Photosynthesis-dependent Ca2+ influx and functional diversity between phospholipases in the formation of cell polarity in migrating cells of red algae

Unicellular spore cells, designated as monospores (also called archeospores), are well known as migrating plant cells, in which establishment of the anterior-posterior axis directs asymmetrical distribution of F-actin. Since the mechanisms of cell polarity formation are not yet fully elucidated in monospores, we investigated the roles of phosphoinositide signaling systems and Ca²⁺ mobilization in migration. Although we have already found the critical involvement of phosphatidylinositol 3-kinase in the establishment of cell polarity, we recently demonstrated the important roles of extracellular Ca²⁺ influx, phospholipase C (PLC) and phospholipase D (PLD). The remarkable characteristics of these factors are that Ca²⁺ influx depends on photosynthetic activity and that PLC and PLD play roles in the establishment and maintenance of cell polarity, respectively. These findings could provide new insight into the regulation of migration in eukaryotic cells.Key words: Ca²⁺ influx, cell polarity, phospholipase C, phospholipase D, photosynthesis, Porphyra yezoensisMonospores are responsible for asexual and clonal propagation of the marine multicellular red algae Porphyra and have an exceptional characteristic as migrating plant cells.^1–5 Monospores possess a round shape just after release from gametophytic blades (Fig. 1A and B), then undergo morphological change during migration. The establishment of cell polarity leads to the determination of anterior-posterior axis and asymmetrical localization of F-actin (Fig. 1C). After migration, monospores adhere to the substratum in which the apical-basal axis has been established for further development (Fig. 1D). Asymmetrical distribution of F-actin is also found in chemotaxic migration of Dictyostelium cells and leukocytes.^6,7 In these cells, reciprocal local accumulation of phosphoinositides, such as phosphatidylinositol-3,4,5-trisphosphate [PtdIns(3,4,5)P₃] at the leading edge and phosphatidylinositol-4,5-bisphosphate [PtdIns(4,5)P₂] at the trailing side, is critical for the establishment of cell polarity. Phosphatidylinositol 3-kinase (PI3K) and PtdIns(3,4,5)P₃-specific D-3-phosphatase PTEN have been identified as key modulators in the establishment of cell polarity, bringing asymmetrical distribution of these two phosphoinositides in plasma membranes.^6,8 Similarly, we found the involvement of PI3K activity in the establishment of cell polarity in migrating monospores,³ suggesting the evolutional conservation of the function of PI3K in migrating eukaryotic cells. On the other hand, the importance of cell wall synthesis has been found in the maintenance of the cell polarity during monospore migration⁴ as reported in Fucus zygotes.^9,10 Therefore, the establishment and maintenance of cell polarity are thought to be regulated separately in monospores of P. yezoensis. In this addendum, further evidence of differential regulation of cell polarity formation by extracellular Ca²⁺ influx and phospholipases in migrating monospores of red algae is documented according to our recent report.⁵Open in a separate windowFigure 1Establishment and maintenance of cell polarity in monospores from the red alga P. yezoensis. (A) Discharge of unicellular monospores from a multicellular gametophytic blade of P. yezoensis strain TU-1. Scale bar = 20 µm. (B–D) Asymmetrical distribution of F-actin during early development of monospores. F-actin was stained with alex Flour 488 phalloidin. (B) Discharged monospore. (C) Migrating monospore. (D) Adhering monospore. Upper and lower photos in each panel show bright-field and fluorescent images, respectively. Arrow in (C) indicates the direction of migration. Scale bars = 5 µm. (e) Schematic representation of our working hypothesis about the formation of cell polarity required for monospore migration. Photosynthesis-dependent [Ca²⁺]_cyt increase regulates PLC and PI3K for the establishment of cell polarity, while PLD is required for the maintenance of the established cell polarity. DG, diacylgycerol; IP3, inositol-1,4,5-trisphosphate; IP3r, IP3 receptor; PC, phosphatidylcholine.     

Regulation of stomatal opening by the guard cell expansin AtEXPA1

Stomatal movement is strictly regulated by various intracellular and extracellular factors in response environmental signals. In our recent study, we found that an Arabidopsis guard cell expressed expansin, AtEXPA1, regulates stomatal opening by altering the structure of the guard cell wall. This addendum proposes a mechanism by which guard cell expansins regulate stomatal movement.Key words: expansin, stomatal movement, AtEXPA1, guard cell, wall looseningStomatal movement is the most popular model system for cellular signaling transduction research. A complicated complex containing many proteins has been proposed to control stomatal responses to outside stimuli. The known regulation factors are primarily located in the nucleus, cytoplasm, plasma membrane and other intracellular organelles.^1,2 Although the cell wall structure of the stomata is different from that of other cells,^3,4 the presence of stomatal movement regulation factors in the cell wall has seldom been reported in reference ⁵. In our previous work, we found that extracellular calmodulin stimulates a cascade of intracellular signaling events to regulate stomatal movement.⁶ The involvement of this signaling pathway is the first evidence that cell wall proteins play an important role in regulation of stomatal opening. Cell wall-modifying factors constitute a major portion of cell wall proteins. However, the role of these factors in the regulation of stomatal movement is not yet known.Expansins are nonenzymatic proteins that participate in cell wall loosening.^7–9 Expansins were first identified as “acid-growth” factors because they have much higher activities at acidic pHs.^10,11 It has been reported that expansins play important roles in plant cell growth, fruit softening, root hair emergence and other developmental processes in which cell wall loosening is involved.^7–9,12,13 Wall loosening is an essential step in guard cell swelling and the role of stomatal expansins was investigated. AtEXPA1 is an Arabidopsis guard-cell-specific expansin.^13,14 Over-expressing AtEXPA1 increases the rate of light-induced stomatal opening,^14,15 while a potential inhibitor of expansin activity, AtEXPA1 antibody, reduces the sensitivity of stomata to stimuli.¹⁴ We showed that the transpiration rate and the photosynthesis rate in plant lines overexpressing AtEXPA1 were nearly two times the rates for wild-type plants (Fig. 1). These in plant data revealed that expansins accelerated stomatal opening under normal physiological conditions. In addition, the increases in the transpiration and photosynthesis rates strongly suggested the possibility of exploiting expansin-regulated stomatal sensitivity to modify plant drought tolerance. Compared with the effect of hydrolytic cell wall enzymes, the destruction of cell wall structures induced by expansins is minimal. In addition, it is very difficult to directly observe the changes in the guard cell wall structure caused by expansins during stomatal movement. Our recent work showed that, in AtEXPA1-overexpressing plants, the volumetric elastic modulus is lower than in wild-type plants,¹⁴ which indicates the wall structure was loosened and that the cell wall was easier to extend. Taken together, our data suggest that expansins participate in the regulation of stomatal movement by modifying the cell walls of guard cells.Open in a separate windowFigure 1Effects of AtEXPA1 overexpression on transpiration rates and photosynthesis rates. The transpiration rate (left) and photosynthesis rate (right) of wild-type and transgenic AtEXPA1 lines were measured at 10:00 AM in the greenhouse after being watered overnight. The illumination intensity was 180 µmol/m²·s. Bars represent the standard error of the mean of at least five plants per line.It is well known that the activation of proton-pumping ATPase (H⁺-ATPase) in the plasma membrane is an early and essential step in stomatal opening.¹⁶ The action of the pump results in an accumulation of H⁺ outside of the cell, increases the inside-negative electrical potential across the plasma membrane and drives potassium uptake through the voltage-gated, inward-rectifying K⁺ channels.^17–19 The main function of the H⁺ pump is well accepted to create an electrochemical gradient across the plasma membrane; however, the other result is the acidification of the guard cell wall, which may also contribute to stomatal opening. A possible mechanism responsible for this effect is as follows. Expansins are in an inactive state when the stomata are in the resting state. Stomatal opening signals induce wall acidification and activate expansins. Then, the expansins move along with cellulose microfibrils and transiently break down hydrogen bonding between hemicellulose and the surface of cellulose microfibrils,^20,21 facilitating the slippage of cell wall polymers under increasing guard cell turgor pressure. The guard cell then swells and the stomata open (Fig. 2).Open in a separate windowFigure 2Model of how guard cell wall expansins regulate stomatal opening. Environmental stimuli, e.g., light, activate guard cell plasma membrane H⁺-ATPases to pump H⁺ into the extracellular wall space. The accumulation H⁺ acidifies the cell wall and induces the activation of expansin. The active expansin disrupts non-covalent bonding between cellulose microfibrils and matrix glucans to enable the slippage of the cell wall. The wall is loosened coincident with guard cell swelling and without substantial breakdown of the structure.Although our results indicate that AtEXPA1 regulates stomatal movement, the biochemical and structural mechanism by which AtEXPA1 loosens the cell wall remains to be discovered. It remains to figure out the existing of other expansins or coordinators involving in this process. In addition, determining the roles of expansins and the guard cell wall in stomatal closing is another main goal of future research.     

Dynamic protein trafficking to the cell wall

Recently we have studied the secretion pattern of a pectin methylesterase inhibitor protein (PMEI1) and a polygalacturonase inhibitor protein (PGIP2) in tobacco protoplast using the protein fusions, secGFP-PMEI1 and PGIP2-GFP. Both chimeras reach the cell wall by passing through the endomembrane system but using distinct mechanisms and through a pathway distinguishable from the default sorting of a secreted GFP. After reaching the apoplast, sec-GFP-PMEI1 is stably accumulated in the cell wall, while PGIP2-GFP undergoes endocytic trafficking. Here we describe the final localization of PGIP2-GFP in the vacuole, evidenced by co-localization with the marker Aleu-RFP, and show a graphic elaboration of its sorting pattern. A working model taking into consideration the presence of a regulated apoplast-targeted secretion pathway is proposed.Key words: cell wall trafficking, endocytosis, GPI-anchor, PGIP2, PMEI1, secretion pathway, vacuole fluorescent markerCell wall biogenesis, growth, differentiation and remodeling, as well as wall-related signaling and defense responses depend on the functionality of the secretory pathway. Matrix polysaccharides, synthesized in the Golgi stacks, and cell wall proteins, synthesized in the ER, are packaged into secretory vesicles that fuse with the plasma membrane (PM) releasing their cargo into the cell wall. Also the synthesis and deposition of cellulose itself are driven by the endomembrane system which controls the assembly, within the Golgi, and the export to the plasma membrane of rosette complexes of cellulose synthase.¹ Secretion to the cell wall has always been considered a default pathway² but recent studies have evidenced a complex regulation of wall component trafficking that does not seem to follow the default secretion model. Recent evidence that several cell wall proteins are retained in the Golgi stacks until specific signals at the N-terminal domain are proteolitically removed is a case in point.^3–5 Moreover, it has previously been reported that secretion of exogenous marker proteins (secGFP and secRGUS) and cell wall polysaccharides reach the PM through different pathways.⁶ More recently, we have reported that cell wall protein trafficking also occurs through mechanisms distinguishable from that of a secreted GFP suggesting that more complex events than the mechanisms of bulk flow control cell wall growth and differentiation.⁷ To follow cell wall protein trafficking we used a Phaseolus vulgaris polygalacturonase inhibitor protein (PGIP2) and an Arabidopsis pectin methylesterase inhibitor protein (PMEI1) fused to GFP (PGIP2-GFP and secGFP-PMEI1). Both apoplastic proteins are involved in the remodeling of pectin network with different mechanisms. PGIP2 specifically inhibits exogenous fungal polygalacturonases (PGs) and is involved in the plant defense mechanisms against pathogenic fungi.^8,9 PMEI1 counteracts endogenous PME and takes part in the physiological synthesis and remodeling of the cell wall during growth and differentiation.^10,11 The specific functions of the two apoplastic proteins seem to be strictly related to the distinct mechanisms that control their secretion and stability in the cell wall. In fact, while secGFP-PMEI1 moves through ER and Golgi stacks linked to a glycosyl phosphatidylinositol (GPI)-anchor, PGIP2-GFP moves as a cargo soluble protein. Furthermore, secGFP-PMEI1 is stably accumulated in the cell wall, while PGIP2-GFP, over the time, is internalized into endosomes and targeted to vacuole, likely for degradation. After reaching the cell wall, the different fate of the two proteins seems to be strictly related to the presence/absence of their physiological counteractors. PMEI regulates the demethylesterification of homogalacturonan by inhibiting pectin methyl esterase (PME) activity through the formation of a reversible 1:1 complex which is stable in the acidic cell wall environment.¹² Stable wall localization of PMEI1 is likely related to its interaction with endogenous PME, always present in the wall. Unlike PMEs, fungal polygalacturonases (PGs), the physiological interactors of PGIP2, are present in the cell wall only during a pathogen attack. The absence of PGs may determine PGIP2 internalization. Internalization events have been already reported for PM proteins,^13–16 while cell wall protein internalization is surely a less well-known event. To date, only internalization of an Arabidopsis pollen-specific PME^4,5,17 and PGIP2 ⁷ has been reported.To further confirm the internalization of PGIP2-GFP and its final localization into the vacuole, we constructed a red fluorescent variant (RFP) of the green fluorescent marker protein that accumulates in lytic or acidic vacuole because of the barley aleurain sorting determinants (Aleu-RFP).¹⁸ The localization of PGIP2-GFP was compared to that of Aleu-RFP by confocal microscopy in tobacco protoplasts transiently expressing both fusions. Sixty hours after transformation, PGIP2-GFP labeled the central vacuole as indicated by complete co-localization with the vacuolar marker (Fig. 1A–D). Instead, at the same time point, secGFP-PMEI1 still labeled the cell wall (Fig. 1E–H) and never reached the vacuolar compartment. To summarize PGIP2-GFP secretion pattern, a graphic elaboration of confocal images is reported describing the sorting of PGIP2GFP in tobacco protoplast (Fig. 1I). The protein transits through the endomembrane system (green) and reaches the cell wall which is rapidly regenerating as evidenced by immunostaining with the red monoclonal antibody JIM7 that binds to methylesterified pectins.¹⁹ PGIP2-GFP is then internalized in endosomes, labeled in yellow because of the co-localization with the styryl dye FM4-64, a red marker of the endocytic pathway.Open in a separate windowFigure 1PGIP2-GFP, but not secGFP-PMEI1, is internalized and reaches the vacuole in tobacco leaf protoplasts. (A) Approximately 60 h after transformation, PGIP2-GFP labeled the central vacuole as indicated by co-localization with the vacuole marker Aleu-RFP (B). (C) Merged image of (A and B). (D) Differential interference contrast (DIC) image of (A–C). On the contrary, secGFP-PMEI1 still labeled cell wall (E). (F) No co-localization is present in the vacuole labeled by Aleu-RFP. (G) Merged image of (E and F). (H) DIC image of (E–G). (I) Graphic elaboration of confocal images describing the sorting of PGIP2. The protein is sorted by the endomembrane system (green) to the cell wall (red) that is regenerated by the protoplast. Lacking the specific ligand, it is then internalized in endosome (yellow). Details are reported in the text.In Figure 2 we propose a model of the mechanism of secGFP-PMEI1 and PGIP2-GFP secretion derived from the different lines of evidence previously reported in reference ⁷. SecGFPPMEI1 (Fig. 2-1), but not PGIP2-GFP (Fig. 2-2), carries a GPI-anchor, required for its secretion to the cell wall. When the anchorage of GPI is inhibited by mannosamine (Fig. 2-a) or by the fusion of GFP to the C-terminus of PMEI1 (Fig. 2-b), the two non-anchored proteins accumulate in the Golgi stacks. Evidence of retention in Golgi stacks has already been reported for other two cell wall proteins.^3–5 Unlike secGFP-PMEI1, PGIP2-GFP is not stably accumulated in the cell wall and undergoes endocytic trafficking (Fig. 2-3). PGIP2-GFP internalization, likely due to the absence of PGs, might also be related with its ability to interact with homogalacturonan and oligogalacturonides,²⁰ which have been reported to internalize^21,22 (Fig. 2-4). Since SYP 121, a Qa-SNARE, is involved in the default secretion of secGFP,²³ but not in secretion of PGIP2-GFP and secGFP-PMEI1, trafficking mechanisms underlying secretion into the apoplast are likely different from those underlying the default route (Figs. 2-5). Taken as a whole, evidence suggests the existence of currently undefined signals that control apoplast-targeted secretion.Open in a separate windowFigure 2Schematic illustration for secGFP-PMEI1 and PGIP2-GFP trafficking. See text for details.     

Myosin light chain kinases: Division of work in cell migration

Cell motility is a highly coordinated multistep process. Uncovering the mechanism of myosin II (MYO2) activation responsible for the contractility underlying cell protrusion and retraction provides clues on how these complementary activities are coordinated. Several protein kinases have been shown to activate MYO2 by phosphorylating the associated myosin light chain (MLC). Recent work suggests that these MLC kinases are strategically localized to various cellular regions during cell migration in a polarized manner. This localization of the kinases together with their specificity in MLC phosphorylation, their distinct enzymatic properties and the distribution of the myosin isoforms generate the specific contractile activities that separately promote the cell protrusion or retraction essential for cell motility.Key words: myosin, MLCK, ROK, MRCK, phosphorylation, cell migrationCell movement is a fundamental activity underlying many important biological events ranging from embryological development to immunological responses in the adult. A typical cell movement cycle entails polarization, membrane protrusion, formation of new adhesions, cell body translocation and finally rear retraction.¹ A precise temporal and spatial coordination of these separate steps that take place in different parts of the cell is important for rapid and efficient movement.²One major event during eukaryotic cell migration is the myosin II (MYO2)-mediated contraction that underlies cell protrusion, traction and retraction.^1,3 An emerging theme from collective findings is that there are distinct myosin contractile modules responsible for the different functions which are separately regulated by local myosin regulatory light chain (MLC) kinases. These kinases contribute to contractile forces that connect adhesion, protrusion and actin organization.² Unraveling the regulation of these contractile modules is therefore pivotal to a better understanding of the coordination mechanism.At the lamellipodium, the conventional calcium/calmodulin-dependent myosin light chain kinase (MLCK) has been shown to play an essential role in a Rac-dependent lamellipodial extension.⁴ Inhibition of calmodulin or MLCK activity by specific photoactivatable peptides in motile eosinophils effectively blocks lamellipodia extension and net movement.⁵ Furthermore, there is a strong correlation between activated MLCK and phosphorylated MLC within the lamellipodia of Ptk-2 cells as revealed by fluorescence resonance energy transfer (FRET) analysis.⁶ More recent studies showed MLCK to regulate the formation of focal complexes during lamellipodia extension.^7,8 Functionally, MLCK is thought to play a critical role in the environment-sensing mechanism that serves to guide membrane protrusion. It mediates contraction that exerts tension on integrin-extracellular matrix (ECM) interaction, which, depending on the rigidity of the substratum, will lead to either stabilization of adhesion resulting in protrusion or destabilization of attachment seen as membrane ruffling on non-permissive surfaces.^8,9As a Rho effector, Rho-associated kinase (ROK/ROCK/Rho-kinase) has been shown to regulate stress fibers and focal adhesion formation by activating myosin, an effect that can be blocked by the specific ROK inhibitor Y-27632.^10,11 Myosin activation by ROK is the effect of two phosphorylation events: the direct phosphorylation on MLC and the inhibition of myosin phosphatase through phosphorylation of its associated myosin-binding subunit (MBS).¹¹ Consistent with this notion of a localization-function relationship, ROK and MBS, which can interact simultaneously with activated RhoA,¹¹ have been shown to colocalize on stress fibers.^12,13 In migrating cells, Rho and ROK activities have been mostly associated with the regulation of tail retraction, as inhibition of their activities often results in trailing tails due to the loss of contractility specifically confined to the cell rear.^14,15 Tail retraction requires high contractile forces to overcome the strong integrin-mediated adhesion established at the rear end, an event which coincides with the strategic accumulation of highly stable and contractile stress fibers that assemble at the posterior region of migrating cells.MRCK was previously shown to phosphorylate MLC and promote Cdc42-mediated cell protrusion.¹⁶ More recently, it was found to colocalize extensively with and regulate the dynamics of a specific actomyosin network located in the lamella and cell center, in a Cdc42-dependent manner but independent of MLCK and ROK.¹⁷ The lamellar actomyosin network physically overlaps with, but is biochemically distinct from the lamellipodial actin meshwork.^9,18 The former network consists of an array of filaments assembled in an arrangement parallel to the leading edge, undergoing continuous retrograde flow across the lamella, with their disassembly occurring at the border of the cell body zone sitting in a deeper region.^17–19 Retrograde flow of the lamellar network plays a significant role in cell migration as it is responsible for generating contractile forces that support sustained membrane protrusion and cell body advancement.^17–19It is therefore conceivable that these three known MLC kinases are regulated by different signaling mechanisms at different locations and on different actomyosin contractile modules. The coordination of the various modules will ensure persistent directional migration (Figure 1). Phosphorylation of MLC by PAK and ZIP kinase has also been reported, but their exact roles in this event have yet to be determined.^20,21 It is also noteworthy that individual kinases can work independently of each other, as amply shown by evidence from inhibitor treatments. This is particularly true for MRCK in the lamella, whose activity on lamellar actomyosin flow is not affected by ML7 and Y-27632, the inhibitors of MLCK and ROK respectively.¹⁷ These findings further indicate that although both ROK and MRCK have been shown to upregulate phosphorylated MLC levels by inhibiting the myosins phosphatases,^11,22 they are likely to act as genuine MLC kinases themselves, without the need of MLCK as previously suggested.¹¹Open in a separate windowFigure 1Upper panel depicts a model for the specific activation of the different MLC kinases at various locations in the cell. In response to upstream signals, MLC kinases MLCK, MRCK and ROK are activated and localized to different regions. In the case of MRCK and ROK, the interaction of the GTP-bound Rho GTPase binding domain will determine the specific action of the downstream kinase, resulting in actomyosin contractility at different locations. The coordination of these signalling events is crucial for directional cell migration. Lower panel shows a typical front-rear location for Myosin 2A and 2B in a migrating U2OS cell.In conjunction with their differences in localization, the three MLC kinases show apparent individual preferences and specificity towards the MYO2 isoforms that they associate with. The two major MYO2 isoforms MYO2A and 2B are known to have distinct intracellular distributions that are linked to their individual functions (Figure 1).^23,24 In motile cells, MYO2A localization that is skewed towards the protruding cell front is consistent with it being the major myosin 2 component of the lamellar filaments regulated by MRCK as well as its regulation by MLCK in lamellipodial contraction.^8,17,19 In contrast, the enrichment of MYO2B at retracting cell rear conforms well with the requirement of thick and stable stress fibers capable of causing tail contraction and prevention of protrusion under the control of Rho/ROK signaling.^23,25 The selection for MYO2B filaments in the cell rear stems from their more contractile and stable nature compared with MYO2A, a consequence of their higher time-averaged association with actin.^26,27 Conversely, the lower tension property of MYO2A filaments suggests that they are more dynamic in nature,^26,27 a characteristic which fits well with the dynamic actomyosin activities at the leading edge and lamella that regulate protrusion.It deserves special mention that the three MLC kinases display subtle differences in their specificity towards MLC. While MLCK and MRCK phosphorylate only a single Ser19 site (monophosphorylation),^18,28 ROK is able to act on both Thr18 and Ser19 residues causing diphosphorylation of MLC,²⁹ MLCK only causes diphosphorylation when present at higher concentrations.³⁰ By further increasing its actin-activated ATPase activity, diphosphorylation of MLC has been shown to induce a higher myosin activation and filament stability.^30–32 The use of specific antibodies that can differentiate between the two populations of phosphorylated MLC has been instrumental in revealing their localization and correlation with the activity of the MLC kinases. The emerging picture from these experiments is that mono and diphosphorylated MLC exhibit distinct distributions in migrating cells, with the monophosphorylated MLC localized more towards the protrusive region, while the diphosphorylated form is more enriched at the posterior end.^21,33 Taking into account their biochemical properties, the polarized distributions of these differentially phosphorylated MLC coincide functionally with the segregation of the MYO2 isoforms and their corresponding regulators. These findings provide further support for the existence of segregated contractile modules in migrating cell and their distinctive regulation.The mechanisms that determine the specific segregation of the contractile modules and their regulation are unclear. However, some clues have emerged from recent studies. It has been shown that the C-terminal coiled-coil region of MYO2B is important for determining its localization in cell rear²⁵ and which requires Rho/ROK activity as their inhibition resulted in the loss of this specific localization.²³ Correspondingly, the inhibition of MRCK activity resulted in the loss of lamella-localized MYO2A.¹⁷ These findings suggest that activation of MYO2 filaments by their upstream regulators is important for their functional segregation and maintenance. It is noteworthy that both ROK and MRCK have distinct regulatory domains including the pleckstrin homology domains which have been shown to be essential for their localization, a process which may involve myosin interaction and lipid-dependent targeting as has been respectively shown for ROK and MRCK.^11,13,16 Further, the specificity of MRCK for lamellar actomyosin is believed to be largely determined by the two proteins it forms a complex with: the adaptor LRAP35a, and the MYO2-related MYO18A. Activation of MYO18A by MRCK, a process bridged by LRAP35a, is a crucial step which facilitates MRCK regulation on lamellar MYO2A.¹⁷The mechanisms responsible for segregating the contractile modules and their regulators may also comprise a pathway that parallels the microtubule-modulatory Par6/aPKC/GSK3β signalling pathway which regulates cellular polarization. This notion is supported by both Cdc42 and Rho being common upstream regulators of these two pathways.³⁴ GTPase activation may determine the localized activities of the separate contractile modules and create an actomyosin-based asymmetry across the cell body, which together with the microtubule-based activities, result in the formation of a front-back axis important for directional movement. The involvement of MRCK in MTOC reorientation and nuclear translocation events,³⁵ and our unpublished observation that LRAP35a has a GSK3β-dependent microtubule stabilizing function are supportive of a possible cross-talk between these two pathways.In conclusion, the complex regulation of contractility in cell migration emphasizes the importance of the localization, specificity and enzymatic properties of the different MLC kinases and myosin isoforms involved. The initial excitement and confusion caused by the emergence of the different MLC kinases are fading, being now overtaken by the curiosity about how they cooperate and are coordinated while promoting cell motility.     

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.     

Putting the brakes on cancer cell migration: JAM-A restrains integrin activation

Junctional Adhesion Molecule A (JAM-A) is a member of the Ig superfamily of membrane proteins expressed in platelets, leukocytes, endothelial cells and epithelial cells. We have previously shown that in endothelial cells, JAM-A regulates basic fibroblast growth factor, (FGF-2)-induced angiogenesis via augmenting endothelial cell migration. Recently, we have revealed that in breast cancer cells, downregulation of JAM-A enhances cancer cell migration and invasion. Further, ectopic expression of JAM-A in highly metastatic MDA-MB-231 cells attenuates cell migration, and downregulation of JAM-A in low-metastatic T47D cells enhance migration. Interestingly, JAM-A expression is greatly diminished as breast cancer disease progresses. The molecular mechanism of this function of JAM-A is beyond its well-characterized barrier function at the tight junction. Our results point out that JAM-A differentially regulates migration of endothelial and cancer cells.Key words: JAM-A, integrin, α_vβ₃, FGF-2, breast cancer, cell migration and invasion, T47D, MDA-MB-231, siRNAEndothelial and epithelial cells exhibit cell polarity and have characteristic tight junctions (TJs) that separate apical and basal surfaces. TJs are composed of both transmembrane and cytoplasmic proteins. The three major families of transmembrane proteins include claudins, occludin and JAM family members.^1–3 Additionally, interaction between the peripheral proteins such as PDS-95/Discs large/ZO family (PDZ) domain-containing proteins in TJs plays an important role in maintaining the junctional integrity.^2,4,5JAMs are type I membrane proteins (Fig. 1) predominately expressed in endothelial and epithelial cell TJs, platelets and some leukocytes.^6–8 The classical JAMs are JAM-A, JAM-B and JAM-C, which can all regulate leukocyte-endothelial cell interaction through their ability to undergo heterophilic binding with integrins α_Lβ₂ or α_vβ₃, α₄β₁ and α_Mβ₂ respectively. The cytoplasmic tail of JAMs contains a type II PDZ-domain-binding motif (Fig. 1) that can interact with the PDZ domain containing cytoplasmic molecules such as ZO-1, ASIP/PAR-3 or AF-6.^9,10 Additionally, consistant with their junctional localization and their tendency to be involved in homophilic interactions, JAMs have been shown to modulate paracellular permeability and thus may play an important role in regulating the epithelial and endothelial barrier.^11,12 In addition, ectopic expression of JAM-A in CHO cells promotes localization of ZO-1 and occludin at points of cell contacts, which suggests a role for JAM-A in TJ assembly.^10,13,14 Recently, it has been shown that JAM-A regulates epithelial cell morphology by modulating the activity of small GTPase Rap1 suggesting a role for JAM-A in intracellular signaling.¹⁵Open in a separate windowFigure 1Schematic representation of the domain structure of JAM family proteins. V, variable Ig domain; C2, constant type 2 Ig domain; TM, transmembrane domain; T-II, Type II PDZ-domain binding motif.We have previously shown that JAM-A is a positive regulator of fibroblast growth factor-2 (FGF-2) induced angiogenesis.¹⁶ Evidence was provided to support the notion that JAM-A forms a complex with integrin α_vβ₃ at the cell-cell junction in quiescent human umbilical cord vein endothelial cells (HUVECs) and FGF-2 dissociates this complex.¹⁶ It was further established that inhibition of JAM-A using a function-blocking antibody also inhibits FGF-2 induced HUVECs migration in vitro and angiogenesis in vivo. Overexpression of JAM-A induced a change in HUVECs morphology similar to that observed when treated with FGF-2.¹⁷ Furthermore, overexpression of JAM-A, but not its cytoplasmic domain deletion mutant, augmented cell migration in the absence of FGF-2.¹⁷ In addition, downregulation of JAM-A in HUVECs using specific siRNA, resulted in reduced FGF-2-induced cell migration and inhibition of mitogen activated protein (MAP) kinase activation.¹⁸ These findings clearly suggested that JAM-A positively regulates FGF-2-induced endothelial cell migration. This was further confirmed in vivo by using JAM-A null mouse in which FGF-2 failed to support angiogenesis.¹⁹It is known that JAM-C, a JAM family member, is involved in the process of tumor cell metastasis.²⁰ However, little is known about JAM-A''s role in cancer progression. We recently found that JAM-A is expressed in breast cancer tissues and cell lines.²¹ Based on our studies with endothelial cells it was felt that JAM-A expression in breast cancer cells may also enhance the migratory ability of these cells. Surprisingly, we found an inverse relation between the expression of JAM-A and the metastatic ability of breast cancer cells. T47D cells, which express high levels of JAM-A, are the least migratory; whereas MDA-MB-231 cells, which are highly migratory, are found to express the least amount of JAM-A.²¹ We also found that overexpression of JAM-A in MDA-MB-231 cells caused a change in cell morphology from spindle-like to rounded shape and formed cobblestone-like clusters.²¹ This is consistent with the previous report, that downregulation of JAM-A expression from epithelial cells using siRNA results in the change of epithelial cell morphology.¹⁵ This change in cell morphology by knockdown of JAM-A was attributed to the disruption of epithelial cell barrier function.¹⁵ It was further shown that knockdown of JAM-A affects epithelial cell morphology through reduction of β₁integrin expression due to decreased Rap1 activity.¹⁵ Our observed effect of JAM-A downregulation in T47D cells, however, is not due to downregulation of β₁integrin, since the level of this integrin was not affected in these cells. Interestingly, overexpression of JAM-A significantly affected both the cell migration and invasion of MDA-MB-231 cells. Furthermore, knockdown of JAM-A using siRNA enhanced invasiveness of MDA-MB-231 cells, as well as T47D cells.²¹ The ability of JAM-A to attenuate cell invasion was found to be due to the formation of functional tight junctions as observed by distinct accumulation of JAM-A and ZO-1 at the TJs and increased transepithelial resistance. These results identify, for the first time, a tight junctional cell adhesion protein as a key negative regulator of breast cancer cell migration and invasion.²¹JAM-A has been shown to be important in maintaining TJ integrity.^15,22–25 Disruption of TJs has been implicated to play a role in cancer cell metastasis by inducing epithelial mesenchymal transition.²⁶ Several laboratories, including ours, have shown that cytokines and growth factors redistribute JAM-A from TJs.^16,27,28 Consistent with this finding, it has been shown that hepatocyte growth factor (HGF) disrupts TJs in human breast cancer cells and downregulates expression of several TJ proteins.²⁹ It is therefore conceivable that the loss of JAM-A in highly metastatic cells is a consequence of disruption of TJs. This was further supported by the findings that overexpression of JAM-A forms functional TJs in MDA-MB-231 cells and attenuates their migratory behavior. Our result is the first report correlating an inverse relationship of JAM-A expression in breast cancer cells to their invasive ability.²¹Using cDNA microarray technology, it has been revealed how genes involved in cell-cell adhesion, including those of the TJ, are under or overexpressed in different carcinomas.^15,30 Cell-cell adhesion molecules have been well documented to regulate cancer cell motility and invasion. Of these, the cadherin family have been studied the most.^31,32 It was proposed that a cadherin switch, that is, the loss of E-cadherin and subsequent expression of N-cadherin, may be responsible for breast cancer cell invasion.^33,34 Although the role of cadherins is well-documented, it remains controversial since some breast cancer cell lines that do not express these proteins still posses highly invasive characteristics.^33,34 However, the observed effect of overexpression of JAM-A does not appear to be simply due to the formation of TJs, since individual cells that express increased JAM-A show reduced migration.²¹ This is not surprising, considering the fact that JAM-A in addition to its function of regulating TJ integrity is also shown to participate in intracellular signaling. JAM-A is capable of interacting homotypically as well as heterotypically on the cell surface.^35,36 It has also been shown that it interacts with several cytoplasmic proteins through its PDZ domain-binding motif and recruits signaling proteins at the TJs.³⁷ Recent findings using site-directed mutagenesis suggest that cis-dimerization of JAM-A is necessary for it to carry out its biological functions.³⁸ Our own observations suggest that a JAM-A function-blocking antibody inhibits focal adhesion formation in endothelial cells (unpublished data), whereas overexpresion of JAM-A in MDA-MB-231 cells show increased and stable focal adhesions.²¹ It is therefore conceivable that in quiescent endothelial/epithelial cells JAM-A associates with integrin to form an inactive complex at the TJ (Fig. 2). Growth factors such as FGF-2 signaling dissociates this complex thus allowing dimerization of JAM-A and activation of integrin augmenting cell migration (Fig. 2). On the contrary, in MDA-MB-231 cancer cells, which express low levels of JAM-A and do not form tight junctions, there may not be efficient inactive complex formation between JAM-A and integrin. Overexpression of JAM-A in these cells however, may promote such inactive complex formation leading to inhibition of integrin activation and JAM-A dimerization, both necessary events for cell migration. We are currently in the process of determining the specificity of interaction of JAM-A with integrins. Further experimentation is ongoing to determine the contribution of JAM-A dependent signaling in cell migration.Open in a separate windowFigure 2Schematic representation of JAM-A regulation of cell migration. JAM-A forms an inactive complex with the integrin and sequesters it at the TJs. Growth factor signaling dissociates this complex, promoting integrin activation and JAM-A dimerization leading to cell migration via MAP kinase activation. Ectopic expression of JAM-A in cancer cells may induce its association with integrin, forming an inactive complex and hence attenuation of migration.JAM-A differentially regulates cell migration in endothelial and cancer cells due to its ability to form inactive complex with integrin, making it a metastasis suppressor. The downregulation of JAM-A in carcinoma cells may be detrimental to the survival of breast cancer patients. It is therefore very important to determine the molecular determinants that are responsible for the downregulation of JAM-A during cancer progression. Thus, JAM-A, a molecule that dictates breast cancer cell invasion, could be used as a prognostic marker for metastatic breast cancer.     

Reactive oxygen species as transducers of sphinganine-mediated cell death pathway

Long chain bases or sphingoid bases are building blocks of complex sphingolipids that display a signaling role in programmed cell death in plants. So far, the type of programmed cell death in which these signaling lipids have been demonstrated to participate is the cell death that occurs in plant immunity, known as the hypersensitive response. The few links that have been described in this pathway are: MPK6 activation, increased calcium concentrations and reactive oxygen species (ROS) generation. The latter constitute one of the more elusive loops because of the chemical nature of ROS, the multiple possible cell sites where they can be formed and the ways in which they influence cell structure and function.Key words: hydrogen peroxide, long chain bases, programmed cell death, reactive oxygen species, sphinganine, sphingoid bases, superoxideA new transduction pathway that leads to programmed cell death (PCD) in plants has started to be unveiled.^1,2 Sphingoid bases or long chain bases (LCBs) are the distinctive elements in this PCD route that naturally operates in the entrance site of a pathogen as a way to contend its spread in the plant tissues.^2,3 This defense strategy has been known as the hypersensitive response (HR).^4,5As a lately discovered PCD signaling circuit, three connected transducers have been clearly identified in Arabidopsis: the LCB sphinganine (also named dihydrosphingosine or d18:0); MPK6, a mitogen activated kinase and superoxide and hydrogen peroxide as reactive oxygen species (ROS).^1,2 In addition, calcium transients have been recently allocated downstream of exogenously added sphinganine in tobacco cells.⁶Contrary to the signaling lipids derived from complex glycerolipid degradation, sphinganine, a metabolic precursor of complex sphingolipids, is raised by de novo synthesis in the endoplasmic reticulum to mediate PCD.^1,2 Our recent work demonstrated that only MPK6 and not MPK3 (commonly functionally redundant kinases) acts in this pathway and is positioned downstream of sphinganine elevation.² Although ROS have been identified downstream of LCBs in the route towards PCD,¹ the molecular system responsible for this ROS generation, their cellular site of formation and their precise role in the pathway have not been unequivocally identified. ROS are produced in practically all cell compartments as a result of energy transfer reactions, leaks from the electron transport chains, and oxidase and peroxidase catalysis.⁷Similar to what is observed in pathogen defense,³ increases in endogenous LCBs may be elicited by addition of fumonisin B1 (FB1) as well; FB1 is a mycotoxin that inhibits ceramide synthase. This inhibition results in an accumulation of its substrate, sphinganine and its modified forms, leading to the activation of PCD.^1,2,8 The application of FB1 is a commonly used approach for the study of PCD elicitation in Arabidopsis.^1,2,9–11An early production of ROS has been linked to an increase of LCBs. For example, an H₂O₂ burst is found in tobacco cells after 2–20 min of sphinganine supplementation,¹² and superoxide radical augmented in the medium 60 min after FB1 or sphinganine addition to Arabidopsis protoplasts (Fig. 1A). In consonance with this timing, both superoxide and H₂O₂ were detected in Arabidopsis leaves after 3–6 h exposure to FB1 or LCBs.¹ However, the source of ROS generation associated with sphinganine elevation seems to not be the same in both species: in tobacco cells, ROS formation is apparently dependent on a NADPH oxidase activity, a ROS source consistently implicated in the HR,^13,14 while in Arabidopsis, superoxide formation was unaffected by diphenyliodonium (DPI), a NADPH oxidase inhibitor (Fig. 1A). It is possible that the latter oxidative burst is due to an apoplastic peroxidase,¹⁵ or to intracellular ROS that diffuse outwards.^16,17 These results also suggest that both tobacco and Arabidopsis cells could produce ROS from different sources.Open in a separate windowFigure 1ROS are produced at early and long times in the FB1-induced PCD in Arabidopsis thaliana (Col-0). (A) Superoxide formation by Arabidopsis protoplasts is NADPH oxidase-independent and occurs 60 min after FB1 or sphinganine (d18:0) exposure. Protoplasts were obtained from a cell culture treated with cell wall lytic enzymes. Protoplasts were incubated with 10 µM FB1 or 10 µM sphinganine for 1 h. Then, cells were vacuum-filtered and the filtrate was used to determine XTT [2,3-bis-(2-methoxy-4-nitro-5-sulfophenyl)-2H-tetrazolium-5-carboxanilide, disodium salt] reduction as described in references ²⁸ and ²⁹. DPI was used at 50 µM. (B) H₂O₂ formation in Arabidopsis wt and lcb2a-1 mutant in the presence and absence of FB1. Arabidopsis seedlings were exposed to 10 µM FB1 and after 48 h seedlings were treated with DA B (3,3-diaminobencidine) to detect H₂O₂ according to Thordal-Christensen et al.³⁰It has been suggested that the H₂O₂ burst associated with the sphinganine signaling pathway leads to the expression of defense-related genes but not to the PCD itself in tobacco cells.¹² It is possible that ROS are involved in the same way in Arabidopsis, since defense gene expression is also induced by FB1 in Arabidopsis.⁹ In this case, it will be important to define how the early ROS that are DPI-insensitive could contribute to the PCD manifestation mediated by sphinganine.The generation of ROS (4–60 min) found in Arabidopsis was associated to three conditions: the addition of sphinganine (Fig. 1A), FB1 (Fig. 1A) or pathogen elicitors.¹⁵ This is consistent with the MPK6 activation time, which is downstream of sphinganine elevation and occurs as early as 15 min of FB1 or sphinganine exposure.² All of them are events that appear as initial steps in the relay pathway that produces PCD.In order to explore a possible participation of ROS at more advanced times of PCD progression, we detected in situ H₂O₂ formation in Arabidopsis seedlings previously exposed to FB1 for 48 h. As shown in Figure 1B, formation of the brown-reddish precipitate corresponding to the reaction of H₂O₂ with 3,3′-diaminobenzidine (DAB) was only visible in the FB1-exposed wild type plants, as compared to the non-treated plants. However, when lcb2a-1 mutant seedlings were used, FB1 exposure had a subtle effect in ROS formation. This mutant has a T-DNA insertion in the gene encoding subunit LCB2a from serine palmitoyltransferase (SPT), which catalyzes the first step in sphingolipid synthesis¹⁸ and the mutant has a FB1-resistant phenotype.² These results indicate that mutations in the LCB1¹ and LCB2a² genes (coding for the subunits of the heterodimeric SPT) that lead to a non-PCD phenotype upon the FB1 treatment, are unable to produce H₂O₂. In addition, they suggest that high levels of hydrogen peroxide are produced at advanced times in the PCD mediated by LCBs in Arabidopsis.Exposure of Arabidopsis to an avirulent strain of Pseudomonas syringae produces an endogenous elevation of LCBs as a way to implement defense responses that include HR-PCD.³ In this condition, we clearly detected H₂O₂ formation inside chloroplasts (Fig. 2A). When ultrastructure of the seedlings tissues exposed to FB1 for 72 h was analyzed, integrity of the chloroplast membrane system was severely affected in Arabidopsis wild-type seedlings exposed to FB1.² Therefore, we suggest that ROS generation-LCB induced in the chloroplast could be responsible of the observed membrane alteration, as noted by Liu et al. who found impairment in chloroplast function as a result of H₂O₂ formation in this organelle from tobacco plants. Interestingly, these plants overexpressed a MAP kinase kinase that activated the kinase SIPK, which is the ortholog of the MPK6 from Arabidopsis, a transducer in the PCD instrumented by LCBs.²Open in a separate windowFigure 2Conditions of LCBs elevation produce H₂O₂ formation in the chloroplast and perturbation in the membrane morphology of mitochondria. (A) Exposure of Arabidopsis leaves to the avirulent strain Pseudomonas syringae pv. tomato DC3000 (avrRPM1) (or Pst avrRPM1) induces H₂O₂ formation in the chloroplast. Arabidopsis leaves were infiltrated with 1 × 10⁸ UFC/ml Pst avrRPM1 and after 18 h, samples were treated to visualize H₂O₂ formation with the DAB reaction. Controls were infiltrated with 10 mM MgCl₂ and then processed for DAB staining. Then, samples were analyzed in an optical photomicroscope Olympus Provis Model AX70. (B) Effect of FB1 on mitochondria ultrastructure. Wild type Arabidopsis seedlings were treated with FB1 for 72 h and tissues were processed and analyzed according to Saucedo et al.² Ch, chloroplast; M, mitochondria; PM, plasma membrane. Arrows show mitochondrial cisternae. Bars show the correspondent magnification.In addition, we have detected alterations in mitochondria ultrastructure as a result of 72 h of FB1 exposure (Fig. 2B). These alterations mainly consist in the reduced number of cristae, the membrane site of residence of the electron transport complexes. In this sense, it has been shown that factors that induce PCD such as the victorin toxin, methyl jasmonate and H₂O₂ produce alterations in mitochondrial morphology.^20–22 In fact, some of these studies propose that ROS are formed in the mitochondria and then diffuse to the chloroplasts.^22–24It is reasonable to envisage that damage of the membrane integrity of these two organelles reflects the effects of vast amounts of ROS produced by the electron transport chains.^25,26 Recent evidence supports the destruction of the photosynthetic apparatus associated to the generation of ROS in the HR.²⁶ At this time of PCD progression, ROS could be contributing to shut down the energy machinery in the cell, which ultimately would become the point of no-return of PCD²⁷ as part of the execution program of the cell death mediated by LCBs.In conclusion, we propose that ROS can display two different functional roles in the PCD process driven by LCBs. These roles depend on the time of ROS expression, the cellular site where they are generated, the enzymes that produce them, and the magnitude in which they are formed.     

A role for pectin-associated arabinans in maintaining the flexibility of the plant cell wall during water deficit stress

One of the main components of pectin, a primary constituent of higher plant cell walls, is rhamnogalacturonan I. This polymer comprised of linked alternating rhamnose and galacturonic acid residues is decorated with side chains composed of arabinose and galactose residues. At present, the function of these side chains is not fully understood. Our research on Southern African resurrection plants, plants that are capable of surviving severe dehydration (desiccation), has revealed that their cell walls are capable of extreme flexibility in response to water loss. One species, Myrothamnus flabellifolia, has evolved a constitutively protected leaf cell wall, composed of an abundance of arabinose polymer side chains, suggested to be arabinans and/or arabinogalactans, associated with the pectin matrix. In this article, we propose a hypothetical model that explains how the arabinan rich pectin found in the leaves of this desiccation-tolerant plant permits almost complete water loss without deleterious consequences, such as irreversible polymer adhesion, from occurring. Recent evidence suggesting a role for pectin-associated arabinose polymers in relation to water dependent processes in other plant species is also discussed.Key words: arabinans, cell wall, desiccation, resurrection, rehydration, rhamnogalacturonan IThe flowering plant cell wall is a composite structure consisting of a skeletal framework of cellulose and hemicellulose embedded within a matrix of pectin polysaccharides and cell wall glycoproteins.^1,2 The pectin matrix, in turn, is composed of three primary types of polysaccharides, these being rhamnogalacturonan I (RGI), rhamnogalacturonan II (RGII) and homogalacturonan (HG).¹ RGII is a complex polysaccharide, consisting of many unusual sugar moieties, and is not present in large amounts in the wall.³ HG is effectively a linear homopolymer of galacturonic acid and is believed to facilitate the formation of tight junctions, ‘egg boxes’, by complexing with calcium ions present in the cell wall.¹ RGI is a polymer composed of a backbone of alternating glycosidically linked rhamnose and galacturonic acid residues.¹ Side chains, consisting of either arabinogalactan polymers or linear chains of arabinans and/or galactans, are then attached to the rhamnose residues of the RGI backbone.¹ The manner with which these polymers are attached or become entangled with each other and cellulosic polymers to form the pectin matrix has been a matter of debate. The classical theory is that the RGI and HG polymers alternate with each other as block polymers and that the side chains interact with neighbouring polysaccharide chains. Recently, this standard theory has been questioned and an argument whereby the HG polymers are actually side chains of a RGI backbone polymer has been advanced.⁴ Nevertheless, the complexity of pectin polysaccharides is such that ascribing definitive functions to this matrix of polysaccharides has proven quite difficult. The physical properties of the pectin matrix suggest a number of possible functions. The water binding properties of the galacturonic acid residues indicate that polymers containing these groups have the capacity to hydrate and swell and so possibly help maintain polymer separation in the wall.⁵ The side chains of RGI include arabinan and galactan polymers which have been shown to be highly mobile^6,7,8 with the potential to interact with each other forming a temporally entangled matrix.⁹ It is also believed that arabinan chains, which have been shown to contain ferulate residues attached to terminal arabinose groups, are able to oxidatively cross-link via the formation of diferulate bridges between arabinan chains that originate on separate RGI polysaccharides.¹⁰ The pectin matrix is now believed to contain sub-domains of RGI, HG and RGII which may interact with different polysaccharide components of the cell wall such as cellulose or xyloglucan.^11,12 Hence, it is possible that the pectin matrix may form these associations with other polysaccharides via covalent⁹ and/or non-covalent¹¹ (e.g., H-bonding) interactions and in so doing ensure the integrity of the wall and its polymer organisation. Although a number of general functions, such as hydration and ion binding, have been proposed for the pectin matrix, in particular the RGI polymer and its neutral side chains, there has been difficulty in elucidating specific functions for these polysaccharides. A number of molecular genetic studies have been performed with the aim of establishing specific functions for the RGI side chains. A recent study showed that genetic removal of the arabinan side chains in the cell walls of Nicotiana plumbaginfolia results in the formation of a non-organogenic callus culture with loosely attached cells.¹³ Furthermore, it has been shown that ‘in muro’ fragmentation of the RG1 backbone in Solanum tuberosum results in abnormal development of the periderm.¹⁴ This suggests that these side chains may play at least some role in normal cell attachment and cell development. However, the real problem is that no obvious phenotypic differences between wild type and mutant plants (in which neutral side chains have been modified) have been observed.^15,16,17 It may be that the conditions under which phenotypic differences between wild type and mutant plants would arise have not yet been investigated. We believe the water binding and attachment properties of the pectin matrix are particularly important. This is especially so given the role pectin plays in the middle lamella ensuring attachment of cells to each other and in the formation of the apoplast where water mediated transport of solutes occurs.¹ Our research has focused on a group of Southern African plants termed ‘Resurrection plants’ because of their unique ability to survive severe dehydration (desiccation) to an almost air-dry state.¹⁸ We have been interested in how the cell walls of angiosperm resurrection plants such as Craterostigma wilmsii^19,20 and Myrothamnus flabellifolia^21,22 may have become adapted to survive this extreme water deficit stress (desiccation). We have shown that in the case of the Myrothamnus flabellifolia leaf cell wall, which becomes considerably folded when dried, does not undergo dramatic changes in composition or polymer location in response to desiccation.²¹ Rather we propose that this plant has evolved a constitutively protected cell wall which is able to undergo repeated cycles of desiccation and rehydration.^21,22 We have observed that the pectin component of the leaf cell wall in this species was unusually rich in arabinose polymers, most likely arabinan and arabinogalactan in nature, which we advanced was the reason that the cell wall of this species was able to tolerate desiccation.²¹ Here we provide a simple model (Fig. 1) whereby the arabinan side chains of the pectin polysaccharides are responsible for possibly buffering/replacing the lost water during desiccation and in so doing prevent the formation of tight junctions (e.g., egg boxes) or strong H-bonding interactions between the normally separate ‘skeletal’ polysaccharides (e.g., cellulose microfibrils and xyloglucan tethers) embedded in the pectin matrix. Our model is supported by the observation that cell wall arabinans play a crucial role in the response of guard cells to turgor pressure.²³ It was shown that removal of arabinans by enzymatic digestion of leaf strips of Commelina communis resulted in locking of the guard cell walls in either the open or closed position.²³ Additional roles for arabinan polymers in cell walls have recently been implied with respect to the salt tolerance of Mesembryanthemum crystallinum,²⁴ ensuring hydration of the seed endosperm of Gleditsia triacanthos during germination²⁵ and the tolerance of tropical legume seeds to dehydration.²⁶ We believe that the arabinan side chains of RGI play a critical role in the ability of cell walls to remain flexible during plant growth and may have important functions in relation to the water content of the cell. Further studies aimed at determining the relationship between wall water content, RGI side chains and cell wall flexibility may reveal hitherto unsuspected functions for these polysaccharides in the life of the plant.Open in a separate windowFigure 1A model proposing the role of arabinose rich pectin polymers in stabilising the cell wall against water loss. (A) Pectin consisting of short arabinan chains in the hydrated state, (B) pectin consisting of short arabinan chains in the dehydrated state; (C) pectin consisting of long arabinan chains in the hydrated state; (D) pectin consisting of long arabinan chains in the dehydrated state. The likelihood of irreversible tight junctions (e.g., egg boxes) forming in arabinan poor cell walls during dehydration is demonstrated in (B) while the reversible buffering effect of arabinan rich cell walls is proposed in (D) as would possibly occur in Myrothamnus flabellifolia. For simplicity arabinan chains not participating in the buffering interactions between the RGI backbone chains have been shortened to two arabinose residues in length. Note in (A) and (B) all arabinan chains are two arabinose residues in length.     

A new callose function: Involvement in differentiation and function of fern stomatal complexes

Callose in polypodiaceous ferns performs multiple roles during stomatal development and function. This highly dynamic (1→3)-β-D-glucan, in cooperation with the cytoskeleton, is involved in: (a) stomatal pore formation, (b) deposition of local GC wall thickenings and (c) the mechanism of stomatal pore opening and closure. This behavior of callose, among others, probably relies on the particular mechanical properties as well as on the ability to form and degrade rapidly, to create a scaffold or to serve as a matrix for deposition of other cell wall materials and to produce fibrillar deposits in the periclinal GC walls, radially arranged around the stomatal pore. The local callose deposition in closing stomata is an immediate response of the external periclinal GC walls experiencing strong mechanical forces induced by the neighboring cells. The radial callose fibrils transiently co-exist with radial cellulose microfibrils and, like the latter, seem to be oriented via cortical MTs.Key words: callose, cytoskeleton, fern stomata, guard cell wall thickening, stomatal function, stomatal pore formationCallose represents a hemicellulosic matrix cell wall component, usually of temporal appearance, which is synthesized by callose synthases, enzymes localized in the plasmalemma and degraded by (1→3)-β-glucanases.^1–4 It consists of triple helices of a linear homopolymer of (1→3)-β-glucose residues.^5–7 The plant cell is able to form and degrade callose in a short time. On the surface of the plasmolyzed protoplast a thin callose surface film may arise within seconds.⁸ Callose is the only cell wall component that is implicated in a great variety of developmental plant processes, like cell plate formation,^9–11 microspore development,^12–14 trafficking through plasmodesmata,^15,16 formation and closure of sieve pores,¹⁶ response of the plant cells to multiple biotic and abiotic stresses,^4,5 establishment of distinct “cell cortex domains”,¹⁷ etc.Despite the widespread occurrence of callose, its general function(s) is (are) not well understood (reviewed in refs. ⁴ and ⁵). It may serve as: a matrix for deposition of other cell wall materials, as in developing cell plates;⁹ a cell wall-strengthening material, as in cotton seed hairs and growing pollen tubes;¹⁸ a sealing or plugging material at the plasma membrane of pit fields, plasmodesmata and sieve plate pores;¹⁶ a mechanical obstruction to growth of fungal hyphae or a special permeability barrier, as in pollen mother cell walls and muskmelon endosperm envelopes.^4,19,20 The degree of polymerization, age and thickness of callose deposits may cause variation in its physical properties.⁵Evidence accumulated so far showed that a significant number of ferns belonging to Polypodiales and some other fern classes forms intense callose deposits in the developing GC wall thickenings.^21–28 This phenomenon has not been observed in angiosperm stomata, although callose is deposited along the whole surface of the young VW and in the VW ends of differentiating and mature stomata (our unpublished data; reviewed in refs ²⁹ and ³⁰).Stomata are specialized epidermal bicellular structures (Fig. 1A) regulating gas exchange between the aerial plant organs and the external environment. Their appearance in the first land plants was crucial for their adaptation and survival in the terrestrial environment. The constituent GCs have the ability to undergo reversible changes in shape, leading to opening and closure of the stomatal pore (stomatal movement). The mechanism by which GCs change shape is based on: (a) the particular mechanical properties of GC walls owed to their particular shape, thickening, fine structure and chemical composition and (b) the reversible changes in vacuole volume, in response to environmental factors, through fairly complicated biochemical pathways.^30–33Open in a separate windowFigure 1(A) Diagrammatic representation of an elliptical stoma. (B–E) Diagram to show the process of stomatal pore formation in angiosperms (B and C) and Polypodiales ferns (D and E). The arrows in (B) indicate the forming stomatal pore. DW, dorsal wall; EPW, external periclinal wall; GC, guard cell; IPW, internal periclinal wall; ISP, internal stomatal pore; PE polar ventral wall end; VW, ventral wall.The present review is focused on the multiple-role of callose in differentiating and functioning fern stomata, as they are substantiated by the available information, including some unpublished data, and in particular in: stomatal pore formation, deposition of GC wall thickenings and opening and closure of the stomatal pore. The mode of deposition of fibrillar callose deposits in GC walls and the mechanism of their alignment are also considered.     

Directional cell migration in vivo: Wnt at the crest

Directional cell migration is essential for almost all organisms during embryonic development, in adult life and contributes to pathological conditions. This is particularly critical during embryogenesis where it is essential that cells end up in their correct, precise locations in order to build a normal embryo. Many cells have solved this problem by following a gradient of a chemoattractant usually secreted by their target tissues. Our recent research has found an alternative, complimentary, mechanism where intracellular signals are able to generate cell polarity and directional migration in absence of any external chemoattactant. We used neural crest cells to study cell migration in vivo, by performing live imagining of the neural crest cell migrating during embryo development. We show that the Planar Cell Polarity (PCP) or non-canonical Wnt signaling pathway interacts with the proteoglycan syndecan-4 to control the direction in which cell protrusions are generated, and in consequence, the direction of migration. By analyzing the activity of the small GTPases using in vivo FRET imaging we showed that PCP signaling activates RhoA, while syndecan-4 inhibits Rac, both at the back of the neural crest cell. Here we discuss a model where these signals are integrated to generate directional migration in vivo.Key words: directional migration, cell migration, syndecan-4, PCP, non-canonical Wnt, neural crest, RhoA, RacThe ability of cells to move in a directed manner is a fundamental requirement for life. In multi-cellular organisms, this requirement begins in the embryo, where morphogenetic processes are dependent on the correct movement of large numbers of cells. In the adult too, cell migration plays a vital role in many systems including the immune system and wound healing. Cell migration defects can contribute to the pathology of many diseases including vascular diseases such as atherosclerosis, and chronic inflammatory diseases like asthma and multiple sclerosis. Likewise, metastasis in cancer is characterized by mis-regulation of the normal cell migration machinery and results in cells that are normally static becoming aggressively motile and invasive.Cell migration requires cell polarization and the formation of protrusions at one end of the cell. Polarization results in a different molecular ensemble at the front of the cell compared to that at the back. Cell protrusion formation at the front of the cell requires reorganization of the actin and microtubule cytoskeleton to produce a protrusion either in the form of a broad sheet-like lamellipodium or spiky filopodium. Small GTPases are well known modulators of these processes (reviewed in ref. ¹).Several mechanism has been proposed as involved in directional migration during embryo development, such as chemotaxis (migration toward an soluble chemoattractant),² haptotaxis (migration toward a substrate-bound chemoattractant),³ population pressure (migration from a region of high towards a region of low cell density)⁴ and contact inhibition of locomotion (change in the direction of migration as a consequence of cell-cell contact),⁵ being chemotaxis the most widely accepted and studied.The correct orientation of the cell and its protrusion is the keystone of directional migration and, in the case of chemotaxis, it is supposed to be controlled by the action of external chemical cues (chemoattractants) that are produced by or near to the target tissue.⁶ One of the best examples for chemoattraction in vivo is the migration of the progenitor germ cells, which are attracted by the chemokine SDF-1.² It has been shown in vitro and in vivo, that upon receiving a chemotactic signal, the cell becomes polarized in the direction of migration. Nevertheless, it is known that cells cultured in vitro can became polarized and exhibit directional migration in absence of extrinsic chemoattractants.⁷ Pankov et al. showed that persistent directional migration in vitro can be achieved solely by modulating the activity of the small GTPase, Rac: high levels of Rac promotes the formation of peripheral lamella during random migration, while slightly lower levels of Rac suppress peripheral lamella and favour the formation of a polarized cell with lamella just at the leading edge.⁷ Is it possible that a similar mechanism of directional migration could occur in vivo?The migration of Neural Crest (NC) cells has been used as a model to study directional cell migration in vivo.^8–10 The neural crest is an embryonic population of cells that are specified at the border between the neural plate and the epidermis.¹¹ Upon induction neural crest cells undergo an epithelial to mesenchymal transition,¹² detach from the neural tube and migrate following defined pathways that eventually allow them to colonize almost the entire embryo.¹³ Finally, after reaching their destination NC cells differentiate to form many different cell types including neurons, glia, cartilage, skeleton and pigment cells.¹⁴ The migration of the NC cells is critical for the proper differentiation of their derivatives and there are several human syndromes associated with failures in this process.The migration of NC cells is a highly ordered process; individual NC cells migrate with high persistence towards the direction of their targets,⁸ but until now it was not known how this directionality is controlled. A number of molecules have been identified as key players in neural crest migration, such as Ephrins, Semaphorins, Slit/Robo, etc. (reviewed in ref. ¹³). However most of these molecules work as inhibitory signals, which are required to restrict the migration of NC cells from prohibited areas. Although chemoattraction has been one of the proposed mechanisms to explain this directional migration, no chemoattractant has thus far been found in the NC.It has been known for many years that NC cells can migrate in vitro with a high directionality even in the absence of external signals.¹⁵ Therefore, our work has been focused on understanding how NC directionality is controlled. Recently, we have unveiled some of the molecules that control this directional migration in vitro. More importantly, we have been able to show that the same molecular machinery controls directional migration in vivo.^9,10One of the key factors that controls directional migration of NC cells is the Planar Cell Polarity (PCP) or non-canonical Wnt signaling pathway.^9,10,16 PCP signaling was first described in Drosophila, where a number of mutations were identified that disrupt the formation of bristles and hairs on the adult cuticle.¹⁷ In the Drosophila wing, epithelial cells are highly polarized, with a single hair outgrowth forming at the distal end of each cell. Mutations in PCP genes cause loss in cell polarity in this tissue with hairs forming in a disorganized pattern.¹⁸ In vertebrates, PCP signaling also regulates cell polarity during a number of different developmental processes including neural tube closure, cochlear hair orientation and ciliogenesis.¹⁹We have shown that the PCP pathway is essential for correct neural crest migration in Xenopus. Injection of dominant negative forms of the intracellular PCP component Dishevelled (Dsh), which inhibit the PCP pathway but not canonical Wnt signaling, block the migration of cranial neural crest cells in vivo.⁹ Recently this role has also been extended to zebrafish where directional migration of neural crest is severely disrupted in the PCP mutant trilobite (strabismus) and in embryos injected with a dominant negative form of Dsh or a morpholino against wnt5a,¹⁰ with no effect in neural crest cell motility.^9,10 Two factors, pescadillo and syndecan-4 that have recently been proposed as modulators of the PCP signaling,^20,21 are also required for NC migration.^10,21 Taken together, these data point to an essential role for PCP signaling in neural crest migration.What is the cellular and molecular mechanism by which PCP signaling controls migration of NC cells? In order to investigate this question we analyzed the direction of neural crest migration and cell polarity in vitro and in vivo after interfering with two elements of the PCP signaling pathway: syndecan-4 and Dsh. One of the key finding of our work was that the inhibition of NC migration through syndecan-4 depletion does not affect the velocity of cell migration, but significantly reduces the directional migration of the cells in vivo (Fig. 1A and B). Consequently, when the orientation of cell protrusions was analyzed we found that syndecan-4 depletion does not affect the formation of cell protrusions, but the direction in which the cell protrusions are generated during migration. More precisely, normal cells extend their lamellipodia at the front of the cell (Fig. 1D), while cells where syndecan-4 is inhibited generate protrusion in all directions (Fig. 1E). A similar analysis was performed for embryos expressing a mutated form of Dsh that works as a dominant negative of PCP signaling and an equivalent effect on directional migration and the orientations of cell protrusions was observed (Fig. 1C and F).Open in a separate windowFigure 1Directional migration of neural crest cells. (A and B) Example of track of a single cell migrating in vivo. (A) Control cell showing persistent directional migration. (B) Cell in which the PCP signaling has been inhibited, showing absence of directional migration. (C) Cell in which syndecan-4 has been inhibited, showing no persistent migration. (D–F) Analysis of cell polarity and model of directional migration. Fn: fibronectin; Syn4: syndecan-4. (D) Control cell. Activation of Fn/Syn4 and PCP/RhoA lead to inhibition of Rac at the back of the cell, with the consequence polarization and directional migration. (E) Inhibition of PCP signaling leads to absence of RhoA activity, and in consequence an increase of Rac activity at the back of the cell. It seems that the inhibition of Rac activity by Syn4 is not sufficient to keep low levels of Rac at the back of the cells. High levels of Rac at the back produce a loss in cell polarity and in directional migration. (F) Inhibition of Syn4 generates high levels of Rac activity by a double mechanism: absence of direct inhibition of Rac and absence of RhoA which is dependent on PCP signaling. High levels of Rac at the back produce a loss of cell polarity and directional migration.As cell protrusions are known to be controlled by small GTPases and as PCP and syndecan-4 signaling regulates the activities of small GTPases,^18,22 we analyzed the activity of cdc42, RhoA and Rac after interfering with Dsh and syndecan-4. We choose to perform FRET analysis of these molecules as it is a technique that allows the visualization of their localized activity. More interestingly we succeeded in performing FRET analysis in cells migrating in vivo for the first time. Our results show that syndecan-4 inhibits Rac activity, while Dsh signaling promotes RhoA activity. In addition, we show that RhoA inhibits Rac in neural crest cells.¹⁰ The regulation of Rac by syndecan-4 is similar to that seen in other cells types in vitro.^23,24The model that emerges from these results to explain directional migration of NC cells in vivo is as follows (Fig. 1D). After delamination NC cells come into contact with fibronectin in the extracellular matrix, which is known to provide the main substrate for neural crest cells during their migration.^25,26 The interaction of fibronectin with syndecan-4 leads to two major changes in the cell: activation of PCP signaling and inhibition of Rac activity. The activated PCP signaling becomes localized at the back of the cell. From here, PCP contributes to the inhibition of Rac at the back of the cell, through the activation of RhoA. The coordinated activities of syndecan-4 and PCP signaling lead to polarised Rac activity across the cell, with Rac enriched at the leading edge, where it promotes the polymerization of actin and formation of lamellipodia, resulting in directional migration (Fig. 1D). Inhibition of PCP signaling produces high levels of Rac all over the cell as Rac, an inhibitor of RhoA in many cell types including neural crest cells, is absent (Fig. 1E). This generates cell protrusions in all directions with the consequent loss of cell polarity. If syndecan-4 is absent, the levels of Rac activity are also high all over the cell as the inhibition of Rac by syndecan-4 is absent (Fig. 1F), which also leads to a loss of cell polarity.Although detailed study of the localized activity of small GTPases has not been performed for other migratory cells in vivo, it is likely that the machinery will be similar to the one described here for NC cells. For example, it is well established in Xenopus, zebrafish and chick embryos that the migration of mesodermal cells during gastrulation requires PCP signaling.^27–29 It has also been shown that gastrulation in Xenopus²⁰ and in zebrafish (unpublished observations) requires the activity of syndecan-4. Thus, it is expected that cell polarity established during the migration of mesodermal cells will be dependent on small GTPases controlled by non-canonical Wnt signaling and syndecan-4.This novel integrated view of PCP, syndecan-4 and small GTPase activity during directional cell migration in vivo is an important advance in our knowledge of cell migration. Nevertheless, how the PCP signaling becomes activated only at the back of the cell, is a key question that needs to be answered. Future studies will be necessary to solve this and other crucial problems.     

Prions: En route from structural models to structures

The prion hypothesis^1–3 states that the prion and non-prion form of a protein differ only in their 3D conformation and that different strains of a prion differ by their 3D structure.^4,5 Recent technical developments have enabled solid-state NMR to address the atomic-resolution structures of full-length prions, and a first comparative study of two of them, HET-s and Ure2p, in fibrillar form, has recently appeared as a pair of companion papers.^6,7 Interestingly, the two structures are rather different: HET-s features an exceedingly well-ordered prion domain and a partially disordered globular domain. Ure2p in contrast features a very well ordered globular domain with a conserved fold, and—most probably—a partially ordered prion domain.⁶ For HET-s, the structure of the prion domain is characterized at atomic-resolution. For Ure2p, structure determination is under way, but the highly resolved spectra clearly show that information at atomic resolution should be achievable.Key words: prion, NMR, solid-state NMR, MAS, structure, Ure2p, HET-sDespite the large interest in the basic mechanisms of fibril formation and prion propagation, little is known about the molecular structure of prions at atomic resolution and the mechanism of propagation. Prions with related properties to the ones responsible for mammalian diseases were also discovered in yeast and funghi^8,9 which provide convenient model system for their studies. Prion proteins described include the mammalian prion protein PrP, Ure2p,¹⁰ Rnq1p,¹¹ Sup35,¹² Swi1,¹³ and Cyc8,¹⁴ from bakers yeast (S. cervisiae) and HET-s from the filamentous fungus P. anserina. The soluble non-prion form of the proteins characterized in vitro is a globular protein with an unfolded, dynamically disordered N- or C-terminal tail.^15–18 In the prion form, the proteins form fibrillar aggregates, in which the tail adopts a different conformation and is thought to be the dominant structural element for fibril formation.Fibrills are difficult to structurally characterize at atomic resolution, as X-ray diffraction and liquid-state NMR cannot be applied because of the non-crystallinity and the mass of the fibrils. Solid-state NMR, in contrast, is nowadays well suited for this purpose. The size of the monomer, between 230 and 685 amino-acid residues for the prions of Figure 1, and therefore the number of resonances in the spectrum—that used to be large for structure determination—is now becoming tractable by this method.Open in a separate windowFigure 1Prions identified today and characterized as consisting of a prion domain (blue) and a globular domain (red).Prion proteins characterized so far were found to be usually constituted of two domains, namely the prion domain and the globular domain (see Fig. 1). This architecture suggests a divide-and-conquer approach to structure determination, in which the globular and prion domain are investigated separately. In isolation, the latter, or fragments thereof, were found to form β-sheet rich structures (e.g., Ure2p(1-89),^6,19 Rnq1p(153-405)²⁰ and HET-s(218-289)²¹). The same conclusion was reached by investigating Sup35(1-254).²² All these fragements have been characterized as amyloids, which we define in the sense that a significant part of the protein is involved in a cross-beta motif.²³ An atomic resolution structure however is available presently only for the HET-s prion domain, and was obtained from solid-state NMR²⁴ (vide infra). It contains mainly β-sheets, which form a triangular hydrophobic core. While this cross-beta structure can be classified as an amyloid, its triangular shape does deviate significantly from amyloid-like structures of smaller peptides.²³Regarding the globular domains, structures have been determined by x-ray crystallography (Ure2p^25,26 and HET-s²⁷), as well as NMR (mammal prions^15,28–30). All reveal a protein fold rich in α-helices, and dimeric structures for the Ure2 and HET-s proteins. The Ure2p fold resembles that of the β-class glutathione S-transferases (GST), but lacks GST activity.²⁵It is a central question for the structural biology of prions if the divide-and-conquer approach imposed by limitations in current structural approaches is valid. Or in other words: can the assembly of full-length prions simply be derived from the sum of the two folds observed for the isolated domains?     

L1 cell adhesion molecules as regulators of tumor cell invasiveness

Fast growing malignant cancers represent a major therapeutic challenge. Basic cancer research has concentrated efforts to determine the mechanisms underlying cancer initiation and progression and reveal candidate targets for future therapeutic treatment of cancer patients. With known roles in fundamental processes required for proper development and function of the nervous system, L1-CAMs have been recently identified as key players in cancer biology. In particular L1 has been implicated in cancer invasiveness and metastasis, and has been pursued as a powerful prognostic factor, indicating poor outcome for patients. Interestingly, L1 has been shown to be important for the survival of cancer stem cells, which are thought to be the source of cancer recurrence. The newly recognized roles for L1CAMs in cancer prompt a search for alternative therapeutic approaches. Despite the promising advances in cancer basic research, a better understanding of the molecular mechanisms dictating L1-mediated signaling is needed for the development of effective therapeutic treatment for cancer patients.Key words: L1CAMs, cancer, metastasis, axon guidance, cancer stem cell, migration, invasionA major obstacle in oncology is the early diagnosis and curative therapeutic intervention of locally invasive cancers that rapidly disseminate from the primary tumor to form metastases. The standard treatment for malignant tumors consists of surgical removal of the tumor mass followed by chemo- and radiotherapy in order to eradicate the remaining cancer cells. Despite such aggressive intervention, a population of resistant cancer cells often remains intact and is thought to be the source of cancer recurrence.During the past decades, cancer basic research has focused on determining the molecular mechanisms underlying cancer initiation and progression that can provide a basis for the development of new and effective therapeutic treatments for cancer patients. An important finding was the discovery that cancer onset and development are often associated with alterations in the expression of cell adhesion molecules, which are likely to stimulate tumor cell invasiveness by signaling mechanisms that enhance cell migration.¹ The L1 family of neural cell adhesion molecules (L1-CAMs), which is comprised of four structurally related transmembrane proteins L1, CHL1, NrCAM and neurofascin (Fig. 1), is now in the spotlight of cancer research due to their upregulation in certain human tumors. L1-CAMs are transmembrane molecules of the immunoglobulin superfamily, characterized by an extracellular region of six immunoglobulin-like domains and four to five fibronectin type III repeats, followed by a highly conserved cytoplasmic domain, which is reversibly linked to the cell cytoskeleton through binding to ankyrin and ERM proteins (ezrin-radixin-moesin).² Its multi-domain structure allows complex heterophilic interactions with diverse cell receptors, although homophilic interactions also have a crucial role in L1-CAMs mediated signaling.Open in a separate windowFigure 1L1-CAMs: All have 6 Ig domains and 4–5 FN domains. The 186 kD Neurofascin isoform has a mucin-like Pro/Ala/Thr-rich (PAT) domain, while the 155 kD has only the 4 FN domains. RGD and DGEA motifs interact with integrins, while the FigQ/AY motif binds to ankyrin. ERM binding sites are indicated. The RSLE motif in L1 recruits AP2/clathrin adaptor for endocytosis.A wealth of studies has revealed L1-CAMs as pivotal components for proper development of the nervous system through regulation of cell-cell interactions. L1-CAMs have critical roles in neuronal migration and survival, axon outgrowth and fasciculation, synaptic plasticity and regeneration after trauma.² Neither CHL1 nor L1 is present on mature astrocytes, oligodendroglia or endothelial blood vessel cells in the brain, but CHL1 is upregulated in astrocytes upon injury³ and is present on oligodendroglial precursors.^4,5 During neural development, L1 plays an important role in the migration of dopaminergic neuronal cell groups in the mesencephalon and diencephalon.⁶ In the cerebellum, L1 is required for the inward migration of granule neurons from the external granular layer and cooperates with NrCAM in regulating neuronal positioning.² Similarly, CHL1 controls area-specific migration and positioning of deep layer cortical neurons in the neocortex.⁷ In addition to its role in neuronal precursor positioning, L1 plays a crucial role in axon guidance, which is governed by repellent and attractive response mechanisms directed by Ephrins and Semaphorins and their receptors (Ephs, Neuropilins, Plexins).² The importance of L1-CAMs in the development and function of the nervous system is exemplified by developmental neuropsychiatric disorders that are associated with mutation or genetic polymorphisms in genes encoding L1 (X-linked mental retardation) and CHL1 (low IQ, speech and motor delay). Polymorphisms in L1 and CHL1 genes are also associated with schizophrenia, and NrCAM gene polymorphisms are linked to autism in some populations.²Recent studies have described upregulation of L1 in a variety of tumor types. Overexpression of L1 correlates with tumor progression and metastasis in certain human gliomas,⁸ melanoma,⁹ ovarian¹⁰ and colon carcinomas.^11–13 Interestingly, L1 was found to be present only in cells at the invasive front of colon cancers but not in the tumor mass.¹² L1 is also associated with micrometastasis to both lymph nodes and bone marrow in patients bearing other cancers, suggesting a potential role in early metastatic spread.¹¹ L1 has now been pursued as both a biomarker and a powerful prognostic factor, indicative of poor outcome for patients as observed for epithelial ovarian carcinoma¹⁰ and colorectal cancer.¹¹ More recently, L1 has been shown to be overexpressed in a small fraction of glioma cells, termed glioma stem cells, which are capable of self-renewal and generate the diverse cells that comprise the tumor.¹⁴ First characterized in acute myeloid leukemia,¹⁵ cancer stem cells have been recently described in a variety of solid tumors, including breast cancer, lung cancer and gastrointestinal tumors.¹⁶ In gliomas, L1 expression was shown to be required for maintaining the growth and survival of glioma stem cells.¹⁴ These findings suggest that L1 may be implicated not only in cancer invasiveness but also in cancer survival. It will be important to determine if L1 is also upregulated in other cancer stem cells as well as to define the role of L1-mediated signaling in other cancers. Although not extensively investigated, NrCAM has also been shown to be overexpressed in glioblastoma cell lines and several cases of high grade astrocytoma¹⁷ and ependymomas.¹⁸ Studies are needed to address whether CHL1 and neurofascin play analogous roles in cancer onset and progression.The molecular mechanisms of L1-mediated signaling that govern the migration of neuronal precursors and guidance of axons during the development of the nervous system may also be used by cancer cells to facilitate invasion and cancer progression. Integrins are well-characterized cooperative partners for L1-CAMs, and signal transduction pathways activated by this complex are known to promote cell adhesion and directional motility. L1/integrin-mediated signaling may converge with growth factor signaling networks to promote motility. Like L1, CHL1 cooperates with integrins to stimulate migration. All L1-CAMs reversibly engage the actin cytoskeleton through a conserved motif FigQ/AY in the cytoplasmic domain that contains a crucial tyrosine residue required for binding the spectrin adaptor ankyrin. Phosphorylation of the FigQY tyrosine decreases ankyrin binding, whereas dephosphorylation promotes L1-ankyrin interaction. Dynamic adhesive interactions controlled by phosphorylation/dephosphorylation of the ankyrin motif in L1 family members may enable a cell to cyclically attach and detach from the ECM substrate or from neighboring cells, thus facilitating migration.¹ Another way L1 promotes cell migration is by stimulating endocytosis of integrins, reducing cell adhesion to the extracellular matrix.¹⁹ Thus, it is reasonable to speculate that upregulation of L1 in cancer may result in increased L1-mediated signaling and, consequently, increased cell migration.L1-CAMs are cleaved by metalloproteases, releasing functionally active ectodomain fragments that are laid down as “tracks” on the extracellular matrix (ECM). These fragments can cause autocrine activation of signal transduction pathways, promoting cell migration through heterophilic binding to integrins.²⁰ Specifically, L1 is cleaved constitutively or inducibly by the ADAM family metalloproteases (a disintegrin and metalloprotease) ADAM10 and ADAM17, which stimulates cell migration and neurite outgrowth during brain development.^20,21 In colon cancer, L1 colocalizes with ADAM 10 at the invasive front of the tumor tissue, suggesting that L1 shedding may play a role in cancer invasiveness.¹² Similarly, CHL1 is shed by ADAM8, which was reported to promote cell migration and invasive activity of glioma cells in vitro and is highly expressed in human brain tumors including glioblastoma multiforme, correlating with invasiveness in vivo.²² Furthermore, NrCAM, found in pancreatic, renal and colon cancers, is subject to ectodomain shedding,²³ but its function in regulating cell migration or invasion has not yet been studied.Given the newly recognized roles of L1 in tumor progression, a growing body of experimental studies has explored novel therapeutic approaches targeting L1-CAMs. Antibody-based therapeutic strategies are being pursued to functionally inhibit homophilic and heterophilic interactions of cell adhesion molecules to suppress tumor invasive motility. L1 monoclonal antibodies reduce in vivo growth of human ovarian and colon carcinoma cells in mouse xenograft models.^13,24,25 L1 targeting using lentiviral-mediated short hairpin RNA (shRNA) interference decreases growth and survival of glioma stem cells in vitro, suppresses tumor growth, and increases survival of tumor-bearing animals.¹⁴ These findings raise the possibility that L1 represents a cancer stem cell-specific therapeutic target for improving the treatment of malignant gliomas and other brain tumors. Cancer stem cells represent a potential target for future treatment of different cancer as these cells are believed to be responsible for cancer recurrence.²⁶ Promoting cancer stem cell differentiation by drug treatment could potentially reduce stem cells properties of self-renewal and proliferation, leading to inhibition of tumor growth.Inhibitors of metalloproteases that block L1-CAM shedding represent a potentially novel approach to curtailing tumor invasiveness. Chemical inhibitors of ADAMS are appealing for glioma therapy due to their diffusability, which circumvents blood-brain barrier limitations. Another novel approach involves the secreted axon repellent protein, Semaphorin 3A (Sema3A). L1-CAMs serve as co-receptors for Sema3A by cis binding in the plasma membrane to Neuropilin-1, important for repellent axon guidance.² Interestingly, Sema3A inhibits invasiveness of prostate cancer cells²⁷ and migration and spreading of breast cancer cells in in vitro assays,²⁸ and thus may also be mediated by L1-CAMs. Such an approach could be potentially useful in mitigating invasion of cancer cells in gliomas and other tumors that are known to express L1 and Neuropilins. However, effective strategies for some types of cancer can promote cancer progression in other types. For example, Sema3A has been shown to contribute to the progression of pancreatic cancer²⁹ and colon cancer.³⁰ Thus, it is imperative that the molecular mechanisms underlying L1-mediated signaling are understood in a tissue specific manner. Despite the promising advances in cancer basic research, much more research is needed to better design strategies for cancer therapy.     

MEK1/2 and p38-like MAP kinase successively mediate H2O2 signaling in Vicia guard cell

As a second messenger, H₂O₂ generation and signal transduction is subtly controlled and involves various signal elements, among which are the members of MAP kinase family. The increasing evidences indicate that both MEK1/2 and p38-like MAP protein kinase mediate ABA-induced H₂O₂ signaling in plant cells. Here we analyze the mechanisms of similarity and difference between MEK1/2 and p38-like MAP protein kinase in mediating ABA-induced H₂O₂ generation, inhibition of inward K⁺ currents, and stomatal closure. These data suggest that activation of MEK1/2 is prior to p38-like protein kinase in Vicia guard cells.Key words: H₂O₂ signaling, ABA, p38-like MAP kinase, MEK1/2, guard cellAn increasing number of literatures elucidate that reactive oxygen species (ROS), especially H₂O₂, is essential to plant growth and development in response to stresses,^1–4 and involves activation of various signaling events, among which are the MAP kinase cascades.^1–3,5 Typically, activation of MEK1/2 mediates NADPH oxidase-dependent ROS generation in response to stresses,^4,6–8 and the facts that MEK1/2 inhibits the expression and activation of antioxidant enzymes reveal how PD98059, the specific inhibitor of MEK1/2, abolishes abscisic acid (ABA)-induced H₂O₂ generation.^6,8,9 It has been indicated that PD98059 does not to intervene on salicylic acid (SA)-stimulated H₂O₂ signaling regardless of SA mimicking ABA in regulating stomatal closure.^2,6,8,10 Generally, activation of MEK1/2 promotes ABA-induced stomatal closure by elevating H₂O₂ generation in conjunction with inactivating anti-oxidases.Moreover, activation of plant p38-like protein kinase, the putative counterpart of yeast or mammalian p38 MAP kinase, has been reported to participate in various stress responses and ROS signaling. It has been well documented that p38 MAP kinase is involved in stress-triggered ROS signaling in yeast or mammalian cells.^11–13 Similar to those of yeast and mammals, many studies showed the activation of p38-like protein kinase in response to stresses in various plants, including Arabidopsis thaliana,^14–16 Pisum sativum,¹⁷ Medicago sativa¹⁸ and tobacco.¹⁹ The specific p38 kinase inhibitor SB203580 was found to modulate physiological processes in plant tissues or cells, such as wheat root cells,²⁰ tobacco tissue²¹ and suspension-cultured Oryza sativa cells.²² Recently, we investigate how activation of p38-like MAP kinase is involved in ABA-induced H₂O₂ signaling in guard cells. Our results show that SB203580 blocks ABA-induced stomatal closure by inhibiting ABA-induced H₂O₂ generation and decreasing K⁺ influx across the plasma membrane of Vicia guard cells, contrasting greatly with its analog SB202474, which has no effect on these events.^23,24 This suggests that ABA integrate activation of p38-like MAP kinase and H₂O₂ signaling to regulate stomatal behavior. In conjunction with SB203580 mimicking PD98059 not to mediate SA-induced H₂O₂ signaling,^23,24 these results generally reveal that the activation of p38-like MAP kinase and MEK1/2 is similar in guard cells.On the other hand, activation of p38-like MAP kinase^23,24 is not always identical to that of MEK1/2^8,25 in ABA-induced H₂O₂ signaling of Vicia guard cells. For example, H₂O₂^- and ABA-induced stomatal closure was partially reversed by SB203580. The maximum inhibition of both regent-induced stomatal closure were observed at 2 h after treatment with SB203580, under which conditions the stomatal apertures were 89% and 70% of the control values, respectively. By contrast, when PD98059 was applied together with ABA or H₂O₂, the effects of both ABA- and H₂O₂-induced stomatal closure were completely abolished (Fig. 1). These data imply that the two members of MAP kinase family are efficient in H₂O₂-stimulated stomatal closure, but p38-like MAP kinase is less susceptive than MEK1/2 to ABA stimuli.Open in a separate windowFigure 1Effects of SB203580 and PD98059 on ABA- and H₂O₂-induced stomatal closure. The experimental procedure and data analysis are according to the previous publication.^8,23,24It has been reported that ABA or NaCl activate p38 MAP kinase in the chloronema cells of the moss Funaria hygrometrica in 2∼10 min.²⁶ Similar to this, SB203580 improves H₂O₂-inhibited inward K⁺ currents after 4 min and leads it to the control level (100%) during the following 8 min (Fig. 2). However, the activation of p38-like MAP kinase in response to ABA need more time, and only recovered to 75% of the control at 8 min of treatment (Fig. 2). These results suggest that control of H₂O₂ signaling is required for the various protein kinases including p38-like MAP kinase and MEK1/2 in guard cells,^1,2,8,23,24 and the ABA and H₂O₂ pathways diverge further downstream in their actions on the K⁺ channels and, thus, on stomatal control. Other differences in action between ABA and H₂O₂ are known. For example, Köhler et al. (2001) reported that H₂O₂ inhibited the K⁺ outward rectifier in guard cells shows that H₂O₂ does not mimic ABA action on guard cell ion channels as it acts on the K⁺ outward rectifier in a manner entirely contrary to that of ABA.²⁷Open in a separate windowFigure 2Effect of SB203580 on ABA- and H₂O₂-inhibited inward K⁺ currents. The experimental procedure and data analysis are according to the previous publication.²⁴ SB203580 directs ABA- and H₂O₂-inactivated inward K⁺ currents across plasma membrane of Vicia guard cells. Here the inward K⁺ currents value is stimulated by −190 mV voltage.Based on the similarity and difference between PD98059 and SB203580 in interceding ABA and H₂O₂ signaling, we speculate the possible mechanism is that the member of MAP kinase family specially regulate signal event in ABA-triggered ROS signaling network,^1–4 and the signaling model as follows (Fig. 3).Open in a separate windowFigure 3Schematic illustration of MAP kinase-mediated H₂O₂ signaling of guard cells. The arrows indicate activation. The line indicates enhancement and the bar denotes inhibition.