Actin bundling via LIM domains

The LIM domain is defined as a protein-protein interaction module involved in the regulation of diverse cellular processes including gene expression and cytoskeleton organization. We have recently shown that the tobacco WLIM1, a two LIM domain-containing protein, is able to bind to, stabilize and bundle actin filaments, suggesting that it participates to the regulation of actin cytoskeleton structure and dynamics. In the December issue of the Journal of Biological Chemistry we report a domain analysis that specifically ascribes the actin-related activities of WLIM1 to its two LIM domains. Results suggest that LIM domains function synergistically in the full-length protein to achieve optimal activities. Here we briefly summarize relevant data regarding the actin-related properties/functions of two LIM domain-containing proteins in plants and animals. In addition, we provide further evidence of cooperative effects between LIM domains by transiently expressing a chimeric multicopy WLIM1 protein in BY2 cells.Key words: Actin-binding proteins, actin-bundling, cysteine-rich proteins, cytoskeleton, LIM domainThe LIM domain is a ≈55 amino acid peptide domain that was first identified in 1990 as a common cystein-rich sequence found in the three homeodomain proteins LIN-11, Isl1 and MEC-3. It has since been found in a wide variety of eukaryotic proteins of diverse functions. Animals possess several families of LIM proteins, with members containing 1–5 LIM domains occasionally linked to other catalytic or protein-binding domains such as homeodomain, kinase and SH3 domains. In contrast, plants only possess two distinct sets of LIM proteins. One is plant-specific and has not been functionally characterized yet. The other one comprises proteins that exhibit the same overall structure as the animal cystein rich proteins (CRPs), i.e., two very similar LIM domains separated by a ≈50 amino acid-long interLIM domain and a relatively short and variable C-terminal domain (Fig. 1A). The mouse CRP2 protein was the first CRP reported to interact directly with actin filaments (AF) and to stabilize the latter.¹ Identical observations were subsequently described for the chicken CRP1 and tobacco WLIM1 proteins.^2,3 In addition, these two proteins were shown to arrange AF into cables both in vitro and in vivo and thus join the list of actin bundlers.Open in a separate windowFigure 1Domain maps for wild-type WLIM1 (A) and GFP-fused chimeric 3xWLIM1 (B). A. WLIM1 basically comprises a short N-terminal domain (Nt), two LIM domains (LIM1 and LIM2), an interLIM spacer (IL) and a C-terminal domain (Ct). B. 3xWLIM1 consists of three tandem WLIM1 copies. This chimeric protein has been fused in C-terminus to GFP and transiently expressed in tobacco BY2 cells.To identify the peptide domains of WLIM1 responsible for its actin-related properties/activities, we generated domain-deleted and single domain variants and submitted them to a series of in vivo and in vitro assays.⁴ Localization experiments established that both LIM domains are required to efficiently target the actin cytoskeleton in tobacco BY2 cells. High-speed (200,000 g) cosedimentation data confirmed that the actin-binding activity of WLIM1 relies on its LIM domains. Indeed, the deletion of either the first or the second LIM domain respectively resulted in a 5-fold and 10-fold decrease of the protein affinity for AF. Importantly, each single LIM domain was found able to interact with AF in an autonomous manner, although with a reduced affinity compared to the wild-type WLIM1. Low-speed (12,500 g) cosedimentation data and electron microscopy observations revealed that the actin bundling activity of WLIM1 is also triggered by its LIM domains. Surprisingly each single LIM domain was able to bundle AF in an autonomous manner, suggesting that WLIM1 has two discrete actin-bundling sites. However, the bundles induced by the variants containing only one LIM domain, i.e., LIM domain-deleted mutants and single LIM domains, differed from those induced by the full-length WLIM1. They appeared more wavy and loosely packed and formed only at relatively high protein:actin ratios. Together these data suggest that LIM domains are autonomous actin-binding and -bundling modules that function in synergy in wild-type WLIM1 to achieve optimal activities.To further assess the mechanism of cooperation between the LIM domains of plant CRP-related proteins, we generated a chimeric protein composed of three WLIM1 copies in tandem (3 × WLIM1, Fig. 1B), and transiently expressed it as a GFP-fusion in tobacco BY2 cells. We anticipated that such a six LIM domain-containing protein displays an even higher actin-bundling activity. (Fig. 2A) shows the typical actin cytoskeleton pattern in an expanding BY2 cell as visualized using the actin marker GFP-fABD2.⁵ As previously reported by Sheahan et al.,⁵ GFP-fABD2 decorated dense, transversely oriented, cortical networks as well as transvacuolar strands connecting the subcortical-perinuclear region to the cortex. Ectopic expression of WLIM1-GFP (BY2 cells normally do not express the WLIM1 gene) induced moderate but perceptible modifications of the actin cytoskeleton structure (Fig. 2B). Most AF are arranged in bundles thicker than those observed in GFP-fABD2 expressing cells and fine AF arrays are less frequently observed. As expected, this phenotype was significantly enhanced in cells transformed with the 3xWLIM1-GFP protein (Fig. 2C). Indeed, cells were almost devoided of fine AF arrays and exhibited very thick actin cables (Fig. 2C) that, at times (≈30 %), form atypical long looped structures (Fig. 2D). The appearance of such structures may result from the increase of cable stability and thickness induced by the 3xWLIM1-GFP protein, as these parameters are likely to determine, at least partially, the maximal length of actin bundles. Together the present observations support earlier data showing that LIM domains work in concert in LIM proteins to regulate actin bundling in plant cells. Strikingly, vertebrate and plant CRPs invariably contain two LIM domains. The lack, in these organisms, of CRP-related proteins combining more than two LIM domains may be explained by the fact that very thick cables, such as those induced by the artificial 3xWLIM1, may be too stable structures incompatible with the necessary high degree of actin cytoskeleton plasticity. As an exception, a muscle CRP-related protein with five LIM domains (Mlp84B) has been identified in Drosophila.⁶ However, rather than decorating actin filaments in an homogenous manner, this protein has been found to concentrate in a specialized region of the Z-discs where it stabilizes, in concert with D-titin, muscle sarcomeres.⁷Open in a separate windowFigure 2Typical actin cytoskeleton patterns in tobacco BY2 cells that have been transiently transformed, using a particle gun, with GFP-fABD2 (A), WLIM1-GFP (B), and 3xWLIM1-GFP (C and D). For each construct, more than 60 cells were analyzed by confocal microscopy. In the case of 3xWLIM1-GFP, two prevalent patterns have been observed (C and D). Bars = 20 µm.The relatively well conserved spacer length (≈50 amino acids) that separates the two LIM domains in vertebrate CRPs and related plant LIM proteins remains an intriguing feature the importance of which in actin cable organization remains to be established. Using electron microscopy we are currently evaluating the effects of the modification of the interLIM domain length on the structural properties of actin cables.     

Tubular actin filaments in tobacco guard cells

The dynamic remodeling of actin filaments in guard cells functions in stomatal movement regulation. In our previous study, we found that the stochastic dynamics of guard cell actin filaments play a role in chloroplast movement during stomatal movement. In our present study, we further found that tubular actin filaments were present in tobacco guard cells that express GFP-mouse talin; approximately 2.3 tubular structures per cell with a diameter and height in the range of 1–3 µm and 3–5 µm, respectively. Most of the tubular structures were found to be localized in the cytoplasm near the inner walls of the guard cells. Moreover, the tubular actin filaments altered their localization slowly in the guard cells of static stoma, but showed obvious remodeling, such as breakdown and re-formation, in moving guard cells. Tubular actin filaments were further found to be colocalized with the chloroplasts in guard cells, but their roles in stomatal movement regulation requires further investigation.Key words: actin dynamics, tubular actin filaments, chloroplast, guard cell, stomatal movementStomatal movement responses to surrounding environment are mediated by guard cell signaling.^1,2 Actin filaments within guard cells are dynamic cytoarchitectures and function in stomatal development and movement.³ Arrays of actin filaments in guard cells that are dependent on different stomatal apertures have also been reported in references ^4–7. For example, the random or longitudinal orientations of actin filaments in closed stomata change to a radial orientation or ring-like array after stomata opening.^5,6,8 The reorganization of the actin architecture during stomatal movement depends on the depolymerization and repolymerization of actin filaments in guard cells. In contrast to the traditional treadmill model of actin dynamic mechanisms, stochastic dynamics of actin have been revealed in plant cells, such as in the epidermal cells of hypocotyl and root, the pavement cells of Arabidopsis cotyledons, and the guard cells of tobacco (Nicotiana tabacum).^9–11 In this alternative system, the short actin fragments generated from severed long filaments can link with each other to form longer filaments by end-joining activity. The actin regulatory proteins, Arp2/3 complex, capping protein and actin depolymerizing factor (ADF)/cofilin, may also be involved in the stochastic dynamics of actin filaments.^12,13Using tobacco GFP-mouse talin expression lines, we have previously analyzed the stochastic dynamics of guard cell actin filaments and their roles in chloroplast displacement during stomatal movement.^6,11 We found from these analyses that another arrangement of actin filaments, i.e., tubular actin filaments, exists in the guard cells of these tobacco lines. We first found the circle-like actin filaments in 82% of the guard cells (counting 320 cells) in tobacco expressing GFPmouse talin when analyzing a single optical section (Fig. 1A). In a previous study of BY-2 cells expressing GFP-Lifeact labeled actin filaments, Smertenko et al. found similar structures, i.e., quoit-like structures or acquosomes in all of the plant tissues examined except growing root hairs.¹⁰ However, in our present analysis of serial sections, we determined that the circle-like actin filaments in the tobacco guard cells were long tubes (Fig. 1A), as the lengths (about 3–5 µm) of these structures were greater than their diameter (about 1–3 µm). Hence, we denoted these structures as tubular actin filaments to distinguish them from the circular conformations of actin filaments observed previously in other plant cell tissues.^10,14–19 About 2.3 of these tubular actin filaments were found per guard cell, which is less than the number of acquosomes reported in BY-2 cells (about 6.7 per cell).¹⁰ Analysis of serial optical sections at the z-axis revealed that the tubular actin filaments localize in the cytoplasm near the inner walls of the guard cells (Fig. 1B), which is similar to the distribution of chloroplasts in guard cells.¹¹ Longitudinal sections further revealed a colocalization of tubular actin filaments and chloroplasts (Fig. 1B).Open in a separate windowFigure 1Tubular actin filaments in the guard cells of a tobacco (Nicotiana tabacum) line expressing GFP-mouse talin. (A) Optical-sections (interval, 1.5 µm) of guard cells in a moving stoma showing tubular actin filaments (arrow heads). Frames (a1) and (a2) are cross sections of 1.5-µm-picture through the yellow and red lines, respectively, revealing the cross section of the circle structures are parallel lines (arrows). (B) Optical-sections of a stoma from the outer periclinal walls to the inner walls of the guard cells (interval, 1 µm). The tubular actin filaments (arrow heads) are localized in the cytoplasm near to the inner periclinal walls of guard cells. Frame (b1) is the guard cell on the right of the frame “4 µm”; (b2) is the cross section of b1 through the red line; and (b3) is a higher magnification image of the area encompassed by the white square in b2. Arrows indicate the colocalization between the tubular actin filaments and the chloroplast (indicated using a red pseudocolor). (C) Time-series imaging showing the movement of tubular actin filaments in the guard cells of static stomata. Frame (c1) comprises three images colored red (0 S), green (40 S) and blue (80 S), that are merged in a single frame to show the translocation of the tubular actin filaments (arrows). (D) Time-series images of the opening stomata showing the breakdown (arrows) and re-formation (arrowheads) of the tubular actin filaments. All images were captured using a Zeiss LSM 510 META confocal laser scanning microscope, as described by Wang et al.¹¹ Bars, 10 µm.We performed time-lapse imaging and found that the translocation of tubular actin filaments is slow in static stomata in which the distance between two tubular actin filaments typically increased from 2.22 to 2.50 µm after 80 sec (Fig. 1C). In moving stomata, however, the tubular actin filaments showed an obvious dynamic reorganization whereby they could be processed into short fragments and also reemerged after they had disintegrated (Fig. 1D). These results indicate that tubular actin filaments have stochastic dynamics that are similar to the long actin filaments of guard cells.¹¹ In our previous study, we found that the stochastic dynamics of actin filaments correlate with light-induced chloroplast movement in guard cells.¹¹ However, whether the dynamics of the tubular actin filaments are also involved in chloroplast movement during stomatal movement remains to be investigated. In cultured mesophyll cells which had been mechanically isolated from Zinnia elegans, Wilsen et al. previously found a close association between fully closed actin rings and chloroplasts.¹⁸ These authors further found that the average percentage of cells with free actin rings increased at the initial culture stage, and then decreased, which indicates that the formation of actin rings might be a response of the actin cytoskeleton to cellular stress or disturbance.¹⁸ The turgor pressure of guard cells is the fundamental basis of stomatal movement leading to changes in the shape, volume, wall structure, and membrane surface of guard cells.^20–24 We speculate from our current data that there is a relationship between tubular actin filaments and the shape changes of guard cells during stomatal movement.     

Plant-Derived Small Molecule Inhibitors of Neuronal NO-Synthase: Potential Effects on Protein Degradation

Cigarette smoking is known to cause a decrease in NO production in man resulting in a variety of pathological effects, including vascular dysfunction. Aqueous extracts of cigarette and cigarette smoke contain chemical inhibitors to NO-synthases, a heme-containing cytochrome P450 enzymes. More recently, it was shown that freshly harvested leaves from the tobacco plant (Nicotiana tabacum, Solanaceae) also contain chemical inhibitors to neuronal NO-synthase (nNOS). Examination of leaves from 32 other plants representing diverse members of the plant kingdom showed that 17 other plants, besides tobacco, contain these chemical inhibitors. Of all these plants, 16 are members of the core asterids flowering plant group and 6 are members of the Solanaceae family. Although the identity of the chemicals is not known, perhaps the closely related plants contain the same or similar compounds that inhibit nNOS. The inhibitory effects are not attributable to nicotine. The discovery of these chemicals and their further characterization may help to explain the loss of nNOS in smokers. In this addendum, we discuss these results in light of the effect of tobacco-derived chemicals in inhibiting P450 cytochromes, as well as our thoughts on how the inactivation of nNOS leads to its selective downregulation through proteolytic degradation.Key Words: nitric oxide, inhibitors, tobacco, natural products, cytochrome P450, nitric oxide synthase, heat shock proteinsNO-synthase (NOS) is a cytochrome P450 like hemeprotein enzyme that catalyzes the production of NO from L-arginine.^1,2 NO is very important cellular signaling molecule, which plays a role in a variety of physiological processes including blood pressure regulation, neurotransmission, and penile erection. Like other P450 cytochromes, the catalytic site of NOS contains a heme prosthetic group that upon reduction and complexation with CO forms a chromophore that absorbs in the 450 nm region. Thus, although not closely related with respect to amino acid sequence to other P450 enzymes, the catalytic core of NOS and the chemistry involved in catalysis by NOS are highly similar to other P450 cytochromes.The cytochrome P450 enzymes comprise a very large family of enzymes that are responsible for metabolism of drugs and other xenobiotics as well as for synthesis of many hormones. There are over 2,000 P450 genes characterized in animals with over 50 genes in humans and over 1,900 genes in plants. The diversity of the P450 family is thought to reflect a complex “battle” between plants and animals.³ That is, plants are thought to have developed new alkaloids with new enzymes to protect themselves against animals whereas the animals developed new enzymes to metabolize these newly created plant toxins. This is thought to explain why the diversity in CYP genes increased dramatically approximately 400 millions years ago when organisms moved from the oceans to land.^3,4 Consistent with this hypothesis, chemicals in plants have been documented to inhibit mammalian P450 cytochromes. More importantly for humans, a variety of plant products have been documented to interact clinically with P450 cytochromes.⁵ For example, furanocoumarins found in grapefruit juice are known to enhance the oral bioavailability of felodipine, a calcium channel blocker, by inhibiting intestinal P450 metabolism of the drug.⁶In the case of tobacco-based cigarettes, hydrophobic chemicals in cigarettes have been found to inhibit P450 cytochromes. In particular, aromatase, the terminal P450 enzyme responsible for estrogen synthesis in humans, has been shown to be inhibited by tobacco smoke.⁷ Acyl derivatives of noricotine and anabasine were found to be active inhibitors of aromatase. In the case of NOS, studies by Heitzer et al.⁸ showed the amelioration of the vascular dysfunction in smokers by tetrahydrobiopterin and the authors concluded that there was a loss of endothelial NO synthesis caused by smoking. Moreover, Xie et al.⁹ showed that exposing rats to cigarette smoke led to the loss of both neuronal NO-synthase (nNOS) activity and protein. This observation reminded us of our earlier studies showing that guanabenz is a time-dependent irreversible inhibitor of nNOS that causes a loss of the penile nNOS activity and protein when given to rats.¹⁰ Thus, based on these findings, we wondered if chemicals in cigarettes could interact with NOS and explain the loss of NOS. We discovered that aqueous extracts of cigarettes and cigarette smoke could inhibit nNOS¹¹ and endothelial NOS.¹² More recently, extracts from freshly harvested tobacco leaves and leaves of various other plants from diverse phylogeny were also examined.¹³Tobacco leaves contained inhibitors to nNOS, indicating that these compounds did not necessarily arise from the curing and processing of the tobacco leaves to make cigarettes or from burning of the cigarettes. Moreover, other extracts made from other plants including 6 from the Solanacea family were found to have inhibitory activity. Thus, we believe that some common chemical or related chemicals exist that inhibits nNOS. The inhibitors are small molecules with hydrophilic and cationic qualities. In comparison, the inhibitors to aromatase are more hydrophobic than those of nNOS, likely reflecting the hydrophobic nature of the active site of aromatase, which accepts steroidal androgens. On the other hand, nNOS accepts L-arginine, a cationic water-soluble compound. The water-soluble nature of the inhibitors has so far hindered the isolation and characterization of the nNOS inhibitors, and thus we do not know the identity of these chemicals.The irreversible nature of the interaction of tobacco inhibitors with nNOS found in our studies may be important in understanding the long-term consequences of smoking. The irreversible inactivation of nNOS by various guanidine-based compounds is known to cause the enhanced proteasomal degradation of nNOS.¹⁴ The dysfunctional nNOS is known to be selectively ubiquitinated¹⁵ by a process involving Hsp70 and CHIP¹⁶ (Fig. 1). The degradation is also accelerated by inhibition of Hsp90, indicating that NOS is also regulated by the Hsp90-based chaperones.¹⁴ The loss of nNOS protein due to cigarette smoke suggests a similar process may be occurring with smokers.Open in a separate windowFigure 1Ubiquitination and Degradation of Inactivated nNOS. The nNOS is inactivated by chemicals found in some plants by alterations directed at the heme-containing active site of nNOS. This dysfunctional form of nNOS is directed for ubiquitination by an E3 ligase, such as CHIP (C-terminal Hsp-interacting protein), hsp70, and E2 ligase enzyme. The ubiquitinated nNOS is then recognized for proteasomal degradation. Closed circles represent ubiquitin molecules.The discovery of nNOS inhibitors in plants could be adequately interpreted in the context of plant-animal warfare. However, in light of the recent discovery of plant NOS,¹⁷ and in particular NOS in tobacco cells,¹⁸ it is possible that these inhibitors are endogenous modulators of the plant NOS. In this respect, an endogenous inhibitor has been identified for mammalian NOS.¹⁹ Furthermore, considering that the Hsp90-and Hsp70-based chaperones, ubiquitin, CHIP, and proteasome are found in plants and serve similar functions,^20–23 the NOS in plants may be similarly regulated. Thus, the mechanism outlined in Figure 1 for nNOS may also be pertinent to regulation of plant NOS. We believe this process reflects a fundamental biological process of protein quality control that is greatly affected by the binding of small molecules to the target protein.     

Filopodial focal complexes direct adhesion and force generation towards filopodia outgrowth

Filopodia are key structures within many cells that serve as sensors constantly probing the local environment. Although filopodia are involved in a number of different cellular processes, their function in migration is often analyzed with special focus on early processes of filopodia formation and the elucidation of filopodia molecular architecture. An increasing number of publications now describe the entire life cycle of filopodia, with analyses from the initial establishment of stable filopodium-substrate adhesion to their final integration into the approaching lamellipodium. We and others can now show the structural and functional dependence of lamellipodial focal adhesions as well as of force generation and transmission on filopodial focal complexes and filopodial actin bundles. These results were made possible by new high resolution imaging techniques as well as by recently developed elastomeric substrates and theoretical models. The data additionally provide strong evidence that formation of new filopodia depends on previously existing filopodia through a repetitive filopodial elongation of the stably adhered filopodial tips. In this commentary we therefore hypothesize a highly coordinated mechanism that regulates filopodia formation, adhesion, protein composition and force generation in a filopodia dependent step by step process.Key words: filopodia, focal adhesion, cell force, filopodial focal complex, actinCell protrusion depends on collaborative interactions of lamellipodia and filopodia.¹ Although filopodia cannot drive cell migration alone, in contrast to lamellipodia, they are essential for many cell biological functions such as guidance of neuronal growth cones² or during angiogenesis.³ Furthermore, filopodia are vital to cell-cell contact establishment as described for epithelial cells⁴ or during dorsal closure in Drosophila,⁵ and are also implicated in cancer cell metastasis.^6,7 Lamellipodia as well as filopodia can be formed independently from each other,⁸ and recent results implicate independent basic mechanisms of cytoskeletal regulation for their formation. While lamellipodia protrusion is a well accepted Arp2/3-dependent process where actin branches constantly form the protrusive force at the leading edge of the lamella,⁹ the details of filopodia formation are far from being understood.^10–13 Although earlier experiments indicated Arp2/3 was also involved in filopodia formation,¹⁴ recent results point to a machinery that is far less dependent, or even possibly independent, of Arp2/3 with formins being the central regulating molecules instead.⁸As soon as filopodia start to form, they constantly sense their environment upon elongation. Transmembrane proteins such as cadherins or integrins^15,16 connect filopodia to surrounding cells, extracellular matrix, or even pathogens to form stable contacts. When filopodial adhesion fails, retraction takes place.¹⁷ Although integrins and talin have been shown to be initially present at these sites in an un-clustered but active state, many additional adhesion proteins take part in filopodial focal complexes (filopodial FXs).^16,18 Starting from a small VASP-containing adhesion spot at the tip of filopodia, proteins such as vinculin, paxillin, talin, tensin and even zyxin form an elongated filopodial FX behind the VASP spot along the filopodium. While integrin as well as VASP transport along the filopodia shaft via myosin-X has been described,¹⁹ it is still unclear whether additional adhesion proteins are also actively transported or whether diffusion takes place. Diffusion is typically a non-limiting process during cytoplasmic protein complex formation. However for filopodia, diffusion could have an important regulatory function as already hypothesized in theoretical models,²⁰ because they are small in width and densely filled with actin filaments. Therefore, local concentrations of soluble adhesion molecules might drop within filopodia upon FX formation resulting in a pure physical regulation of filopodial length as well as filopodial FX size.The almost complete focal adhesion site specific protein inventory of filopodia FXs^16,18 as indicated above provided further indications for a dependency of lamellipodial focal adhesions (FAs) on filopodial FXs. This hypothesis was confirmed using fluorescent live cell imaging to identify the transition of filopodial FXs into fully assembled FAs upon FX contact with the leading edge of the lamellipodium. While filopodial FXs were responsible for only a sub-fraction of FAs in fish fibroblasts,¹⁸ stable FAs of human keratinocytes were formed almost exclusively by enlargement of existing filopodial FXs¹⁶ (see scheme, Fig. 1).Open in a separate windowFigure 1Filopodia determine the fate of lamellipodial structures. Filopodia are formed by actin polymerization at their tip. Upon stable adhesion, a small but fully assembled filopodial focal complex (FX) is formed. This FX becomes enlarged in size upon lamellipodial contact to form focal adhesions. In parallel, the filopodial actin cross-linker fascin becomes exchanged by palladin and α-actinin as soon as the filopodial actin bundles are incorporated into the lamellipodium. In a next step, α-actinin becomes partially exchanged by myosin II, leading to enhanced force values applied at filopodial-originated FA sites bound to the substrate. The tight interaction between FAs and filopodial actin bundles reduces the actin retrograde flow within the filopodium in front of the FA (lower inlay) compared to filopodia lacking stable FAs in the lamellipodium (not shown). Adhesion sites formed in the lamellipodium lack connections to distinct actin bundles leading to low force application at these sites and short lifetimes (upper inlay).The structural dependency of lamellipodial complexes on filopodial protein aggregates could be also shown for actin bundles. Here, parallel oriented actin filaments become cross-linked by proteins such as fascin or IRSp53-Eps8-complex upon filopodia formation.^21,22 These tightly packed bundles of 15–30 single actin filaments originate from the lamellipodial actin meshwork.²³ Interestingly, filopodial actin bundles in turn also affect lamellipodial actin structures independent of whether the filopodium adheres in a stable manner or looses contact. Nemethova et al.¹⁸ described the contribution of non-adhering filopodia to the construction of concave bundles of actin filaments within the lamellipodium of fish fibroblasts. These bundles often extended in length and interconnected with adjacent bundles. Similar observations were found for fibroblasts of chicken embryos and neuronal growth cones.^24,25 Here, filopodial actin bundles were clearly shown to be the origin of nearly 85% of all actin bundles found in the lamella. These actin filaments typically pointed towards the direction of migration. Additionally, myosin II was associated with these filopodial derived actin filaments to form polarized actin bundles. Of equal importance are findings presented by Schäfer et al. in this issue. The authors analyzed the fate of stably adhered filopodia and identified a stepwise exchange of filopodial fascin against the actin cross-linker proteins palladin and especially α-actinin in areas where filopodia were just overgrown by the lamellipodial leading edge (schematically presented in Fig. 1). α-Actinin further induced incorporation of myosin II into filopodial actin bundles in the lamellipodium. The authors additionally found that FAs displayed an enhanced lifetime when adhered to these myosin containing actin filaments. Therefore, these findings could also explain the unusual stability of filopodial actin filaments in neuronal growth cones observed by Mallavarapu and Mitchison.¹⁷ For keratinocytes, filopodia-dependent actin bundles are the only myosin containing actin structures oriented in the direction of movement within the lamellipodium and the lamella. Sensitivity and resolution improvements in cell force analyses further proved that these actin bundles were responsible for almost the entire force transmitted from the lamellipodium of migrating keratinocytes to the substrate. These forces were transferred at FA sites emerging from filopodial FXs, proving the importance of filopodia in lamellipodial structures and functions. Although filopodia-independent adhesion sites are also formed in keratinocytes right behind the leading edge, these sites are neither connected to detectable actin filament bundles nor do they transmit significant forces (see scheme, Fig. 1). Consequently, their sizes and life spans are strongly reduced (Schäfer et al., this issue).Recent results in keratinocytes additionally close the circle from stably adhered filopodia to the generation of new ones. Our original observations indicated that new filopodia were mainly formed in a direct extension of focal adhesions. Since these adhesion sites also depended on previously adhered filopodial FXs, a closer look revealed a consecutive outgrowth of the same filopodia.¹⁶ These cycles were only interrupted when outgrowing filopodia did not adhere in a stable manner between outgrowth cycles. Present results suggest that the same tip complex is present in all subsequently formed filopodia with a VASP tip signal remaining in place during successive filopodial elongations. As a result, well aligned, consecutive elongated focal adhesions can be found in keratinocytes. We can only speculate whether such an Arp2/3-independent mechanism describes a basic principle in filopodia formation at this point, but similar results have been observed for fish fibroblasts with a repetitive and alternating transition between filopodia and microspikes as filopodia-like structures barely extending over the lamellipodial leading edge.¹⁸The strong interdependency between lamellipodial FAs and stably adhered filopodia is also highlighted by actin retrograde flow analyses in keratinocytes (Schäfer et al. this issue). Retrograde actin flow is generated by actin polymerization at the cell front and myosin activity pulling the filaments rearwards. The interaction of actin with FAs is known to dampen flow rates in front of lamellipodial FAs.²⁶ Furthermore, filamentous-actin dynamics measured in lung epithelial cells showed a fast retrograde actin flow at the leading edge compared to rates within the lamellae. The highest flow rates were in the range of 0.3–0.5 µm/min.²⁷ Interestingly, keratocytes exhibited ten times slower flow rates at the leading edge,²⁸ indicating that retrograde flow strongly depends on the cell type analyzed. Actin filaments polymerizing at the tips of filopodia also undergo retrograde flow, but these flow rates are much faster compared to those found in lamellipodia,²⁴ as shown by bleaching experiments in chick embryo fibroblasts with flow rates approximately two-fold faster in filaments derived from filopodia compared to flow rates measured within the lamellipodium. These flow rates of approximately 1.3 µm/min were similar to those found for filopodia in other studies.²² Furthermore, we could show that this retrograde flow rate strongly depends on stable FAs formed behind the filopodium (Schäfer et al. this issue and Fig. 1). In the absence of these FAs, actin retrograde flow is doubled once more to rates of approximately 2.5 µm/min in filopodia. Therefore, although rates of FAs containing filopodia are still much higher than those found in lamellipodia, these rates are still slowed down indicating an effective connection between FAs and filopodial actin. These results further imply that myosin II incorporation into filopodial-originated actin bundles is responsible for enhanced retrograde flow rates in filopodia compared to rates found in the lamellipodium and that myosin II incorporation does not depend on stably adhered FAs directly behind filopodia. These data also strongly support the hypothesis that new filopodia form in front of stable lamellipodial FAs. It will be an intriguing question for future studies to analyze whether the reduced retrograde flow speeds in front of lamellipodial FAs might even be a prerequisite for efficient assembly and stable adhesion of small filopodial FXs, or perhaps even for filopodia formation in general.Taking into account all the currently known functions of filopodia, the presented results finally indicate that filopodia might be characterized best not only by one but actually two main functions. The first function is environmental sensing. Various transmembrane proteins can be involved leading to various roles for filopodia such as formation of cell-cell or cell-matrix interactions.^5,15 Although these functions in environmental sensing seem to be highly diverse, force generation along filopodial-originated actin bundles as the second function for filopodia might be of universal importance independent of the cell type that forms them. Force transmission along cell-pathogen interacting filopodia have been observed,²⁹ and the formation of adherens junctions after filopodia mediated cell-cell interaction is also a cell force dependent process.⁵ Therefore, these observations fit well to the currently presented data by Schäfer et al. (this issue) proving the importance of filopodia-dependent cell matrix interactions in cell force generation in the direction of migration (see scheme, Fig. 1).Present in almost every moving cell type, filopodia are therefore much more than just sensors for environmental conditions. In fact, these needle-like structures are the starting point for essential structures of adhesion and movement. Independent of whether they adhere stably or not, filopodia define the position of cellular adhesion sites, actin bundles, cell force generation and application, and, finally, the new filopodia to be formed.     

Arabidopsis LIM Proteins: A Family of Actin Bundlers with Distinct Expression Patterns and Modes of Regulation

Recently, a number of two LIM-domain containing proteins (LIMs) have been reported to trigger the formation of actin bundles, a major higher-order cytoskeletal assembly. Here, we analyzed the six Arabidopsis thaliana LIM proteins. Promoter-β-glucuronidase reporter studies revealed that WLIM1, WLIM2a, and WLIM2b are widely expressed, whereas PLIM2a, PLIM2b, and PLIM2c are predominantly expressed in pollen. LIM-green fluorescent protein (GFP) fusions all decorated the actin cytoskeleton and increased actin bundle thickness in transgenic plants and in vitro, although with different affinities and efficiencies. Remarkably, the activities of WLIMs were calcium and pH independent, whereas those of PLIMs were inhibited by high pH and, in the case of PLIM2c, by high [Ca²⁺]. Domain analysis showed that the C-terminal domain is key for the responsiveness of PLIM2c to pH and calcium. Regulation of LIM by pH was further analyzed in vivo by tracking GFP-WLIM1 and GFP-PLIM2c during intracellular pH modifications. Cytoplasmic alkalinization specifically promoted release of GFP-PLIM2c but not GFP-WLIM1, from filamentous actin. Consistent with these data, GFP-PLIM2c decorated long actin bundles in the pollen tube shank, a region of relatively low pH. Together, our data support a prominent role of Arabidopsis LIM proteins in the regulation of actin cytoskeleton organization and dynamics in sporophytic tissues and pollen.     

Interior decoration: Tropomyosin in actin dynamics and cell migration

Cell migration and invasion requires the precise temporal and spatial orchestration of a variety of biological processes. Filaments of polymerized actin are critical players in these diverse processes, including the regulation of cell anchorage points (both cell-cell and cell-extracellular matrix), the uptake and delivery of molecules via endocytic pathways and the generation of force for both membrane protrusion and retraction. How the actin filaments are specialized for each of these discrete functions is yet to be comprehensively elucidated. The cytoskeletal tropomyosins are a family of actin associating proteins that form head-to-tail polymers which lay in the major groove of polymerized actin filaments. In the present review we summarize the emerging isoform-specific functions of tropomyosins in cell migration and invasion and discuss their potential roles in the specialization of actin filaments for the diverse cellular processes that together regulate cell migration and invasion.Key words: tropomyosin, actin, migration, invasion, cytoskeleton, actin dynamics, adhesionActin is the most abundant protein in eukaryotic cells and is critical for maintaining structural integrity. The polymerization of globular (G)-actin monomers forms actin filaments (F-actin),¹ which play a role in diverse and complex cellular functions including intercellular transport of organelles and vesicles,^2,3 cytokinesis,⁴ apoptosis⁵ and cell motility.⁶ Intricate details describing the molecular scale interactions between regulatory proteins and actin have been extensively investigated but the mechanistic control of diverse actin filament functions remain largely unclear. Recent improvements in analysis techniques⁷ and the use of physiologically relevant models of 3D cell culturing⁸ have now begun to reveal mechanisms of actin cytoskeleton regulation. Accruing evidence suggests that the actin decorating protein tropomyosin is a key regulator of actin filament specialization. Of particular interest is the impact that tropomyosin regulation has on actin filament activity during cell migration and invasion that underpins immunological cell homing, development, wound healing and metastasis.     

Regulation of MAPK signaling and cell death by MAPK phosphatase MKP2

Mitogen-activated protein kinase (MAPK) pathways play crucial roles in developmental and adaptive responses. Depending on the stimulus, MAPK activation regulates a wide variety of plant cell responses, such as proliferation, differentiation and cell death, which normally require precise spatial and temporal control. In this context, protein phosphatases play important roles by regulating the duration and magnitude of MAPK activities. During infection by non-host and incompatible host microorganisms, MAPK activity can promote a local cell death mechanism called hypersensitive response (HR), which is part of the plant defence response. HR-like responses require sustained MAPK activity and correlate with oxidative burst. We recently showed that MAPK phosphatase MKP2 positively controls biotic and abiotic stress responses in Arabidopsis. MKP2 interacts with MPK6 in HR-like responses triggered by fungal elicitors, suggesting that MKP2 protein is part of the mechanism involved in MAPK regulation during HR. Here we discuss the interplay of MAPK and MKP2 phosphatase signaling during cell death responses elicited by host-pathogen interactions.Key words: Arabidopsis, hypersensitive response (HR), MAPK, MPK6, MKP2, ROSDifferent studies have identified conserved components of MAPK pathways in plants and have provided evidence that MAPK signaling regulates a wide variety of plant biological responses.¹ For example, MAPK signaling is required for the regulation of stomatal functions,^2–4 hormone signaling^5,6 and innate immunity responses.^7–9 An increasing number of reports indicate that plant MAPKs, in particular tobacco SIPK/Ntf4 and WIPK and their Arabidopsis orthologs, MPK6 and MPK3, are converging points for signals elicited by different pathogens and play regulatory roles in disease responses.¹⁰One of the most efficient and immediate immune responses dependent on MAPK signaling is a mechanism of cell death called hypersensitive response (HR). HR is a rapid, localized cell death process at the site of pathogen infection, which is associated with specific molecular effects such as the generation of reactive oxygen species (ROS) and protein phosphorylation.¹¹ The best evidence implicating MAPK activity in HR comes from gain-of-function studies overexpressing SIPK/Ntf4 and WIPK in tobacco leaves. In these experiments, activation of SIPK/Ntf4 kinases efficiently induces HR-like cell death,^12,13 but the absence of endogenous WIPK function causes delayed induction of this HR phenotype, suggesting that WIPK activity facilitates or potentiates the SIPK signal.¹⁴ Similarly, overexpression analyses of Arabidopsis MPK3 and MPK6 proteins, either alone or co-expressed with activated upstream regulators (MKK proteins), also triggers a cell death phenotype,¹⁵ suggesting a coordinated role of MKK/MAPK signaling modules in HR.¹⁵ Thus, the involvement of MAPK activities such as SIPK/MPK6 in HR cell death responses is supported by different studies; however their regulation by phosphatases remains less understood.The main regulators of MAPKs are specific phosphatases belonging to various families, including PP2C Ser/Thr phosphatases, Tyr phosphatases (PTPs) or dual specificity phosphatases (DSPs) such as the MAPK phosphatase (MKP) subgroup.^16,17 In general, dephosphorylation of MAPKs inactivates their function in many metabolic, developmental or adaptive responses. In the context of HR, we have recently shown that Arabidopsis MKP phosphatase MKP2 interacts with MPK6 in the response triggered by fungal elicitors. In particular, co-expression of MPK6 and MKP2 proteins in infected tobacco leaves significantly attenuates the cell death phenotype produced by expressing MPK6 alone, suggesting that MKP2 negatively regulates MAPK activities in this process.¹⁸     

AtAGP18, a lysine-rich arabinogalactan protein in Arabidopsis thaliana,functions in plant growth and development as a putative co-receptor for signal transduction

Arabinogalactan-proteins (AGPs) are a class of hyperglycosylated, hydroxyproline-rich glycoproteins that are widely distributed in the plant kingdom. AtAGP17, 18 and 19 are homologous genes encoding three classical lysine-rich AGPs in Arabidopsis. We observed subcellular localization of AtAGP18 at the plasma membrane by expressing a translational fusion gene construction of AtAGP18 attached to a green fluorescent protein (GFP) tag in Arabidopsis plants. We also overexpressed AtAGP18 without the GFP tag in Arabidopsis plants, and the resulting transgenic plants had a short, bushy phenotype. Here we discuss putative roles of AtAGP18 as a glycosylphosphatidylinositol (GPI)-anchored protein involved in a signal transduction pathway regulating plant growth and development.Key words: Arabidopsis thaliana, arabinogalactan-proteins, co-receptor, glycosylphosphatidylinositol, lipid rafts, overexpressionArabinogalactan-proteins (AGPs) are plant cell surface glycoproteins or proteoglycans which are thought to play important roles in various aspects of plant growth and development, such as somatic embryogenesis, cell proliferation and elongation, pattern formation and hormone signaling.¹ The lysine-rich classical AGP subfamily in Arabidopsis contains three members: AtAGP17, 18 and 19. The subcellular localization of AtAGP17 and AtAGP18 was previously studied in our laboratory by expressing GFP-AtAGP17/18 fusion proteins in tobacco cell cultures.^2,3 In a recent report, we used Arabidopsis plants to overexpress GFP-AtAGP17/18/19 fusion proteins to observe subcellular localization of the lysine-rich AGPs in planta, in contrast to our previous plant cell culture work.⁴ Moreover, the lysine-rich AGPs alone (i.e., AtAGP17/18/19 without the GFP tag) were overexpressed in Arabidopsis plants, and only AtAGP18 overexpressors had a distinctive phenotype. This phenotype included shorter stems, more branches and less seeds, indicating a role for AtAGP18 in plant growth and development.⁴ In this addendum, we further discuss the putative biological role of AtAGP18 on a molecular level and its possible mode of action in cellular signaling.Classical AGPs are frequently predicted to have a glycosylphosphatidylinositol (GPI) anchor, which would allow for the localization of such AGPs to the outer surface of the plasma membrane. Biochemical analyses were carried out to support this hypothesis in tobacco, pear,⁵ rose⁶ and Arabidopsis.⁷ The lysine-rich classical AGPs, AtAGP17 and 18, were predicted to have a GPI anchor.⁸ To test this idea, tobacco cell cultures expressing GFP-AtAGP17/18 fusion proteins were plasmolyzed and GFP fluorescence was observed on the plasma membrane.^2,3 To corroborate this finding in planta, GFP-AtAGP17/18 were expressed in Arabidopsis plants and leaf trichome cells were plasmolyzed. Enhanced GFP fluorescence was observed at the plasma membrane of these transgenic trichome cells, indicating the presence of GFP-AtAGP17/18 at the plasma membrane.⁴ The localization of these lysine-rich classical AGPs at the plasma membrane suggests possible biological roles in sensing extracellular signals. They are likely associated with lipid rafts involved in cell signaling for the following reasons. In plants as well as animals, there are sterol-enriched, detergent-resistant plasma membrane microdomains called lipid rafts. Lipid rafts are known to be involved in signal transduction and are enriched in transmembrane receptors and GPI-anchored proteins, including AGPs.^9–11 The accumulation of these proteins in such microdomains may allow for interactions between these proteins in sensing extracellular signals which lead to various intracellular events. Interestingly, a recent study shows that lipid rafts from hybrid aspen cells contain callose synthase and cellulose synthase, and these enzymes are active since in vitro polysaccharide synthesis by the isolated detergent-resistant membranes was observed. These results demonstrate that lipid rafts are involved in cell wall polysaccharide biosynthesis.¹² In addition, an Arabidopsis pnt mutant study shows GPI-anchored proteins are required in cell wall synthesis and morphogenesis.¹³ These observations, coupled with previous observations that cellulose synthases as well as AGPs interact with microtubules, suggest that AGPs in lipid rafts may have a role in signal events, including those regulating cellulose and/or callose biosynthesis or deposition.^14,15To examine the role of LeAGP-1, a lysine-rich AGP in tomato, transgenic tomato plants were produced which expressed GFP-LeAGP-1 under the control of the cauliflower mosaic virus 35S promoter.¹⁶ The tomato LeAGP-1 overexpressors and Arabidopsis AtAGP18 overexpressors both have a bushy phenotype similar to transgenic tobacco plants overproducing cytokinins.^4,16,17 Cytokinins are an important class of plant hormones involved in many plant growth and development processes, such as cell growth and division, differentiation and other physiological processes.¹⁸ Therefore, Sun et al. proposed that LeAGP-1 might function in concert with the cytokinin signal transduction pathway.¹⁶ Since the overexpression phenotypes of AtAGP18 are similar to those of LeAGP-1, AtAGP18 is also likely associated with the cytokinin signal transduction pathway. The prevailing model for cytokinin signaling in Arabidopsis is similar to the two-component system in bacteria and yeast. In this model, the cytokinin receptor contains an extracellular domain, a kinase domain and a receiver domain. When the cytokinin receptor senses cytokinin signals, it auto-phosphorylates at a His residue in the kinase domain. The phosphoryl group is then transferred to an Asp residue in the receiver domain. Subsequently, the phosphoryl group is transferred to a His residue in the histidine phosphotransfer protein (Hpt) and the Hpt translocates to the nucleus and transfers the phosphoryl group to an Asp residue in a downstream response regulator to activate it.¹⁹ This model is consistent with our hypothesis since the cytokinin receptor in this model is a receptor kinase located in the plasma membrane with an extra-cellular domain that can potentially interact with AtAGP18. AtAGP18 may function as a co-receptor that first binds to cytokinins, then either directly interacts with cytokinin receptors or brings the cytokinins to cytokinin receptors in the plasma membrane. The first scenario is analogous to the interaction of contactin and contactin-associated protein (Caspr) in neurons. In this model, contactin is a GPI-anchored protein on the cell surface that binds to signal molecules and interacts with the transmembrane receptor Caspr to transmit signals to the cell interior.²⁰ The second scenario is analogous to fibroblast growth factor (FGF) signal activation in which heparan sulfate proteoglycans bind to FGF molecules and bring them to the FGF receptor.²¹Based on all the above observations and findings, a hypothetical model for AtAGP18 function is proposed in Figure 1. The model shows AtAGP18 located on the outer surface of the plasma membrane in lipid rafts where it could act as a co-receptor to sense extracellular signals (such as cytokinin) and interact with transmembrane proteins, possibly receptor kinases or ion channels, in the lipid rafts to initiate signaling by triggering various intracellular events. Interestingly, receptor tyrosine kinases and ion channels are known to be present in lipid rafts.^9,22 Moreover, AGPs are likely associated with ion channels since addition of the AGP-binding reagent Yariv phenylglycoside resulted in elevated cytoplasmic calcium concentrations in tobacco cells and lily pollen tubes.^15,23,24 Clearly, additional work will be required to verify such a model, and to better understand how AtAGP18 might sense extracellular signals and interact with the transmembrane proteins in the lipid rafts.Open in a separate windowFigure 1Model for atAGP18 functioning in cellular signaling to control plant growth and development. In this model, lipid rafts are enriched in glycosphingolipids, sterols, transmembrane proteins (such as receptors, receptor kinases and ion channel proteins) and GPI-anchored proteins including AtAGP18. (a) AtAGP18 acts as a co-receptor by binding to signaling molecules and directly interacting with transmembrane proteins in the lipid rafts. (B) AtAGP18 acts as a co-receptor by binding to signaling molecules and bringing the signaling molecules to transmembrane proteins in the lipid rafts. Upon activation by the extracellular signals, the transmembrane proteins initiate signaling and lead to various intracellular events (e.g., phosphorylation similar to the two-component signaling system, influx of calcium ions). The different components of the AtAGP18 molecule and the various lipid components of lipid rafts and plasma membrane are shown in the boxed inset. Hpt, histidine phosphotransfer protein.     

A Novel MAPKK Involved in Cell Death and Defense Signaling

A high-throughput in planta overexpression screen of a Nicotiana benthamiana cDNA library identified a mitogen activated protein kinase kinase (MAPKK), NbMKK1, as a potent inducer of hypersensitive response (HR)-like cell death. NbMKK1-mediated cell death was attenuated in plants whereby expression of NbSIPK, an ortholog of tobacco SIPK and Arabidopsis AtMPK6, was knocked down by virus-induced gene silencing (VIGS), suggesting that NbMKK1 functions upstream of NbSIPK. In accordance with this result, NbMKK1 phosphorylated NbSIPK in vitro, and furthermore NbMKK1 and NbSIPK physically interacted in yeast two-hybrid assay. VIGS of NbMKK1 in N. benthamiana resulted in a delay of Phytophthora infestans INF1 elicitin-mediated HR as well as in the reduction of resistance against a non-host pathogen Pseudomonas cichorii. Our data of NbMKK1, together with that of LeMKK4,¹ demonstrate the presence of a novel defense signaling pathway involving NbMKK1/LeMKK4 and SIPK.Key Words: MAPK, defense, cell death, in planta screenMitogen activated protein kinase (MAPK) cascades are highly conserved signaling pathways in eukaryotes, comprising three tiered classes of protein kinase, MAPKKK (MAPKK kinase), MAPKK and MAPK, that sequentially relay phosphorylation signals.² The Arabidopsis genome carries genes for 20 MAPKs, 10 MAPKKs³ and more than 25 MAPKKKs.⁴ In plants, MAPK signaling is known to function in various biotic^4,5 and abiotic⁶ stress responses and cytokinesis.⁷ In defense signaling, extensive research has been carried out for two tobacco MAPKs, SIPK⁸ (salicylic-acid-induced protein kinase; hereafter designated as NtSIPK) and WIPK⁹ (wound-induced protein kinase = NtWIPK), and their orthologs in Arabidopsis¹⁰ (AtMPK6 and ATMPK3, respectively), partly because kinase activities of these two MAPKs are easy to detect by an in gel kinase assay using myeline basic protein (MBP) as substrate.¹¹ Both NtSIPK and NtWIPK are activated by the interaction between host resistance (R)- gene and cognate avirulence gene of pathogen^11,12 and elicitor perception by host cells.^13,14 Shuqun Zhang and his group showed that an upstream kinase of both NtSIPK and NtWIPK is NtMEK2.¹⁵ Transient overexpression of constitutively active NtMEK2 caused phosphorylation of NtSIPK and NtWIPK, resulting in rapid HR-like cell death in tobacco leaves.¹⁵ Later, the same lab showed that overexpression of NtSIPK alone also caused HR-like cell death.¹⁶ The downstream target proteins of NtSIPK and AtMPK6 are being identified and include 1-aminocyclopropane-1-carboxylic acid sythase-6 (ACS-6).^17,18 Although recent studies identified another MAPK cascade (NtMEK1 → Ntf6) involved in defense responses^19,20 we can still say that the current research focus of MAPK defense signaling centers around the cascade comprising [NtMEK2→ NtSIPK/NtWIPK→ target proteins] of tobacco and its orthologous pathways in other plant species.In an effort to search for plant genes involved in HR-like cell death, we have been employing a high-throughput in planta expression screen of N. benthamiana cDNA libraries. In this experimental system, a cDNA library was made in a binary potato virus X (PVX)-based expression vector pSfinx.²¹ The cDNA library was transferred to Agrobacterium tumefaciens, and 40,000 of the bacterial colonies were individually inoculated by toothpicks onto leaf blades of N. benthamiana leaves. The phenotype around the inoculated site was observed 1–2 weeks following the inoculation. This rapid screen identified 30 cDNAs that caused cell death after overexpression, including genes coding for ubiquitin proteins, RNA recognition motif (RRM) containing proteins, a class II ethylene-responsive element binding factor (EREBP)-like protein²² and a MAPKK protein (this work). Such an in planta screening technique has been used before for the isolation of fungal²¹ and oomycete^23,24 elicitors and necrosis inducing genes, but not for isolation of plant genes. Overexpression screening of cDNA libraries is a common practice in prokaryotes, yeast and amimal cells,^25,26 so it is a surprise that this approach has not been systematically applied in plants. Given its throughput, we propose that this virus-based transient overexpression system is a highly efficient way to isolate novel plant genes by functional screen.²⁷ Since overexpression frequently causes non-specific perturbation of signaling, genes identified by overexpression should be further validated by loss-of-function assays, for instance, VIGS.²⁸Overexpression of the identified MAPKK gene, NbMKK1, triggered a rapid generation of H₂O₂, followed by HR-like cell death in N. benthamiana leaves (this work). NbMKK1-GFP fusion protein overexpression also caused cell death, and curiously NbMKK1-GFP was shown to localize consistently in the nucleus. Sequence comparison classified NbMKK1 to the Group D of MAPKKs about which little information is available. So far, a MAPKK, LeMKK4, from tomato belonging to the Group D MAPKKs, was shown to cause cell death after overexpression.¹ Based on amino acid sequence similarity and phylogenetic analyses, LeMKK4 and NbMKK1 seem to be orthologs. To see whether NbMKK1 transduces signals through SIPK and WIPK, we performed NbMKK1 overexpression in N. benthamiana plants whereby the expression of either NbSIPK or NbWIPK (WIPK ortholog in N. benthamiana) was silenced by VIGS. NbMKK1 did not induce cell death in NbSIPK-silenced plants, suggesting that the NbMKK1 cell death signal is transmitted through NbSIPK. Indeed, NbMKK1 phosphorylated NbSIPK in vitro, and NbMKK1 and NbSIPK physically interacted in yeast two-hybrid assay. These results suggest that NbMKK1 interacts with NbSIPK, most probably with its N-terminal docking domain, and phosphorylates NbSIPK in vivo to transduce the cell death signal downstream.NbMKK1 exhibits constitutive expression in leaves. To determine the function of NbMKK1 in defense, we silenced NbMKK1 by VIGS, and such plants were challenged with Phytophthora infestans INF1 elicitin²⁹ and Pseudomonas cichorii, a non-host pathogen. INF1-mediated HR cell death was remarkably delayed in NbMKK1-silenced plants. Likewise, plant defense against P. cichorii was compromised in NbMKK1-silenced plants. These results indicate that NbMKK1 is an important component of signaling of INF1-mediated HR and non-host resistance to P. cichorii.Together, our analyses of NbMKK1 and independent work from Greg Martin''s lab on LeMKK4¹ suggest that a Group D MAPKK, NbMKK1/LeMKK4, functions upstream of SIPK and transduces defense signals in these solanaceous plants (Fig. 1). In plants as well as in other eukaryotes, it is common that kinases have multiple partners. The work on these kinases fits this concept. A single MAPK (e.g., SIPK) is phosphorylated by multiple MAPKKs (e.g., NtMEK2 and NbMKK1), and a single MAPKK (e.g., NtMEK2) can phosphorylate multiple MAPKs (e.g., NtSIPK and NtWIPK).Open in a separate windowFigure 1Defense signaling through NbMKK1/LeMKK4. Two defense signal pathways involving NtMEK2 (indicated as MEK2) → WIPK/SIPK and NtMEK1(indicated as MEK1) → Ntf6 are well documented. By our and Pedley and Martin''s¹ works, another novel MAPKK, NbMKK1/LeMKK4 was demonstrated to participate in defense signaling by phosphorylation of SIPK.     

ARG1 and ARL2 form an actin-based gravity-signaling chaperone complex in root statocytes?

Plants are acutely sensitive to the directional information provided by gravity. They have evolved statocytes, which are specialized cells that sense gravity and, upon integration of the corresponding information with that of other environmental stimuli, control the growth behavior of their organs. The cellular mechanisms that allow statocytes to sense and transduce gravitational information likely involve detecting the sedimentation of, or the tension/pressure exerted by, starch-filled amyloplasts—the presumptive statoliths—within their cytoplasm. Gravity signaling in root statocytes controls the direction of transport of signaling compounds, especially auxin, across the root cap, establishing a lateral gradient that is transmitted to cells in the elongation zone and results in gravitropic curvature. The Arabidopsis J-domain proteins ARG1 and ARL2 function as gravity-signal transducers in root statocytes. In the January issue of The Plant Journal, we reported that ARG1 and ARL2 function non-redundantly in a common gravity signaling pathway required for accumulation of the auxin efflux facilitator PIN3 on the new bottom side of statocytes following gravity stimulation, and lateral redistribution of auxin toward the new lower flank of stimulated roots. Here we present data suggesting that ARG1 physically associates with ARL2, the J-domain co-chaperone HSC70, and actin in vivo. We briefly discuss potential mechanisms by which ARG1 and ARL2 might function in gravity signaling in light of this information.Key words: gravitropism, statocytes, actin, HSC70/HSP70, auxinGravitropic growth in plants requires the ability to sense an organ''s orientation within the gravity field, transduce that information into a biochemical signal, and mount a differential cellular-elongation response that corrects growth to follow a defined gravity set point angle. Gravity sensation in roots occurs mainly in statocytes of the root cap columella, either via detection of the position or movement of, or the pressure exerted by, amyloplasts within these cells, or by perceiving the pressure exerted by the statocyte''s protoplast within its cell wall.^1,2 Several models address the mechanism by which statocytes use amyloplasts as gravity susceptors. One model postulates that gravity signal transduction initiates when sedimenting amyloplasts promote the opening of mechano-sensitive ion channels, either directly or through interaction with the actin cytoskeleton. Alternatively, signal transduction may initiate upon sedimentation, when amyloplast-borne ligands interact with receptors located on sensitive structures within the statocytes.¹ Nevertheless, the activated pathway promotes a fast and transient cytoplasmic alkalinization and redistribution of the auxin efflux carrier PIN3 to the lower membrane of the statocytes. These changes contribute to lateral, downward, redistribution of auxin across the cap, resulting in the formation of a lateral auxin gradient that, upon transmission to the elongation zone, promotes tip curvature (reviewed in ref. ¹).Along with experiments demonstrating the importance of amyloplasts in gravity susception, genetic analysis in Arabidopsis has identified ARG1 and ARL2 as gravity signal transducers in root and hypocotyl statocytes.^3–6 ARG1 and ARL2 function in a gravity-signaling pathway that links gravistimulation to the accumulation of PIN3 within the PM at the lower side of the statocytes and a redistribution of auxin across the cap.^4–6 ARG1 and ARL2 are membrane-associated proteins. While ARG1 localizes throughout the endosomal/secretory pathway, ARL2 associates primarily with the PM, and both proteins are found at the cell plate during cytokinesis.^4,6 ARG1 and ARL2 contain J-domains near their N-termini. J-domain proteins typically function as molecular co-chaperones by interacting with HSC70, coupling substrate binding to HSC70 ATPase activity.¹⁰ A direct role for HSC70 in gravity signaling has not been reported, though proteomics approaches have identified cytosolic HSC70 isoforms as gravity-regulated proteins within Arabidopsis root tips.^7,8The C-termini of ARG1 and ARL2 contain putative coiled-coil domains with similarity to predicted coiled-coils found in proteins that interact with the cytoskeleton, including Rho-associated kinases, myosin, heavy chain kinesins and tropomyosins.³ Interestingly, a biochemical fractionation experiment detected possible interaction between ARG1 and polymerized actin in plant extracts.⁴ Furthermore, proteomic analyses of root gravitropism also identified actin as a gravity-regulated protein^7,8 whereas other studies strongly suggest a role for the actin cytoskeleton in the modulation of gravity signaling in roots.⁹To investigate possible in vivo interactions between ARG1, ARL2, HSC70 and/or actin, we generated transgenic plants expressing GFP fusions to ARG1 or ARL2, in the arg1-2 or arl2-3 mutant backgrounds (Ws alleles), and used them in co-immunoprecipitation experiments. All constructs were tested for functional rescue prior to use. Functional GFP fusions to ARG1 or ARL2 were isolated from plant extracts using immobilized anti-GFP antibodies. Initial experiments indicated that both N and C-terminal GFP fusions to ARG1 and C-terminal fusions to ARL2 could be purified by this method (Fig. 1A). However, we consistently noticed lower relative abundance of ARL2-GFP purified from plants compared to ARG1-GFP, even when both were expressed under the same promoter (Fig. 1A). This is consistent with lower fluorescence intensity and fusion protein abundance of ARL2-GFP compared to ARG1-GFP in the many transgenic lines analyzed (data not shown).Open in a separate windowFigure 1HSC70, actin and ARL2 co-purify with ARG1 from plant extracts. Total protein extracts from four-week-old liquid grown plants of the indicated genotypes (top) were affinity-purified using anti-GFP antibodies (gift of Richard Vierstra, UW-Madison) cross-linked to Protein A-Sepharose beads and detected with the indicated antibodies. In (A) the affinity-purified eluate was probed with anti-GFP antibody to indicate the presence of both full-length GFP fusions and breakdown products. In (B) western blots of total protein extracts (Input, In) and affinity-purified eluates (E) were probed with anti-HSC70 (diluted 1:10,000; Stressgen, San Diego, CA), monoclonal anti-actin (diluted 1:1000; clone C4 from Chemicon Int., Temecula, CA), or anti-a-tubulin antibodies (Sigma, St Louis, MO). In (C) western blots of affinity-purified eluates (top) were probed with two anti-ARG1 antibodies raised in rabbits against a bacterially expressed His6-ARG1 protein fusion (anti-ARG1 I) and against a peptide corresponding to the 15 C-terminal residues of ARG1 (anti-ARG1 II), both used at 1:1000 dilutions, as well as with a polyclonal anti-ARL2 antibody raised in rabbits against a synthetic ARL2 peptide, used at 1:3000 dilution. Note that anti-ARG1 II did not recognize the ARG1-GFP protein probably because the corresponding C-terminal epitope was masked in this C-terminal fusion. A western blot of total protein extracts used for affinity purification (In, bottom) was probed with anti-HSP70 antibody (gift of Elizabeth Craig, UW-Madison; 1:20,000 dilution) to show relative amounts of protein prior to extraction. Experimental procedures. Plants expressing ARG1::GFP-ARG1 and 35S::ARL2-GFP are described elsewhere (reviewed in refs. ⁴ and ⁶, respectively). Plants expressing 35S::ARG1-GFP were generated by transformation with a T-DNA derived from the binary vector pK7FWG2²⁵ containing the ARG1 cDNA, whereas plants expressing AtMDR1::GFP were a gift from Edgar Spalding (UW-Madison). For immunoprecipitation, anti-GFP serum was cross-linked to Protein A Sepharose (Amersham, Piscataway, NJ) using the cross-linking reagent disuccinimidyl suberate according to the manufacturer''s recommendations (Pierce, Rockford, Ill.). Extracts from liquid-grown plants were generated by grinding in 1:1 (vol:wt) ice cold grinding buffer [20 mM Tris, 1 mM EDTA, 100 mM NaCl, 0.1% Triton X-100 pH7.5, containing 1:100 dilution of protease inhibitor cocktail (Sigma, or Calbiochem, San Diego, CA) and 1 mM phenylmethylsulphonyl fluoride] in a mortar, then adding more buffer to a ratio of 4:1 (vol:wt). Homogenates were cleared by filtering through Miracloth (Calbiochem) and spinning twice at 13,000 g for 15 min each to pellet cellular debris. Protein concentration in the resulting supernatant was then quantified by a modified Lowry assay, using BSA as a standard. Extracts were gently rocked with anti-GFP beads for one hour at room temperature. The beads were washed extensively (five times with ≥5× bead vol.) in extraction buffer, and the bound protein was eluted with low pH (0.1 M glycine pH 2.5) into a small amount of neutralizing 1 M tris buffer (pH 8.0), precipitated with TCA, separated by SDS-PAGE, blotted to PVDF membrane (Millipore, Billerica, MA), and probed with the appropriate antibodies. Positive signals were detected using HRP-conjugated secondary antibodies at 1:20,000 (Sigma).Proteins that interact with ARG1 or ARL2 are expected to bind to the anti-GFP-coated beads in this assay, along with their GFP-ARG1 or GFP-ARL2 partner, and to co-elute with them upon washing the beads with a low-pH solution. Their identity can be revealed by western blot analysis of bead eluates, using specific antibodies. Figure 1B shows that HSC70 co-purifies with ARG1-GFP and GFP-ARG1 when extracts from plants expressing these fusions are tested in this assay. However, HSC70 was not present in the eluate when plants expressing GFP alone under the control of the CaMV35S or AtMDR1 ectopic promoters were tested (Fig. 1B). Interestingly, we did not detect HSC70 in bead eluates when ARL2-GFP-expressing plants were subjected to this co-immunoprecipitation assay, suggesting that ARG1 and ARL2 differ in their affinity for HSC70. It should however be noted that the low amount of ARL2-GFP precipitated in these experiments does not allow for direct comparison (Fig. 1B). These results indicate an in vivo interaction between ARG1 and HSC70, suggesting that ARG1 is a bona fide co-chaperone.Figure 1B also shows that actin is present specifically in immunocomplexes containing ARG1. In a parallel experiment, ARL2-GFP precipitates did not contain detectable actin (Fig. 1B). However, here again, the low amount of ARL2 in these precipitates does not allow direct comparison to ARG1. Together these data suggest that ARG1 physically associates with actin in vivo, even though they do not address whether this association involves monomeric or polymerized actin. Interestingly, the biochemical fractionation experiments reported by Boonsirichai et al. (2003) suggested that some ARG1 might associate with polymerized actin in vivo.Plants with mutations in ARG1, ARL2, or both, result in very similar gravity signaling defects, demonstrating that ARG1 and ARL2 function as non-redundant members of the same signaling pathway.⁵ One interpretation of this data is that ARG1 and ARL2 function in the same signaling complex, the presence of both being required for complex activity in gravity signaling. In support of this model, we have detected an in vivo interaction between ARG1 and ARL2 by immunoprecipitation. Figure 1C shows that endogenous ARG1 (∼45 kDa) co-immunoprecipitates with ARL2-GFP from extracts derived from plants expressing 35S::ARL2-GFP, but not from extracts derived from negative control plants. We were not surprised by the failure to detect ARG1 in precipitates from plants expressing ARL2-GFP under the control of its endogenous promoter (ARL2::ARL2-GFP). Indeed, its expression is restricted to a few cells—therefore, the protein is present at low abundance in whole-plant extracts.⁶ Similarly, native ARL2 was not detected in ARG1 immunoprecipitates likely due to the low level of endogenous ARL2 expression. Together, these results suggest that ARG1 and ARL2 associate in a complex in vivo, consistent with the function of these proteins in a common signaling pathway.⁵The sub-cellular localization of a putative ARG1- and ARL2-containing complex is unknown, but is likely associated with the PM of root statocytes, where ARL2-GFP localizes in cells expressing ARL2::ARL2-GFP.⁶ Such a complex may serve as a membrane-associated signaling module that recruits cytosolic HSC70 to a membrane site in a gravity-regulated manner. In agreement with this model, we found that HSP70 increases in abundance in a biochemical fraction that includes membrane-associated proteins extracted from Arabidopsis root tips within 10 min of a gravistimulus.⁸ A similar recruitment model has been proposed for J-protein function in several HSC70-mediated processes, including the regulation of vesicle dynamics and signaling by HSC70.¹⁰ In animal systems, HSC70 activity is required for several steps in clatherin-dependent vesicle recycling,¹¹ a process that appears to mediate PIN protein trafficking¹². Recruitment of HSC70 activity to clatherin-coated vesicles is mediated mainly by the J-domain protein auxillin, which has paralogs other than ARG1 and ARL2 in plants.¹⁰ HSC70 function is also required for Ca²⁺-induced exocytosis in Drosophila, and depends on the J-domain-containing cysteine-string protein, apparent homologs of which are absent from plants.¹³ GRV2, a locus required for normal shoot tropisms, encodes a membrane-associated J-domain protein homologous to animal RME-8.¹⁴ RME-8 is a membrane-associated J-domain protein that interacts with cytosolic HSC70 and is required for endocytosis and endosomal trafficking in D. melanogaster and C. elegans.^15,16 GRV2 appears to regulate dynamics of late vacuolar endosomes,¹⁷ which is important for gravity signaling in shoots but not roots.¹⁸ GRV2/RME-8, auxillin, and cysteine-string proteins are each specific for endosomal trafficking involving particular molecules or vesicles, thus illustrating the specificity that J-domain proteins can confer to HSC70-mediated processes.One possibility is that the putative ARG1/ARL2 complex recruits HSC70 to alter actin dynamics in response to gravistimulation. The actin network in root statocytes appears to be intimately associated with the mechanism of gravity sensation, as treatments with low concentrations of the actin-depolymerizing drug Latrunculin B enhances gravity signaling events including amyloplast sedimentation, alkalinization of the statocyte cytoplasm, with subsequent enhancement of both auxin redistribution and organ curvature.⁹ arg1-2 mutants are deficient in gravity-induced statocyte alkalinization and auxin redistribution, suggesting that ARG1 and microfilaments regulate this gravity signaling response.⁴The lateral polarization of the statocytes upon gravistimulation likely involves dramatic changes in vesicle trafficking. Mechanisms that lead to accumulation of PIN3 upon the new bottom side of root statocytes are unknown, but require ARG1 and ARL2 either directly or as upstream signaling components. The prominent localization of ARG1 and ARL2 to the cell plate during cytokinesis suggests that they are present in areas of intense vesicle dynamics. The recent discovery that members of an Arabidopsis retromer complex are required for cell polarization, PIN trafficking, and gravitropic growth suggests they may function in redirection of auxin following gravistimulation.^19,20 Animal homologs of these retromer components have been identified in association with clathrin-coated vesicles,²¹ and disruption of Drosophila Vps35 or human SNX9 affect endocytosis and actin organization.^22,23 The mechanisms of clathrin-dependent endocytosis are unresolved, but appear to depend on dynamic cortical microfilaments in animals and yeast suggesting that similar mechanisms may function in plants.²⁴ ARG1 and ARL2 may mediate statocyte polarization via regulating actin and/or vesicle dynamics. Of course, we have not yet excluded the possibility that ARG1 and ARL2 might, in fact, transduce the gravity signal in an HSC70 and actin-independent manner through interactions with yet unidentified molecules. In this case, the interactions identified in this study would be relevant to other, yet uncharacterized, functions potentially associated with ARG1 and ARL2 in plants.^4,5 Work aimed at identifying additional ARG1 and ARL2-interacting proteins is underway.     

Why have chloroplasts developed a unique motility system?

Organelle movement in plants is dependent on actin filaments with most of the organelles being transported along the actin cables by class XI myosins. Although chloroplast movement is also actin filament-dependent, a potential role of myosin motors in this process is poorly understood. Interestingly, chloroplasts can move in any direction and change the direction within short time periods, suggesting that chloroplasts use the newly formed actin filaments rather than preexisting actin cables. Furthermore, the data on myosin gene knockouts and knockdowns in Arabidopsis and tobacco do not support myosins'' XI role in chloroplast movement. Our recent studies revealed that chloroplast movement and positioning are mediated by the short actin filaments localized at chloroplast periphery (cp-actin filaments) rather than cytoplasmic actin cables. The accumulation of cp-actin filaments depends on kinesin-like proteins, KAC1 and KAC2, as well as on a chloroplast outer membrane protein CHUP1. We propose that plants evolved a myosin XI-independent mechanism of the actin-based chloroplast movement that is distinct from the mechanism used by other organelles.Key words: actin, Arabidopsis, blue light, kinesin, myosin, organelle movement, phototropinOrganelle movement and positioning are pivotal aspects of the intracellular dynamics in most eukaryotes. Although plants are sessile organisms, their organelles are quickly repositioned in response to fluctuating environmental conditions and certain endogenous signals. By and large, plant organelle movements and positioning are dependent on actin filaments, although microtubules play certain accessory roles in organelle dynamics.^1,2 Actin inhibitors effectively retard the movements of mitochondria,^3–6 peroxisomes,^5,7–11 Golgi stacks,^12,13 endoplasmic reticulum (ER),^14,15 and nuclei.^16–18 These organelles are co-aligned and associated with actin filaments.^{5,7,8,10–12,15,18} Recent progress in this field started to reveal the molecular motility system responsible for the organelle transport in plants.¹⁹Chloroplast movement is among the most fascinating models of organelle movement in plants because it is precisely controlled by ambient light conditions.^20,21 Weak light induces chloroplast accumulation response so that chloroplasts can capture photosynthetic light efficiently (Fig. 1A). Strong light induces chloroplast avoidance response to escape from photodamage (Fig. 1B).²² The blue light-induced chloroplast movement is mediated by the blue light receptor phototropin (phot). In some cryptogam plants, the red light-induced chloroplast movement is regulated by a chimeric phytochrome/phototropin photoreceptor neochrome.^23–25 In a model plant Arabidopsis, phot1 and phot2 function redundantly to regulate the accumulation response,²⁶ whereas phot2 alone is essential for the avoidance response.^27,28 Several additional factors regulating chloroplast movement were identified by analyses of Arabidopsis mutants deficient in chloroplast photorelocation.^29–32 In particular, identification of CHUP1 (chloroplast unusual positioning 1) revealed the connection between chloroplasts and actin filaments at the molecular level.²⁹ CHUP1 is a chloroplast outer membrane protein capable of interacting with F-actin, G-actin and profilin in vitro.^29,33,34 The chup1 mutant plants are defective in both the chloroplast movement and chloroplast anchorage to the plasma membrane,^22,29,33 suggesting that CHUP1 plays an important role in linking chloroplasts to the plasma membrane through the actin filaments. However, how chloroplasts move using the actin filaments and whether chloroplast movement utilizes the actin-based motility system similar to other organelle movements remained to be determined.Open in a separate windowFigure 1Schematic distribution patterns of chloroplasts in a palisade cell under different light conditions, weak (A) and strong (B) lights. Shown as a side view of mid-part of the cell and a top view with three different levels (i.e., top, middle and bottom of the cell). The cell was irradiated from the leaf surface shown as arrows. Weak light induces chloroplast accumulation response (A) and strong light induces the avoidance response (B).Here, we review the recent findings pointing to existence of a novel actin-based mechanisms for chloroplast movement and discuss the differences between the mechanism responsible for movement of chloroplasts and other organelles.     

Flowering time regulation by the SUMO E3 ligase SIZ1

Flowering is a developmental process, which is influenced by chemical and environmental stimuli. Recently, our research established that the Arabidopsis SUMO E3 ligase, AtSIZ1, is a negative regulator of transition to flowering through mechanisms that reduce salicylic acid (SA) accumulation and involve SUMO modification of FLOWERING LOCUS D (FLD). FLD is an autonomous pathway determinant that represses the expression of FLOWERING LOCUS C (FLC), a floral repressor. This addendum postulates mechanisms by which SIZ1-mediated SUMO conjugation regulates SA accumulation and FLD activity.Key words: SIZ1, SA, flowering, SUMO, FLD, FLCSUMO conjugation and deconjugation are post-translational processes implicated in plant defense against pathogens, abscisic acid (ABA) and phosphate (Pi) starvation signaling, development, and drought and temperature stress tolerance, albeit only a few of the modified proteins have been identified.^1–8 The Arabidopsis AtSIZ1 locus encodes a SUMO E3 ligase that regulates floral transition and leaf development.^8,9 siz1 plants accumulate substantial levels of SA, which is the primary cause for dwarfism and early short-day flowering exhibited by these plants.^1,9 How SA promotes transition to flowering is not yet known but apparently, it is through a mechanism that is independent of the known floral signaling pathways.^9,10 Exogenous SA reduces expression of AGAMOUS-like 15 (AGL15), a floral repressor that functions redundantly with AGL18.^11,12 A possible mechanism by which SA promotes transition to flowering may be by repressing expression of AGL15 and AGL18 (Fig. 1).Open in a separate windowFigure 1Model of how SUMO conjugation and deconjugation regulate plant development in Arabidopsis. SIZ1 and Avr proteins regulate biosynthesis and accumulation of SA, a plant stress hormone that is involved in plant innate immunity, leaf development and regulation of flowering time. SA promotes transition to flowering may through AGL15/AGL18 dependent and independent pathways. FLC expression is activated by FRIGIDA but repressed by the autonomous pathway gene FLD, and SIZ1-mediated sumoylation of FLD represses its activity. Lines with arrows indicate upregulation (activation), and those with bars identify downregulation (repression).siz1 mutations also cause constitutive induction of pathogenesis-related protein genes leading to enhanced resistance against biotrophic pathogens.¹ Several bacterial type III effector proteins, such as YopJ, XopD and AvrXv4, have SUMO isopeptidase activity.^13–15 PopP2, a member of YopJ/AvrRxv bacterial type III effector protein family, physically interacts with the TIR-NBS-LRR type R protein RRS1, and possibly stabilizes the RRS1 protein.¹⁶ Phytopathogen effector and plant R protein interactions lead to increased SA biosynthesis and accumulation, which in turn activates expression of pathogenesis-related proteins that facilitate plant defense.¹⁷ SIZ1 may participate in SUMO conjugation of plant R proteins to regulate Avr and R protein interactions leading to SA accumulation, which, in turn, affects phenotypes such as diseases resistance, dwarfism and flowering time (Fig. 1).Our recent work revealed also that AtSIZ1 facilitates FLC expression, negatively regulating flowering.⁹ AtSIZ1 promotes FLC expression by repressing FLD activity.⁹ Site-specific mutations that prevent SUMO1/2 conjugation to FLD result in enhanced activity of the protein to represses FLC expression, which is associated with reduced acetylation of histone 4 (H4) in FLC chromatin.⁹ FLD, an Arabidopsis ortholog of Lysine-Specific Demethylase 1 (LSD1), is a floral activator that downregulates methylation of H3K4 in FLC chromatin and represses FLC expression.^18,19 Interestingly, bacteria expressing recombinant FLD protein did not demethylate H3K4me2, inferring that the demethylase activity requires additional co-factors as are necessary for LSD1.^18,20 Together, these results suggest that SIZ1-mediated SUMO modification of FLD may affect interactions between FLD and co-factors, which is necessary for FLC chromatin modification.Despite our results that implicate SA in flowering time control, how SIZ1 regulates SA accumulation and the identity of the effectors involved remain to be discovered. In addition, it remains to be determined if SIZ1 is involved in other mechanisms that modulate FLD activity and FLC expression, or the function of other autonomous pathway determinants.     

Interplay between adhesion turnover and cytoskeleton dynamics in the control of growth cone migration

The migration of neuronal growth cones, driving axon extension, is a fascinating process which has been subject of intense investigation over several decades. Many of the key underlying molecules, in particular adhesion proteins at the cell membrane which allow for target recognition and binding, and cytoskeleton filaments and motors which power locomotion have been identified. However, the precise mechanisms by which growth cones coordinate, in time and space, the transmission of forces generated by the cytoskeleton to the turnover of adhesion proteins are still partly unresolved. To get a better grasp at these processes, we put here in relation the turnover rate of ligand/receptor adhesions and the degree of mechanical coupling between cell adhesion receptors and the actin rearward flow. These parameters were obtained recently for N-cadherin and IgCAM based adhesions using ligand-coated microspheres in combination with optical tweezers and photo-bleaching experiments. We show that the speed of growth cone migration requires both a fairly rapid adhesion dynamics and a strong physical connection between adhesive sites and the cytoskeleton.Key words: actin retrograde flow, molecular clutch, myosin, N-cadherin, IgCAMGrowth cones are motile structures at the distal extremity of axons responsible for pathfinding and neurite extension during nervous system development and repair (Fig. 1A). Growth cone advance relies on two coupled processes. First, an internal dynamics of the cytoskeletal network, with actin polymerization occurring at the leading edge, depolymerization in the central region, and myosin activity pulling on lamellipodial actin filaments.¹ These integrated mechanisms altogether result in a continuous retrograde flow of actin (Fig. 1B). This flow provides the mechanical tension that drives axonal extension, through a connection to the dynamic array of microtubules that fills the axon and invades the growth cone central domain.² Second, there is repeated formation and dissociation of transient contacts between growth cones and the extracellular matrix or adjacent cells. These contacts are mediated by trans-membrane cell adhesion molecules (CAMs), e.g. integrins,³ immunoglobulin CAMs (IgCAMs)^4,5 and cadherins,^6,7 which form specific ligand/receptor bonds with variable lifetimes. A still open question is how these two processes, i.e. actin flow and adhesion dynamic, are coordinated at the growth cone level and contribute to set migration speed. A thorough understanding of these mechanisms is important both from a fundamental perspective and for the design of new compounds to foster axon regeneration after injury.Open in a separate windowFigure 1Growth cone advance and actin flow. (A) Growth cone from a 2 DIV rat hippocampal neuron plated on N-cadherin coated glass. This growth cone moved forward at a speed of about 1 µm/min (B) Raw fluorescence image of transfected actin-GFP. (C) Sequential actin-GFP images were subtracted, giving rise to intensity variations that display the movement of newly assembled actin (black). Note the rapid retrograde movement of actin spots (arrowheads), at a velocity of several µm/min.The coupling between actin-based motility and substrate adhesion has been shown for certain adhesion molecules such as NCAM and N-cadherin to involve a “molecular clutch” (Fig. 2). This mechanism implies a direct transmission of traction forces from the cytoskeleton to the substrate through a strong physical connection between the actin flow and ligand-bound adhesion receptors.^8,9 The connection is likely provided by adaptor proteins that can make transient bridges between actin filaments and the cytoplasmic domain of adhesion molecules, i.e. α-and β-catenin in the case of N-cadherin,¹⁰ ankyrin and ezrin in the case of IgCAMs such as L1.^11–14 These purely mechanical connections can also be accompanied by signalling events such as Rac-1 activation by N-cadherin liganding¹⁵ and phosphorylation of the L1 intracellular tail that regulates binding to ankyrin.^11,12 When only few molecular bonds are formed, e.g. at low ligand density, coupling to the actin flow is not strong enough, resulting in “slippage.” In this process, transient bonds can be formed and broken repeatedly between ligand-occupied adhesion receptors and the actin network. This is how the speed of growth cone translocation usually reaches at most 1 µm/min, whereas the internal actin flow rate proceeds at a rate of several µm/min (Fig. 1A and B). Such slippage is best demonstrated by the use of optical tweezers to impose low forces on ligand-coated microspheres presented to the growth cone dorsal surface (Fig. 3A). Beads tend to move rearward as they couple to the actin flow, and then suddenly snap back into the trap center, when receptor-cytoskeleton bonds break¹⁶ (the force of optical tweezers is usually not enough to rupture ligand-receptor bonds, which remain intact at the cell surface). Thus, a step in which a nucleating cluster of adhesion receptors recruits a minimal number of intracellular partners allowing coupling to the actin flow, can be a rate-limiting factor in growth cone progression.Open in a separate windowFigure 2Molecular components involved in growth cone migration. (A) Top view diagram showing filopodia which sense the environment, a flat lamellipodium which is the site of actin dynamics and the thicker central domain and axon which contain dynamic microtubules. The plus signs are sites of actin polymerization and the minus signs indicate actin depolymerization. (B) Side view showing the life cycle of ligand/receptor adhesions.Open in a separate windowFigure 3Optical tweezers and FRAP experiments to measure ligand/receptor and receptor/cytoskeleton dynamics. (A) Optical tweezers experiments performed on ligand-coated beads placed on the growth cone dorsal surface.¹⁶ (B) The distance traveled rearward with respect to the trap center is measured. A 2 min trajectory is indicated in red. A pooled parameter called coupling index taking into account the latency for bead escape, as well as the mean velocity and lateral diffusion of the bead, measures the strength of receptor/cytoskeleton interactions. (C) FRAP experiments on membrane GFP-tagged molecules accumulated at ligand-coated microspheres having sedimented on growth cones. The fluorescence intensity is normalized to represent the receptor enrichment level at the bead contact. (D) The recovered intensity is fit by a diffusion/reaction model, which yields a collective equilibrium turnover rate of ligand/receptor bonds. In red is the average of a series of individual curves (grey).In contrast, when strong connection is formed and if the substrate is resistant enough, then the molecular clutch engages and the cell reacts. In growth cones from Aplysia bag cells, forces were imposed on microspheres coated with ApCAM (the homolog of vertebrate NCAM) using a microneedle to locally block the retrograde actin flow. This was systematically followed by a protrusion of the microtubule-rich central domain towards those stiff contacts and forward expansion of the actin-rich lamellipodium.⁹ These phenomena were later shown to be controlled by a src protein kinase.¹⁷ In the case of rat hippocampal neurons, a dramatic accumulation of actin at N-cadherin coated microspheres is observed when the latter are restrained from moving rearward by a microneedle.¹⁶ This phenomenon is mediated by a connection between N-cadherin and α-catenin, likely triggering local actin polymerization. By careful analysis of the bead trajectories at varying ligand densities and computation of the latency for bead escape when the optical trap is applied continuously, one can extract a quantitative index of receptor-cytoskeleton coupling (Fig. 3B). Overall, a strong correlation was observed between such coupling index and the velocity of growth cone migration on N-cadherin substrates, both by varying N-cadherin ligand density and by expressing mutated N-cadherin molecules, supporting the clutch concept.¹⁶ This mechanism is consistent with in vivo experiments showing that overexpression of the N-cadherin intracellular tail in retinal ganglion cells results in severely impaired axon outgrowth.¹⁸ As a negative example of the clutch model, beads coated with fibronectin (our unpublished data) or anti-α1 integrin antibodies³ couple weakly to the actin flow in growth cones while, in parallel, the migration of growth cones on fibronectin- or collagen-coated substrates is rather limited.^6,19Molecular mechanisms parallel to the “clutch” can also be involved in growth cone migration. For example, IgCAM adhesions can not only couple to the rearward actin flow but also to static components of the cytoskeleton. Indeed, a 30% fraction of TAG-1 or anti-L1 coated beads can stay immobile on the growth cone surface.^12,20 These contrasting behaviors are likely mediated by interactions between the IgCAM intracellular domain and different binding partners (ankyrin vs. ERM),¹³ and may be responsible for the pauses which alternate with phases of growth cone advance. Also, homophilic adhesions between molecules of cadherin-11 couple very weakly to the actin flow, but promote substantial growth cone migration when cadherin-11 is presented as a substrate. This effect seems to be mediated by an independent interaction with the FGF receptor, which triggers actin dynamic through a signaling cascade.^21,22As growth cones migrate, adhesion sites must be recycled at a rate that somehow matches the speed of migration. Adhesion turnover can be schematically decomposed in several sequential phases (Fig. 2B). (1) Initiation of a first single ligand/receptor bond powered by membrane diffusion²³ and followed by trapping through a key/lock interaction; (2) Formation of small adhesion clusters through the recruitment of more ligand/receptor pairs, and possibly stabilized by cis-oligomerization (cadherins through the same interface as the trans-dimer, IgCAMs through FnIII domains). These clusters might form very transiently and serve as sites of actin recruitment, as demonstrated for N-cadherin;¹⁶ (3) contact maturation and possible reinforcement by connection to the cytoskeleton (as demonstrated for integrins in fibroblasts²⁴); (4) Adhesion rupture, which can proceed through ligand/receptor dissociation triggered by cytoskeleton tension. Indeed, the intrinsic lifetime of ligand/receptor bonds such as cadherins, is sensitive to the mechanical force applied on them.²⁵ Furthermore, the loosening of receptor/cytoskeketon connections can cause inside-out rupture of ligand/receptor bonds. This was demonstrated for fibronectin/integrin interactions by the fact that when fibronectin coated-beads reach the base of a fibroblast lamellipodium, they spontaneously detach from the cell surface.²⁶ In the case of very sticky ligand/receptor interactions such as SynCAM homophilic adhesions,²⁷ this process can actually be a limiting step that slows down growth cone advance. Indeed, SynCAM couples very well to the actin flow, but is unable to support growth cone migration.¹⁶ Finally, adhesion rupture might also proceed through membrane rupture, the adhesion receptors being extracted from the cell membrane and left behind on the substrate (demonstrated for integrins at the tail of fibroblasts²⁸).These basic processes can be accompanied by more complex and active phenomena, e.g. involving forward surface transport as shown for NCAM²⁹ or internal trafficking in the case of L1.³⁰ By interacting with the clathrin adaptor AP-2 through a specific RSLE motif in its intracellular tail, L1 can undergo endocytosis in the central domain and exocytosis at the periphery of the growth cone.³⁰ This mechanism generates a density gradient of L1 molecules which accelerates the formation of bonds with a variety of ligands, including L1 itself. The use of an L1-GFP construct in which the N-terminal GFP could be rapidly cleaved off by thrombin, together with L1-Fc microspheres manipulated by optical tweezers showed that local exocytosis of L1-rich vesicles at the growth cone periphery indeed participates in enhancing the formation of L1 homophilic contacts.³¹ We did not observe such internal traffic for N-cadherin within the growth cone, partly because of a difficulty to introduce a fluorescent protein tag in the ectodomain, which otherwise perturbs the adhesive function. However, the use of an N-cadherin molecule with triple mutation in the juxta-membrane domain that abolishes binding to p120 catenin, involved in the export of N-cadherin to the cell surface, suggested that recycling events might also play a role.³²By measuring the fluorescence recovery after photobleaching (FRAP) of GFP-tagged receptors transiently trapped at ligand-coated microspheres and analyzing the curves using a diffusion/reaction model, we were able to compute the equilibrium turnover rates of ligand/receptor pairs in controlled adhesive contacts involving many simultaneous bonds (Fig. 3C and D). We found that mature L1 homophilic adhesions recycle fast compared to other IgCAMs such as TAG-1/NrCAM adhesions,²⁰ likely owing to the specific internalization motif present in L1. Indeed, the recycling rate was reduced by a factor of 3 after truncation of the L1 intracellular tail, which prevented endocytosis.³¹ N-cadherin homophilic adhesions have an intermediate turnover rate, which is sensitive to the binding to catenin partners.³² Using these measurements as well as data from the literature, we plotted the impact of both receptor-cytoskeleton coupling and adhesion turnover rate on neurite outgrowth (Fig. 4A and B), which is strongly proportional to growth cone velocity.¹⁶ The graphs show that a strong coupling between ligand-occupied receptors and the actin flow is necessary, but not sufficient for neurite extension (Fig. 4C). Another requirement is that the turnover of ligand/receptor adhesions lies in an optimal range: not too high, otherwise bonds detach before coupling can occur, and not too slow either, since sticky bonds which do not rupture paralyze growth cone progression (Fig. 4D). A similar bell-shape curve between the strength of cell-substrate adhesion and cell migration speed was demonstrated for fibroblasts³³ and keratocytes,³⁴ indicating that these coupled mechanisms are fundamental to cell migration. To fully understand the quantitative relationship between adhesion turnover and the clutch process, it would be helpful to add data to this preliminary graph. For example, the extracellular matrix molecule laminin is known to support axon growth very efficiently but, to our knowledge, neither the coupling to the actin flow in growth cones or the adhesive turnover rate of integrins has been evaluated yet. Conversely, NCAM was shown to couple well to the actin flow³⁵ and induce neurite outgrowth,^4,36 but measurements of the lifetime of NCAM homophilic adhesions within growth cones are still lacking.Open in a separate windowFigure 4Impact of ligand/receptor turnover rate and receptor/cytoskeletal coupling on neurite outgrowth. (A and B) Example of 2 DIV rat hippocampal neurons plated on N-cadherin-Fc coated substrate and transfected at 1 DIV with N-cadherin-GFP. (A) DIC image. (B) Fluorescence image. The longest neurite, most likely the axon, is outlined by arrowheads. (C and D) In both graphs, the y-axis represents the longest neurite length after two days plating on ligand-coated glass. (Red) Rat hippocampal neurons transfected with either wild type or mutated N-cadherin molecules, interacting with purified N-cadherin ligands.^16,32 The scale in red intensity represents from dark to light: wild type N-cadherin, N-cadherin deleted of the whole ectodomain, N-cadherin truncated in the C-terminal region binding to β-catenin, N-cadherin with triple mutation in the juxta-membrane domain interacting with p120, and wild type N-cadherin in the presence of cytochalasin D. (Blue) Neurons transfected with either wild type L1 (dark) or L1 truncated in the intracellular tail (light), interacting with purified L1.³¹ Neurite growth on L1 was estimated from references.^7,14,30 (Grey) Interaction between endogenous SynCAM1 molecules expressed on growth cones and SynCAM-Fc ligands coated on microspheres or flat glass.¹⁶ The turnover of SynCAM homophilic interactions was estimated from SynCAM-coated Quantum dots detaching from neurons transfected with SynCAM1.⁴² (Green) Neuroblastoma cells expressing NrCAM-GFP in contact with TAG-1 coated microspheres.²⁰ DRG neurite growth on TAG-1 was taken from reference.⁴³ (Orange) The coupling index was taken from optical tweezers experiments using anti-β1 integrin coated beads interacting with DRG neurons,³ the turnover rate was inferred from FRAP experiments on fibroblast focal contacts⁴⁴ and neurite growth on fibronectin was estimated from reference.¹⁹ We omitted statistics for clarity. The SEM are usually in the order of 5–15% of the mean, for sample sizes of typically 20–30 beads (coupling index and turnover rate) and 40–100 transfected cells (neurite length).One important question is how these observations obtained from simplified in vitro systems using stiff substrates of well-defined geometry coated with specific purified proteins at controlled density, translate to the in vivo situation. There, the 3D substrate is comprised of extracellular matrix and multiple cell types, co-expressing many different CAMs that can bind simultaneously in various stoechiometries and also generating local gradients of chemo-attractant and chemo-repulsive signals.³⁷ Substrate flexibility is probably an important factor, since axons grow more slowly when neurons are plated on a layer of fibroblasts expressing CAMs²¹ than when molecules are immobilized on a substrate.^16,30 This preference for cells to move on stiff substrates, called durotaxis, has been well described for fibroblasts.³⁸ Another specific feature of the in vivo situation is the existence of decision points, often correlated with the presence of guidepost cells where growth cones make a pause and often change shape and reorient before turning to another direction.³⁹ This type of behavior has been successfully mimicked in vitro using artificial guideposts made of fibronectin or laminin coated microspheres.⁴⁰ Whereas growth cones display a fairly continuous displacement on a homogeneous substrate,¹⁶ the presence of these guideposts make growth cones either slow down, pause or even collapse, or conversely accelerate, depending on the CAM grafted on the bead.^40,41 Finally, the shape itself of the growth cone can be an indicator of its motile state:³⁹ this is also true in vitro where small growth cones are often the most rapid whereas large and flat growth cones stay rather immobile. Thus, although the in vivo situation seems at first sight awfully complex, some general trends can be explained given a small number of interacting molecular species and rather simple bio-chemical and mechanical models.In conclusion, the dynamic regulation of growth cone advance can take place at several levels: (1) the actin-associated proteins controlling actin dynamics (nucleation, polymerization, sequestering, branching); (2) the activity of motors pulling on the actin network, generating the retrograde flow; (3) the intracellular adaptor proteins that link actin to the CAMs; (4) the membrane delivery and retrieval of CAMs; (5) the ligand/receptor interaction properties themselves; and (6) the processes regulating microtubule assembly and microtubule/actin interactions at the base of the growth cone. The orchestration in time and space of all these processes generates the movement and reactivity of growth cones necessary to lead axons to their target cells.