1 Department of Physiology, Nagoya University Graduate School of Medicine, 65 Tsurumai Showa-ku, Nagoya Aichi 4668550, Japan
2 Department of Physical Therapy, Nagoya University School of Health Sciences, 1-1-20 Daikominami Higashi-ku, Nagoya Aichi 4618673, Japan
3 ICORP, Cell Mechanosensing Project, Japan Science and Technology Corporation, 65 Tsurumai Showa-ku, Nagoya Aichi 4668550, Japan
*Author for correspondence (e-mail: msokabe{at}med.nagoya-u.ac.jp)
Accepted May 22, 2001
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SUMMARY |
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Key words: Focal contact, Evanescent light microscopy, Integrin, Cytoskeleton, Endothelial cells
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INTRODUCTION |
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Two types of cellular mechanism have been proposed for integrin clustering. The first model suggests that the clustering is dependent on the lateral diffusion of integrins in the plasma membrane (Sheets et al., 1995; Miyamoto et al., 1995; Regen and Horwitz, 1992). It is conceivable that unoccupied integrins that have been released from the substrate at the rear of the cell diffuse laterally in the plasma membrane and arrive at the leading edge to become a source for making fresh FCs. However, it is unclear in this scheme why they would not attach to the substrate again before reaching the leading edge, where they are needed. The second model suggests that intracellular integrins, which have already been internalized from the cell membrane, are translocated to the site where FCs are newly formed (Lawson and Maxfield, 1995; Bretscher, 1989). Exocytosis of vesicles carrying integrins was suggested to be a mechanism to supply integrins for newly forming FCs at the leading edge of migrating cells (Stossel, 1993; Lawson and Maxfield, 1995). It is known that circulating proteins such as transferrin, low density lipoprotein and ferritin receptors are returned to the cell surface at the leading edge of spreading or moving cells (Bretscher, 1983; Bretscher and Thomson, 1983; Ekblom et al., 1983). However, many of these studies used morphological observations of fixed samples and/or biochemical evaluations. Recently, there have been several studies of the morphological changes of FCs (Regen and Horwitz, 1992; Smilenov et al., 1999; Zamir et al., 2000; Pankov et al., 2000). These observed the movement of the cell surface integrins in migrating fibroblasts and demonstrated the relationship between the movement of FCs and stress fibers. However, the precise cellular mechanisms of integrin clustering are not clearly understood during the formation of the FCs in living human umbilical vein endothelial cells (HUVECs).
High-resolution imaging of the clustering process of integrins in living cells is crucial to understanding the molecular mechanism of integrin clustering during FC formation. Total-internal-reflection fluorescence (TIRF) microscopy is potentially useful to visualize the cell-substrate contact region (Schwartz et al., 1980; Tatsumi et al., 1999). Because this contact region is selectively illuminated by evanescent light in TIRF microscopy, fluorescence from fluorophores both in the bulk solution and inside the cells are significantly suppressed compared with epifluorescence (EpiF) microscopy. Using this TIRF imaging system, we have analyzed the process of integrin clustering during FC formation in HUVECs.
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MATERIAL AND METHODS |
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Fluorescent staining of integrins in living HUVECs
HUVECs removed from a culture dish were transferred onto a handmade chamber for TIRF microscopy (described below) and were incubated with FITC-labeled monoclonal antibodies against the extracellular domain of integrin ß1 subunit (FITC-anti-ß1-integrin; CD29, ENDOGEN, USA) for 30-90 minutes (1:50 dilution). After washing the dye three times, the chamber was moved onto a stage of a multimode imaging microscope (described below). In some experiments, HUVECs were incubated without the above antibody for 2 hours and were fixed with 2% paraformaldehyde. The specimen was labeled with the same monoclonal antibody against the extracellular domain as above or with that against the intracellular domains of the ß1 subunit (CD29, DAKO, Denmark). The latter antibody was further labeled with secondary rhodamine-labeled antibody. The staining patterns of FCs labeled with the antibodies were exactly the same with or without preincubation with the antibody against the extracellular domain of the ß1 subunit. This result demonstrated that the antibody used for the live imaging did not perturb the integrin clustering and that integrin dynamics can be imaged by an FITC-labeled antibody against extracellular domains of the ß1 subunit in living cells.
For the simultaneous live imaging of exocytosis and integrin clustering, cells were treated with both anti-ß1-integrin antibody and FM4-64 (1 µM, Molecular Probes, USA), a fluorescent styryl dye, that can selectively label transport vesicles (Henkel and Betz, 1995). The simultaneous observation was made 1 hour after plating the cells.
To examine co-localization of ß1 integrin with vinculin, FITC anti-ß1-integrin antibody-labeled HUVECs were cultured for 2 hours and fixed with 2% paraformaldehyde, permeabilized by 0.5% Triton X-100 and blocked with unlabeled goat anti-mouse IgG (Chemicon International) to suppress the nonspecific staining. This sample was incubated with anti-vinculin (mouse monoclonal, clone V284, Upstate Biotechnology) and TRITC anti-mouse IgG (DAKO, Denmark). No TRITC staining was seen without anti-vinculin treatment.
Multimode microscopy
TIRF optics were incorporated into an inverted microscope (Axiovert 135M; Zeiss, Germany) with differential interference contrast (DIC), EpiF illumination, confocal laser-scanning fluorescence (CLSF) and reflection interference contrast (RIC) optics. Video-enhanced DIC imaging was used to examine the details of the cell structures and EpiF imaging to monitor the intracellular distribution of FITC-antibody-labeled integrins and transport vesicles. A CLSF microscope (Radiance 2000; Bio-Rad, USA) was used to analyze the three-dimensional distribution of the integrins, particularly those near the submembrane region (1 µm above the substrate). For RIC imaging, the EpiF mirror was simply replaced with a 50% mirror and each specimen was illuminated with red filtered (610 nm) light. RIC microscopy was used to view FCs to clarify whether the TIRF images of integrins originated from FCs.
The chamber for the multimode imaging microscope consisted of a 0.25-0.35 mm soda-lime-glass cover slip (No. 3, Matsunami, Japan) and a rectangular prism. The laser beam (473 nm, 3-18 mW, attenuated to 10 mW (Solid State 473, HK5510, Shimadzu, Japan) or 532 nm, 5 mW, (DPSS532, Coherent, USA)) was directed into the rectangular prism (2 mm height BK-7 glass prism, refractive index=1.522, Nihon Ryokyo, Japan). The incident beam propagates in the glass cover slip towards the optical axis of the microscope via multiple internal reflections. The diameter of the each spot of laser light at each internal reflection was 500 µm. The incident angle (
) in our experimental condition was 70°. The depth (Dp) of the evanescent light penetration (surface light intensity divided by e) was estimated to be 75 nm for 473 nm light using Eqn 1 (Burmeister et al., 1994; Zhu et al., 1986):
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where is the wavelength of the laser light, n1 and n2 are the refractive indices of glass and water, respectively. Because the actual penetration depth depends on the refractive index of the medium and the flatness of the substratum, the penetration depth of evanescent light was experimentally examined in a previous study (Tatsumi et al., 1999). TIRF microscope images were focused on a cooled-CCD camera (Micromax, Princeton Instruments, USA) after passing through a 510 nm or 560 nm long-wave-path filter (for green fluorescence from labeled integrins), or a 590 nm long-wave-path filter (for red fluorescence from labeled actins or transport vesicles). The images obtained were processed with the programs MetaMorph (Universal Imaging, USA) and Photoshop (Adobe). During the experiment, the temperature of the chamber was maintained at
37°C.
In the bleaching and recovery experiments, a higher power of laser (18 mW, 30 seconds exposure) was used to bleach the FITC fluorescence only in the evanescent field, and the recovery of the fluorescence was examined with smaller power and short exposure of the laser light (1 mW, 1 second).
Drugs
The drugs used were cytochalasin D (Sigma), butanedione monoxime (BDM; Sigma), ML9 (Biomol, USA), ML7 (Biomol, USA), colchicine (Sigma); bodipy-labeled phalloidin (Molecular Probes, USA), antibodies for integrin (FITC-anti-ß1 integrin; anti-human CD29, ENDOGEN, USA or anti-human CD29, DAKO, Denmark), octadecyl rhodamine-B chloride (O246, Molecular Probes, USA), FM4-64 (Molecular Probes, USA), goat anti-mouse IgG (Chemicon International, USA), anti-vinculin (mouse monoclonal, clone V284, Upstate Biotechnology, USA) and TRITC-anti-mouse IgG (DAKO, Denmark). Other drugs were from Sigma (USA) or Wako (Japan).
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RESULTS |
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Time-lapse imaging of integrin cluster formation at FCs
The dynamic process of the formation of integrin clusters at FCs was examined in living cells by time-lapse TIRF microscopy. TIRF images were recorded every 3 minutes from 1 hour after plating for 15 minutes, because cells showed most active integrin clustering at FCs between 1 hour and 2 hours after plating. Analysis of the integrin clustering at FCs (329 FCs from 10 cells) during the first 30 minutes showed that 10% of FCs were elongated, 3% were shortened and 87% were unchanged. The average length of the unchanged FCs at 2 hours after plating was 7.01±1.64 µm (n=50). This value was similar to that of FCs at 6 hours after plating, suggesting that the FCs with this length were matured. There were no differences in the size and density of the FCs between the cells that were incubated with the antibody against extracellular integrin and the cells without incubation with the antibody (see Methods). The period necessary for FC formation (2 hours) was the same in control HUVECs and anti-integrin-antibody-treated HUVECs. These observations suggest that the anti-ß1-integrin antibody was non-perturbing.
Unidirectional elongation of integrin clusters was usually observed over the entire ventral surface of the cells (Fig. 2). The elongation rate and direction of individual integrin clusters were calculated from time-lapse images. The values obtained are summarized in Fig. 2A, demonstrating a clear tendency for the elongation rate in the central region of the cell to be lower than that in the marginal region of the cell. The elongation rate of integrin clusters within 10 µm of the cell margin was 0.42±0.29 µm minute-1 (n=15) and that in the central region was 0.17±0.06 µm minute-1 (n=15), which is significantly lower (P<0.01) (Fig. 2B).
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The intensity profile of the fluorescence of individual elongating integrin clusters at FCs were measured to clarify the details of the FC elongation (Fig. 3). The fluorescence intensity at the extending tip of the integrin clusters in the central region of the cell was relatively low compared with that of the middle part of the integrin cluster and became higher during elongation (Fig. 3A). However, the intensity of fluorescence at the extending tip of the integrin clusters at the marginal region of the cell was always high during elongation (Fig. 3B, arrowheads). These results suggest that a relatively small amount of integrin was supplied at the extending tip of FCs in the central region, whereas a large amount of integrin was supplied at the tip of FCs at the marginal region of the cell.
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Source and mechanism of integrin supply to the FC formation
The distribution of integrins slightly above the integrin cluster at FCs was imaged by CLSF microscopy to examine the source of integrin supplied to the extending tip of FCs. Time-lapse recordings of integrin clusters at FCs were carried out by interchangeable TIRF and CLSF microscopy at 1 hour after plating (Fig. 5). CLSF images at 1 µm above the basal surface of the cell showed the punctate spots of integrins remained near the extending tip of the integrin clusters at FCs in 32% (n=110) of the cases observed. In the remaining cases, diffuse fluorescence images of integrins were observed near the integrin clusters at FCs. These findings suggested that intracellular integrins near the tip of FCs could be a source of integrins that can be used for the FC formation.
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DISCUSSION |
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Two hypotheses for the construction of new FCs have been proposed. First, lateral diffusion of integrins in a local region of the cell membrane (Sheets et al., 1995), which was suggested to be necessary for FC formation (Miyamoto et al., 1995). Second, vesicles that contain integrins translocate from the intracellular space to FC-forming sites (Lawson and Maxfield, 1995). The present findings support the latter hypothesis, because the photobleaching and recovery experiments by evanescent illumination strongly suggested that integrins were supplied from the intracellular space to the elongating site of FCs. Analysis of the intensity profile of integrin fluorescence and simultaneous imaging of elongating FCs by CLSF and TIRF microscopy also support the latter hypothesis. Furthermore, the EpiF (or CLSF) imaging of integrins demonstrated that most were condensed near the nucleus after the endocytosis of the integrins at 30 minutes after the plating and, at 6 hours, these punctate spots of integrins around the nucleus decreased (or disappeared) while those at FCs increased, suggesting that most of the integrins were supplied from intracellular space. Simultaneous imaging of exocytosis of integrin-containing vesicles and elongation of FCs also suggested that the supply of integrins to FCs was mediated by exocytosis. Fig. 8 summarizes our hypothesis for integrin translocation and FC formation. The extension of integrins towards the cell centre might be due to a direct incorporation of integrins by exocytosis to the tip of FCs and/or the lateral translocation of integrins that originated from exocytosis near the tip of FCs. However, we do not exclude the possibility that part of the integrins incorporated in the tip of the elongating FCs could arise from the bleached cell-surface integrins that could not be seen after photobleaching. It is plausible that surface integrins will turn over for a long time. The integrins supplied at the tip of FCs could also be recycled slowly because the site of adhesion could turn over.
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The importance of stress fibers for the FC formation has been reported in several studies. Inhibition of FCs formation by cytochalasin D or BDM has been shown in previous studies (Folsom and Sakaguchi, 1997; Folsom and Sakaguchi, 1999) as well as in the present study. We also showed that vesicles containing integrins were associated with stress fibers and that the stress fibers terminated near FCs within 100 nm. Because the vesicle diameter is 50-300 nm (Heuser, 1980; Orci et al., 1986), the stress-fiber-dependent translocating system could bring vesicles containing integrins to the vicinity of FCs.
Myosin V is associated with punctate structures that are presumably Golgi-derived cytoplasmic vesicles (Espreafico et al., 1992) and/or with small organelles that are colocalized with actin filaments or microtubules (Evans et al., 1997). Under certain conditions, myosin V attached to synaptic vesicles moves along actin filaments (Evans et al., 1998). Inhibition of serine and threonine phosphorylation by H-7 or ML-7 (both block the activity of myosin-light-chain kinase) also resulted in the destruction of actin bundles and FCs (Zamir et al., 2000). The inhibitory action of BDM and ML-9 (and ML-7) on FC formation in this study agrees with the idea that vesicles containing integrins are translocated to FCs in an actomyosin-dependent way (Fig. 8).
The rate of microtubule-based transportation (20-600 µm minute-1) (Vale et al., 1985; Bloom, 1992) was generally higher than that of actomyosin-based transportation (0.1-10 µm minute-1) (Smilenov et al., 1999; Choquet et al., 1997). In the present study, the elongation rate of integrin clusters at FCs was 0.29 µm minute-1 and colchicine had no inhibitory effect on the formation of integrin clusters at FCs, again suggesting the presence of actomyosin-dependent translocation of integrins. The integrin-cluster elongation at FCs in this study was towards the cell centre. The actin-dependent transport was reported to be towards the cell centre in many cases (Palecek et al., 1996; Pankov et al., 2000; Zamir et al., 2000). These observations also agree with the idea that elongation of FCs is dependent on the actin fibers.
The mechanisms of integrin incorporation to the extending end of FCs remain unclear. The present observations using FM4-64 strongly suggested that the incorporation of integrin was mediated by exocytosis. Several studies suggested that an actomyosin-based transport step was involved in the exocytosis, because exocytosis at the base of growing microvilli (Fath and Burgess, 1993) in sea urchin eggs was inhibited by BDM (Bi et al., 1997). The inhibitory action of BDM on FC formation in the present study might be due to the inhibition of exocytosis of integrin-containing vesicles in addition to the inhibition of actomyosin-dependent translocation.
The processes of integrin translocation have been studied in migrating cells (Stossel, 1993; Lawson and Maxfield, 1995; Palecek et al., 1996). In these studies, integrins were inserted at the leading edge of migrating cells. However, the observations in the present study showed that FCs were formed independent of cell migration or shape changes. The FCs in HUVECs in the present study elongated towards the center of the cell in a straightforward way (probably along actin fibers). The elongation of FCs does not directly contribute to the extending process of HUVECs. Formations of FCs in HUVECs would strengthen the cell-substrate contact and stabilize the morphology of the cells.
Recent studies (Regen and Horwitz, 1992; Smilenov et al., 1999; Pankov et al., 2000; Zamir et al., 2000) showed the translocation of surface integrins in the fibloblasts. The present study demonstrated that the elongation of integrin cluster ceased when it reached a length of 7 µm and that the elongated integrin clusters did not move laterally. Therefore, we consider that the FCs we observed here are different from the movable extracellular matrix contacts reported by Pankov et al. (Pankov et al., 2000).
The present study, which focused on the process of FC formation in HUVECs, suggested that the extension of integrins towards the centre is due to the exocytosis of integrin-containing vesicles at FCs. However, the present findings do not exclude the presence of the lateral translocation of surface integrins. The lateral movement of integrins towards the integrin cluster and our model of FC formation are not exclusive; these two mechanisms might both work during FC formation. However, the EpiF (and CLSF) imaging of integrins in the present study showed that most were condensed near the nucleus after the endocytosis of integrins, which will be translocated to FCs, suggesting that most integrins were supplied from the intracellular space in HUVECs. Another possible explanation is that the dynamics of integrin translocation during FC formation in HUVECs might differ from that of the integrin translocation in fibroblasts. The multimode live imaging used in this study will be a powerful tool to clarify the molecular mechanism of FC formation.
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ACKNOWLEDGMENTS |
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