Structural basis of transmembrane domain interactions in integrin signaling

Cell surface receptors of the integrin family are pivotal to cell adhesion and migration. The activation state of heterodimeric αβ integrins is correlated to the association state of the single-pass α and β transmembrane domains. The association of integrin αIIbβ3 transmembrane domains, resulting in an inactive receptor, is characterized by the asymmetric arrangement of a straight (αIIb) and tilted (β3) helix relative to the membrane in congruence to the dissociated structures. This allows for a continuous association interface centered on helix-helix glycine-packing and an unusual αIIb(GFF) structural motif that packs the conserved Phe-Phe residues against the β3 transmembrane helix, enabling αIIb(D723)β3(R995) electrostatic interactions. The transmembrane complex is further stabilized by the inactive ectodomain, thereby coupling its association state to the ectodomain conformation. In combination with recently determined structures of an inactive integrin ectodomain and an activating talin/β complex that overlap with the αβ transmembrane complex, a comprehensive picture of integrin bi-directional transmembrane signaling has emerged.Key words: cell adhesion, membrane protein, integrin, platelet, transmembrane complex, transmembrane signalingThe communication of biological signals across the plasma membrane is fundamental to cellular function. The ubiquitous family of integrin adhesion receptors exhibits the unusual ability to convey signals bi-directionally (outside-in and inside-out signaling), thereby controlling cell adhesion, migration and differentiation.^1–5 Integrins are Type I heterodimeric receptors that consist of large extracellular domains (>700 residues), single-pass transmembrane (TM) domains, and mostly short cytosolic tails (<70 residues). The activation state of heterodimeric integrins is correlated to the association state of the TM domains of their α and β subunits.^6–10 TM dissociation initiated from the outside results in the transmittal of a signal into the cell, whereas dissociation originating on the inside results in activation of the integrin to bind ligands such as extracellular matrix proteins. The elucidation of the role of the TM domains in integrin-mediated adhesion and signaling has been the subject of extensive research efforts, perhaps commencing with the demonstration that the highly conserved GFFKR sequence motif of α subunits (Fig. 1), which closely follows the first charged residue on the intracellular face, αIIb(K989), constrains the receptor to a default low affinity state.¹¹ Despite these efforts, an understanding of this sequence motif had not been reached until such time as the structure of the αIIb TM segment was determined.¹² In combination with the structure of the β3 TM segment¹³ and available mutagenesis data,^6,9,10,14,15 this has allowed the first correct prediction of the overall association of an integrin αβ TM complex.¹² The predicted association was subsequently confirmed by the αIIbβ3 complex structure determined in phospholipid bicelles,¹⁶ as well as by the report of a similar structure based on molecular modeling using disulfide-based structural constraints.¹⁷ In addition to the structures of the dissociated and associated αβ TM domains, their membrane embedding was defined^{12,13,16,18,19} and it was experimentally recognized that, in the context of the native receptor, the TM complex is stabilized by the inactive, resting ectodomain.¹⁶ These advances in integrin membrane structural biology are complemented by the recent structures of a resting integrin ectodomain and an activating talin/β cytosolic tail complex that overlap with the αβ TM complex,^20,21 allowing detailed insight into integrin bi-directional TM signaling.Open in a separate windowFigure 1Amino acid sequence of integrin αIIb and β3 transmembrane segments and flanking regions. Membrane-embedded residues^{12,13,16,18,19} are enclosed by a gray box. Residues 991–995 constitute the highly conserved GFFKR sequence motif of integrin α subunits.     

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.     

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?     

Numb: A new player in EMT

Epithelial to mesenchymal transition (EMT) is a critical event in embryogenesis and plays a fundamental role in cancer progression and metastasis.¹ Numb has been shown to play an important role in the proper functions of Par protein complex and in cell-cell junctions,^2,3 both of which are associated with EMT.^4,5 However, the role of Numb in EMT has not been fully elucidated. Recently, we showed that Numb is capable of binding to both Par3 and E-cadherin. Intriguingly, the interaction of Numb with E-cadherin or the Par protein complex is dynamically regulated by tyrosine phosphorylation induced by HGF or Src. Knockdown of Numb by shRNA in MDCK cells led to a lateral to apical translocation of E-cadherin and β-catenin, active F-actin polymerization, mis-localization of Par3 and aPKC, a decrease in cell-cell adhesion and an increase in cell migration and proliferation. These data suggest a diverse role for Numb in regulating cell-cell adhesion, polarity and migration during EMT.⁶Key words: Numb, E-cadherin, tyrosine kinase, cell polarity, adhesion, EMTNumb was originally identified as a gene required for cell fate determination during neuroblast division and sensory organogenesis.⁷ Recently, a number of proteins involved in cell polarity, cell-cell adhesion and tumorigenesis have been identified as binding partners for Numb. These include the Par3-Par6-aPKC polarity complex, E-cadherin, integrin, Notch, WNT and p53.^8–10 Although these new data implicate Numb in multiple signaling pathways, questions remain as to how the various interactions are regulated and in which biological context they occur. Interestingly, most binding partners of Numb are involved in one way or another in the onset and/or progression of cancer. For instance, the Par complex is involved in regulating the formation and stability of tight junction whereas E-cadherin is a key component of the adherens junction in epithelial cells. Understandably, deregulation of the Par protein complex and/or E-cadherin is implicated in EMT. A variety of stimuli have been identified to induce EMT,¹ including transforming growth factor-β (TGFβ), hepatocyte growth factor (HGF), fibroblast growth factor (FGF) and activation of tyrosine kinase Src. During the progression of EMT, non-motile epithelial cells gradually lose their apical-basal polarity and cell-cell junctions and become mesenchymal cells with an ability to migrate away from the primary site to surrounding tissues.¹ Therefore, the study of Numb interactions with the Par protein complex and E-cadherin in the context of EMT provides a good point of entry to decode the complex signaling network mediated by Numb.Based on data obtained from Madin-Darby canine kidney (MDCK) cells,⁶ we propose a model in which Numb regulates epithelial polarity and cell-cell adhesion in EMT (Fig. 1). In epithelial cells, Numb binds to E-cadherin or the Par protein complex via Par3 under a normal physiological condition to stabilize adherens and tight junctions. Under the influence of an extracellular cue, such as HGF, the Met receptor recruits and activates one of its downstream components, c-Src. C-Src subsequently phosphorylates the DNVYYY motif on E-cadherin, resulting in Numb dissociation from phosphorylated E-cadherin. Numb then binds and sequesters phosphorylated aPKC and Par6, while phosphorylated Par3 is released from the Par complex. The Numb-aPKC-Par6 complex remains on the plasma membrane or in the cytoplasm, whereas Par3 is transported into the nucleus (by an unknown mechanism). Phosphorylated E-cadherin is relocated to an apico-lateral domain accompanied with active F-actin polymerization. Enhanced F-actin polymerization, together with reduced cell-cell adhesion and increased cell proliferation, promotes cell migration (Fig. 1).Open in a separate windowFigure 1A model depicting the role of Numb in epithelial-mesenchymal transition (EMT). In epithelial cells, Numb stabilizes both E-cadherin based adherens junctions through a PTB domain binding to NVYY motif on E-cadherin, and Par protein complex on tight junctions by interacting with Par3. During the early stage of EMT, c-Met recruits the tyrosine kinase c-Src upon its activation by HGF. C-Src phosphorylates E-cadherin on the NVYY motif, which leads to dissociation of Numb from E-cadherin and the translocation of E-cadherin to an apical domain. Numb forms a complex with phosphorylated aPKC and Par6, whereas phosphorylated Par3 is transported into the nucleus. Enhanced F-actin polymerization, together with reduced cell-cell adhesions, promotes the transition to mesenchymal cells.While our work in MDCK epithelial cells identifies a key role for Numb in regulating the sub-cellular localizationsand functions of E-cadherin and the Par protein complex, it raises a number of unaddressed questions. First, biochemical data obtained from temperature sensitive src mutant (ts-src) MDCK cell line suggest that tryosine phosphorylation plays a critical role in regulating the dynamic interactions of Numb with E-cadherin or the Par protein complex. Using a peptide array screen, we found that a conserved DNVYYY motif in E-cadherin is the binding site for phosphotyrosine binding (PTB) domain of Numb. Interestingly, peptide analogs in which a Tyr residue in the YYY triad is replaced by a pTyr were deficient in binding. This suggests that the Numb PTB domain may act in a phosphotyrosine-independent manner in EMT signal transduction events, and that the interaction of Numb with VIEW E-cadherin is negatively regulated by tyrosine kinase signaling. An intriguing question is why E-caderin harbors a conserved YYY triad⁶ when phosphorylation of one Tyr is sufficient to eliminate Numb binding. Furthermore, when and where are the YYY triad phosphorylated during EMT? It is likely that one or more tyrosine residues are phosphorylated by Src, depending on the duration of stimulus that a cell receives during EMT. Phosphorylated E-cadherin may be targeted to an alternative location in the cell by the E3 ligase Hakai which mediates the endocytosis and degradation of E-cadherin¹¹ after its phosphorylation and dissociation from Numb. To complicate the issue, Par3 and aPKC themselves are also shown to be tyrosine phosphorylated by HGF treatment or Src activation in our studies, although the precise phosphorylation sites on either protein remain to be determined. In a previous report, a high-throughput phosphoproteomic screen has identified multiple tyrosine phosphorylation sites in the carboxyl terminus of Par-3.¹² Phosphorylation of Y1127 in Par-3 reduced its interaction with LIMK2 and delayed tight junction assembly in mammalian epithelial cells with a constitutive Src activation or under EGF treatment.¹² A recent study, which shows that the SH2 domain of C-terminal Src kinase (Csk) binds to the Y1127 site on Par3, adds another layer of complexity to the scenario.¹³ In the same vein, Src has been shown to associate with aPKC and phosphorylate the latter in PC12 cells.¹⁴ Par6 appears also to be subjected to Src regulation as recruitment of Src by Pals leads to Pals1 and Par6 to be sequestered away from JAM-C of desmosome adhesions in the blood-testis-barrier, causing a disruption in cell adhesion.¹⁵ It will be of interest to determine whether Par6 is directly phosphorylated by Src in this case.¹⁵ Together, these studies suggest a role for tyrosine phosphorylation, besides serine/threonine phosphorylation, in regulating the functions of various polarity proteins.Second, our biochemical and immunfluorescence data suggest that the interactions of Numb with E-cad/Par complex are spatially and temporally regulated in response to HGF treatment. However, the molecular mechanisms for some remarkable phenotypes remain poorly defined. For instance, shRNA-mediated Numb knockdown caused a dramatic apicolateral mis-localization of E-cadherin and β-catenin. Dynamic cellular localization may be regulated by endocytosis and post-translational modifications such as phosphorylation. In keeping with this, Numb can directly bind to several endocytic proteins, including AP2 and Eps15, through its conserved DPF and NPF motifs.¹⁶ Our data suggest that Numb is responsible for targeting E-cadherin to correct localization on the basolateral membrane, but we cannot rule out the possibility that other proteins may be involved in this process. One candidate that may facilitate E-cadherin localization is Rab11. Previous studies revealed that in newly polarized MDCKII cells, Rab11 mutation causes an apical mis-localization of E-cadherin and aberrant actin localization, while leaving ZO-1 localization unchanged. This mirrors the phenotype we observed in the Numb shRNA cells.¹⁷ In support of the notion that Rab11 and Numb may be functionally related, Rab11 and Numb are segregated selectively in the pIIb daughter cell during the division of a sensory organ precursor (SOP).¹⁸ It is likely that the mis-localization of E-cadherin is caused by an abnormal enodocytosis of E-cadherin in the absence of Numb.In contrast, the Par protein complex provides critical spatial information in the formation of tight junctions. Treatment of MDCK cells by HGF caused Par3 to dissociate from Par6-aPKC, suggesting that the Par3-Par6-aPKC complex¹⁹ is dynamically regulated. Additionally, both Par3 and aPKC are found to translocate to the nucleus following HGF treatment. A key question, therefore, is how the localizations of these polarity proteins themselves are regulated during EMT. Through a PB1-PB1 domain heterodimarization,²⁰ Par-6 binds to aPKC and thereby inhibits its catalytic. Moreover, Par6 also helps recruit substrates to aPKC,²⁰ one of which is Par3. Par6 binds to Par3, through a PDZ-PDZ interaction, but aPKC can also bind directly to Par3 through its kinase domain, and phosphorylates a Ser residue on Par3.²¹ Importantly, depending on the cellular context, Par3, Par6 and aPKC may not form a constitutive complex.²⁰ For example, in Drosophila neuroblasts and embryonic epithelial cells, Par3 apical localization is independent of aPKC,²² while in mammalian epithelial cells, Par3 is not apical but is associated with tight junctions.²³ Interactions of these polarity proteins are dynamically regulated by multiple protein kinases, small GTPases, or competition from other binding partners.¹⁹ The dynamic change of components of the Par complex leads to the different sub-cellular distribution of these individual polarity proteins, and thereby alters their functions.¹⁹ We showed that activation of Src kinase or HGF treatment reduces the association of Par3 with aPKC, but does not change the binding between aPKC and Par6. In support of this, a similar observation is made in MDCK cells with activation of tyrosine kinase ErbB2, although neither Par3 nor Par6 is a substrate of ErbB2.²⁴ The dynamic assembly and dissolution of the Par3-Par6-aPKC complex in response to intracellular cues or extracellular stimuli may be a general mechanism used by metazoan to control cell polarity and movement.Lastly, active F-actin polymerization is one of most striking morphologies we observed in the Numb-shRNA cells. Given that the Numb-shRNA cells exhibit a faster rate of migration and wound healing than control cells, it is likely that the active F-actin polymerization is caused by aberrant activation of the Rho family GTPase activity. Small GTPases of the Rho family control organization of cytoskeleton, cell motility, cell growth, morphogenesis, cytokinesis and trafficking.²⁵ The most common members of small Rho GTPases are RhoA, Rac1 and Cdc42. RhoA is responsible for the activation of stress fibers and cell contractility.²⁵ Rac1 activation leads to polymerization of filamentous actin, which results in lamellipodium formation and membrane ruffling at the leading edge of migrating cells.²⁵ Cdc42 activation causes the formation of filopodia, long finger-like protrusions at the edges of lamellipodia.²⁵ A Rho family of GTPase exists in two states: a GDP-bound inactive state and a GTP-bound active state. The switch between the two states is regulated by a large group of guanine nucleotide exchange factors (GEFs) and GTPase activating proteins (GAPs).²⁵ GEFs and GAPs typically contain interaction domains which direct the enzymes to specific sub-cellular locations and help recruit upstream/downstream partners to affect processes such as actin cytoskeleton, cell polarity, microtubule dynamics and membrane transport.²⁵ It is often difficult to pinpoint exactly which member of the Rho family GTPases leads to a specific phenotype due to the overlapping functions of the Rho GTPase members. However, several observations have suggested that certain small Rho GTPases are essential for the establishment of the apico-basal polarity and likely interplay with Numb. Deletion of Cdc42 abolished normal localization of aPKC, Par6 and Numb in neuroepithelium.²⁶ Par6 binds to GTP-bound Cdc42 through a Rho GTPase-bindingCdc42/Rac-interactive binding (CRIB) domain.^23,27 Another line of evidence is that Numb binds to intersectin, a Cdc42 GEF, and enhances the GEF activity of intersectin, leading to activation of Cdc42 in vivo.²⁸ It appears that Numb both regulates and is regulated by Cdc42 temporally and spatially. Par3 has also been reported to sequester Tiam1, a Rac GEF, to inhibit Rac activation in hippocampal neurons.²⁹ Nevertheless, Par3 recruits Tiam1 to activate Rac in keratinocytes.³⁰ Thus, the dynamic interplay between Rho GTPases and the Par protein complex may be cell type-dependent. It will also be of interest in the future to determine whether other GEFs/GAPs interact with the Par protein complex, and whether and how these interactions are modulated by Numb.A more detailed understanding of the role of Numb in EMT will undoubtedly provide valuable insights into the molecular basis of cancer metastasis and suggest novel therapeutic strategies for cancer.     

v-Crk regulates membrane dynamics and Rac activation

Cell migration is an integrated process that involves cell adhesion, protrusion and contraction. We recently used CAS (Crk-associated substrate, 130CAS)-deficient mouse embryo fibroblasts (MEFs) to examined contribution made to v-Crk to that process via its interaction with Rac1. v-Crk, the oncogene product of avian sarcoma virus CT10, directly affects membrane ruffle formation and is associated with Rac1 activation, even in the absence of CAS, a major substrate for Crk. In CAS-deficient MEFs, cell spreading and lamellipodium dynamics are delayed; moreover, Rac activation is significantly reduced and it is no longer targeted to the membrane. However, expression of v-Crk by CAS-deficient MEFs increased cell spreading and active lamellipodium protrusion and retraction. v-Crk expression appears to induce Rac1 activation and its targeting to the membrane, which directly affects membrane dynamics and, in turn, cell migration. It thus appears that v-Crk/Rac1 signaling contributes to the regulation of membrane dynamics and cell migration, and that v-Crk is an effector molecule for Rac1 activation that regulates cell motility.Key words: v-Crk, Rac, lamellipodia dynamics, cell migration, p130CASCell migration is a central event in a wide array of biological and pathological processes, including embryonic development, inflammatory responses, angiogenesis, tissue repair and regeneration, cancer invasion and metastasis, osteoporosis and immune responses.^1,2 Although the molecular basis of cell migration has been studied extensively, the underlying mechanisms are still not fully understood. It is known that cell migration is an integrated process that involves formation of cell adhesions and/or cell polarization, membrane protrusion in the direction of migration (e.g., filopodium formation and lamellipodium extension), cell body contraction and tail detachment.^1–3 Formation of cell adhesions, including focal adhesions, fibrillar adhesions and podosomes are the first step in cell migration. Cell adhesions are stabilized by attachment to the extracellular matrix (ECM) mediated by integrin transmembrane receptors, which are also linked to various cytoplasmic proteins and the actin cytoskeleton, which provide the mechanical force necessary for migration.^2,4 The next steps in the process of cell migration are filopodium formation and lamellipodium extension. These are accompanied by actin polymerization and microtubule dynamics, which also contribute to the control of cell adhesion and migration.⁵Focal adhesions are highly dynamic structures that form at sites of membrane contact with the ECM and involve the activities of several cellular proteins, including vinculin, focal adhesion kinase (FAK), Src family kinase, paxillin, CAS (Crk-associated substrate, p130CAS) and Crk.⁶ A deficiency in focal adhesion protein is associated with the severe defects in cell motility and results in embryonic death. For example, FAK deficiency disrupts mesoderm development in mice and delays cell migration in vitro,⁷ which reflects impaired assembly and disassembly the focal adhesions.⁸ In addition, mouse embryonic fibroblasts (MEFs) lacking Src kinase showed a reduced rate of cell spreading that resulted in embryonic death.⁹ Taken together, these findings strongly support the idea that cell adhesion complexes play crucial roles in cell migration.CAS is a hyperphosphorylated protein known to be a major component of focal adhesion complexes and to be involved in the transformation of cells expressing v-Src or v-Crk.¹⁰ CAS-deficient mouse embryos die in utero and show marked systematic congestion and growth retardation,⁴ while MEFs lacking CAS show severely impaired formation and bundling of actin stress fibers and delayed cell motility.^4,11,12 Conversely, transient expression of CAS in COS7 cells increases cell migration.¹¹ Crk-null mice also exhibit lethal defects in embryonic development,¹³ which is consistent with the fact that CAS is a major substrate for v-Crk, and both CAS and v-Crk are necessary for induction of cell migration.¹⁴ v-Crk consists of a viral gag sequence fused to cellular Crk sequences, which contain Src homology 2 (SH2) and SH3 domains but no kinase domain, and both CAS and paxillin bind to SH2 domains.^12,15,16 Despite the absence of a kinase domain, cell expressing v-Crk show upregulation of tyrosine phosphorylation of CAS, FAK and paxillin, which is consistent with v-Crk functioning as an adaptor protein.¹⁷ Moreover, this upregulation of tyrosine phosphorylation correlates well with the transforming activity of v-Crk.¹⁷ By contrast, tyrosine phosphorylation of FAK and CAS is diminished in Src kinase-deficient cells expressing v-Crk, and they are not targeted to the membrane, suggesting v-Crk signaling is Src kinase-dependent. After formation of the CAS/v-Crk complex, v-Crk likely transduces cellular signaling to Src kinase and FAK.¹² Notably, tyrosine phosphorylation of FAK and cell migration and spreading are all enhanced when v-Crk is introduced into CAS-deficient MEFs.¹² We therefore suggest that v-Crk activity, but not cellular Crk activity, during cell migration and spreading is CAS-independent.Membrane dynamics such as lamellipodium protrusion and membrane ruffling reportedly involve Rac1,¹⁸ α4β1 integrin,¹⁹ Arp2/3,⁶ and N-WASP,²⁰ and are enhanced in v-Crk-expressing CAS-deficient MEFs.²¹ Moreover, expression in those cells of N17Rac1, a dominant defective Rac1 mutant, abolished membrane dynamics at early times and delayed cell migration.²¹ v-Crk-expressing, CAS-deficient MEFs transfected with N17Rac1 did not begin spreading until one hour after being plated on fibronectin, and blocking Rac activity suppressed both membrane dynamics and cell migration. We therefore suggest that v-Crk is involved in cell attachment and spreading, and that this process is mediated by Rac1 activation. In addition, v-Crk expression apparently restores lamellipodium formation and ruffle retraction in CAS-deficient MEFs. Thus v-Crk appears to participate in a variety cellular signaling pathways leading to cell spreading, Rac1 activation, membrane ruffling and cell migration, even in the absence of CAS, its major substrate protein.In fibroblasts, the Rho family of small GTP-binding proteins (e.g., Cdc42, Rac and Rho) functions to control actin cytoskeleton turnover, including filopodium extension, lamellipodium formation and generation of actin stress fibers and focal adhesions.²² These GTPases function in a cascade, such that activation of Cdc42 leads to activation of Rac1, which in turn activates Rho.²² Once activated, Rho controls cell migration. Cell adhesion to ECM leads to the translocation of Rac1 and Cdc42 from the cytosol to the plasma membrane,²³ where they regulate actin polymerization at the leading edge.^19,24 Dominant negative Rac and Cdc42 mutants inhibit the signaling to cell spreading initiated by the interaction of integrin with ECM.²⁴ The fact that cellular levels of activated Rac are higher in cells adhering to ECM than in suspended cells further suggests that activation of Rac and Cdc42 is a critical step leading to membrane protrusion and ruffle formation. It is noteworthy in this regard that v-Crk is able to induce Rac activation and its translocation to plasma membrane.²¹Overall, the findings summarized in this article demonstrate that v-Crk participates in several steps leading to cell adhesion and spreading (Fig. 1), and the targeting of v-Crk to focal adhesion sites appears to be a prerequisite for regulation of cell migration and spreading via Rac activation. To fully understand its function, however, it will be necessary to clarify the role of v-Crk in Rac1 and Cdc42 activation initiated by integrin-ECM interactions.Open in a separate windowFigure 1Schematic diagram of v-Crk signaling in MEFs. Cell adhesion signaling initiated by the integrin-ECM interaction triggers v-Crk signaling mediated by Src kinase, after which focal adhesion proteins are tyrosine phosphorylated. These events lead to translocation of Rac from the cytosol to the membrane, where it promotes membrane protrusion and ruffle formation. Under basal conditions, Rac is bound with GDP and is inactive. Upon stimulation, Rac activation is mediated by guanine nucleotide exchange factors (GEFs) that stimulate the release of bound GDP and the binding of GTP. Activation of Rac is transient, however, as it is inactivated by GTPase activating protein (GAP).     

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.     

Significance of microtubule catastrophes at focal adhesion sites

Directional cell migration requires cell polarization and asymmetric distribution of cell signaling. Focal adhesions and microtubules are two systems which are essential for these. It was shown that these two systems closely interact with each other. It is known that microtubule targeting stimulates focal adhesion dissociation. Our recent study shows that focal adhesions, in turn, specifically induce microtubule catastrophe via a biochemical mechanism. We were able to track down one of the focal adhesion proteins paxillin which is involved in this process. Paxillin phosphorylation was previously shown to be the key component in the regulation of focal adhesion assembly or disassembly. Since microtubule catastrophe dynamic differs at the leading edge and cell rear, similar to paxillin phosphorylation levels, we suggest a model connecting asymmetric distribution of focal adhesions and asymmetric distribution of microtubule catastrophes at adhesion sites as a feedback loop.Key words: microtubule catastrophe, focal adhesion, microtubules, paxillin, cell motilityCell migration is important for many biological processes. It requires organized asymmetric dynamics of focal adhesions (FAs), sites where cells interact with extra cellular matrix. FAs appear at the leading edge as small transient dot-like structures termed focal complexes (FXs).^1,2 FX assembly and disassembly is regulated by phosphorilation status of paxillin a major FX protein.^3,4 Most of FXs form and disassemble rapidly. However, some adhesions mature in a force-dependent manner, into larger late adhesions. This process, involves both an increase in size and change in molecular composition^3,5 and is accompanied by a reduction in local paxillin phosphorylation.⁴ Late adhesions are more stable, immobile and undergo forced disassembly by multiple microtubule targeting events⁶ only underneath the approaching cell body or transform into fibrillar adhesions by a Src-dependent mechanism.⁷Similarly to the leading edge, proper adhesion patterns at the cell rear are also essential. Most trailing adhesions are initiated in protrusions at the rear and flanks of the cell as FX rapidly mature in response to tension and transform into sliding trailing adhesions.⁸ The process of sliding is complex. While adhesion proteins coupled with the actin cytoskeleton can be translocated relative to substratum, those that are associated with the membrane are thought to undergo treadmilling within the adhesion site.^9,10 Treadmilling, which includes disassembly of adhesion proteins at the distal end and reassembly at the proximal end,¹⁰ is accompanied by fusion with new adhesions formed in front of the sliding one.⁶ Thus, despite a protein composition similar to late adhesions, sliding adhesions are more dynamic. Not surprisingly, sliding adhesions have high paxillin phosphorylation at the distal end of the adhesion site, indicating very dynamic assembly/disassembly rates.⁴Several mechanisms have been proposed for the regulation of adhesion turnover (reviewed in ref. ¹¹). However, these have not accounted for the observed asymmetry of adhesion turnover. Understanding this requires examining the connection with another asymmetric intracellular system, the microtubule network. This dynamic network closely interacts with FAs. Microtubules play an essential role in cell migration and polarized distribution of signals within the cell. Multiple microtubule targeting to FA leads to their disassembly both at the leading edge and at the cell rear.⁶Unlike microtubule growth in other cell regions, growth at its leading edge is persistent, characterized by short periods of shrinkage.⁸ Simultaneous observation of microtubules and FAs show that microtubules specifically target adhesion sites.¹² More detailed analysis of microtubule dynamics reveals that FAs are preferable sites for microtubule catastrophes.¹³ Although FAs cover only about 5% of cell area more than 40% of catastrophes occur at these sites. The likelihood of microtubule catastrophe is seven times higher when a microtubule grows through a FA rather than through an adhesion-free area¹³ and about 90% of microtubules approaching adhesion sites undergo catastrophe. Although most of the catastrophes occur at late adhesions, due to their increased stability and lifespan, there is no difference in efficiency of catastrophe induction between small focal complexes and large rigid late adhesions.¹³ As FX do not have dense adhesion or actin plaque, it is likely that microtubule catastrophe is triggered by a biochemical mechanism rather than mechanical rigidity. This is also supported by the fact that mechanical obstacles in a cell do not necessarily cause microtubule catastrophe.¹³At the cell rear, microtubule dynamics differ from those at the leading edge. Microtubules spend less time in a growing phase and more time in pauses and shrinkage.⁸ Polymerization and depolymerization occur within a very limited area close to the cell edge.⁸ Live-cell imaging of cells expressing both microtubule and focal adhesion markers show that this complex dynamic sequence often happens within a single sliding adhesion. Microtubules that are captured at the proximal end of adhesion undergo multiple repetitive catastrophes at the distal end (Fig. 1) accompanied by rescue at the capture site. Thus, the capture mechanism significantly increases the lifetime of a microtubule and ensures that repetitive catastrophes occur at the single adhesion. This scenario leads to high catastrophe frequency at the cell rear, resulting in intensive catastrophe-dependent regulation in this cell region.Open in a separate windowFigure 1Multiple microtubule catastrophes at the sliding adhesion. (A) Frame from TIRF video sequence of a fish fibroblast cell (CAR) co-transfected with GFP-tubulin (green) to visualize microtubules and Cherry-Zyxin (red) to mark focal adhesions. The boxed region is presented in the kymograph in (B). Bar, 10 µm. (B) Kymograph of microtubule dynamics at a trailing end focal adhesion. Top panel shows microtubule (MT) only. Bottom panel shows life history plot of MT (green line shows movement of MT end) in relation to focal adhesions (red). Arrows show catastrophes at the distal end of adhesion, arrowheads show capture at the proximal end of adhesion.Detailed analysis of microtubule catastrophe localization shows that they occur at the areas of FAs where paxillin is enriched and highly phosphorylated.^4,13 Paxillin was shown to interact with microtubules through its Lim2/Lim3 domain.¹⁴ Purified GST-Lim2/Lim3 fragment injected into the cell localizes to FAs, displacing endogenous paxillin.¹³ This leads to a 40% decrease in the number of microtubule catastrophe events at adhesion sites,¹³ indicating that paxillin is needed for catastrophe initiation.In summary, we conclude that microtubule catastrophes at focal adhesions are specific events that are triggered by a biochemical mechanism. This process involves the focal adhesion protein paxillin, which may serve as a docking site for microtubules and/or microtubule catastrophe factors. The nature of catastrophe factors remains to be clarified. Possible mechanisms include molecules which induce microtubule catastrophe directly, such as stathmin,¹⁵ or molecules which regulate catastrophe-inducing factors activity. Alternatively, catastrophe factors at adhesion sites could act by removing stabilizing factors from microtubule tips. Thus, allowing already active catastrophe-inducing molecules such as kinesin-13 family member MCAK^16,17 to complete their function. Furthermore, microtubule catastrophe at paxillin-enriched areas, followed by release of microtubule-associated factors, may be involved in paxillin phosphorylation. This local regulation of adhesion disassembly would close the feed-back loop to microtubule regulation of FA turnover.In this model, asymmetric distribution of microtubule catastrophes is tightly linked to asymmetric regulation of FA. Since asymmetric FA dynamics in a cell are critical for organization of the actin cytoskeleton, tensile force distribution and directional cell migration, we conclude that microtubule catastrophes serve as important regulatory events for asymmetric signaling and dynamics of the whole cell (Fig. 2).Open in a separate windowFigure 2Model for asymmetric focal adhesion and microtubule dynamics. Focal complexes at the leading edge either disassemble or mature in response to tension. Microtubules undergo catastrophe both at focal complexes and late adhesions. Late adhesions disassemble in response to multiple microtubule targeting. At the cell rear a microtubule is captured at the proximal end of sliding adhesion and undergoes multiple catastrophes at its distal end, supporting disassembly of this region.     

Leaf carbohydrate metabolism during defense: Intracellular sucrose-cleaving enzymes do not compensate repression of cell wall invertase

The significance of cell wall invertase (cwINV) for plant defense was investigated by comparing wild type (wt) tobacco Nicotiana tabacum L. Samsun NN (SNN) with plants with RNA interference-mediated repression of cwINV (SNN::cwINV) during the interaction with the oomycetic phytopathogen Phytophthora nicotianae. We have previously shown that the transgenic plants developed normally under standard growth conditions, but exhibited weaker defense reactions in infected source leaves and were less tolerant to the pathogen. Here, we show that repression of cwINV was not accompanied by any compensatory activities of intracellular sucrose-cleaving enzymes such as vacuolar and alkaline/neutral invertases or sucrose synthase (SUSY), neither in uninfected controls nor during infection. In wt source leaves vacuolar invertase did not respond to infection, and the activity of alkaline/neutral invertases increased only slightly. SUSY however, was distinctly stimulated, in parallel to enhanced cwINV. In SNN::cwINV SUSY-activation was largely repressed upon infection. SUSY may serve to allocate sucrose into callose deposition and other carbohydrate-consuming defense reactions. Its activity, however, seems to be directly affected by cwINV and the related reflux of carbohydrates from the apoplast into the mesophyll cells.Key words: cell wall invertase, apoplastic invertase, alkaline invertase, neutral invertase, sucrose synthase, plant defense, Nicotiana tabacum, Phytophthora nicotianaePlant defense against pathogens is costly in terms of energy and carbohydrates.^1,2 Sucrose (Suc) and its cleavage products glucose and fructose are central molecules for metabolism and sensing in higher plants (reviewed in refs. ³ and ⁴). Rapid mobilization of these carbohydrates seems to be an important factor determining the outcome of plant-pathogen interactions. In particular in source cells reprogramming of the carbon flow from Suc to hexoses may be a crucial process during defense.^1,2There are two alternative routes of sucrolytic carbohydrate mobilization. One route is reversible and involves an uridine 5′-diphosphate (UDP)-dependent cleavage catalyzed by sucrose synthase (SUSY). Its activity is limited by the concentrations of Suc and UDP in the cytosol, as the affinity of the enzyme to its substrate is relatively low (K_m for Suc 40–200 mM). The other route is the irreversible, hydrolytic cleavage by invertases (INVs), which exhibit high affinity to Suc (K_m 7–15 mM).⁵Plants possess three different types of INV isoenzymes, which can be distinguished by their solubility, subcellular localization, pH-optima and isoelectric point. Usually, they are subdivided into cell wall (cwINV), vacuolar (vacINV), and alkaline/neutral (a/nINVs) INVs.cwINV, also referred to as extracellular or apoplastic INV, is characterized by a low pH-optimum (pH 3.5–5.0) and usually ionically bound to the cell wall. It is the key enzyme of the apoplastic phloem unloading pathway and plays a crucial role in the regulation of source/sink relations (reviewed in refs. ^{3, 6–8}). A specific role during plant defense has been suggested, based on observations that cwINV is often induced during various plant-pathogen interactions, and the finding that overexpression of a yeast INV in the apoplast increases plant resistance.^6,8–10 It was shown, that a rapid induction of cwINV is, indeed, one of the early defense-related reactions in resistant tobacco source leaves after infection with Phytophthora nicotianae (P. nicotianae).¹¹ Finally, the whole infection area in wt leaves was covered with hypersensitive lesions, indicating that all cells had undergone hypersensitive cell death (Fig. 1A).^1,11 When the activity of cwINV was repressed by an RNAi construct, defense-related processes were impaired, and the infection site exhibited only small spots of hypersensitive lesions. Finally, the pathogen was able to sporulate, indicating a reduced resistance of these transgenic plants (Fig. 1A).¹Open in a separate windowFigure 1Defense-induced changes in the activity of intracellular sucrose-cleaving enzymes and their contribution to defense. (A) The repression of cwINV in source leaves of tobacco leads to impaired pathogen resistance and can not be compensated by other sucrose-cleaving enzymes. The intensity of defense reactions is amongst others indicated by the extent of hypersensitive lesions. (B and C) Absolute activity of vacuolar (B) and alkaline/neutral (C) INVs at the infection site (white symbols, control; black symbols, infection site). (D) Increase in SUSY activity at the infection site. All data points taken from noninfected control parts of the plants in each individual experiment and each point along the time scale of an experiment are set as 0%. At least three independent infections are averaged and their means are presented as percentage changes ± SE (circles, SNN; triangles, SNN::cwINV). Insets show the means of the absolute amount of activities (white symbols, control; black symbols, infection site). Material and methods according to Essmann, et al.¹vacINV, also labeled as soluble acidic INV, is characterized by a pH optimum between pH 5.0–5.5. Among others it determines the level of Suc stored in the vacuole and generates hexose-based sugar signals (reviewed in refs. ³ and ¹²). Yet, no specific role of vacINV during pathogen response has been reported. Although vacINV and cwINV are glycoproteins with similar enzymatic and biochemical properties and share a high degree of overall sequence homology and two conserved amino acid motifs,⁴ the activity of vacINV in tobacco source leaves was not changed due to the repression of the cwINV (Fig. 1B).¹ After infection with P. nicotianae the activity of vacINV in wt SNN did not respond under conditions where cwINV was stimulated.¹ There was also no significant change in the transgenic SNN::cwINV (Fig. 1B). This suggests that during biotic stress, there is no crosstalk between the regulation of cwINV and vacINV.a/nINVs exhibit activity maxima between pH 6.5 and 8.0, are not glycosylated and thought to be exclusively localized in the cytosol. But recent reports also point to a subcellular location in mitochondria and chloroplasts.^13,14 Only a few a/nINVs have been cloned and characterized, and not much is known about their physiological functions (reviewed in refs. ^{4, 14} and ¹⁵). Among other things they seem to be involved in osmotic or low-temperature stress response.^14,15 During the interaction between tobacco and P. nicotianae the activity of a/nINVs rose on average 17% in the resistant wt SNN between 1 to 9 hours post infection (Fig. 1C). By contrast, in SNN::cwINV the a/nINVs activities remained unchanged in control leaves and even after infection (Fig. 1C). This suggests that the defense related stimulation in a/nINVs activities is rather a secondary phenomenon, possibly in response to the enhanced cwINV activity and the related carbohydrate availability in the cytosol.SUSY can be found as a soluble enzyme in the cytosol, bound to the inner side of the plasma membrane or the outer membrane of mitochondria, depending on the phosphorylation status. It channels hexoses into polysaccharide biosynthesis (i.e., starch, cellulose and callose) and respiration.^12,16 There is also evidence that SUSY improves the metabolic performance at low internal oxygen levels¹⁷ but little is known about its role during plant defense. Callose formation is presumably one of the strongest sink reactions in plant cells.^1,18 Defense-related SUSY activity may serve to allocate Suc into callose deposition and other carbohydrate-consuming defense reactions. In fact, in the resistant wt the activity of SUSY increased upon interaction with P. nicotianae in a biphasic manner (Fig. 1D). The time course is comparable to that of cwINV activity and correlates with callose deposition and enhanced respiration.^1,11 However, repression of cwINV leads in general to a reduction of SUSY activity in source leaves of tobacco.¹ After infection the activation of SUSY was also significantly impaired (Fig. 1D). At the same time, the early defense-related callose deposition in infected mesophyll cells of SNN::cwINV plants is substantially delayed.¹ It is known that expression of SUSY isoforms is differentially controlled by sugars,¹² and there is evidence that hexoses generated by the defense-induced cwINV activity deliver sugar signals to the infected cells.¹ In this sense, the reduction of defense-related, cwINV-generated sugar signals could be responsible for the repression of SUSY activity in SNN::cwINV plants after infection with P. nicotianae.Only limited hexoses or hexose-based sugar signals could be generated by cytoplasmic Suc cleavage.¹² The reduction of soluble carbohydrates for sugar signaling and also as fuel for metabolic pathways that support defense reactions could be responsible for the impaired resistance in SNN::cwINV plants (Fig. 1A).Obviously, neither intracellular INV isoforms, nor SUSY can compensate for the reduced carbohydrate availability due to cwINV repression during plant defense. The data also suggest that the activity of SUSY is affected by cwINV and related reflux of carbohydrates. It is known that SUSY activity can be controlled, e.g., by sugar-mediated phosphorylation¹² and one may speculate that posttranslational modulation of the protein is affected by the defense-related carbohydrate status of the cell.     

Conformational stability and aggregation of therapeutic monoclonal antibodies studied with ANS and thioflavin T binding

Characterization of aggregation profiles of monoclonal antibodies (mAb) is gaining importance because an increasing number of mAb-based therapeutics are entering clinical studies and gaining marketing approval. To develop a successful formulation, it is imperative to identify the critical biochemical properties of each potential mAb drug candidate. We investigated the conformational change and aggregation of a human IgG1 using external dye-binding experiments with fluorescence spectroscopy and compared the aggregation profiles obtained to the results of size-exclusion chromatography. We show that using an appropriate dye at selected mAb concentration, unfolding or aggregation can be studied. In addition, dye-binding experiments may be used as conventional assays to study therapeutic mAb stability.Key words: therapeutic monoclonal antibody, protein aggregation, conformational change, stability and shelf-life prediction, accelerated studiesMonoclonal antibodies (mAbs) have emerged as a novel class of protein drugs and are utilized for a variety of mostly incurable and debilitating diseases such as cancer and rheumatoid arthritis.^1–4 For treatment of chronic diseases, it is desirable for these drugs to be administered subcutaneously, in which case high protein concentrations (>100 mg/mL) are generally needed.^5,6 Protein-based drugs containing mAbs must contain minimum amounts of aggregation and fragmentation and conserve their structural integrity during storage because degraded or aggregated protein may induce immunogenicity or reduce efficacy. Currently, size-exclusion chromatography-high performance liquid chromatography (SEC-HPLC) is the most commonly used method to characterize mAb aggregation profiles;⁷ however it is time consuming, expensive and requires expertise. SEC-HPLC cannot be used to obtain accurate biophysical profiles of mAbs at high concentrations because dilution during the experiment might lead to reversible aggregation. Furthermore, the potential interaction of aggregates with surfaces, e.g., needle, tubing, column, will lead to the loss of sample and thus an inaccurate analysis.^8,9 Additional drawbacks of the technique are that different conformations such as partially unfolded monomers also cannot be distinguished by SEC-HPLC and large aggregates may be totally excluded during the injection into the column.External dye binding assays have been used to characterize protein stability and aggregation,^10–12 and studies involving biopharmaceuticals have been reported recently, e.g., for thermostability screening¹⁰ and detection of aggregation.^11–14 These methods are not limited by protein quantity and are more sensitive because they are fluorescence-based. We studied the accelerated unfolding of an IgG1 mAb with the hydrophobic dye 1-anilino-8-naphthale-nesulfonate (ANS), and its accelerated aggregation with aggregate specific Thioflavin T (ThT). We have also conducted accelerated aggregation studies with SEC-HPLC7 and compared the findings to the ThT binding results. We hypothesize that key structures formed during mAb aggregation can be probed selectively by the appropriate dyes (Fig. 1) with specific mAb concentrations.Open in a separate windowFigure 1Key structures of the mAb probed by fluorescent dyes. N and U are native and unfolded monomers, respectively. “n” reactive monomers form aggregates.     

ARF-GTPase as a Molecular Switch for Polar Auxin Transport Mediated by Vesicle Trafficking in Root Development

Polar auxin transport (PAT), which is controlled precisely by both auxin efflux and influx facilitators and mediated by the cell trafficking system, modulates organogenesis, development and root gravitropism. ADP-ribosylation factor (ARF)-GTPase protein is catalyzed to switch to the GTP-bound type by a guanine nucleotide exchange factor (GEF) and promoted for hybridization to the GDP-bound type by a GTPase-activating protein (GAP). Previous studies showed that auxin efflux facilitators such as PIN1 are regulated by GNOM, an ARF-GEF, in Arabidopsis. In the November issue of The Plant Journal, we reported that the auxin influx facilitator AUX1 was regulated by ARF-GAP via the vesicle trafficking system.¹ In this addendum, we report that overexpression of OsAGAP leads to enhanced root gravitropism and propose a new model of PAT regulation: a loop mechanism between ARF-GAP and GEF mediated by vesicle trafficking to regulate PAT at influx and efflux facilitators, thus controlling root development in plants.Key Words: ADP-ribosylation factor (ARF), ARF-GAP, ARF-GEF, auxin, GNOM, polar transport of auxinPolar auxin transport (PAT) is a unique process in plants. It results in alteration of auxin level, which controls organogenesis and development and a series of physiological processes, such as vascular differentiation, apical dominance, and tropic growth.² Genetic and physiological studies identified that PAT depends on efflux facilitators such as PIN family proteins and influx facilitators such as AUX1 in Arabidopsis.Eight PIN family proteins, AtPIN1 to AtPIN8, exist in Arabidopsis. AtPIN1 is located at the basal side of the plasma membrane in vascular tissues but is weak in cortical tissues, which supports the hypothesis of chemical pervasion.³ AtPIN2 is localized at the apical side of epidermal cells and basally in cortical cells.^1,4 GNOM, an ARF GEF, modulates the localization of PIN1 and vesicle trafficking and affects root development.^5,6 The PIN auxin-efflux facilitator network controls root growth and patterning in Arabidopsis.⁴ As well, asymmetric localization of AUX1 occurs in the root cells of Arabidopsis plants,⁷ and overexpression of OsAGAP interferes with localization of AUX1.¹ Our data support that ARF-GAP mediates auxin influx and auxin-dependent root growth and patterning, which involves vesicle trafficking.¹ Here we show that OsAGAP overexpression leads to enhanced gravitropic response in transgenic rice plants. We propose a model whereby ARF GTPase is a molecular switch to control PAT and root growth and development.Overexpression of OsAGAP led to reduced growth in primary or adventitious roots of rice as compared with wild-type rice.¹ Gravitropism assay revealed transgenic rice overxpressing OsAGAP with a faster response to gravity than the wild type during 24-h treatment. However, 1-naphthyl acetic acid (NAA) treatment promoted the gravitropic response of the wild type, with no difference in response between the OsAGAP transgenic plants and the wild type plants (Fig. 1). The phenotype of enhanced gravitropic response in the transgenic plants was similar to that in the mutants atmdr1-100 and atmdr1-100/atpgp1-100 related to Arabidopsis ABC (ATP-binding cassette) transporter and defective in PAT.⁸ The physiological data, as well as data on localization of auxin transport facilitators, support ARF-GAP modulating PAT via regulating the location of the auxin influx facilitator AUX1.¹ So the alteration in gravitropic response in the OsAGAP transgenic plants was explained by a defect in PAT.Open in a separate windowFigure 1Gravitropism of OsAGAP overexpressing transgenic rice roots and response to 1-naphthyl acetic acid (NAA). (A) Gravitropism phenotype of wild type (WT) and OsAGAP overexpressing roots at 6 hr gravi-stimulation (top panel) and 0 hr as a treatment control (bottom panel). (B) Time course of gravitropic response in transgenic roots. (C and D) results correspond to those in (A and B), except for treatment with NAA (5 × 10⁻⁷ M).The polarity of auxin transport is controlled by the asymmetric distribution of auxin transport proteins, efflux facilitators and influx carriers. ARF GTPase is a key member in vesicle trafficking system and modulates cell polarity and PAT in plants. Thus, ARF-GDP or GTP bound with GEF or GAP determines the ARF function on auxin efflux facilitators (such as PIN1) or influx ones (such as AUX1).ARF1, targeting ROP2 and PIN2, affects epidermal cell polarity.⁹ GNOM is involved in the regulation of PIN1 asymmetric localization in cells and its related function in organogenesis and development.⁶ Although VAN3, an ARF-GAP in Arabidopsis, is located in a subpopulation of the trans-Golgi transport network (TGN), which is involved in leaf vascular network formation, it does not affect PAT.¹⁰ OsAGAP possesses an ARF GTPase-activating function in rice.¹¹ Specifically, our evidence supports that ARF-GAP bound with ARF-GTP modulates PAT and gravitropism via AUX1, mediated by vesicle trafficking, including the Golgi stack.¹Therefore, we propose a loop mechanism between ARF-GAP and GEF mediated by the vascular trafficking system in regulating PAT at influx and efflux facilitators, which controls root development and gravitropism in plants (Fig. 2). Here we emphasize that ARF-GEF catalyzes a conversion of ARF-bound GDP to GTP, which is necessary for the efficient delivery of the vesicle to the target membrane.¹² An opposite process of ARF-bound GDP to GTP is promoted by ARF-GTPase-activating protein via binding. A loop status of ARF-GTP and ARF-GDP bound with their appurtenances controls different auxin facilitators and regulates root development and gravitropism.Open in a separate windowFigure 2Model for ARF GTPase as a molecular switch for the polar auxin transport mediated by the vesicle traffic system.