Calreticulin Affects Focal Contact-dependent but Not Close Contact-dependent Cell-substratum Adhesion*

Marc P. FadelDagger , Ewa DziakDagger , Chun-Min Lo§, Jack Ferrier§, Nasrin Mesaeli, Marek Michalakparallel , and Michal OpasDagger **

From the Dagger  Department of Anatomy and Cell Biology, University of Toronto, Toronto, Ontario M5S 1A8, Canada, the § Medical Research Council Group in Periodontal Physiology, University of Toronto, Toronto, Ontario M5S 1A8, Canada, and the  Medical Research Council Group in Molecular Biology of Membranes, Department of Biochemistry, University of Alberta, Edmonton, Alberta T6G 2H7, Canada

    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

We used two cell lines expressing fast (RPEfast) and slow (RPEslow) attachment kinetics to investigate mechanisms of cell-substratum adhesion. We show that the abundance of a cytoskeletal protein, vinculin, is dramatically decreased in RPEfast cells. This coincides with the diminished expression level of an endoplasmic reticulum chaperone, calreticulin. Both protein and mRNA levels for calreticulin and vinculin were decreased in RPEfast cells. After RPEfast cells were transfected with cDNA encoding calreticulin, both the expression of endoplasmic reticulum-resident calreticulin and cytoplasmic vinculin increased. The abundance of other adhesion-related proteins was not affected. RPEfast cells underexpressing calreticulin displayed a dramatic increase in the abundance of total cellular phosphotyrosine suggesting that the effects of calreticulin on cell adhesiveness may involve modulation of the activities of protein tyrosine kinases or phosphatases which may affect the stability of focal contacts. The calreticulin and vinculin underexpressing RPEfast cells lacked extensive focal contacts and adhered weakly but attached fast to the substratum. In contrast, the RPEslow cells that expressed calreticulin and vinculin abundantly developed numerous and prominent focal contacts slowly, but adhered strongly. Thus, while the calreticulin overexpressing RPEslow cells "grip" the substratum with focal contacts, calreticulin underexpressing RPEfast cells use close contacts to "stick" to it.

    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Cells can form many different types of adhesion to the substratum, however, the best known are focal contacts and close contacts (1). Both focal and close contacts have been defined in terms of the closeness of the cell's ventral surface to substratum by interference reflection microscopy (IRM)1 (1, 2). Focal contacts have been extensively characterized, both at the biochemical and microscopic level (for reviews, see Refs. 3 and 4). When viewed by IRM (1, 2), focal contacts appear as discrete (ca. 10 by 0.5 µm) black patches and represent the areas of closest approach of the ventral cell membrane to the substratum (10-15 nm or less) (1, 5). Structurally, focal contacts associate with termini of stress fibers (5) and are the sites of linkage of the actin cytoskeleton to the extracellular matrix via integrins (6). Many "focal adhesion proteins" such as focal adhesion kinase (pp125FAK), paxillin, talin, alpha -actinin, and vinculin have been shown to localize there (3, 4, 7). It is thought that these cytoplasmic proteins have a role in stabilizing the focal contact after its formation and in signal transduction through integrins clustered there (3, 8). Relatively little is known about close contacts. The ventral cell membrane is separated from the substratum by a distance of 20-50 nm in close contacts and they appear as broad gray areas by IRM (1). Close contacts do not contain vinculin or talin nor do they associate with the termini of bundled actin microfilaments (9), but instead they may associate with a cortical meshwork of actin microfilaments (10). Their only distinguishing feature suggested so far may be an accumulation of alpha -actinin (11). Close contacts are the main form of adhesion to the substratum in cells lacking focal contacts. Finally, non-adherent areas of a cell's underside appear white by IRM and are characterized by a cell-substratum separation distance of 100 nm or more.

To investigate the mechanisms of cell adhesiveness, we used retinal pigmented epithelial cell lines displaying fast (RPEfast) or slow (RPEslow) kinetics of attachment to the substratum. Here we show that differential adhesiveness correlates inversely with the expression level of mRNA and protein for the focal contact-associated cytoskeletal protein, vinculin, and the endoplasmic reticulum (ER) resident chaperone, calreticulin. Calreticulin is a multifunctional Ca2+-binding protein of the ER shown to be important in Ca2+ homeostasis (12-16), modulation of gene expression (17-21), correct folding of glycosylated proteins (22), and cell adhesion (16, 23). The RPEslow cells develop prominent focal contacts which likely account for the high strength of adhesion, whereas the RPEfast have no focal contacts but extensive close contacts and consequently require smaller forces to detach. Furthermore, we show that an inverse relationship exists between the level of calreticulin and the level of total cellular phosphotyrosine (Tyr(P)) such that the cells underexpressing calreticulin display a dramatic increase in the abundance of total cellular Tyr(P). In either cell type spatial distributions of calreticulin and Tyr(P) are complementary, with the former being confined to the ER and the latter being found outside of it. This suggests that the effects of calreticulin on cell adhesiveness may involve modulation of the activities of protein-tyrosine kinases or phosphatases.

    EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Materials-- Tissue culture media, fetal bovine serum (FBS), trypsin, trypsin/EDTA, geneticin (G-418 sulfate), and restriction endonuclease were from Life Technologies, Inc. (Canadian Life Technologies, Burlington, ON, Canada). Goat serum, bovine serum albumin, trypan blue, TRITC-conjugated concanavalin A (ConA), nocodazole, sphingosine, thapsigargin, and fibronectin were from Sigma. Chelex 100 resin and Coomassie Brilliant Blue were from Bio-Rad (Mississauga, ON, Canada). All the electrophoresis reagents were purchased from Bio-Rad. The nitrocellulose membranes (0.22 µm pore size) used in immunodetection were from Micron Separations Inc. (Westboro, MA). Chemiluminescence ECL Western blotting system was from Amersham (Oakville, ON, Canada). DNA Molecular Weight Markers III were from Roche Molecular Biochemicals (Mannheim GmbH, Germany). Hoechst 33258 was from Molecular Probes (Eugene, OR). Vinol 205S was from St. Lawrence Chemical (Toronto, ON, Canada). All chemicals were of the highest grade commercially available.

Antibodies-- A well characterized goat anti-calreticulin antibody (24-27) and a rabbit antibody against calreticulin recognizing six carboxyl-terminal amino acids of the protein (QAKDEL) were used to detect calreticulin. Rabbit anti-calreticulin (designated anti-CRT283) was raised against the synthetic peptide QAKDEL (28), which encodes amino acids 386-401 of rabbit calreticulin (29). This antibody is specific for the ER-resident form of calreticulin and it does not recognize other KDEL-containing proteins such as protein-disulfide isomerase, Grp78 (BiP), ERp72, Grp96 (endoplasmin), nor does it recognize calreticulin without the KDEL sequence (28). A mouse monoclonal antibody against vinculin was from ICN ImmunoBiologicals (Montreal, PQ, Canada), a monoclonal mouse anti-Tyr(P) (PY-20) and a rabbit polyclonal antibody against pp125FAK were from Santa Cruz Biotechnology Inc. (Santa Cruz, CA), mouse monoclonal antibodies against actin was from Sigma. Anti-beta 1 integrin was from Dr. M. Ginsberg (Scripps Research Institute) and anti-alpha 5 and anti-beta 1 integrins were from Dr. B. Chan (University of Western Ontario). All secondary antibodies were from Jackson ImmunoResearch Laboratories (West Grove, PA).

Cell Culture-- The RPEslow and RPEfast cells were generated from previously characterized retinal pigment epithelial cell lines, RPE-J and RPE-J+EC (30, 31). The RPE-J cells were immortalized by infection with temperature-sensitive tsA SV40 virus (30). The RPE-J+EC cells were generated by transfection of RPE-J cells with full-length canine E-cadherin DNA (31). Both cell lines were re-cloned using efficacy of cell-substratum adhesiveness as a screening criterion. Two subclones were selected that attached with high and average efficacy, and designated RPEfast and RPEslow, respectively. The cells were grown in high glucose Dulbecco's modified Eagle's medium supplemented with 10% FBS and minimal essential medium non-essential amino acids (Canadian Life Technologies, Burlington, ON, Canada).

Transfections-- Cells were grown in 10-cm dishes and transfected by the DEAE-dextran method (32). Plasmid DNA was purified by QIAGEN column chromatography. Cells were transiently transfected with calreticulin expression vector and 5 µg of pS65T-CI GFP plasmid (CLONTECH, Palo Alto, CA) as described (19). The time of transient transfection was 72 h. Following transfection cells were trypsinized and sorted by flow cytometry using a fluorescence-activated cell sorter (Epics Elite, Coulter Electronics, Hialeah, FL). 40,000 GFP-positive and 40,000 GFP-negative cells were collected, pelleted by centrifugation, washed with PBS followed by incubation for 15 min at room temperature with the lysis buffer containing 1% Nonidet P-40, 0.25% sodium deoxycholate, 1 mM EGTA, 150 mM NaCl, and 50 mM Tris, pH 7.4. The lysis buffer also contained a mixture of protease inhibitors including 1 mM benzamidine, 1 mM phenylmethylsulfonyl fluoride, 0.1 µg of aprotinin/ml, 0.5 µg of leupeptin/ml, 0.5 µg of (N-[N-(L-3-trans-carboxyoxirane-2-carbonyl)-L-leucyl]agmatine/ml, 0.5 µg of pepstatin/ml, 50 µg of L-1-chloro-3-(4-tosylamido)-7-amino-2-heptanone hydrochloride and L-1-chloro-3-(4-tosylamido)-4-phenyl-2-butanone/ml, 0.1 µg of (4-ami-dinophenyl)-methansulfonyl fluoride/ml, 50 µg of phosphoramidon/ml. Cells extracts were centrifuged on spin columns (Bio-Rad) to remove DNA from lysates and subjected in quadruplicate to SDS-PAGE as described (14).

SDS-PAGE and Western Blotting-- Protein samples (40,000 cells/lane for extracts and 2 µg/lane for molecular weight markers) were subjected to SDS-PAGE and Western blotting as described by Mery et al. (14). The primary antibodies were used at the following dilutions in PBS: anti-actin, 1:100; anti-pp125FAK, 1:200; anti-integrins, 1:1000; anti-vinculin, 1:1500; CRT283, 1:300; goat anti-calreticulin 1:300; and anti-Tyr(P) 1:5000. The protein bands in each blot were scanned two-dimensionally using a densitometer (UltroScanXL, Pharmacia) and areas under the curves were calculated using Gel Scan XL software.

RNA Isolation and Northern Blotting-- RNA extraction and Northern blotting was performed as described previously (17). For vinculin mRNA, 572 base pairs of EcoRI fragment of Vinc 1020 cDNA (generously donated by Dr. Sue Craig, Johns Hopkins University) were used as a probe (33). The blots were normalized by probing with human glyceraldehde 3-phosphate dehydrogenase cDNA. The relative abundance of mRNA was determined using a Fujiex BAS1000 PhosphorImager.

Immunolabeling and Microscopy-- For immunolocalization, the cells on coverslips were fixed in 3.7% formaldehyde in PBS for 10 min. After washing (3 times for 5 min) in PBS, the cells were permeabilized with 0.1% Triton X-100 in buffer containing 100 mM PIPES, 1 mM EGTA, and 4% (w/v), polyethylene glycol 8000 (pH 6.9) for 2 min, washed 3 times for 5 min in PBS, and then incubated either anti-calreticulin antibody (diluted 1:50 in PBS), anti-vinculin antibody (diluted 1:50 in PBS), or anti-Tyr(P) (diluted 1:50 in PBS) for 30 min at room temperature. After washing (3 times for 5 min) in PBS the cells were stained with appropriate secondary antibodies for 30 min at room temperature. The secondary antibodies were: fluorescein isothiocyanate-conjugated donkey anti-mouse IgG(H+L) (diluted 1:30 in PBS), Texas Red-conjugated donkey anti-mouse (F(ab')2 used at 1:30 dilution), and dichlorotriazinylamino fluorescein-conjugated donkey anti-goat IgG(H+L) (diluted 1:30 in PBS). TRITC-conjugated ConA was used at 20 µg/ml in PBS. For double labeling incubations with appropriate antibodies were done sequentially. After the final wash (3 times 5 min) the slides were mounted in Vinol 205S which contained 0.25% 1,4-diazabicyclo-(2,2,2)-octane and 0.002% p-phenylenediamine to prevent photobleaching. For actin staining, stock solution of 3.3 mM rhodamine phalloidin in methanol was diluted 1:10 in PBS and incubated with fixed and permeabilized cells for 20 min at room temperature. A Bio-Rad MRC-600 confocal fluorescence microscope equipped with a krypton/argon laser was used for fluorescence and phase-contrast microscopy. For IRM, cells were plated on 15-mm square coverslips and allowed to attach for ~36 h. Cells were fixed with 4% glutaraldehyde for 10 min and washed with PBS and mounted in PBS. IRM was conducted as described previously (1, 34, 35) using a Zeiss IM 35 invertoscope. IRM takes advantage of the interference of wavefronts reflected at the phase boundaries created by the thin layer of culture medium which separates a cell and the glass substratum to which that cell adheres. In monochromatic light, gray levels in the resultant fringe image is indicative of the distance of separation between the cell and the substratum (1, 2, 36, 37). In general, white indicates a separation distance over 100 nm, light-to-dark greys are generated by a separation of 70-20 nm, and black is generated by a cell-substratum gap of less than 15 nm. For further analysis, the brightness of IRM images was normalized using Adobe Photoshop such that all cell-free areas were all displaying a similar gray level range. Next, the images were analyzed in terms of the percentage of area of cell underside of each cell image, which fell under one of four gray level ranges. To do so, the number of pixels falling within a specified gray level range was quantified with Image Pro Plus software. The ranges were designated as follows: "black," "dark gray," "light gray," and "white." For focal contact counting, the cells were labeled with anti-vinculin antibody as described above and the vinculin positive patches were manually counted.

Attachment Assays-- For attachment assays, cells were trypsinized with EDTA-free trypsin, and plated onto tissue culture dishes at an initial density of ca. 40,000 cells per 35-mm dish. Some experiments were conducted on fibronectin-coated culture dishes. Briefly, tissue culture dishes were incubated for 90 min with a fibronectin solution (1 µg/ml in PBS) at 37 °C, then rinsed with cold PBS and used without drying. At selected time intervals, non-adherent cells were washed off the dishes with a brief rinse of cold medium and the remaining adherent cells were trypsinized and counted (in triplicate) on a Coulter Counter (Coulter Electronics, Hialeah, FL). For calcium sensitivity of cell attachment, Ca2+-free medium was prepared in the same manner as high glucose Dulbecco's modified Eagle's medium except for the omission of CaCl2, replacement of vitamin D-Ca2+ pantothenate by Na+-pantothenate, and the use of chelated distilled water. Chelation of water was carried out with Chelex 100 resin at 5 g/10 liter for 1 h with stirring. FBS was chelated with 5 g of Chelex per 100 ml for 1 h with stirring, after which Chelex was filtrated and pH adjusted to 7.2. Ca2+ concentration was measured with F-2000 Hitachi fluorescence spectrophotometer using 1.2 mM Indo-1 Na+ dissolved in dimethyl sulfoxide. Calibration curve of Ca2+ concentrations (0.0-39.8 mM) was made using Calcium Calibration Kit 1. The final Ca2+ concentration of total medium (high glucose Dulbecco's modified Eagle's medium plus FBS) was calculated to be approximately 20 nM.

Electron Microscopy-- Cells were plated at a density of 10,000 cells per 35-mm cell culture dish and allowed to grow for over a week until they were well packed. The culture dishes were washed with PBS (3 × 5 min) before fixing with 2% glutaraldehyde and 2% paraformaldehyde for 30 min. After washing with PBS cells were treated with 0.5% osmium tetroxide for 30 min. Cells were rinsed with distilled water after the next set of PBS washes to prepare them for 30 min en bloc secondary fixation with 1% uranyl acetate. Cells were then rinsed and dehydrated with 5-min single washes with 10, 25, and 50% alcohol, and 2 times 5-min washes with 70, 80, 90, and 100% alcohol and then embedded in Epon. 80-90-nm sections were cut with diamond knife on a microtome and placed on nickel grids. Structural staining with saturated uranyl acetate, washing, and lead citrate staining were 15 min each. The sections were then analyzed on a Hitachi-7000 electron microscope.

Electric Cell-substrate Impedance Sensing (ECIS)-- The apparatus, the model equations, and the frequency scan impedance measurements were as previously reported (38-41). Electrode arrays, relay bank, lock-in amplifier, and software for data analysis were obtained from Applied BioPhysics (Troy, New York). Cells were cultured on a small gold electrode, with a much larger gold electrode serving as the counterelectrode. The total measured resistance is composed of the resistance at the electrode-electrolyte interface, plus the resistance of the current pathway in the electrolyte between the ventral surface of the cells and the electrode, plus the resistance of the current pathway in the electrolyte between the cells. Therefore a difference in measured resistance will indicate a difference in under-the-cells resistance and/or a difference in between-the-cells resistance. The measured capacitance as a function of frequency reflects the rate at which charge moves within the electrode-electrolyte boundary layer, and depends on resistance in the current pathways, the capacitance falling off more rapidly with frequency as the total resistance increases. The data are expressed as normalized resistance (measured resistance in the presence of a cell layer divided by measured resistance of the electrode without cells) versus frequency, and normalized capacitance versus frequency. A difference in the frequency at which the peak of the normalized resistance occurs indicates a difference in the under-the-cells resistance, and thus a difference in cell-substratum separation. As distance between the ventral surface of the cells and the electrode decreases, the peak of the normalized resistance will shift toward lower frequencies (38-42).

Centrifugal Detachment Assay-- To evaluate the forces necessary to detach the RPE cells we modified a centrifugal detachment assay described by Hertl et al. (43). Briefly, 100,000 cells were plated in 12-mm Costar Transwell Clear tissue culture inserts (Corning Costar, Cambridge, MA) and allowed to attach overnight. The following day individual tissue culture inserts were submerged upside down (such that the cell layer is now upside down) in centrifugation media (high glucose Dulbecco's modified Eagle's + 10% FBS + 10 mM HEPES) contained in a 25 × 29-mm Beckman Ultra-Clear centrifuge tube (Palo Alto, CA) with a No. 3 rubber stopper at the bottom. Care was taken to ensure that no air bubbles were trapped at the cell-centrifugation medium interface. Two tubes at a time, each containing a different cell type, were centrifuged using a SW-28 swing out bucket rotor in a Beckman XL-70 Ultra Centrifuge. Tubes were either spun at 30,000 or 60,000 × g for 15 min at 22 °C. Control tubes were left on the counter (1 × g) for 15 min at room temperature. After the application of force the tissue culture inserts were removed and replaced in an upright position. The cells remaining attached were then trypsinized and counted using a Coulter counter. Cells in tissue culture inserts were viewed before and after force application by phase-contrast microscopy to examine cell morphology. Data were expressed as the number of cells remaining attached after the application of force, and pooled from four independent experiments.

    RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Characterization of Adhesive Cell Phenotype of RPE Cells

To characterize the adhesive properties of both RPE cell types, we utilized assays to measure different aspects of cell adhesion, including attachment efficacy, closeness of ventral cell membrane to the substratum, and force required for detachment of cells from the substratum.

RPEfast Cells Attach More Efficiently than RPEslow Cells-- We first examined the attachment kinetics of the cells in normal and low extracellular Ca2+ conditions ([Ca2+] ~20 nM). Initial attachment of the RPEfast cells was unaffected by the low extracellular Ca2+ conditions. Specifically, almost all of the RPEfast cells, both in normal and low extracellular Ca2+ conditions had attached 4 h after plating, but only a little more than half of the RPEslow cells in normal conditions, and only about one-third of those in low extracellular Ca2+ had attached (Fig. 1A). This trend continued when the efficacy of attachment was examined over a longer time course. The RPEfast cells showed no difference in their efficacies of attachment overnight in normal and low extracellular Ca2+, whereas the RPEslow cells did not attach well in low extracellular Ca2+ (Fig. 1B). It was apparent from the efficacies of attachment after 4 h and from the time course of attachment overnight, that RPEfast cells attached overall more effectively than RPEslow cells. The same trend was maintained on fibronectin-coated substrata, but the difference in attachment efficacy between RPEfast and RPEslow cells was less dramatic due to improved attachment of RPEslow cells (data not shown).


View larger version (13K):
[in this window]
[in a new window]
 
Fig. 1.   RPEfast cells attach more effectively than RPEslow cells. Left, attachment efficacy of RPEslow cells and RPEfast cells after 4 h in normal (2 mM) and low (<= 20 nM) extracellular Ca2+ concentration. Data is expressed as the percentage of cells originally plated which attached after 4 h ± S.D. Right, time course of attachment of RPEslow cells and RPEfast cells in normal and low extracellular Ca2+ conditions over 16 h. Data is expressed as number of adherent cells ± S.D. RPEfast cells display similar attachment kinetics over the long term irrespective of extracellular Ca2+ concentration.

RPEfast Cells Have No Focal Contacts-- Examination of the cells by IRM revealed that RPEslow cells had many focal contacts, whereas RPEfast cells had few or no focal contacts (Fig. 2, A and A'). Instead, RPEfast cells had extensive areas of close contacts. As the gray level of an IRM image is indicative of cell-substratum separation distance, we quantified the gray levels in IRM images of both cell types. To do so, the gray levels in micrographs of several cells from each type were analyzed in terms of the percentage of the cell underside which fell under the category of black, dark gray, light gray, and white (Fig. 2B). RPEslow cells had about twice as much black area, but significantly less gray areas than RPEfast cells. To relate the ventral cell membrane closeness to the substratum and the number of vinculin-rich adhesions, which normally colocalize with focal contacts visualized by IRM (44-46), the number of vinculin-positive patches in immunostained cells was quantified for each cell type. There were twice as many vinculin-positive patches in RPEslow cells as in RPEfast cells, which corresponded closely to the 2-fold difference in the percentage of cell areas generating zero order interference (black in IRM) between the two cells types (Fig. 2C).


View larger version (54K):
[in this window]
[in a new window]
 
Fig. 2.   Topography of the underside is much more diversified in RPEslow cells that in RPEfast cells. A, IRM images of RPEslow (A) cells and RPEfast (A') cells. Notice the presence of numerous discrete black focal contacts in RPEslow cells (white arrowheads in A) and the uniform distribution of gray corresponding to close contacts in RPEfast cells. Black fringes at the periphery of all cells are caused by reflection of light from the dorsal cell surface and are unrelated to cell adhesion. B, quantification of gray levels and vinculin positive adhesion patches in RPEslow (n = 13) and RPEfast (n = 12) cells. The bars indicate the average percentage of the imaged cell (± S.E.) belonging to a particular pixel gray level category. The inset shows the average number of vinculin positive adhesion patches (± S.E.) for RPEslow cells and RPEfast cells. Scale bar, 25 µM.

Interestingly, although RPEslow cells had about twice as many focal contacts as RPEfast cells, they also had more white areas, corresponding to distant, non-adherent areas of the ventral membrane. The underside of RPEfast cells on the other hand was more of a uniform gray in the IRM, with less of either extreme in gray levels. This indicates that the ventral membrane of RPEslow cells is undulating, whereas the ventral membrane of RPEfast cells is relatively uniformly separated from the substratum.

RPEfast Cells Have a Uniformly Flat Ventral Membrane-- To examine the topography of the ventral cell membrane more closely, we examined sections cut perpendicularly to the substratum by transmission EM (Fig. 3). RPEslow cells displayed a ventral membrane which varied continuously in its separation from the substratum, averaging 43.0 ± 51.4 nm over a total measured length of 10.90 µm (n = 13) of cell underside. On the other hand, the ventral membrane of RPEfast cells was uniform in its separation from the substratum, which remained at 10.5 ± 1.3 nm over a total measured length of RPEfast cell's underside of 8.43 µm (n = 10). Hence, the EM results fully support our IRM findings in that RPEslow cells have an undulating ventral membrane, whereas RPEfast cells have a uniformly flat underside.


View larger version (101K):
[in this window]
[in a new window]
 
Fig. 3.   Electron microscopy images of RPEslow and RPEfast cells. The cells were sectioned at an orientation perpendicular to the substratum. RPEslow cells have an undulating ventral membrane, whereas RPEfast cells have a ventral membrane that is uniformly flat. Scale bar, 500 nM.

RPEfast Cells Have a Lower Average Cell-substratum Separation than RPEslow Cells-- Fig. 4 shows typical results for normalized resistance and normalized capacitance as a function of frequency from the ECIS frequency scan measurements (n = 8). The RPEfast cells displayed a significantly higher peak value for resistance (4-fold higher), which occurred at a significantly lower frequency (more than 4-fold lower) than for the RPEslow cells. In addition, the capacitance curve for RPEfast cells was shifted to significantly lower frequencies (about 16-fold lower), relative to the capacitance curve for RPEslow cells. The higher peak resistance, and the shift to lower frequencies of the capacitance curve for the RPEfast cells, indicated that the sum of the resistance under the cells and between the cells was higher for the RPEfast cells than for the RPEslow cells. Furthermore, the occurrence of the resistance peak at a lower frequency with RPEfast cells indicated that the resistance under the cells was higher for these cells, and therefore the average distance between the ventral membrane and the gold electrode substratum was smaller for the RPEfast cells than for the RPEslow cells.


View larger version (11K):
[in this window]
[in a new window]
 
Fig. 4.   ECIS properties of RPEslow cells and RPEfast cells. A, normalized resistance; and B, normalized capacitance, as a function of frequency from the ECIS frequency scan measurements (n = 8). The resistance peak (A) for RPEfast cells is significantly higher and occurs at a much lower frequency than for RPEslow cells. Notice that the capacitance curve (B) for RPEfast cells is also shifted to lower frequencies compared with RPEslow cells. Data is expressed in normalized units.

RPEslow Cells Require a Larger Force to Detach than RPEfast Cells-- Finally, in order to quantify the actual adhesive strengths of these two cells types, we carried out a centrifugal detachment assay. The cells were centrifuged at various g forces such that the direction of the force was oriented perpendicular to the cell monolayer. The number of cells remaining attached after the application of differing forces, were then counted. Specifically, each cell type was subjected to forces of 1, 30,000, and 60,000 × g (Fig. 5). We detected no differences between the two cell types in their ability to remain attached after experiencing centrifugal forces of 1 × g (control) and 30,000 × g. After 60,000 × g, however, there were very few RPEfast cells which remained attached (~5% of control), whereas most of the RPEslow cells (~80% of control) were still attached. Thus, we have shown that strength of adhesion was smaller for the RPEfast cells than for the RPEslow cells.


View larger version (19K):
[in this window]
[in a new window]
 
Fig. 5.   RPEfast cells require a smaller centrifugal force to detach than RPEslow cells. Cells were exposed to centrifugal forces of 30,000 and 60,000 × g. At 60,000 × g, RPEslow cells resisted detachment, whereas most of the RPEfast cells have detached. Data is expressed as average number of cells remaining attached ± S.E.

The Cell Adhesion Phenotype Correlates with the Abundance of Calreticulin and Vinculin

Using Northern and Western blot analysis we evaluated the levels of mRNA and protein for calreticulin and vinculin in the RPE cells. RPEfast cells had much lower levels of mRNA and protein for both calreticulin and vinculin than RPEslow cells (Fig. 6, A and B). Densitometric analysis of the bands revealed that RPEfast cells express approximately half the levels of calreticulin mRNA and protein, and vinculin mRNA and protein, respectively, of RPEslow cells. Thus far, we have clearly seen a difference between the level of expression of calreticulin and vinculin and the adhesive phenotype of the RPE cells, such that the cells expressing calreticulin and vinculin more abundantly were less "adhesive." This is contrary to previous results showing that fibroblasts overexpressing calreticulin attached more effectively than cells underexpressing calreticulin (16, 23). Next, we addressed a possible mechanism for the observed differences in adhesive ability of these cells and we examined the expression levels of several other known adhesion proteins. Specifically, we examined the protein levels of actin, alpha 5 and beta 1 integrin, and of pp125 focal adhesion kinase (FAK), known components and regulators of focal contacts. We did not find any significant difference in the expression levels of these proteins between the two cell types (Fig. 6C).


View larger version (20K):
[in this window]
[in a new window]
 
Fig. 6.   Differential expression of calreticulin is accompanied by a differential expression of vinculin. A, identification of calreticulin and vinculin mRNAs in RPEslow cells and RPEfast cells. Note the lower levels of both calreticulin and vinculin mRNAs in RPEfast cells as compared with RPEslow cells. Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) mRNA was also identified to ensure equal loading. B, Western blot analysis of calreticulin and vinculin protein levels in RPEslow and RPEfast cells. Western blot with anti-tubulin antibodies shows uniformity of loading. C, changes in the level of calreticulin expression do not affect the abundance of several other adhesion-related proteins.

Is the Differential Expression of Calreticulin Related to the Differential Expression of Vinculin?-- To address this, the RPEfast cells, underexpressing calreticulin, were transfected for 72 h with a calreticulin expression vector encoding the full-length protein (with signal sequence and KDEL ER retrieval signal). Expression and localization of the recombinant calreticulin was monitored with anti-calreticulin antibodies. Western blot analysis of cellular extracts clearly showed that the level of the ER-resident calreticulin has increased in transfected RPEfast cells. The averaged densitometric analysis of the bands revealed an over 6-fold increase in calreticulin upon transfection of RPEfast cells with a calreticulin expression vector. Increasing the expression of calreticulin in these cells brought about a concomitant increase (approximately 2-fold) in the level of expression of vinculin (Fig. 7A). These findings are further supported by the immunolocalization of vinculin in cells differentially expressing either native or recombinant calreticulin. RPEslow cells had vinculin-rich adhesions which are more numerous and prominent than those in RPEfast cells. Importantly, the number and prominence of these vinculin-rich adhesions increased in RPEfast cells after transfection with the calreticulin vector (Fig. 7B). Microfluorimetry indicated that both the cytoplasmic pool of vinculin and the cytoskeleton-bound vinculin present in cell adhesions were more abundant in the RPEfast cells after up-regulation of calreticulin. The measurements shown are based on the distribution of intensities of fluorescence signal coming from a single confocal optical section and, albeit far from quantitative, give a rough estimation of the distribution of antigen abundance throughout the imaged section. Thus, we have shown that cells expressing higher levels of calreticulin also express higher levels of vinculin, and conversely, cells expressing lower levels of calreticulin also express lower levels of vinculin. Furthermore, an increase in the expression of vinculin is induced by raising intracellular levels of calreticulin by transfection.


View larger version (58K):
[in this window]
[in a new window]
 
Fig. 7.   Overexpression of calreticulin increases the abundance of vinculin and the number and prominence of vinculin containing adhesions. A, immunodetection of vinculin and calreticulin in RPEfast cells transfected with a calreticulin expression vector. Notice that overexpression of calreticulin is paralleled by an increase in the expression of vinculin. Each lane corresponds to a different experiment. B, immunolocalization of vinculin in RPEslow (B), RPEfast (B'), and RPEfast cells transfected with calreticulin expression vector (B''). The graphs show the density of vinculin label (in gray levels) measured along the line indicated in each photograph. In RPEfast cells vinculin-rich attachments are sparse, and the cytoplasmic presence of vinculin is not prominent. Both types of vinculin distributions are restored in transfected RPEfast cells. The average density of cytoplasmic label (in gray levels) is: 52 in RPEslow cells, 31 in RPEfast cells, and 66 in transfected RPEfast cells. Scale bar, 25 µM.

Focal Contacts Can Be Induced to Form in RPEfast Cells

Are RPEfast Cells Lacking Components Necessary in the Assembly of Focal Contacts, or Is It the Stimulus for Assembly Which Is Missing?-- In order to address this question, we treated the RPEfast cells with agents that have previously been shown to induce focal contact formation. Nocodazole, an inhibitor of microtubule polymerization, appears to induce an increase in intracellular contractility (47-49), which is paralleled by a stimulation of focal contact formation (48, 50, 51). Sphingosine is a breakdown product of sphingolipids and it induces Tyr phosphorylation of several substrates, including pp125FAK, which stimulates focal contact formation (52, 53). Treatment of RPEfast cells for 16 h with either nocodazole (1 µg/ml) or sphingosine (10 µM) stimulated formation of prominent stress fibers (not shown) and a marked restoration of numerous vinculin-positive focal contacts (Fig. 8). Western blot analysis of cell extracts from RPEfast cells treated with nocodazole or sphingosine revealed that there was an over 2-fold increase in the abundance of vinculin (Fig. 8A). The abundance of calreticulin did not change appreciably in nocodazole-treated cells but it increased by 50% above untreated cells in cells that were treated with sphingosine (Fig. 8A). We next asked if induction of calreticulin by stressing the ER is also associated with an increased abundance of vinculin. To do so we exposed the calreticulin underexpressing RPEfast cells to thapsigargin. Thapsigargin is an inhibitor of the Ca2+-ATPase responsible for refilling the ER Ca2+ stores after the Ca2+ release (54). The calreticulin gene promoter is activated upon treatment of cells with thapsigargin, which causes increased calreticulin synthesis (55). Western blot analysis of cell extracts from RPEfast cells treated with thapsigargin revealed an increase in the level of expression of calreticulin by 50% above untreated cells. This was paralleled by a similar increase in the expression level of vinculin (Fig. 8). Immunofluorescence microscopy (Fig. 8) demonstrated that the newly induced vinculin assembled into focal contacts, albeit much less effectively than after treatment with either nocodazole or sphingosine. This indicates that the elements necessary for construction of focal contacts are present in the RPEfast cells.


View larger version (110K):
[in this window]
[in a new window]
 
Fig. 8.   Focal contacts can be induced in RPEfast cells. A, Western blot analysis of cell extracts from RPEfast cells treated with nocodazole (Noco), sphingosine (Sphin), and thapsigargin (TG) revealed that treatments with sphingosine and thapsigargin, but not with nocodazole, induced increases in the levels of calreticulin and vinculin relative to the untreated cells (Ctrl). Immunofluorescence shows that the number and prominence of vinculin positive adhesions is restored after treatment of the RPEfast cells by nocodazole (C) and sphingosine (D). The thapsigargin treatment (E) produces an increase in cytoplasmic pool of vinculin but a less prominent increase in focal contacts. B shows untreated RPEfast cells. Scale bar, 25 µM.

Changes in Levels of Calreticulin Expression Inversely Correlate with the Abundance of Total Cellular Phosphotyrosine

It is well established that calreticulin is an ER-resident protein and that vinculin is a cytoplasmic protein. For calreticulin to affect expression of vinculin, there must be an ER signaling pathway involved. The signaling mechanism may sense the level of calreticulin in the ER and convey that information to the nucleus to up-regulate or attenuate gene transcription, in this case, of vinculin. We began examining the possible role of calreticulin as a signaling molecule from inside the ER, with tyrosine kinases/phosphatases as cytoplasmic mediators of calreticulin's effects. A correlation between cell adhesiveness and the overall intracellular Tyr(P) level has been well established (56, 57). We therefore asked if there was a relationship between the level of Tyr(P) and the level of calreticulin in the cell. Western blotting with anti-Tyr(P) antibodies (Fig. 9) clearly demonstrated that the overall Tyr(P) level was much lower in RPEslow cells than in RPEfast cells. We also localized the Tyr(P) signal using indirect immunofluorescence in respect to the ER visualized with TRITC-ConA. ConA has been shown to preferentially accumulated in the region of the rough ER, hence it has become an accepted, albeit nonspecific marker of the ER (58, 59). In either cell type the Tyr(P) labeling was localized outside of the ER and it delineated focal contacts in RPEslow cells and the regions of intercellular contacts in the RPEfast cells (Fig. 9).


View larger version (119K):
[in this window]
[in a new window]
 
Fig. 9.   Total level of cellular Tyr(P) correlates inversely with the abundance of calreticulin. A, RPEslow cells have a much lower level of total cellular Tyr(P) as compared with RPEfast cells as detected by Western blotting with PY-20 antibody. The position of molecular marker proteins is indicated. B and C, localization of Tyr(P) in calreticulin overexpressing RPEslow and calreticulin underexpressing RPEfast cells in relationship to the ER by double labeling immunofluorescence. B, in RPEfast cells, which have no focal contacts, the Tyr(P) signal is concentrated in cell-cell junctions. C, in RPEslow cells the Tyr(P) staining is concentrated mainly in focal contacts (C), but is also detectable in cell-cell junctions (C'). C and C' are two confocal sections taken 3 µm apart. Notice that the Tyr(P) signal is not associated with the ER in any cell (B' and C''). Scale bar, 25 µM.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Work from several laboratories strongly suggests that the abundance of calreticulin and vinculin correlate with cell "adhesiveness," such that cells with more calreticulin and vinculin typically "attach better" (16, 23, 60-63).

Calreticulin and Cell Adhesion

How Can Cells Differing in Abundance of ER-resident Calreticulin Have Such Different Adhesive Properties?-- An attractive working hypothesis was that overexpression of calreticulin caused more effective processing of transmembrane components of focal contacts. So far, however, no difference in the abundance of any of the integrins has been reported for any of the cell types differentially expressing calreticulin. Of focal contact components, a correlation has been shown between the abundance of calreticulin and the abundance of vinculin in both fibroblasts (16) and epithelial cells (present work). The effect of calreticulin on vinculin expression is specific as transfection of RPEfast cells with calreticulin cDNA, which causes restoration of calreticulin protein levels, also causes a corresponding increase in vinculin protein levels. Importantly, induction of calreticulin by Ca2+ deprivation of the ER due to thapsigargin treatment is also paralleled by an increase in vinculin abundance. These effects are unlikely to be due to an ER response to a general stress, as overexpression of calreticulin does not induce changes typical of those caused by the stressed ER. This was shown by Western blotting of cell extracts from cells differentially expressing calreticulin, in which we did not detect any changes in expression of SERCA2b, BiP, calnexin, protein-disulfide isomerase, ER proteins Erp72 and ERp61 (not shown). The increase in vinculin levels and the reappearance of vinculin-positive focal contacts in RPEfast cells after transfection with calreticulin cDNA is indicative of a functional role of calreticulin in the control of cell adhesion.

Is This Evidence for a Putative ER Signaling Pathway Dependent on Calreticulin?-- Calreticulin is a Ca2+-binding protein, whose ubiquity and conservation implies an important functional role. Cells overexpressing calreticulin develop prominent focal contacts, likely due to the up-regulation of vinculin. The changes in cell adhesiveness are coincident with changes in the levels of protein Tyr phosphorylation in cells differentially expressing calreticulin. It has previously been shown that the level of cellular Tyr(P) influences the ability of a cell to form focal contacts (4, 64-66). Here we show an inverse relationship between the level of total cellular Tyr(P) and the level of calreticulin expression. Possibly, an increase in the abundance of calreticulin in the ER inactivates cytoplasmic kinases or activates cytoplasmic phosphatases, causing a lowering of the levels of total cellular Tyr(P). It is also possible although that the raising of cellular Tyr(P) is downstream of the lowering of vinculin level. Recent work from Adamson and collaborators (67) shows that in vinculin-deficient mouse embryonic fibroblasts the level of cellular Tyr(P) is greatly increased over wild type fibroblasts. The opposite scenario is also plausible in which changes in cellular Tyr(P) are upstream of the changes in structure and function of cell adhesions. Crowley and Horwitz (68) showed that artificially raising the level of tyrosine phosphorylation in a cell caused the destabilization of focal contacts, including the dissociation of vinculin, and conversely, lowering this level of Tyr(P) by adding exogenous tyrosine phosphatase caused a stabilization of focal contacts. The changes in cellular Tyr(P) due to differential levels of expression of calreticulin may be indicative of alterations in the phosphorylation status of signaling proteins which affect several signaling cascades.

Adhesiveness Versus Adhesion: Stick and Grip Model

Here, our first measure of adhesion was efficacy of attachment, which showed that RPEfast cells attached with higher efficacy than RPEslow cells, and did so in a Ca2+ independent manner. Morphologically, both the IRM and EM work demonstrated that RPEfast cells have a very smooth ventral membrane, whereas RPEslow cells have an extensively undulating ventral membrane. The ECIS data2 showed that the ventral membrane of RPEfast cells was on average closer to the substratum than that of RPEslow cells.

Is a Smoother and Closer Approach of the Ventral Membrane to the Substratum Indicative of Stronger Adhesion?-- Thus far, the various adhesion assays conducted revealed that the RPEfast cells would conventionally be considered the "more adhesive" cell type. However, the results of the centrifugal assay show that RPEfast cells require less force to be detached than RPEslow cells do. This suggests that although RPEfast cells may attach faster, RPEslow cells develop a stronger physical attachment to the substratum.

How Could More "Adhesive" Cells be Less "Adherent"?-- The semiquantitative IRM analysis of calreticulin underexpressing RPEfast and calreticulin overexpressing RPEslow cells revealed they differed with respect to adhesion in one important way: RPEfast cells had almost no focal contacts and adhered to the substratum with extensive close contacts instead. In contrast, RPEslow cells adhered to the substratum with numerous focal contacts. Focal contacts link the extracellular matrix to the actin cytoskeleton via integrins which require the presence of extracellular Ca2+ for formation of the extracellular matrix-cytoskeletal connection (69, 70). Thus, if attachment in RPEfast cells occurs in a focal contact-independent way, and hence an integrin-independent way, it may be less susceptible to changes in extracellular Ca2+ than RPEslow cells. Furthermore, close contacts provide a fast means of adhesion during cell spreading and, in fact primary chick heart fibroblasts adhere only via close contacts but are still well spread and have "good adhesion characteristics" (10, 71). Similarly, epithelial cells during the first 24 h in culture adhere exclusively via close contacts (72). Norton and Izzard (73) reported a fibroblast adhesion mutant that formed close contacts, but no focal contacts, and was still able to attach and spread without impairment. While close contacts are crucial for initial attachment and cell spreading, the presence of focal contacts has been shown to be instrumental in developing adhesion of a high strength. Theoretical modeling has shown that an applied force will detach cells from a substratum via peeling of the adhesion bonds (74, 75). Formation of rigid clusters of adhesion bonds (focal contacts) will greatly increase the force required to produce detachment, relative to having the same number of independent adhesion bonds (74-76). This is no doubt why the RPEfast cells, devoid of focal contacts, attach efficiently, i.e. are very adhesive, but do not adhere strongly enough to sustain a high detachment force. In contrast, strong adhesion based on cytoskeletal linkages in RPEslow cells takes time to develop, so these cells are not very adhesive.

Do RPEfast Cells Lack Any of the Components Necessary in the Formation of Focal Contacts or Is Only the Stimulus Missing?-- The assembly of focal contacts depends on regulated interactions between (i) receptors and their extracellular ligands, (ii) receptors and the cytoskeleton, and (iii) different cytoskeletal components (4, 65). The theoretical work of Ward and Hammer (77) shows that the formation of clusters of transmembrane cytoskeleton-receptor-ligand links can be greatly affected by affinities between participating cytoskeletal proteins. They suggest that the control of assembly of focal contacts occurs mainly via "modulation of specific cytoskeletal proteins to solidify cytoskeleton-cytoskeleton connections within precursor focal contact structures." In the present case of calreticulin underexpressing RPEfast cells, both the decreased abundance of vinculin and increased level of cellular Tyr phosphorylation may account for their relative inability to stabilize the cytoskeleton associated with focal contacts. Thus, based on the present data, we cannot discern if it is the paucity of vinculin, altered interactions between focal contact components, or signaling that is affected by the differing level of calreticulin expression in RPEslow and RPEfast cells. Cells exposed to sphingosine and thapsigargin up-regulated both calreticulin and vinculin. Focal contact formation, however, was not very efficient in thapsigargin-treated cells. Sphingosine, in addition to its effects on phosphorylation of pp125FAK and paxillin (52, 53), also causes the Ca2+ release from the ER (78, 79) via inositol 1,4,5-trisphosphate-independent mechanisms (80-82). Thus it is likely that sphingosine induces calreticulin by Ca2+ deprivation from the ER in a manner analogous to thapsigargin (55). Thapsigargin, in contrast to sphingosine, is not known to induce any changes in the phosphorylation status of components of focal contacts: this may explain the relatively modest increase in focal contact number after the thapsigargin treatment. Finally, the treatment of RPEfast cells with nocodazole increased the level of vinculin over control, but did not alter the abundance of calreticulin. This suggests that vinculin expression can be induced by factors that act independently of calreticulin, indicating multiple pathways in which focal contacts can be induced.

Over 20 years ago, it was proposed by Rees et al. (83) that cell adhesion comprises "stick," referring to receptor-ligand interactions occurring at the cell surface, and "grip," involving the formation of bonds between the ligated receptors and the cytoskeleton. This intuitive proposal has been elegantly formalized by Ward and Hammer (76) into a working hypothesis that encompasses a range of cell adhesive behaviors (84-86). Here we show that cells underexpressing calreticulin and vinculin stick to the substratum with close contacts, while cells abundantly expressing these proteins grip it with focal contacts.

    ACKNOWLEDGEMENT

We are grateful to Dr. Reinhart Reithmeier for critical reading of the manuscript and Dr. E. Rodriguez-Boulan for providing RPE-J cells. We thank Drs. S. Craig and D. Critchley for vinculin DNA probes and Dr. M. Ginsberg for anti-beta 1 integrin antibodies. The gererosity of Dr. B. Chan in providing various anti-alpha -integrins is gratefully acknowledged.

    FOOTNOTES

* The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

parallel Supported by grants-in-aid from Heart and Stroke Foundation of Alberta and the Medical Research Council and is a Medical Research Council Senior Scientist and Alberta Heritage Foundation for Medical Research Medical Scientist.

** Supported by grants-in-aid from the Heart and Stroke Foundation of Ontario and the Medical Research Council. To whom correspondence should be addressed: Dept. of Anatomy & Cell Biology, University of Toronto, Medical Sciences Building, Toronto, Ontario M5S 1A8, Canada. Tel.: 416-978-8947; Fax: 416-978-3954; E-mail: m.opas{at}utoronto.ca.

2 ECIS has been used to evaluate cell motility and cell attachment and spreading (38-42, 87). Our results show, however, that ECIS data for cell-substratum separation cannot be used as an indication of the degree of focal contact formation. In some cases a decrease in measured cell-substratum separation is correlated with an increase in focal contact area, as can occur when temperature is increased (41, 42) and, in fact, it has been assumed that an increase in ECIS measured total resistance indicates an increase in focal contact formation (88). Here we show that the RPEfast cells which are almost devoid of focal contacts, have a smaller average cell-substratum separation than the RPEslow cells. The underside of RPEslow cells undulates greatly covering extremes in cell-substratum separation, hence the average cell-substratum separation, as indicated by the resistance of the cell layer, is larger that that of RPEfast cells, even though RPEslow cells have numerous focal contacts. Thus, the assumption made by Moy et al. (88) appears not valid. An increase in total resistance may reflect an increased resistance between the cells and under the cells, but an increase in the latter resistance (indicating a decrease in cell-substratum separation) may not necessarily be correlated with an increase in focal contact area.

    ABBREVIATIONS

The abbreviations used are: IRM, interference reflection microscopy; ConA, concanavalin A; ECIS, electric cell-substrate impedance sensing; ER, endoplasmic reticulum; FBS, fetal bovine serum; Tyr(P), phosphotyrosine; PBS, phosphate-buffered saline; RPE, retinal pigment epithelium; PAGE, polyacrylamide gel electrophoreses; TRITC, tetramethyl rhodamine isothiocyanate; FAK, focal adhesion kinase; PIPES, 1,4-piperazinediethanesulfonic acid.

    REFERENCES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
  1. Izzard, C. S., and Lochner, L. R. (1976) J. Cell Sci. 21, 129-159[Abstract]
  2. Curtis, A. S. G. (1964) J. Cell Biol. 20, 199-215[Abstract/Free Full Text]
  3. Burridge, K., and Chrzanowska-Wodnicka, M. (1996) Annu. Rev. Cell Dev. Biol. 12, 463-518[CrossRef][Medline] [Order article via Infotrieve]
  4. Burridge, K., Chrzanowska-Wodnicka, M., and Zhong, C. L. (1997) Trends Cell Biol. 7, 342-347[CrossRef]
  5. Heath, J. P., and Dunn, G. A. (1978) J. Cell Sci. 29, 197-212[Medline] [Order article via Infotrieve]
  6. Chen, W.-T., Hasegawa, E., Hasegawa, T., Weinstock, C., and Yamada, K. M. (1985) J. Cell Biol. 100, 1103-1114[Abstract]
  7. Jockusch, B. M., Bubeck, P., Giehl, K., Kroemker, M., Moscher, J., Rothkegel, M., Rüdiger, M., Schlüter, K., Stanke, G., and Winkler, J. (1995) Annu. Rev. Cell Dev. Biol. 11, 379-416[CrossRef][Medline] [Order article via Infotrieve]
  8. Hynes, R. O. (1992) Cell 69, 11-25[Medline] [Order article via Infotrieve]
  9. Katoh, K., Kano, Y., Masuda, M., and Fujiwara, K. (1996) Cell Struct. Funct. 21, 27-39[Medline] [Order article via Infotrieve]
  10. Couchman, J. R., and Rees, D. A. (1979) J. Cell Sci. 39, 149-165[Abstract]
  11. Chen, W.-T., and Singer, S. J. (1982) J. Cell Biol. 95, 205-222[Abstract]
  12. Bastianutto, C., Clementi, E., Codazzi, F., Podini, P., De Giorgi, F., Rizzuto, R., Meldolesi, J., and Pozzan, T. (1995) J. Cell Biol. 130, 847-855[Abstract]
  13. Liu, N., Fine, R. E., Simons, E., and Johnson, R. J. (1994) J. Biol. Chem. 269, 28635-28639[Abstract/Free Full Text]
  14. Mery, L., Mesaeli, N., Michalak, M., Opas, M., Lew, D. P., and Krause, K.-H. (1996) J. Biol. Chem. 271, 9332-9339[Abstract/Free Full Text]
  15. Fasolato, C., Pizzo, P., and Pozzan, T. (1998) Mol. Biol. Cell 9, 1513-1522[Abstract/Free Full Text]
  16. Opas, M., Szewczenko-Pawlikowski, M., Jass, G. K., Mesaeli, N., and Michalak, M. (1996) J. Cell Biol. 135, 1913-1923[Abstract]
  17. Burns, K., Duggan, B., Atkinson, E. A., Famulski, K. S., Nemer, M., Bleackley, R. C., and Michalak, M. (1994) Nature 367, 476-480[CrossRef][Medline] [Order article via Infotrieve]
  18. Dedhar, S., Rennie, P. S., Shago, M., Leung-Hagesteijn, C.-Y., Yang, H., Filmus, J., Hawley, R. G., Bruchovsky, N., Cheng, H., Matusik, R. J., and Giguère, V. (1994) Nature 367, 480-483[CrossRef][Medline] [Order article via Infotrieve]
  19. Michalak, M., Burns, K., Andrin, C., Mesaeli, N., Jass, G. H., Busaan, J. L., and Opas, M. (1996) J. Biol. Chem. 271, 29436-29445[Abstract/Free Full Text]
  20. Desai, D., Michalak, M., Singh, N. K., and Niles, R. M. (1996) J. Biol. Chem. 271, 15153-15159[Abstract/Free Full Text]
  21. Shago, M., Flock, G., Hagesteijn, C. Y. L., Woodside, M., Grinstein, S., Giguère, V., and Dedhar, S. (1997) Exp. Cell Res. 230, 50-60[CrossRef][Medline] [Order article via Infotrieve]
  22. Helenius, A., Trombetta, E. S., Hebert, D. N., and Simons, J. F. (1997) Trends Cell Biol. 7, 193-200[CrossRef]
  23. Leung-Hagesteijn, C.-Y., Milankov, K., Michalak, M., Wilkins, J., and Dedhar, S. (1994) J. Cell Sci. 107, 589-600[Abstract/Free Full Text]
  24. Michalak, M., and MacLennan, D. H. (1980) J. Biol. Chem. 255, 1327-1334[Abstract/Free Full Text]
  25. Milner, R. E., Baksh, S., Shemanko, C., Carpenter, M. R., Smillie, L., Vance, J. E., Opas, M., and Michalak, M. (1991) J. Biol. Chem. 266, 7155-7165[Abstract/Free Full Text]
  26. Opas, M., Dziak, E., Fliegel, L., and Michalak, M. (1991) J. Cell. Physiol. 149, 160-171[Medline] [Order article via Infotrieve]
  27. Fliegel, L., Burns, K., Opas, M., and Michalak, M. (1989) Biochim. Biophys. Acta 982, 1-8[Medline] [Order article via Infotrieve]
  28. Andrin, C., Pinkoski, M. J., Burns, K., Atkinson, E. A., Krahenbuhl, O., Fraser, F., Winkler, U., Tschopp, J., Opas, M., Bleackley, R. C., and Michalak, M. (1998) Biochemistry 37, 10386-10396[CrossRef][Medline] [Order article via Infotrieve]
  29. Fliegel, L., Burns, K., MacLennan, D. H., Reithmeier, R. A. F., and Michalak, M. (1989) J. Biol. Chem. 264, 21522-21528[Abstract/Free Full Text]
  30. Nabi, I. R., Mathews, A. P., Cohen-Gould, L., Gundersen, D., and Rodriguez-Boulan, E. (1993) J. Cell Sci. 104, 37-49[Abstract/Free Full Text]
  31. Marrs, J. A., Andersson-Fisone, C., Jeong, M. C., Cohen-Gould, L., Zurzolo, C., Nabi, I. R., Rodriguez-Boulan, E., and Nelson, W. J. (1995) J. Cell Biol. 129, 507-519[Abstract]
  32. Ausubel, F. M., Brent, R., Kingston, R. E., Moore, D. D., Seidman, J. C., Smith, J. A., and Struhl, K. (1995) Current Protocols in Molecular Biology, Wiley-Interscience, New York
  33. Coutu, M. D., and Craig, S. W. (1988) Proc. Natl. Acad. Sci. U. S. A. 85, 8535-8539[Abstract]
  34. Opas, M., and Kalnins, V. I. (1984) Eur. J. Cell Biol. 33, 60-65[Medline] [Order article via Infotrieve]
  35. Opas, M., and Kalnins, V. I. (1986) Invest. Ophthalmol. Vis. Sci. 27, 1622-1633[Abstract]
  36. Bereiter-Hahn, J., Fox, C. H., and Thorell, B. (1979) J. Cell Biol. 82, 767-779[Abstract]
  37. Gingell, D., and Todd, I. (1979) Biophys. J. 26, 507-526[Abstract]
  38. Giaever, I., and Keese, C. R. (1991) Proc. Natl. Acad. Sci. U. S. A. 88, 7896-7900[Abstract]
  39. Mitra, P., Keese, C. R., and Giaever, I. (1991) BioTechniques 11, 504-510[Medline] [Order article via Infotrieve]
  40. Lo, C. M., Keese, C. R., and Giaever, I. (1995) Biophys. J. 69, 2800-2807[Abstract]
  41. Lo, C. M., and Ferrier, J. (1998) Physiol. Rev. E 57, 6982-6987[CrossRef]
  42. Lo, C. M., Glogauer, M., Rossi, M., and Ferrier, J. (1998) Eur. Biophys. J. 27, 9-17[CrossRef][Medline] [Order article via Infotrieve]
  43. Hertl, W., Ramsey, W. S., and Nowlan, E. D. (1984) In Vitro 20, 796-801[Medline] [Order article via Infotrieve]
  44. Geiger, B. (1979) Cell 18, 193-205[Medline] [Order article via Infotrieve]
  45. Geiger, B. (1983) Biochim. Biophys. Acta 737, 305-341[Medline] [Order article via Infotrieve]
  46. Badley, R. A., Woods, A., Carruthers, L., and Rees, D. A. (1980) J. Cell Sci. 43, 379-390[Abstract]
  47. Danowski, B. A. (1989) J. Cell Sci. 93, 255-266[Abstract]
  48. Bershadsky, A., Chausovsky, A., Becker, E., Lyubimova, A., and Geiger, B. (1996) Curr. Biol. 6, 1279-1289[Medline] [Order article via Infotrieve]
  49. Pletjushkina, O. J., Belkin, A. M., Ivanova, O. J., Oliver, T., Vasiliev, J. M., and Jacobson, K. (1998) Cell Adhes. Commun. 5, 121-135[Medline] [Order article via Infotrieve]
  50. Enomoto, T. (1996) Cell Struct. Funct. 21, 317-326[Medline] [Order article via Infotrieve]
  51. Liu, B. P., Chrzanowska-Wodnicka, M., and Burridge, K. (1998) Cell Adhes. Commun. 5, 249-255[Medline] [Order article via Infotrieve]
  52. Seufferlein, T., and Rozengurt, E. (1994) J. Biol. Chem. 269, 27610-27617[Abstract/Free Full Text]
  53. Wang, F., Nobes, C. D., Hall, A., and Spiegel, S. (1997) Biochem. J. 324, 481-488[Medline] [Order article via Infotrieve]
  54. Thastrup, O., Cullen, P. J., Drobak, B. K., Hanley, M. R., and Dawson, A. P. (1990) Proc. Natl. Acad. Sci. U. S. A. 87, 2466-2470[Abstract]
  55. Waser, M., Mesaeli, N., Spencer, C., and Michalak, M. (1997) J. Cell Biol. 138, 547-557[Free Full Text]
  56. Daniel, J. M., and Reynolds, A. B. (1997) BioEssays 19, 883-891[Medline] [Order article via Infotrieve]
  57. Hanks, S. K., and Polte, T. R. (1997) BioEssays 19, 137-145[Medline] [Order article via Infotrieve]
  58. Virtanen, I., Ekblom, P., and Laurila, P. (1980) J. Cell Biol. 85, 429-434[Abstract]
  59. Tartakoff, A. M., and Vassalli, P. (1983) J. Cell Biol. 97, 1243-1248[Abstract]
  60. Rodríguez Fernández, J. L., Geiger, B., Salomon, D., and Ben-Ze'ev, A. (1992) Cell Motil. Cytoskeleton 22, 127-134[Medline] [Order article via Infotrieve]
  61. Coppolino, M., Leung-Hagesteijn, C., Dedhar, S., and Wilkins, J. (1995) J. Biol. Chem. 270, 23132-23138[Abstract/Free Full Text]
  62. Ezzell, R. M., Goldmann, W. H., Wang, N., Parasharama, N., and Ingber, D. E. (1997) Exp. Cell Res. 231, 14-26[CrossRef][Medline] [Order article via Infotrieve]
  63. Coppolino, M. G., Woodside, M. J., Demaurex, N., Grinstein, S., St-Arnaud, R., and Dedhar, S. (1997) Nature 386, 843-847[CrossRef][Medline] [Order article via Infotrieve]
  64. Otey, C. A. (1996) Int. Rev. Cytol. 167, 161-183[Medline] [Order article via Infotrieve]
  65. Craig, S. W., and Johnson, R. P. (1996) Curr. Opin. Cell Biol. 8, 74-85[CrossRef][Medline] [Order article via Infotrieve]
  66. Ilic, D., Damsky, C. H., and Yamamoto, T. (1997) J. Cell Sci. 110, 401-407[Abstract/Free Full Text]
  67. Xu, W. M., Baribault, H., and Adamson, E. D. (1998) Development 125, 327-337[Abstract/Free Full Text]
  68. Crowley, E., and Horwitz, A. F. (1995) J. Cell Biol. 131, 525-537[Abstract]
  69. Buck, C. A., and Horwitz, A. F. (1987) Annu. Rev. Cell Biol. 3, 179-205[CrossRef]
  70. Sjaastad, M. D., and Nelson, W. J. (1997) BioEssays 19, 47-55[Medline] [Order article via Infotrieve]
  71. Couchman, J. R., and Rees, D. A. (1979) Cell Biol. Int. Rep. 3, 431-439[Medline] [Order article via Infotrieve]
  72. Cottler-Fox, M., Sparring, K. M., Zetterberg, A., and Fox, C. H. (1979) Exp. Cell Res. 118, 414-418[Medline] [Order article via Infotrieve]
  73. Norton, E. K., and Izzard, C. S. (1982) Exp. Cell Res. 139, 463-467[Medline] [Order article via Infotrieve]
  74. Ward, M. D., Dembo, M., and Hammer, D. A. (1994) Biophys. J. 67, 2522-2534[Abstract]
  75. Ward, M. D., Dembo, M., and Hammer, D. A. (1995) Ann. Biomed. Eng. 23, 322-331[Medline] [Order article via Infotrieve]
  76. Ward, M. D., and Hammer, D. A. (1993) Biophys. J. 64, 936-959[Abstract]
  77. Ward, M. D., and Hammer, D. A. (1994) J. Math. Biol. 32, 677-704[Medline] [Order article via Infotrieve]
  78. Ghosh, T. K., Bian, J., and Gill, D. L. (1990) Science 248, 1653-1656[Medline] [Order article via Infotrieve]
  79. Zhang, H., Desai, N. N., Olivera, A., Seki, T., Brooker, G., and Spiegel, S. (1991) J. Cell Biol. 114, 155-167[Abstract]
  80. Desai, N. N., Carlson, R. O., Mattie, M. E., Olivera, A., Buckley, N. E., Seki, T., Brooker, G., and Spiegel, S. (1993) J. Cell Biol. 121, 1385-1395[Abstract]
  81. Tornquist, K., and Ekokoski, E. (1994) Biochem. J. 299, 213-218[Medline] [Order article via Infotrieve]
  82. Mattie, M., Brooker, G., and Spiegel, S. (1994) J. Biol. Chem. 269, 3181-3188[Abstract/Free Full Text]
  83. Rees, D. A., Lloyd, C. W., and Thom, D. (1977) Nature 267, 124-128
  84. Lotz, M. M., Burdsal, C. A., Erickson, H. P., and McClay, D. R. (1989) J. Cell Biol. 109, 1795-1805[Abstract]
  85. Massia, S. P., and Hubbell, J. A. (1991) J. Cell Biol. 114, 1089-1100[Abstract]
  86. Opas, M. (1995) Biochem. Cell Biol. 73, 311-316[Medline] [Order article via Infotrieve]
  87. Giaever, I., and Keese, C. R. (1993) Nature 366, 591-592[CrossRef][Medline] [Order article via Infotrieve]
  88. Moy, A. B., Van Engelenhoven, J., Bodmer, J., Kamath, J., Keese, C., Giaever, I., Shasby, S., and Shasby, D. M. (1996) J. Clin. Invest. 97, 1020-1027[Abstract/Free Full Text]


Copyright © 1999 by The American Society for Biochemistry and Molecular Biology, Inc.