Advertisement
JBC

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Originally published In Press as doi:10.1074/jbc.M402944200 on June 28, 2004

J. Biol. Chem., Vol. 279, Issue 37, 38277-38286, September 10, 2004
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
279/37/38277    most recent
M402944200v1
Right arrow Submit a Letter to Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Zwartz, G. J.
Right arrow Articles by Sklar, L. A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Zwartz, G. J.
Right arrow Articles by Sklar, L. A.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

Real-time Analysis of Very Late Antigen-4 Affinity Modulation by Shear*

Gordon J. Zwartz{ddagger}, Alexandre Chigaev{ddagger}, Denise C. Dwyer, Terry D. Foutz, Bruce S. Edwards, and Larry A. Sklar§

From the Department of Pathology and Cancer Research and Treatment Center, University of New Mexico, Albuquerque, New Mexico 87131

Received for publication, March 16, 2004 , and in revised form, June 16, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Shear promotes endothelial recruitment of leukocytes, cell activation, and transmigration. Mechanical stress on cells caused by shear can induce a rapid integrin conformational change and activation, followed by an increase in binding to the extracellular matrix. The molecular mechanism of increased avidity is unknown. We have shown previously that the affinity of the {alpha}4{beta}1 integrin, very late antigen-4 (VLA-4), measured with an LDV-containing small molecule, varies with cellular avidity, measured from cell disaggregation rates. In this study, we measured in real time affinity changes of VLA-4 in response to shear. The resulting affinity was comparable with the state mediated by receptor signaling and corresponded in time with intracellular Ca2+ responses. Ca2+ ionophores and N,N'-[1,2-ethanediyl-bis(oxy-2,1-phenylene)]bis[N-[2-[(acetyloxy)methoxy]-2-oxoethyl]]-, bis[(acetyloxy)methyl]ester demonstrate that the affinity regulation of VLA-4 in the presence of shear was related to Ca2+ signaling. Pertussis toxin treatment implicates Gi in an unknown pathway that connects shear, Ca2+ elevation, VLA-4 affinity, and cell avidity.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Leukocytes are recruited to endothelial cells in a multistep process using selectin and integrin adhesion molecules (1, 2). These molecules allow a cell to tether, roll, adhere, and transmigrate along and across an endothelial layer. Selectin and some integrin molecules and their associated ligands mediate tethering and rolling interactions. Firm adhesion is mediated by vascular ligands of the immunoglobulin superfamily such as vascular cell adhesion molecule 1 (VCAM-1)1 and their associated integrins (1, 2). The adhesive strength or avidity (3) of cells expressing integrins can be rapidly modulated by chemokines and chemoattractants, which also regulate leukocyte recruitment and migration across vascular endothelium. The rapid changes in avidity have been attributed to changes in the number of interacting molecules or valency due to molecular redistribution or clustering and to changes in the affinity of the individual receptor-ligand bonds (3–10).

Physiological shear can also regulate leukocyte traffic by stimulating mechanosensors on neutrophils, monocytes, lymphocytes, erythrocytes, and platelets (see Ref. 11 and references therein). Shear arises from bifurcating blood vessels or rapid changes in blood vessel diameters. Shear acting on leukocytes, bound to endothelial cells, produces mechanical stress on the cells or their receptors, regulating cell growth and proliferation, protein synthesis, gene expression, and blood cell recruitment (12, 13). Integrins (such as {alpha}v{beta}3, {alpha}5{beta}1, {alpha}4{beta}1, and {alpha}2{beta}1) on endothelial cells can act as mechanosensors to changes in blood flow (13, 14) and trigger an intracellular signaling pathway involving focal adhesion kinase and mitogen-activated protein kinase cascades. How shear specifically induces blood cell adhesiveness or recruitment through mechanosensors is unknown. Indirect evidence shows that increased integrin binding to the extracellular matrix occurs when shear acts on cells or their mechanosensors to induce intracellular signaling. For example, intracellular signaling leads to conformational changes and activation of {alpha}v{beta}3 on endothelial cells and {alpha}4{beta}1 and {alpha}5{beta}1 integrins on monocytic cells (15, 16). Shear acting on endothelial cells affects the GTPase Rho signaling pathway and in monocytic cells induces inositol 1,4,5-trisphosphate-sensitive Ca2+ release that affects cell adhesion avidity.

We have used an LDV-containing small molecule fluorescent probe to determine whether mechanical stress generated by shear can affect the affinity of VLA-4 by monitoring in real time the changes in VLA-4 affinity on live cells (17). We examined the contribution of intracellular signaling mechanisms to VLA-4 activation by shear. We found that VLA-4 affinity induced by shear was intermediate in affinity between the resting state and the Mn2+-activated affinity state and similar to the physiologically activated receptor state generated using "inside-out" signaling (17). We found a temporal correlation between the intracellular Ca2+ response and the higher VLA-4 affinity. We used Ca2+ ionophores (A23187 [GenBank] and ionomycin) and BAPTA-AM to show that VLA-4 affinity regulation in response to shear was related to intracellular Ca2+ signaling. Finally, we pretreated cells with pertussis toxin (PTX) to block Gi signaling) and observed that VLA-4 activation was inhibited in the presence of shear. Our data suggest that shear regulates cell adhesion avidity by changing VLA-4 affinity and involves an incompletely characterized inside-out signaling pathway.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Materials—The VLA-4-specific 4-(N'-2-methyphenyl)ureido)-phenylacetyl-L-leucyl-L-aspartyl-L-valyl-L-alanyl-L-lysine (LDV-containing small molecule) and its FITC-labeled analog were synthesized at Commonwealth Biotechnologies, Inc. (Richmond, VA). Binding and dissociation of the LDV-FITC probe were described previously (17, 18). Intracellular Ca2+ was chelated using 5,5'-dimethyl-BAPTA-AM (acetoxymethyl ester) (Molecular Probes, Inc., Eugene, OR) according to the manufacturer's instructions. A23187 [GenBank] Ca2+ ionophore was purchased from Sigma and used at 1 µM concentration. Ionomycin was purchased from Calbiochem and used at 1 µM concentration. Fura Red AM and Fluo-4 AM were purchased from Molecular Probes. FITC-conjugated monoclonal antibody, 44H6, against CD49d was purchased from Serotec (Raleigh, NC). All other reagents were from Sigma.

Cell Lines and Transfectant Construct—Human monoblastoid U937 cells were purchased from ATCC (Manassas, VA). Cells were grown in RPMI 1640 (Invitrogen) supplemented with 10% heat-inactivated fetal bovine serum, 100 units/ml penicillin, 100 µg/ml streptomycin, 10 mM HEPES, pH 7.4, 100 µg/ml ciprofloxacin, 2 mM L-glutamine, at 37 °Cin a humidified atmosphere of 5% CO2 and 95% air. Site-directed mutants of formyl peptide receptor in the human monoblastoid line U937 constitutively expressing human VLA-4 integrin were prepared as described (19). High expressors were selected using the MoFlo Flow Cytometer (Cytomation, Inc., Fort Collins, CO). VLA-4 expression was measured with FITC-44H6 and quantified by comparison with a standard curve generated with Quantum Simply Cellular microspheres (Flow Cytometry Standards, San Juan, Puerto Rico) stained in parallel with the same monoclonal antibody. This produces an estimate of the total monoclonal antibody-binding sites/cell. Typically, we find 40,000–60,000 VLA-4 sites/U937 cell.

LDV-FITC Probe—The VLA-4 probe (20–22) was initially optimized from the ILDV binding sequence of the alternatively spliced connecting segment 1 of fibronectin. This sequence is homologous and isosteric with the QIDS peptide found in the VCAM-1-binding site (23). The peptide sequence (Leu-Asp-Val-Pro-Ala-Ala-Lys-FITC) of the probe was based on structure-activity relationships of a potent VLA-4 binding inhibitor (compound 13 in Ref. 22). The specificity of the molecule for the VCAM-1/VLA-4 interaction was examined previously in cell adhesion and ligand binding assays (17). The binding characterization showed that molecular dissociation rates of the LDV-FITC probe from VLA-4 on U937 were homogenous (i.e. single exponential) regardless of the cation present (18).

Cell Preparation—U937 cells (10 x 106 cells/ml) for shear experiments were loaded with 6 µM Fura Red or 200 nM Fluo-4, for 30–60 min at 37 °C and gently mixed every 10 min. Then the cells were washed with complete RPMI and resuspended in phenol red-deficient RPMI (supplemented with 0.1% human serum albumin (Bayer Corp., Elkhart, IN) and 1.5 mM Ca2+). Cells were kept on ice after staining and washing. Typically, 5 min prior to each experiment, 4 nM LDV-FITC probe was added as a ligand to 1 x 106 cells/ml, and the sample was incubated in a 37 °C water bath. Cells were illuminated with the 488-nm argon laser from a Becton-Dickinson FACScan flow cytometer (BD Immunocytometry Systems, San Jose, CA). Emission fluorescence was detected using a 585-nm band pass filter for Fura Red (FL2) and 530-nm band pass filter for Fluo-4 (FL1). Fura Red fluorescence decreased when the indicator bound to free Ca2+. Changes in the affinity state of VLA-4 were monitored using the LDV-FITC probe. The probe was added 5 min prior to each experiment usually at 4 nM and incubated in a 37 °C water bath. Detailed analysis of real time binding and dissociation of the LDV-FITC probe was previously described in Refs. 17 and 18. In several experiments (where the extracellular Ca2+ concentration varied), Hepes buffer (110 mM NaCl, 10 mM KCl, 10 mM glucose, and 30 mM HEPES, pH 7.4) supplemented with 0.1% human serum albumin was used. Cell density was determined using a Z2-Coulter counter (Coulter Corp., Miami, FL).

Intracellular Calcium Calibration—Molecular Probes calcium calibration kit 1 was used to generate a series of free calcium buffers that were used to obtain an intracellular cellular calcium calibration curve (Fig. 1). The kits contain two 50-ml solutions, one solution containing 10 mM K2EGTA and the other 10 mM CaEGTA. Both solutions contained 100 mM KCl, 30 mM MOPS, pH 7.2. Intermediate free calcium concentrations between 0 and 39 µM were obtained by cross-diluting the two buffers. Before adding U937 to each of the prepared buffers, the cells were stained with the intracellular calcium indicator Fura Red. Prior to each experiment, 1 x 106 U937 cells were added to 1 ml of a specific free calcium buffer. Then the solution was incubated for 5 min in a 37 °C water bath. A base line was established during the first 2 min of sampling with a FACScan to measure the resting state of the cells. Then 10 ng/ml of a calcium ionophore (A23187 [GenBank] ) was added and mixed gently, and sampling was resumed. Measurements of intracellular calcium were obtained when the Fura Red signal equilibrated. Fig. 1 shows that changes in the mean channel fluorescence (MCF) corresponded to logarithmic changes in the intracellular calcium levels. The intracellular Ca2+ calibration curve depended on Fura Red staining efficiency, viability of U937 cells, sensitivity of cells to external activation, and flow cytometer voltage and gain settings. The Fura Red MCF values for cellular resting states between 550 and 650 correspond to intracellular calcium concentrations between 100 and 10 nM. MCF values of ~400 after stimulation indicate an intracellular calcium concentration of ≥1000 nM.



View larger version (11K):
[in this window]
[in a new window]
  		FIG. 1.
Intracellular Ca2+ calibration. 10 ng/ml calcium ionophore (A23187 [GenBank] ) was added to the cell solutions containing U937 cells stained with Fura Red (a calcium indicator) in their respective calcium buffers. The intracellular calcium concentration (x axis) and the mean channel fluorescence (y axis) were obtained at each equilibrium point. A solid line is used to guide the eye.

 
Creating Fluid Shear—Fluid shear was initially generated using a Fischer Scientific minivortexer (Fischer Scientific, Hampton, NH) set to 3200 rpm. The shear rate was estimated to be ~200–12,000 s–1, comparing the vortexed fluid motion inside a 12 (outer diameter) x 75-mm tube with the fluid motion inside a Couette viscometer. The maximum (Smax) and minimum (Smin) wall shear rate for a given rotational velocity was approximated (24) as follows,

(Eq. 1)

(Eq. 2)
where RI (ranging from ~0.535 to 0.25 cm) and RO (0.55 cm) represent radii of the inner fluid and outer fluid surfaces, and {Omega} is the angular speed of the inner cylinder.

Before being subjected to shear, U937 cells were incubated for 5 min ina37 °C water bath. Each sample was gently mixed to resuspend cells, and a tube was attached to a flow cytometer. Data were acquired for 1–3 min to establish a base line for resting cells, and then each sample was removed from the flow cytometer to be exposed to shear for 5–30 s using a minivortexer. Samples were reattached to the flow cytometer, and data sampling was resumed.

A minivortexer generates turbulent fluid flow. To reduce this variability, we used a computer-driven syringe (Alitea, Bellevue, WA) to push samples through a 50-cm-long 0.03-inch (762-µm) inner diameter fluorinated ethylene propylene (FEP) tubing (Upchurch Scientific, Oak Harbor, WA) at flow rates of 33, 100, 200, and 400 µl/s. The capillary wall shear rates (Swall) were calculated using the following,

(Eq. 3)
where Q represents the flow rate, and r is the tube radius (the corresponding wall shear rates were calculated to be 750, 2300, 4600, and 9200 s–1, respectively), which was the maximal shear rate cells would experience (24). For deformable particulates, such as cells, there is a net radial hydrodynamic force moving particulates toward the flow axis (25), even at a low Reynolds number. Thus, not all cells flow along a capillary wall (maximal shear rate) or capillary axis (zero shear rate). Consequently, there was a range of shear experienced by flowing cells, and the maximal shear rate does not represent the shear experienced by all cells. For a simple approximation, we have assumed an average shear rate (between the maximum and minimum shear rate experienced by cells) for the four flow rates to be 375, 1150, 2300, and 4600 s–1.

Fig. 2 shows a schematic of capillary shear. Typically, a 1-ml sample containing 1 x 106 U937 cells was aspirated into a 1-ml computer-driven syringe. After the sample was loaded into a syringe, the sample was pushed into a FEP tube at one of the four flow rates to generate shear. When that cycle was completed, the sample was aspirated into the same syringe. This cycle was repeated five times. After the fifth cycle, a computer-operated solenoid valve (NResearch, Caldwell, NJ), used to separate the shear FEP line from an FEP line leading to a FACScan, was switched to allow samples to be pushed toward a FAC-Scan at 1 µl/s.



View larger version (13K):
[in this window]
[in a new window]
  		FIG. 2.
A schematic of capillary shear. A computer-driven syringe (Syringe) pushes and aspirates cell samples through a three-way computer-controlled solenoid valve (Valve; 0.04-inch inner diameter) and into 0.03-inch inner diameter FEP tubing (Capillary).

 
Flow Cytometry and Data Analysis—Flow cytometric analysis was done on a Becton-Dickinson FACScan flow cytometer (BD Biosciences). Data acquisition was performed using CellQuest (BD Biosciences). Data were analyzed offline using the Windows Multiple Document Interface Flow Cytometry Interface (Scripps, La Jolla, CA). Time and fluorescence information were extracted from the data using FacsQuery software, developed by Bruce Edwards. Peak analysis and data fitting were done using PeakFit version 4.11 (Systat, Point Richmond, CA) and GraphPad Prism 4 (GraphPad, San Diego, CA), respectively.

A ligand dissociation analysis would not readily distinguish heterogeneity in the affinity of resting and activated receptors on a given cell as compared with heterogeneity in the distribution of receptors on activated and resting cells. However, the distribution of the amount of ligand bound would distinguish cells that had activated receptors from cells that did not. Thus, we have analyzed cell distributions before and after activation as shown in Fig. 3, regions A and B. The same principles were used for the analysis of ligand binding and Ca2+ response. For this analysis, a Gaussian curve was fitted to the mean channel fluorescence distribution obtained from region A, the resting state of cells. Region B was fitted with two Gaussian curves. One fit used the peak centroid and the full-width half maximum of Region A. The peak height was allowed to vary. This component represents resting cells. A second Gaussian curve was fitted to the remainder of the distribution in which the centroid and peak height were allowed to vary but full-width half-maximum was fixed using the fit values obtained from Region A. The second curve represented activated cells. A simultaneous two-Gaussian fit to the mean channel fluorescence distribution obtained from Region B was done. The ratio of the total events under the two histograms was taken to estimate the fraction of cells activated under shear ((resting–activated)/activated).



View larger version (35K):
[in this window]
[in a new window]
  		FIG. 3.
Effect of capillary shear on the intracellular Ca2+ and LDV-FITC probe binding to VLA-4 at different shear rates. A, intracellular calcium signaling in U937 cells was examined at 2300 s–1 (200 µl/s; open circles), 1150 s–1 (100 µl/s; open squares), and 375 s–1 (33 µl/s; open triangles). Data were adjusted to the same base line and start time for flow into the FACScan. Resting and activated cell populations were selected from two time-gated regions denoted as Region B and A, respectively (see "Results") (analysis of resting and activated cells is shown in C). B, same as A except LDV-FITC binding to VLA-4 data was examined (analysis of resting and activated cells is shown in D). C, normalized histogram of MCF calcium signaling data at 200 (2300 s–1), 100 (1150 s–1), and 33 (375 s–1) µl/s obtained from A. The histograms were generated by selecting Region B in A (85–95 s; shear-activated) and Region A (12–22 s; resting state; in A). The resting state histogram was obtained by averaging histograms from Region A of 200 (2300 s–1), 100 (1150 s–1), and 33 (375 s–1) µl/s data. The remaining histograms were obtained from Region B for the three flow conditions. All data were normalized to the largest value in the mean channel fluorescence distribution. D, same as C, except histograms of LDV-FITC probe binding to VLA-4 data were examined. E, percentage of U937 cells that were activated under shear conditions. A hyperbolic equation fit is shown for the LDV-FITC probe (filled circles) and Ca2+ response (open circles). The percentage of activated cells was calculated using data presented in A and B.

 

   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Fluid Forces Increase the Affinity of the {alpha}4{beta}1 Integrin in Real Time—Studies were conducted in a turbulent fluid flow environment using a Fischer Scientific minivortexer. To determine whether shear can affect the affinity of VLA-4, we used the LDV-FITC probe (17). Prior to applying shear, U937 cells (1 x 106 cells/ml) were equilibrated with 4 nM probe. The concentration chosen for the experiments was below the dissociation constant (Kd of ~12 nM) for probe binding to resting VLA-4 and above the Kd for the physiologically activated receptor (Kd of ~1–2nM) (17). Therefore, the transition from the low affinity to the high affinity receptor leads to an increased binding of the probe (from ~25 to ~75% of receptor occupancy). Fig. 4 shows the rapid and transient increase in probe binding to sheared cells. The binding of the probe was detected after data acquisition was re-established, indicating that seconds were needed to induce cell activation. The binding reached a peak at 40–60 s after vortexing and decreased to the basal level after another 40–60 s. For comparison, we show the conformational state induced by 1 mM Mn2+ (Kd ~0.5 nM and occupancy ~90%) in the buffer containing 1 mM Mn2+ and 1 mM Ca2+ (18). Fig. 4 shows that Mn2+ increased probe binding above the level detected for shear.



View larger version (17K):
[in this window]
[in a new window]
  		FIG. 4.
Binding and dissociation kinetics of the LDV-FITC probe on U937 cells in response to cell vortexing and cell activation by Mn2+. Samples were analyzed for 30–120 s to establish a base line and then vortexed for 5 s at 3200 rpm. 200 s later, 1 mM Mn2+ was added to induce a high VLA-4 affinity. To induce dissociation, unlabeled LDV probe (2 µM) was added. The MCF value at the end of the dissociation curve corresponds to the nonspecific binding of the LDV-FITC probe plus the cell autofluorescence.

 
Affinity Changes in a Controlled Fluid Force Environment— The range of shear rates was narrowed with computer-driven syringes and capillary tubes (see "Experimental Procedures"). Fig. 3, A and B, shows the kinetics of intracellular Ca2+ (determined using Fluo-4) and VLA-4 probe binding in response to the different levels of shear rates. The percentage of activated cells was calculated from Regions A and B in Fig. 3, A and B, as described under "Experimental Procedures." The resting state and shear histograms were normalized to the largest value in each of their distributions. Fig. 3, C and D, show how shear affected the number of activated cells. Those results are quantified in Fig. 3E, where the fraction of activated cells versus shear stress, fit to a hyperbolic equation, were comparable for both the LDV-FITC probe and intracellular Ca2+ responses.

Simultaneous Observation of Integrin Activation and Intracellular Ca2+ Elevation in Response to Fluid Forces—To follow VLA-4 affinity changes simultaneously with intracellular Ca2+ responses, the cells were stained with both Fura Red and the LDV-FITC probe. In several experiments, we used Fluo-4 to detect intracellular Ca2+ and VLA-4 activity in parallel. Fig. 5A shows Ca2+ and LDV-FITC binding responses after U937 cells were vortexed at 3200 rpm for 5, 15, and 30 s. The Fura Red fluorescent signal decreased as the intracellular Ca2+ concentration increased (the Fura Red axis in Fig. 5A is inverted). A transient and dose-dependent increase in intracellular Ca2+ was accompanied by an increase in the binding of the LDV-FITC probe. The kinetics of probe binding was similar, but the amplitude of signal was dependent on the duration of shear, reflecting differences in the number of activated cells (see "Affinity Changes in a Controlled Fluid Force Environment" and Fig. 3).



View larger version (22K):
[in this window]
[in a new window]
  		FIG. 5.
Correlation between LDV-FITC binding and Ca2+ response after using a vortexer and capillary flow to shear cells. A, simultaneous response of VLA-4 activation (filled circles) and intracellular Ca2+ signaling (open circles) in U937 cells that were subjected to 5, 15, and 30 s of shear using a vortexer at 3200 rpm. The right axis (Fura Red MCF) was inverted. A decrease in Fura Red fluorescent intensity corresponds to an increase in intracellular calcium. B, correlation between LDV-FITC probe binding to VLA-4 and Ca2+ response after shearing cells in a FEP capillary tube. Simultaneous response of VLA-4 activation (filled circles) and intracellular Ca2+ signaling (open circles) in U937 cells to shear generated by flowing 400 µl/s (4600 s–1) through a 50-cm length 0.03-inch (762-µm) inner diameter FEP tube. The right axis (Fura Red MCF) was inverted.

 
Fig. 5B shows a representative fluid flow experiment using a computer-driven syringe to produce a maximum wall shear rate of 9200 s–1 (average shear rate of 4600 s–1; see "Creating Fluid Forces"). Resting cells were delivered to the flow cytometer at 1 µl/s for 1–2 min to obtain a base line. Then cells were sheared for ~30 s and delivered to the flow cytometer at 1 µl/s. Both the LDV-FITC probe binding and the intracellular Ca2+ signal increased and returned to their resting state at similar rates. The signal decay kinetics was significantly longer after capillary shear than for vortexing (compare Figs. 5B and 3, A and B, with Figs. 5A and 4). The kinetics of intracellular Ca2+ signaling as well as binding and dissociation of the LDV-FITC probe, observed simultaneously, vary in parallel in response to vortexing or capillary fluid flow.

Intracellular Ca2+ and Integrin Affinity Changes—To show the effect of intracellular Ca2+ on VLA-4 affinity, we activated cells through their G-protein-coupled receptors (GPCR), added Ca2+ ionophores (ionomycin and A23187 [GenBank] ), and chelated intracellular Ca2+ with BAPTA. It is known that VLA-4 can be activated through formyl peptide, CXCR2, CXCR4, and CCR3 receptors (17). Here, we took advantage of nucleotide receptors constitutively expressed on U937 cells (P2Y2 and P2Y6) (26–28) that bind ATP to mediate a rapid and transient increase in intracellular Ca2+ (28). Fig. 6 shows that the addition of 1 µM ATP results in rapid increases in the Ca2+ signal with slower LDV-FITC probe binding of amplitude similar to 30 s of vortexing. The binding of the LDV-FITC probe to the cells was limited by the rate of probe binding (kon ~3–5 x 106 M–1 s–1) and was somewhat slower than the actual VLA-4 activation rate (for comparison, see the probe binding kinetics in response to Mn2+) (Fig. 4) (17, 18).



View larger version (20K):
[in this window]
[in a new window]
  		FIG. 6.
Simultaneous response of VLA-4 activation and intracellular Ca2+ signaling in U937 cells to shear and the addition of 1 µM ATP. U937 cell samples were vortexed at 3200 rpm for 30 s, and 1 µM ATP was added later. The VLA-4 activation (filled circles) and intracellular Ca2+ signaling (open circles) were simultaneously observed. The right axis (Fura Red MCF) was inverted.

 
The dissociation of the LDV-FITC probe followed the slow decrease in the intracellular Ca2+ measured using Fura Red. This slow decay (~50 s) reflected the kinetics of restoration of VLA-4 basal activity and was slower than probe dissociation from the resting state ~0.06 s–1 (half-life of ~11 s) (17). Thus, the kinetics of VLA-4 activation on U937 cells coincides with the kinetics of intracellular Ca2+ signaling when the cell was activated through GPCR. The data were consistent with a resting Ca2+ concentration between 10 and 100 nM with elevation to ~1000 nM following activation.

We used the Ca2+ ionophore ionomycin to increase the intracellular Ca2+ concentration. Ionomycin acts as a mobile ion carrier across membranes and was used as a Ca2+-mobilizing agent (29). After establishing a sample base line for 1 min, ionophores (1 µM ionomycin in Fig. 7A and 10 µg/ml A23187 [GenBank] in Fig. 7, B and C) were added. Cell activation was prevented during mixing by gently inverting the sample. Fig. 7A shows that ionomycin activated VLA-4 in the presence of 1 and 10 mM extracellular Ca2+, and the time course of the Ca2+ elevation was similar to the time course of VLA-4 activation. An increase in the extracellular Ca2+ concentration alone did not change the total binding of the LDV-FITC probe. Since both intracellular Ca2+ conditions led to similar total probe binding, it was likely that the two conditions had the same affinity state. However, the decay phase of the integrin activation was ~3 times longer in 10 mM Ca2+, suggesting that VLA-4 activation was strongly intracellular Ca2+-dependent.



View larger version (20K):
[in this window]
[in a new window]
  		FIG. 7.
VLA-4 activation and intracellular Ca2+ signaling in U937 cells treated with calcium ionophores (ionomycin and A23187 [GenBank] ) and BAPTA. A, effect of the extracellular buffer Ca2+ concentrations on VLA-4 activation in U937 cells. The inset shows the Ca2+ response (Fluo-4) to the addition of 1 µM ionomycin (solid line) and to a control for 1 mM extracellular Ca2+. The arrows indicate the time ionomycin was added. B, LDV-FITC response for the U937 cells incubated without (filled circles) or with (open squares) 100 µM BAPTA (incubated 30 min prior to each experiment at 37 °C). C, Ca2+ response (Fura Red) for cells that were incubated without (filled circles) or with (open squares) 100 µM BAPTA.

 
Intracellular Ca2+ was chelated by incubating cells with BAPTA. Then A23187 [GenBank] was added to elevate intracellular free Ca2+ (Fig. 7, B and C) and detected as a decrease in Fura Red fluorescence corresponding to an alteration from resting to elevated (~1000 nM) Ca2+ levels. The binding of the LDV-FITC probe increased at the same time (Fig. 7B). Buffering intracellular Ca2+ with BAPTA allowed A23187 [GenBank] to induce a slow increase in the intracellular Ca2+ and LDV-FITC probe binding. Thus, the amount of the BAPTA (100 µM) loaded inside the cells was nearly sufficient to completely buffer Ca2+ influx. The slow increase in the binding of the LDV-FITC probe coincides with a slow increase in the intracellular free Ca2+.

Effect of Fluid Forces on the LDV-FITC Probe Dissociation Rate—We measured LDV-FITC dissociation rates of vortexed cells to characterize VLA-4 affinity under conditions where the duration of VLA-4 activation corresponds to the duration of the intracellular Ca2+ response (~100 s). Cells were incubated in 10 mM Ca2+, where the calcium signal lasts long enough to measure the LDV-FITC probe dissociation rate under shear (see Fig. 7A). The results are summarized in Table I and compared with a range of values found for other modes of VLA-4 activation. We found that the dissociation behavior after 10 s of vortexing required two exponential curves (fast and slow components) to fit the data. The fraction of the sites that appeared to remain in the resting state (fast component) was 24%, whereas the remaining sites exhibited a dissociation rate 4 times slower (activated state). The rate was comparable with the physiological GPCR activation pathway or divalent cation conditions (10 mM Ca2+ and 1 mM Mn2+). Intracellular pathways activated through extracellular stimuli (N-formyl-Met-Leu-Phe-Phe, interleukin-5, or IgE) all lead to VLA-4 of a similar affinity (17) and presumably in an extended conformational state of higher avidity. Our data suggest that intracellular signaling also occurs when cells are subjected to shear. Consequently, VLA-4 is activated to a similar affinity state as those generated from physiological stimuli.


View this table:
[in this window]
[in a new window]
  		TABLE I
LDV-FITC probe dissociation rates for U937 cells treated with different concentrations of divalent cation, N-formyl-Met-Leu-Phe-Phe, or exposed to shear

Experiments were performed with U937 cells expressing a nondesensitizing/noninternalizing formyl peptide receptor mutant. 106 cells/ml were equilibrated for 5 min in a 37 °C water bath with 12 or 16 nM LDV-FITC probe in HEPES buffer (supplemented with 0.1% human serum albumin and 10 mM Ca2+ + 1 mM Mg2+). After obtaining a base line, the sample was removed from the flow cytometer and vortexed or not for 10 s, and the LDV-FITC probe was added. Data were averaged for two experiments. The numbers in parenthesis represent the fraction of slow (76%) and fast (24%) dissociation in response to shear. Fractions were obtained from fitting dissociation data to a two-exponential fit, with the fast rate fixed to the unvortexed resting state.

 
Chelation of Intracellular Ca2+ Prevents Integrin Affinity Changes in Response to Fluid Forces— Fig. 8 shows the simultaneous LDV-FITC probe and intracellular Ca2+ response to shear in cells incubated with and without BAPTA. Cells that were treated with BAPTA do not respond to shear, whereas untreated cells do. Our results indicate that VLA-4 activation in response to shear was downstream of Ca2+ signaling and that an increase in intracellular Ca2+ was associated with activation of VLA-4.



View larger version (18K):
[in this window]
[in a new window]
  		FIG. 8.
Simultaneous response of VLA-4 activation and intracellular Ca2+ signaling in U937 cells that were treated with and without BAPTA (chelates intracellular Ca2+) and subjected to 10 s of shear using a vortexer at 3200 rpm. A, LDV-FITC response for cells that were incubated without (filled circles) and with (open squares) 100 µM BAPTA and vortexed for 10 s at 3200 rpm. B, Ca2+ response (Fura Red) for cells that were incubated without (filled circles) and with (open squares) 100 µM BAPTA.

 
Pertussis Toxin Effect on Ca2+ Signaling and Integrin Affinity in Response to Fluid Force—Heterotrimeric G-proteins are part of a pathway that activates integrins (30). To determine whether heterotrimeric G-proteins were involved in the VLA-4 response to shear, U937 cells were treated with PTX. After establishing a base line of LDV-FITC probe binding, the sample was vortexed for 10 s, and sampling resumed (Fig. 9A). Treatment of the cells with PTX nearly abrogated the activation of VLA-4 by shear, suggesting that G{alpha}i-related signaling can be an intermediate step in a mechanosensing pathway for VLA-4 activation. To test this hypothesis, we activated the same PTX-treated cells using P2Y receptors, constitutively expressed on U937 cells (26, 27). These receptors are coupled to the G{alpha}q subunit/phospholipase C{beta}2 pathway (31–33), which is PTX-resistant (34). To promote Ca2+ signaling and VLA-4 activation, 1 µM ATP was applied to PTX-treated and -untreated U937 cells. Whereas VLA-4 activation was reduced (Fig. 9B), the intracellular Ca2+ response was retained. Thus, a functional G{alpha}i subunit was required for VLA-4 activation in response to shear but was not required for the intracellular Ca2+ response.



View larger version (20K):
[in this window]
[in a new window]
  		FIG. 9.
VLA-4 activation in U937 cells treated with (filled circles) and without (open squares) 10 ng/ml PTX. Cells were activated by vortexing for 10 s at 3200 rpm (A) or by adding 1 µM ATP without vortexing (B). Cells were incubated at 37 °C with 10 ng/ml PTX 24 h prior to the experiments. 5 min prior to each experiment, 4 nM LDV-FITC were added, and the sample was incubated in a water bath at 37 °C.

 
To determine whether PTX-treated cells lose viability as represented by their capacity to respond through the G{alpha}q pathway, we examined the Ca2+ dose-response curve for ATP (Fig. 10). A quantitative analysis was obtained by measuring the peak height of the Ca2+ response (measured with respect to a base line defined to be the time course before the addition of ATP) after the addition of ATP. The time courses of the ATP dose curve for cells treated with and without PTX were the same. Thus, the data indicate that cells treated with PTX were not adversely affected when compared with untreated cells.



View larger version (11K):
[in this window]
[in a new window]
  		FIG. 10.
VLA-4 activation on U937 cells stimulated with different concentrations of ATP and treated with (filled squares) and without (open squares) 10 ng/ml PTX 24 h prior to each experiment and then stained with Fura Red. Intracellular Ca2+ peak heights (y axis) were measured by subtracting the MCF Fura Red base line from the MCF Fura Red peak.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
Fluid Forces, Intracellular Ca2+, and VLA-4 Affinity—We have previously detected the real time regulation of VLA-4 affinity by divalent cations, physiological signaling, and reducing agents. Here we have shown that VLA-4 affinity was elevated in the presence of shear and that the effect was rapid and transient (Figs. 3, 4, 5, 6, 8, and 9). A significant fraction of the cells, correlating with the receptors on them, responded to shear. The affinity of VLA-4 produced by this pathway was indistinguishable from the affinity produced by GPCR signaling.

The kinetics of intracellular Ca2+ signaling also corresponded to the time course of LDV-FITC binding to VLA-4 (Figs. 5 and 6) in the presence of shear. It was conceivable that the shorter vortex duration-induced responses (Fig. 4) as compared with the response to capillary fluid flow was due to shear produced during delivery through 0.03-inch internal diameter tubing that may preserve cells in an activated state for a longer period of time. In the absence of shear, Ca2+ ionophores (ionomycin and A23187 [GenBank] ) regulated VLA-4 affinity. Moreover, increased intracellular Ca2+ was always associated in time with increased LDV-FITC probe binding. The relevance of intracellular Ca2+ response in the presence of shear was further demonstrated with BAPTA-AM, which abolished the VLA-4 response to shear (Fig. 8).

Response Pathways for VLA-4 Activation—GPCR stimulation affects cell adhesive avidity through a G{alpha}i dependent process (10). Since VLA-4 activation can involve G{alpha}i pathways (35–38) and shear activates VLA-4 (Figs. 4, 5, 6) (39–42), we examined the role of G{alpha}i response to shear (Fig. 9A) by pretreating cells with PTX. Because VLA-4 activation was associated with intracellular Ca2+ signaling, we used ATP to initiate a Ca2+ response for cells treated with PTX. Those cells were activated through P2Y receptors (Fig. 11 (V)), which were coupled to G{alpha}q (PTX-resistant (31–34)). Based on experiments with PTX, we observed that a functional G{alpha}i subunit was required for VLA-4 activation by shear but not for ATP (Fig. 10) or intracellular Ca2+ signaling induced by shear (data not shown).



View larger version (41K):
[in this window]
[in a new window]
  		FIG. 11.
Possible mechanisms of VLA-4 activation by fluid flow and other mechanical factors. The bent and extended conformation of VLA-4 is shown; a more extended conformation has higher affinity for ligand (76). I, mechanical extension of VLA-4 can be accomplished by pulling a counterstructure ligand (VCAM-1). II, VLA-4 is extended directly by the flow of fluid. III, an unknown "shear sensor" transducing a signal through the G{alpha}i subunit and/or generating intracellular free Ca2+ results in inside-out signaling and VLA-4 conformational change. IV, signaling leads to the activation of other integrin molecules ("inside-out" signaling). V, elevation of intracellular free Ca2+ and VLA-4 activation through the G{alpha}q pathway. Crossed out arrows, integrin activation via the G{alpha}i pathway was blocked using PTX.

 
Integrins are one of four classes of mechanosensors (43, 44) that include ion channels (45), G-protein receptors (46), and tyrosine kinase receptors (47). Each can be associated with intracellular Ca2+ signaling pathways (48–50). Connections among the classes are illustrated by Gi and G12/G13 signaling pathways that are sufficient to activate {alpha}IIb{beta}3 receptors on platelets (30). Whereas G{alpha}q-mediated signaling is not essential for {alpha}IIb{beta}3 activation but is for Ca2+ mobilization (Fig. 11 (V)), the overall mechanism connecting G protein receptors to integrin activation in platelets is unknown.

Fluid flow generated by a vortexer can affect suspended cells in several ways. Turbulent fluid motion produced stress on a cell membrane as a result of differential fluid velocities that can activate mechanosensors. In principle, fluid vortex motion can cause cells to collide in a nonbinding manner and activate receptors or the cell membrane. Alternatively, colliding U937 cells, potentially forming homotypic aggregates (doublets or triplets) through engagement of integrins and their ligands, would be subject to mechanical stress that would pull the aggregates apart and could initiate a cell signaling sequence and/or molecular extension. Cellular aggregates between VLA-4 and a U937 cellular ligand would be inhibited by the presence of LDV peptides binding specifically to VLA-4 (17). Whereas U937 homotypic aggregation involves {beta}2 integrin (51, 52), our previous data (53) have shown that U937 at 3 x 106 cells/ml in a shear environment exhibited no homotypic aggregates even in the absence of anti-{beta}2 antibodies. We have examined whether blocking CD18 binding using anti-bodies (TS1/18; Endogen, Woburn, MA) to block {beta}2 integrin-dependent adhesion would affect intracellular Ca2+ signaling and VLA-4. No signal reduction was observed (data not shown). Thus, formation of cellular aggregates and engagement of integrins was unlikely to account for significant outside-in signaling in our study.

A schematic diagram of potential mechanisms that may induce a higher integrin affinity state in the presence of shear is shown in Fig. 11. An integrin can be stretched under force by its counterstructure (Fig. 11 (I)) or directly by fluid flow (Fig. 11 (II)) and may increase its bond adhesion strength in a catch bond mechanism (54). The latter remains a viable option, since integrins are known to be flexible (55), and shear may lead to an extension similar to the extended chain conformation observed for von Willebrand factor (56). It is worth noting that the integrin binding partners talin and paxillin that regulate cell adhesion, migration, and integrin conformation (57–63) could provide a means of mechanotransduction. That signaling has been documented with a magnetic drag force (64) to extend integrin molecules, generating an intracellular calcium response, gene transcription (65) and tyrosine phosphorylation (66–68).

Another mechanism could involve an outside-in signaling pathway and a mechanoreceptor (Fig. 11 (III)), such as an ion channel, tyrosine kinase, or G-protein-coupled receptors. Two lines of research support the existence of integrin activation through shear signaling (Fig. 11 (II and III)). First, shear rapidly stimulated {alpha}v{beta}3 via a small GTPase Rho signaling pathway (16) and caused an increase in the avidity of {alpha}v{beta}3 and {alpha}5{beta}1 integrin bearing cells to the extracellular matrix (69). Further, shear promoted lymphocyte migration across vascular endothelium in an {alpha}4{beta}1- and {alpha}L{beta}2-dependent manner, and shear-induced signal was coupled to G{alpha}i (70). The participation of intracellular Ca2+ pathways in integrin activation (Fig. 11 (IV)) was shown for {beta}1 integrin activation (71), for intracellular Ca2+ elevation induced by shear (72), and through identification of a novel calcium and integrin binding in {beta}3 integrin activation (73).

Catch Bond: A Cellular Braking Mechanism—Our results were consistent with shear-induced mechanotransduction resulting in intracellular Ca2+ signaling and VLA-4 activation. The new VLA-4 affinity state observed under fluid flow was the same one induced by GPCR signaling, which was shown previously to increase the length of the VLA-4 molecule, to decrease the cellular avidity, and to decrease the ligand dissociation rate (17, 18). These VLA-4 structural and functional changes appear to parallel the global conformational rearrangement of the extracellular domains induced by ligands and divalent cation (74) and the switchblade model for the {alpha}v{beta}3 integrin based on electron microscopy, NMR, and epitope exposure data (75). Using fluorescence resonance energy transfer and the LDV-FITC probe (21), we found a striking correlation between the degree of VLA-4 extension and its affinity (76). The prediction that force could increase adhesion bond strengths, catch bond (54), was verified by atomic force microscopy of P-selectin binding to P-selectin glycoprotein ligand-1 (77). We hypothesize that extension of an integrin could also be part of a braking system in leukocyte rolling (78) and that shear could play a role in the pathways shown in Fig. 11 (I, II, and III). We have obtained direct evidence for the first of these.2


   FOOTNOTES
 
* This work was supported by National Institutes of Health Grants RR14175/EB02022 and HL56384 (to L. A. S.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

{ddagger} These two authors contributed equally to this work. Back

§ To whom correspondence and reprint requests should be addressed: Dept. of Pathology and Cancer Research and Treatment Center, MSC 08-4630, 1 University of New Mexico, Albuquerque, NM 87131-0001. Tel.: 505-272-6892; Fax: 505-272-6995; E-mail: lsklar{at}salud.unm.edu.

1 The abbreviations used are: VCAM-1, vascular cell adhesion molecule 1; BAPTA-AM, N,N'-[1,2-ethanediylbis(oxy-2,1-phenylene)]bis[N-[2-[(acetyloxy)methoxy]-2-oxoethyl]]-bis[(acetyloxy)methyl]ester; FEP, fluorinated ethylene propylene; GPCR, G-protein-coupled receptor; LDV-containing small molecule, 4-((N'-2-methylphenyl)ureido)-phenylacetyl-L-leucyl-L-aspartyl-L-valyl-L-prolyl-L-alanyl-L-alanyl-L-lysine; FITC, fluorescein isothiocyanate; LDV-FITC probe, 4-((N'-2-methylphenyl)ureido)-phenylacetyl-L-leucyl-L-aspartyl-L-valyl-L-prolyl-L-alanyl-L-alanyl-L-lysine-FITC; MCF, mean channel fluorescence; PTX, pertussis toxin; VLA-4, very late antigen-4({alpha}4{beta}1 integrin); MOPS, 4-morpholinepropanesulfonic acid; BAPTA, N,N'-[1,2-ethanediyl-bis(oxy-2,1-phenylene)]bis[N-[2-[(acetyloxy)methoxy]-2-oxoethyl]]; AM, bis[(acetyloxy)methyl]ester. Back

2 G. J. Zwartz, A. Chigaev, T. D. Foutz, B. S. Edwards, and L. A. Sklar, unpublished data. Back



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Springer, T. A. (1994) Cell 76, 301–314[CrossRef][Medline] [Order article via Infotrieve]
  2. Springer, T. A. (1995) Annu. Rev. Physiol. 57, 827–872[CrossRef][Medline] [Order article via Infotrieve]
  3. Carman, C. V., and Springer, T. A. (2003) Curr. Opin. Cell Biol. 15, 547–556[CrossRef][Medline] [Order article via Infotrieve]
  4. Chan, J. R., Hyduk, S. J., and Cybulsky, M. I. (2003) J. Immunol. Methods 273, 43–52[CrossRef][Medline] [Order article via Infotrieve]
  5. Faull, R. J., and Ginsberg, M. H. (1995) Stem Cells 13, 38–46[Abstract]
  6. Grabovsky, V., Feigelson, S., Chen, C., Bleijs, D. A., Peled, A., Cinamon, G., Baleux, F., Arenzana-Seisdedos, F., Lapidot, T., van Kooyk, Y., Lobb, R. R., and Alon, R. (2000) J. Exp. Med. 192, 495–506[Abstract/Free Full Text]
  7. Laudanna, C., Kim, J. Y., Constantin, G., and Butcher, E. (2002) Immunol. Rev. 186, 37–46[CrossRef][Medline] [Order article via Infotrieve]
  8. Sanchez-Mateos, P., Campanero, M. R., Balboa, M. A., and Sanchez-Madrid, F. (1993) J. Immunol. 151, 3817–3828[Abstract]
  9. Stewart, M., and Hogg, N. (1996) J. Cell. Biochem. 61, 554–561[CrossRef][Medline] [Order article via Infotrieve]
  10. Weber, C., Kitayama, J., and Springer, T. A. (1996) Proc. Natl. Acad. Sci. U. S. A. 93, 10939–10944[Abstract/Free Full Text]
  11. Moazzam, F., DeLano, F. A., Zweifach, B. W., and Schmid-Schonbein, G. W. (1997) Proc. Natl. Acad. Sci. U. S. A. 94, 5338–5343[Abstract/Free Full Text]
  12. Ingber, D. E. (2002) Circ. Res. 91, 877–887[Abstract/Free Full Text]
  13. Lehoux, S., and Tedgui, A. (1998) Hypertension 32, 338–345[Abstract/Free Full Text]
  14. Chien, S., Li, S., and Shyy, Y. J. (1998) Hypertension 31, 162–169[Abstract/Free Full Text]
  15. Ashida, N., Takechi, H., Kita, T., and Arai, H. (2003) J. Biol. Chem. 278, 9327–9331[Abstract/Free Full Text]
  16. Tzima, E., del Pozo, M. A., Shattil, S. J., Chien, S., and Schwartz, M. A. (2001) EMBO J. 20, 4639–4647[CrossRef][Medline] [Order article via Infotrieve]
  17. Chigaev, A., Blenc, A. M., Braaten, J. V., Kumaraswamy, N., Kepley, C. L., Andrews, R. P., Oliver, J. M., Edwards, B. S., Prossnitz, E. R., Larson, R. S., and Sklar, L. A. (2001) J. Biol. Chem. 276, 48670–48678[Abstract/Free Full Text]
  18. Chigaev, A., Zwartz, G., Graves, S. W., Dwyer, D. C., Tsuji, H., Foutz, T. D., Edwards, B. S., Prossnitz, E. R., Larson, R. S., and Sklar, L. A. (2003) J. Biol. Chem. 278, 38174–38182[Abstract/Free Full Text]
  19. Kew, R. R., Peng, T., DiMartino, S. J., Madhavan, D., Weinman, S. J., Cheng, D., and Prossnitz, E. R. (1997) J. Leukocyte Biol. 61, 329–337[Abstract]
  20. Chen, C., Mobley, J. L., Dwir, O., Shimron, F., Grabovsky, V., Lobb, R. R., Shimizu, Y., and Alon, R. (1999) J. Immunol. 162, 1084–1095[Abstract/Free Full Text]
  21. Chen, L. L., Whitty, A., Lobb, R. R., Adams, S. P., and Pepinsky, R. B. (1999) J. Biol. Chem. 274, 13167–13175[Abstract/Free Full Text]
  22. Lin, K., Ateeq, H. S., Hsiung, S. H., Chong, L. T., Zimmerman, C. N., Castro, A., Lee, W. C., Hammond, C. E., Kalkunte, S., Chen, L. L., Pepinsky, R. B., Leone, D. R., Sprague, A. G., Abraham, W. M., Gill, A., Lobb, R. R., and Adams, S. P. (1999) J. Med. Chem. 42, 920–934[CrossRef][Medline] [Order article via Infotrieve]
  23. Wang, J. H., Pepinsky, R. B., Stehle, T., Liu, J. H., Karpusas, M., Browning, B., and Osborn, L. (1995) Proc. Natl. Acad. Sci. U. S. A. 92, 5714–5718[Abstract/Free Full Text]
  24. Turitto, V. T., and Goldsmith, H. L. (1996) in Vascular Medicine (Loscalzo, J., Creager, M. A., and Dzau, V. J., eds) p. 141, Little, Brown, and Company, Boston
  25. Chien, S., Usami, S., and Skalak, R. (1984) in Handbook of Physiology (Renkin, E. M., Michel, C. C., and Geiger, S. R., eds) p. 225, American Physiological Society, Bethesda, MD
  26. Di Virgilio, F., Chiozzi, P., Ferrari, D., Falzoni, S., Sanz, J. M., Morelli, A., Torboli, M., Bolognesi, G., and Baricordi, O. R. (2001) Blood 97, 587–600[Abstract/Free Full Text]
  27. Jin, J., Dasari, V. R., Sistare, F. D., and Kunapuli, S. P. (1998) Br. J. Pharmacol. 123, 789–794[CrossRef][Medline] [Order article via Infotrieve]
  28. Kunapuli, S. P., and Daniel, J. L. (1998) Biochem. J. 336, 513–523[Medline] [Order article via Infotrieve]
  29. Wilson, B. S., Pfeiffer, J. R., Smith, A. J., Oliver, J. M., Oberdorf, J. A., and Wojcikiewicz, R. J. (1998) Mol. Biol. Cell 9, 1465–1478[Abstract/Free Full Text]
  30. Nieswandt, B., Schulte, V., Zywietz, A., Gratacap, M. P., and Offermanns, S. (2002) J. Biol. Chem. 277, 39493–39498[Abstract/Free Full Text]
  31. Berridge, M. J. (1993) Nature 361, 315–325[CrossRef][Medline] [Order article via Infotrieve]
  32. Bodor, E. T., Waldo, G. L., Hooks, S. B., Corbitt, J., Boyer, J. L., and Harden, T. K. (2003) Mol. Pharmacol. 64, 1210–1216[Abstract/Free Full Text]
  33. Sromek, S. M., and Harden, T. K. (1998) Mol. Pharmacol. 54, 485–494[Abstract/Free Full Text]
  34. Fields, T. A., and Casey, P. J. (1997) Biochem. J. 321, 561–571[Medline] [Order article via Infotrieve]
  35. Alon, R., Aker, M., Feigelson, S., Sokolovsky-Eisenberg, M., Staunton, D. E., Cinamon, G., Grabovsky, V., Shamri, R., and Etzioni, A. (2003) Blood 101, 4437–4445[Abstract/Free Full Text]
  36. Alon, R., Grabovsky, V., and Feigelson, S. (2003) Microcirculation 10, 297–311[CrossRef][Medline] [Order article via Infotrieve]
  37. von Andrian, U. H., Chambers, J. D., McEvoy, L. M., Bargatze, R. F., Arfors, K. E., and Butcher, E. C. (1991) Proc. Natl. Acad. Sci. U. S. A. 88, 7538–7542[Abstract/Free Full Text]
  38. Warnock, R. A., Campbell, J. J., Dorf, M. E., Matsuzawa, A., McEvoy, L. M., and Butcher, E. C. (2000) J. Exp. Med. 191, 77–88[Abstract/Free Full Text]
  39. Jackson, S. P., Nesbitt, W. S., and Kulkarni, S. (2003) J. Thromb. Haemost. 1, 1602–1612[CrossRef][Medline] [Order article via Infotrieve]
  40. Martinez-Lemus, L. A., Wu, X., Wilson, E., Hill, M. A., Davis, G. E., Davis, M. J., and Meininger, G. A. (2003) J. Vasc. Res. 40, 211–233[CrossRef][Medline] [Order article via Infotrieve]
  41. Nesbitt, W. S., Giuliano, S., Kulkarni, S., Dopheide, S. M., Harper, I. S., and Jackson, S. P. (2003) J. Cell Biol. 160, 1151–1161[Abstract/Free Full Text]
  42. Shyy, J. Y., and Chien, S. (2002) Circ. Res. 91, 769–775[Abstract/Free Full Text]
  43. Fisher, A. B., Chien, S., Barakat, A. I., and Nerem, R. M. (2001) Am. J. Physiol. 281, L529–L533
  44. Ingber, D. E. (2003) J. Cell Sci. 116, 1397–1408[Abstract/Free Full Text]
  45. Hamill, O. P., and Martinac, B. (2001) Physiol. Rev. 81, 685–740[Abstract/Free Full Text]
  46. Labrador, V., Chen, K. D., Li, Y. S., Muller, S., Stoltz, J. F., and Chien, S. (2003) Biorheology 40, 47–52[Medline] [Order article via Infotrieve]
  47. Chen, K. D., Li, Y. S., Kim, M., Li, S., Yuan, S., Chien, S., and Shyy, J. Y. (1999) J. Biol. Chem. 274, 18393–18400[Abstract/Free Full Text]
  48. Berk, B. C., Corson, M. A., Peterson, T. E., and Tseng, H. (1995) J. Biomech. 28, 1439–1450[CrossRef][Medline] [Order article via Infotrieve]
  49. Mazzucato, M., Pradella, P., Cozzi, M. R., De Marco, L., and Ruggeri, Z. M. (2002) Blood 100, 2793–2800[Abstract/Free Full Text]
  50. Tseng, H., Peterson, T. E., and Berk, B. C. (1995) Circ. Res. 77, 869–878[Abstract/Free Full Text]
  51. Cho, J. Y., Chain, B. M., Vives, J., Horejsi, V., and Katz, D. R. (2003) Exp. Cell Res. 290, 155–167[CrossRef][Medline] [Order article via Infotrieve]
  52. Weeks, B. S., Goldman, S., Touma, S., Payne, M., Cadet, P., and Stefano, G. B. (2001) J. Cell. Physiol. 189, 179–188[CrossRef][Medline] [Order article via Infotrieve]
  53. Zwartz, G., Chigaev, A., Foutz, T., Larson, R. S., Posner, R., and Sklar, L. A. (2004) Biophys. J. 86, 1243–1252[Medline] [Order article via Infotrieve]
  54. Dembo, M., Torney, D. C., Saxman K., and Hammer, D. (1988) Proc. R. Soc. Lond. B Biol. Sci. 234, 55–83[Medline] [Order article via Infotrieve]
  55. Bazzoni, G., Ma, L., Blue, M. L., and Hemler, M. E. (1998) J. Biol. Chem. 273, 6670–6678[Abstract/Free Full Text]
  56. Siedlecki, C. A., Lestini, B. J., Kottke-Marchant, K. K., Eppell, S. J., Wilson, D. L., and Marchant, R. E. (1996) Blood 88, 2939–2950[Abstract/Free Full Text]
  57. Cowan, K. J., Law, D. A., and Phillips, D. R. (2000) J. Biol. Chem. 275, 36423–36429[Abstract/Free Full Text]
  58. Garcia-Alvarez, B., Bobkov, A., Sonnenberg, A., and de Pereda, J. M. (2003) Structure (Camb.) 11, 615–625[Medline] [Order article via Infotrieve]
  59. Garcia-Alvarez, B., de Pereda, J. M., Calderwood, D. A., Ulmer, T. S., Critchley, D., Campbell, I. D., Ginsberg, M. H., and Liddington, R. C. (2003) Mol. Cell 11, 49–58[CrossRef][Medline] [Order article via Infotrieve]
  60. Goldfinger, L. E., Han, J., Kiosses, W. B., Howe, A. K., and Ginsberg, M. H. (2003) J. Cell Biol. 162, 731–741[Abstract/Free Full Text]
  61. Han, J., Rose, D. M., Woodside, D. G., Goldfinger, L. E., and Ginsberg, M. H. (2003) J. Biol. Chem. 278, 34845–34853[Abstract/Free Full Text]
  62. Tadokoro, S., Shattil, S. J., Eto, K., Tai, V., Liddington, R. C., de Pereda, J. M., Ginsberg, M. H., and Calderwood, D. A. (2003) Science 302, 103–106[Abstract/Free Full Text]
  63. Zell, T., Warden, C. S., Chan, A. S., Cook, M. E., Dell, C. L., Hunt, S. W., III, and Shimizu, Y. (1998) Curr. Biol. 8, 814–822[CrossRef][Medline] [Order article via Infotrieve]
  64. Pommerenke, H., Schreiber, E., Durr, F., Nebe, B., Hahnel, C., Moller, W., and Rychly, J. (1996) Eur. J. Cell Biol. 70, 157–164[Medline] [Order article via Infotrieve]
  65. Chen, J., Fabry, B., Schiffrin, E. L., and Wang, N. (2001) Am. J. Physiol. 280, C1475–C1484
  66. Danen, E. H., Lafrenie, R. M., Miyamoto, S., and Yamada, K. M. (1998) Cell Adhes. Commun. 6, 217–224[Medline] [Order article via Infotrieve]
  67. Martin, K. H., Slack, J. K., Boerner, S. A., Martin, C. C., and Parsons, J. T. (2002) Science 296, 1652–1653[Abstract/Free Full Text]
  68. Schmidt, C., Pommerenke, H., Durr, F., Nebe, B., and Rychly, J. (1998) J. Biol. Chem. 273, 5081–5085[Abstract/Free Full Text]
  69. Jalali, S., del Pozo, M. A., Chen, K., Miao, H., Li, Y., Schwartz, M. A., Shyy, J. Y., and Chien, S. (2001) Proc. Natl. Acad. Sci. U. S. A. 98, 1042–1046[Abstract/Free Full Text]
  70. Cinamon, G., Shinder, V., and Alon, R. (2001) Nat. Immunol. 2, 515–522[CrossRef][Medline] [Order article via Infotrieve]
  71. Rowin, M. E., Whatley, R. E., Yednock, T., and Bohnsack, J. F. (1998) J. Cell. Physiol. 175, 193–202[CrossRef][Medline] [Order article via Infotrieve]
  72. Jen, C. J., Jhiang, S. J., and Chen, H. I. (2000) J. Appl. Physiol 89, 1657–1662[Abstract/Free Full Text]
  73. Tsuboi, S. (2002) J. Biol. Chem. 277, 1919–1923[Abstract/Free Full Text]
  74. Takagi, J., Petre, B. M., Walz, T., and Springer, T. A. (2002) Cell 110, 599–611[CrossRef][Medline] [Order article via Infotrieve]
  75. Beglova, N., Blacklow, S. C., Takagi, J., and Springer, T. A. (2002) Nat. Struct. Biol. 9, 282–287[CrossRef][Medline] [Order article via Infotrieve]
  76. Chigaev, A., Buranda, T., Dwyer, D. C., Prossnitz, E. R., and Sklar, L. A. (2003) Biophys. J. 85, 99046–99058
  77. Marshall, B. T., Long, M., Piper, J. W., Yago, T., McEver, R. P., and Zhu, C. (2003) Nature 423, 190–193[CrossRef][Medline] [Order article via Infotrieve]
  78. Chen, S., and Springer, T. A. (1999) J. Cell Biol. 144, 185–200[Abstract/Free Full Text]

Add to CiteULike CiteULike   Add to Complore Complore   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us   Add to Digg Digg   Add to Reddit Reddit   Add to Technorati Technorati    What's this?


This article has been cited by other articles:


Home page
 J. Immunol.Home page
M. Phillipson, B. Heit, S. A. Parsons, B. Petri, S. C. Mullaly, P. Colarusso, R. M. Gower, G. Neely, S. I. Simon, and P. Kubes
Vav1 Is Essential for Mechanotactic Crawling and Migration of Neutrophils out of the Inflamed Microvasculature
J. Immunol., June 1, 2009; 182(11): 6870 - 6878.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Cell Physiol.Home page
D. A. Dikeman, L. A. Rivera Rosado, T. A. Horn, C. S. Alves, K. Konstantopoulos, and J. T. Yang
{alpha}4{beta}1-Integrin regulates directionally persistent cell migration in response to shear flow stimulation
Am J Physiol Cell Physiol, July 1, 2008; 295(1): C151 - C159.
[Abstract] [Full Text] [PDF]

Home page
 JEMHome page
R. Pasvolsky, S. W. Feigelson, S. S. Kilic, A. J. Simon, G. Tal-Lapidot, V. Grabovsky, J. R. Crittenden, N. Amariglio, M. Safran, A. M. Graybiel, et al.
A LAD-III syndrome is associated with defective expression of the Rap-1 activator CalDAG-GEFI in lymphocytes, neutrophils, and platelets
J. Exp. Med., July 9, 2007; 204(7): 1571 - 1582.
[Abstract] [Full Text] [PDF]

Home page
 J. Immunol.Home page
A. Chigaev, A. Waller, G. J. Zwartz, T. Buranda, and L. A. Sklar
Regulation of Cell Adhesion by Affinity and Conformational Unbending of {alpha}4beta1 Integrin
J. Immunol., June 1, 2007; 178(11): 6828 - 6839.
[Abstract] [Full Text] [PDF]

Home page
 J. Immunol.Home page
J. A. DiVietro, D. C. Brown, L. A. Sklar, R. S. Larson, and M. B. Lawrence
Immobilized Stromal Cell-Derived Factor-1{alpha} Triggers Rapid VLA-4 Affinity Increases to Stabilize Lymphocyte Tethers on VCAM-1 and Subsequently Initiate Firm Adhesion
J. Immunol., March 15, 2007; 178(6): 3903 - 3911.
[Abstract] [Full Text] [PDF]

Home page
 BloodHome page
S. J. Hyduk, J. R. Chan,, S. T. Duffy, M. Chen, M. D. Peterson, T. K. Waddell, G. C. Digby, K. Szaszi, A. Kapus, and M. I. Cybulsky
Phospholipase C, calcium, and calmodulin are critical for {alpha}4{beta}1 integrin affinity up-regulation and monocyte arrest triggered by chemoattractants
Blood, January 1, 2007; 109(1): 176 - 184.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
279/37/38277    most recent
M402944200v1
Right arrow Submit a Letter to Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Zwartz, G. J.
Right arrow Articles by Sklar, L. A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Zwartz, G. J.
Right arrow Articles by Sklar, L. A.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
 All ASBMB Journals   Molecular and Cellular Proteomics 
 Journal of Lipid Research   ASBMB Today 
Copyright © 2004 by the American Society for Biochemistry and Molecular Biology.
Advertisement
spacer
Advertisement
Advertisement