CLE genes may act in a variety of tissues/cells and involve other signaling cascades in addition to CLV3-WUS-like pathways

CLE, which is the term for the CLV3/ESR-related gene family, is thought to participate in CLAVATA3-WUSCHEL (CLV3-WUS) and CLV3-WUS-like signaling pathways to regulate meristem activity in plant. Although some CLE genes are expressed in meristems, many CLE genes appear to express in a variety of tissues/cells. Here we report that CLE14 and CLE20 express in various specific tissues/cells outside the shoot/root apical meristem (SAM/RAM), including in highly differentiated cells, and at different developmental stages. Overexpressing CLE14 or CLE20 also causes multiple phenotypes, which is consistent with its expression pattern in Arabidopsis. These results suggest that CLE genes may play multiple roles and involve other signaling cascades in addition to the CLV3-WUS and CLV3-WUS-like pathways.Key words: CLE, CLAVATA3-WUSCHEL, cell signaling and development, root apical meristem, arabidopsisIntercellular communication and coordination between adjacent cell populations are critical for cell-fate specification, as well as for meristem organization and maintenance. In the shoot apical meristem (SAM), local signaling, which involves the CLAVATA3-WUSCHEL (CLV3-WUS) negative feedback loop, controls stem cell homeostasis and SAM activity.¹ As well, it has been suggested that a CLV3-WUS-like negative feedback pathway operates to control root apical meristem (RAM) activity. This view is supported by the facts that a WUS-related homeobox gene, WOX5, is expressed in cells of the quiescent center (QC) in the RAM, and that loss-of-function of WOX5 in the QC leads to the differentiation of the adjacent root cap initials (RCI), whereas gain-of-function blocks the differentiation of derivatives of the RCI in the root.² Additional support for the function in the RAM of a CLV3-WUS-like pathway, comes from observations that CLE genes (collectively referred to as the CLV3/ESR-relate gene family) are not only expressed in the RAM,^3,4 but also, that overexpression of some CLE genes triggers premature termination of the RAM.⁵ In this regard it has been recently reported that CLE40, which expresses in the differentiating daughter cells of the distal root stem cells, restricts WOX5 expression and promotes differentiation of stem cells in the RAM.⁶ Taken together these data suggest a CLV3-WUS-like feedback loop acts to negatively regulate RAM activity in plants.Our previous results have shown that CLE14 and CLE20 express in specific cells of roots, and that overexpression of CLE14 or CLE20 in Arabidopsis triggers early termination of the RAM in a CLAVATA1 (CLV1)-independent, but CLAVATA2 (CLV2)-dependent manner.^7,8 We also showed that both CLE14 and CLE20 peptides inhibit, irreversibly, root growth by reducing cell division rates in the RAM.⁷ CLV2 and CRN (a receptor-like protein kinase, also known as SOL2, isolated as a suppressor of root-specific overexpression of CLE19) are required for CLE14 and CLE20 peptide functions in vitro.^9,10 Using computational modeling approaches we further demonstrated that 12-amino-acid CLE14 and CLE20 peptides may function through a potential heterodimer/heterotetramer CLV2-CRN complex.⁷CLV3 expresses exclusively in the stem cells of the SAM, and it has been consistently shown that the CLV3 peptide is required for homeostasis of the stem cells and for the maintenance of the SAM.¹ Although some CLE genes are found to express in meristems, many CLE genes appear to express in an array of tissues and cells, including highly differentiated tissues/cells.^3,4 In this report we show that CLE14 and CLE20 express in specific tissues outside the RAM and SAM of Arabidopsis, including highly differentiated cells, and at different developmental stages. Overexpressing CLE14 or CLE20 also causes multiple phenotypes, which is consistent with its expression pattern in Arabidopsis. These results suggest that CLE genes may play multiple roles in regulating the developmental fate of cells, which includes, but is not limited to, stem cells, and also may be involved in other signaling cascades in addition to the CLV3-WUS pathway.     

Environmental stimuli and intestinal stem cell behavior

Comment on: Wong VWY, et al. Nat Cell Biol 2012; 14:401-8.The intestine carries out important functions related to digestion and absorption. It is composed of three distinct layers, an outer muscle layer, a mesenchymal layer and the epithelial layer. The epithelial layer forms the protective barrier that faces the luminal content of the intestine. In order to maintain barrier function the epithelial layer needs constant replenishment. This is ensured by continuous cellular replication in proliferative crypt compartments. Following exit from the crypt, cells adopt fates along either secretory or absorptive lineage and will, after three to four days, be exfoliated into the lumen of the intestine from the tips of the villi. Intestinal stem cells located at the bottom of the proliferative crypt compartment ensure lifelong maintenance of the organ (Fig. 1A).Open in a separate windowFigure 1. Diagram of the intestinal stem cell niche. (A) Lgr5-expressing columnar-based crypt cells (CBCs) intercalated between Paneth cells are indicated in green. Stem cells located in position +4 are yellow. Lrig1 is expressed in a gradient along the niche axis with highest expression in the CBCs indicated with the thickness of the red line. Proliferation in the stem cell niche ensures continuous replenishment of the transit-amplifying (TA) compartment. (B) Within the stem cell niche, Lgr5-expressing CBCs are actively dividing and will give rise to both HopX-expressing +4 cells and TA cells. HopX-expressing cells, which are less mitotically active, will give rise to fewer TA cells and occasionally an Lgr5-expressing stem cell. Lrig1 expression in the stem cell niche reduces the amplitude of ErbB activation and is essential for controlling stem cell proliferation.Adult stem cell niches are far more heterogeneous than previously anticipated.¹ The intestinal stem cell niche can be subdivided by the relative position within the crypt. Stem cells located in position +4, just above secretory Paneth cells, express HopX, Bmi1 and Tert. These cells are generally less mitotically active than Lgr5-expressing stem cells located at the bottom of the proliferative crypts intercalated between Paneth cells (Fig. 1A).^2,3 It has been argued that both populations represent the most primitive stem cell; however, recent studies suggest that stem cells can interconvert between the two states (Fig. 1B).³ Fate mapping from cells in position 4 and at the bottom of the crypt supports this.^2,4 The positional cues responsible for cellular sorting into different functional stem cell compartments are poorly characterized. The only known effector of cellular positioning is Wnt (wingless-related MMTV integration site) signaling.⁵ Wnt is highly expressed by Paneth cells along with other mitotic factors, such as ErbB and Notch ligands.⁶ This could functionally account for the differences observed in proliferative potential along the stem cell axis. The discrete expression patterns of Lgr5 and HopX also support the existence of distinct microenvironments that supports cellular identities. A thorough characterization of the factors responsible for stem cell identity will help delineate and define the functional relationship between the distinct stem cell populations.Tissue homeostasis is governed by balanced loss and gain of cells. The stem cell niche supports constant proliferation via pro-mitotic stimuli. In order to control the amplitude of signaling strength, many pathways have developed negative feedback loops. Lrig1 (Leucine-rich repeats and immunoglobulin-like domains 1) is a negative feedback regulator of ErbB-mediated growth factor signaling.⁷ Lrig1 marks stem cells in various epithelial tissues including the intestinal epithelium, where it is expressed within the entire stem cell niche including the +4 and Lgr5-expressing cells (Fig. 1).^8,9 The functional relevance of Lrig1 and negative feedback regulation is clear from the pronounced expansion of the intestinal stem cell compartment observed in the Lrig1-KO mouse model.⁹ This is mediated via increased ErbB signaling and demonstrates the importance of balanced signaling strength within the stem cell niche.⁹ Moreover, an independent study reveals that Lrig1-KO animals have a higher incidence of colorectal cancer, suggesting that unbalanced stem cell proliferation increases tumor susceptibility.¹⁰ Future studies will address whether additional feedback regulators control signaling strength within the intestinal stem cell niche and how homeostasis within the stem cell compartment affects tumor susceptibility.     

Orientation of vascular cell divisions in Arabidopsis

Orientation of cell division is essential for plant development as the direction of growth is determined by the direction of cell expansion and orientation of cell division. We have demonstrated that cell division orientation in vascular tissue is regulated by the interactions between a receptor kinase (PXY) expressed in dividing cells and its peptide ligand (CLE41) that is localized to adjacent phloem cells. Given that other receptor kinases have been identified as orienting the cell division plane in several developmental processes, we suggest that localized signaling from adjacent cells may be a general mechanism for defining the plane of cell division.Key words: xylem, phloem, cell division orientation, procambium, cambiumThroughout the life of plants, new organs are generated from meristems which contain stem cells at their center. Meristematic cells divide in regulated processes resulting in displacement of daughter cells to the periphery of the meristem where they differentiate, taking on new cell identities.¹ Vascular meristems (cambium and procambium) are responsible for radial growth and are the main source of plant biomass.² Their regulation has a come under increasing scrutiny as biomass is likely to play an increasing role in generation of renewable energy.³Arabidopsis vascular tissue is organized into discrete collateral bundles in stems,⁴ whereas in hypocotyls, vasculature forms in a continuous ring, much like that of trees.⁵ In both cases spatially separated xylem and phloem are formed along the stem mediolateral axis and are populated with cells derived from the procambium or cambium (Fig. 1). Vascular initials displaced from the meristematic zone towards the center of the stem differentiate into xylem whereas those displaced towards the outside of the stem differentiate into phloem. This organization occurs because vascular meristematic cells are long and thin and divide periclinally down their long axis, perpendicular to the mediolateral axis. Because these are highly ordered divisions, vascular tissue is characterized by long files of cells. Until recently regulatory factors which influence the highly ordered nature of these divisions—and therefore plant vascular tissue organization were entirely unknown.Open in a separate windowFigure 1Arabidopsis vascular tissue at the base of inflorescence stems (A) and hypocotyls (B). The mediolateral axes are marked with arrows, x is xylem, ph is phloem, pc is procambium, c is cambium. Scale bars are 50 µm.     

“Double hit” makes the difference

Comment on: Menendez JA, et al. Cell Cycle 2012; 11:2782–92.Metformin (N’, N’-dimethylbiguanide) is an anti-diabetic drug prescribed to more than 100 million patients in the world. In addition to its efficacy for the treatment of diabetes, several recent studies have shown that it has anti-tumoral properties.¹ We and others have shown that metformin targets cancer cell metabolism by inhibiting mitochondrial complex 1 activity.^2,3 This energetic stress leads to a decrease of intracellular ATP concentration, and cancer cells will increase their rate of glycolysis.² This compensatory response is not sufficient to restore ATP levels, but is adequate to maintain viable cells in most of the cancer cells. Indeed, metformin blocks cell growth but can also induce apoptosis in some cancer cell models.⁴ The increase of glycolysis induced by metformin is somehow inconsistent with the observed inhibition of proliferation, since cancer cells use preferentially glycolysis to grow faster. This switch to glycolysis, also known as the “Warburg effect,” is linked to oncogenic transformation⁵ and is accompanied by the hyperactivation of the mTOR pathway. In cancer cells, the increase of glycolysis induced by metformin is associated with a strong inhibition of the mTOR pathway via the AMPK. This new metabolic order established by metformin may explain the paradoxical effect of metformin. In view of the above scenario, Menendez et al. decided to test the synthetic lethality of metformin and combined metformin treatment with glucose starvation. They showed that the treatment of breast cancer cells with metformin alone does not induce apoptosis but arrests cells in G₀/G₁. Glucose starvation by itself induces few apoptosis, but the combination of metformin with the absence of glucose induces massive apoptosis. This is not altogether surprising, since the dual action of metformin and glucose starvation block the two main ways of production of ATP (i.e., mitochondrial respiration and glycolysis) (Fig. 1). This is an interesting observation, which could be valuable for future anticancer therapy; however, glucose starvation is not therapeutically feasible. Thus, the use 2-deoxyglucose (2-DG), an inhibitor of glycolysis, could be useful. We and others found that the combination of 2-DG and metformin inhibits prostate cancer cell proliferation and breast tumor growth in xenograft models.^2,6 Although it induces a slight apoptotic response in vitro, 2-DG alone is not efficient in vivo to alter tumor growth⁶ but improves the curative action of radiotherapy;⁷ similarly, it reinforces metformin action. Another interesting issue raised by Menendez et al. is the use of such dual therapy to target cancer stem cells. Metformin has been shown to selectively kill cancer stem cells and the chemotherapy-resistant subpopulation of cancer stem cells.^8,9 Cancer stem cells greatly depend on aerobic glycolysis to sustain their stemness and immortality. The synthetic lethality induced by metformin and glucose starvation may help to improve chemotherapy action and avoid cancer relapse. In conclusion, targeting cancer cell metabolism with a “dual hit therapy” opens new avenues for the future treatment of cancer.Open in a separate windowFigure 1. The combination of metformin and glucose starvation induces a strong energetic stress. Metformin inhibits the mitochondrial complex 1 and glucose starvation, or 2-DG inhibits ATP production from glycolysis. The combination of the two energetic stresses induces a massive energetic stress and leads to a strong apoptotic response.     

A systems biology approach to dissect the contribution of brassinosteroid and auxin hormones to vascular patterning in the shoot of Arabidopsis thaliana

Systems biology can foster our understanding of hormonal regulation of plant vasculature. One such example is our recent study on the role of plant hormones brassinosteroids (BRs) and auxin in vascular patterning of Arabidopsis thaliana (Arabidopsis) shoots. By using a combined approach of mathematical modelling and molecular genetics, we have reported that auxin and BRs have complementary effects in the formation of the shoot vascular pattern. We proposed that auxin maxima, driven by auxin polar transport, position vascular bundles in the stem. BRs in turn modulate the number of vascular bundles, potentially by controlling cell division dynamics that enhance the number of provascular cells. Future interdisciplinary studies connecting vascular initiation at the shoot apex with the established vascular pattern in the basal part of the plant stem are now required to understand how and when the shoot vascular pattern emerges in the plant.Key words: Arabidopsis, vascular, auxin, brassinosteroids, mathematical model, computer simulationsThe plant vascular system is responsible for the long-distance transport of water, solutes and molecules throughout the plant, being essential for plant growth and development. It is formed by two different functional tissues: the xylem, which transports water from roots to aerial organs, and the phloem, through which nutrients and photosynthetic products and signaling molecules are transported.During embryogenesis, the vasculature is characterized as an undifferentiated procambial tissue in the innermost part of the plant embryo.¹ Later in development, the procambium (i.e., a group of pluripotent stem cells²) begins to divide and differentiate into xylem and phloem tissues through oriented cell divisions. In the shoot, procambium generates xylem tissue centripetally and phloem tissue centrifugally, driving the formation of collateral vascular bundles around it.^3,4 In the inflorescence stem of the model plant Arabidopsis, the radial pattern of the vasculature exhibits a periodic organization made by the alternation of vascular bundles and interfascicular fibers, which altogether form the vascular ring (Fig. 1A).Open in a separate windowFigure 1Vascular patterning in Arabidopsis shoot inflorescence stem. (A) Radial section of DR5::GUS expression at the base of the inflorescence stem in Arabidopsis Col-0 plants. (B) Computer simulation result for auxin concentration ([Auxin]) in arbitrary units (a.u.) along a ring of cells; x and y stand for spatial coordinates. Auxin is distributed in maxima which, according to the model hypothesis, position vascular bundles. (C) Longitudinal section of Arabidopsis Col-0 wild-type plant at the most apical zone, immediately below the shoot apical meristem. Arrows point to xylem strains coming from the lateral organs.Previous studies have documented the importance of plant hormones such as auxin and BRs in vascular cell differentiation and patterning.⁵ Defective polar auxin transport distorts shoot vascular patterning^6,7 and BR loss-of-function mutants exhibit few vascular bundles.^8,9 But how do these hormones control shoot vascular patterning? In order to answer this question, we used both quantitative measurements of vascular phenotypes and computational modeling.¹⁰     

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.     

Do Plant Cells Secrete Exosomes Derived from Multivesicular Bodies?

Multivesicular bodies (MVBs) are spherical endosomal organelles containing small vesicles formed by inward budding of the limiting membrane into the endosomal lumen. In mammalian red cells and cells of immune system, MVBs fuse with the plasma membrane in an exocytic manner, leading to release their contents including internal vesicles into the extracellular space. These released vesicles are termed exosomes. Transmission electron microscopy studies have shown that paramural vesicles situated between the plasma membrane and the cell wall occur in various cell wall-associated processes and are similar to exosomes both in location and in morphology. Our recent studies have revealed that MVBs and paramural vesicles proliferate when cell wall appositions are rapidly deposited beneath fungal penetration attempts or during plugging of plasmodesmata between hypersensitive cells and their intact neighboring cells. This indicates a potential secretion of exosome-like vesicles into the extracellular space by fusion of MVBs with the plasma membrane. This MVB-mediated secretion pathway was proposed on the basis of pioneer studies of MVBs and paramural vesicles in plants some forty years ago. Here, we recall the attention to the occurrence of MVB-mediated secretion of exosomes in plants.Key Words: cell wall, endocytosis, endosome, exocytosis, exosome, multivesicular body, paramural bodyMultivesicular bodies (MVBs) are spherical endosomal organelles containing a number of small vesicles formed by inward budding of the limiting membrane into the endosomal lumen.¹ MVBs contain endocytosed cargoes and deliver them into lysosomal/vacuolar compartments for degradation. They also incorporate newly synthesized proteins destined for lysosomal/vacuolar compartments.² In mammalian cells of hematopoietic origin, endosomal MVBs function in removal of endocytosed surface proteins in an exocytic manner. They are redirected to the plasma membrane, where they release their contents including internal vesicles into the extracellular space by membrane fusion. The released vesicles are termed exosomes.³ During reticulocyte maturation to erythrocyte, a group of surface proteins, such as the transferrin receptor, become obsolete and are discarded via MVB-mediated secretion.³ Time-course transmission electron microscopy (TEM) first revealed that colloidal gold-transferrin was internalized into MVBs via receptor-mediated endocytosis and then transferrin together with its receptor were delivered into the extracellular space via the fusion of MVBs with the plasma membrane of reticulocytes.⁴ Some other cell types of hematopoietic origin, such as activated platelets, cytotoxic T cells and antigen-presenting cells, also secrete exosomes. Exosomes thus may play a role in various physiological processes other than discarding obsolete proteins.³Our recent TEM studies provided ultrastructural evidence on the enhanced vesicle trafficking in barley leaf cells attacked by the biotrophic powdery mildew fungus. Multivesicular compartments including MVBs, intravacuolar MVBs, and paramural bodies turned out to proliferate in intact host cells during formation of cell wall appositions (papilla response), in the hypersensitive response, and during accommodation of haustoria.^5,6 MVBs proliferated in the cytoplasm of haustorium-containing epidermal cells during compatible interactions and near sites of cell wall-associated oxidative microburst either during the papilla response or during the hypersensitive response. Because MVBs in plant cells have been demonstrated to be endosomal compartments,^7–9 they may participate in internalization of nutrients from the apoplast of intact haustorium-containing epidermal cells and sequestration of damaged membranes and deleterious materials originating from the oxidative microburst.^5,6 The presence of intravacuolar MVBs with double limiting membranes (Fig. 1A) indicates an engulfment of MVBs by the tonoplast and a vacuole-mediated autophagy of MVBs.^5,6 MVBs, as prevacuolar compartments in plant cells,⁹ thus probably deliver their contents into the central vacuole via both the fusion with the tonoplast and the engulfment by the tonoplast (Fig. 2A and B). On the other hand, paramural bodies, in which small vesicles are situated between the cell wall and the plasma membrane, were associated with cell wall appositions deposited beneath fungal penetration attempts (Fig. 1B) or around hypersensitive cells including sites of plugged plasmodesmata (Fig. 1C and D).^5,6 Because paramural vesicles are similar to exosomes both in location and in morphology, we speculated that MVBs fuse with the plasma membrane in an exocytic manner to form paramural bodies.^5,6 Endocytosed cell surface materials in endosomal MVBs may be reused and delivered together with newly synthesized materials in Golgi apparatus-derived vesicles to cell wall appositions, which are deposited rapidly to prevent fungal penetration (Fig. 2A) or to contain hypersensitive cell death (Fig. 2B). MVBs thus may be driven along two distinct pathways to deliver their contents into either central vacuole or extracellular space.Open in a separate windowFigure 1Multivesicular compartments in intact cells in barley leaves attacked by the barley powdery mildew fungus. (A) An intravacuolar multivesicular body (MVB) with double limiting membranes in an intact epidermal cell (EC) adjacent to a hypersensitive epidermal cell (EC*). The arrows point to the outer limiting membrane, which is seemingly derived from the tonoplast. Note that neighboring intravacuolar vesicles (in between two arrowheads) may result from degradation of double limiting membranes of intravacuolar MVBs or may be delivered into the vacuole by MVB-fusion with the tonoplast. (B) Paramural vesicles (arrowheads) in a paramural body associated with cell wall appositions (asterisk) deposited by an intact epidermal cell. (C) A multivesicular body (MVB) in contact with a paramural body (PMB) (a nonmedian section) associated with cell wall appositions (asterisk) deposited by an intact mesophyll cell adjacent to a hypersensitive mesophyll cell. Note that cell wall appositions deposit beside an intercellular space (IS). The arrows point to the tonoplast. (D) A paramural body (PMB) associated with cell wall appositions (asterisks) blocking plasmodesmata (in between two arrowheads) at the side of an intact mesophyll cell (MC) underlying a hypersensitive epidermal cell (EC*). The arrows point to the tonoplast. CV, central vacuole; CW, cell wall; MB, microbody. Bars, 1µm.Open in a separate windowFigure 2Hypothetical diagram of delivery of endocytosed cell surface materials via MVBs into the central vacuole or the extracellular space where intact barley cells deposit cell wall appositions. (A) Deposition of cell wall appositions (asterisk) beneath powdery mildew penetration attempts. AGT, appressorial germ tube; PP, penetration peg. (B) Deposition of cell wall appositions (asterisks) against constricted plasmodesmata (PD) between a hypersensitive epidermal cell (EC) penetrated by the powdery mildew fungus and an underlying mesophyll cell (MC). H, haustorium. Arrows and numbers show pathways of vesicle trafficking. 1, Secretion of Golgi-derived vesicles containing newly synthesized materials; G, Golgi body; TGN, trans-Golgi network; 2, Endocytosis of cell surface materials from coated pits (coated open circles) via coated vesicles (coated circles) to multivesicular bodies (MVB); 3, Delivery of endocytosed materials for degradation inside the central vacuole (CV) via membrane fusion between MVBs and the tonoplast (T); small broken circles, vesicles in degradation; 4, Delivery of endocytosed materials for degradation inside the central vacuole via engulfment of MVBs by the tonoplast; large broken circles; MVB limiting membranes in degradation; 5, delivery of endocytosed materials into the extracellular space for deposition of cell wall appositions (asterisks) via membrane fusion between MVBs and the plasma membrane (PM). CW, cell wall; PMB, paramural body. PD0, 1, 2, 3 and 4 represent stages of plugging plasmodesmata. PD0, open plasmodesmata between two intact mesophyll cells (MC) subjacent to the hypersensitive epidermal cell (EC); PD1, constriction of plasmodesmata by callose (grey dots) deposition at plasmodesmal neck region; PD2, constricted plasmodesmata associated with plasmodesma-targeted secretion; PD3, further blocking of plasmodesmata by deposition of cell wall appositions; PD4, completely blocked plasmodesmata.Earlier than the discovery in animal cell systems,⁴ it was proposed in two independent papers in 1967 that the fusion of MVBs with the plasma membrane might result in the release of small vesicles into the extracellular space in fungi and in higher plants.^10,11 Several lines of evidence support the occurrence of MVB-mediated secretion of exosome-like vesicles in plants. First, vesicles of the same morphology as MVB internal vesicles have been observed in extracellular spaces or paramural spaces in various types of plant cells in various plant species by TEM.¹² An early study on endocytosis by soybean protoplasts also showed small extracellular vesicles attaching on the plasma membrane.⁸ Second, cooccurrence of MVBs and paramural vesicles has been observed in processes of cell proliferation, cell differentiation, and cell response to abiotic and biotic stress. Examples are cell plate formation,^13,14 secondary wall thickening,^15,16 cold hardness,^17,18 and deposition of cell wall appositions upon pathogen attack.^5,6,19–21 Third, identical molecular components, such as arabinogalactan proteins^22,23 and peroxidases,⁶ have been immunolocalized in both MVBs and paramural bodies. Despite these pieces of evidence, a conclusive demonstration of MVB-mediated secretion of exosomes in plants requires further exploration.The presently available experimental systems, approaches, and membrane markers may allow future demonstration of MVB-mediated secretion of exosomes in plants. Recent in vivo real-time observation and colocalization of cell surface and endosomal markers have already revealed that endosomes filled with endocytosed preexisting cell wall and plasma membrane materials are rapidly delivered to cytokinetic spaces to form cell plates in dividing tobacco, Arabidopsis, and maize cells.²⁴ Because TEM observed paramural bodies attaching to cell plates¹³ and MVBs in the vicinity of cell plates during all stages of cell plate formation,^14,25,26 MVBs and paramural bodies may participate in delivery of endocytosed building blocks to cell plates. Jiang''s and Robinson''s labs together developed a transgenic tobacco BY-2 cell line stably expressing a YFP-labeled vacuolar sorting receptor protein and antibodies against the vacuolar sorting receptor protein localized to the limiting membrane of MVBs.⁹ These tools together with live cell imaging and immunoelectron microscopy may allow visualization of MVB-fusion to the new plasma membrane, of vacuolar sorting receptors in both the limiting membrane of MVBs and the new plasma membrane, and of identical cell plate components in both internal vesicles of MVBs and paramural vesicles.In spite of obvious differences in plant and animal cytokinesis, the generation of cell plates by cell-plate-directed fusion of endosomes resembles the plugging of midbody canals by midbody-directed endosomes to separate daughter cells at the terminal phase of animal cytokinesis.²⁷ Likely, functional similarities of the fusion between endosomal MVBs and the plasma membrane to eliminate unwanted cell contents may also exist in maturation of mammalian red blood cells and plant sieve elements in the sense that the fusion of MVBs with the plasma membrane may occur during maturation of the latter.²⁸ On the other hand, although plant cells may secrete MVB-derived exosomes in defense response upon pathogen attack,^5,6 plant cell walls rule out the direct intercellular communication during the immune response mediated by exosomes in the circulation of mammals.³ In contrast, plasmodesma-directed secretion of exosomes would block the cell-to-cell communication between hypersensitive cells and their neighboring cells during hypersensitive response.⁵ Further exploration will lead us to a better understanding of similarities and differences of exosome secretion between plants and animals.     

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.     

Tubular actin filaments in tobacco guard cells

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