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1 Department of Biological Science and Technology, Faculty of Industrial Science and Technology, and 2 Tissue Engineering Research Centre, Research Institute of Biological Science, Tokyo University of Science, Yamazaki 2641, Noda, Chiba, 278-8510, Japan
3 Department of Biomolecular Sciences, Graduate School of Life Sciences, Tohoku University, Sendai, Miyagi 980-8578, Japan
4 Department of Clinical Molecular Biology, Faculty of Medicine, Kyoto University, Kyoto 606-8507, Japan
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Introduction |
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4 or ß1 chains causes serious deficiencies in both haematopoiesis and embryogenesis (Fassler et al. 1995; Hirsch et al. 1996; Arroyo et al. 1996). Neutralizing antibodies against these molecules blocked cell adhesion of HS/PCs to stromal cells in vitro and the homing of HS/PCs into haematopoietic organs, such as bone marrow and spleen in vivo (Craddock et al. 1997). However, molecular mechanisms underlying the movement of HS/PCs into haematopoietic microenvironments following cell adhesion have not been fully elucidated. Reorganization of the actin cytoskeleton is essential for cell adhesion, movement and migration (Pantaloni et al. 2001) and several types of actin-binding proteins were identified that regulate actin polymerization and depolymerization (Chen et al. 2000; Moon & Drubin 1995). Cofilin, well known to be expressed ubiquitously in all eukaryotic cell types and to be essential for cell motility (Nishida et al. 1984; Nishida et al. 1987; Abe et al. 1996; Chen et al. 2001; Moon & Drubin 1995), is a member of the actin depolymerizing regulatory molecules, and controls actin reorganization by depolymerizing and severing actin filaments (Rosenblatt & Mitchison 1998). Previously, we and other investigators provided experimental evidence that these activities of cofilin are reversibly regulated by a serine/threonine kinase, LIM kinase 1 (LIMK1), and a protein phosphatase, Slingshot, through phosphorylation and dephosphorylation of Ser-3, with the phosphorylated form being inactive (Mizuno et al. 1994; Arber et al. 1998; Yang et al. 1998; Nagata et al. 1999; Niwa et al. 2002).
Rho-related small GTPases, including Rho, Rac and Cdc42, are known to play a central role in adhesion, cell shape formation and motility in a variety of mammalian cell types, such as fibroblasts, neural cells and haematopoietic cells (Hall 1998; Benard et al. 1999; Sander et al. 1999). They regulate actin cytoskeletal reorganization through various effector proteins, which interact with active GTP-bound forms of the Rho family, in their signal transduction pathways (Bar-Sagi & Hall 2000). LIMK1 is activated through phosphorylation at Thr-508 within the activation loop in its kinase domain by the serine/threonine kinases, p21-activated kinase (PAK) and Rho-associated kinase (ROCK), which are downstream effectors of Rac/Cdc42 and Rho, respectively (Edwards et al. 1999; Maekawa et al. 1999; Ohashi et al. 2000). Therefore, both Rac/Cdc42-PAK and Rho-ROCK signal transduction pathways activate LIMK1, which in turn phosphorylates cofilin, thus regulating actin reorganization (Rosenblatt & Mitchison 1998; Arber et al. 1998; Yang et al. 1998; Toshima et al. 2001; Nishita et al. 2002).
Recently, we developed a coculture system involving HS/PCs and the bone marrow stromal cell types, HESS-5 and HESS-M28 (Tsuji et al. 1996, 1999a, 1999b; Nakamura et al. 1999; Tsuji et al. 2000; Wang et al. 2003). HESS-5 and HESS-M28 cells possess long-term haematopoietic-supportive abilities for not only CD45+/CD34+ cells but also CD45+/CD34– Lineage marker-negative cells. Additionally, they can promote the proliferation of HS/PCs, which have the potential for long-term reconstitution and multiple differentiation in non-obese diabetic SCID mice (NOD/SCID mice: Tsuji et al. 1999b; Nakamura et al. 1999; Tsuji et al. 2000; Wang et al. 2003). The human erythroleukaemia (HEL) cell line is thought to be derived from HS/PCs, and HEL cells were stimulated to proliferate via the CD18 integrin by coculture with HESS-5 cells (Tsuji et al. 1998). In these culture conditions, HS/PCs migrate and proliferate underneath the stromal cell layer making it a useful and suitable model to investigate molecular mechanisms of self-renewal of HSCs, and the proliferation, migration and homing of HS/PCs.
In the present study, we investigated the molecular mechanisms of actin cytoskeletal reorganization underlying morphological changes in HEL cells during the course of migration. We have found that HEL cell migration underneath HESS-M28 cells is initiated by cell adhesion via the CD29 integrin on HEL cells. Following cell adhesion of HEL cells to HESS-M28 cells, filamentous actin localizes to the contact site between them and a membrane protrusion is formed and extended for directional movement. We also provide evidence that LIMK1 activation and cofilin phosphorylation mediated by the Rho-ROCK and Rac/Cdc42-PAK pathways are critical for HEL cell migration.
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Results |
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Previous studies have shown that HS/PCs have potent migration properties beneath stromal cell layers (Tsuji et al. 1999a, 1999b; Nakamura et al. 1999). It is difficult to investigate and understand the molecular mechanisms underlying the migration of either CD34+ cells or CD34+/CD38– cells freshly isolated from haematopoietic organs, because these populations are known to be heterogeneous. Therefore, we examined the migration ability of haematopoietic cell lines underneath a confluent layer of haematopoietic-supportive HESS-M28 cells (Fig. 1A). Amongst the haematopoietic cell lines, HEL cells have the most potent migration properties underneath HESS-M28 cells and this shows good correlation with differentiation levels. Differentiation of HEL cells was thought to be the early blast and/or erythroblast stage as they can differentiate into macrophage-like cells and gpIIb/IIIa-positive megakaryocytes, upon stimulation with 12-O-tetradecanoylphorbol 13-acetate or erythrocyte precursors following exposure to hemin (Martin & Papayannopoulou 1982; Koeffler 1983). Thus, HEL cells have multipotent differentiation properties, at least in a myeloid-erythroid lineage, and are thought to be a useful tool for studying the proliferation and differentiation of HS/PCs.
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Actin cytoskeletal reorganization during HEL cell migration
It has been established that reorganization of the actin cytoskeleton is both essential and central to the process of cell migration and is also an early cellular response to cell adhesion and cell shape change (Hall 1998; Rosenblatt & Mitchison 1998; Bar-Sagi & Hall 2000; Pantaloni et al. 2001; Etienne-Manneville & Hall 2002). We therefore investigated both morphological changes and actin cytoskeletal reorganization during HEL cell migration beneath a HESS-M28 cell layer. For fluorescent microscopy we generated stable transformants of HEL and HESS-M28 cells expressing yellow-fluorescent protein (YFP) and cyan-fluorescent protein (CFP), respectively. Filamentous actin was stained with Alexa Fluor 594-conjugated phalloidin (red signal).
YFP-HEL cells (green signal) were seeded on to a confluent layer of CFP-HESS-M28 cells (blue signal). After 1 h of coculturing, cells were fixed and stained with Alexa Fluor 594-phalloidin. Time-course images of YFP-HEL cell migration underneath CFP-HESS-M28 cells were selected and viewed by x–z projections of the entire complement of optical sections from the coculture using confocal laser microscopy (Fig. 2). First, YFP-HEL cells adhered to CFP-HESS-M28 cells and filamentous actin (yellow signal) was localized to a specific area at the bottom of the HEL cells (Fig. 2A,a–e). In the initial stages of HEL cell migration, the cells moved to (or along) the cell edges of HESS-M28 cells, in which stress-fibre formation was observed (red), and filamentous actin was reorganized to generate a small frontal protrusion in the direction of migration advancement, to the cell edges of HESS-M28 cells (Fig. 2B,a–d). At this point, HEL cells appear on a stress-fibre (red image in Fig. 2d,e) at the cell edges of HESS-M28 cells (Fig. 2Be). Subsequently, the front protrusion was elongated and its length was then longer than that of the cell body (Fig. 2Ca). HEL cells migrated from the cell edges of HESS-M28 cells, its protrusion being under the stress-fibers of the HESS-M28 cells, whereas the HEL cell bodies remained on the outside of the HESS-M28 cell edges (Fig. 2C,a–c). Filamentous actin was localized at the leading edge of HEL cell protrusions (Fig. 2Ca,e) and following migration, HEL cells were flattened under the HESS-M28 cell layers with the loss of filamentous actin (Fig. 2D,a–e).
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Migration of HEL cells beneath HESS-M28 cell layers is predicted to be triggered by interactions between surface molecules of both cells. Among the known adhesion molecules, the role of integrins in early haematopoiesis has been well studied and some integrin family members have been shown to regulate the adhesion, migration and proliferation of HS/PCs (Giancotti & Ruoslahti 1999). Furthermore, our previous studies have demonstrated that many kinds of integrins are expressed on the cell surface of HEL cells (Tsuji et al. 1998). To identify surface molecules on HEL cells that trigger migration, we examined the inhibitory effects on HEL cell migration of neutralizing monoclonal antibodies (mAbs) against integrin family members. HEL cell migration was remarkably reduced when the cells were pretreated with anti-CD29 (integrin ß1 chain) mAb (Fig. 3, clone Lia1/2), suggesting that it is regulated by adhesion, via the CD29 integrin, to cell surface molecules on HESS-M28 cells.
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We next examined the localization of CD29 on HEL cells in the initial step of HEL cell migration (Fig. 4A). After the coculture between HEL and HESS-M28 cells for 15 min, cells were fixed and stained with anti-CD29 mAb (clone K20) and Alexa Fluor 594-phalloidin for filamentous actin. Filamentous actin was detected at the front of membrane protrusion of HEL cells and CD29 was localized in the protrusion of migrating HEL cells (Fig. 4A). These results suggest that adhesion via the CD29 integrin is involved in HEL cell migration by stimulating actin cytoskeletal reorganization.
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Rho GTPases and their downstream effectors mediate HEL cell migration
It is generally recognized that reorganization of the actin cytoskeleton in most mammalian cells is regulated by the Rho-GTPase family members Rho, Rac and Cdc42, and by their downstream effectors. To investigate the possible involvement of these Rho-GTPases in HEL cell migration, we expressed dominant negative forms (RhoN19, RacN17, Cdc42N17) in the cells and examined their inhibitory effects (Fig. 5). HEL cells expressing YFP-fused dominant-negative forms of Rho GTPases were isolated using a flow cytometer, following a deprivation procedure for dead cells, with over 95% purity (data not shown). Expression of RacN17 dramatically reduced the relative ratio of HEL cells adhered to HESS-M28 cells (Fig. 5A). In contrast, expression of both RhoN19 and Cdc42N17 significantly inhibited HEL cell migration (Fig. 5B), although the numbers of adherent cells were unaffected (Fig. 5A).
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LIMK1 specifically phosphorylates cofilin at Ser-3 and induces actin reorganization by inhibiting the actin-depolymerization activity of cofilin. The coordinate regulation of actin reorganization participates in biological responses such as cell adhesion, movement and migration (Hall 1998; Pantaloni et al. 2001; Etienne-Manneville & Hall 2002). To investigate the possible involvement of LIMK1 in HEL cell migration, we examined the effect of a cell permeable peptide inhibitor for LIMK1, S3-peptide, which contains the phosphorylation site of cofilin and a cell-permeable sequence motif of penetratin (Nishita et al. 2002). As a control, we used the RV peptide, which contains the reverse sequence of cofilin and the penetratin sequence. HEL cells were treated with various concentrations of these peptides for 30 min and then seeded on to a confluent layer of HESS-M28 cells. S3 peptide, but not RV peptide, significantly inhibited HEL cell migration in a dose-dependent manner (Fig. 8A). Next, we examined the effect of active and inactive mutants of cofilin (Cofilin S3A and S3E, in which Ser-3 is replaced by alanine and glutamic acid, respectively) on HEL cell migration which was markedly decreased by expression of both mutants (Fig. 8B).
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The previous analyses suggested that cofilin phosphorylation by LIMK1 participates in the regulation of HEL cell migration through actin cytoskeletal reorganization. The levels of endogenous cofilin in mammalian cells are very high and any increase in phosphorylation levels is likely to be limited to the site of actin reorganization in response to external stimuli (Toshima et al. 2001; Nishita et al. 2002). Therefore, we directly measured phosphorylated cofilin, during the process of HEL cell migration, by x–z projections of the entire complement of optical sections from the cocultures using confocal laser microscopy. YFP-HEL cells were seeded on to a confluent layer of CFP-HESS-M28 cells. After 1 h of coculturing, cells were fixed and stained with anti-phosphorylated cofilin antibody. The different images of the protrusion formation during HEL cell migration were selected and analysed by x–z projections of the entire range of optical sections and by x–y projections of 4 optical sections at various distances from the bottom (Fig. 9A–C). HEL cells in the initial migratory phase, in which large amounts of filamentous actin accumulate in the front of the membrane protrusions (Fig. 2B). Phosphorylated cofilin (yellow signal) also localized predominantly at the front of the membrane protrusion of YFP-HEL cells (green image) underneath CFP-HESS-M28 cells (blue image) during migration. At the end of the migration process, phosphorylated cofilin could not be detected in the migrating HEL cells, which did not form the protrusion. However, the migrating HEL cells are able to move underneath HESS-M28 cells. In this case, phosphorylated cofilin was also detected at the front at the membrane protrusion of HEL cells (data not shown). Additionally, phosphorylated cofilin (red signal) was detected at the cell edge of HESS-M28 cells, where large amounts of stress fibers had formed.
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Discussion |
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The actin cytoskeleton mediates a variety of essential biological functions in all eukaryotic cells and provides the driving force for dynamic aspects of cell behaviour such as cell migration, phagocytosis, cytokinesis, polarization, axon guidance and branching. In the present study, we developed a coculturing system, composed of YFP-HEL and CFP-HESS-M28 cells, and successfully observed both dramatic morphological changes and actin reorganization of HEL cells in three-dimensional analyses using confocal laser microscopy (Fig. 2). Filamentous actin was observed at the front of migrating cells in the initial phase of locomotion, and HEL cells then migrated underneath the stromal cell layer from the cell edge of stromal cells, where actin stress fibers of HESS-M28 cells can also be observed. Interestingly, actin reorganization, detected with phalloidin, in HEL cells was observed only at the specific areas at the bottom of HEL cells that adhered to HESS-M28 cells and then also in the small protrusions during the initial phase of migration. This observation suggests that direct cellular interaction between HEL cells and HESS-M28 cells is essential for triggering actin cytoskeleton reorganization during HEL cell migration.
Among the molecules known to direct cell-cell contact, the integrin family members play important roles in the regulation of cell adhesion, morphological changes, migration and homing (Giancotti & Ruoslahti 1999). In HEL cells, many integrin family members are expressed on the cell surface (Tsuji et al. 1998) and the treatment of neutralizing antibody against the integrin ß1 chain, CD29, significantly inhibits migration of HEL cells underneath HESS-M28 cells (Fig. 3). Furthermore, CD29 localized in membrane protrusion of the migrating HEL cells and CD29-ligation induced both actin filament assembly and production of short protrusions (Fig. 4). These findings suggest that actin reorganization induced by CD29 signalling is critical for induction of a polarized morphology and the directional movement of HEL cells in the migration process.
Integrin-induced and chemokine-induced signals are known to stimulate activation of members of the Rho family of small GTPases and also successive actin cytoskeletal reorganization (Clark et al. 1998; Hall 1998; Rosenblatt & Mitchison 1998; Bar-Sagi & Hall 2000; Pantaloni et al. 2001; Etienne-Manneville & Hall 2002). CD29-mediated signalling has an essential role in cell migration (Figs 3 and 4) and expression of dominant negative forms of Rho, Cdc42 and Rac in HEL cells significantly inhibited membrane protrusion formation induced by CD29-ligation and, consequently, HEL cell migration (Figs 5 and 6). Rho is required for adhesion and movement and it has been suggested that its activity might be restricted to the rear of the cell in order to generate the retraction forces required to move the cell body during macrophage chemotaxis (Allen et al. 1998; Bar-Sagi & Hall 2000). In the present study, the polarity formation induced by CD29-ligation and the resulting successive directional movements are essential for HEL cell migration (Figs 2 and 4). Rac is also crucial for generating the filamentous actin-rich lamellipodial protrusions that are thought to have major roles in driving cell movement (Etienne-Manneville & Hall 2002). In our present study, we showed that expression of dominant negative form of Rac caused HEL cell detachment from HESS-M28 cells (Fig. 5A), suggesting that the Rac pathway is essential for adhesion of HEL cells to stromal cells.
A Rho-associated kinase, ROCK, is a serine/threonine kinase that is implicated in Rho-mediated actin reorganization such as formation of stress fibers and focal adhesions (Leung et al. 1996; Amano et al. 1997). Two substrates, myosin light chain (MLC) and an interacting phosphatase, are known to regulate the assembly of actin-myosin filament bundles (Machesky & Hall 1997). ROCK leads to an increase in MLC phosphorylation and the inhibition of MLC phosphatase, and thereby induces actomyosin-based contractility (Kimura et al. 1996). In contrast, a downstream effector of Rac/Cdc42, PAK, leads to a decrease in MLC phosphorylation through inhibition of MLC-kinase and thereby induces the loss of actomyosin-based contractility, an opposing event to that induced by Rho activation (Sanders et al. 1999). HEL cell migration beneath HESS-M28 cells was reduced by exposure of the cells to Y-27632, a specific inhibitor of ROCK (Fig. 7A). Moreover, the expression of PAK-AI, which inhibits the activation of endogenous PAK (Frost et al. 1998), also significantly blocked the migration of HEL cells (Fig. 7B). These results suggest that the extent of MLC phosphorylation, which is oppositely regulated by the Rho-ROCK and Rac-PAK pathways, is necessary for HEL cell migration through a mutual interplay between these pathways.
Actin reorganization regulated by the LIMK1-cofilin pathway is known to play an essential role in the rapid turnover of actin filaments and contribute to various cell responses such as cell adhesion, movement, morphogenesis and chemotaxis (Rosenblatt & Mitchison 1998; Chen et al. 2000; Toshima et al. 2001; Nishita et al. 2002). In contrast to MLC contractility, which undergoes opposing regulation, the Rho-ROCK and Rac-PAK pathways activate LIMK1, which results in a convergence of these signals to increase cofilin phosphorylation at Ser-3 (Agnew et al. 1995; Moriyama et al. 1996). Phosphorylation of cofilin blocks its interaction with actin, thereby inhibiting its actin-depolymerizing and severing activities (Condeelis 2001). Treatment of HEL cells with S3 peptide, which specifically inhibits the kinase activity of LIMK1, significantly reduces HEL cell migration in a dose-dependent manner (Fig. 8). Furthermore, the expression of not only cofilin S3A but also cofilin S3E significantly down-regulates HEL cell migration (Fig. 8B). It has been suggested that cofilin S3A has increased actin-binding and causes successive depolymerizing of actin whereas the S3E mutant mimics phosphorylated cofilin. Dissociated phosphorylated cofilin can promote actin filament turnover following dephosphorylation in a specific feedback pathway, probably via Slingshot phosphatases that target cofilin (Niwa et al. 2002). Transduced cofilin S3E may sequester endogenous cofilin phosphatase activity, thereby leading to inactivation of the endogenous protein and inhibition of the rapid turnover of actin filaments needed for successive cell movements. In contrast, LIMK1 would play a role in stabilizing actin filaments by inactivating cofilin and promoting formation of directional protrusions for migration and lamellipodia and contractile structures for cell movement.
Phosphorylated cofilin is localized at a specific site and is thought to regulate actin reorganization in response to biological stimuli, and is therefore a very low percentage of the total endogenous protein (Toshima et al. 2001; Nishita et al. 2002). In the initial stages of HEL cell migration, filamentous actin was strongly induced in the short protrusions where phosphorylated cofilin was also localized (Figs 2Ba and 9Aa). In HEL cells with a long rod-like stem, filamentous actin was detected at the bottom and was also observed in large amounts at the front of the protrusion but not in whole region of the rod-like stem (Fig. 2C). These morphological features suggest that the flattened protrusions of HEL cells following migration are maintained by physiological pressure from the thin spaces under the areas of adhesion to HESS-M28 cells and that filamentous actin, which is required for the directional movement and elongation of the protrusion, is therefore accumulated at the front of the protrusion. The morphology of HEL cell protrusions also resembles filamentous actin-rich lamellipodia rather than filopodia. In this phase of migration, phosphorylated cofilin is also localized at the front of the protrusion rather than the stem of the protrusion (Fig. 9C). These observations therefore suggest that the LIMK1-cofilin pathway is essential for the migration of HEL cells.
The evidence presented in this study provides novel findings that the migration of HEL cells, a multipotent haematopoietic progenitor cell line, is regulated by actin reorganization through a Rho-GTPase/LIMK1/cofilin pathway. Such findings greatly increase our understanding of the migration, movement and homing of HS/PCs into haematopoietic microenvironments.
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FuGENE 6 Transfection Reagent was purchased form Roche Diagnostics (Manheim, Germany). G418 solution was purchased from PAA (Linz, Austria). The Dead Cell Removal Kit was purchased from Miltenyi Biotec (Bergisch Gladbach, Germany). Y-27632 was purchased from Calbiochem (San Diego, CA, USA). Synthetic S3 and RV peptides were designed and synthesized as previously described (Aizawa et al. 2001). Rabbit polyclonal antibody against Ser-3 phosphorylated cofilin (phosphorylated-cofilin) was prepared as previously described (Toshima et al. 2001). Fluorescein isothiocyanate (FITC)-conjugated F(ab')2 fragment to mouse IgG was purchased from ICN Pharmaceuticals (Aurora, OH, USA). Rhodamine-conjugated anti-rabbit IgG antibody was purchased from Chemicon (Temucula, CA, USA). Alexa Flour 594-conjugated phalloidin was purchased from Molecular Probes (Eugene, OR, USA). Mouse monoclonal antibody against CD29 (clone K20) was purchased from Immunotech (Cedex, France). Neutralizing monoclonal antibodies against human CD29 (clone Lia1/2), CD49d (clone HP2/1), CD49f (clone GoH3), CD18 (clone 7E4), CD11c (clone BU15) and CD41 (clone P2) were purchased from Immunotech. Neutralizing monoclonal antibodies against human CD18 (clone MHM23), CD11a (clone MHM24), CD11b (clone 2LPM19c) and CD11c (clone KB90) were purchased from DAKO (Glostrup, Denmark). Neutralizing monoclonal antibody to human CD11c (clone B-l6) were purchased from Pharmingen (San Diego, CA, USA). A mouse IgG fraction, as a control antibody, was purchased from Zymed Laboratories Inc (San Francisco, CA, USA).
Plasmids pEYFP-C1 and pECFP-C1 vectors were purchased from Clontech (Palo Alto, CA, USA). Expression plasmids for RhoN19, RacN17, Cdc42N17 and PAK-AI, cofilin S3A, and cofilin S3E in the pEYFP-C1 vector were constructed as previously described (Nishita et al. 2002).
Cells and transfection
The haematopoietic-supportive stromal cell line, HESS-M28, was derived from murine bone marrow as previously described (Tsuji et al. 2000) and maintained in minimal essential medium (Sigma, St. Louis, MO, USA) supplemented with 10% horse serum (JRH, Lenexa, KA, USA). The human erythroleukaemia cell line, HEL, was purchased from the Japan Cell Research Bank and maintained in RPMI-1640 medium (Sigma) supplemented with 5% foetal bovine serum (FBS: JRH). Human monocytic leukaemia cell line, U937, human EBV-transformed B lymphoblastoid cell line, IM-9, and human leukaemia T cell line, Jurkat, were purchased from Japan Cell Research Bank, and maintained in RPMI-1640 medium (Sigma) supplemented with 10% FBS (JRH). For transient expression in HEL cells, 2 x 107 cells were incubated with 10 µg of expression plasmids in 400 µL of electroporation medium (RPMI-1640 medium containing 10% FBS) and electroporated at 350 V and 700µF, using a Gene Pulser II (Bio-Rad, Hercules, CA, USA), then cultured for 8-10 h in outgrowth medium (RPMI-1640 medium containing 10% FBS). For the isolation of stable transformants, YFP-transduced HEL cells were selected under 0.8 µg/ml G418 and single clones were isolated by flow cytometry using the EPICS ALTRA (Beckman Coulter, Fullerton, CA, USA). To isolate a CFP-stable expressing clone of HESS-M28, cells were transfected with pECFP-C1 expression plasmids using FuGene, selected under 0.8 µg/ml G418, and a single cell colony was purified by flow cytometry.
Cell staining
HEL cells were seeded on to confluent layers of HESS-M28 cells on 22 mm-glass coverslips and incubated for various periods. To analyse the effect on filamentous actin formation by ligation to anti-CD29 antibody, HEL cells were also plated on to 8-well culture slides (Becton Dickinson) that had been precoated with either 1 µg/ml anti-CD29 antibody (clone Lia1/2) or 1 µg/ml control anti-CD18 antibody (clone 7E4). To detect filamentous actin, cells were fixed with 4% formaldehyde and permeabilized with PBS containing 0.1% Triton X-100. After blocking with PBS containing 1% BSA for 30 min, cells were incubated with Alexa Fluor 594-conjugated phalloidin for 30 min at room temperature. To detect the localization of CD29 and filamentous actin, cells were fixed with 4% formaldehyde and permeabilized with PBS containing 0.1% Triton X-100. After blocking with PBS containing 1% BSA for 1 h, cells were incubated with anti-CD29 antibody (clone K20) for 1 h at room temperature. After being washed with PBS, cells were incubated with FITC-conjugated F(ab')2 fragment to mouse IgG and Alexa Fluor 594-conjugated phalloidin for 30 min at room temperature. To detect phosphorylated cofilin, cells were fixed with 4% formaldehyde and cold methanol. After blocking with PBS containing 1% BSA for 1 h, cells were then incubated with anti-phosphorylated cofilin antibody for 1 h at room temperature. After being washed with PBS containing 0.05% Tween-20, cells were incubated with rhodamine-conjugated anti-rabbit IgG antibody for 1 h at room temperature. Cells were observed by confocal microscopy using a TCS SP2 AOBS (Leica Microsystems, Tokyo, Japan) or by fluorescent microscopy using an Axiovert S100 (Carl Zeiss, Oberkochen, Germany). The images acquired by confocal microscopy were processed with Leica Confocal Software (Leica). Image acquisition from the Zeiss inscribe was made with a cooled CCD camera using Quantix (Photometrics), and the images were processed with Metamorph software (Universal Imaging Corp.).
Migration assay
For analysis of the effects of specific antibodies against different integrin adhesion molecules, HEL cells (2 x 105) were pre-incubated for 30 min in RPMI-1640 medium containing 10% FBS and the antibody solution at a concentration of 10 µg/ml and then seeded on to a confluent layer of HESS-M28 cells in a 24-well plate. After 2 h of culture, cells were fixed with 3% formaldehyde and washed with PBS. The migration ratios were calculated from the numbers of migrated and adhered HEL cells.
For analysis of the effects of Y-27632, HEL cells (3 x 105) were pre-incubated for 30 min in RPMI-1640 containing 10% FBS, in the presence or absence of the indicated amounts of Y-27632 and then seeded on to a confluent layer of HESS-M28 cells in a 12-well plate. After 2 h of culture, cells were fixed in 3% formaldehyde and washed with PBS. The migration ratios were calculated from the numbers of migrated and adhered HEL cells.
For analysis of the effects on HEL cell migration underneath HESS-M28 cells by RhoN19, RacN17, Cdc42N17, PAK-AI, cofilin S3A and cofilin S3E, specific expression plasmids were transfected by electroporation as described above. After culturing for 8–10 h, live cells were purified using a Dead Cell Removal Kit (Miltenyi Biotec), and EYFP-expressing cells were sorted using a flow cytometer (EPICS ALTRA, Beckman Coulter). The purity of the isolated cells was over 95%. Purified cells were pre-incubated for more than 1 h in RPMI-1640 medium containing 10% FBS and plated on to a confluent layer of HESS-M28 cells in 24-well glass-bottom plates (Asahi Techno Glass, Tokyo, Japan). After 3 h of culture, cells were fixed in 3% formaldehyde, washed with PBS and photographed on an Axiovert S100 fluorescent microscope (Carl Zeiss, Oberkochen, Germany). Adhesion ratios were calculated from the numbers of input and adhered EYFP-positive HEL cells. Migration ratios were calculated from the numbers of migrated and adhered EYFP-positive HEL cells. Then, relative ratios for HEL cell adhesion and migration were calculated from the ratios of mock and each Rho GTPase family members.
To analyse the effects of S3 peptide and RV peptide, HEL cells (3 x 105) were pre-incubated for 30 min in RPMI-1640 containing 10% FBS, in the presence or absence of the indicated amounts of peptide and then seeded on to a confluent layer of HESS-M28 cells in a 12-well plate. After 2 h of culture, cells were fixed with 3% formaldehyde and washed with PBS. The migration ratios were calculated from the numbers of migrated and adhered HEL cells.
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Acknowledgements |
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Footnotes |
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* Correspondence: E-mail: t-tsuji{at}nifty.com, t-tsuji{at}rs.noda.tus.ac.jp
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Received: 6 December 2003
Accepted: 22 January 2004
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