Lymphocyte binding to VCAM-1 activates endothelial cell NADPH oxidase, resulting in the generation of 1 μM H2O2. This is required for VCAM-1-dependent lymphocyte migration. In this study, we identified a role for protein kinase Cα (PKCα) in VCAM-1 signal transduction in human and mouse endothelial cells. VCAM-1-dependent spleen cell migration under 2 dynes/cm2 laminar flow was blocked by pretreatment of endothelial cells with dominant-negative PKCα or the PKCα inhibitors, Rö-32-0432 or Gö-6976. Phosphorylation of PKCαThr638, an autophosphorylation site indicating enzyme activity, was increased by Ab cross-linking of VCAM-1 on endothelial cells or by the exogenous addition of 1 μM H2O2. The anti-VCAM-1-stimulated phosphorylation of PKCαThr638 was blocked by scavenging of H2O2 and by inhibition of NADPH oxidase. Furthermore, anti-VCAM-1 signaling induced the oxidation of endothelial cell PKCα. Oxidized PKCα is a transiently active form of PKCα that is diacylglycerol independent. This oxidation was blocked by inhibition of NADPH oxidase. In summary, VCAM-1 activation of endothelial cell NADPH oxidase induces transient PKCα activation that is necessary for VCAM-1-dependent transendothelial cell migration.

Vascular cell adhesion molecule-1 expression is induced on endothelial cells in inflammatory tissues during atherosclerosis, immune challenge, and transplantation (1, 2, 3). VCAM-1 is required for eosinophil infiltration into the lung in experimental OVA-induced asthma (3) and T cell infiltration across the blood-brain barrier in experimental allergic encephalomyelitis (4). VCAM-1 also functions in combination with other adhesion molecules during chronic inflammation and tumor metastasis. Moreover, the VCAM-1 knockout is a mouse embryonic lethal (5). Therefore, understanding VCAM-1 signaling has important implications for disease intervention.

Leukocyte binding to VCAM-1 on endothelial cells activates endothelial cell signaling required for lymphocyte migration (6, 7, 8). We have reported that binding to VCAM-1 activates endothelial cell NADPH oxidase (6, 8, 9). NADPH oxidase generates superoxide that dismutates to hydrogen peroxide, yielding 1 μM H2O2 (9, 10). Extracellular H2O2 is generated by cell membrane NADPH oxidase as VCAM-1-mediated signals are blocked by exogenous addition of catalase (6, 11). H2O2 can then diffuse through membranes at 100 μm/s (12). The VCAM-1-induced generation of H2O2 activates localized endothelial cell actin structural changes and is required for VCAM-1-dependent lymphocyte migration (6). Mechanisms for reactive oxygen species (ROS)3-induced intracellular endothelial cell signals during VCAM-1-dependent lymphocyte migration have not been defined.

ROS can stimulate activation of protein kinase C (PKC). PKCα activity is most often described as requiring the cofactors Ca2+ and phosphatidylserine or diacylglycerol (DAG). PKCα can also become activated by H2O2 oxidation of its regulatory domain (13). Moreover, PKCα prepared from 5 mM H2O2-treated COS-7 cells did not require its cofactors Ca2+, phosphatidylserine, or DAG (14). However, this 5 mM H2O2 is much higher than the 1 μM H2O2 generated by VCAM-1 signaling (9, 10). It has not been reported whether PKCα is activated by VCAM-1-stimulated ROS production.

PKC activation by phorbol esters (PMA) or poly-l-arginine has also been shown to regulate cell shape and permeability in monolayers of endothelial or epithelial cells, respectively (15, 16, 17). Endothelial cell monolayer permeability is increased by PMA stimulation of PKCα in HUVECs (15). PMA stimulation induces contraction of bovine pulmonary artery endothelial cells and increases permeability to albumin (18, 19). Increases in vascular permeability and increases in leukocyte transendothelial migration occur in inflammatory sites. Whether VCAM-1 “outside-in” signals modulate PKC activity has not been reported.

In this study, we demonstrate that VCAM-1-stimulated endothelial cell NADPH oxidase activity results in transient activation of PKCα in endothelial cell lines and in cultures of human lung microvascular endothelial cells. In addition, we demonstrate that PKCα activity is required for VCAM-1-dependent transendothelial spleen cell migration.

The endothelial cell line mHEVa cells was previously derived from BALB/c mouse axillary lymph nodes and cultured as described (6, 9, 11, 20, 21, 22). The mHEVa cells have been spontaneously immortalized but are not transformed (20). Human microvascular endothelial cells from the lung (HMEC-Ls) (Clonetics) were grown in endothelial growth medium (Clonetics) plus 5% FCS and were used at passage 1–4. For spleen cells, single-cell suspensions were obtained from spleens of male 6- to 8-wk-old BALB/c mice (Harlan Industries) as previously described (6) and the RBC were lysed by hypotonic shock (20). The animal procedures were reviewed and approved by the Animal Care and Use Committee at Northwestern University (Chicago, IL).

Apocynin was from Acros Organics. Diphenyleneiodonium chloride (DPI), Gö-6976, Rö-32-0432, and rabbit anti-PKCα (catalog no. SA-144) were obtained from Biomol. The [5, 6, 8, 9, 11, 12, 14, 15-[3H] (N)]-arachidonic acid (60–100 Ci/mM) was obtained from PerkinElmer. Catalase was from ICN Biomedicals. Rat anti-mouse VCAM-1 (clone MVCAM.A), mouse anti-human VCAM-1 (clone 51-10C9), and rat IgG (isotype Ab, clone R35-95) were obtained from BD Pharmingen. Goat anti-mouse IgG1 (catalog no. 1070-01) and goat anti-rat IgG (catalog no. 3050-01) were from Southern Biotechnology Associates. Rabbit anti-phospho-PKCαβ Thr638 (catalog no. 9375), and mouse anti-phosphotyrosine (catalog no. 9411) were from Cell Signaling Technology. Rabbit anti-phosphoserine (catalog no. 61-8100) were from Zymed Laboratories. Mouse anti-β-actin (catalog no. ab6276) was obtained from Abcam. HRP-conjugated donkey anti-rabbit Ab was obtained from Amersham Biosciences. HRP-conjugated goat anti-mouse IgG was obtained from Bio-Rad. Dominant-negative (DN) PKCα in the plasmid pCMV (vector) was a gift from A. Descoteaux (University of Québec, Québec, Canada). This inactive transdominant mutant PKCα has the lysine in the ATP-binding domain replaced (23). Iodoacetamidofluorescein (IAF) (catalog no. I9271), anti-FITC (catalog no. F5636), DTT (catalog no. D-9779), DMSO (catalog no. 154938), and H2O2 (catalog no. H-1009) were obtained from Sigma-Aldrich.

The parallel plate flow chamber was used to examine migration under conditions of laminar flow. Spleen cells were used as a source of cells contiguous with the blood stream that could then migrate across endothelial cells. Spleen cell migration across the mHEV cell lines is stimulated by mHEV cell constitutive production of the chemokine MCP-1 (22) and is dependent on adhesion to VCAM-1 (6). We have previously reported that, after migration across the mHEV cells, the spleen cells are 65–70% B cells, 12–15% CD4+ cells, and 5–8% CD8+ cells (10). For this migration assay, endothelial cells were grown to confluence on slides and then the slide was placed in a parallel plate flow chamber (24). In vivo, in the absence of inflammation, the rapid fluid dynamics of the blood result in blood cells located midstream of the vascular flow (25). However, during inflammation, there is a change of fluid dynamics (25, 26, 27). With inflammation, vascular permeability increases yielding fluid flow from the blood into the tissues which likely contributes to contact of blood cells with the endothelium (“margination”) (25, 27). There is also cell contact as the blood cells leave the capillaries and enter the postcapillary venules (26). Therefore, spleen cells (3 × 106) were added to the flow chamber (3.5 cm2) at 2 dynes/cm2. Next, to initiate spleen cell contact with the endothelial cells in vitro, the spleen cells were allowed to settle in the chamber as monitored by microscopy and then 2 dynes/cm2 was applied for the 15 min laminar flow assay. We have observed by microscopy that during the assay under laminar flow, the spleen cells in contact with the endothelial cells either roll, roll and detach, or roll, firmly attach, and migrate. After cell contact, the focus of the studies is on mechanisms of transendothelial migration under conditions of laminar flow. For this assay, the coculture was exposed to laminar flow at 2 dynes/cm2 at 37°C for 3 min to examine cell association or for 15 min to examine cell migration. After the 3 or 15 min at 2 dynes/cm2, the cells are washed at 2 dynes/cm2 with PBS supplemented with 0.2 mM CaCl2 and 0.1 mM MgCl2 because cations are required for cell adhesion. These cells were fixed with 3% paraformaldehyde for 1 h. To quantify migrated spleen cells at 15 min, phase contrast microscopy was used to count migrated cells that are phase dark (28). It has been reported that the transendothelial migration of an individual leukocyte, after it has rolled to a site of migration, occurs in 2 min (20). However, transendothelial migration of leukocytes is asynchronous. In the laminar flow assay, spleen cell migration is detected by 15 min. The number of spleen cells that were associated but not migrated (phase light cells) at 15 min is low because in 15 min, the majority of nonmigrating cells roll off the monolayer of endothelial cells as determined by microscopy (data not shown). Therefore, the number of spleen cells associated with the endothelial cells at 3 min of laminar flow are those cells that mediated cell-cell contact.

mHEVa were grown to 90% confluence in 12-well plates or on slides. The cells were transfected with 1 μg of DN PKCα or the vector pCMV per well. Each of these transfections were performed in the presence of 2 μl/well LipofectAMINE 2000 Reagent (Invitrogen Life Technologies) according to instructions in culture medium without gentamicin, penicillin, or streptomycin. After 3.5 h, the medium was removed and replaced with medium containing antibiotics for 24 h.

Endothelial cells in 12-well plates were washed with PBS and lysed in 100 μl/well radioimmunoprecipitation buffer (1% Triton X-100, 0.5% sodium deoxycholate, 0.2% SDS, 150 mM NaCl, 10 mM, HEPES (pH 7.3), 2 mM EDTA, 10 mg/ml leupeptin, 10 mg/ml aprotinin, 100 mg/ml iodoacetamide, and 1 mM PMSF). Triplicate samples were combined, and lysates were incubated on ice for 20 min. Lysates were sonicated and then centrifuged for 10 min at 15,000 × g at 4°C. Equal volumes of this lysate were incubated with 50 μl of protein G beads and 5 μg/ml of indicated Ab overnight at 4°C with gentle rocking. These beads were washed three times with radioimmunoprecipitation buffer and one time each with 10 mM NaCl followed by 10 mM Tris (pH 7.4). Beads were heated for 5 min at 100°C in SDS sample buffer.

Cell lysates or immunoprecipitates in SDS sample buffer were analyzed by 7.5 or 12% SDS-PAGE and transferred to polyvinylidene difluoride membranes according to manufacturer’s instructions (Bio-Rad) (300 mA for 3 h). The membranes were blocked in 5% nonfat dried milk or in 5% BSA for phosphoserine Abs in TBS plus 0.1% Tween 20 (TBST) for 1 h at room temperature and washed three times for 5 min in TBST. Membranes were incubated with primary Abs in TBST plus 5% BSA overnight, washed three times for 5 min in TBST, incubated with secondary Abs in TBST plus 5% BSA for 1 h, washed three times for 10 min in TBST, and examined for detection with ECL (Amersham Biosciences) and autoradiography. Equal protein loading was verified by stripping the membrane with Restore Western Blot Stripping Buffer (Pierce) for 15 min and then labeling with rabbit anti-PKCα or mouse anti-β-actin. Densitometry was performed using Image J software from the National Institutes of Health. The data are presented as the fold increase in the ratio of relative intensity of the band/the relative intensity of band for the loading control (total PKCα or β-actin).

Endothelial cells were stimulated with anti-VCAM-1, isotype control Ab, or 1 μM H2O2. These cells were lysed in the presence of 10 μM IAF (pH 5.5–6) and PKCα was immunoprecipitated. Fluorescein or total PKCα was detected by Western blot. IAF reacts with nonoxidized cysteines (30). Thus, loss of IAF reactivity indicates oxidation. Controls include 10 mM DTT-reduced lysates for 10 min or lysates oxidized with 200 μM H2O2 for 20 min before addition of IAF.

The mHEV cells were prelabeled with 1.5 μCi/well [3H]arachidonic acid for 48 h. Where indicated, the cells were pretreated for 20 min with inhibitors. Cells were stimulated with H2O2 or by cross-linking VCAM-1 with 27 μg/ml rat anti-mouse VCAM-1 plus 15 μg/ml goat anti-rat IgG Ab. The stimulants and inhibitors were added to medium from the endothelial cell monolayers so that there was no addition of fresh serum-containing medium which stimulates phospholipase C activity. The H2O2 or VCAM-1 stimulation of endothelial cells was not done in serum-free conditions because serum is required for the mHEVa cell viability and the endothelial cell promotion of spleen cell migration (data not shown). The H2O2 or VCAM-1 stimulation was terminated with ice-cold methanol/concentrated HCl (100:1, v/v) and lipids were extracted by the method of Bligh and Dyer (31). After centrifugation at 2000 × g for 20 min, the lower chloroform phase was collected and evaporated under nitrogen. Samples as well as standards for DAG, phosphatidic acid, and arachidonic acid were resuspended in 15 μl of chloroform and spotted onto Silica Gel 60 TLC plates (200 μm; catalog no. 10028; Selecto Scientific). Plates were developed in a solvent system of hexane:ether:acetic acid (70:30:1) for the separation of DAG. The lipids were visualized by iodine vapor and then individual spots that comigrated with lipid standards were scraped and quantified by liquid scintillation spectrometry.

Data were analyzed by a one-way ANOVA followed by Tukey’s multiple comparisons test (SigmaStat; Jandel Scientific).

VCAM-1 is a member of the Ig superfamily. Another member of the Ig superfamily, ICAM-1, signals through PKCα in endothelial cells (32, 33). It is not known whether PKCα is involved in VCAM-1 signaling for VCAM-1-mediated lymphocyte migration. Therefore, we have used cultures of endothelial cells to determine whether endothelial PKCα is required for VCAM-1-dependent spleen cell migration under laminar flow. It has been previously reported that VCAM-1 can support leukocyte rolling, firm adhesion, and migration (6, 21, 34). Spleen cells were chosen as a source of cells contiguous with the blood. We chose an endothelial cell model that provides a method to examine VCAM-1-mediated lymphocyte migration, while avoiding endothelial cell outside-in signals from lymphocyte binding to other adhesion molecules on endothelial cells (6, 9, 10, 20). In this model, the cell line mHEVa constitutively expresses VCAM-1 and does not express other ligands for leukocyte migration such as ICAM-1, PECAM-1, and E-selectin as determined by immunofluorescence labeling (21), cDNA microarray analysis or leukocyte adhesion assays with blocking Abs (data not shown) (21). Spleen cell migration across these endothelial cells requires adhesion to VCAM-1 (6, 21). The endothelial cell line also constitutively produces MCP-1 for the stimulation of lymphocyte chemotaxis (22). To examine PKCα function, confluent monolayers of mHEVa cells were untreated or pretreated for 20 min with two inhibitors of PKCα, Rö-32-0432 (100 nM), or Gö-6976 (2.3 nM). The IC50 for Rö-32-0432 is 18–72 nM for purified rat brain PKCα, β, γ (35). Gö-6976 inhibits PKCα and PKCβI with IC50 values in the nanomolar range, whereas up to 3 μM Gö-6976 has no effect on the activity of PKC δ, ε, ζ (36, 37). After treatment with the PKCα inhibitors, the cells were washed five times. The last wash was added to a set of untreated cells to determine whether effective concentrations of free inhibitor were removed. Spleen cells were added at 2 dynes/cm2 laminar flow. After 15 min, the monolayers were washed and fixed. Spleen cell migration was quantified by phase contrast microscopy (28). Rö-32-0432 and Gö-6976 blocked spleen cell migration under laminar flow (Fig. 1 A).

FIGURE 1.

Inhibition of endothelial cell PKCα blocks VCAM-1-dependent spleen cell migration. A, Confluent monolayers of mHEVa cells were nontreated (NT) or treated for 20 min with the PKCα-selective inhibitors 2.3 nM Gö-6976 and 100 nM Rö-32-0432 and then washed five times. In addition, medium from the last washes was added to nontreated spleen cells to ensure that the inhibitor was sufficiently removed. Spleen cells were added to the endothelial monolayer in the presence or absence of anti-VCAM-1 (14 μg/cm2 endothelial cell monolayer) at 2 dynes/cm2, allowed to briefly settle to mediate cell contact and then exposed to 2 dynes/cm2 laminar flow for 15 min. Cells were washed and fixed in 3% paraformaldehyde for 1 h. Spleen cell migration was examined by phase contrast microscopy (28 ). Nonmigrated spleen cells are phase-light and migrated spleen cells appear as phase dark (28 ). The cells that migrated are >88% lymphocytes (10 ). Gö-6976 and Rö-32-0432 had no affect on cell viability, as determined by trypan blue exclusion (data not shown). Data are from three to five experiments. *, p < 0.05 compared with nontreated, DMSO treated, and last washes. B, Confluent monolayers of mHEVa cells were treated as in A with the inhibitors indicated, 2 dynes/cm2 laminar flow was applied for 3 min and then the cells were washed and fixed. The total number of spleen cells associated with the endothelial monolayer was determined. Data are from three experiments. *, p < 0.05 compared with cells without anti-VCAM-1 Abs.

FIGURE 1.

Inhibition of endothelial cell PKCα blocks VCAM-1-dependent spleen cell migration. A, Confluent monolayers of mHEVa cells were nontreated (NT) or treated for 20 min with the PKCα-selective inhibitors 2.3 nM Gö-6976 and 100 nM Rö-32-0432 and then washed five times. In addition, medium from the last washes was added to nontreated spleen cells to ensure that the inhibitor was sufficiently removed. Spleen cells were added to the endothelial monolayer in the presence or absence of anti-VCAM-1 (14 μg/cm2 endothelial cell monolayer) at 2 dynes/cm2, allowed to briefly settle to mediate cell contact and then exposed to 2 dynes/cm2 laminar flow for 15 min. Cells were washed and fixed in 3% paraformaldehyde for 1 h. Spleen cell migration was examined by phase contrast microscopy (28 ). Nonmigrated spleen cells are phase-light and migrated spleen cells appear as phase dark (28 ). The cells that migrated are >88% lymphocytes (10 ). Gö-6976 and Rö-32-0432 had no affect on cell viability, as determined by trypan blue exclusion (data not shown). Data are from three to five experiments. *, p < 0.05 compared with nontreated, DMSO treated, and last washes. B, Confluent monolayers of mHEVa cells were treated as in A with the inhibitors indicated, 2 dynes/cm2 laminar flow was applied for 3 min and then the cells were washed and fixed. The total number of spleen cells associated with the endothelial monolayer was determined. Data are from three experiments. *, p < 0.05 compared with cells without anti-VCAM-1 Abs.

Close modal

The last washes, that were added to nontreated cells, had no effect on spleen cell migration (Fig. 1,A). The inhibitors did not affect cell viability as determined by trypan blue exclusion (data not shown). Gö-6976 did not affect the total number of spleen cells that were associated with the endothelial cells under laminar flow (Fig. 1,B). Furthermore, Gö-6976 did not affect VCAM-1-mediated interaction of spleen cells with the endothelial cells (Fig. 1 B). The order of magnitude of cells migrated/cm2 in 15 min is similar to previous studies by us and others (6, 38, 39, 40, 41) when the data are adjusted for the experimental parameters such as well area and time course (data not shown). In summary, although there was no effect of inhibition of PKCα on association of the spleen cells with the endothelial cells, the spleen cells were unable to complete migration across the endothelial cells.

It was then determined whether transient transfection of endothelial cells with a DN PKCα blocks VCAM-1-dependent spleen cell migration. mHEVa cells on slides were Lipofectamine-transfected with 1 μg of DN PKCα plasmid or vector pCMV for 3.5 h, washed, and cultured for 24 h. These cells were examined for increased PKCα expression by Western blot as well as examined for their ability to promote spleen cell migration. Migration was examined in the parallel plate flow chamber assay at 2 dynes/cm2 laminar flow for 15 min (28). The transfection did not affect mHEVa cell expression of VCAM-1 as determined by immunolabeling and flow cytometry (data not shown). As expected, transfection with the DN PKCα increased total PKCα expression (Fig. 2,A). Importantly, transfection with the DN PKCα inhibited spleen cell migration as compared with the vector control (Fig. 2,B). The DN PKCα did not alter the number of spleen cells associated with the endothelial cells (Fig. 2 C). Therefore, taking together the data, VCAM-1-dependent spleen cell migration across monolayers of the mHEVa cell lines used endothelial cell PKCα activity.

FIGURE 2.

DN PKCα blocks VCAM-1-dependent spleen cell migration. mHEVa were grown to 90% confluence and transfected with 1 μg of vector with DN PKCα or vector pCMV for 3.5 h, washed, and cultured for 24 h. A, The cells were examined for total PKCα expression by Western blot. B, Spleen cells were added to the endothelial cell monolayer and the coculture was exposed to 2 dynes/cm2 laminar flow for 15 min as in Fig. 1. Cells were washed and fixed in 3% paraformaldehyde for 1 h. Spleen cell migration was examined by phase contrast microscopy. C, Spleen cells were added to the endothelial cell monolayer and the coculture was exposed to 2 dynes/cm2 laminar flow for 3 min as in Fig. 1. Cells were washed and fixed in 3% paraformaldehyde for 1 h. Total number of spleen cells associated with the endothelial cells was examined by microscopy. The DN PKCα had no affect on cell viability, as determined by trypan blue exclusion (data not shown). Data are from three experiments. *, p < 0.05 compared with nontreated.

FIGURE 2.

DN PKCα blocks VCAM-1-dependent spleen cell migration. mHEVa were grown to 90% confluence and transfected with 1 μg of vector with DN PKCα or vector pCMV for 3.5 h, washed, and cultured for 24 h. A, The cells were examined for total PKCα expression by Western blot. B, Spleen cells were added to the endothelial cell monolayer and the coculture was exposed to 2 dynes/cm2 laminar flow for 15 min as in Fig. 1. Cells were washed and fixed in 3% paraformaldehyde for 1 h. Spleen cell migration was examined by phase contrast microscopy. C, Spleen cells were added to the endothelial cell monolayer and the coculture was exposed to 2 dynes/cm2 laminar flow for 3 min as in Fig. 1. Cells were washed and fixed in 3% paraformaldehyde for 1 h. Total number of spleen cells associated with the endothelial cells was examined by microscopy. The DN PKCα had no affect on cell viability, as determined by trypan blue exclusion (data not shown). Data are from three experiments. *, p < 0.05 compared with nontreated.

Close modal

We previously reported that anti-VCAM-1 cross-linking of VCAM-1 mimics spleen cell activation of endothelial cell NADPH oxidase (6, 9, 11). Anti-VCAM-1 Abs are also used to demonstrate that binding to VCAM-1 can stimulate the signal rather than signals generated by a subsequent cell-cell interaction. Therefore, we determined whether anti-VCAM-1 stimulates endothelial cell PKCα activity. This was examined in endothelial cell lines (mHEVa cells) and cultures of nonimmortalized endothelial cells (HMEC-L). The endothelial cell lines constitutively express VCAM-1 whereas the HMEC-L cells require cytokine stimulation for expression of adhesion molecules. We stimulated adhesion molecule expression on HMEC-L cells with TNF-α overnight. After TNF-α stimulation, adhesion molecules including VCAM-1 were expressed by HMEC-L cells as determined by immunolabeling and fluorescence microscopy (data not shown). VCAM-1 on monolayers of mHEVa cells or TNF-α-treated HMEC-Ls was cross-linked with anti-VCAM-1 plus a secondary Ab for 10 min. Anti-CD98 was used as a control primary Ab because it binds to the mHEVa cells (data not shown) but CD98 does not signal through PKCα (42, 43, 44). After stimulation, the cells were washed, lysed, and examined by Western blot for the phosphorylation of PKCα Thr638. This Thr638 is an autophosphorylation site indicative of activation of PKCα (45). At the optimal time of 10 min (Fig. 3,A), Ab cross-linking of VCAM-1 significantly increased the phosphorylation of PKCα Thr638 as compared with nontreated cells, isotype Ab-treated cells, or anti-CD98-treated cells in mHEVa cells (Fig. 3, B and C). Anti-VCAM-1 also stimulated phosphorylation of PKCα Thr638 in cytokine-treated cultures of HMEC-L cells (Fig. 3,D). Furthermore, VCAM-1 stimulation did not increase total PKCα (Fig. 3 and see Fig. 6). In summary, stimulation of VCAM-1 increased PKCα activity in an endothelial cell line and in cultures of HMEC-L cells.

FIGURE 3.

Anti-VCAM-1 stimulation induces an increase in phosphorylation of PKCα Thr638 that is dependent on ROS generation. Confluent monolayers of A–C mHEVa cells and D TNF-α (1 ng/ml)-treated HMEC-L cells in 12-well plates were nontreated (NT) or incubated for 30 min with apocynin (4 mM), with catalase (5000 U/ml), or with DPI (5 μM). These endothelial cells were stimulated with 27 μg/ml anti-VCAM-1 Ab or control Ab plus 15 μg/ml of a secondary Ab for (A) 5–60 min or (B–D) 10 min. In D, the HMEC-L cells were also stimulated with 1 μM H2O2 for 10 min. Upper micrographs in each panel are representative Western blots using rabbit anti-phospho PKCα Thr638 (1/1000), rabbit anti-PKCα (1/100), or anti-β-actin (1/5000) followed by HRP-conjugated donkey anti-rabbit secondary Ab (1/4000) and ECL detection. The phosphorylation status of PKCα Thr638 is presented as the fold increase in the ratio of the relative intensity of phospho-PKCα Thr638/the relative intensity of the loading control (total PKCα or β-actin). The apocynin, catalase, DPI, and H2O2 had no effect on endothelial cell viability as determined by trypan blue exclusion (data not shown). Data presented are the mean ± SE from three experiments. *, p < 0.05 compared with NT, isotype Ab, or inhibitor-treated cells.

FIGURE 3.

Anti-VCAM-1 stimulation induces an increase in phosphorylation of PKCα Thr638 that is dependent on ROS generation. Confluent monolayers of A–C mHEVa cells and D TNF-α (1 ng/ml)-treated HMEC-L cells in 12-well plates were nontreated (NT) or incubated for 30 min with apocynin (4 mM), with catalase (5000 U/ml), or with DPI (5 μM). These endothelial cells were stimulated with 27 μg/ml anti-VCAM-1 Ab or control Ab plus 15 μg/ml of a secondary Ab for (A) 5–60 min or (B–D) 10 min. In D, the HMEC-L cells were also stimulated with 1 μM H2O2 for 10 min. Upper micrographs in each panel are representative Western blots using rabbit anti-phospho PKCα Thr638 (1/1000), rabbit anti-PKCα (1/100), or anti-β-actin (1/5000) followed by HRP-conjugated donkey anti-rabbit secondary Ab (1/4000) and ECL detection. The phosphorylation status of PKCα Thr638 is presented as the fold increase in the ratio of the relative intensity of phospho-PKCα Thr638/the relative intensity of the loading control (total PKCα or β-actin). The apocynin, catalase, DPI, and H2O2 had no effect on endothelial cell viability as determined by trypan blue exclusion (data not shown). Data presented are the mean ± SE from three experiments. *, p < 0.05 compared with NT, isotype Ab, or inhibitor-treated cells.

Close modal
FIGURE 6.

Anti-VCAM-1 and H2O2 induces oxidation of PKCα. mHEVa cells were treated with apocynin and then treated with anti-VCAM-1 or control Ab for 10 min as described in Fig. 3. A, The lysates were nontreated or reduced with 10 mM DTT for 10 min. As a positive control for oxidation, a lysate from NT cells was oxidized with 200 μM H2O2 for 20 min. The lysates were then examined for cysteine oxidation status by determining reactivity with 10 μM IAF (20 min), which reacts with nonoxidized cysteines. PKCα was immunoprecipitated and examined by Western blot using anti-fluorescein or anti-total PKCα Abs (1/1000). There was no change in total PKCα. B, mHEVa cells were stimulated for the time indicated with anti-VCAM-1 and examined for IAF reactivity as described in A. Data presented are the mean ± SD from three experiments. *, p < 0.05 as compared with NT. **, p < 0.05 for the comparison indicated.

FIGURE 6.

Anti-VCAM-1 and H2O2 induces oxidation of PKCα. mHEVa cells were treated with apocynin and then treated with anti-VCAM-1 or control Ab for 10 min as described in Fig. 3. A, The lysates were nontreated or reduced with 10 mM DTT for 10 min. As a positive control for oxidation, a lysate from NT cells was oxidized with 200 μM H2O2 for 20 min. The lysates were then examined for cysteine oxidation status by determining reactivity with 10 μM IAF (20 min), which reacts with nonoxidized cysteines. PKCα was immunoprecipitated and examined by Western blot using anti-fluorescein or anti-total PKCα Abs (1/1000). There was no change in total PKCα. B, mHEVa cells were stimulated for the time indicated with anti-VCAM-1 and examined for IAF reactivity as described in A. Data presented are the mean ± SD from three experiments. *, p < 0.05 as compared with NT. **, p < 0.05 for the comparison indicated.

Close modal

Adhesion to VCAM-1 stimulates endothelial cell H2O2 generation that is dependent on endothelial cell NADPH oxidase (9, 11) but not NO synthase, xanthine oxidase, or cytochrome P450 (6). NADPH oxidase generates superoxide that dismutates to H2O2. The VCAM-1-stimulated H2O2 generation is dependent on endothelial cell NADPH oxidase as it is blocked by apocynin (an NADPH oxidase inhibitor), by DPI (an irreversible inhibitor of flavoproteins such as NADPH oxidase) or by antisense to the catalytic subunit of NADPH oxidase (6, 9, 11). Therefore, we determined whether the anti-VCAM-1-stimulated generation of ROS activates PKCα. mHEVa cells were pretreated for 30 min with the H2O2 scavenger catalase (5000 U/ml), with apocynin (4 mM), or with DPI (5 μM). In addition, TNF-α-activated HMEC-Ls were pretreated for 30 min with 4 mM apocynin. We have previously reported that these concentrations of inhibitors block VCAM-1-stimulated generation of ROS and block VCAM-1-dependent lymphocyte migration without affecting lymphocyte adhesion to endothelial cells or endothelial cell viability (6, 10). After pretreatment, the cells were stimulated with anti-VCAM-1 or a control Ab and a secondary Ab. Anti-CD98 or isotype control Abs did not stimulate PKCα (Fig. 3). DPI, apocynin, and catalase inhibited anti-VCAM-1-stimulated phosphorylation of PKCα Thr638 in the mHEVa cells (Fig. 3, B and C). Apocynin also blocked VCAM-1-stimulated PKCα phosphorylation in cultures of HMEC-L cells (Fig. 3 D). In summary, PKCα has a function downstream of NADPH oxidase that is required for endothelial cell promotion of VCAM-1-dependent spleen cell migration.

Binding to VCAM-1 stimulates endothelial cell NADPH oxidase, resulting in the generation of 1 μM H2O2 (10). H2O2 diffuses through cell membranes at 100 μm/s (12), suggesting that the H2O2 produced at the endothelial cell membrane upon VCAM-1 stimulation may modulate intracellular signaling pathways. Therefore, it was determined whether exogenous H2O2 (0.1–5 μM) activated autophosphorylation of endothelial cell PKCα Thr638. At 10 min, 1 μM H2O2 significantly increased autophosphorylation of PKCα Thr638 in mHEVa cells (Fig. 4) and cultures of cytokine-treated HMEC-L cells (Fig. 3,D). A 5-fold increase in H2O2 (5 μM) had less activation of PKCα (Fig. 4). In summary, VCAM-1 activates endothelial cell NADPH oxidase resulting in the generation of low concentrations of H2O2 that then can activate endothelial cell PKCα. This H2O2 activation of PKCα was transient (Fig. 4 B). Therefore, H2O2, at concentrations that are generated by VCAM-1-outside-in signals (10), stimulated a significant increase in PKCα Thr638 phosphorylation.

FIGURE 4.

Exogenous 1 μM H2O2 activates endothelial cell-associated PKCα. A, mHEVa cells were incubated with low concentrations of exogenous H2O2 (0.1, 1, and 5 μM) for 10 min. Phospho-PKCα Thr638 was examined by Western blot and analyzed as described in Fig. 3. *, p < 0.05 as compared with 0, 0.1 and 5 μM H2O2. **, p < 0.05 as compared with 0 and 0.1 μM H2O2. B, mHEVa cells were incubated with 1 μM H2O2 for 1, 5, 10, or 15 min. *, p < 0.05 as compared with 1, 5, and 15 min. **, p < 0.05. A significant decrease compared with 10 min. Data presented are the mean ± SE from three experiments.

FIGURE 4.

Exogenous 1 μM H2O2 activates endothelial cell-associated PKCα. A, mHEVa cells were incubated with low concentrations of exogenous H2O2 (0.1, 1, and 5 μM) for 10 min. Phospho-PKCα Thr638 was examined by Western blot and analyzed as described in Fig. 3. *, p < 0.05 as compared with 0, 0.1 and 5 μM H2O2. **, p < 0.05 as compared with 0 and 0.1 μM H2O2. B, mHEVa cells were incubated with 1 μM H2O2 for 1, 5, 10, or 15 min. *, p < 0.05 as compared with 1, 5, and 15 min. **, p < 0.05. A significant decrease compared with 10 min. Data presented are the mean ± SE from three experiments.

Close modal

PKCα is a classical PKC that uses calcium and DAG as cofactors for activity. During cell stimulation of DAG-dependent PKCα, phospholipase generation of DAG is increased (46, 47). PKCα can also be activated by oxidation with 4 mM H2O2 and this activity is DAG independent (13, 14). Therefore, we determined whether binding to VCAM-1-stimulated generation of DAG. mHEV cells were prelabeled with 1.5 μCi/well [3H]arachidonic acid for 48 h during which the endothelial cells grew to confluence. Fresh culture medium was not added during anti-VCAM-1 or 1 μM H2O2 stimulation, as we observed that fresh serum in culture medium stimulates phospholipase activity in the endothelial cells (data not shown). An increase in free [3H]DAG was not detected after stimulation of endothelial cells by Ab cross-linking of VCAM-1 or treatment with 1 μM H2O2 (Fig. 5). Ab cross-linking of VCAM-1 or 1 μM H2O2 induced a slight decrease in DAG (Fig. 5). In summary, binding to VCAM-1 or 1 μM H2O2 did not stimulate an increase in DAG.

FIGURE 5.

Anti-VCAM-1 and H2O2 stimulation does not increase endothelial cell DAG. Phospholipase C activity was assessed by measuring the production of [3H]DAG by mHEV cells prelabeled with (A) 1.5 μCi/well or (B) 0.5 μCi/well [3H]arachidonic acid for 48 h. A, Endothelial cells were stimulated with 27 μg/ml rat anti-mouse VCAM-1 Ab plus 15 μg/ml goat anti-rat IgG Ab for 10 min. B, Endothelial cells were stimulated with 1 μM H2O2 for 30–600 s. *, p < 0.05. Data presented are the mean ± SE from three experiments.

FIGURE 5.

Anti-VCAM-1 and H2O2 stimulation does not increase endothelial cell DAG. Phospholipase C activity was assessed by measuring the production of [3H]DAG by mHEV cells prelabeled with (A) 1.5 μCi/well or (B) 0.5 μCi/well [3H]arachidonic acid for 48 h. A, Endothelial cells were stimulated with 27 μg/ml rat anti-mouse VCAM-1 Ab plus 15 μg/ml goat anti-rat IgG Ab for 10 min. B, Endothelial cells were stimulated with 1 μM H2O2 for 30–600 s. *, p < 0.05. Data presented are the mean ± SE from three experiments.

Close modal

Oxidation of PKCα has been reported to induce a transient activation of PKCα that is independent of DAG (13, 14). Because binding to VCAM-1 activated a transient increase in phosphorylation of PKCα Thr638 (Fig. 3,A) and no increase in free DAG in the cells (Fig. 5), it was determined whether binding to VCAM-1 induced oxidation of PKCα. Confluent monolayers of mHEVa cells were nontreated, treated with apocynin or treated with the 0.1% DMSO solvent control. The cells were nonstimulated, stimulated with anti-VCAM-1 or an isotype Ab plus a secondary Ab, or stimulated with 1 μM H2O2 for 10 min. The lysates from these cells were nontreated or as a control, they were reduced with 10 mM DTT for 10 min. A positive control for oxidation was a NT cell lysate incubated with 200 μM H2O2 for 20 min. To examine oxidation of cysteines, the lysates were incubated for 20 min with 10 μM IAF which reacts with nonoxidized cysteines (30). PKCα was immunoprecipitated and examined by Western blot for fluorescein or total PKCα. Anti-VCAM-1 induced a decrease in IAF reactivity (Fig. 6), indicating an oxidation of PKCα (30). PKCα from 1 μM H2O2-treated cells also had reduced reactivity with IAF (Fig. 6). These decreases in IAF reactivity were blocked by apocynin and were reversed by reduction of lysates with DTT (Fig. 6). As expected, the positive control lysates that were exposed to high levels of H2O2 exhibited reduced IAF reactivity (Fig. 6). The oxidation of PKCα in anti-VCAM-1-stimulated cells was transient (Fig. 6,B), which is consistent with the transient nature of PKCα activity after oxidation and the transient autophosphorylation at PKCα Thr638 (Fig. 3). In summary, anti-VCAM-1 and the VCAM-1-signaling intermediate, 1 μM H2O2, induced oxidation of PKCα and transient activation of PKCα.

In this study, we demonstrate that endothelial cell PKCα activity is an intracellular signal during VCAM-1-dependent cell migration. We previously reported that lymphocyte migration across mHEVa cells is blocked by Ab inhibition of VCAM-1 binding (6, 10) or by inhibition of endothelial cell NADPH oxidase activity (6, 9). We have shown here that this VCAM-1-dependent migration is blocked by inhibition of PKCα. The activation of PKCα is downstream of VCAM-1-stimulated NADPH oxidase in endothelial cell lines and cultures of HMEC-L cells. The endothelial cell NADPH oxidase-generated ROS oxidize and activate endothelial cell PKCα. Moreover, the exogenous addition of 1 μM H2O2, which corresponds to the levels of H2O2 produced after VCAM-1 stimulation (6, 10, 20), mimicked VCAM-1-induced phosphorylation of PKCα Thr638 and the oxidation of PKCα. The phosphorylation of PKCα is required for enzyme activity and Thr638 is an autophosphorylation site (45, 48). Thus, we demonstrate that VCAM-1 stimulates ROS-mediated activation of PKCα that is required for VCAM-1-dependent spleen cell migration. This is the first report on VCAM-1 signaling through PKCα and we identified an oxidation mechanism for this activation.

PKCα is a classical PKC that uses the cofactors calcium and DAG. We have previously reported that VCAM-1 stimulates release of intracellular calcium and calcium channels (9). However, in addition to DAG activation of PKCα, PKCα can also be activated independent of DAG when PKCα is oxidized by H2O2. The majority of studies on ROS modulation of PKC activity or endothelial cell function focus on high levels of ROS for damage to endothelial cells (49). It has been reported that 5–10 mM H2O2 induces translocation of PKCα to the membrane of human saphenous vein endothelial cells (50) or induces an increase in activity of an oxidatively modified PKCα that is independent of Ca2+ and phospholipids in transfected COS-7 cells, NIH3T3 cells, C6 glioma, or B16 melanoma cells (13, 14). H2O2 (5 mM) activates PKCα by oxidation of the PKC regulatory domain in 3–10 min (13). However, beyond 10 min, these high concentrations of H2O2 induce inactivation of the PKCα, presumably through additional oxidation (13). In addition, it has been reported that, when PKCα is oxidized by high concentrations of H2O2, it then induces activation of PLD for generation of phosphatidic acid in leukemia cells (51). They also reported that <200 μM H2O2 does not induce PKCα-mediated activation of PLD in the leukemia cells (51). Consistent with this, in our studies, PLD activity, as measured by free phosphatidic acid generation, was not increased in 1 μM H2O2-treated endothelial cells or in anti-VCAM-1-stimulated endothelial cells that generate 1 μM H2O2 (data not shown). We did demonstrate that low concentrations of H2O2 (1–5 μM) activate and oxidize PKCα in 10 min. However, in contrast to the later oxidative inactivation of PKCα in studies with high concentrations of H2O2, we found that, after 10 min, PKCα oxidation was reduced to baseline and its enzymatic activation was reversed. Therefore, with low concentrations of H2O2, there is reversible oxidation/activation of PKCα. We also demonstrated that there was not an increase in generation of free DAG under conditions that avoided addition of fresh serum-containing medium because fresh serum induced an increase in DAG (data not shown). Therefore, these data are consistent with transient DAG-independent PKCα activity downstream of VCAM-1 activation of NADPH oxidase. This transient activation of PKCα is in concordance with the rapid transendothelial migration of leukocytes as after migration, endothelial cell junctions are rapidly reformed. Furthermore, it is likely that the VCAM-1-stimulated production of a maximum of 1–2 μM H2O2 is tightly controlled so that leukocyte migration is obtained without vascular damage.

Localization of signals during leukocyte transendothelial migration is important as endothelial cells retract only at the site of contact with a leukocyte. Therefore, it is interesting to discuss how extracellular production of H2O2 could activate localized signals for endothelial cells. Cell membrane NADPH oxidase generates extracellular superoxide which rapidly dismutates to H2O2. Therefore, how would extracellular H2O2, which rapidly diffuses as well as passes through cell membranes at 100 μm/s (12), result in stimulation of localized endothelial function to allow endothelial cell shape changes at the site of leukocyte contact and VCAM-1-dependent leukocyte passage? Also, is not extracellular ROS just washed away by blood flow? Our current and previous studies suggest that, because exogenous addition of catalase scavenges the H2O2 and blocks VCAM-1-dependent responses (6, 11), VCAM-1 engagement results in extracellular H2O2. Specifically, we have reported that catalase scavenging of extracellular H2O2 blocks VCAM-1-dependent lymphocyte migration (6) and VCAM-1/ROS-dependent activation of endothelial cell-associated matrix metalloproteinases (MMPs) (11). Furthermore, this VCAM-1-dependent endothelial cell ROS oxidation and activation of endothelial cell-associated MMPs was not altered under static conditions verus conditions of laminar flow (11). Thus, a working model is that ROS produced by localized endothelial cell membrane NADPH oxidase: 1) oxidize localized proteins such as membrane-associated MMPs or 2) diffuse into the cell at 100 μm/s and act on localized intracellular PKCα, thereby affecting localized endothelial cell functions. In the cell, as the concentration of H2O2 is only 1 μM (9, 10), H2O2 likely oxidizes localized targets but, as the H2O2 diffuses, it would be too dilute to have a functional effect. In addition, extracellular ROS that is washed away by the blood flow would also be diluted and therefore have little functional effect at distant sites. The detection of localized VCAM-1-dependent changes in endothelial cell junctions is an exciting target recently under investigation in our laboratory and beyond the scope of this manuscript.

When addressing oxidative activation of PKC, it is important to include that PKCα activity is also regulated by phosphatases. Inhibition of Ser/Thr phosphatases (PPs) increases PKCα phosphorylation in bovine pulmonary microvascular endothelial cells (52). An inhibition of Ser/Thr PPs can occur by oxidation of the phosphatase active site (53, 54, 55, 56). Furthermore, the sensitivity of PKCα to Ser/Thr PPs is reversible (57), providing an additional potential mechanism for transient PKC activity during endothelial cell retraction. Alternatively, active phosphatases can dephosphorylate PKCα and thus may participate in down-regulation of PKCα activity, thereby contributing to the transient nature of the PKCα activity during VCAM-1 signaling. Whether VCAM-1-stimulated PKCα is also modulated by Ser/Thr PPs is an interesting area for future research. Nevertheless, we identified that ROS generated by anti-VCAM-1-stimulation induced oxidation of PKCα and activation of PKCα autophosphorylation. Both the oxidation and phosphorylation of PKCα was transient. Furthermore, the oxidative activation of PKCα was required for VCAM-1-dependent spleen cell migration.

There are many stimulants that increase vascular permeability through the regulation of PKCα and phosphatases. During inflammation, there is an increase in leukocyte migration across endothelium and an increase in vascular permeability. DN PKCα blocks histamine-induced increases in endothelial permeability (58). PKCα-dependent hyperpermeability of endothelium is also induced by the chemokine CCL2 (59) or the cytokine TNF-α (16). In addition, the inhibition of phosphatases induces cell rounding, cytoskeletal disorganization, and cellular detachment in smooth muscle cells, HUVECs, and HMEC-L cells (60, 61). Thus, during inflammatory processes, PKCα regulates increases in endothelial cell permeability and VCAM-1 signaling in endothelial cells.

During leukocyte passage into inflamed tissue, the migration of leukocytes across the endothelium can be inhibited by Abs against a limited number of endothelial cell adhesion molecules or by a combination of Abs against multiple endothelial cell adhesion molecules. With regards to VCAM-1-dependent inflammation in vivo, anti-VCAM-1 Abs block infiltration of eosinophils into the lung in experimental asthma and block T cell infiltration into the brain in experimental allergic encephalomyelitis (3, 4). Furthermore, Abs against the ligand for VCAM-1, α4 integrin, have been used in multiple sclerosis and inflammatory bowel disease patients (62, 63, 64, 65). Therefore, in our studies on identification of VCAM-1-dependent signals, we focused on VCAM-1-mediated models. The mHEVa cells are a model for lymphocyte triggering of VCAM-1 whereas examination of lymphocyte triggering of a complex mix of adhesion molecules on TNF-α-activated HMEC-L cells does not facilitate identification of signaling mechanisms for VCAM-1. Furthermore, we have reported that in vivo treatment with the ROS scavenger bilirubin blocks VCAM-1-dependent leukocyte migration but not the migration of leukocytes that use multiple other endothelial cell adhesion molecules (66). Therefore, in these studies, the VCAM-1 on HMEC-L cells expressing multiple adhesion molecules was specifically triggered by anti-VCAM-1 Abs rather than by lymphocyte binding as lymphocytes would trigger a complex set of intracellular signals through multiple adhesion molecules on the TNF-α-activated HMEC-L cells. For the same reasons, the migration of spleen cells across cytokine-activated HMEC-L cells was not examined to address VCAM-1-mediated signals as the HMEC-L cells express multiple receptors that support leukocyte migration and these receptors have different signaling pathways (8, 66). Nevertheless, our studies performed with HMEC-L cells demonstrate that VCAM-1 on microvascular endothelial cells activates PKCα through VCAM-1-triggered endothelial cell NADPH oxidase and ROS. The identification of mechanisms for VCAM-1 signaling is important for proposing intervention of VCAM-1-dependent processes in vivo such as VCAM-1-dependent lung eosinophilia, VCAM-1-dependent T cell migration into the brain in multiple sclerosis, or VCAM-1-dependent T cell migration into the bowel in inflammatory bowel disease (3, 4, 62, 63, 64, 65).

In summary, during VCAM-1 signaling, ROS oxidize, and transiently activate PKCα. This activation of PKCα is required for VCAM-1-dependent spleen cell migration. Thus, we have defined a novel mechanism for VCAM-1 signaling and identified a function for oxidized PKCα during VCAM-1-dependent spleen cell migration.

This manuscript is dedicated in memory of Norman Cook who constructed the parallel plate flow chamber and was an excellent teacher, builder, pilot, friend, and father.

The authors have no financial conflict of interest.

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.

1

This work was supported by National Institutes of Health Grant RO1 HL68171 (to J.M.C.-M.).

3

Abbreviations used in this paper: ROS, reactive oxygen species; PKC, protein kinase C; DAG, diacylglycerol; HMEC-L, human microvascular endothelial cells from the lung; DPI, diphenyleneiodonium chloride; IAF, iodoacetamidofluorescein; DN, dominant negative; MMP, matrix metalloproteinase; PP, phosphatase.

1
Iiyama, K., L. Hajra, M. Iiyama, H. Li, M. DiChiara, B. D. Medoff, M. I. Cybulsky.
1999
. Patterns of vascular cell adhesion molecule-1 and intercellular adhesion molecule-1 expression in rabbit and mouse atherosclerotic lesions and at sites predisposed to lesion formation.
Circ. Res.
85
:
199
-207.
2
Mueller, J. P., M. J. Evans, R. Cofiell, R. P. Rother, L. A. Matis, E. A. Elliott.
1995
. Porcine vascular cell adhesion molecule (VCAM) mediates endothelial cell adhesion to human T cells: development of blocking antibodies specific for porcine VCAM.
Transplantation
60
:
1299
-1306.
3
Chin, J. E., C. A. Hatfield, G. E. Winterrowd, J. R. Brashler, S. L. Vonderfecht, S. F. Fidler, R. L. Griffin, K. P. Kolbasa, R. F. Krzesicki, L. M. Sly, et al
1997
. Airway recruitment of leukocytes in mice is dependent on α4-integrins and vascular cell adhesion molecule-1.
Am. J. Physiol.
272
:
L219
-L229.
4
Baron, J. L., J. A. Madri, N. H. Ruddle, G. Hashim, C. A. Janeway, Jr.
1993
. Surface expression of α4 integrin by CD4 T cells is required for their entry into brain parenchyma.
J. Exp. Med.
177
:
57
-68.
5
Gurtner, G. C., V. Davis, H. Li, M. J. McCoy, A. Sharpe, M. I. Cybulsky.
1995
. Targeted disruption of the murine VCAM1 gene: essential role of VCAM-1 in chorioallantoic fusion and placentation.
Genes Dev.
9
:
1
-14.
6
Matheny, H. E., T. L. Deem, J. M. Cook-Mills.
2000
. Lymphocyte migration through monolayers of endothelial cell lines involves VCAM-1 signaling via endothelial cell NADPH oxidase.
J. Immunol.
164
:
6550
-6559.
7
van Wetering, S., J. D. van Buul, S. Quik, F. P. J. Mul, E. C. Anthony, J.-P. ten Klooster, J. G. Collard, P. L. Hordijk.
2002
. Reactive oxygen species mediate Rac-induced loss of cell-cell adhesion in primary human endothelial cells.
J. Cell Sci.
115
:
1837
-1846.
8
Cook-Mills, J. M., T. L. Deem.
2005
. Active endothelial cell function during inflammation.
J. Leukocyte Biol.
77
:
487
-495.
9
Cook-Mills, J. M., J. D. Johnson, T. L. Deem, A. Ochi, L. Wang, Y. Zheng.
2004
. Calcium mobilization and Rac1 activation are required for VCAM-1 (vascular cell adhesion molecule-1) stimulation of NADPH oxidase activity.
Biochem. J.
378
:
539
-547.
10
Tudor, K. S. R. S., K. L. Hess, J. M. Cook-Mills.
2001
. Cytokines modulate endothelial cell intracellular signal transduction required for VCAM-1-dependent lymphocyte transendothelial migration.
Cytokine
15
:
196
-211.
11
Deem, T. L., J. M. Cook-Mills.
2004
. Vascular cell adhesion molecule-1 (VCAM-1) activation of endothelial cell matrix metalloproteinases: role of reactive oxygen species.
Blood
104
:
2385
-2393.
12
Mathai, J. C., V. Sitaramam.
1994
. Stretch sensitivity of transmembrane mobility of hydrogen peroxide through voids in the bilayer: role of cardiolipin.
J. Biol. Chem.
269
:
17784
-17793.
13
Gopalakrishna, R., W. B. Anderson.
1989
. Ca2+- and phospholipid-independent activation of protein kinase C by selective oxidative modification of the regulatory domain.
Proc. Natl. Acad. Sci. USA
86
:
6758
-6762.
14
Konishi, H., M. Tanaka, Y. Takemura, H. Matsuzaki, Y. Ono, U. Kikkawa, Y. Nishizuka.
1997
. Activation of protein kinase C by tyrosine phosphorylation in response to H2O2.
Proc. Natl. Acad. Sci. USA
94
:
11233
-11237.
15
Dang, L., J. P. Seale, X. Qu.
2004
. Reduction of high glucose and phorbol-myristate-acetate-induced endothelial cell permeability by protein kinase C inhibitors LY379196 and hypocrellin A.
Biochem. Pharmacol.
67
:
855
-864.
16
Koss, M., G. R. Pfeiffer, II, Y. Wang, S. T. Thomas, M. Yerukhimovich, W. A. Gaarde, C. M. Doerschuk, Q. Wang.
2006
. Ezrin/radixin/moesin proteins are phosphorylated by TNF-α and modulate permeability increases in human pulmonary microvascular endothelial cells.
J. Immunol.
176
:
1218
-1227.
17
Ohtake, K., T. Maeno, H. Ueda, M. Ogihara, H. Natsume, Y. Morimoto.
2003
. Poly-l-arginine enhances paracellular permeability via serine/threonine phosphorylation of ZO-1 and tyrosine dephosphorylation of occludin in rabbit nasal epithelium.
Pharm. Res.
20
:
1838
-1845.
18
Garcia, J. G., H. W. Davis, C. E. Patterson.
1995
. Regulation of endothelial cell gap formation and barrier dysfunction: role of myosin light chain phosphorylation.
J. Cell. Physiol.
163
:
510
-522.
19
Stasek, J. E., Jr, C. E. Patterson, J. G. Garcia.
1992
. Protein kinase C phosphorylates caldesmon77 and vimentin and enhances albumin permeability across cultured bovine pulmonary artery endothelial cell monolayers.
J. Cell. Physiol.
153
:
62
-75.
20
Cook-Mills, J. M., J. S. Gallagher, T. L. Feldbush.
1996
. Isolation and characterization of high endothelial cell lines derived from mouse lymph nodes.
In Vitro Cell Dev. Biol.
32
:
167
-177.
21
Tudor, K.-S. R. S., T. L. Deem, J. M. Cook-Mills.
2000
. Novel α4-integrin ligands on an endothelial cell line.
Biochem. Cell Biol.
78
:
99
-113.
22
Qureshi, M. H., J. Cook-Mills, D. E. Doherty, B. A. Garvy.
2003
. TNF-α-dependent ICAM-1- and VCAM-1-mediated inflammatory responses are delayed in neonatal mice infected with Pneumocystis carinii.
J. Immunol.
171
:
4700
-4707.
23
St-Denis, A., F. Chano, P. Tremblay, Y. St-Pierre, A. Descoteaux.
1998
. Protein kinase C-α modulates lipopolysaccharide-induced functions in a murine macrophage cell line.
J. Biol. Chem.
273
:
32787
-32792.
24
Lawrence, M. B., L. McIntire, V., and S. G. Eskin. 1987. Effect of flow on polymorphonuclear leukocyte/endothelial cell adhesion. Blood 70: 1284–1290.
25
Nobis, U., A. R. Pries, G. R. Cokelet, P. Gaehtgens.
1985
. Radial distribution of white cells during blood flow in small tubes.
Microvasc. Res.
29
:
295
-304.
26
Smith, M. L., D. S. Long, E. R. Damiano, K. Ley.
2003
. Near-wall micro-PIV reveals a hydrodynamically relevant endothelial surface layer in venules in vivo.
Biophys. J.
85
:
637
-645.
27
Lipowsky, H. H..
2005
. Microvascular rheology and hemodynamics.
Microcirculation
12
:
5
-15.
28
Ager, A., S. Mistry.
1988
. Interaction between lymphocytes and cultured high endothelial cells: an in vitro model of lymphocyte migration across high endothelial venule endothelium.
Eur. J. Immunol.
18
:
1265
-1274.
29
Chen, Y.-H., Q. Lu, D. A. Goodenough, B. Jeansonne.
2002
. Nonreceptor tyrosine kinase c-Yes interacts with occludin during tight junction formation in canine kidney epithelial cells.
Mol. Biol. Cell
13
:
1227
-1237.
30
Wu, Y., K. S. Kwon, S. G. Rhee.
1998
. Probing cellular protein targets of H2O2 with fluorescein-conjugated iodoacetamide and antibodies to fluorescein.
FEBS Lett.
440
:
111
-115.
31
Bligh, E. G., W. J. Dyer.
1959
. A rapid method for total lipid extraction and purification.
Can. J. Biochem. Physiol.
37
:
911
-917.
32
Darley-Usmar, V. M., A. Severn, V. J. O’Leary, M. Rogers.
1991
. Treatment of macrophages with oxidized low-density lipoprotein increases their intracellular glutathione content.
Biochem. J.
278
:
429
-434.
33
Park, Y.-H., T. Suzuki, M. Miyama-Inaba, T. Masuda, Y. Yoshida, H. Uchino.
1986
. Feedback regulation of antibody formation by suppressive B cell factor: preferential suppression of high-affinity antibody production by memory B lymphocytes.
Intl. Arch. Allergy Appl. Immunol.
81
:
156
-164.
34
Alon, R., P. D. Kassner, M. W. Carr, E. B. Finger, M. E. Hemler, T. A. Springer.
1995
. The integrin VLA-4 supports tethering and rolling in flow on VCAM-1.
J. Cell Biol.
128
:
1243
-1253.
35
Wilkinson, S. E., P. J. Parker, J. S. Nixon.
1993
. Isoenzyme specificity of bisindolylmaleimides, selective inhibitors of protein kinase C.
Biochem. J.
294
:
335
-337.
36
Martiny-Baron, G., M. G. Kazanietz, H. Mischak, P. M. Blumberg, G. Kochs, H. Hug, D. Marme, C. Schachtele.
1993
. Selective inhibition of protein kinase C isozymes by the indolocarbazole Go 6976.
J. Biol. Chem.
268
:
9194
-9197.
37
Ma, R., P. E. Kudlacek, S. C. Sansom.
2002
. Protein kinase Cα participates in activation of store-operated Ca2+ channels in human glomerular mesangial cells.
Am. J. Physiol.
283
:
C1390
-C1398.
38
Pietschmann, P., J. J. Cush, P. E. Lipsky, N. Oppenheimer-Marks.
1992
. Identification of subsets of human T cells capable of enhanced transendothelial migration.
J. Immunol.
149
:
1170
-1178.
39
Yamamoto, H., J. B. Sedgwick, W. W. Busse.
1998
. Differential regulation of eosinophil adhesion and transmigration by pulmonary microvascular endothelial cells.
J. Immunol.
161
:
971
-977.
40
Smith, W. B., L. Noack, Y. Khew-Goodall, S. Isenmann, M. A. Vadas, J. R. Gamble.
1996
. Transforming growth factor-β1 inhibits the production of IL-8 and the transmigration of neutrophils through activated endothelium.
J. Immunol.
157
:
360
-368.
41
May, M. J., A. Ager.
1992
. ICAM-1-independent lymphocyte transmigration across high endothelium: differential up-regulation by interferon γ, tumor necrosis factor-α and interleukin 1β.
Eur. J. Immunol.
22
:
219
-226.
42
Kakugawa, K., M. Hattori, N. Beauchemin, N. Minato.
2003
. Activation of CEA-CAM-1-mediated cell adhesion via CD98: involvement of PKCδ.
FEBS Lett.
552
:
184
-188.
43
Cho, J. Y., K. M. Skubitz, D. R. Katz, B. M. Chain.
2003
. CD98-dependent homotypic aggregation is associated with translocation of protein kinase Cδ and activation of mitogen-activated protein kinases.
Exp. Cell Res.
286
:
1
-11.
44
Rintoul, R. C., R. C. Buttery, A. C. Mackinnon, W. S. Wong, D. Mosher, C. Haslett, T. Sethi.
2002
. Cross-linking CD98 promotes integrin-like signaling and anchorage-independent growth.
Mol. Biol. Cell
13
:
2841
-2852.
45
Keranen, L. M., E. M. Dutil, A. C. Newton.
1995
. Protein kinase C is regulated in vivo by three functionally distinct phosphorylations.
Curr. Biol.
5
:
1394
-1403.
46
Selvatici, R., S. Flazarano, A. Mollica, S. Spisani.
2006
. Signal transduction pathways triggered by selective formylpeptide analogues in human neutrophils.
Eur. J. Pharmacol.
534
:
1
-11.
47
Becker, K. P., Y. A. Hannun.
2005
. Protein kinase C and phospholipase D: intimate interactions in intracellular signaling.
Cell. Mol. Life Sci.
62
:
1448
-1461.
48
Cazaubon, S. M., P. J. Parker.
1993
. Identification of the phosphorylated region responsible for the permissive activation of protein kinase C.
J. Biol. Chem.
268
:
17559
-17563.
49
van der Goes, A., D. Wouters, S. M. van der Pol, R. Huizinga, E. Ronken, P. Adamson, J. Greenwood, C. D. Dijkstra, H. E. de Vries.
2001
. Reactive oxygen species enhance the migration of monocytes across the blood-brain barrier in vitro.
FASEB J.
15
:
1852
-1854.
50
Chen, C. C., C. S. Liau, Y. T. Lee.
1996
. Tumor necrosis factor-α, platelet-activating factor, and hydrogen peroxide activate protein kinase C subtypes α and ε in human saphenous vein endothelial cells.
J. Cardiovasc. Pharmacol.
28
:
240
-244.
51
Lee, B. D., J. H. Kim, S. D. Lee, Y. Kim, P. G. Suh, S. H. Ryu.
2000
. Hydrogen peroxide-induced phospholipase D2 activation in lymphocytic leukemic L1210 cells.
J. Leukocyte Biol.
67
:
630
-636.
52
Lum, H., J. L. Podolski, M. E. Gurnack, I. T. Schulz, F. Huang, O. Holian.
2001
. Protein phosphatase 2B inhibitor potentiates endothelial PKC activity and barrier dysfunction.
Am. J. Physiol.
281
:
L546
-L555.
53
Thannickal, V. J., B. L. Fanburg.
2000
. Reactive oxygen species in cell signaling.
Am. J. Physiol.
279
:
L1005
-L1028.
54
Forman, H. J., M. Torres.
2002
. Reactive oxygen species and cell signaling: respiratory burst in macrophage signaling.
Am. J. Resp. Crit. Care Med.
166
:
S4
-S8.
55
Sommer, D., S. Coleman, S. A. Swanson, P. M. Stemmer.
2002
. Differential susceptibilities of serine/threonine phosphatases to oxidative and nitrosative stress.
Arch. Biochem. Biophys.
404
:
271
-278.
56
Pieri, L., S. Dominici, B. Del Bello, E. Maellaro, M. Comporti, A. Paolicchi, A. Pompella.
2003
. Redox modulation of protein kinase/phosphatase balance in melanoma cells: the role of endogenous and γ-glutamyltransferase-dependent H2O2 production.
Biochim. Biophys. Acta
1621
:
76
-83.
57
Ricciarelli, R., A. Azzi.
1998
. Regulation of recombinant PKC α activity by protein phosphatase 1 and protein phosphatase 2A.
Arch. Biochem. Biophys.
355
:
197
-200.
58
Kolosova, I. A., S.-F. Ma, D. M. Adyshev, P. Wang, M. Ohba, V. Natarajan, J. G. N. Garcia, A. D. Verin.
2004
. Role of CPI-17 in the regulation of endothelial cytoskeleton.
Am. J. Physiol.
287
:
L970
-L980.
59
Stamatovic, S. M., O. B. Dimitrijevic, R. F. Keep, A. V. Andjelkovic.
2006
. Protein kinase C-α:RhoA cross talk in CCL-2-induced alterations in brain endothelial permeability.
J. Biol. Chem.
281
:
8379
-8388.
60
Gabel, S., J. Benefield, J. Meisinger, G. J. Petruzzelli, M. R. Young.
1999
. Protein phosphatases 1 and 2A maintain endothelial cells in a resting state, limiting the motility that is needed for the morphogenic process of angiogenesis.
Otolaryngol. Head Neck Surg.
121
:
463
-468.
61
Hosoya, N., M. Mitsui, F. Yazama, H. Ishihara, H. Ozaki, H. Karaki, D. J. Hartshorne, H. Mohri.
1993
. Changes in the cytoskeletal structure of cultured smooth muscle cells induced by calyculin-A.
J. Cell Sci.
105
:
883
-890.
62
Cree, B..
2006
. Emerging monoclonal antibody therapies for multiple sclerosis.
Neurologist
12
:
171
-178.
63
Bennett, J. L..
2006
. Natalizumab and progressive multifocal leukoencephalopathy: migrating towards safe adhesion molecule therapy in multiple sclerosis.
Neurol. Res.
28
:
291
-298.
64
Niino, M., C. Bodner, M.-L. Simard, S. Alatab, D. Gano, H. J. Kim, M. Trigueiro, D. Racicot, C. Guerette, J. P. Antel, et al
2006
. Natalizumab effects on immune cell responses in multiple sclerosis.
Ann. Neurol.
59
:
748
-754.
65
Lanzarotto, F., M. Carpani, R. Chaudhary, S. Ghosh.
2006
. Novel treatment options for inflammatory bowel disease targeting α4 integrin.
Drugs
66
:
1179
-1189.
66
Keshavan, P., T. L. Deem, S. J. Schwemberger, G. F. Babcock, J. M. Cook-Mills, S. D. Zucker.
2005
. Unconjugated bilirubin inhibits VCAM-1-mediated transendothelial leukocyte migration.
J. Immunol.
174
:
3709
-3718.