Enhancement of Migration by Protein Kinase Calpha and Inhibition of Proliferation and Cell Cycle Progression by Protein Kinase Cdelta in Capillary Endothelial Cells*

(Received for publication, September 27, 1996, and in revised form, December 17, 1996)

Elizabeth O. Harrington , Joachim Löffler , Peter R. Nelson , K. Craig Kent , Michael Simons and J. Anthony Ware Dagger

From the Vascular Biology Unit, Departments of Medicine and Vascular Surgery, Beth Israel Hospital and Harvard Medical School, Boston, Massachusetts 02215

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
REFERENCES


ABSTRACT

Activation of protein kinase C (PKC) induces angiogenesis, migration, and proliferation of endothelial cells (EC), but can also prevent growth factor-induced EC proliferation. To determine whether these disparate effects are mediated by substrates of individual PKC isoenzymes, PKCalpha and PKCdelta were overexpressed in rat microvascular EC. Basal and stimulated migration were enhanced in PKCalpha EC compared with either PKCdelta or control EC. Serum-induced growth of PKCdelta EC was decreased, while that of PKCalpha cells was similar to control EC. Phorbol ester markedly inhibited PKCdelta EC growth but enhanced growth of PKCalpha and control EC. To determine possible causes for this altered proliferation, the effect of PKCdelta on adhesion, mitogen-activated protein kinase activity, and cell cycle progression was measured. Adherence of PKCdelta EC to vitronectin was significantly enhanced. Serum-induced extracellular signal-regulated kinase-2 activity was increased equally in both PKCalpha and PKCdelta EC above that of control, while extracellular signal-regulated kinase-1 activity was similar in all EC. Cell cycle analysis suggested that PKCdelta EC entered S phase inappropriately and were delayed in passage through S phase. Thus, PKCalpha may mediate some proangiogenic effects of PKC activation; conversely, PKCdelta may direct antiangiogenic aspects of overall PKC activation, including slowing of the cell cycle progression.


INTRODUCTION

The formation of new blood vessels and the repair of those damaged by disease or injury depend upon endothelial migration and proliferation (1, 2). Several external agents that promote or inhibit proliferation and migration have been identified (3, 4), but the intracellular messengers that mediate these processes are less clear.

Activation of the serine-threonine kinase protein kinase C (PKC)1 by phorbol esters induces migration, proliferation (5), and tube formation of cultured endothelial cells (6, 7) and causes angiogenesis in vivo (7-9). In addition, chemical inhibitors of PKC or the down-regulation of PKC by prolonged treatment with phorbol esters abrogates the proliferative effects induced by growth factors and mitogens (10, 11) and also enhances endothelial permeability (12, 13) and alters the expression level of several fibrinolytic enzymes and their inhibitors (14). In contrast, treatment of endothelial cells with direct activators of PKC alters some responses that are usually associated with stimulation by physiologic agonists (15) and, under some conditions, can prevent growth factor-induced proliferation (16). This apparent paradox might be explained by the fact that the PKC family is composed of related but structurally distinct isoenzymes, each a product of separate genes and with discrete cofactor requirements, substrate specificity, and tissue distribution (16-18). Since phorbol esters activate multiple isoenzymes of PKC, the possibility is raised that each PKC isoenzyme may selectively mediate separate, and perhaps opposing, effects within stimulated endothelial cells.

Preliminary studies in our laboratory revealed that rat capillary endothelial cells expressed several isoenzymes of PKC, including PKCalpha , -delta , -eta , -theta , and -zeta . Of these isoenzymes, previous investigations have found that overexpression of PKCalpha or PKCdelta in various cultured cells could affect their proliferation (19-21). Overexpression of PKCalpha in fibroblasts promoted their proliferation, while proliferation was inhibited in human breast cancer cells and other cell lines (20, 22-24); similar alterations of cell growth have been observed in cells overexpressing PKCdelta (20, 21, 25). Thus, the possibility exists that activation of either of these isoenzymes mediates the inhibitory component of PKC activation on endothelial growth, while the other promotes one or more processes essential to endothelial repair and angiogenesis. Such an effect would be presumably mediated by isoenzyme-specific substrates, of which a few have been identified (e.g. elongation factor eEF-1alpha (26)). In the present study, we began by testing the effect of overexpression of PKCalpha and PKCdelta on endothelial migration and proliferation, both of which are essential processes for angiogenesis and wound healing. When initial experiments revealed an inhibitory effect of PKCdelta on endothelial proliferation, we then examined potential underlying causes.


EXPERIMENTAL PROCEDURES

Construction of Rat Fat Pad Epididymal Endothelial Cell (RFPEC) Cell Lines Overexpressing PKCalpha and PKCdelta

The RFPEC were a generous gift from R. D. Rosenberg (MIT) (27, 28) and were propagated in M199 medium (Life Technologies, Inc.) supplemented with 2 mM L-glutamine, penicillin (10 units ml-1), streptomycin (10 units ml-1), and amphotericin B (250 ng ml-1). To obtain the RFPEC that stably express vector (control), PKCalpha , or PKCdelta , pcDNA-Neo (Invitrogen, Inc.), pcDNA-bPKCalpha , or pcDNA-hPKCdelta constructs, respectively, were transfected into early passage RFPEC cells by the calcium phosphate precipitation technique. Following selection for resistance to Geneticin, a number of vector-transfected, PKCalpha -transfected, and PKCdelta -transfected clones of endothelial cells (designated as control EC, PKCalpha EC, and PKCdelta EC) were isolated and expanded, and the mRNA was examined by Northern blot analysis for expression of the respective transcripts.

Determination of Protein Kinase C Activity

Cells were removed from subconfluent cultures of stably transfected RFPEC by trypsin and washed with PBS. Total cell counts were determined using a Coulter counter. Equivalent numbers of cells were pelleted, resuspended in homogenization buffer (50 mM Tris, pH 7.5, 5 mM EDTA, 10 mM EGTA, 50 mg ml-1 Nalpha -p-tosyl-L-chloromethyl ketone, 100 mg ml-1 N-tosyl-L-phenylalanine chloromethyl ketone, 100 mg ml-1 phenylmethylsulfonyl fluoride, 2 mg ml-1 leupeptin, 0.3% beta -mercaptoethanol), and briefly sonicated. The cytosolic and cytoskeletal (the latter defined by its lack of solubility in 1% Triton X-100) fractions were obtained by differential centrifugation. PKC kinase activity was determined by measuring phosphorylation of a PKC-specific peptide substrate, based on a conserved region of the epidermal growth factor receptor, using the protein kinase C enzyme assay system purchased from Amersham Life Science, Inc.

RNA Analysis

The EC were removed from the culture dish with trypsin and washed with PBS. The cell pellet was resuspended in lysis buffer (140 mM NaCl, 1.5 mM Mg2Cl, 0.5% Triton X-100, 15 mM Tris, pH 8.3), vortexed for 30 s, and incubated on ice for 10 min. The nuclei were pelleted, and the supernatant was transferred to a fresh tube. An equivalent volume of buffer containing 25 mM EDTA, 300 mM NaCl, 2% SDS, 200 mg ml-1 proteinase K, and 200 mM Tris, pH 7.5, was added to the supernatant and incubated at 65 °C for 1 h. Total RNA was extracted with phenol/chloroform, precipitated with ethanol, and resuspended in DEPC-treated double distilled H2O. For Northern transfer analysis, 20 µg of total RNA was loaded per lane, subjected to electrophoresis on a 2% formaldehyde-agarose gel, and transferred to GeneScreen Plus membrane according to the manufacturer's recommendations. The blot was UV-cross-linked (Stratalinker, Stratagene) and hybridized with random primed cDNA probes at 65 °C for 3 h in Quik-Hyb solution (Stratagene). Blots were washed under high stringency conditions. Audioradiographs were analyzed using a scanning densitometer.

Immunoblot Analysis

Vector control, PKCalpha , and PKCdelta EC were removed from the culture dish with trypsin and washed with PBS. The cell pellet was resuspended in dissociation buffer (20 mM Tris-Cl, pH 6.8, 50 mM NaCl, 5 mM EDTA, 0.5% Triton X-100, 0.5% deoxycholate, 50 mg ml-1 Nalpha -p-tosyl-L-chloromethyl ketone, 100 mg ml-1 N-tosyl-L-phenylalanine chloromethyl ketone), sonicated briefly, and incubated at 37 °C for 30 min. Total crude cellular protein was suspended in 4 × Laemmli buffer (1 × Laemmli buffer 0.25 M Tris-Cl, pH 6.8, 8% SDS, 40% glycerol, 10% beta -mercaptoethanol, 0.004% bromphenol blue) and resolved on a 10% SDS-polyacrylamide separating gel with a 4% polyacrylamide stacking gel. The proteins were subsequently transferred to Immobilon-P membranes (Millipore, Inc.) according to the manufacturer's recommendations. Immunoblot analysis of the stably expressing clones was performed using the polyclonal antibodies for PKCalpha and PKCdelta obtained from Santa Cruz Biotechnology, Inc.

Immunohistochemistry

Stably transfected cells were propagated on coverslips placed in 12-well dishes and grown in the absence of serum for 24 h prior to immunofluorescence analysis of the PKC isoenzymes. The cells were washed with PBS and fixed in 2% paraformaldehyde in PBS for 10 min. The cells were then washed with PBS supplemented with 1% bovine serum albumin (PBS-BSA) and rendered permeable in 0.1% Triton X-100, PBS-BSA solution for 10 min. The cells were then washed and incubated in 10% goat serum, PBS-BSA solution for 30 min at room temperature. The solution was removed, and the cells were incubated in the presence of an optimal concentration of primary antibody (a 1:50 dilution of the PKCalpha , PKCdelta (Santa Cruz Biotechnology), or histone (Accurate Biochemicals, Inc.) antibodies) in 2% goat serum, PBS-BSA solution for 1 h. The cells were washed with 0.05% Triton X-100 in PBS-BSA. The biotinylated anti-rabbit or anti-mouse IgG (Vector Laboratories) was added at a 1:200 final dilution and incubated at room temperature for 1 h. The cells were washed, and streptavidin-Texas Red (Amersham Life Sciences) was added at a 1:200 dilution and incubated for 30 min at room temperature. Cells stained for the Golgi apparatus were treated with 0.5 µM BODIPY TR ceramide (Molecular Probes) for 1 h. Cells were washed once, and the coverslips were placed face-down on a slide with FluorSave (Calbiochem). The cells were visualized under × 40 magnification by fluorescence microscopy.

Cell Migration Assay

Stably transfected cells were grown to confluency and then incubated in serum-free M199 for 24 h prior to the start of the assay. The cells were removed from the culture dish with trypsin, washed once in PBS, and resuspended in serum-free M199 at a final concentration of 1 × 106 cells/ml of medium. Chemotactic agents were placed in the lower wells of a 48-well microchemotaxis chamber (Neuro Probe, Inc.) at indicated concentrations. An 8-µm porous, polyvinylpyrrolidone free, polycarbonate membrane (Poretics Corp.), precoated with collagen type I, was placed between the upper and lower wells of the chemotaxis chamber. The cell suspension was then added to the upper wells of the chamber at a density of 5 × 104 cells/well. Chemotaxis was assayed over 4 h at 37 °C in a CO2 incubator, under both unstimulated (basal) and stimulated (25 ng of hepatocyte growth factor/scatter factor per ml of medium) conditions. The membrane was removed from the chamber, fixed in 70% ethanol for 20 min at -20 °C, and stained in hematoxylin overnight. The upper surface of the stained membrane was scraped using a cotton swab, leaving only the cells that migrated to the undersurface. Migration was assessed by counting the number of cells on the lower surface of the membrane at a × 200 magnification by light microscopy.

Proliferation Assay

Endothelial cells were seeded at equivalent densities in six-well culture dishes and allowed to adhere overnight in complete medium. Following a wash with PBS, the cells were incubated for 24 h in M199 without serum. The cells were subsequently stimulated by the addition of enriched serum concentrations (either 1 or 15%) in the presence or absence of 1 µM phorbol 12-myristate 13-acetate (PMA). At the indicated times following stimulation, the cells were treated with trypsin, and the cells were counted with a Coulter counter.

Cell Adhesion Assay

The determination of endothelial cell adhesion was performed as described previously (29) with some modification. Briefly, Corning 96-well enzyme-linked immunosorbent assay plates were coated with 1 mg of purified human vitronectin resuspended in Ca2+, Mg2+-free PBS (pH 7.4) for 1 h at 37 °C. The plates were rinsed with the same buffer, coated with 1% heat-denatured BSA in the same buffer, and incubated at room temperature for 30-60 min. Actively growing (50-80% confluent) cultures of RFPEC were removed from the culture plate with trypsin, washed, and resuspended in M199 supplemented with 0.5% BSA. Cells were plated in each matrix-coated well at 2.5 × 104 cells/well and incubated at 37 °C for 60-90 min. The unadhered cells were removed, and the wells were gently washed. Adherent cells were detected by incubating in the presence of 6 mg ml-1 p-nitrophenylphosphate (Sigma) in 50 mM sodium acetate (pH 5), 1% Triton X-100 for 1 h at 37 °C, followed by the addition of N NaOH, and the absorbance was determined at 405 nm using an enzyme-linked immunosorbent assay plate reader.

Flow Cytometry

The cells were seeded, synchronized by serum deprivation for 72 h, and stimulated as described for the proliferation assay. DNA flow cytometric analysis of stably overexpressing EC cells was performed using a technique described by Tennenbaum et al. (30). The endothelial cells were removed from the culture plate by treating with 0.003% trypsin in sample buffer (3.4 mM sodium citrate, 0.1% Nonidet P-40, 1.5 mM spermine HCl, 0.5 mM Tris-Cl, pH 7.6). The reaction was stopped by the addition of 0.05% trypsin inhibitor, 0.01% ribonuclease A in sample buffer. The cells were subsequently treated with ice-cold 0.042% propidium iodide, 0.116% spermine HCl in sample buffer. The cells were kept on ice and in aluminum foil until analyzed. Flow cytometry was performed on a FACStar plus flow cytometer at an excitation of 488-nm wavelength and 630DF22 emission, and the data were analyzed using the Verity MODFIT software.

Stably transfected control, PKCalpha , and PKCdelta EC were removed with trypsin from the culture plate and resuspended in PBS at 5 × 106 cells ml-1 for the flow cytometric analysis of the surface expression of beta 3 or beta 5 integrins. Fifty-microliter aliquots of the cell suspensions were incubated in the presence of optimal concentrations of the primary antibody (1 µg of rabbit anti-rat beta 3 IgG fluorescein isothiocyanate-conjugated (Pharmingen, Inc.) or 0.5 µl of rabbit anti-human beta 5 polyclonal antibody (Chemicon, Inc.) per 50 µl reaction) at room temperature for 15 min. The cells were pelleted at 1000 × g for 5 min at 4 °C. When necessary, the cells were resuspended in 50 µl of PBS and incubated with appropriate volumes of the secondary antibody (1.2 mg ml-1 donkey anti-rabbit IgG fluorescein isothiocyanate-conjugated (Jackson Immunochemicals, Inc.)) for 15 min at room temperature. The cells were washed twice with PBS and resuspended in 200 µl of PBS. Fluorescence bound to cells was detected with a FACStar plus flow cytometer set at a 488-nm excitation wavelength and 530DF30 emission.

MAP Kinase Activity Assay

Vector control, PKCalpha , and PKCdelta EC were rendered quiescent for 24 h prior to the assay. The cells were stimulated with M199 supplemented with 15% FBS or 1 µM PMA for the indicated times. The cells were then harvested by removing the medium, washing once with ice-cold PBS, and incubating in radioimmune precipitation buffer (50 mM Hepes, pH 7.4, 150 mM NaCl, 5 mM EGTA, 5 mM EDTA, 20 mM NaF, 20 mM sodium pyrophosphate, 1% Triton X-100, 1 mM sodium orthovanadate, 1 mM phenylmethylsulfonyl fluoride) on a rocking incubator at 4 °C for 30 min. The lysed cells were scraped from the culture dish and transferred to a microcentrifuge tube. Large cellular debris was removed from the protein suspension by centrifugation at 15800 × g for 5 min at 4 °C. The cleared protein extracts were transferred to a fresh microcentrifuge tube, and total protein concentration was determined by means of the bicinchoninic acid assay (Pierce). MAP kinase was immunoprecipitated using extracellular signal-regulated kinase (ERK)-1 or ERK-2 antibody (Santa Cruz Biotechnology) from extract containing equal amounts of protein. After the sample volumes were adjusted with radioimmune precipitation buffer, the ERK-1 or ERK-2 antibody (1 µg of antibody/250 µg total protein) was added. The extracts were incubated on a rocking incubator at 4 °C overnight. The immune complexes were pelleted with protein A-agarose (Life Technologies, Inc.), washed three times in radioimmune precipitation buffer, and suspended in 50 µl of kinase buffer (10 mM Tris-Cl, pH 7.5, 150 mM NaCl, 10 mM MgCl2). ERK-2 activity was assayed by incubating 20 µl of each sample with 20 µl of the reaction mixture (8 mg of myelin basic protein (Sigma), 0.5 µCi of [gamma -32P]ATP (specific activity 3000 mCi/mmol) (DuPont NEN), and 10 µM ATP) for 30 min at 25 °C. The reaction was quenched with 15 µl of 4 × Laemmli buffer. The phosphorylated myelin basic protein was then resolved on a 15% SDS-polyacrylamide separating gel with a 4% polyacrylamide stacking gel and visualized by autoradiography.


RESULTS

Isolation and Characterization of Overexpressing Cell Lines

To investigate the role of PKCalpha and PKCdelta isoenzymes in relation to angiogenesis, stably transfected microvascular RFPEC were established that overexpressed the eukaryotic expression vector pcDNA-Neo containing the complete cDNA sequence of PKC isoenzyme PKCalpha or PKCdelta or, as a control, the expression vector without an inserted gene (PKCalpha EC, PKCdelta EC, and control EC, respectively). The stably expressing RFPEC cell lines were selected by neomycin resistance and screened by Northern blot analysis for gene expression (Fig. 1).


Fig. 1. Isolation and analysis of RNA from PKCdelta and PKCalpha EC. Overexpression of PKCdelta and PKCalpha was confirmed by analyzing the mRNA levels of stably transfected cell lines.
[View Larger Version of this Image (51K GIF file)]


The stably transfected RFPEC displayed the cobblestone morphology typical of endothelial cells and were not visibly altered by transfection or overexpression. Immunoblot analysis demonstrated increased protein production of the corresponding PKC isoenzymes. The enzymatic activity of PKC was determined in several cell lines. Following this initial analysis, two clones of each type (PKCalpha EC and PKCdelta EC) were chosen for further study on the basis of similar levels of total kinase activity. Table I summarizes the PKC activity of the control and the two selected PKCalpha and PKCdelta EC lines, revealing total PKC activity that was increased in both the cytosolic and cytoskeletal fractions and was comparable between PKCalpha and PKCdelta EC.

Table I.

Enhancement of PKC activity in stable cell lines


Clone Cellular fraction PKC activity Increase

pmol 32P/min -fold
Control EC Cytosolic 17.3 1
Cytoskeletal 14.9 1
PKCdelta EC Cytosolic 73.73 4.26
Cytoskeletal 31.7 2.13
PKCalpha EC Cytosolic 89.25 5.16
Cytoskeletal 27.44 1.84

To ensure that overexpression of PKCalpha and PKCdelta isoenzymes in the endothelial cells did not cause abnormal subcellular localization, we assessed the intracellular location of these PKC isoenzymes by immunofluorescence in both quiescent and PMA-treated EC. Experiments in PMA-treated EC were performed because activation of some PKC isoenzymes is associated with their redistribution into distinct subcellular locations (31). In quiescent control EC, PKCalpha could be detected primarily within the cytoplasm and nucleus (Fig. 2A). Staining of these cells with antibodies directed against histone proteins or with a fluorescently tagged ceramide confirmed the nuclei and Golgi apparatus structures (Fig. 2, I and J), respectively. Following a 10-min incubation with PMA, PKCalpha could still be seen in the nucleus, but also at the periphery of the cell along the plasma membrane (Fig. 2B). Interestingly, PKCalpha translocated primarily to regions of the plasma membrane in which there was cell-cell contact. A similar cellular distribution of PKCalpha was noted in both the PKCalpha EC (Fig. 2, C and D) and in the PKCdelta EC (data not shown). Immunofluorescent staining for PKCdelta demonstrated the nuclear and cytosolic location for this isoenzyme in serum-starved control EC (Fig. 2E). PKCdelta redistributed to the plasma membrane and the nuclear membrane upon activation with PMA (Fig. 2F). A similar pattern of staining for PKCdelta was noted in the PKCalpha (data not shown) and PKCdelta (Fig. 2, G and H) EC. Thus, constitutive overexpression of PKCalpha or PKCdelta did not affect normal cellular localization in either stimulated or quiescent endothelial cells, although enzymatic activity was increased.


Fig. 2. Intracellular location of PKCalpha and delta  in vector, PKCalpha , and PKCdelta EC. In vector control EC (A, B, E, and F), locations of both PKCalpha (A and B) and PKCdelta (E and F) are shown under quiescent (A and E) and PMA-stimulated (B and F) conditions. In PKCalpha EC (C and D), the location of PKCalpha under quiescent (C) and PMA-stimulated (D) conditions is shown. In G and H, the location of PKCdelta in PKCdelta EC is shown both before (G) and after (H) stimulation with PMA. Vector control EC were also stained for histone protein (I) to identify nuclei; with fluorescently labeled ceramide (J) to identify the Golgi apparatus; and with secondary antibody alone (anti-mouse and anti-rabbit antibodies; K and L) as negative controls.
[View Larger Version of this Image (83K GIF file)]


Effect of PKC Isoenzymes on Endothelial Cell Migration

To determine the role of PKCalpha and delta  in endothelial cell migration, the respective stably transfected cell lines were seeded in a microchemotaxis chamber, and the number of endothelial cells that migrated through the polycarbonate membrane was determined as described under "Experimental Procedures." When hepatocyte growth factor (HGF or scatter factor), a powerful stimulus for migration and angiogenesis (32, 33), was utilized as the agonist, PKCalpha EC traversed the membrane at a significantly greater rate than did PKCdelta or control EC (Fig. 3), suggesting a migratory response mediated by PKCalpha to this stimulus. The basal rate of migration (i.e. that occurring in the absence of any chemotactic agent) of PKCalpha was also consistently greater than that of the control EC or PKCdelta EC (Fig. 3), further implicating a specific role for PKCalpha in enhancing endothelial cell migration. Thus, both basal and agonist-stimulated endothelial migration differed between PKCalpha and PKCdelta EC, and PKCalpha EC migration was enhanced from the response seen in control EC.


Fig. 3. Enhanced migration in PKCalpha EC. The mean and S.D. are shown of the number of EC that migrated in response to stimulation with HGF, 25 ng/ml medium, or under basal conditions (without stimulation by HGF or serum) (n = 9; *, p <=  0.01 versus control or PKCdelta EC).
[View Larger Version of this Image (27K GIF file)]


Effect of PKCalpha and PKCdelta on Endothelial Cell Proliferation

Stimulation of quiescent PKCalpha EC with low (1%) concentrations of serum induced a growth rate similar to that of the vector control EC (Fig. 4A). In contrast, PKCdelta EC exhibited much less proliferation in response to serum stimulation than did either the control or PKCalpha EC. Proliferation of the control EC and PKCalpha EC in response to PMA was mildly enhanced (7.5%) above EC treated with serum alone at 72 h (Fig. 4B), but the growth rate of PKCdelta EC was significantly inhibited further by PMA, with an inhibition of 46.2 ± 8.8% below that of non-PMA-treated PKCdelta EC at 72 h (p < 0.05) (Fig. 4C). This inhibition of serum-induced growth to serum stimulation was noted in both clones of PKCdelta EC tested; neither of the PKCalpha EC clones exhibited altered growth. Qualitatively similar responses were seen when quiescent cells were stimulated with higher (15%) serum concentrations in the presence or absence of PMA (data not shown). Thus, overexpression and stimulation of one isoenzyme (PKCdelta ) blocked endothelial proliferation, a response not seen when PKCalpha was overexpressed to a similar degree.


Fig. 4. Overexpression of PKCdelta inhibits cellular proliferation in EC. All EC were rendered quiescent by serum deprivation and then stimulated with media containing 1% serum alone (A) or with PMA 1 µM (B). Cell counts were determined at the indicated number of hours after the initiation of serum stimulation. Data are expressed as the mean and S.D. of the percentage of change in the number of EC from that present before serum stimulation (at 0 h) in three determinations in a single experiment, which was repeated three times with similar results. C summarizes data of three experiments from a single clone of control (open circle ) and PKCdelta (black-square) EC, demonstrating the percentage of inhibition of endothelial cell growth upon PMA stimulation. The data are qualitatively similar to results obtained testing the other clones of each type and are presented as the mean and S.D. (*, p <=  0.05 versus control).
[View Larger Version of this Image (18K GIF file)]


Effects of PKCalpha and PKCdelta Overexpression on Endothelial Cell Adhesion

In order to better understand the cause of the decreased proliferation in PKCdelta EC, we next asked whether PKC would lessen adhesion to extracellular matrices, prevent mitogen-activated protein kinase (MAP kinase) activation, or alter cell cycle progression in EC. To determine whether integrin-mediated endothelial adhesion, an event that is required for endothelial proliferation and angiogenesis (29, 34, 35), was lessened by PKCdelta , we examined the adhesion of subconfluent cultures of control, PKCalpha , and PKCdelta EC seeded on vitronectin-coated plates. Rather than being diminished, the ability of the PKCdelta EC to adhere to the extracellular matrix was significantly enhanced above that seen with the control EC (Fig. 5), with a mean increase in adherence of 32.4% (p <=  0.005). In contrast, increased PKCalpha expression did not significantly alter the ability of the endothelial cells to adhere to vitronectin. Preincubation of these cells with a synthetic peptide, GRGDSP, which corresponds to the vitronectin protein sequence that directly interacts with the integrin receptor binding domain, abolished adherence of these cells, thus demonstrating that the base line adherence of the cells, plus that enhanced in PKCdelta EC, was specific for the integrin receptors. To determine whether the enhanced adhesion resulted from increased expression of integrin receptors on the cell surface, we analyzed the cellular surface expression level of alpha vbeta 3 and alpha vbeta 5 by immunofluorescence using flow cytometry. Neither the overall cellular surface expression level of beta 3 nor that of beta 5 was significantly enhanced in PKCdelta EC as compared with PKCalpha or control EC (data not shown). Thus, enhanced adhesion in PKCdelta EC most probably resulted from increased affinity modulation of the integrin receptors. These results, therefore, demonstrate that the reduction in cell growth in PKCdelta did not result from impaired adhesion to extracellular matrices.


Fig. 5. Effect of PKCdelta and PKCalpha on EC adhesion to vitronectin. Equivalent numbers of stably transfected endothelial cells were seeded on enzyme-linked immunosorbent assay plates coated with vitronectin and incubated for 1 h. Adherent cells were detected as described under "Experimental Procedures." Results shown are the combined mean and S.D. of the OD (n = 7, control EC; n = 15, PKCdelta EC; n = 12, PKCalpha EC; *, p <=  0.005 versus control or PKCalpha EC).
[View Larger Version of this Image (25K GIF file)]


Serum-induced ERK-2 Activation in PKCalpha and PKCdelta EC

We next tested the possibility that impaired PKCdelta EC growth resulted from impaired activation of one of the MAP kinases, ERK-1 or ERK-2, that are known to be activated following overall PKC activation (36, 37) in EC.

Serum stimulation of vector (control) EC that had been rendered quiescent demonstrated a rapid increase in ERK-2 activity by 10 min, with a gradual diminution of the kinase activity by 2-4 h after stimulation (Fig. 6, A and B). ERK-2 activity also increased within 10 min following serum stimulation of the PKCalpha and PKCdelta EC; however, the activity was enhanced above control and remained elevated above the basal kinase activity even at 4 h following stimulation. Phorbol ester treatment of the stably transfected cells resulted in similar levels of ERK-2 activity in PKCalpha or PKCdelta EC as compared with control, with the overall ERK-2 activity diminishing at a more rapid rate in control EC (i.e. 4 h) (Fig. 6, A and C). Similar analyses demonstrated very low overall ERK-1 activity in all cells tested. Serum stimulation resulted in mild enhanced ERK-1 activity; however, there were no noticeable differences between the control, PKCalpha , and PKCdelta EC (data not shown). Thus, PKCdelta and PKCalpha appeared to be equally effective in activating ERK-2alpha , and thus the decrease in proliferation in PKCdelta EC appeared to result from a mechanism independent of ERK-1 or ERK-2 activity.


Fig. 6. Analysis of ERK-2 activity in control (bullet ), PKCdelta (triangle ), and PKCalpha (square ) EC upon serum or PMA stimulation. All EC were rendered quiescent by serum deprivation and then stimulated with 15% serum or 1 µM PMA. The cells were harvested, ERK-2 was immunoprecipitated, and kinase activity was assayed by the ability to phosphorylate myelin basic protein. The phosphorylated myelin basic protein was resolved on SDS-polyacrylamide gel electrophoresis, and a representative autoradiograph is shown in A. The audioradiographs were quantitated using a scanning densitometer, and the results are presented in B (serum-stimulated) and C (PMA-stimulated).
[View Larger Version of this Image (22K GIF file)]


Effect of PKCdelta and PKCalpha on EC Cell Cycle Progression

To determine if an alteration in cell cycle progression might explain the decrease in cell growth in PKCdelta EC, cell cycle analysis was performed on serum-deprived and stimulated PKCdelta , PKCalpha , and control EC. After 72 h of serum starvation, 19.8 ± 4.2% of control EC and 17.1 ± 0.4% PKCalpha EC were in S phase, with 74.2 ± 5.5% and 77.3 ± 0.9% in G0/G1, respectively (Fig. 7A). In contrast, 26.3 ± 2.2% of the PKCdelta cells were in S phase, with 66.5 ± 1.9% in G0/G1. These data indicate that an abnormally high percentage of PKCdelta EC entered S phase inappropriately, i.e. under conditions of serum deprivation. Stimulation with serum caused control EC and PKCalpha EC to reenter the cell cycle normally (Fig. 7, B and C). PKCdelta EC, on the other hand, after an initial increase in the percentage of cells in G2/M phase at 6 h, followed by an increase in cells in G0/G1 at 12 h after serum, returned to a very high percentage of cells in S phase up to 60 h (Fig. 7D). The prolonged time that a high percentage of PKCdelta EC could be found in S phase suggested that these cells required an abnormal amount of time to complete S phase.


Fig. 7. Cell cycle analysis of control, PKCdelta , and PKCalpha EC. In A, representative flow cytometric tracings are shown of the distribution of control (upper) and PKCdelta (lower) EC populations, demonstrating a higher percentage of cells in S phase (solid area) and a correspondingly lower percentage in G0/G1 (hatched area) in the PKCdelta EC. Dotted area, G2/M phase. In B, C, and D, quantification of an experiment is presented in which vector control (B), PKCalpha (C), and PKCdelta (D) EC rendered quiescent by serum starvation for 72 h were stimulated with serum. At the indicated times following stimulation, the cells were collected and stained with propidium iodide, and the percentage of cells in each phase of the cell cycle was determined on FACStar flow cytometer. These results are representative of two experiments performed in two separate clonal populations. The solid bar represents G0/G1 phase, the hatched bar represents G2/M phase, and the open bar represents S phase.
[View Larger Version of this Image (37K GIF file)]



DISCUSSION

The two major findings of this study are that overexpression of two different PKC isoenzymes normally expressed in microvascular endothelial cells exert distinct effects on endothelial proliferation, migration, and adhesion to extracellular matrix and that PKCdelta -mediated inhibition of endothelial growth results from a defect in S phase of the endothelial cell cycle. The observation that overexpression of PKCdelta , but not PKCalpha , prevents proliferation of microvascular endothelial cells, while PKCalpha enhances their migration in response to HGF (scatter factor), suggests that these isoenzymes phosphorylate different substrates in these cells with different physiologic effects. The disparity between the effects of overexpression of the two isoenzymes is heightened by PKC-activating phorbol esters in that treatment of PKCalpha EC exerts a mitogenic effect similar to that in control cells, while PKCdelta EC were even more strongly inhibited by similar treatment. Thus, these data suggest that activation of each of these isoenzymes by angiogenic stimuli, such as HGF or those contained in serum, may mediate distinct aspects of several processes that are required for vascular repair and angiogenesis. Stimulation of endothelium with phorbol esters, which activate both PKCalpha and PKCdelta , as well as several other isoenzymes expressed in endothelium (5, 38), has been noted to have both stimulatory and inhibitory effects on endothelial proliferation and angiogenesis (5, 10, 39, 40) that can be temporally dispersed within the same cells (41). The results from this study suggest that PKCdelta might mediate those aspects of PKC activation that are inhibitory for endothelial repair and angiogenesis, while an important component of the proangiogenic effect, endothelial migration, might be mediated by PKCalpha .

Repair of small defects in endothelium are accomplished largely by endothelial migration, rather than proliferation (3, 42). Migration is an important component of angiogenesis as well (3). The present study's finding that endothelial migration is enhanced in PKCalpha EC might result from enhancement of cytoskeletal reorganization in response to stimuli, which is a necessary component of cell locomotion; overall PKC stimulation has been associated with promotion of cytoskeletal reorganization of endothelial cells (3, 43). Our results suggest that PKCalpha , but not PKCdelta , may be at least one mediator of migration response to HGF (scatter factor), a powerful angiogenic agent that is present along with its receptor in a substantial amount in the vasculature (4, 32, 44, 45).

We considered several possible explanations for the PKCdelta -mediated inhibition of endothelial growth, including a reduction in endothelial adhesion to matrix, a failure to activate downstream mediators such as ERK-1 or ERK-2, and a defect in progression of endothelial cells through the cell cycle. Of these explanations, only the latter appears to be the case. Regarding adhesion, both endothelial cell growth and migration are thought to require attachment of the cell to matrix via its integrin surface receptors (42, 46, 47). Blocking the vitronectin integrin receptors alpha vbeta 3 or alpha vbeta 5 inhibits neovascularization in the cornea or chick chorioallantoic membrane models (35), suggesting the importance of these two integrins for endothelial cell proliferation. In this study, microvascular endothelial cells in which PKCdelta was overexpressed demonstrated markedly enhanced adhesion to vitronectin or fibrinogen matrices, and thus a decrease in integrin mediated adhesion was not the cause of the decrease in proliferation of PKCdelta EC. This enhancement of adhesion could result from either a PKC-mediated alteration of the number of these receptors or an increase in avidity by either a direct effect on integrin conformation (so-called "inside-out" signaling) (48) or as an amplification step following cell adhesion ("outside-in") that prevents detachment (48). Overall PKC activation has been linked with promotion of integrin avidity for soluble fibrinogen and solid matrices in several cell types (49, 50). Neither PKCalpha EC nor PKCdelta EC demonstrated increased expression of these receptors when compared with control; thus, a PKCdelta -mediated effect on affinity of these integrins for vitronectin is a likely explanation for our results.

A cascade of signaling events merging at the MAP kinase family of proteins, ERK-1 and ERK-2, is involved in many of the intracellular signaling pathways that lead to endothelial cell growth, migration, and adhesion (51). Activation of overall PKC leads to activation of MAP kinase; PKCalpha , at least, has been shown to phosphorylate Raf kinase (40), an upstream mediator of MAP kinase (36). In our studies, both PKCalpha and PKCdelta enhance ERK-1 and ERK-2 kinase activity with a resultant prolongation of ERK-2 activity in stably transfected endothelial cells. The pathway by which PKCdelta blocks proliferation and cell cycle progression, however, must be distal or parallel to that leading to ERK-2 or ERK-1. In addition, our results suggest that effective activation of ERK-2 activity by PKCdelta is not sufficient for endothelial cell proliferation. This finding bolsters those of Hirai et al. (25), who found that PKCdelta inhibited growth of Ras-transformed NIH 3T3 cells despite activating AP-1, a component of the signaling pathway downstream from MAP kinase.

Inhibition of endothelial cell proliferation by PKCdelta appears to result from a specific defect in endothelial cell cycle progression, in which the cells enter S phase inappropriately and require additional time to complete this phase. Because of the distal nature of this defect and the intranuclear location of PKCdelta , it seems likely that the isoenzyme interacts directly with one of the cyclin-cyclin-dependent kinase/inhibitor complexes that regulates entry and completion of S phase. Inappropriate S phase entry has been associated with apoptosis of vascular smooth muscle cells treated with basic fibroblast growth factor antisense oligonucleotides (52), but such apoptosis was not found in PKCdelta EC.2 Manipulation of PKC activity has not been previously associated with this S phase defect and only rarely with any abnormality of cell cycle function. Overexpression of PKCdelta , but not PKCalpha or PKCzeta , in Chinese hamster ovary fibroblasts causes an arrest in G2/M phase of the cell cycle, but only after activation with PMA (21); no defects were seen in S phase. In that study, inhibition of cell cycle progression by PKCdelta was attributed to an isoenzyme-specific effect; since the PKC enzymatic activity was much higher in the PKCdelta -transfected cells than in the other transfectants, however, it is possible that the observed effect merely reflected an increase in overall enzyme activity. In this study, however, the similarities of enzymatic activity make it likely that the defect in cell cycle progression, as well as differences in adhesion and migration, resulted from interaction of PKCdelta with isoenzyme-specific substrates. Thus, in addition to its isoenzyme-specific character, this arrest in S phase appears to be somewhat specific for endothelial cells.

Specific effects of individual PKC isoenzymes on proliferation are known to vary according to cell types. For example, overexpression of PKCalpha in murine fibroblast cells inhibits proliferation and does not lead to cell transformation (53, 54), but in NIH 3T3 and human breast cancer cell lines, increases in PKCalpha expression altered the cell morphology, enhanced proliferation, and increased tumorigenicity (23, 55). Thus, the tissue-specific effects of individual PKC isoenzymes are likely to be mediated by substrates or effectors with restricted tissue or subcellular expression. Even within an individual cell type, the substrates with which each isoenzyme interacts differ. A full understanding of the mechanism by which these two individual PKC isoenzymes mediate either enhanced adhesion or migration or decrease proliferation of endothelial cells will require identification of their selective downstream targets. Such identification, together with the assignment of selective endothelial functions to individual PKC isoenzymes provided by this study, would provide essential details critical to our understanding of reendothelialization and angiogenesis.

Acknowledgments

We thank Malay Raychowdhury, James D. Chang, and Masao Yukawa for technical assistance and advice.


FOOTNOTES

*   This work was supported by National Institutes of Health (NIH) Grants HL 38820, HL 47032, and HL51043 (to J. A. W.), NIH Grant HL 53793 (to M. S.), American Heart Association Grant GIA 95007560 (to M. S. and E. O. H.), and an NIH National Research Service Award (to E. O. H.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Dagger    To whom correspondence should be addressed: Cardiovascular Division, Albert Einstein College of Medicine, 1300 Morris Park Ave., Forch. Bldg. G-42, Bronx, NY 10461. Tel.: 718-430-3087; Fax: 718-430-8989; E-mail: jaware{at}aecom.yu.edu.
1   The abbreviations used are: PKC, protein kinase C; RFPEC, rat fat pad epididymal endothelial cell(s); EC, endothelial cell(s); PBS, phosphate-buffered saline; BSA, bovine serum albumin; MAP, mitogen-activated protein; ERK, extracellular signal-regulated kinase; PMA, phorbol 12-myristate 13-acetate; HGF, hepatocyte growth factor.
2   E. O. Harrington and J. A. Ware, unpublished observations.

REFERENCES

  1. Yang, E. Y., and Moses, H. L. (1990) J. Cell Biol. 111, 731-741 [Abstract]
  2. Ausprunk, D. H., and Folkman, J. (1977) Microvasc. Res. 14, 53-65 [Medline] [Order article via Infotrieve]
  3. Camussi, G., Montrucchio, G., Lupia, E., De Martino, A., Perona, L., Arese, M., Vercellone, A., Toniolo, A., and Bussolino, F. (1995) J. Immunol. 154, 6492-6501 [Abstract/Free Full Text]
  4. Grant, D. S., Kleinman, H. K., Goldberg, I. D., Bhargava, M. M., Nickoloff, B. J., Kinsella, J. L., Polverini, P., and Rosen, E. M. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 1937-1941 [Abstract]
  5. Kent, K. C., Mii, S., Harrington, E. O., Chang, J. D., Mallette, S., and Ware, J. A. (1995) Circ. Res. 77, 231-238 [Abstract/Free Full Text]
  6. Kinsella, J. L., Grant, D. S., Weeks, B. S., and Kleinman, H. K. (1992) Exp. Cell Res. 199, 56-62 [Medline] [Order article via Infotrieve]
  7. Montesano, R., and Orci, L. (1985) Cell 42, 469-477 [Medline] [Order article via Infotrieve]
  8. Hu, D. E., and Fan, T. P. (1995) Inflammation 19, 39-54 [Medline] [Order article via Infotrieve]
  9. Wright, P. S., Cross-Doersen, D., Miller, J. A., Jones, W. D., and Bitonti, A. J. (1992) J. Cell. Physiol. 152, 448-457 [Medline] [Order article via Infotrieve]
  10. Presta, M., Tiberio, L., Rusnati, M., Dell'Era, P., and Ragnotti, G. (1991) Cell Regul. 2, 719-26 [Medline] [Order article via Infotrieve]
  11. Daviet, I., Herbert, J. M., and Maffrand, J. P. (1990) FEBS Lett. 259, 315-317 [CrossRef][Medline] [Order article via Infotrieve]
  12. Lynch, J. J., Ferro, T. J., Blumenstock, F. A., Brockenauer, A. M., and Malik, A. B. (1990) J. Clin. Invest. 85, 1991-1998 [Medline] [Order article via Infotrieve]
  13. Nagpala, P. G., Malik, A. B., Vuong, P. T., and Lum, H. (1996) J. Cell Physiol. 166, 249-255 [CrossRef][Medline] [Order article via Infotrieve]
  14. Langer, D. J., Kuo, A., Kariko, K., Ahuja, M., Klugherz, B. D., Ivanics, K. M., Hoxie, J. A., Williams, W. V., Liang, B. T., Cines, D. B., et al. (1993) Circ. Res. 72, 330-340 [Abstract]
  15. Murphy, H. S., Maroughi, M., Till, G. O., and Ward, P. A. (1994) Am. J. Physiol. 267, L145-L151 [Abstract/Free Full Text]
  16. Ahmed, A., Plevin, R., Shoaibi, M. A., Fountain, S. A., Ferriani, R. A., and Smith, S. K. (1994) Am. J. Physiol. 266, C206-C212 [Abstract/Free Full Text]
  17. Newton, A. C. (1995) J. Biol. Chem. 270, 28495-28498 [Free Full Text]
  18. Nishizuka, Y. (1992) Science 258, 607-614 [Medline] [Order article via Infotrieve]
  19. Eldar, H., Zisman, Y., Ullrich, A., and Livneh, E. (1990) J. Biol. Chem. 265, 13290-13296 [Abstract/Free Full Text]
  20. Mischak, H., Goodnight, J., Kolch, W., Martiny-Baron, G., Schaechtle, C., Kazanietz, M. G., Blumberg, P. M., Pierce, J. H., and Mushinski, J. F. (1993) J. Biol. Chem. 268, 6090-6096 [Abstract/Free Full Text]
  21. Watanabe, T., Ono, Y., Taniyama, Y., Hazama, K., Igarashi, K., Ogita, K., Kikkawa, U., and Nishizuka, Y. (1992) Proc. Natl. Acad. Sci. U. S. A. 89, 10159-10163 [Abstract]
  22. Kindregan, H. C., Rosenbaum, S. E., Ohno, S., and Niles, R. M. (1994) J. Biol. Chem. 269, 27756-27761 [Abstract/Free Full Text]
  23. Persons, D. A., Wilkison, W. O., Bell, R. M., and Finn, O. J. (1988) Cell 52, 447-458 [Medline] [Order article via Infotrieve]
  24. Ways, D. K., Kukoly, C. A., deVente, J., Hooker, J. L., Bryant, W. O., Posekany, K. J., Fletcher, D. J., Cook, P. P., and Parker, P. J. (1995) J. Clin. Invest. 95, 1906-1915 [Medline] [Order article via Infotrieve]
  25. Hirai, S., Izumi, Y., Higa, K., Kaibuchi, K., Mizuno, K., Osada, S., Suzuki, K., and Ohno, S. (1994) EMBO J. 13, 2331-2340 [Abstract]
  26. Kielbassa, K., Muller, H.-J., Meyer, H. E., Marks, F., and Gschwendt, M. (1995) J. Biol. Chem. 270, 6156-6162 [Abstract/Free Full Text]
  27. Kojima, T., Leone, C. W., Marchildon, G. A., Marcum, J. A., and Rosenberg, R. D. (1992) J. Biol. Chem. 267, 4859-4869 [Abstract/Free Full Text]
  28. Marcum, J. A., and Rosenberg, R. D. (1985) Biochem. Biophys. Res. Commun. 126, 365-372 [Medline] [Order article via Infotrieve]
  29. Brooks, P. C., Clark, R. A., and Cheresh, D. A. (1994) Science 264, 569-571 [Medline] [Order article via Infotrieve]
  30. Tennenbaum, T., Giloh, H., Fusenig, N. E., and Kapitulnik, J. (1988) J. Invest. Dermatol. 90, 857-860 [Abstract]
  31. Goodnight, J., Mischak, H., Kolch, W., and Mushinski, J. F. (1995) J. Biol. Chem. 270, 9991-10001 [Abstract/Free Full Text]
  32. Bussolino, F., Di Renzo, M. F., Ziche, M., Bocchietto, E., Olivero, M., Naldini, L., Gaudino, G., Tamagnone, L., Coffer, A., and Comoglio, P. M. (1992) J. Cell Biol. 119, 629-641 [Abstract]
  33. Morimoto, A., Okamura, K., Hamanaka, R., Sato, Y., Shima, N., Higashio, K., and Kuwano, M. (1991) Biochem. Biophys. Res. Commun. 179, 1042-1049 [Medline] [Order article via Infotrieve]
  34. Brooks, P. C., Montgomery, A. M., Rosenfeld, M., Reisfeld, R. A., Hu, T., Klier, G., and Cheresh, D. A. (1994) Cell 79, 1157-1164 [Medline] [Order article via Infotrieve]
  35. Friedlander, M., Brooks, P. C., Shaffer, R. W., Kincaid, C. M., Varner, J. A., and Cheresh, D. A. (1995) Science 270, 1500-1502 [Abstract]
  36. Marquardt, B., Frith, D., and Stabel, S. (1994) Oncogene 9, 3213-3218 [Medline] [Order article via Infotrieve]
  37. Tseng, H., Peterson, T. E., and Berk, B. C. (1995) Circ. Res. 77, 869-878 [Abstract/Free Full Text]
  38. Bussolino, F., Silvagno, F., Garbarino, G., Costamagna, C., Sanavio, F., Arese, M., Soldi, R., Aglietta, M., Pescarmona, G., Camussi, G., and Bosia, A. (1994) J. Biol. Chem. 269, 2877-2886 [Abstract/Free Full Text]
  39. Kosaka, C., Sasaguri, T., Zen, K., Masuda, J., Shimokado, K., and Ogata, J. (1995) Ann. N. Y. Acad. Sci. 748, 538-540 [Medline] [Order article via Infotrieve]
  40. Sozeri, O., Vollmer, K., Liyanage, M., Frith, D., Kour, G., Mark, G. E. d., Stabel, S., and Mark, G. E. (1992) Oncogene 7, 2259-2262 [Medline] [Order article via Infotrieve]
  41. Zhou, W., Takuwa, N., Kumada, M., and Takuwa, Y. (1993) J. Biol. Chem. 268, 23041-23048 [Abstract/Free Full Text]
  42. Basson, C. T., Knowles, W. J., Bell, L., Albelda, S. M., Castronovo, V., Liotta, L. A., and Madri, J. A. (1990) J. Cell Biol. 110, 789-801 [Abstract]
  43. Tang, D. G., Diglio, C. A., and Honn, K. V. (1993) Prostaglandins 45, 249-267 [Medline] [Order article via Infotrieve]
  44. Maher, J. J. (1993) J. Clin. Invest. 91, 2244-2252 [Medline] [Order article via Infotrieve]
  45. Nakamura, Y., Morishita, R., Higaki, J., Kida, I., Aoki, M., Moriguchi, A., Yamada, K., Hayashi, S., Yo, Y., Matsumoto, K., et al. (1995) Biochem. Biophys. Res. Commun. 215, 483-488 [CrossRef][Medline] [Order article via Infotrieve]
  46. Leavesley, D. I., Schwartz, M. A., Rosenfeld, M., and Cheresh, D. A. (1993) J. Cell Biol. 121, 163-170 [Abstract]
  47. Liaw, L., Lindner, V., Schwartz, S. M., Chambers, A. F., and Giachelli, C. M. (1995) Circ. Res. 77, 665-672 [Abstract/Free Full Text]
  48. Clark, E. A., Shattil, S. J., and Brugge, J. S. (1994) Trends Biochem. Sci. 19, 464-469 [CrossRef][Medline] [Order article via Infotrieve]
  49. Haimovich, B., Kaneshiki, N., and Ji, P. (1996) Blood 87, 152-161 [Abstract/Free Full Text]
  50. Saitoh, M., Salzman, E. W., Smith, M., and Ware, J. A. (1989) Blood 74, 2001-2006 [Abstract]
  51. Sa, G., Murugesan, G., Jaye, M., Ivashchenko, Y., and Fox, P. L. (1995) J. Biol. Chem. 270, 2360-2366 [Abstract/Free Full Text]
  52. Fox, J. C., and Shanley, J. R. (1996) J. Biol. Chem. 271, 12578-12584 [Abstract/Free Full Text]
  53. Borner, C., Filipuzzi, I., Weinstein, I. B., and Imber, R. (1991) Nature 353, 78-80 [CrossRef][Medline] [Order article via Infotrieve]
  54. Borner, C., Ueffing, M., Jaken, S., Parker, P. J., and Weinstein, I. B. (1995) J. Biol. Chem. 270, 78-86 [Abstract/Free Full Text]
  55. Finkenzeller, G., Marme, D., and Hug, H. (1992) Cell. Signalling 4, 163-177 [CrossRef][Medline] [Order article via Infotrieve]

©1997 by The American Society for Biochemistry and Molecular Biology, Inc.