymerization and controls the production of transport intermediates from endosome.¹¹ Unlike other WASP family members, WASH has distinct N-terminal domains, termed WASH homology domain 1 (WHD1) and tubulin-binding region (TBR).¹² Moreover, WASH has been shown to regulate recycling of many surface receptors via endosomal trafficking in activated T cells.¹³In our previous works, we found embryonic lethality and extensive autophagy in WASH-deficient mice. WASH recruits RNF2 to ubiquitinate AMBRA1 and inhibits the ubiquitination of Beclin1, a well-known moderator in autophage.^{14, 15} Of interest, WASH is located in cell nucleus and participates in hematopoietic stem cell differentiation through recruiting NURF complex to c-Myc promoter.¹⁶ However, the role and mechanism of WASH in NK cell function remain poorly understood.In this study, we show that inhibition of WASH expression with RNA interference or an inducible gene targeting system severely impair NK cell cytotoxicity through blockade of lytic granule polarization. In addition, Src family kinase Lck can interact with and induce tyrosine phosphorylation of WASH protein in human NK cells. These analyses provide the cellular and molecular mechanisms involved in the regulation of WASH function during NK cell activation.     

The Association of the Arabidopsis Actin-Related Protein2/3 Complex with Cell Membranes Is Linked to Its Assembly Status But Not Its Activation

In growing plant cells, the combined activities of the cytoskeleton, endomembrane, and cell wall biosynthetic systems organize the cytoplasm and define the architecture and growth properties of the cell. These biosynthetic machineries efficiently synthesize, deliver, and recycle the raw materials that support cell expansion. The precise roles of the actin cytoskeleton in these processes are unclear. Certainly, bundles of actin filaments position organelles and are a substrate for long-distance intracellular transport, but the functional linkages between dynamic actin filament arrays and the cell growth machinery are poorly understood. The Arabidopsis (Arabidopsis thaliana) “distorted group” mutants have defined protein complexes that appear to generate and convert small GTPase signals into an Actin-Related Protein2/3 (ARP2/3)-dependent actin filament nucleation response. However, direct biochemical knowledge about Arabidopsis ARP2/3 and its cellular distribution is lacking. In this paper, we provide biochemical evidence for a plant ARP2/3. The plant complex utilizes a conserved assembly mechanism. ARPC4 is the most critical core subunit that controls the assembly and steady-state levels of the complex. ARP2/3 in other systems is believed to be mostly a soluble complex that is locally recruited and activated. Unexpectedly, we find that Arabidopsis ARP2/3 interacts strongly with cell membranes. Membrane binding is linked to complex assembly status and not to the extent to which it is activated. Mutant analyses implicate ARP2 as an important subunit for membrane association.In plant cells, the actin cytoskeleton forms an intricate network of polymers that organizes the cytoplasm and defines the long-distance intracellular trafficking patterns of the cell. The actin network is critical not only for tip-growing cells (for review, see Cole and Fowler, 2006; Lovy-Wheeler et al., 2007) but also during the coordinated cell expansion that occurs in cells that utilize a diffuse growth mechanism (for review, see Wasteneys and Galway, 2003; Smith and Oppenheimer, 2005). For example, the polarized diffuse growth of leaf trichomes is highly sensitized to actin cytoskeleton disruption (Mathur et al., 1999; Szymanski et al., 1999), and a recent analysis of Arabidopsis (Arabidopsis thaliana) ACTIN mutants revealed widespread cell swelling and isotropic expansion in numerous cell types in the root and shoot (Kandasamy et al., 2009). The actin network is dynamic. The array reorganizes during cell morphogenesis (Braun et al., 1999; Szymanski et al., 1999) and in response to endogenous (Lemichez et al., 2001) and external (Hardham et al., 2007) cues. A major research goal is to better understand not only how plant cells convert G-actin subunits to particular actin filament arrays but also how the actin network interacts with the cellular growth machinery during cell expansion.This is a difficult problem to solve, because in expanding vacuolated cells the actin array adopts numerous configurations and consists of dense meshworks of cortical actin filaments and bundles (Baluska et al., 2000), thick actin bundles that penetrate the central vacuole (Higaki et al., 2006), and meshworks of filaments and bundles that surround the nucleus and chloroplasts (Kandasamy and Meagher, 1999; Collings et al., 2000). The spatial relationships between these actin networks and localized cell expansion are not obvious. Certainly, the plasma membrane-cell wall interface is a critical location for the regulated delivery and fusion of vesicles containing cell wall polysaccharides. Frequent reports of localized domains of enriched cortical actin signal at regions of presumed localized cell expansion have led to the widely held view that the cortical actin array creates local tracks for vesicle-mediated secretion (for review, see Smith and Oppenheimer, 2005; Hussey et al., 2006). In one study, the dynamics of actin filaments were analyzed in living hypocotyl epidermal cells that utilize a diffuse growth mechanism (Staiger et al., 2009). In this case, individual actin filaments are very unstable and randomly oriented; therefore, the precise relationships between cortical F-actin, vesicle delivery, and cell shape change remain obscure. The best known function for the actin cytoskeleton is that of a track for myosin-dependent vesicle and organelle trafficking (Shimmen, 2007). The actin bundle network mediates the transport of cargo between endomembrane compartments (Geldner et al., 2001; Kim et al., 2005) and the long-distance actomyosin transport of a variety of organelles, including the Golgi (Nebenfuhr et al., 1999; Peremyslov et al., 2008; Prokhnevsky et al., 2008). Generation of distributed (Gutierrez et al., 2009; Timmers et al., 2009) and localized (Wightman and Turner, 2008) actin bundle networks appears to define early steps in the trafficking of Golgi-localized cellulose synthase complexes to the sites of primary and secondary wall synthesis, respectively.Plant cells employ diverse collections of G-actin-binding proteins, actin filament nucleators, and actin-bundling and cross-linking proteins to generate and remodel the F-actin network (for review, see Staiger and Blanchoin, 2006). One actin filament nucleator, termed the Actin-Related Protein2/3 (ARP2/3) complex, controls numerous aspects of plant morphogenesis and development. The vertebrate complex consists of the actin-related proteins ARP2 and ARP3 and five other unrelated proteins termed ARPC1 to ARPC5, in order of decreasing mass. ARP2/3 in isolation is inactive, but in the presence of proteins termed nucleation-promoting factors such as WAVE/SCAR (for WASP family Verprolin homologous/Suppressor of cAMP Repressor), ARP2/3 is converted into an efficient actin filament-nucleating machine (for review, see Higgs and Pollard, 2001; Welch and Mullins, 2002). In mammalian cells, ARP2/3 activities are linked to membrane dynamics. Keratocytes that crawl persistently on a solid substrate appear to use ARP2/3-generated dendritic actin filament networks at the leading edge to either drive or consolidate plasma membrane protrusion (Pollard and Borisy, 2003; Ji et al., 2008). In many vertebrate cell types, ARP2/3 has a strong punctate intracellular localization (Welch et al., 1997; Strasser et al., 2004), which could reflect hypothesized activities at the Golgi (Stamnes, 2002) or late endosomal (Fucini et al., 2002; Holtta-Vuori et al., 2005) compartment.Genetic studies in plants reveal nonessential but widespread functions for ARP2/3. In the moss Physcomitrella patens, the ARPC4 and ARPC1 subunit genes are critical during tip growth of protonemal filaments (Harries et al., 2005; Perroud and Quatrano, 2006). In Arabidopsis, loss of either ARP2/3 subunit gene or mutations in WAVE complex genes that positively regulate ARP2/3 cause complicated syndromes, including the loss of polarized diffuse growth throughout the shoot epidermis, defective cell-cell adhesion, and decreased hypocotyl elongation (for review, see Szymanski, 2005). Altered responses to exogenous Suc (Li et al., 2004; Zhang et al., 2008) and reduced root elongation (Dyachok et al., 2008) are also reported for wave and arp2/3 strains. In higher plants, the involvement of ARP2/3 in tip growth and root hair development is more subtle. In Lotus japonicus, mutation of NAP1 and PIR1, known positive regulators of ARP2/3 (Basu et al., 2004; Deeks et al., 2004; El-Assal et al., 2004a), causes incompletely penetrant root hair phenotypes, but in the presence of symbiotic bacteria, the mutants have defective infection threads and reduced root nodule formation. Arabidopsis arp2/3 mutants do not have obvious tip growth defects in pollen tubes or root hairs, but in the presence of GFP:TALIN (Mathur et al., 2003b) and in double mutant combinations with the actin-binding protein CAP1 (Deeks et al., 2007), the effects of ARP2/3 on root hair growth are unmasked.In Arabidopsis, the genetics of the positive regulation of ARP2/3 are well characterized and appear to occur solely through another heteromeric complex termed WAVE (Eden et al., 2002; for review, see Szymanski, 2005). The putative WAVE/SCAR complex contains five subunits, one of which is the ARP2/3 activator SCAR. Plant SCARs contain conserved N-terminal and C-terminal domains that mediate interactions with other WAVE complex proteins and ARP2/3 activation, respectively (Frank et al., 2004; Basu et al., 2005). In nonplant systems, the regulatory relationships between WAVE and ARP2/3 appear to vary between cell types and species (for review, see Bompard and Caron, 2004; Stradal and Scita, 2006). However, in Arabidopsis, double mutant analyses indicate that WAVE is the sole pathway for ARP2/3 activation and that all subunits positively regulate ARP2/3 (Deeks et al., 2004; Basu et al., 2005; Djakovic et al., 2006). SCAR quadruple mutants are indistinguishable from arp2/3 null plants (Zhang et al., 2008). In moss, BRICK1 and ARP2/3 mutants have similar phenotypes, suggesting conserved regulatory relationships between WAVE and ARP2/3 in the plant kingdom (Harries et al., 2005; Perroud and Quatrano, 2006, 2008).Despite extensive molecular genetic knowledge about the ARP2/3 pathway and the strong actin cytoskeleton and growth phenotypes of arp2/3 plants, there are few direct data on the existence of the plant complex and its cellular function. There are reports of ARP2/3 localization based on the behavior of individual subunits (Le et al., 2003). In some cases, the results are weakened by the unknown specificity of heterologous ARP2/3 antibodies (Van Gestel et al., 2003; Fiserova et al., 2006). A specific antibody was raised against Silvetia ARP2 (Hable and Kropf, 2005). In developing zygotes, rhizhoid emergence is an early and actin-dependent developmental event, and at this stage a broad subcortical cone of ARP2 signal extends from the nuclear envelope toward the rhizhoid apex (Hable and Kropf, 2005). Double labeling experiments detected considerable overlap between ARP2 and actin, but surprisingly, there was a broad cortical domain of putative organelle-associated distal ARP2 that did not overlap with actin. In tip-growing P. patens chloronema cells, ARPC4 also appears to be membrane associated and localizes to a broad subcortical apical zone (Perroud and Quatrano, 2006). For these localization and genetic studies that rely on individual ARP2/3 subunits, it is important to prove that a plant ARP2/3 complex exists to test for an association of the complex with endomembrane compartments.In this paper, we provide several lines of evidence for an evolutionarily conserved pathway for ARP2/3 complex assembly in plant cells. These studies are based in part on genetic and biochemical analyses of the putative ARP2/3 subunit gene ARPC4. We found that disruption of the ARPC4 gene caused catastrophic disassembly of the complex and an array of phenotypes that were indistinguishable from known arp2/3 mutants. Chromatography experiments clearly revealed that functional hemagglutinin (HA)-tagged ARPC4 and endogenous ARP3 subunits assemble fully into ARP2/3 complexes. Surprisingly, much of the cellular pool of the plant ARP2/3 complex is membrane associated. An analysis of an extensive collection of wave and arp2/3 mutants allowed us to conclude that the normal association with membranes depended on the presence of ARP2 and the assembly status of the complex but not on the existence of an active pool of ARP2/3 in the cell.     

Actin-Binding Proteins Implicated in the Formation of the Punctate Actin Foci Stimulated by the Self-Incompatibility Response in Papaver

The actin cytoskeleton is a key target for signaling networks and plays a central role in translating signals into cellular responses in eukaryotic cells. Self-incompatibility (SI) is an important mechanism responsible for preventing self-fertilization. The SI system of Papaver rhoeas pollen involves a Ca²⁺-dependent signaling network, including massive actin depolymerization as one of the earliest cellular responses, followed by the formation of large actin foci. However, no analysis of these structures, which appear to be aggregates of filamentous (F-)actin based on phalloidin staining, has been carried out to date. Here, we characterize and quantify the formation of F-actin foci in incompatible Papaver pollen tubes over time. The F-actin foci increase in size over time, and we provide evidence that their formation requires actin polymerization. Once formed, these SI-induced structures are unusually stable, being resistant to treatments with latrunculin B. Furthermore, their formation is associated with changes in the intracellular localization of two actin-binding proteins, cyclase-associated protein and actin-depolymerizing factor. Two other regulators of actin dynamics, profilin and fimbrin, do not associate with the F-actin foci. This study provides, to our knowledge, the first insights into the actin-binding proteins and mechanisms involved in the formation of these intriguing structures, which appear to be actively formed during the SI response.The ability to perceive and integrate signals into networks is essential for all eukaryotic cells. The actin cytoskeleton is a major target and integrator of signaling networks in eukaryotic cells. In plants, many extracellular stimuli lead to rapid structural changes in the actin cytoskeleton (Staiger, 2000; Hussey et al., 2006). Although many of the signaling intermediates that regulate actin dynamics are well defined in animal cells and yeast (Iden and Collard, 2008; Thomas et al., 2009), considerably less is known for plants. However, it is generally accepted that actin-binding proteins (ABPs) function as transducers of cellular stimuli into changes in cellular architecture (Hussey et al., 2006; Staiger and Blanchoin, 2006; Thomas et al., 2009). This includes three abundant monomer-binding proteins, profilin, actin-depolymerizing factor (ADF), and cyclase-associated protein (CAP), which function synergistically to stimulate actin turnover in vitro (Chaudhry et al., 2007). Bundling and cross-linking proteins, such as fimbrin, function to stabilize actin filaments into higher order structures (Kovar et al., 2000b; Thomas et al., 2009). These and other regulators of actin turnover are likely targets for signal-mediated changes in actin architecture in response to biotic and abiotic stresses.Self-incompatibility (SI) is a genetically controlled system to prevent self-fertilization in flowering plants. SI is controlled by a multiallelic S-locus; S-specific pollen rejection results from the interaction of pollen S- and pistil S-determinants that have matching alleles (Franklin-Tong, 2008). In Papaver rhoeas, the pistil S-determinants (previously called S proteins, recently renamed PrsS; Foote et al., 1994; Wheeler et al., 2009) act as ligands, interacting with the pollen S-determinant PrpS (Wheeler et al., 2009), triggering increases in calcium influx and increases in cytosol-free calcium in incompatible pollen (Franklin-Tong et al., 1993, 1997, 2002). The Ca²⁺-mediated signaling network results in rapid inhibition of incompatible pollen tube growth and triggers programmed cell death (PCD) involving several caspase-like activities (Thomas and Franklin-Tong, 2004; Bosch and Franklin-Tong, 2007).The SI response and the Ca²⁺-signaling pathway in Papaver stimulate rapid reorganization and massive depolymerization of actin filaments in incompatible pollen tubes (Geitmann et al., 2000; Snowman et al., 2002). Moreover, it has been demonstrated that changes in actin dynamics are necessary and sufficient for PCD initiation (Thomas et al., 2006). Intriguingly, there seems to be cross talk between actin and microtubule cytoskeletons in mediating PCD in pollen (Poulter et al., 2008). Thus, there is compelling evidence for signaling to the actin cytoskeleton in mediating PCD during SI (for a recent review, see Bosch et al., 2008). SI also triggers further changes to the actin cytoskeleton. Small F-actin foci are formed, and these increase in size within the first hour after SI stimulus and remain observable for at least 3 h (Geitmann et al., 2000; Snowman et al., 2002). These aggregates contain F-actin, as they stain with rhodamine phalloidin. The formation of the small actin foci and the larger F-actin structures occurs after cessation of pollen tube growth, so they are unlikely to play a role in pollen inhibition.Punctate F-actin foci are unusual structures, and there appears to be a paucity of examples of their formation in any eukaryotic cell type. Actin patches are associated with endocytosis in normally growing yeast (Pelham and Chang, 2001; Kaksonen et al., 2003; Ayscough, 2004; Young et al., 2004), and “actin nodules” are formed during filipodia formation in platelets (Calaminus et al., 2008). Actin bodies are formed when yeast cells enter the quiescent cycle (Sagot et al., 2006), and Hirano bodies are observed in animal cells and Dictyostelium undergoing stress or in the disease state (Hirano, 1994; Maselli et al., 2002). Large star-shaped actin arrays have been observed in pollen tubes growing in vivo (Lord, 1992), but their nature and function are unknown. When we first described the SI-induced structures, we avoided the terminology of patches or bodies, as it was not known whether they were either structurally or functionally comparable to any of these previously characterized actin-based structures.Studies on the SI-mediated actin responses to date have focused on the initial phase of depolymerization, and no analysis of what is involved in the formation of the large punctate actin foci has been made. Here, we show that the formation of punctate actin foci requires actin polymerization, but once formed they are unusually stable. Moreover, we find that their formation correlates with changes in the intracellular localization of two ABPs, CAP and ADF, but not with two other key regulators of actin dynamics, profilin and fimbrin.     

Heterodimeric Capping Protein from Arabidopsis Is a Membrane-Associated,Actin-Binding Protein

The actin cytoskeleton is a major regulator of cell morphogenesis and responses to biotic and abiotic stimuli. The organization and activities of the cytoskeleton are choreographed by hundreds of accessory proteins. Many actin-binding proteins are thought to be stimulus-response regulators that bind to signaling phospholipids and change their activity upon lipid binding. Whether these proteins associate with and/or are regulated by signaling lipids in plant cells remains poorly understood. Heterodimeric capping protein (CP) is a conserved and ubiquitous regulator of actin dynamics. It binds to the barbed end of filaments with high affinity and modulates filament assembly and disassembly reactions in vitro. Direct interaction of CP with phospholipids, including phosphatidic acid, results in uncapping of filament ends in vitro. Live-cell imaging and reverse-genetic analyses of cp mutants in Arabidopsis (Arabidopsis thaliana) recently provided compelling support for a model in which CP activity is negatively regulated by phosphatidic acid in vivo. Here, we used complementary biochemical, subcellular fractionation, and immunofluorescence microscopy approaches to elucidate CP-membrane association. We found that CP is moderately abundant in Arabidopsis tissues and present in a microsomal membrane fraction. Sucrose density gradient separation and immunoblotting with known compartment markers were used to demonstrate that CP is enriched on membrane-bound organelles such as the endoplasmic reticulum and Golgi. This association could facilitate cross talk between the actin cytoskeleton and a wide spectrum of essential cellular functions such as organelle motility and signal transduction.The cellular levels of membrane-associated lipids undergo dynamic changes in response to developmental and environmental stimuli. Different species of phospholipids target specific proteins and this often affects the activity and/or subcellular localization of these lipid-binding proteins. One such membrane lipid, phosphatidic acid (PA), serves as a second messenger and regulates multiple developmental processes in plants, including seedling development, root hair growth and pattern formation, pollen tube growth, leaf senescence, and fruit ripening. PA levels also change during various stress responses, including high salinity and dehydration, pathogen attack, and cold tolerance (Testerink and Munnik, 2005, 2011; Wang, 2005; Li et al., 2009). In mammalian cells, PA is critical for vesicle trafficking events, such as vesicle budding from the Golgi apparatus, vesicle transport, exocytosis, endocytosis, and vesicle fusion (Liscovitch et al., 2000; Freyberg et al., 2003; Jenkins and Frohman, 2005).The actin cytoskeleton and a plethora of actin-binding proteins (ABPs) are well-known targets and transducers of lipid signaling (Drøbak et al., 2004; Saarikangas et al., 2010; Pleskot et al., 2013). For example, several ABPs have the ability to bind phosphoinositide lipids, such as phosphatidylinositol 4,5-bisphosphate [PtdIns(4,5)P₂]. The severing or actin filament depolymerizing proteins such as villin, cofilin, and profilin are inhibited when bound to PtdIns(4,5)P₂. One ABP appears to be strongly regulated by another phospholipid; human gelsolin binds to lysophosphatidic acid and its filament severing and barbed-end capping activities are inhibited by this biologically active lipid (Meerschaert et al., 1998). Gelsolin is not, however, regulated by PA (Meerschaert et al., 1998), nor are profilin (Lassing and Lindberg, 1985), α-actinin (Fraley et al., 2003), or chicken CapZ (Schafer et al., 1996).The heterodimeric capping protein (CP) from Arabidopsis (Arabidopsis thaliana) also binds to and its activity is inhibited by phospholipids, including both PtdIns(4,5)P₂ and PA (Huang et al., 2003, 2006). PA and phospholipase D activity have been implicated in the actin-dependent tip growth of root hairs and pollen tubes (Ohashi et al., 2003; Potocký et al., 2003; Samaj et al., 2004; Monteiro et al., 2005a; Pleskot et al., 2010). Exogenous application of PA causes an elevation of actin filament levels in suspension cells, pollen, and Arabidopsis epidermal cells (Lee et al., 2003; Potocký et al., 2003; Huang et al., 2006; Li et al., 2012; Pleskot et al., 2013). Capping protein (CP) binds to the barbed end of actin filaments with high (nanomolar) affinity, dissociates quite slowly, and prevents the addition of actin subunits at this end (Huang et al., 2003, 2006; Kim et al., 2007). In the presence of phospholipids, AtCP is not able to bind to the barbed end of actin filaments (Huang et al., 2003, 2006). Furthermore, capped filament ends are uncapped by the addition of PA, allowing actin assembly from a pool of profilin-actin (Huang et al., 2006). Collectively, these data lead to a simple model whereby CP, working in concert with profilin-actin, serves to maintain tight regulation of actin assembly at filament barbed ends (Huang et al., 2006; Blanchoin et al., 2010; Henty-Ridilla et al., 2013; Pleskot et al., 2013). Furthermore, the availability of CP for filament ends can be modulated by fluxes in signaling lipids. Genetic evidence for this model was recently obtained by analyzing the dynamic behavior of actin filament ends in living Arabidopsis epidermal cells after treatment with exogenous PA (Li et al., 2012). Specifically, changes in the architecture of cortical actin arrays and dynamics of individual actin filaments that are induced by PA treatment were found to be attenuated in cp mutant cells (Li et al., 2012; Pleskot et al., 2013).Structural characterization of chicken CapZ demonstrates that the α- and β-subunits of the heterodimer form a compact structure resembling a mushroom with pseudo-two-fold rotational symmetry (Yamashita et al., 2003). Actin- and phospholipid-binding sites are conserved on the C-terminal regions, sometimes referred to as tentacles, which comprise amphipathic α-helices (Cooper and Sept, 2008; Pleskot et al., 2012). Coarse-grained molecular dynamics (CG-MD) simulations recently revealed the mechanism of chicken and AtCP association with membranes (Pleskot et al., 2012). AtCP interacts specifically with lipid bilayers through interactions between PA and the amphipathic helix of the α-subunit tentacle. Extensive polar contacts between lipid headgroups and basic residues on CP (including K278, which is unique to plant CP), as well as partial embedding of nonpolar groups into the lipid bilayer, are observed (Pleskot et al., 2012). Moreover, a glutathione S-transferase fusion protein containing the C-terminal 38 amino acids from capping protein α subunit (CPA) is sufficient to bind PA-containing liposomes in vitro (Pleskot et al., 2012). Collectively, these findings lead us to predict that AtCP will behave like a membrane-associated protein in plant cells.Additional evidence from animal and microbial cells supports the association of CP with biological membranes. In Acanthamoeba castellanii, CP is localized primarily to the hyaline ectoplasm in a region of the cytoplasm just under the plasma membrane that contains a high concentration of actin filaments (Cooper et al., 1984). Localization of CP with regions rich in actin filaments and with membranes was supported by subcellular fractionation experiments, in which CP was associated with a crude membrane fraction that included plasma membrane (Cooper et al., 1984). Further evidence demonstrates that CP localizes to cortical actin patches at sites of new cell wall growth in budding yeast (Saccharomyces cerevisiae), including the site of bud emergence. By contrast, CP did not colocalize with actin cables in S. cerevisiae (Amatruda and Cooper, 1992). CP may localize to these sites by direct interactions with membrane lipids, through binding the ends of actin filaments, or by association with another protein different from actin. In support of this hypothesis, GFP-CP fusion proteins demonstrate that sites of actin assembling in living cells contain both CP and the actin-related protein2/3 (Arp2/3) complex, and CP is located in two types of structures: (1) motile regions of the cell periphery, which reflect movement of the edge of the lamella during extension and ruffling; and (2) dynamic spots within the lamella (Schafer et al., 1998). CP has been colocalized to the F-actin patches in fission yeast (Schizosaccharomyces pombe; Kovar et al., 2005), which promotes Arp2/3-dependent nucleation and branching and limits the extent of filament elongation (Akin and Mullins, 2008). These findings lend additional support for a model whereby CP cooperates with the Arp2/3 complex to regulate actin dynamics (Nakano and Mabuchi, 2006). Activities and localization of other plant ABPs are linked to membranes. Membrane association has been linked to the assembly status of the ARP2/3 complex, an actin filament nucleator, in Arabidopsis (Kotchoni et al., 2009). SPIKE1 (SPK1), a Rho of plants (Rop)-guanine nucleotide exchange factor (GEF) and peripheral membrane protein, maintains the homeostasis of the early secretory pathway and signal integration during morphogenesis through specialized domains in the endoplasmic reticulum (ER; Zhang et al., 2010). Furthermore, Nck-associated protein1 (NAP1), a component of the suppressor of cAMP receptor/WASP-family verprolin homology protein (SCAR/WAVE) complex, strongly associates with membranes and is particularly enriched in ER membranes (Zhang et al., 2013a). Finally, a superfamily of plant ABPs, called NETWORKED proteins, was recently discovered; these link the actin cytoskeleton to various cellular membranes (Deeks et al., 2012; Hawkins et al., 2014; Wang et al., 2014).In this work, we demonstrate that CP is a membrane-associated protein in Arabidopsis. To our knowledge, this is the first direct evidence for CP-membrane association in plants. This interaction likely targets CP to cellular compartments such as the ER and Golgi. This unique location may allow CP to remodel the actin cytoskeleton in the vicinity of endomembrane compartments and/or to respond rapidly to fluxes in signaling lipids.     

The Apical Actin Fringe Contributes to Localized Cell Wall Deposition and Polarized Growth in the Lily Pollen Tube

In lily (Lilium formosanum) pollen tubes, pectin, a major component of the cell wall, is delivered through regulated exocytosis. The targeted transport and secretion of the pectin-containing vesicles may be controlled by the cortical actin fringe at the pollen tube apex. Here, we address the role of the actin fringe using three different inhibitors of growth: brefeldin A, latrunculin B, and potassium cyanide. Brefeldin A blocks membrane trafficking and inhibits exocytosis in pollen tubes; it also leads to the degradation of the actin fringe and the formation of an aggregate of filamentous actin at the base of the clear zone. Latrunculin B, which depolymerizes filamentous actin, markedly slows growth but allows focused pectin deposition to continue. Of note, the locus of deposition shifts frequently and correlates with changes in the direction of growth. Finally, potassium cyanide, an electron transport chain inhibitor, briefly stops growth while causing the actin fringe to completely disappear. Pectin deposition continues but lacks focus, instead being delivered in a wide arc across the pollen tube tip. These data support a model in which the actin fringe contributes to the focused secretion of pectin to the apical cell wall and, thus, to the polarized growth of the pollen tube.Pollen tubes provide an excellent model for studying the molecular and physiological processes that lead to polarized cell growth. Because all plant cell growth results from the regulated yielding of the cell wall in response to uniform turgor pressure (Winship et al., 2010; Rojas et al., 2011), the cell wall of the pollen tube must yield only at a particular spot: the cell apex, or tip. To accomplish the extraordinary growth rates seen in many species, and to balance the thinning of the apical wall due to rapid expansion, the pollen tube delivers prodigious amounts of wall material, largely methoxylated pectins, to the tip in a coordinated manner. Recent studies suggest that the targeted exocytosis increases the extensibility of the cell wall matrix at the tip, which then yields to the existing turgor pressure, permitting the tip to extend or grow (McKenna et al., 2009; Hepler et al., 2013). There are many factors that influence exocytosis in growing pollen tubes; in this study, we investigate the role of the apical actin fringe.For many years, it has been known that an actin structure exists near the pollen tube tip, yet its exact form has been a matter of some contention (Kost et al., 1998; Lovy-Wheeler et al., 2005; Wilsen et al., 2006; Cheung et al., 2008; Vidali et al., 2009; Qu et al., 2013). The apical actin structure has been variously described as a fringe, a basket, a collar, or a mesh. Using rapid freeze fixation of lily (Lilium formosanum) pollen tubes followed by staining with anti-actin antibodies, the structure appears as a dense fringe of longitudinally oriented microfilaments, beginning 1 to 5 µm behind the apex and extending 5 to 10 µm basally. The actin filaments are positioned in the cortical cytoplasm close to the plasma membrane (Lovy-Wheeler et al., 2005). More recently, we used Lifeact-mEGFP, a probe that consistently labels this palisade of longitudinally oriented microfilaments in living cells (Vidali et al., 2009; Fig. 1A, left column). For the purposes of this study, we will refer to this apical organization of actin as a fringe.Open in a separate windowFigure 1.The actin fringe and the thickened pollen tube tip wall are stable, although dynamic, structures during pollen tube growth. A, The left column shows a pollen tube transformed with Lifeact-mEGFP imaged with a spinning-disc confocal microscope. Maximal projections from every 15 s are shown. The right column shows epifluorescence images of a pollen tube stained with PI. Again, images captured every 15 s are shown. Bars = 10 μm. B, The data from the pollen tube in A expressing Lifeact-mEGFP were subjected to kymograph analysis using an 11-pixel strip along the image’s midline. C, The first three frames from the pollen tube in A and B were assigned the colors red, blue, and green, respectively, and then overlaid. Areas with white show the overlap of all three. The fringe is stable, but most of its constituent actin is not shared between frames.Many lines of evidence demonstrate that actin is required for pollen tube growth. Latrunculin B (LatB), which blocks actin polymerization, inhibits pollen tube growth and disrupts the cortical fringe at concentrations as low as 2 nm. Higher concentrations are needed to block pollen grain germination and cytoplasmic streaming (Gibbon et al., 1999; Vidali et al., 2001). Actin-binding proteins, including actin depolymerizing factor-cofilin, formin, profilin, and villin, and signaling proteins, such as Rho-of-Plants (ROP) GTPases and their effectors (ROP interacting crib-containing proteins [RICs]), also have been shown to play critical roles in growth and actin dynamics (Fu et al., 2001; Vidali et al., 2001; Allwood et al., 2002; Chen et al., 2002; Cheung and Wu, 2004; McKenna et al., 2004; Gu et al., 2005; Ye et al., 2009; Cheung et al., 2010; Staiger et al., 2010; Zhang et al., 2010a; Qu et al., 2013; van Gisbergen and Bezanilla, 2013).Our understanding of the process of exocytosis and pollen tube elongation has been influenced by ultrastructural images of pollen tube tips, which reveal an apical zone dense with vesicles (Cresti et al., 1987; Heslop-Harrison, 1987; Lancelle et al., 1987; Steer and Steer, 1989; Lancelle and Hepler, 1992; Derksen et al., 1995). It has long been assumed that these represent exocytotic vesicles destined to deliver new cell wall material. This model of polarized secretion has been challenged in recent years in studies using FM dyes. Two groups have suggested that exocytosis occurs in a circumpolar annular zone (Bove et al., 2008; Zonia and Munnik, 2008). However, other studies, using fluorescent beads attached to the cell surface, indicate that the maximal rate of expansion, and of necessity the greatest deposition of cell wall material, occurs at the apex along the polar axis of the tube (Dumais et al., 2006; Rojas et al., 2011). Similarly, our experiments with propidium iodide (PI; McKenna et al., 2009; Rounds et al., 2011a) and pectin methyl esterase fused to GFP (McKenna et al., 2009) show that the wall is thickest at the very tip and suggest that wall materials are deposited at the polar axis, consistent with the initial model of exocytosis (Lancelle and Hepler, 1992). Experiments using tobacco (Nicotiana tabacum) pollen and a receptor-like kinase fused to GFP also indicate that exocytosis occurs largely at the apical polar axis (Lee et al., 2008).Many researchers argue that apical actin is critical for exocytosis (Lee et al., 2008; Cheung et al., 2010; Qin and Yang, 2011; Yan and Yang, 2012). More specifically, recent work suggests that the fringe participates in targeting vesicles and thereby contributes to changes in growth direction (Kroeger et al., 2009; Bou Daher and Geitmann, 2011; Dong et al., 2012). In this article, using three different inhibitors, namely brefeldin A (BFA), LatB, and potassium cyanide (KCN), we test the hypothesis that polarized pectin deposition in pollen tubes requires the actin fringe. Our data show that during normal growth, pectin deposition is focused to the apex along the polar axis of the tube. However, when growth is modulated, different end points arise, depending on the inhibitor. With BFA, exocytosis stops completely, and the fringe disappears, with the appearance of an actin aggregate at the base of the clear zone. LatB, as shown previously (Vidali et al., 2009), incompletely degrades the actin fringe and leaves a rim of F-actin around the apical dome. Here, we show that, in the presence of LatB, pectin deposition continues, with the focus of this activity shifting in position frequently as the slowly elongating pollen tube changes direction. With KCN, the actin fringe degrades completely, but exocytosis continues and becomes depolarized, with pectin deposits now occurring across a wide arc of the apical dome. This dome often swells as deposition continues, only stopping once normal growth resumes. Taken together, these results support a role for the actin fringe in controlling the polarity of growth in the lily pollen tube.     

The Endoplasmic Reticulum Is a Reservoir for WAVE/SCAR Regulatory Complex Signaling in the Arabidopsis Leaf

During plant cell morphogenesis, signal transduction and cytoskeletal dynamics interact to locally organize the cytoplasm and define the geometry of cell expansion. The WAVE/SCAR (for WASP family verprolin homologous/suppressor of cyclic AMP receptor) regulatory complex (W/SRC) is an evolutionarily conserved heteromeric protein complex. Within the plant kingdom W/SRC is a broadly used effector that converts Rho-of-Plants (ROP)/Rac small GTPase signals into Actin-Related Protein2/3 and actin-dependent growth responses. Although the components and biochemistry of the W/SRC pathway are well understood, a basic understanding of how cells partition W/SRC into active and inactive pools is lacking. In this paper, we report that the endoplasmic reticulum (ER) is an important organelle for W/SRC regulation. We determined that a large intracellular pool of the core W/SRC subunit NAP1, like the known positive regulator of W/SRC, the DOCK family guanine nucleotide-exchange factor SPIKE1 (SPK1), localizes to the surface of the ER. The ER-associated NAP1 is inactive because it displays little colocalization with the actin network, and ER localization requires neither activating signals from SPK1 nor a physical association with its W/SRC-binding partner, SRA1. Our results indicate that in Arabidopsis (Arabidopsis thaliana) leaf pavement cells and trichomes, the ER is a reservoir for W/SRC signaling and may have a key role in the early steps of W/SRC assembly and/or activation.The W/SRC (for WASP family verprolin homologous/suppressor of cAMP receptor regulatory complex) and Actin-Related Protein (ARP)2/3 complex are part of an evolutionarily conserved Rho-of-Plants (ROP)/Rac small GTPase signal transduction cascade that controls actin-dependent morphogenesis in a wide variety of tissues and developmental contexts (Smith and Oppenheimer, 2005; Szymanski, 2005; Yalovsky et al., 2008). Many of the components and regulatory relationships among the complexes were discovered based on the stage-specific cell-swelling and -twisting phenotypes of the distorted class of Arabidopsis (Arabidopsis thaliana) trichome mutants (Szymanski et al., 1999; Zhang et al., 2005, 2008; Djakovic et al., 2006; Le et al., 2006; Uhrig et al., 2007). However, in both maize (Zea mays) and Arabidopsis, W/SRC and/or ARP2/3 are required for normal pavement cell morphogenesis (Frank and Smith, 2002; Mathur et al., 2003b; Brembu et al., 2004). Compared with other Arabidopsis pavement cell mutants, the shape defects of the distorted group are relatively mild. However, the distorted mutants and spike1 (spk1) differ from most other morphology mutants in that they display gaps in the shoot epidermis, most frequently at the interface of pavement cells and stomata (Qiu et al., 2002; Le et al., 2003; Li et al., 2003; Mathur et al., 2003b; Zhang et al., 2005; Djakovic et al., 2006). The cell gaps may reflect either uncoordinated growth between neighboring cells or defective cortical actin-dependent secretion of polysaccharides and/or proteins that promote cell-cell adhesion (Smith and Oppenheimer, 2005; Hussey et al., 2006; Leucci et al., 2007).In tip-growing cells, there is a strict requirement for actin to organize the trafficking and secretion activities of the cell to restrict growth to the apex. In Arabidopsis, the W/SRC-ARP2/3 pathway is not an essential tip growth component, because null alleles of both W/SRC and ARP2/3 subunits do not cause noticeable pollen tube or root hair phenotypes (Le et al., 2003; Djakovic et al., 2006). However, reverse genetic analysis of the W/SRC subunit BRK1 and ARP2/3 in the tip-growing protonemal cells of Physcomitrella patens revealed the obvious importance of this pathway (Harries et al., 2005; Perroud and Quatrano, 2008). Along similar lines, in two different legume species, W/SRC subunits are required for a normal root nodulation response to symbiotic bacteria (Yokota et al., 2009; Miyahara et al., 2010), indicating a conditional importance for this pathway in root hair growth. These genetic studies centered on the W/SRC and ARP2/3 pathways, in addition to those that involve a broader collection of actin-based morphology mutants (Smith and Oppenheimer, 2005; Blanchoin et al., 2010), are defining important cytoskeletal proteins and new interactions with the endomembrane system during morphogenesis. However, it is not completely clear how unstable actin filaments and actin bundle networks dictate the growth patterns of cells (Staiger et al., 2009).The difficulty of understanding the functions of specific actin arrays can be explained, in part, by the fact that plant cells that employ a diffuse growth mechanism have highly unstable cortical actin filaments and large actin bundles that do not have a geometry that obviously relates to the direction of growth or a specific subcellular activity (Blanchoin et al., 2010). This is in contrast to the cortical endocytic actin patches in yeast (Saccharomyces cerevisiae; Evangelista et al., 2002; Kaksonen et al., 2003) and cortical meshworks in the lamellipodia of crawling cells (Pollard and Borisy, 2003) that reveal subcellular locations where actin works to locally control membrane dynamics. In thick-walled plant cells, the magnitude of the forces that accompany turgor-driven cell expansion exceed those that could be generated by actin polymerization by orders of magnitude (Szymanski and Cosgrove, 2009). Localized cell wall loosening or the assembly of an anisotropic cell wall generates asymmetric yielding responses to turgor-induced stress (Baskin, 2005; Cosgrove, 2005). Therefore, the actin-based control of cell boundary dynamics is indirect, and the actin cytoskeleton influences cell shape change, in part, by actin and/or myosin-dependent trafficking of hormone transporters (Geldner et al., 2001) and organelles (Prokhnevsky et al., 2008), including those that control the localized delivery of protein complexes and polysaccharides that pattern the cell wall (Leucci et al., 2007; Gutierrez et al., 2009). In this scheme for actin-based growth control, the actin network dynamically rearranges at spatial scales that span from approximately 1- to 10-µm subcellular domains that may locally position organelles (Cleary, 1995; Gibbon et al., 1999; Szymanski et al., 1999) to the more than 100-µm actin bundle networks that operate at the spatial scales of entire cells (Gutierrez et al., 2009; Dyachok et al., 2011). It is clear from the work of several laboratories that the W/SRC and ARP2/3 protein complexes are required to organize cortical actin and actin bundle networks in trichomes (Szymanski et al., 1999; Le et al., 2003; Deeks et al., 2004; Zhang et al., 2005) and cylindrical epidermal cells (Mathur et al., 2003b; Dyachok et al., 2008, 2011). A key challenge now is to understand how plant cells deploy these approximately 10- to 20-nm heteromeric protein complexes to influence the patterns of growth at cellular scales.The genetic and biochemical control of ARP2/3 is complicated, but this is a tractable problem in plants, because the pathway is relatively simple compared with most other species in which it has been characterized. For example, in organisms ranging from yeast to humans, there are multiple types of ARP2/3 activators, protein complexes, and pathways that activate ARP2/3 (Welch and Mullins, 2002; Derivery and Gautreau, 2010). However, the maize and Arabidopsis genomes encode only WAVE/SCAR homologous proteins that can potently activate ARP2/3 (Frank et al., 2004; Basu et al., 2005). Detailed genetic and biochemical analyses of the WAVE/SCAR gene family in Arabidopsis demonstrated that the plant activators function interchangeably within the context of the W/SRC and define the lone pathway for ARP2/3 activation (Zhang et al., 2008). Bioinformatic analyses are consistent with a prominent role for W/SRC in the angiosperms, because in general, WASH complex subunits, which are structurally similar to WAVE/SCAR proteins, are largely absent from the higher plant genomes, while WAVE/SCAR genes are highly conserved (Kollmar et al., 2012).The components and regulatory schemes of the W/SRC-ARP2/3 pathway in Arabidopsis and P. patens are conserved compared with vertebrate species that employ these same protein complexes (Szymanski, 2005). For example, mutant complementation tests indicate that human W/SRC and ARP2/3 complex subunits can substitute for the Arabidopsis proteins (Mathur et al., 2003b). Furthermore, biochemical assays of Arabidopsis W/SRC (Basu et al., 2004; El-Assal et al., 2004; Frank et al., 2004; Le et al., 2006; Uhrig et al., 2007) and ARP2/3 assembly (Kotchoni et al., 2009) have shown that the binary interactions among W/SRC subunits and ARP2/3 complex assembly mechanisms are indistinguishable from those that have been observed for human W/SRC (Gautreau et al., 2004) and yeast ARP2/3 (Winter et al., 1999). After an initial period of controversy concerning the biochemical control of W/SRC, it is now apparent that vertebrate W/SRC (Derivery et al., 2009; Ismail et al., 2009), like the ARP2/3 complex (Machesky et al., 1999), is intrinsically inactive and requires positive regulation by Rac and other factors to fully activate ARP2/3 (Ismail et al., 2009; Lebensohn and Kirschner, 2009; Chen et al., 2010). Although overexpression of dominant negative ROP mutants causes trichome swelling and a reduced trichome branch number (Fu et al., 2002), the involvement of ROPs in trichome morphogenesis has been difficult to prove with a loss-of-function ROP allele because so many ROPs are expressed in this cell type (Marks et al., 2009). Existing reports on ROP loss-of-function mutants demonstrate the importance of pavement cell morphogenesis but do not document a trichome phenotype (Fu et al., 2005; Xu et al., 2010). A recent report describes a clever strategy to generate ROP loss-of-function lines that used the ectopic expression of ROP-specific bacterial toxins. There was a strong association between inducible expression of the toxins and the appearance of trichomes with severe trichome swelling and reduced branch number phenotypes (Singh et al., 2012). Although the exact mechanism of ROP-dependent control of W/SRC remains to be determined, the results described above in combination with the detection of direct interactions between the ROPGEF SPK1, active forms of ROP, and W/SRC subunits (Basu et al., 2004, 2008; Uhrig et al., 2007) strongly suggest that W/SRC is a ROP effector complex.The major challenge in the field now is to better understand the cellular control of W/SRC and how the complex is partitioned into active and inactive pools. In mammalian cells that crawl on a solid substrate, current models propose that a cytosolic pool of inactive WAVE/SCAR proteins and W/SRC is locally recruited and activated at specific plasma membrane surfaces in response to signals from some unknown Rac guanine nucleotide-exchange factor (GEF), protein kinase, and/or lipid kinase (Oikawa et al., 2004; Lebensohn and Kirschner, 2009; Chen et al., 2010). However, in Drosophila melanogaster neurons (Bogdan and Klämbt, 2003) and cultured human melanoma cells (Steffen et al., 2004), there are large pools of W/SRC with a perinuclear or organelle-like punctate localization that has no obvious relationship to cell shape or motility, raising uncertainty about the cellular mechanisms of W/SRC activation and the importance of different subcellular pools of the complex.In plants, cell fractionation experiments indicate that SCAR1 and ARP2/3 have an increased association with membranes compared with their animal counterparts (Dyachok et al., 2008; Kotchoni et al., 2009). In tip-growing moss protonemal cells, both the W/SRC subunit BRK1 and ARP2/3 localize to a population of unidentified organelles within the apical zone (Perroud and Quatrano, 2008). Similar live-cell imaging experiments in Arabidopsis reported a plasma membrane localization for SCAR1 and BRK1 in a variety of shoot epidermal and root cortex, and their accumulation at young trichome branch tips and at three-way cell wall junctions may define subcellular domains for W/SRC-ARP2/3-dependent actin filament nucleation at the plasma membrane (Dyachok et al., 2008). However, to our knowledge, active W/SRC, defined here as the fraction of W/SRC that colocalizes with ARP2/3 or actin, has not been reported in plants, and the plasma membrane is not necessarily the only organelle involved in W/SRC regulation. For example, the reported accumulation of BRK1 and SCAR1 at three-way cell wall junctions has a punctate appearance at the cell cortex that may not simply correspond to the plasma membrane (Dyachok et al., 2008). Also, in young stage 4 trichomes, there was an uncharacterized pool of intracellular SCAR1, but not BRK1, that localized to relatively large punctate structures (Dyachok et al., 2008). The endoplasmic reticulum (ER) may also be involved in W/SRC regulation. The ER-localized DOCK family ROPGEF SPK1 (Zhang et al., 2010) physically associates with multiple W/SRC proteins (Uhrig et al., 2007; Basu et al., 2008) and, based on genetic criteria, is an upstream, positive regulator of the W/SRC-ARP2/3 pathway (Basu et al., 2008). In the leaf, one function of SPK1 is to promote normal trafficking between the ER and Golgi; however, arp2/3 mutants do not share ER-stress phenotypes with spk1 (Zhang et al., 2010), making it unclear if SPK1 and the ER are directly involved in W/SRC signaling.This paper focuses on the localization and control of the W/SRC subunit NAP1/GNARLED/NAPP/HEM1/2. Arabidopsis NAP1 directly interacts with the ROP/Rac effector subunit SRA1/PIROGI/KLUNKER/PIRP (Basu et al., 2004; El-Assal et al., 2004; Uhrig et al., 2007). Based on the equally severe syndrome of nap1 and arp2/3 null phenotypes, and double mutant analyses, the only known function of NAP1 is to positively regulate ARP2/3 (Brembu et al., 2004; Deeks et al., 2004; El-Din El-Assal et al., 2004; Li et al., 2004). The vertebrate SRA1-NAP1 dimer is important for W/SRC assembly (Gautreau et al., 2004) and forms an extended physical surface that trans-inhibits the C-terminal ARP2/3-activating domain of WAVE/SCAR (Chen et al., 2010). The plant NAP1 and SRA1 proteins share end-to-end amino acid conservation with their vertebrate homologs and may form a heterodimer with similar functions (Basu et al., 2004; El-Assal et al., 2004; Uhrig et al., 2007). We report here that Arabidopsis NAP1 is strongly associated with ER membranes. In a detailed series of localization experiments, we detect a complicated intracellular distribution of NAP1 among the ER, the nucleus, and unidentified submicrometer punctae. A large pool of ER-associated NAP1 is inactive, based on the low level of colocalization with actin.Its accumulation on the ER does not require activating signals from either SPK1 or SRA1. These data indicate that the ER is a reservoir for W/SRC signaling and suggest that early steps in the positive regulation of NAP1 and the W/SRC occur on the ER surface.     

Interdependence of Endomembrane Trafficking and Actin Dynamics during Polarized Growth of Arabidopsis Pollen Tubes

During polarized growth of pollen tubes, endomembrane trafficking and actin polymerization are two critical processes that establish membrane/wall homeostasis and maintain growth polarity. Fine-tuned interactions between these two processes are therefore necessary but poorly understood. To better understand such cross talk in the model plant Arabidopsis (Arabidopsis thaliana), we first established optimized concentrations of drugs that interfere with either endomembrane trafficking or the actin cytoskeleton, then examined pollen tube growth using fluorescent protein markers that label transport vesicles, endosomes, or the actin cytoskeleton. Both brefeldin A (BFA) and wortmannin disturbed the motility and structural integrity of ARA7- but not ARA6-labeled endosomes, suggesting heterogeneity of the endosomal populations. Disrupting endomembrane trafficking by BFA or wortmannin perturbed actin polymerization at the apical region but not in the longitudinal actin cables in the shank. The interference of BFA/wortmannin with actin polymerization was progressive rather than rapid, suggesting an indirect effect, possibly due to perturbed endomembrane trafficking of certain membrane-localized signaling proteins. Both the actin depolymerization drug latrunculin B and the actin stabilization drug jasplakinolide rapidly disrupted transport of secretory vesicles, but each drug caused distinct responses on different endosomal populations labeled by ARA6 or ARA7, indicating that a dynamic actin cytoskeleton was critical for some steps in endomembrane trafficking. Our results provide evidence of cross talk between endomembrane trafficking and the actin cytoskeleton in pollen tubes.Pollen tubes of flowering plants are specialized cells that deliver immotile sperm to the proximity of female gametes for successful reproduction (Johnson and Preuss, 2002). The growth of pollen tubes is both polar and directional (Hepler et al., 2001); many cellular activities contribute to such growth, the most important being the dynamics of the actin cytoskeleton system, targeted exocytosis, and endocytosis (Hepler et al., 2001).Pollen tubes contain longitudinal actin cables along the shank, which are important for providing structural support and acting as tracks for the movement of large organelles (Staiger et al., 1994). The apical area of pollen tubes instead contains dynamic filamentous actin (F-actin), as shown by fluorescently labeled actin-binding proteins (Kost et al., 1999; Fu et al., 2001; Chen et al., 2002; Wilsen et al., 2006). The dynamics of F-actin are critical for the polarized growth of pollen tubes. Genetically manipulating the activities of the small GTPases ROP (Kost et al., 1999; Fu et al., 2001; Cheung et al., 2008) and Rab (de Graaf et al., 2005), or of actin-binding proteins such as profilin and formin (Staiger et al., 1994; Chen et al., 2002; Cheung and Wu, 2004), disrupted F-actin dynamics and inhibited tube growth and caused apical bulges. Application of drugs such as latrunculin B (LatB) and jasplakinolide (Jas) showed similar effects (Gibbon et al., 1999; Vidali et al., 2001; Cardenas et al., 2005; Hörmanseder et al., 2005; Chen et al., 2007).Targeted exocytosis delivers building materials for cell membranes and cell walls and therefore is critical for maintaining growth polarity and directionality of growing pollen tubes (Hepler et al., 2001). Because targeted exocytosis brings more membrane and wall materials than needed to the apex of a pollen tube, an active endocytic system exists to retrieve excess secreted materials. In addition to this nonselective bulk membrane retrieval, pollen tubes may have selective and regulated endocytic trafficking pathways. For example, experiments using charged gold particles indicated the existence of two distinct endocytic pathways in tobacco (Nicotiana tabacum) pollen tubes (Moscatelli et al., 2007), and other studies showed that pollen tubes are able to take in materials from the extracellular matrix (Lind et al., 1996; Goldraij et al., 2006). The axis of targeted exocytosis correlated with the direction of tube growth and it asymmetrically changed toward the new apex during tube reorientation (Camacho and Malho, 2003; de Graaf et al., 2005). Disruption of membrane trafficking altered growth trajectories (de Graaf et al., 2005). Both suggest that membrane trafficking is a critical part of polarity maintenance and reorientation.As two important cellular processes in pollen tube growth, membrane trafficking and actin polymerization are conceivably dependent on each other. For example, several studies demonstrated that dynamic actin polymerization was essential for membrane trafficking (Hörmanseder et al., 2005; Wang et al., 2005; Chen et al., 2007; Lee et al., 2008), while others explored whether membrane trafficking affected actin polymerization (de Graaf et al., 2005; Hörmanseder et al., 2005). These studies, however, were mostly done with rapidly growing pollen tubes from tobacco or lily (Lilium longiflorum). For the model plant Arabidopsis (Arabidopsis thaliana), whose pollen tubes grow slower, little is known in this regard. Given a robust protocol for Arabidopsis pollen germination (Boavida and McCormick, 2007), it is now possible to investigate the interactions between these two cellular activities.In this study, we analyzed the effects of drug treatments on Arabidopsis pollen tubes expressing fluorescent protein probes for transport vesicles, endosomes, or the actin cytoskeleton. We show that perturbing actin dynamics by LatB or Jas treatments disrupted the V-shaped distribution of transport vesicles, caused aggregation, and finally dissipation of a subpopulation of endosomes, indicating that actin dynamics are critical at some steps of endomembrane trafficking. On the other hand, disturbing endomembrane trafficking with brefeldin A (BFA) or wortmannin abolished the F-actin structure at the apical region without affecting the longitudinal actin cables at the shank. These results provide evidence that endomembrane trafficking and actin dynamics interact at certain steps during polarized growth of Arabidopsis pollen tubes.     

Actin Reorganization Underlies Phototropin-Dependent Positioning of Nuclei in Arabidopsis Leaf Cells

In epidermal and mesophyll cells of Arabidopsis (Arabidopsis thaliana) leaves, nuclei become relocated in response to strong blue light. We previously reported that nuclear positions both in darkness and in strong blue light are regulated by the blue light receptor phototropin2 in mesophyll cells. Here, we investigate the involvement of phototropin and the actin cytoskeleton in nuclear positioning in epidermal cells. Analysis of geometrical parameters revealed that, in darkness, nuclei were distributed near the center of the cell, adjacent to the inner periclinal wall, independent of cell shape. Dividing the anticlinal wall into concave, convex, and intermediate regions indicated that, in strong blue light, nuclei became relocated preferably to a concave region of the anticlinal wall, nearest the center of the cell. Mutant analyses verified that light-dependent nuclear positioning was regulated by phototropin2, while dark positioning of nuclei was independent of phototropin. Nuclear movement was inhibited by an actin-depolymerizing reagent, latrunculin B, but not by a microtubule-disrupting reagent, propyzamide. Imaging actin organization by immunofluorescence microscopy revealed that thick actin bundles, periclinally arranged parallel to the longest axis of the epidermal cell, were associated with the nucleus in darkness, whereas under strong blue light, the actin bundles, especially in the vicinity of the nucleus, became arranged close to the anticlinal walls. Light-dependent changes in the actin organization were clear in phot1 mutant but not in phot2 and phot1phot2 mutants. We propose that, in Arabidopsis, blue-light-dependent nuclear positioning is regulated by phototropin2-dependent reorganization of the actin cytoskeleton.Positioning organelles is essential for cellular activities. The nucleus changes its position in a programmatic way during development and the cell cycle (Britz, 1979; Nagai, 1993; Chytilova et al., 2000). For example, before asymmetrical divisions that give rise to the formation of root hair cells or guard mother cells, the nucleus migrates to the future division plane (Britz, 1979). In elongating root hair cells of Arabidopsis (Arabidopsis thaliana), the nucleus is maintained at a fixed distance from the apex (Ketelaar et al., 2002).While the nuclear migrations before mitosis and in root hairs are developmental, nuclear positioning is also regulated environmentally. In the fern, Adiantum capillus-veneris, nuclei in prothallial cells change their intracellular positions in response to light (Kagawa and Wada, 1993, 1995). The nuclei are located along the anticlinal walls in darkness and move toward the outer periclinal walls in weak light and to the anticlinal walls in strong light (Kagawa and Wada, 1993, 1995; Tsuboi et al., 2007). This response is called light-dependent nuclear positioning. Since the response is induced in cells that exhibit neither cell division nor expansion, it is believed to have a physiological role, distinct from the nuclear positioning associated with development.Recently, light-dependent nuclear positioning was reported in the spermatophyte Arabidopsis (Iwabuchi et al., 2007). In epidermal and mesophyll cells of dark-treated leaves, nuclei are distributed along the inner periclinal wall. Under strong light, they become located along the anticlinal walls. In mesophyll cells, nuclear movement from inner periclinal to anticlinal walls is induced repeatedly and specifically by blue light of high-fluence rate (more than 50 μ mol m⁻² s⁻¹) and is regulated by the blue light receptor phototropin2. Interestingly, mesophyll cells of the phot2 mutant have aberrantly positioned nuclei even in darkness. By contrast, the involvement of phototropins in nuclear positioning has not yet been examined for epidermal cells.Phototropin is a blue light receptor containing two light oxygen voltage domains at the N terminus, which bind an FMN chromophore, and a Ser/Thr kinase domain at the C terminus, which undergoes blue-light-dependent autophosphorylation (Briggs et al., 2001a; Christie, 2007). Arabidopsis possesses phototropins1 and 2 (Huala et al., 1997; Jarillo et al., 2001; Kagawa et al., 2001; Sakai et al., 2001). Phototropins are shown microscopically and biochemically to localize to the plasma membrane region (Briggs et al., 2001b; Sakamoto and Briggs, 2002; Kong et al., 2006) and mediate several responses, including phototropism (Liscum and Briggs, 1995; Sakai et al., 2001), stomatal opening (Kinoshita et al., 2001), and chloroplast movements (Jarillo et al., 2001; Kagawa et al., 2001; Sakai et al., 2001). In general, phototropin1 is more sensitive to light than its paralog and mediates low-fluence-rate light responses, whereas phototropin2 functions predominantly under higher fluence rates (Sakai et al., 2001).While the photoreceptor eliciting these nuclear movements has been revealed, the motile system responsible for moving the nuclei is still unknown. In general, organelle movements depend on the cytoskeleton, with the specific roles for actin and microtubules dependent on the organelle and species (Wada and Suetsugu, 2004). In land plants, the actin cytoskeleton plays a pivotal role in positioning organelles, including nuclei, chloroplasts, mitochondria, and peroxisomes (Wada and Suetsugu, 2004; Takagi et al., 2009).The role of the cytoskeleton in developmental nuclear movements has been investigated. In growing root hairs of Arabidopsis, the nuclear movements are driven along actin filaments (Ketelaar et al., 2002), whereas, in tobacco (Nicotiana tabacum) BY-2 cells, the cell-cycle-based nuclear migration before mitosis is found to depend on microtubules (Katsuta et al., 1990). In interphase Spirogyra crassa cells, centering of nuclei is regulated by both actin filaments and microtubules, but in distinct ways (Grolig, 1998). To the best of our knowledge, the cytoskeletal basis of environmentally induced nuclear movements in land plants has not been elucidated.The best-characterized organelle movements are the light-induced orientation movements of chloroplasts, and although exceptions have been reported, this movement depends on actin (Britz, 1979; Takagi, 2003; Wada et al., 2003). Under weak light, chloroplasts gather at the periclinal walls, perpendicular to the direction of light (accumulation response), whereas under strong light, they become positioned along the anticlinal walls, parallel to the direction of light (avoidance response). Recently, for Arabidopsis, Kadota et al. (2009) characterized the nature of the actin filaments probably involved in these movements. With the onset of either accumulation or avoidance response, short actin filaments appear at the leading edge of each chloroplast.In Arabidopsis, light-dependent nuclear positioning shows similarities to the chloroplast avoidance response, with regard to the direction of movement, relevant photoreceptor (phototropin2), and effective fluence rate (Iwabuchi and Takagi, 2008). On the other hand, nuclei are larger than chloroplasts and might require thicker, more rigid actin bundles for effective motility. Here, we investigate the involvement of the actin cytoskeleton as well as phototropin in regulatory system for nuclear positioning in epidermal cells of Arabidopsis leaves.     

Involvement of the Sieve Element Cytoskeleton in Electrical Responses to Cold Shocks

This study dealt with the visualization of the sieve element (SE) cytoskeleton and its involvement in electrical responses to local cold shocks, exemplifying the role of the cytoskeleton in Ca²⁺-triggered signal cascades in SEs. High-affinity fluorescent phalloidin as well as immunocytochemistry using anti-actin antibodies demonstrated a fully developed parietal actin meshwork in SEs. The involvement of the cytoskeleton in electrical responses and forisome conformation changes as indicators of Ca²⁺ influx was investigated by the application of cold shocks in the presence of diverse actin disruptors (latrunculin A and cytochalasin D). Under control conditions, cold shocks elicited a graded initial voltage transient, ΔV₁, reduced by external La³⁺ in keeping with the involvement of Ca²⁺ channels, and a second voltage transient, ΔV₂. Cytochalasin D had no effect on ΔV₁, while ΔV₁ was significantly reduced with 500 nm latrunculin A. Forisome dispersion was triggered by cold shocks of 4°C or greater, which was indicative of an all-or-none behavior. Forisome dispersion was suppressed by incubation with latrunculin A. In conclusion, the cytoskeleton controls cold shock-induced Ca²⁺ influx into SEs, leading to forisome dispersion and sieve plate occlusion in fava bean (Vicia faba).It has been argued for a long time that sieve elements (SEs) are devoid of a cytoskeleton (Parthasarathy and Pesacreta, 1980; Thorsch and Esau, 1981; Evert, 1990), but more recent biochemical and cytological studies favor the opposite view. Actin as well as profilin were detected in phloem exudates of various monocot and dicot species (Schobert et al., 1998, 2000), while immunocytochemical tests showed the presence of actin and tubulin in phloem exudates of pumpkin (Cucurbita maxima; Kulikova and Puryaseva, 2002). Proteome analyses gave further credence to the occurrence of microfilaments in SEs in castor bean (Ricinus communis; profilin; Barnes et al., 2004), pumpkin (actin; Walz et al., 2004), canola (Brassica napus; actin, profilin1 and profilin2, actin-depolymerizing factor4; Giavalisco et al., 2006), and rice (Oryza sativa; actin1, actin-depolymerizing factor2, actin depolymerizing-factor3, and actin-depolymerizing factor6; Aki et al., 2008). Moreover, cytological evidence suggests residues of a cytoskeleton in SEs; fluorescent immunolabeling identified an actin/myosin system at the sieve plates (Chaffey and Barlow, 2002).Theoretical considerations also call for the presence of a cytoskeleton in SEs. Turnover and addressing of macromolecules (Fisher et al., 1992; Leineweber et al., 2000) requires a local distribution network in SEs. This function was attributed to an endoplasmic reticulum (ER) continuous to the ER strands running through pore plasmodesma units (Blackman et al., 1998) into the companion cells. Although such a mechanism is essentially conceivable, an interaction between the ER and cytoskeleton would provide a more conventional mode of intracellular distribution (Hepler et al., 1990; Boevink et al., 1998; Ueda et al., 2010; Yokota et al., 2011; Chen et al., 2012). Moreover, macromolecular trafficking through pore plasmodesma units (Lucas et al., 2001) was proposed to be executed by actin and myosin (Oparka, 2004), implying the presence of a cytoskeleton in SEs. Despite the massive circumstantial evidence, however, a complete cytoskeleton network and its spatial distribution in SEs have not been visually documented thus far.The existence of an SE cytoskeleton would raise questions regarding its task(s) in this highly specialized cell type. In other plant cells, the cytoskeleton was proposed to be engaged, among others, in ion channel operation and intracellular signaling (Trewavas and Malho, 1997; Mazars et al., 1997, and refs. therein; Thuleau et al., 1998; Örvar et al., 2000; Sangwan et al., 2001; Drøbak et al., 2004; Davies and Stankovic, 2006), as in animal cells (Janmey, 1998; Lange and Gartzke, 2006). For instance, K⁺ fluxes are regulated by actin dynamics (Hwang et al., 1997; Liu and Luan, 1998; Chérel, 2004), while Ca²⁺ influx into the cytoplasm appears to be mediated by voltage-dependent Ca²⁺-permeable channels associated with microtubules (Mazars et al., 1997; Thion et al., 1998) or by mechanosensitive channels possibly associated with microfilaments (Wang et al., 2004; Zhang et al., 2007).Both types of Ca²⁺-permeable channels probably reside in the SE plasma membrane (Knoblauch et al., 2001; Hafke et al., 2007, 2009; Furch et al., 2009), where they are likely involved in Ca²⁺-dependent systemic signaling (Furch et al., 2009; Hafke et al., 2009; van Bel et al., 2011; Hafke and van Bel, 2013). These channels are also putative initiators of Ca²⁺-induced signal transduction in SEs, leading to sieve-plate occlusion in response to local cold shocks (Thorpe et al., 2010). In fava bean (Vicia faba), Ca²⁺-dependent sieve tube occlusion by dispersion of special phloem-specific proteins (P-proteins) known as forisomes has been studied intensely (Knoblauch et al., 2001; Furch et al., 2007, 2009; Thorpe et al., 2010). Thus, apart from its distributive tasks, a cytoskeleton may be of major importance for intracellular signaling cascades in the highly specialized, sparsely equipped SEs.Our objective was to investigate the existence and spatial distribution of an SE cytoskeleton and its engagement in local signaling through Ca²⁺ influx brought about by cold shocks. This study dealt with the visualization of cytoskeletal components in intact sieve tubes using microinjection of fluorescent phalloidin and immunocytochemistry. Confocal laser-scanning micrography (CLSM) and transmission electron microscopy unequivocally showed a parietally located cylindrical actin meshwork. We demonstrated the engagement of the network in local cold shock-induced electrical responses and its association with Ca²⁺ influx, since we found effects of the Ca²⁺ channel blocker La³⁺ and of the cytoskeleton disruptor latrunculin A (LatA) on electrical signatures triggered by cold shocks and, by consequence, on forisome conformation changes.     

The nuclease activity of DNA2 promotes exonuclease 1–independent mismatch repair

The DNA mismatch repair (MMR) system is a major DNA repair system that corrects DNA replication errors. In eukaryotes, the MMR system functions via mechanisms both dependent on and independent of exonuclease 1 (EXO1), an enzyme that has multiple roles in DNA metabolism. Although the mechanism of EXO1-dependent MMR is well understood, less is known about EXO1-independent MMR. Here, we provide genetic and biochemical evidence that the DNA2 nuclease/helicase has a role in EXO1-independent MMR. Biochemical reactions reconstituted with purified human proteins demonstrated that the nuclease activity of DNA2 promotes an EXO1-independent MMR reaction via a mismatch excision-independent mechanism that involves DNA polymerase δ. We show that DNA polymerase ε is not able to replace DNA polymerase δ in the DNA2-promoted MMR reaction. Unlike its nuclease activity, the helicase activity of DNA2 is dispensable for the ability of the protein to enhance the MMR reaction. Further examination established that DNA2 acts in the EXO1-independent MMR reaction by increasing the strand-displacement activity of DNA polymerase δ. These data reveal a mechanism for EXO1-independent mismatch repair.

The mismatch repair (MMR) system has been conserved from bacteria to humans (1, 2). It promotes genome stability by suppressing spontaneous and DNA damage-induced mutations (1, 3, 4, 5, 6, 7, 8, 9, 10, 11). The key function of the MMR system is the correction of DNA replication errors that escape the proofreading activities of replicative DNA polymerases (1, 4, 5, 6, 7, 8, 9, 10, 12). In addition, the MMR system removes mismatches formed during strand exchange in homologous recombination, suppresses homeologous recombination, initiates apoptosis in response to irreparable DNA damage caused by several anticancer drugs, and contributes to instability of triplet repeats and alternative DNA structures (1, 4, 5, 7, 8, 9, 10, 11, 13, 14, 15, 16, 17, 18). The principal components of the eukaryotic MMR system are MutSα (MSH2-MSH6 heterodimer), MutLα (MLH1-PMS2 heterodimer in humans and Mlh1-Pms1 heterodimer in yeast), MutSβ (MSH2-MSH3 heterodimer), proliferating cell nuclear antigen (PCNA), replication factor C (RFC), exonuclease 1 (EXO1), RPA, and DNA polymerase δ (Pol δ). Loss-of-function mutations in the MSH2, MLH1, MSH6, and PMS2 genes of the human MMR system cause Lynch and Turcot syndromes, and hypermethylation of the MLH1 promoter is responsible for ∼15% of sporadic cancers in several organs (19, 20). MMR deficiency leads to cancer initiation and progression via a multistage process that involves the inactivation of tumor suppressor genes and action of oncogenes (21).MMR occurs behind the replication fork (22, 23) and is a major determinant of the replication fidelity (24). The correction of DNA replication errors by the MMR system increases the replication fidelity by ∼100 fold (25). Strand breaks in leading and lagging strands as well as ribonucleotides in leading strands serve as signals that direct the eukaryotic MMR system to remove DNA replication errors (26, 27, 28, 29, 30). MMR is more efficient on the lagging than the leading strand (31). The substrates for MMR are all six base–base mismatches and 1 to 13-nt insertion/deletion loops (25, 32, 33, 34). Eukaryotic MMR commences with recognition of the mismatch by MutSα or MutSβ (32, 34, 35, 36). MutSα is the primary mismatch-recognition factor that recognizes both base–base mismatches and small insertion/deletion loops whereas MutSβ recognizes small insertion/deletion loops (32, 34, 35, 36, 37). After recognizing the mismatch, MutSα or MutSβ cooperates with RFC-loaded PCNA to activate MutLα endonuclease (38, 39, 40, 41, 42, 43). The activated MutLα endonuclease incises the discontinuous daughter strand 5′ and 3′ to the mismatch. A 5'' strand break formed by MutLα endonuclease is utilized by EXO1 to enter the DNA and excise a discontinuous strand portion encompassing the mismatch in a 5''→3′ excision reaction stimulated by MutSα/MutSβ (38, 44, 45). The generated gap is filled in by the Pol δ holoenzyme, and the nick is ligated by a DNA ligase (44, 46, 47). DNA polymerase ε (Pol ε) can substitute for Pol δ in the EXO1-dependent MMR reaction, but its activity in this reaction is much lower than that of Pol δ (48). Although MutLα endonuclease is essential for MMR in vivo, 5′ nick-dependent MMR reactions reconstituted in the presence of EXO1 are MutLα-independent (44, 47, 49).EXO1 deficiency in humans does not seem to cause significant cancer predisposition (19). Nevertheless, it is known that Exo1^-/- mice are susceptible to the development of lymphomas (50). Genetic studies in yeast and mice demonstrated that EXO1 inactivation causes only a modest defect in MMR (50, 51, 52, 53). In agreement with these genetic studies, a defined human EXO1-independent MMR reaction that depends on the strand-displacement DNA synthesis activity of Pol δ holoenzyme to remove the mismatch was reconstituted (54). Furthermore, an EXO1-independent MMR reaction that occurred in a mammalian cell extract system without the formation of a gapped excision intermediate was observed (54). Together, these findings implicated the strand-displacement activity of Pol δ holoenzyme in EXO1-independent MMR.In this study, we investigated DNA2 in the context of MMR. DNA2 is an essential multifunctional protein that has nuclease, ATPase, and 5''→3′ helicase activities (55, 56, 57). Previous research ascertained that DNA2 removes long flaps during Okazaki fragment maturation (58, 59, 60), participates in the resection step of double-strand break repair (61, 62, 63), initiates the replication checkpoint (64), and suppresses the expansions of GAA repeats (65). We have found in vivo and in vitro evidence that DNA2 promotes EXO1-independent MMR. Our data have indicated that the nuclease activity of DNA2 enhances the strand-displacement activity of Pol δ holoenzyme in an EXO1-independent MMR reaction.     

Profilin-Dependent Nucleation and Assembly of Actin Filaments Controls Cell Elongation in Arabidopsis

Actin filaments in plant cells are incredibly dynamic; they undergo incessant remodeling and assembly or disassembly within seconds. These dynamic events are choreographed by a plethora of actin-binding proteins, but the exact mechanisms are poorly understood. Here, we dissect the contribution of Arabidopsis (Arabidopsis thaliana) PROFILIN1 (PRF1), a conserved actin monomer-binding protein, to actin organization and single filament dynamics during axial cell expansion of living epidermal cells. We found that reduced PRF1 levels enhanced cell and organ growth. Surprisingly, we observed that the overall frequency of nucleation events in prf1 mutants was dramatically decreased and that a subpopulation of actin filaments that assemble at high rates was reduced. To test whether profilin cooperates with plant formin proteins to execute actin nucleation and rapid filament elongation in cells, we used a pharmacological approach. Here, we used Small Molecule Inhibitor of Formin FH2 (SMIFH2), after validating its mode of action on a plant formin in vitro, and observed a reduced nucleation frequency of actin filaments in live cells. Treatment of wild-type epidermal cells with SMIFH2 mimicked the phenotype of prf1 mutants, and the nucleation frequency in prf1-2 mutant was completely insensitive to these treatments. Our data provide compelling evidence that PRF1 coordinates the stochastic dynamic properties of actin filaments by modulating formin-mediated actin nucleation and assembly during plant cell expansion.The actin cytoskeleton provides tracks for the deposition of cell wall materials and plays important roles during many cellular processes, such as cell expansion and morphogenesis, vesicle trafficking, and the response to biotic and abiotic signals (Baskin, 2005; Smith and Oppenheimer, 2005; Szymanski and Cosgrove, 2009; Ehrhardt and Bezanilla, 2013; Rounds and Bezanilla, 2013). Plant cells respond to diverse internal and external stimuli by regulating the turnover and rearrangement of actin cytoskeleton networks in the cytoplasm (Staiger, 2000; Pleskot et al., 2013). How these actin rearrangements sense the cellular environment and what accessory proteins modulate specific aspects of remodeling remain an area of active investigation (Henty-Ridilla et al., 2013; Li et al., 2014a, 2015).Using high spatial and temporal resolution imaging afforded by variable-angle epifluorescence microscopy (VAEM; Konopka and Bednarek, 2008), we quantified the behavior of actin filaments in Arabidopsis (Arabidopsis thaliana) hypocotyl epidermal cells (Staiger et al., 2009). There are two types of actin filament arrays in the cortical cytoplasm of epidermal cells: bundles and single filaments. Generally, actin bundles are stable with higher pixel intensity values, whereas individual actin filaments are fainter, more ephemeral, and constantly undergo rapid assembly and disassembly through a mechanism that has been defined as “stochastic dynamics” (Staiger et al., 2009; Henty et al., 2011; Li et al., 2012, 2015). Elongating actin filaments in the cortical cytoskeleton originate from three distinct locations: the ends of preexisting actin filaments, the side of filaments or bundles, and de novo in the cytoplasm. Plant actin filaments elongate at rates of 1.6 to 3.4 μm/s, which is the fastest assembly reported in eukaryotic cells. Distinct from the mechanism of treadmilling and fast depolymerization in vitro, however, the disassembly of single actin filaments occurs predominately through prolific severing activity (Staiger et al., 2009; Smertenko et al., 2010; Henty et al., 2011). A commonly held view is that the dynamic actin network in plant cells is regulated by the activities of conserved and novel actin-binding proteins (ABPs). Through reverse-genetic approaches and state-of-the-art imaging modalities, we and others have demonstrated that several key ABPs are involved in the regulation of stochastic actin dynamic properties in a wide variety of plants and cell types (Thomas, 2012; Henty-Ridilla et al., 2013; Li et al., 2014a, 2015). Through these efforts, the field has developed a working model for the molecular mechanisms that underpin actin organization and dynamics in plant cells (Li et al., 2015).Profilin is a small (12–15 kD), conserved actin-monomer binding protein present in all eukaryotic cells (dos Remedios et al., 2003). Profilin binds to actin by forming a 1:1 complex with globular (G-)actin, suppresses spontaneous actin nucleation, and inhibits monomer addition at filament pointed ends (Blanchoin et al., 2014). The consequences of profilin activity on actin filament turnover differ based on cellular conditions and the presence of other ABPs. In vitro studies show that the profilin-actin complex associates with the barbed ends of filaments and promotes actin polymerization by lowering the critical concentration and increasing nucleotide exchange on G-actin (Pollard and Cooper, 1984; Pantaloni and Carlier, 1993). When barbed ends are occupied by capping protein, profilin acts as an actin-monomer sequestering protein. These opposing effects of profilin might be a regulatory mechanism for profilin modulation of actin dynamics in cells. In addition to actin, profilin interacts with Pro-rich proteins, as well as polyphosphoinositide lipids in vitro (Machesky et al., 1994). Formin is an ABP that mediates both actin nucleation and processive elongation using the pool of profilin-actin complexes (Blanchoin et al., 2010). The primary sequence of formin includes a Pro-rich domain, named Formin Homology1 (FH1). Evidence from fission and budding yeast shows that profilin can increase filament elongation rates by binding to the FH1 domain (Kovar et al., 2003; Moseley and Goode, 2005; Kovar, 2006). The FH1 domain of Arabidopsis FORMIN1 (AtFH1) is also reported to modulate actin nucleation and polymerization in vitro (Michelot et al., 2005). Recently, two groups reported that profilin functions as a gatekeeper during the construction of different actin networks generated by formin or ARP2/3 complex in yeast and mammalian cells (Rotty et al., 2015; Suarez et al., 2015). These studies highlight the importance of profilin regulation in coordinating the different actin arrays present in the same cytoplasm of eukaryotic cells. However, direct evidence for how profilin facilitates formin-mediated actin nucleation or barbed end elongation in cells remains to be established.Genomic sequencing and isolation of PROFILIN (PRF) cDNAs from plants reveal that profilin is encoded by a multigene family. For example, moss (Physcomitrella patens) has three isovariants (Vidali et al., 2007) and maize (Zea mays) has five (Staiger et al., 1993; Kovar et al., 2001). In Arabidopsis, at least five PRF genes have been identified (Christensen et al., 1996; Huang et al., 1996; Kandasamy et al., 2002). Studies in maize show that the biochemical properties of profilin isoforms differ in vitro (Kovar et al., 2000). Moreover, the localization of profilin isoforms reveals organ-specific expression patterns. Detection of protein levels in vivo with isovariant-specific profilin antibodies demonstrate that Arabidopsis PRF1, PRF2, and PRF3 are constitutively expressed in vegetative tissues, whereas PRF4 and PRF5 are expressed mainly in flower and pollen tissues (Christensen et al., 1996; Huang et al., 1996; Ma et al., 2005).Several genetic studies on the functions of profilin in plants have been conducted. Reduction of profilin levels in P. patens results in the inhibition of tip growth, disorganization of F-actin, and formation of actin patches (Vidali et al., 2007). Moreover, it was shown that the interaction between profilin and actin or Pro-rich ligands is critical for tip growth in moss. Arabidopsis PRF1 has been demonstrated to be involved in cell elongation, cell shape maintenance, and control of flowering time through overexpression and antisense PRF1 transgenic plants, and further, the reduction of PRF1 inhibits the growth of hypocotyls (Ramachandran et al., 2000). However, investigation of a prf1-1 mutant, which contains a T-DNA insertion in the promoter region of the PRF1 gene, indicates that cell expansion of seedlings is promoted and that protein levels of PRF1 are regulated by light (McKinney et al., 2001). Recently, Müssar et al. (2015) reported a new Arabidopsis T-DNA insertion allele, prf1-4, that shows an obvious dwarf seedling phenotype. To date, however, there has not been a critical examination of the impact of the loss of profilin on the organization and dynamics of bona-fide single actin filaments in vivo.Here, we use a combination of genetics and live-cell imaging to investigate the role of PRF1 in the control of actin dynamics and its effect on axial cell expansion. We observed a significant decrease in the overall filament nucleation frequency in prf1 mutants, which is opposite to expectations if profilin suppresses spontaneous nucleation. Through a pharmacological approach, we found that nucleation frequency in wild-type cells treated with a formin inhibitor, SMIFH2, phenocopied prf1 mutants. We also analyzed the dynamic turnover of individual filaments in prf1 mutants and observed a significant decrease in the rate of actin filament elongation and maximum length of actin filaments. Specifically, we found that PRF1 favors the growth of a subpopulation of actin filaments that elongate at rates greater than 2 μm/s and similar results were obtained in cells after SMIFH2 treatment. Our results provide compelling evidence that Arabidopsis PRF1 contributes to stochastic actin dynamics by modulating formin-mediated actin nucleation and filament elongation during axial cell expansion.     

New-Onset Diabetes Mellitus After Transplantation in a Cynomolgus Macaque (Macaca fasicularis)

A 5.5-y-old intact male cynomolgus macaque (Macaca fasicularis) presented with inappetence and weight loss 57 d after heterotopic heart and thymus transplantation while receiving an immunosuppressant regimen consisting of tacrolimus, mycophenolate mofetil, and methylprednisolone to prevent graft rejection. A serum chemistry panel, a glycated hemoglobin test, and urinalysis performed at presentation revealed elevated blood glucose and glycated hemoglobin (HbA1c) levels (727 mg/dL and 10.1%, respectively), glucosuria, and ketonuria. Diabetes mellitus was diagnosed, and insulin therapy was initiated immediately. The macaque was weaned off the immunosuppressive therapy as his clinical condition improved and stabilized. Approximately 74 d after discontinuation of the immunosuppressants, the blood glucose normalized, and the insulin therapy was stopped. The animal''s blood glucose and HbA1c values have remained within normal limits since this time. We suspect that our macaque experienced new-onset diabetes mellitus after transplantation, a condition that is commonly observed in human transplant patients but not well described in NHP. To our knowledge, this report represents the first documented case of new-onset diabetes mellitus after transplantation in a cynomolgus macaque.Abbreviations: NODAT, new-onset diabetes mellitus after transplantationNew-onset diabetes mellitus after transplantation (NODAT, formerly known as posttransplantation diabetes mellitus) is an important consequence of solid-organ transplantation in humans.^{7-10,15,17,19,21,25-28,31,33,34,37,38,42} A variety of risk factors have been identified including increased age, sex (male prevalence), elevated pretransplant fasting plasma glucose levels, and immunosuppressive therapy.^{7-10,15,17,19,21,25-28,31,33,34,37,38,42} The relationship between calcineurin inhibitors, such as tacrolimus and cyclosporin, and the development of NODAT is widely recognized in human medicine.^{7-10,15,17,19,21,25-28,31,33,34,37,38,42} Cynomolgus macaques (Macaca fasicularis) are a commonly used NHP model in organ transplantation research. Cases of natural and induced diabetes of cynomolgus monkeys have been described in the literature;^14,43,45 however, NODAT in a macaque model of solid-organ transplantation has not been reported previously to our knowledge.     

Class XI Myosins Move Specific Organelles in Pollen Tubes and Are Required for Normal Fertility and Pollen Tube Growth in Arabidopsis

Pollen tube growth is an essential aspect of plant reproduction because it is the mechanism through which nonmotile sperm cells are delivered to ovules, thus allowing fertilization to occur. A pollen tube is a single cell that only grows at the tip, and this tip growth has been shown to depend on actin filaments. It is generally assumed that myosin-driven movements along these actin filaments are required to sustain the high growth rates of pollen tubes. We tested this conjecture by examining seed set, pollen fitness, and pollen tube growth for knockout mutants of five of the six myosin XI genes expressed in pollen of Arabidopsis (Arabidopsis thaliana). Single mutants had little or no reduction in overall fertility, whereas double mutants of highly similar pollen myosins had greater defects in pollen tube growth. In particular, myo11c1 myo11c2 pollen tubes grew more slowly than wild-type pollen tubes, which resulted in reduced fitness compared with the wild type and a drastic reduction in seed set. Golgi stack and peroxisome movements were also significantly reduced, and actin filaments were less organized in myo11c1 myo11c2 pollen tubes. Interestingly, the movement of yellow fluorescent protein-RabA4d-labeled vesicles and their accumulation at pollen tube tips were not affected in the myo11c1 myo11c2 double mutant, demonstrating functional specialization among myosin isoforms. We conclude that class XI myosins are required for organelle motility, actin organization, and optimal growth of pollen tubes.Pollen tubes play a crucial role in flowering plant reproduction. A pollen tube is the vegetative cell of the male gametophyte. It undergoes rapid polarized growth in order to transport the two nonmotile sperm cells to an ovule. This rapid growth is supported by the constant delivery of secretory vesicles to the pollen tube tip, where they fuse with the plasma membrane to enlarge the cell (Bove et al., 2008; Bou Daher and Geitmann, 2011; Chebli et al., 2013). This vesicle delivery is assumed to be driven by the rapid movement of organelles and cytosol throughout the cell, a process that is commonly referred to as cytoplasmic streaming (Shimmen, 2007). Cytoplasmic streaming in angiosperm pollen tubes forms a reverse fountain: organelles moving toward the tip travel along the cell membrane, while organelles moving away from the tip travel through the center of the tube (Heslop-Harrison and Heslop-Harrison, 1990; Derksen et al., 2002). Drug treatments revealed that pollen tube cytoplasmic streaming and tip growth depend on actin filaments (Franke et al., 1972; Mascarenhas and Lafountain, 1972; Heslop-Harrison and Heslop-Harrison, 1989; Parton et al., 2001; Vidali et al., 2001). Curiously, very low concentrations of actin polymerization inhibitors can prevent growth without completely stopping cytoplasmic streaming, indicating that cytoplasmic streaming is not sufficient for pollen tube growth (Vidali et al., 2001). At the same time, however, drug treatments have not been able to specifically inhibit cytoplasmic streaming; thus, it is unknown whether cytoplasmic streaming is necessary for pollen tube growth.Myosins are actin-based motor proteins that actively transport organelles throughout the cell and are responsible for cytoplasmic streaming in plants (Shimmen, 2007; Sparkes, 2011; Madison and Nebenführ, 2013). Myosins can be grouped into at least 30 different classes based on amino acid sequence similarity of the motor domain, of which only class VIII and class XI myosins are found in plants (Odronitz and Kollmar, 2007; Sebé-Pedrós et al., 2014). Class VIII and class XI myosins have similar domain architecture. The N-terminal motor domain binds actin and hydrolyzes ATP (Tominaga et al., 2003) and is often preceded by an SH3-like (for sarcoma homology3) domain of unknown function. The neck domain, containing IQ (Ile-Gln) motifs, acts as a lever arm and is bound by calmodulin-like proteins that mediate calcium regulation of motor activity (Kinkema and Schiefelbein, 1994; Yokota et al., 1999; Tominaga et al., 2012). The coiled-coil domain facilitates dimerization (Li and Nebenführ, 2008), and the globular tail functions as the cargo-binding domain (Li and Nebenführ, 2007). Class VIII myosins also contain an N-terminal extension, MyTH8 (for myosin tail homology8; Mühlhausen and Kollmar, 2013), and class XI myosins contain a dilute domain in the C-terminal globular tail (Kinkema and Schiefelbein, 1994; Odronitz and Kollmar, 2007; Sebé-Pedrós et al., 2014). Recently, Mühlhausen and Kollmar (2013) proposed a new nomenclature for plant myosins based on a comprehensive phylogenetic analysis of all known plant myosins that clearly identifies paralogs and makes interspecies comparisons easier (Madison and Nebenführ, 2013).The localization of class VIII myosins, as determined by immunolocalization and the expression of fluorescently labeled full-length or tail constructs, has implicated these myosins in cell-to-cell communication, cell division, and endocytosis in angiosperms and moss (Reichelt et al., 1999; Van Damme et al., 2004; Avisar et al., 2008; Golomb et al., 2008; Sattarzadeh et al., 2008; Yuan et al., 2011; Haraguchi et al., 2014; Wu and Bezanilla, 2014). On the other hand, class XI myosin mutants have been studied extensively in Arabidopsis (Arabidopsis thaliana), which revealed roles for class XI myosins in cell expansion and organelle motility (Ojangu et al., 2007, 2012; Peremyslov et al., 2008, 2010; Prokhnevsky et al., 2008; Park and Nebenführ, 2013). Very few studies have examined the reproductive tissues of class XI myosin mutants. In rice (Oryza sativa), one myosin XI was shown to be required for normal pollen development under short-day conditions (Jiang et al., 2007). In Arabidopsis, class XI myosins are required for stigmatic papillae elongation, which is necessary for normal fertility (Ojangu et al., 2012). Even though pollen tubes of myosin XI mutants have not been examined, the tip growth of another tip-growing plant cell has been thoroughly examined in myosin mutants. Root hairs are tubular outgrowths of root epidermal cells that function to increase the surface area of the root for water and nutrient uptake. Two myosin XI mutants have shorter root hairs, of which the myo11e1 (xik; myosin XI K) mutation has been shown to be associated with a slower root hair growth rate and reduced actin dynamics compared with the wild type (Ojangu et al., 2007; Peremyslov et al., 2008; Park and Nebenführ, 2013). Higher order mutants have a further reduction in root hair growth and have altered actin organization (Prokhnevsky et al., 2008; Peremyslov et al., 2010). Disruption of actin organization was also observed in myosin XI mutants of the moss Physcomitrella patens (Vidali et al., 2010), where these motors appear to coordinate the formation of actin filaments in the apical dome of the tip-growing protonemal cells (Furt et al., 2013). Interestingly, organelle movements in P. patens are much slower than in angiosperms and do not seem to depend on myosin motors (Furt et al., 2012).The function of myosins in pollen tubes is currently not known, although it is generally assumed that they are responsible for the prominent cytoplasmic streaming observed in these cells by associating with organelle surfaces (Kohno and Shimmen, 1988; Shimmen, 2007). Myosin from lily (Lilium longiflorum) pollen tubes was isolated biochemically and shown to move actin filaments with a speed of about 8 µm s⁻¹ (Yokota and Shimmen, 1994) in a calcium-dependent manner (Yokota et al., 1999). Antibodies against this myosin labeled small structures in both the tip region and along the shank (Yokota et al., 1995), consistent with the proposed role of this motor in moving secretory vesicles to the apex.In Arabidopsis, six of 13 myosin XI genes are highly expressed in pollen: Myo11A1 (XIA), Myo11A2 (XID), Myo11B1 (XIB), Myo11C1 (XIC), Myo11C2 (XIE), and Myo11D (XIJ; Peremyslov et al., 2011; Sparkes, 2011). The original gene names (Reddy and Day, 2001) are given in parentheses. Myo11D is the only short-tailed myosin XI in Arabidopsis (Mühlhausen and Kollmar, 2013) and lacks the typical myosin XI globular tail involved in cargo binding (Li and Nebenführ, 2007). The remaining genes have the same domain architecture as the conventional class XI myosins that have been shown to be involved in the elongation of trichomes, stigmatic papillae, and root hairs (Ojangu et al., 2007, 2012; Peremyslov et al., 2008, 2010; Prokhnevsky et al., 2008; Park and Nebenführ, 2013). Therefore, we predicted that these five pollen-expressed, conventional class XI myosins are required for the rapid elongation of pollen tubes. In this study, we examined transfer DNA (T-DNA) insertion mutants of Myo11A1, Myo11A2, Myo11B1, Myo11C1, and Myo11C2 for defects in fertility and pollen tube growth. Organelle motility and actin organization were also examined in myo11c1 myo11c2 pollen tubes.     

Endometrial Decidualization and Deciduosis in Aged Rhesus Macaques (Macaca mulatta)

Superficial decidualization of the endometrial stroma is an essential feature of the implantation stage of pregnancy in rhesus macaques and other primates. Decidualization involves proliferation of the endometrial stromal cells, with differentiation into morphologically distinct decidual cells. Previous reports involving nonpregnant rhesus monkeys have described localized and widespread endometrial decidualization in response to administration of progesterone and synthetic progestogens. Ectopic decidua or ‘deciduosis’ describes the condition in which groups of decidual cells are located outside of the endometrium, most often in the ovaries, uterus and cervix but also in various other organs. In humans, most cases of deciduosis are associated with normal pregnancy, and ectopic decidua can be found in the ovary in nearly all term pregnancies. Here we describe pronounced endometrial decidualization in 2 rhesus macaques. Both macaques had been treated long-term with medroxyprogesterone acetate for presumed endometriosis, which was confirmed in one of the macaques at postmortem examination. In one animal, florid extrauterine and peritoneal serosal decidualization was admixed multifocally with carcinomatosis from a primary colonic adenocarcinoma. Cells constituting endometrial and serosal decidualization reactions were immunopositive for the stromal markers CD10, collagen IV, smooth muscle actin, and vimentin and immunonegative for cytokeratin. In contrast, carcinomatous foci were cytokeratin-positive. To our knowledge, this report describes the first cases of serosal peritoneal decidualization in rhesus macaques. The concurrent presentation of serosal peritoneal decidualization with carcinomatosis is unique.Abbreviations: GnRH, gonadotropin-releasing hormone; PAS, periodic acid–Schiff; SMA, smooth-muscle actinSuperficial decidualization of the endometrial stroma is an essential feature of the implantation stage of pregnancy in rhesus macaques and other primates.^13,27,29,37 This process typically begins, and is most prominent, adjacent to the spiral arteries, eventually expanding to affect the endometrium uniformly.³⁵ The endometrial stroma surrounds and supports the endometrial glands and is composed mainly of endometrial stromal cells and blood vessels.³⁵ Decidualization involves proliferation of the endometrial stromal cells, with differentiation into morphologically distinct decidual cells.^7,27,38 Endometrial stromal cells transform into large, polyhedral, cytoplasm-rich cells with large amounts of stored glycogen and are often binucleated or polyploid in character.^{6,13,27,30,35} Ultrastructurally, decidualized cells have numerous ribosomes, prominent rough endoplasmic reticulum and Golgi complexes, and cytoplasmic accumulation of glycogen and lipid droplets.^13,35 Consistent with their stromal origin, decidualized cells express mesenchymal immunohistochemical markers, such as vimentin, desmin, and muscle-specific actin.^{6,7,14,16,20,22}Initiation of decidualization by attachment of the blastocyst to the uterine epithelium depends on previous sensitization by progesterone secretion, after a brief priming by estrogen.^12,13,27 Estrogen and progesterone regulate a series of complex interactions at the interface between the developing embryo and the cells in the stromal compartment, leading to the formation of a differentiated maternal tissue (decidua) that supports embryo growth and maintains early pregnancy.²⁷ Postovulatory levels of circulating progesterone increase and help maintain the differentiation of decidual cells.^{7,13,33,37,38}Ectopic decidua or ‘deciduosis’ describes the condition in which groups of decidual cells reside outside of the endometrium, most often in the ovaries, uterus, and cervix; the fallopian tubes, peritoneum, omentum, diaphragm, liver, skin, spleen, appendix, abdominal–pelvic lymph nodes, renal pelvis, and lungs of women have also been reported as affected.^{6,14,18,20,22,28,29,38} In humans, most cases of deciduosis are associated with normal pregnancy, and ectopic decidua have been reported in the ovary in 90.5% to 100% of term pregnancies.^{6-8,14,20,22,28-30,38} Occasional cases in nonpregnant or postmenopausal women have been attributed to progesterone-secreting active corpora lutea, progesterone secretion by the adrenal cortex, trophoblastic disease, exogenous progestational agents, and pelvic irradiation.^{6-8,14,18,20,22,28,38} Deciduosis is usually an incidental finding that regresses postpartum within 4 to 6 wk; rarely, florid reactions have been reported to cause peritonitis, adhesions, hydronephrosis and hematuria, acute bowel obstruction or perforation (or both), abdominal pain mimicking appendicitis, massive and occasionally fatal hemoperitoneum, vaginal bleeding, and pneumothorax.^{6,7,14,18,20,22,28,29,31}Previous reports involving nonpregnant rhesus macaques have described localized and widespread endometrial decidualization in response to the administration of progesterone, synthetic progestogens, or progesterone-releasing bioactive intrauterine devices and intravaginal rings and have referred to these changes as ‘pseudodecidualization’ to indicate the absence of pregnancy in these animals.^12,33,35,37 In macaques given low (but superphysiologic) levels of progestogens, decidual changes have been noted in localized regions (around spiral arteries and underneath superficial epithelium), whereas high doses of progesterone or synthetic progestagens can cause a more pronounced and extensive reaction.³⁵In cynomolgus macaques, extrauterine decidual cell plaques are rare histologic findings in the subcoelomic mesenchyme of the ovarian cortex.^8,30 Despite the frequency of the condition in women, deciduosis is postulated to be a rarely documented lesion in primates because it is most often observed in conjunction with pregnancy, and pregnant cynomolgus macaques are seldom used in toxicity studies.⁸ Here we describe the pronounced endometrial decidualization of 2 rhesus macaques, one of which also had florid extrauterine and peritoneal decidualization that was admixed multifocally with carcinomatosis. Both macaques had been treated long-term with medroxyprogesterone acetate for presumed endometriosis, which was confirmed in one of the macaques at postmortem examination. To our knowledge, this report describes the first cases of peritoneal decidualization in rhesus macaques as well as the concurrent occurrence of carcinomatosis, endometriosis and peritoneal decidualization in a macaque. The extensive intermixing of the cell populations presented a diagnostic challenge at pathologic examination, and accurate diagnosis was achieved only through the use of multiple immunohistochemical markers.     

Characterization of the Possible Roles for B Class MADS Box Genes in Regulation of Perianth Formation in Orchid

To investigate sepal/petal/lip formation in Oncidium Gower Ramsey, three paleoAPETALA3 genes, O. Gower Ramsey MADS box gene5 (OMADS5; clade 1), OMADS3 (clade 2), and OMADS9 (clade 3), and one PISTILLATA gene, OMADS8, were characterized. The OMADS8 and OMADS3 mRNAs were expressed in all four floral organs as well as in vegetative leaves. The OMADS9 mRNA was only strongly detected in petals and lips. The mRNA for OMADS5 was only strongly detected in sepals and petals and was significantly down-regulated in lip-like petals and lip-like sepals of peloric mutant flowers. This result revealed a possible negative role for OMADS5 in regulating lip formation. Yeast two-hybrid analysis indicated that OMADS5 formed homodimers and heterodimers with OMADS3 and OMADS9. OMADS8 only formed heterodimers with OMADS3, whereas OMADS3 and OMADS9 formed homodimers and heterodimers with each other. We proposed that sepal/petal/lip formation needs the presence of OMADS3/8 and/or OMADS9. The determination of the final organ identity for the sepal/petal/lip likely depended on the presence or absence of OMADS5. The presence of OMADS5 caused short sepal/petal formation. When OMADS5 was absent, cells could proliferate, resulting in the possible formation of large lips and the conversion of the sepal/petal into lips in peloric mutants. Further analysis indicated that only ectopic expression of OMADS8 but not OMADS5/9 caused the conversion of the sepal into an expanded petal-like structure in transgenic Arabidopsis (Arabidopsis thaliana) plants.The ABCDE model predicts the formation of any flower organ by the interaction of five classes of homeotic genes in plants (Yanofsky et al., 1990; Jack et al., 1992; Mandel et al., 1992; Goto and Meyerowitz, 1994; Jofuku et al., 1994; Pelaz et al., 2000, 2001; Theißen and Saedler, 2001; Pinyopich et al., 2003; Ditta et al., 2004; Jack, 2004). The A class genes control sepal formation. The A, B, and E class genes work together to regulate petal formation. The B, C, and E class genes control stamen formation. The C and E class genes work to regulate carpel formation, whereas the D class gene is involved in ovule development. MADS box genes seem to have a central role in flower development, because most ABCDE genes encode MADS box proteins (Coen and Meyerowitz, 1991; Weigel and Meyerowitz, 1994; Purugganan et al., 1995; Rounsley et al., 1995; Theißen and Saedler, 1995; Theißen et al., 2000; Theißen, 2001).The function of B group genes, such as APETALA3 (AP3) and PISTILLATA (PI), has been thought to have a major role in specifying petal and stamen development (Jack et al., 1992; Goto and Meyerowitz, 1994; Krizek and Meyerowitz, 1996; Kramer et al., 1998; Hernandez-Hernandez et al., 2007; Kanno et al., 2007; Whipple et al., 2007; Irish, 2009). In Arabidopsis (Arabidopsis thaliana), mutation in AP3 or PI caused identical phenotypes of second whorl petal conversion into a sepal structure and third flower whorl stamen into a carpel structure (Bowman et al., 1989; Jack et al., 1992; Goto and Meyerowitz, 1994). Similar homeotic conversions for petal and stamen were observed in the mutants of the AP3 and PI orthologs from a number of core eudicots such as Antirrhinum majus, Petunia hybrida, Gerbera hybrida, Solanum lycopersicum, and Nicotiana benthamiana (Sommer et al., 1990; Tröbner et al., 1992; Angenent et al., 1993; van der Krol et al., 1993; Yu et al., 1999; Liu et al., 2004; Vandenbussche et al., 2004; de Martino et al., 2006), from basal eudicot species such as Papaver somniferum and Aquilegia vulgaris (Drea et al., 2007; Kramer et al., 2007), as well as from monocot species such as Zea mays and Oryza sativa (Ambrose et al., 2000; Nagasawa et al., 2003; Prasad and Vijayraghavan, 2003; Yadav et al., 2007; Yao et al., 2008). This indicated that the function of the B class genes AP3 and PI is highly conserved during evolution.It has been thought that B group genes may have arisen from an ancestral gene through multiple gene duplication events (Doyle, 1994; Theißen et al., 1996, 2000; Purugganan, 1997; Kramer et al., 1998; Kramer and Irish, 1999; Lamb and Irish, 2003; Kim et al., 2004; Stellari et al., 2004; Zahn et al., 2005; Hernandez-Hernandez et al., 2007). In the gymnosperms, there was a single putative B class lineage that duplicated to generate the paleoAP3 and PI lineages in angiosperms (Kramer et al., 1998; Theißen et al., 2000; Irish, 2009). The paleoAP3 lineage is composed of AP3 orthologs identified in lower eudicots, magnolid dicots, and monocots (Kramer et al., 1998). Genes in this lineage contain the conserved paleoAP3- and PI-derived motifs in the C-terminal end of the proteins, which have been thought to be characteristics of the B class ancestral gene (Kramer et al., 1998; Tzeng and Yang, 2001; Hsu and Yang, 2002). The PI lineage is composed of PI orthologs that contain a highly conserved PI motif identified in most plant species (Kramer et al., 1998). Subsequently, there was a second duplication at the base of the core eudicots that produced the euAP3 and TM6 lineages, which have been subject to substantial sequence changes in eudicots during evolution (Kramer et al., 1998; Kramer and Irish, 1999). The paleoAP3 motif in the C-terminal end of the proteins was retained in the TM6 lineage and replaced by a conserved euAP3 motif in the euAP3 lineage of most eudicot species (Kramer et al., 1998). In addition, many lineage-specific duplications for paleoAP3 lineage have occurred in plants such as orchids (Hsu and Yang, 2002; Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009; Mondragón-Palomino et al., 2009), Ranunculaceae, and Ranunculales (Kramer et al., 2003; Di Stilio et al., 2005; Shan et al., 2006; Kramer, 2009).Unlike the A or C class MADS box proteins, which form homodimers that regulate flower development, the ability of B class proteins to form homodimers has only been reported in gymnosperms and in the paleoAP3 and PI lineages of some monocots. For example, LMADS1 of the lily Lilium longiflorum (Tzeng and Yang, 2001), OMADS3 of the orchid Oncidium Gower Ramsey (Hsu and Yang, 2002), and PeMADS4 of the orchid Phalaenopsis equestris (Tsai et al., 2004) in the paleoAP3 lineage, LRGLOA and LRGLOB of the lily Lilium regale (Winter et al., 2002), TGGLO of the tulip Tulipa gesneriana (Kanno et al., 2003), and PeMADS6 of the orchid P. equestris (Tsai et al., 2005) in the PI lineage, and GGM2 of the gymnosperm Gnetum gnemon (Winter et al., 1999) were able to form homodimers that regulate flower development. Proteins in the euAP3 lineage and in most paleoAP3 lineages were not able to form homodimers and had to interact with PI to form heterodimers in order to regulate petal and stamen development in various plant species (Schwarz-Sommer et al., 1992; Tröbner et al., 1992; Riechmann et al., 1996; Moon et al., 1999; Winter et al., 2002; Kanno et al., 2003; Vandenbussche et al., 2004; Yao et al., 2008). In addition to forming dimers, AP3 and PI were able to interact with other MADS box proteins, such as SEPALLATA1 (SEP1), SEP2, and SEP3, to regulate petal and stamen development (Pelaz et al., 2000; Honma and Goto, 2001; Theißen and Saedler, 2001; Castillejo et al., 2005).Orchids are among the most important plants in the flower market around the world, and research on MADS box genes has been reported for several species of orchids during the past few years (Lu et al., 1993, 2007; Yu and Goh, 2000; Hsu and Yang, 2002; Yu et al., 2002; Hsu et al., 2003; Tsai et al., 2004, 2008; Xu et al., 2006; Guo et al., 2007; Kim et al., 2007; Chang et al., 2009). Unlike the flowers in eudicots, the nearly identical shape of the sepals and petals as well as the production of a unique lip in orchid flowers make them a very special plant species for the study of flower development. Four clades (1–4) of genes in the paleoAP3 lineage have been identified in several orchids (Hsu and Yang, 2002; Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009; Mondragón-Palomino et al., 2009). Several works have described the possible interactions among these four clades of paleoAP3 genes and one PI gene that are involved in regulating the differentiation and formation of the sepal/petal/lip of orchids (Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009). However, the exact mechanism that involves the orchid B class genes remains unclear and needs to be clarified by more experimental investigations.O. Gower Ramsey is a popular orchid with important economic value in cut flower markets. Only a few studies have been reported on the role of MADS box genes in regulating flower formation in this plant species (Hsu and Yang, 2002; Hsu et al., 2003; Chang et al., 2009). An AP3-like MADS gene that regulates both floral formation and initiation in transgenic Arabidopsis has been reported (Hsu and Yang, 2002). In addition, four AP1/AGAMOUS-LIKE9 (AGL9)-like MADS box genes have been characterized that show novel expression patterns and cause different effects on floral transition and formation in Arabidopsis (Hsu et al., 2003; Chang et al., 2009). Compared with other orchids, the production of a large and well-expanded lip and five small identical sepals/petals makes O. Gower Ramsey a special case for the study of the diverse functions of B class MADS box genes during evolution. Therefore, the isolation of more B class MADS box genes and further study of their roles in the regulation of perianth (sepal/petal/lip) formation during O. Gower Ramsey flower development are necessary. In addition to the clade 2 paleoAP3 gene OMADS3, which was previously characterized in our laboratory (Hsu and Yang, 2002), three more B class MADS box genes, OMADS5, OMADS8, and OMADS9, were characterized from O. Gower Ramsey in this study. Based on the different expression patterns and the protein interactions among these four orchid B class genes, we propose that the presence of OMADS3/8 and/or OMADS9 is required for sepal/petal/lip formation. Further sepal and petal formation at least requires the additional presence of OMADS5, whereas large lip formation was seen when OMADS5 expression was absent. Our results provide a new finding and information pertaining to the roles for orchid B class MADS box genes in the regulation of sepal/petal/lip formation.