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The Apolipoprotein(a) Component of Lipoprotein(a) Stimulates Actin Stress Fiber Formation and Loss of Cell-Cell Contact in Cultured Endothelial Cells*

      Elevated plasma concentrations of lipoprotein(a) (Lp(a)) are a risk factor for a variety of atherosclerotic disorders including coronary heart disease. In the current study, we report that incubation of cultured human umbilical vein or coronary artery endothelial cells with Lp(a) elicits a dramatic rearrangement of the actin cytoskeleton characterized by increased central stress fiber formation and redistribution of focal adhesions. These effects are mediated by the apolipoprotein(a) (apo(a)) component of Lp(a) since incubation of apo(a) with the cells evoked similar cytoskeletal rearrangements, while incubation with low density lipoprotein had no effect. Apo(a) also produced a time-dependent increase in transendothelial permeability. The cytoskeletal rearrangements evoked by apo(a) were abolished by C3 transferase, which inhibits Rho, and by Y-27632, an inhibitor of Rho kinase. In addition to actin cytoskeleton remodeling, apo(a) was found to cause VE-cadherin disruption and focal adhesion molecule reorganization in a Rho- and Rho kinase-dependent manner. Cell-cell contacts were found to be regulated by Rho and Rac but not Cdc42. Apo(a) caused a transient increase in the extent of myosin light chain phosphorylation. Finally apo(a) did not evoke increases in intracellular calcium levels, although the effects of apo(a) on the cytoskeleton were found to be calcium-dependent. We conclude that the apo(a) component of Lp(a) activates a Rho/Rho kinase-dependent intracellular signaling cascade that results in increased myosin light chain phosphorylation with attendant rearrangements of the actin cytoskeleton. We propose that the resultant increase in endothelial permeability caused by Lp(a) may help explain the atherosclerotic risk posed by elevated concentrations of this lipoprotein.
      The vascular endothelium acts as a pivotal regulator in vessel wall homeostasis by forming a selective barrier between components of the blood and extravascular tissues. Increasing evidence suggests that atypical endothelial function is a key event in the initial stages of atherosclerosis development. More specifically, enhanced endothelial cell permeability and the expression of a procoagulant, antifibrinolytic, and proinflammatory phenotype by the endothelium is thought to be a crucial event in the onset of this disease (
      • Poredos P.
      ). Various physiological agents have been identified that elicit some manifestations of endothelial dysfunction. These include growth factors, inflammatory cytokines such as tumor necrosis factor-α (
      • Wojciak-Stothard B.
      • Entwistle A.
      • Garg R.
      • Ridley A.J.
      ), vasoactive substances such as thrombin (
      • Lum H.
      • Ashner J.L.
      • Phillips P.G.
      • Fletcher P.W.
      • Malik A.B.
      ), and mildly oxidized, but not native, low density lipoprotein (LDL)
      The abbreviations used are: LDL, low density lipoprotein; MLC, myosin light chain; Lp(a), lipoprotein(a); apo(a), apolipoprotein(a); HUVEC, human umbilical vein endothelial cell; HCAEC, human coronary artery endothelial cell; BAPTA-AM, 1,2-bis(2-aminophenoxy)ethane-N,N,N′,N′-tetraacetic acid tetrakis (acetoxymethyl ester); PBS, phospate-buffered saline; TRITC, tetramethylrhodamine isothiocyanate; PIPES, 1,4-piperazinediethanesulfonic acid; TUNEL, terminal deoxynucleotidyl transferase dUTP nick-end labeling.
      1The abbreviations used are: LDL, low density lipoprotein; MLC, myosin light chain; Lp(a), lipoprotein(a); apo(a), apolipoprotein(a); HUVEC, human umbilical vein endothelial cell; HCAEC, human coronary artery endothelial cell; BAPTA-AM, 1,2-bis(2-aminophenoxy)ethane-N,N,N′,N′-tetraacetic acid tetrakis (acetoxymethyl ester); PBS, phospate-buffered saline; TRITC, tetramethylrhodamine isothiocyanate; PIPES, 1,4-piperazinediethanesulfonic acid; TUNEL, terminal deoxynucleotidyl transferase dUTP nick-end labeling.
      (
      • Essler M.
      • Retzer M.
      • Bauer M.
      • Heemsker J.W.
      • Aepfelbacher M.
      • Seiss W.
      ).
      Many of these agents alter the permeability of the endothelium by stimulating cell contraction, thereby increasing the size of intercellular gaps and facilitating entry of inflammatory cells and atherogenic lipoproteins. The mechanism by which these agents are able to alter properties of the endothelium has received much attention. Endothelial cell contraction is mediated by interactions between actin filaments and non-muscle myosin II. The activity of non-muscle myosin II is largely regulated by phosphorylation of its regulatory myosin light chain (MLC) at Ser-19 (
      • Garcia J.G.
      • Davis H.W.
      • Patterson C.E.
      ). MLC phosphorylation activates myosin thus enabling it to associate with filamentous actin (F-actin), resulting in the assembly of stress fibers, the formation of mature focal adhesions, and cell contraction (
      • Goeckler Z.M.
      • Wysolmerski R.B.
      ,
      • Chrzanowska-Wodnicka M.
      • Burridge K.
      ). The MLC phosphorylation state is a function of the balance between MLC kinases (principally the Ca2+-calmodulin-dependent myosin light chain kinase) and phosphatases (principally type 1 myosin-associated protein phosphatase) (
      • Verin A.D.
      • Patterson C.E.
      • Day M.A.
      • Garcia J.G.
      ,
      • Somlyo A.P.
      • Somlyo A.V.
      ). Rho kinase, an effector of the small GTPase Rho, has both direct and indirect effects on MLC phosphorylation (
      • Amano M.
      • Ito M.
      • Kimura K.
      • Fukata Y.
      • Chihara K.
      • Nakano T.
      • Matsuura M.
      • Kaibuchi K.
      ). Rho kinase has been found to directly phosphorylate the MLC at Ser-19 albeit at a much lower rate than MLC kinase. In addition, Rho kinase effectively enhances MLC phosphorylation through inactivation of myosin phosphatase (
      • Kimura K.
      • Ito M.
      • Amano M.
      • Chihara K.
      • Fukata Y.
      • Nakafuku M.
      • Yamamori B.
      • Feng J.
      • Nakano T.
      • Okawa K.
      • Iwamatsu A.
      • Kaibuchi K.
      ,
      • Kawano Y.
      • Fukata Y.
      • Oshiro N.
      • Amano M.
      • Nakamura T.
      • Ito M.
      • Matsumura F.
      • Inagaki M.
      • Kaibuchi K.
      ). It has been demonstrated that the ability of tumor necrosis factor-α and mildly oxidized LDL to elicit stress fiber formation, increase MLC phosphorylation, and promote retraction in cultured endothelial cells is mediated by a Rho- and Rho kinase-dependent signaling pathway (
      • Wojciak-Stothard B.
      • Entwistle A.
      • Garg R.
      • Ridley A.J.
      ,
      • Essler M.
      • Retzer M.
      • Bauer M.
      • Heemsker J.W.
      • Aepfelbacher M.
      • Seiss W.
      ).
      In addition to Rho, the GTP-binding proteins Rac and Cdc42 also play central roles in controlling cytoskeletal reorganization. While Rho is the predominant regulator of actin stress fiber formation (
      • Ridley A.J.
      • Hall A.
      ), Rac and Cdc42 are involved in lamellipodia and filopodia formation, respectively, and promote the assembly of small, peripheral adhesion complexes (
      • Bishop A.L.
      • Hall A.
      ). Members of the Rho family of GTPases have also been found to regulate intercellular junctions and thus permeability (
      • Wojciak-Stothard B.
      • Entwistle A.
      • Garg R.
      • Ridley A.J.
      ,
      • Nobes C.D.
      • Hall A.
      ,
      • Wojciak-Stothard B.
      • Potempa S.
      • Eichholtz T.
      • Ridley A.J.
      ).
      Both case-control and prospective studies have shown that lipoprotein(a) (Lp(a)) is a risk factor for coronary heart disease (
      • Marcovina S.M.
      • Koschinsky M.L.
      ,
      • Marcovina S.M.
      • Koschinsky M.L.
      ,
      • Scanu A.M.
      ). Lp(a) is similar to LDL in terms of lipid composition and the presence of the apolipoprotein B-100 moiety. However, Lp(a) is clearly distinguishable from LDL by the presence of the unique glycoprotein apolipoprotein(a) (apo(a)) that is covalently linked to apolipoprotein B-100 by a single disulfide bond (
      • Brunner C.
      • Kraft H.G.
      • Utermann G.
      • Muller H.J.
      ,
      • Koschinsky M.L.
      • Côté G.P.
      • Gabel B.
      • van der Hoek Y.Y.
      ). Apo(a) shares extensive homology with the fibrinolytic zymogen plasminogen and contains multiple repeats of a sequence that is similar to plasminogen kringle IV as well as a single copy of sequences similar to the kringle V and protease regions of plasminogen (
      • McLean J.W.
      • Tomlinson J.E.
      • Kuang W.
      • Eaton D.L.
      • Chen E.
      • Fless G.
      • Scanu A.
      • Lawn R.M.
      ). Although Lp(a) has been recognized as a risk factor for vascular diseases, the nature of the role of Lp(a) in atherosclerosis remains unclear. Potential roles for Lp(a) in lipid deposition and smooth muscle cell proliferation in developing lesions and inhibition of fibrinolysis as well as stimulation of endothelial dysfunction have been documented (
      • Marcovina S.M.
      • Koschinsky M.L.
      ). However, the effect of Lp(a) on endothelial cell morphology and barrier function has yet to be assessed.
      In the present study, we have examined the role of the apo(a) component of Lp(a) in cultured endothelial cell cytoskeleton reorganization. Our findings indicate the following. (i) Apo(a) induces F-actin stress fiber formation, vascular endothelial cadherin (VE-cadherin) dispersion, and focal adhesion molecule reorganization in a Rho- and Rho kinase-dependent manner. (ii) Rho and Rac but not Cdc42 are implicated in apo(a)-mediated actin stress fiber formation and VE-cadherin disruption. (iii) Apo(a) induces a time-dependent increase in MLC phosphorylation that is regulated by Rho and Rho kinase. (iv) Apo(a) does not result in increases in cytosolic calcium levels, but its effects on cytoskeletal rearrangement are calcium-dependent. Taken together, these results suggest a novel mechanism by which Lp(a) can promote endothelial cell dysfunction during atherogenesis.

      EXPERIMENTAL PROCEDURES

      Purification of Proteins—Recombinant apo(a) containing 17 kringle IV-like domains and corresponding to a physiologically relevant isoform size (
      • Koschinsky M.L.
      • Tomlinson J.E.
      • Zioncheck T.F.
      • Schwartz K.
      • Eaton D.L.
      • Lawn R.M.
      ) was purified from conditioned medium harvested from human embryonic kidney (HEK 293) cells stably transfected with the corresponding expression plasmid using lysine-Sepharose affinity chromatography as described previously (
      • Sangrar W.
      • Gabel B.R.
      • Boffa M.B.
      • Walker J.B.
      • Hancock M.A.
      • Marcovina S.M.
      • Horrevoets A.J.G.
      • Nesheim M.E.
      • Koschinsky M.L.
      ). Human plasminogen was purified from fresh-frozen plasma using lysine-Sepharose affinity chromatography as described previously (
      • Castellino F.J
      • Powell J.R.
      ). Human Lp(a) was purified from fresh plasma by sequential ultracentrifugation and gel filtration chromatography as described previously (
      • Edelstein C.
      • Hinman J.
      • Marcovina S.
      • Scanu A.M.
      ). Human LDL was purified from fresh plasma by sequential flotation as described previously (
      • Gabel B.R.
      • McLeod R.S.
      • Yao Z.
      • Koschinsky M.L.
      ).
      Cell Culture—Human umbilical vein endothelial cells (HUVECs) and human coronary artery endothelial cells (HCAECs) were obtained from Clonetics and grown in endothelial growth medium EGM-2 (Clonetics), which contained 2% fetal calf serum unless otherwise indicated. Cells were fed every 2nd day and used at passages 2–5.
      Immunofluorescence—For double immunofluorescence studies, cells were plated onto gelatin-precoated (1 h, 0.1% gelatin (Fisher Scientific) at 37 °C) glass coverslips at a density of 20,000 cells/well in 24-well tissue culture dishes and grown to near confluence. Before treatment with apo(a) and Lp(a), cells were serum-starved for 15 min at 37 °C in physiological saline (145 mm NaCl, 5 mm KCl, 1 mm MgSO4, 1 mm CaCl2, 10 mm HEPES, 10 mm glucose, pH 7.4, containing 0.5% bovine serum albumin). This solution was then replaced with fresh physiological saline containing 400 nm apo(a), 400 nm Lp(a), 100 μg/ml LDL, or 400 nm plasminogen, and the cells were incubated at 37 °C for different time periods. In some experiments, cells were pretreated with C3 transferase from Clostridium botulinum (Cytoskeleton, Inc.) or Y-27632 (Calbiochem-Novabiochem). For C3 transferase, the enzyme was included in the complete culture medium at a concentration of 5 μg/ml for 24 h prior to the 15-min serum starvation. For Y-27632, the cells were serum-starved for 15 min as described above and then were treated for 30 min with Y-27632 at a final concentration of 10 μm in physiological saline; at this time, the solution was removed, and fresh solution containing apo(a) or Lp(a) was added. In other experiments, cells were serum-starved for 15 min and then pretreated with BAPTA-AM (Sigma) or ML-7 (Calbiochem) (25 μm, 1 h) prior to apo(a) treatment. Cells were then prepared for double immunofluorescence as follows. Cells were fixed with 3.7% paraformaldehyde solution in phosphate-buffered saline (PBS) for 5 min, washed once with PBS, fixed and permeabilized with 1.4% formaldehyde containing 0.1% Nonidet P-40 for 1.5 min, and then washed three times with PBS. For F-actin staining, cells were incubated with TRITC-phalloidin diluted 1:100 in saponin buffer (0.1% saponin, 20 mm KPO4, 10mm PIPES, 5 mm EGTA, 2 mm MgCl2, pH 6.8). For vinculin staining, cells were first blocked with 0.1% bovine serum albumin for 20 min at room temperature. Coverslips were then incubated with anti-vinculin monoclonal antibody (Sigma) diluted 1:300 in saponin buffer for 1 h at room temperature. Following three washes with PBS, cells were stained for 1 h with 1:500-diluted goat anti-mouse Alexa488-conjugated antibody (Molecular Probes) and TRITC-phalloidin (Sigma) in saponin buffer. For VE-cadherin staining, cells were treated as indicated for vinculin except using 1:350-diluted anti-VE-cadherin monoclonal antibody (Research Diagnostics). Coverslips were mounted to slides using an anti-fade mounting solution (Dako) and examined using a Zeiss Axiovert S100 inverted fluorescence microscope equipped with a 40× oil immersion lens. Images were captured using a high sensitivity Cooke SensiCam and SlideBook software (Intelligent Imaging Innovations Inc.).
      Apoptosis Detection (TUNEL Assay)—HUVECs were grown on glass coverslips as described above. Apoptosis was examined using an in situ cell death assay kit (Roche Applied Science) according to the manufacturer's instructions. As a positive control, 0.1 mg/ml DNase I was added for 15 min at room temperature to non-starved cells that were previously fixed and permeabilized.
      Transendothelial Permeability Assay—HUVECs were seeded at a final concentration of 25,000 cells/ml onto Transwell inserts (3.0-μm pore size; BD Biosciences) precoated with 7 μg/ml fibronectin (1 h, 37 °C) and placed into 24-well Transwell companion plates (BD Biosciences) containing 600 μl of EGM-2 medium. Cells were grown for 3 days with one change of medium before the experiment. To evaluate the effect of apo(a) on endothelial permeability, medium in the top well was replaced with EGM-2 containing 400 nm apo(a) and 1 mg/ml fluorescein isothiocyanate-dextran (molecular weight, 40,000) in a final volume of 100 μl. The medium in the bottom well was replaced with 600 μl of fresh EGM-2. Control cells were treated identically but in the absence of apo(a). At specific time points, 50 μl of medium from the bottom well was removed and replaced with 50 μl of fresh EGM-2. The 50-μl sample was then diluted with 950 μl of PBS, and fluorescence intensity was evaluated with a fluorometer (PerkinElmer Life Sciences LS-50B) using an excitation wavelength of 492 nm and emission wavelength of 520 nm.
      Transient Transfections—The expression plasmids pRK5mycN19RhoA (encoding a Myc-tagged, dominant negative RhoA), pRK5mycN17Rac (encoding a Myc-tagged, dominant negative Rac), pMT90mycN17Cdc-42 (encoding a Myc-tagged, dominant negative Cdc42), and pMT90V12-Cdc42 (encoding a constitutively active Cdc42) were the kind gifts of Dr. Alan Hall (University College, London). Each expression plasmid contains the cytomegalovirus promoter. HUVECs were seeded on 0.1% gelatin-precoated glass coverslips at 100,000–200,000 cells/ml. The following day, cells were transfected with 0.4 μg of the respective plasmids using the LipofectAMINE Plus transfection kit (Invitrogen). Briefly, 0.4 μg of plasmid and 2 μl of Plus reagent in a final volume of 25 μl of serum- and antibiotic-free EGM-2(-) medium and, separately, 1 μl of LipofectAMINE in a final volume of 25 μl of EGM-2(-) medium were incubated at room temperature for 15 min; the solutions were then combined and incubated for an additional 15 min. The final transfection reaction was added to a well of HUVECs grown in 200 μl of EGM-2(-) medium. Transfection proceeded for 3 h at 37 °C after which cells were washed three times with EGM-2(-) medium and fed complete EGM-2. The following day, cells were fixed and permeabilized as before. To detect transfected cells, HUVECs were stained with anti-c-Myc polyclonal antibody (Santa Cruz Biotechnology) diluted 1:300 in saponin buffer and washed three times using PBS. Coverslips were then incubated for 1 h with 1:200-diluted goat anti-rabbit Alexa350 secondary antibody (Molecular Probes).
      Myosin Light Chain Phosphorylation—MLC phosphorylation was analyzed by SDS-PAGE followed by Western blotting. HUVECs were grown to confluence in 6-well culture dishes (precoated with 0.1% gelatin) and treated with 400 nm apo(a) for different times. In some experiments, cells were pretreated with C3 transferase or Y-27632 as described above prior to treatment with apo(a). The reactions were terminated by immediate addition of 1.5 ml of ice-cold 10% trichloroacetic acid. Cells were scraped into microcentrifuge tubes and then centrifuged for 20 min at 14,000 × g. Supernatants were discarded, and pellets were washed twice with water to remove residual trichloroacetic acid. Resulting pellets were resuspended in 1% SDS and then sonicated for 4–5 h. Samples were subjected to SDS-PAGE on a 15% polyacrylamide gel, and resolved proteins were transferred to Immobilon P (Millipore) in Tris-glycine buffer (25 mm Tris, 192 mm glycine, 0.02% SDS). Membranes were blocked with 5% skim milk powder for 2 h at 4 °C, washed twice in Tween-Tris-buffered saline (TTBS; 0.05% Tween 20, 20 mm Tris, pH 7.4, 0.15 m NaCl), and probed for either phosphorylated MLC (Ser-19) with 1 μg/ml anti-phospho-MLC antibody (pS19, a generous gift from BIOSOURCE) or for total MLC with 1:100-diluted total MLC antibody (A-20, Santa Cruz Biotechnology) at 4 °C. Membranes were then washed three times with TTBS and incubated with 1:3000 dilutions of the appropriate horseradish peroxidase-conjugated secondary antibodies (Bio-Rad) in 5% milk for 2 h at 4 °C. Finally, membranes were developed with enhanced chemiluminescence Western blotting detection reagents (Amersham Biosciences) and exposed to x-ray film. Blots were scanned using a flatbed laser scanner, and the density of the immunoreactive bands was determined using Corel Photopaint Version 8. The amount of phosphorylated MLC was normalized to the total MLC signal in the respective samples.
      Detection of Intracellular Calcium Transients—HUVECs were seeded on glass coverslips precoated with 0.1% gelatin (1 h, 37 °C) at 25,000 cells/ml and grown for 3 days. Cells were loaded with 5 μm Fura-2/AM (Sigma) for 45 min at 37 °C, washed five times, and incubated in HEPES buffer (20 mm HEPES, 120 mm NaCl, 2.7 mm KCl, 1.4 mm MgSO4, 0.5 mm CaCl2, 1.4 mm KH2PO4, 25 mm NaHCO3, 10 mm glucose, pH 7.4). Coverslips were submerged in HEPES buffer contained in a heated mounting dish and maintained at 37 °C. Cells were allowed to equilibrate in the dish for 5 min. Calcium imaging was captured every 9 s using SlideBook software. Apo(a) was added at a final concentration of 400 nm after 18 s. As a positive control, 1 unit/ml thrombin was added at the end of the calcium imaging. Calcium-dependent fluorescence was detected using a Zeiss Axiovert S100 inverted fluorescent microscope using excitation wavelengths of 340 nm and 380 nm and an emission wavelength of 510 nm. Intracellular calcium concentration was expressed as the ratio of fluorescence intensities at 510 nm corresponding to calcium bound (340 nm) and calcium unbound (380 nm) excitation wavelengths (ratio 340/380).

      RESULTS

      Lp(a), through Its Apo(a) Moiety, Causes a Time-dependent Increase in F-actin Stress Fiber Formation and Transendothelial Permeability—Following a brief 15-min serum starvation, confluent HUVECs displayed few actin stress fibers (Fig. 1A). F-actin was concentrated along the cell borders and was generally absent in the central regions of the cell. Treatment with 400 nm apo(a) caused a dramatic increase in the number of F-actin stress fibers traversing the cells. Stress fiber formation was apparent within 5 min of stimulation and appeared to saturate between 10 and 20 min. Cell retraction, accompanied by increased gap formation between the cells, occurred as early as 2 min after apo(a) treatment. The effects of Lp(a) treatment on HUVEC F-actin organization were similar to that observed for apo(a). The appearance of stress fibers was observable after 2 min of stimulation. In addition, maximal stress fiber formation occurred between 5 and 10 min after which time the number of stress fibers appeared to diminish. Thus, Lp(a) appeared to be slightly more potent than apo(a) in eliciting actin stress fiber formation. The risk threshold for plasma concentrations of Lp(a) is taken to be 30 mg/dl or ∼40 nm for an Lp(a) isoform of the size utilized in these studies. Although plasma concentrations of Lp(a) of 400 nm are rare in the population (
      • Marcovina S.M.
      • Albers J.J.
      • Wijsman E.
      • Zhang Z.
      • Chapman N.H.
      • Kennedy H.
      ), we have observed stress fiber stimulation in HUVECs by apo(a) concentrations as low as 25 nm (data not shown).
      Figure thumbnail gr1
      Fig. 1The apo(a) component of Lp(a) causes a time-dependent increase in F-actin stress fibers. A, HUVECs were serum-starved for 15 min and then treated with 400 nm Lp(a) or 400 nm apo(a) for the indicated time periods. Cells were then fixed, permeabilized, and stained for F-actin using TRITC-phalloidin. B, HUVECs were serum-starved for 15 min and then treated with 400 nm apo(a), 100 μg/ml LDL, or 400 nm plasminogen for 5 min. Cells were then fixed, permeabilized, and stained for F-actin using TRITC-phalloidin.
      To determine whether the LDL moiety of Lp(a) had an influence on stress fiber formation, HUVECs were treated with 100 μg/ml LDL for 5 min. LDL produced no detectable change in F-actin organization (Fig. 1B), an observation that is consistent with previous findings (
      • Essler M.
      • Retzer M.
      • Bauer M.
      • Heemsker J.W.
      • Aepfelbacher M.
      • Seiss W.
      ). These data indicate that the moiety on Lp(a) that mediates the effects of this lipoprotein on actin cytoskeletal rearrangements is apo(a). Plasminogen (a kringle-containing protein homologous to apo(a)) was tested to examine whether the effects of apo(a) on the actin cytoskeleton were specific for the kringles in apo(a). Treatment of HUVECs with 400 nm plasminogen for 5 min produced no significant increase in stress fiber formation (Fig. 1B).
      Previous results have implicated Rho proteins in endothelial cell apoptotic cell death (
      • Hippenstiel S.
      • Schmeck B.
      • N′Guessan P.D.
      • Seybold J.
      • Krull M.
      • Preissner K.
      • Eichel-Streiber C.V.
      • Suttorp N.
      ). In situ TUNEL staining was used to examine whether the Rho-dependent cytoskeletal changes observed following apo(a) treatment were accompanied by the characteristic features of apoptosis. TUNEL staining showed that apo(a)-treated HUVECs displayed no signs of DNA fragmentation (data not shown). Therefore, these results indicate that neither the brief 15-min serum starvation nor the exposure for short periods of time to apo(a) caused apoptosis in HUVECs.
      We hypothesized that the increase in gap formation and loss of cell-cell contact following apo(a) exposure would lead to enhanced transendothelial permeability. Consistent with this hypothesis, HUVEC monolayers treated with 400 nm apo(a) displayed a time-dependent increase in transendothelial diffusion of fluorescein isothiocyanate-dextran (Fig. 2). HUVEC monolayers became ∼2-fold more permeable compared with controls. These results confirm that cytoskeletal changes elicited by apo(a) are correlated with increased cellular contractility sufficient to form intercellular gaps.
      Figure thumbnail gr2
      Fig. 2Apo(a) results in a time-dependent increase in transendothelial permeability. HUVECs were treated with medium (control) or medium containing 400 nm apo(a) for the indicated time periods. Transendothelial permeability was determined fluorometrically as described under “Experimental Procedures.” Results represent the mean ± S.E. of four measurements.
      Actin Stress Fiber Stimulation by Apo(a) Leads to Dispersion of VE-cadherin and Is Mediated by Rho and Rho Kinase—To determine whether apo(a)-induced stress fiber formation involves Rho and Rho kinase, HUVECs were pretreated with their respective inhibitors, C3 transferase and Y-27632. Preincubation with C3 transferase or Y-27632 had little or no effect on the F-actin distribution of control serum-starved HUVECs (data not shown) but completely abolished the increase in stress fiber formation induced by apo(a) (Fig. 3, A–D). These results strongly suggest that stimulation of actin stress fiber formation by apo(a) occurs through a Rho/Rho kinase-dependent pathway.
      Figure thumbnail gr3
      Fig. 3Apo(a) results in a Rho/Rho kinase-dependent disruption of VE-cadherin organization. HUVECs were not stimulated (A and E), treated with 400 nm apo(a) for 40 min (B and F), pretreated with 5 μg/ml C3 transferase for 24 h and then treated with apo(a) (C and G), or pretreated with 10 μm Y-27632 for 30 min and then treated with apo(a) (D and H). Cells were fixed, permeabilized, and stained for actin and VE-cadherin.
      Our previous results indicate that apo(a) treatment leads to increased cell contractility and subsequent endothelial permeability. We theorized that loss of cell-cell contacts would result in changes in distribution of the adhesion molecule VE-cadherin. VE-cadherin/cadherin-5 is a junctional protein that is unique to endothelial cells. This class of cadherin is present at sites of cell-cell contact and as such acts as a suitable marker for gap formation and ultimately increased vascular permeability (
      • Lampugnani M.G.
      • Corada M.
      • Caveda L.
      • Breviario F.
      • Ayalon O.
      • Geiger B.
      • Dejana E.
      ). Serum-starved control cells possessed VE-cadherin organized as slender lines along the margins of the cells (Fig. 3E). In contrast, HUVECs treated with apo(a) had a dispersed VE-cadherin appearance that occurred concomitantly with enhanced F-actin stress fiber formation (Fig. 3, B and F). Loss of VE-cadherin at sites of intercellular contact coincided with gap formation between endothelial cells (Fig. 3F). In addition, inhibition of Rho and its effector Rho kinase abolished the effects of apo(a) on VE-cadherin distribution (Fig. 3, G and H).
      Rho/Rho Kinase-mediated Changes in Focal Adhesion Molecule Organization by Apo(a)—Focal adhesions act as attachment sites for stress fibers and play an essential role in linking the actin cytoskeleton to the extracellular matrix. We investigated the effects of apo(a) on the distribution of focal adhesions in HUVECs (Fig. 4) by immunofluorescence microscopy using the characteristic focal adhesion protein vinculin as a marker. In serum-starved cells, vinculin-containing adhesive structures were concentrated at the cell periphery (Fig. 4E). Treatment with apo(a) (Fig. 4F) caused a dramatic loss of peripheral adhesions and an increase in typical oval-shaped focal adhesions distributed throughout the cell body. Therefore, enhanced F-actin stress fiber formation due to apo(a) treatment occurs with a concomitant redistribution of vinculin and formation of focal adhesions. Consistent with the key role of the Rho/Rho kinase pathway in mediating the effect of apo(a) and Lp(a) on stress fiber formation, the inhibitors C3 transferase and Y-27632 completely abrogated the ability of apo(a) to elicit changes in the distribution of focal adhesions (Fig. 4, G and H).
      Figure thumbnail gr4
      Fig. 4Apo(a) causes reorganization of focal adhesions in HUVECs treated with apo(a) that is mediated by Rho and Rho kinase. HUVECs were not stimulated (A and E), treated with 400 nm apo(a) for 40 min (B and F), pretreated with 5 μg/ml C3 transferase for 24 h and then treated with apo(a) (C and G), or pretreated with 10 μm Y-27632 for 30 min and then treated with apo(a) (D and H). Cells were fixed, permeabilized, and stained for actin and vinculin.
      Lp(a) and Apo(a) Promote Actin Stress Fiber Formation in Cultured Arterial Endothelial Cells—Considering the known heterogeneity among endothelial cells isolated from different vascular compartments (
      • Garlanda C.
      • Dejana E.
      ), we examined the ability of apo(a) and Lp(a) to effect cytoskeletal rearrangements in cultured HCAECs. This endothelial cell type is more relevant to the effects of apo(a)/Lp(a) on the initiation of atherosclerosis. Treatment with Lp(a) and apo(a) (each at 400 nm) for 5 min evoked the formation of actin stress fibers in a manner qualitatively very similar to that in HUVECs (Fig. 5, A, D, and G; compare with Fig. 1). As in HUVECs, the effect of Lp(a)/apo(a) in HCAECs was Rho/Rho kinase-dependent (Fig. 5). In addition, a similar effect of apo(a) on the redistribution of focal adhesions was observed as visualized by immunofluorescence microscopy with vinculin as a marker (data not shown).
      Figure thumbnail gr5
      Fig. 5Apo(a) elicits Rho/Rho kinase-dependent actin cytoskeleton reorganization in cultured human coronary artery endothelial cells. Serum-starved HCAECs were not stimulated (A–C), treated with 400 nm apo(a) for 40 min (D–F), or treated with 400 nm Lp(a) for 5 min (G–I). In some cases, cells were pretreated with 5 μg/ml C3 transferase for 24 h (B, E, and H) or pretreated with 10 μm Y-27632 for 30 min (C, F, and I) prior to stimulation. Cells were then fixed, permeabilized, and stained for actin.
      Actin Cytoskeleton Remodeling and VE-cadherin Disruption Mediated by Apo(a) Is Regulated by Rho and Rac but Not Cdc42—We also wished to establish whether other Rho-related GTP-binding proteins may be involved in endothelial cytoskeletal reorganization and loss of cell-cell contact initiated by apo(a). To investigate this possibility, HUVECs were transiently transfected with dominant negative forms of Rho (RhoN19), Rac (RacN17), and Cdc42 (Cdc42N17), treated with apo(a), and subsequently analyzed for changes in actin stress fiber and VE-cadherin organization. Expression of dominant negative RhoN19 and RacN17 did not significantly alter the actin or VE-cadherin distribution in quiescent HUVECs (transfected cells are visualized as Myc-positive) (Fig. 6, A, B, G, and H). HUVECs transfected with dominant negative RhoN19 and RacN17 did not assemble actin stress fibers following apo(a) addition (Fig. 6, D and J). In addition, VE-cadherin was not disrupted in RhoN19- and RacN17-transfected cells treated with apo(a) (Fig. 6, E and K). These findings demonstrate that enhanced stress fiber formation and VE-cadherin reorganization caused by apo(a) depend on activation of both Rho and Rac.
      Figure thumbnail gr6
      Fig. 6Stress fiber formation and loss of cell-cell contacts by apo(a) is regulated by Rac and Rho. HUVECs were transiently transfected with dominant negative RhoN19 (A–F) or RacN17 (G–L). Cells were not stimulated (A–C and G–I) or stimulated with 400 nm apo(a) for 40 min (D–F and J–L). Cells were fixed, permeabilized, and stained for actin and VE-cadherin as well as for c-Myc to identify transfected cells.
      Transfection of serum-starved HUVECs with dominant negative Cdc42N17 resulted in a dramatic increase in the content of stress fibers (Fig. 7A) but, interestingly, did not disrupt VE-cadherin organization (Fig. 7B). Addition of apo(a) did not noticeably increase the already high levels of stress fibers present in serum-starved HUVECs expressing Cdc42N17 (Fig. 7D) but resulted in a more disorganized distribution of VE-cadherin (Fig. 7E). To determine the effects of functional Cdc42, HUVECs were transiently transfected with constitutively active Cdc42 (Cdc42V12). HUVECs transfected with Cdc42V12 possessed few stress fibers, appeared smaller than control cells, and extended numerous long, thin filopodia (Fig. 7G). VE-cadherin staining at the cell periphery was not markedly altered in cells transfected with Cdc42V12, but large punctate accumulations of VE-cadherin were visible in the cytoplasm (Fig. 7H). Treatment with apo(a) did not noticeably alter the pattern of effects elicited by Cdc42V12 (Fig. 7, J and K). Specifically, an increase in central stress fibers was not observed, and the transfected cells retained their highly contracted morphology and filopodia. Moreover apo(a) treatment did not appear to cause disruptions in VE-cadherin distribution in the transfected cells.
      Figure thumbnail gr7
      Fig. 7Cdc42 does not regulate stress fiber formation and VE-cadherin reorganization by apo(a). HUVECs were transiently transfected with dominant negative Cdc42N17 and were not stimulated (A–C) or stimulated (D–F) with 400 nm apo(a) for 40 min. In other experiments, HUVECs were transiently transfected with constitutively active Cdc42V12 and were not stimulated (G–I) or stimulated (J–L) with 400 nm apo(a) for 40 min. Cells were fixed, permeabilized, and stained for actin and VE-cadherin as well as for c-Myc to identify transfected cells.
      Enhancement of MLC Phosphorylation by Apo(a) Is Time-dependent and Regulated by Rho and Rho Kinase—Phosphorylation of the MLC is necessary for myosin II activation and stress fiber formation and thus for cell contraction (
      • Chrzanowska-Wodnicka M.
      • Burridge K.
      ). To establish a role for apo(a) in MLC phosphorylation, HUVECs were exposed to apo(a), and the extent of MLC phosphorylation was assessed by Western blot analysis using an antibody specific for the phosphorylated (on Ser-19) form of MLC. Maximal MLC phosphorylation (∼3-fold higher than untreated control cells) occurred after 5 min of apo(a) stimulation (Fig. 8, A and B); the extent of MLC phosphorylation then decreased but plateaued at a level higher than that present in untreated cells. Interestingly, this time course is broadly coincident with the time-dependent effects of apo(a) on stress fiber formation (Fig. 1A). Consistent with the results described above, pretreatment of the cells with the Rho inhibitor C3 transferase or the Rho kinase inhibitor Y-27632 abolished the ability of apo(a) to enhance the extent of MLC phosphorylation (Fig. 8C).
      Figure thumbnail gr8
      Fig. 8Apo(a) stimulates MLC phosphorylation in HUVECs in a Rho/Rho kinase-dependent manner. HUVECs were serum-starved prior to treatment with apo(a) (400 nm) for 0, 2, 5, 10, and 20 min. Total cellular protein was harvested and subjected to Western blot analysis using an anti-phospho-MLC antibody to determine the extent of MLC phosphorylation. A, representative Western blot showing increased MLC phosphorylation following apo(a) treatment. B, quantification of MLC phosphorylation time course by densitometry (data represent mean ± range of two independent experiments); the data are normalized to the signal for total MLC from blots performed in parallel using the same protein samples. C, representative Western blot showing decreased apo(a)-mediated MLC phosphorylation with pretreatment using C3 transferase and Y-27632. Cells were serum-starved for 15 min, treated with apo(a) (400 nm, 5 min), pretreated with C3 transferase (5 μg/ml, 24 h) and then treated with apo(a), or pretreated with Y-27632 (10 μm, 30 min) and then treated with apo(a). Total cellular proteins were harvested and subjected to Western blot analysis using an anti-phospho-MLC antibody. MLC-P, phosphorylated MLC.
      Apo(a) Treatment Does Not Result in Calcium Transients—The results described in the preceding sections indicate roles for GTP Rho-binding proteins in endothelial cytoskeletal rearrangement evoked by apo(a). We further wished to explore whether a calcium-dependent pathway may be acting in parallel to potentiate the effects of the GTP Rho-binding proteins. HUVECs were loaded with the calcium-sensitive fluorophore Fura-2/AM to monitor changes in intracellular calcium levels. HUVECs treated with apo(a) did not evoke an increase in intracellular calcium levels (Fig. 9A). These results suggest that apo(a) is mediating its effects predominantly through a Rho GTPase-dependent pathway rather than via activation of myosin light chain kinase in a Ca2+/calmodulin-dependent manner.
      Figure thumbnail gr9
      Fig. 9Apo(a) treatment does not lead to increases in intracellular calcium concentration, but the effect is calcium-dependent. A, HUVECs were loaded with Fura-2/AM and stimulated with 400 nm apo(a). Shown is a representative single cell imaging trace of the ratio of fluorescence intensity at 510 nm corresponding to excitation at 340 nm or 380 nm (ratio 340/380). B, HUVECs were not stimulated (a and e), stimulated with 400 nm apo(a) for 40 min (b and f), pretreated with BAPTA-AM (25 μm, 1 h) and then treated with apo(a) (c and g), or pretreated with ML-7 (25 μm, 1 h) and then treated with apo(a) (d and h). Cells were fixed, permeabilized, and stained for actin and VE-cadherin as indicated.
      To confirm this finding, HUVECs were preincubated with the calcium-selective chelator BAPTA-AM prior to apo(a) treatment. Surprisingly, chelation of intracellular calcium by BAPTA-AM completely abolished the increase in F-actin stress fibers produced by apo(a) (Fig. 9B). Therefore, despite the fact that apo(a) does not potentiate intracellular calcium concentration, the effects of apo(a) are dependent on basal calcium levels. This calcium dependence is likely due to activation of the MLC kinase that results in MLC phosphorylation and contraction (
      • Verin A.D.
      • Lazar V.
      • Torry R.J.
      • Labarrere C.A.
      • Patterson C.E.
      • Garcia J.G.
      ,
      • Norwood N.
      • Moore T.M.
      • Dean D.A.
      • Bhattacharjee R.
      • Li M.
      • Stevens T.
      ). To examine this possible role of endothelial MLC kinase, HUVECs were preincubated with the MLC kinase-specific inhibitor ML-7 followed by apo(a) treatment. HUVECs displayed an F-actin cytoskeletal pattern that was consistent with that in cells pretreated with BAPTA-AM (Fig. 9B). Furthermore VE-cadherin appeared as highly organized fine strands when HUVECs were pretreated with BAPTA-AM and ML-7 (Fig. 9B). We therefore conclude that the calcium-activated MLC kinase is necessary for apo(a) to elicit its effects on the endothelial cytoskeleton and promote intercellular gap formation.

      DISCUSSION

      Although numerous studies have demonstrated that Lp(a) is a risk factor for vascular diseases, the exact mechanisms by which it exerts its pathogenic effects remain unclear. Lp(a) or its distinguishing component apo(a) have been shown to affect endothelial cell function through a variety of mechanisms. Clinical studies have demonstrated impaired endothelium-dependent vasodilation in hypercholesterolemic children with high Lp(a) (
      • Sorensen K.E.
      • Celermajer D.S.
      • Georgakopoulos D.
      • Hatcher G.
      • Betteridge D.J.
      • Deanfield J.E.
      ). Similarly, elevation in Lp(a) levels has been associated with impairment of receptor-mediated endothelial vasodilation in adult subjects (
      • Tsurumi Y.
      • Nagashima H.
      • Ichikawa K.
      • Sumiyoshi T.
      • Hosoda S.
      ,
      • Schachinger V.
      • Halle M.
      • Minners J.
      • Berg A.
      • Zeiher A.M.
      ). In vitro studies also support a role for Lp(a) in endothelial dysfunction. For example, the apo(a) component of Lp(a) has been shown to induce monocyte chemotactic activity through the induction of the CC chemokine I-309 and to induce intracellular adhesion molecule-1 expression in HUVECs (
      • Haque N.S.
      • Zhang X.
      • French D.L.
      • Li J.
      • Poon M.
      • Fallon J.T.
      • Gabel B.R.
      • Taubman M.B.
      • Koschinsky M.L.
      • Harpel P.C.
      ,
      • Takami S.
      • Yamashita S.
      • Kihara S.
      • Ishigami M.
      • Takemura K.
      • Kume N.
      • Kita T.
      • Matsuzawa Y.
      ). Additionally, Lp(a), but not apo(a), has been shown to increase the expression of vascular cell adhesion molecule-1 and E-selectin in HCAECs (
      • Allen S.
      • Khan S.
      • Tam S.-P.
      • Koschinsky M.L.
      • Taylor P.
      • Yacoub M.
      ).
      In the present study, we have uncovered an entirely novel mechanism by which Lp(a) can contribute to a dysfunctional endothelial phenotype. We found that Lp(a), as well as its distinguishing component apo(a), stimulates the formation of actin stress fibers and redistribution of focal adhesions in cultured endothelial cells. This effect is manifested by an apo(a)-mediated increase in MLC phosphorylation and ultimately can be demonstrated to result in marked endothelial cell contraction and increased transendothelial permeability. The actin cytoskeleton plays an essential role in endothelial cell structure maintenance (
      • Shasby D.M.
      • Shasby S.S.
      • Sullivan J.M.
      • Peach M.J.
      ,
      • Gotlieb A.I.
      • Langille B.L.
      • Wong M.K.
      • Kim D.W.
      ). Whereas peripheral F-actin serves to preserve endothelial cell monolayer integrity, central F-actin stress fibers regulate adhesion to the substratum. Focal adhesions serve to mediate the physical linking of actin stress fibers to the extracellular matrix through a complex assortment of cytoplasmic proteins including vinculin, α-actinin, talin, and transmembrane adhesive glycoproteins of the integrin family (
      • Lum H.
      • Malik A.B.
      ). In addition to anchoring the cell to its substratum, focal adhesion molecules may play a crucial role in the regulation of endothelial cell permeability (
      • Bishop A.L.
      • Hall A.
      ). We speculate that the presumptive compromised endothelial barrier function in individuals with elevated Lp(a) levels could explain a component of the enhanced risk for atherosclerotic diseases observed in this population.
      We have also begun to elucidate the signaling cascade that is responsible for the cytoskeleton remodeling by apo(a) (Fig. 10). The ability of apo(a)/Lp(a) to evoke these dramatic cytoskeletal rearrangements is mediated through a Rho/Rho kinase-dependent signaling pathway since inhibition of these intracellular effectors abolishes the ability of apo(a) and Lp(a) to increase stress fiber formation and to promote changes in the distribution of focal adhesions. Similarly, inhibition of Rac, an upstream effector of Rho in endothelial cells (
      • Wojciak-Stothard B.
      • Potempa S.
      • Eichholtz T.
      • Ridley A.J.
      ) and Swiss 3T3 cells (
      • Ridley A.J.
      • Hall A.
      ), also abolished these effects of apo(a). In contrast, apo(a) induces its effects on the endothelial cytoskeleton through a pathway that is apparently independent of Cdc42, an upstream effector of Rac in endothelial cells (
      • Wojciak-Stothard B.
      • Potempa S.
      • Eichholtz T.
      • Ridley A.J.
      ) and in Swiss 3T3 cells (
      • Kozma R.
      • Ahmed S.
      • Best A.
      • Lim L.
      ). In the present study, apo(a) treatment disrupted VE-cadherin even in the presence of dominant negative Cdc42 (although the effect of apo(a) on stress fibers could not be ascertained), and apo(a) treatment in quiescent serum-starved HUVECs did not produce any detectable filopodia as seen in cells overexpressing constitutively active Cdc42. Expression of constitutively active Cdc42 appeared to abolish the ability of apo(a) to cause formation of actin stress fibers and redistribution of VE-cadherin. The extreme morphological effects of constitutively active Cdc42 likely overwhelmed any effect of Rac and Rho activation by apo(a).
      Figure thumbnail gr10
      Fig. 10Proposed model demonstrating signaling events involved in apo(a)-mediated endothelial cell contraction and loss of cell-cell contact. Binding of apo(a) to a putative receptor results in activation of Rac, Rho, and Rho kinase. Cellular contraction occurs through phosphorylation of the MLC by Ca2+/calmodulin-activated MLC kinase (MLCK). Rho kinase may enhance MLC phosphorylation through inactivation of the MLC phosphatase type 1 myosin-associated protein phosphatase (PP1M) by direct phosphorylation. Other effectors of endothelial cell contraction, such as tumor necrosis factor-α (TNF-α), activate Rac through Cdc42. The signaling pathway activated by Lp(a), however, appears to bypass Cdc42.
      Our data demonstrate that apo(a) stimulation does not result in an increase in cytosolic calcium concentration (Fig. 9). Interestingly, mildly oxidized LDL has also been shown not to increase intracellular calcium levels (
      • Essler M.
      • Retzer M.
      • Bauer M.
      • Heemsker J.W.
      • Aepfelbacher M.
      • Seiss W.
      ). We have, however, shown that the effect of apo(a) on actin stress fiber formation is dependent on intracellular calcium. This calcium dependence is likely related to the requirement for a basal level of MLC kinase activity (
      • Verin A.D.
      • Lazar V.
      • Torry R.J.
      • Labarrere C.A.
      • Patterson C.E.
      • Garcia J.G.
      ,
      • Norwood N.
      • Moore T.M.
      • Dean D.A.
      • Bhattacharjee R.
      • Li M.
      • Stevens T.
      ). It has been reported that Rho kinase is capable of phosphorylating MLC on Ser-19 (
      • Amano M.
      • Ito M.
      • Kimura K.
      • Fukata Y.
      • Chihara K.
      • Nakano T.
      • Matsuura M.
      • Kaibuchi K.
      ), which may also account, at least in part, for the effect of apo(a) on MLC phosphorylation and hence cellular contraction. However, the requirement for calcium suggests against this scenario since MLC kinase is calcium-dependent, whereas Rho kinase is not.
      Based on the results from the present study, we propose that apo(a) stimulation results in the activation of a signaling cascade involving Rac, Rho, and Rho kinase (Fig. 10). Activated Rho kinase is then capable of phosphorylating and inhibiting the MLC phosphatase type 1 myosin-associated protein phosphatase that would also result in increased MLC phosphorylation (
      • Essler M.
      • Retzer M.
      • Bauer M.
      • Heemsker J.W.
      • Aepfelbacher M.
      • Seiss W.
      ,
      • Essler M.
      • Amano M.
      • Kruse H.J.
      • Kaibuchi K.
      • Weber P.C.
      • Aepfelbacher M.
      ).
      Importantly, we show conclusively that the effects of Lp(a) on endothelial cell cytoskeletal rearrangements are mediated specifically by the apo(a) moiety. Apo(a) and Lp(a) had a similar effect on endothelial cells, and native LDL had no effect. These findings are significant since previous work has found that mildly oxidized LDL also causes cytoskeletal rearrangements via a Rho/Rho kinase-dependent pathway (
      • Essler M.
      • Retzer M.
      • Bauer M.
      • Heemsker J.W.
      • Aepfelbacher M.
      • Seiss W.
      ). In this case, the lysophosphatidic acid component of mildly oxidized LDL was implicated via its interaction with the G protein-coupled lysophosphatidic acid receptor (
      • Essler M.
      • Retzer M.
      • Bauer M.
      • Heemsker J.W.
      • Aepfelbacher M.
      • Seiss W.
      ). Activation of G protein-coupled receptors is thought to promote the GTP-bound, and thus activated, form of Rho through the activation of guanine nucleotide exchange factors (
      • Bishop A.L.
      • Hall A.
      ). The identities of the factors upstream of Rac that mediate the ability of apo(a) to activate the signaling pathway involving Rac, Rho, and Rho kinase have yet to be identified.
      Lp(a) and apo(a) have been shown to compete with plasminogen binding to endothelial cells (
      • Hajjar K.A.
      • Gavish D.
      • Breslow J.L.
      • Nachman R.L.
      ), suggesting that apo(a) may bind plasminogen receptors such as annexin II (
      • Hajjar K.A.
      • Krishnan S.
      ) and α-enolase (
      • Miles L.A.
      • Fless G.M.
      • Scanu A.M.
      • Baynham P.
      • Sebald M.T.
      • Skocir P.
      • Curtiss L.K.
      • Levin E.G.
      • Hoover-Plow J.L.
      • Plow E.F.
      ). The competition of apo(a) for plasminogen binding to the cell surface also has been speculated to result in inhibition of pericellular plasminogen activation (
      • Hajjar K.A.
      • Gavish D.
      • Breslow J.L.
      • Nachman R.L.
      ). This has been shown to inhibit the plasmin-mediated conversion of latent transforming growth factor-β to active transforming growth factor-β (
      • Grainger D.J.
      • Kirschenlohr H.L.
      • Metcalfe J.C.
      • Weissberg P.L.
      • Wade D.P.
      • Lawn R.M.
      ), itself a potent signaling molecule. Note, however, that our experiments were performed on extensively washed monolayers of cells and in the absence of serum as a source of plasminogen. This makes it unlikely that our observations reflect inhibition by apo(a) of transforming growth factor-β activation. That the effect of apo(a) is rapid (within 2 min) and occurs in the absence of serum is consistent with a direct, receptor-mediated effect of apo(a) on intracellular signaling pathways. We cannot at this time, however, exclude the possibility that apo(a) might somehow activate Rho after receptor-mediated internalization.
      Our study constitutes the first demonstration of an effect of the apo(a) component of Lp(a) on endothelial cell intracellular signaling pathways. As such, our findings reveal a novel paradigm for the potential atherogenic effects of Lp(a). It might be hypothesized that Lp(a), through its effects on intracellular signaling pathways, evokes a general dysfunctional program in the endothelium that involves not only the cytoskeletal rearrangements reported here but also the previously described increases in monocyte chemoattractant and cell surface adhesion molecule expression and impairment of receptor-dependent dilation responses. The mechanistic basis of these effects of Lp(a) will be a fruitful area of future research.

      Acknowledgments

      Plasmids used for transfections were generously provided by Alan Hall (Medical Research Council Laboratory for Molecular Cell Biology and Cell Biology Unit, Cancer Research UK Oncogene and Signal Transduction Group, University College, London). Use of the microscope facility was subsidized by the Queen's University Protein Function Discovery Facility.

      References

        • Poredos P.
        Clin. Appl. Thromb. Hemost. 2001; 7: 276-280
        • Wojciak-Stothard B.
        • Entwistle A.
        • Garg R.
        • Ridley A.J.
        J. Cell. Physiol. 1998; 176: 150-165
        • Lum H.
        • Ashner J.L.
        • Phillips P.G.
        • Fletcher P.W.
        • Malik A.B.
        Am. J. Physiol. 1992; 263: L219-L225
        • Essler M.
        • Retzer M.
        • Bauer M.
        • Heemsker J.W.
        • Aepfelbacher M.
        • Seiss W.
        J. Biol. Chem. 1999; 274: 30361-30364
        • Garcia J.G.
        • Davis H.W.
        • Patterson C.E.
        J. Cell. Physiol. 1995; 163: 510-522
        • Goeckler Z.M.
        • Wysolmerski R.B.
        J. Cell Biol. 1995; 130: 613-627
        • Chrzanowska-Wodnicka M.
        • Burridge K.
        J. Cell Biol. 1996; 133: 1403-1415
        • Verin A.D.
        • Patterson C.E.
        • Day M.A.
        • Garcia J.G.
        Am. J. Physiol. 1995; 269: L99-L108
        • Somlyo A.P.
        • Somlyo A.V.
        Nature. 1994; 372: 231-236
        • Amano M.
        • Ito M.
        • Kimura K.
        • Fukata Y.
        • Chihara K.
        • Nakano T.
        • Matsuura M.
        • Kaibuchi K.
        J. Biol. Chem. 1996; 271: 20246-20249
        • Kimura K.
        • Ito M.
        • Amano M.
        • Chihara K.
        • Fukata Y.
        • Nakafuku M.
        • Yamamori B.
        • Feng J.
        • Nakano T.
        • Okawa K.
        • Iwamatsu A.
        • Kaibuchi K.
        Science. 1996; 273: 245-248
        • Kawano Y.
        • Fukata Y.
        • Oshiro N.
        • Amano M.
        • Nakamura T.
        • Ito M.
        • Matsumura F.
        • Inagaki M.
        • Kaibuchi K.
        J. Cell Biol. 1999; 147: 1023-1038
        • Ridley A.J.
        • Hall A.
        Cell. 1992; 70: 389-399
        • Bishop A.L.
        • Hall A.
        Biochem. J. 2000; 348: 241-255
        • Nobes C.D.
        • Hall A.
        Biochem. Soc. Trans. 1995; 23: 456-459
        • Wojciak-Stothard B.
        • Potempa S.
        • Eichholtz T.
        • Ridley A.J.
        J. Cell Sci. 2001; 114: 1343-1355
        • Marcovina S.M.
        • Koschinsky M.L.
        Curr. Cardiol. Rep. 1999; 1: 105-111
        • Marcovina S.M.
        • Koschinsky M.L.
        Semin. Vasc. Med. 2002; 2: 335-344
        • Scanu A.M.
        Curr. Atheroscler. Rep. 2003; 5: 106-113
        • Brunner C.
        • Kraft H.G.
        • Utermann G.
        • Muller H.J.
        Proc. Natl. Acad. Sci. U. S. A. 1993; 90: 11643-11647
        • Koschinsky M.L.
        • Côté G.P.
        • Gabel B.
        • van der Hoek Y.Y.
        J. Biol. Chem. 1993; 268: 19819-19825
        • McLean J.W.
        • Tomlinson J.E.
        • Kuang W.
        • Eaton D.L.
        • Chen E.
        • Fless G.
        • Scanu A.
        • Lawn R.M.
        Nature. 1987; 330: 132-137
        • Koschinsky M.L.
        • Tomlinson J.E.
        • Zioncheck T.F.
        • Schwartz K.
        • Eaton D.L.
        • Lawn R.M.
        Biochemistry. 1991; 30: 5044-5051
        • Sangrar W.
        • Gabel B.R.
        • Boffa M.B.
        • Walker J.B.
        • Hancock M.A.
        • Marcovina S.M.
        • Horrevoets A.J.G.
        • Nesheim M.E.
        • Koschinsky M.L.
        Biochemistry. 1997; 36: 10353-10363
        • Castellino F.J
        • Powell J.R.
        Methods Enzymol. 1981; 80: 365-378
        • Edelstein C.
        • Hinman J.
        • Marcovina S.
        • Scanu A.M.
        Anal. Biochem. 2001; 288: 201-208
        • Gabel B.R.
        • McLeod R.S.
        • Yao Z.
        • Koschinsky M.L.
        Arterioscler. Thromb. Vasc. Biol. 1998; 18: 1738-1744
        • Marcovina S.M.
        • Albers J.J.
        • Wijsman E.
        • Zhang Z.
        • Chapman N.H.
        • Kennedy H.
        J. Lipid Res. 1996; 37: 2569-2585
        • Hippenstiel S.
        • Schmeck B.
        • N′Guessan P.D.
        • Seybold J.
        • Krull M.
        • Preissner K.
        • Eichel-Streiber C.V.
        • Suttorp N.
        Am. J. Physiol. 2002; 283: L830-L838
        • Lampugnani M.G.
        • Corada M.
        • Caveda L.
        • Breviario F.
        • Ayalon O.
        • Geiger B.
        • Dejana E.
        J. Cell Biol. 1995; 129: 203-217
        • Garlanda C.
        • Dejana E.
        Arterioscler. Thromb. Vasc. Biol. 1997; 17: 1193-1202
        • Verin A.D.
        • Lazar V.
        • Torry R.J.
        • Labarrere C.A.
        • Patterson C.E.
        • Garcia J.G.
        Am. J. Respir. Cell Mol. Biol. 1998; 19: 758-766
        • Norwood N.
        • Moore T.M.
        • Dean D.A.
        • Bhattacharjee R.
        • Li M.
        • Stevens T.
        Am. J. Physiol. 2000; 279: L815-L824
        • Sorensen K.E.
        • Celermajer D.S.
        • Georgakopoulos D.
        • Hatcher G.
        • Betteridge D.J.
        • Deanfield J.E.
        J. Clin. Investig. 1994; 93: 50-55
        • Tsurumi Y.
        • Nagashima H.
        • Ichikawa K.
        • Sumiyoshi T.
        • Hosoda S.
        J. Am. Coll. Cardiol. 1995; 26: 1242-1250
        • Schachinger V.
        • Halle M.
        • Minners J.
        • Berg A.
        • Zeiher A.M.
        J. Am. Coll. Cardiol. 1997; 30: 927-934
        • Haque N.S.
        • Zhang X.
        • French D.L.
        • Li J.
        • Poon M.
        • Fallon J.T.
        • Gabel B.R.
        • Taubman M.B.
        • Koschinsky M.L.
        • Harpel P.C.
        Circulation. 2000; 102: 786-792
        • Takami S.
        • Yamashita S.
        • Kihara S.
        • Ishigami M.
        • Takemura K.
        • Kume N.
        • Kita T.
        • Matsuzawa Y.
        Circulation. 1998; 97: 721-728
        • Allen S.
        • Khan S.
        • Tam S.-P.
        • Koschinsky M.L.
        • Taylor P.
        • Yacoub M.
        FASEB J. 1998; 12: 1765-1776
        • Shasby D.M.
        • Shasby S.S.
        • Sullivan J.M.
        • Peach M.J.
        Circ. Res. 1982; 51: 657-661
        • Gotlieb A.I.
        • Langille B.L.
        • Wong M.K.
        • Kim D.W.
        Lab. Investig. 1991; 65: 123-137
        • Lum H.
        • Malik A.B.
        Am. J. Physiol. 1994; 267: L223-L241
        • Kozma R.
        • Ahmed S.
        • Best A.
        • Lim L.
        Mol. Cell. Biol. 1995; 15: 1942-1952
        • Essler M.
        • Amano M.
        • Kruse H.J.
        • Kaibuchi K.
        • Weber P.C.
        • Aepfelbacher M.
        J. Biol. Chem. 1998; 273: 21867-21874
        • Hajjar K.A.
        • Gavish D.
        • Breslow J.L.
        • Nachman R.L.
        Nature. 1989; 339: 303-305
        • Hajjar K.A.
        • Krishnan S.
        Trends Cardiovasc. Med. 1999; 9: 128-138
        • Miles L.A.
        • Fless G.M.
        • Scanu A.M.
        • Baynham P.
        • Sebald M.T.
        • Skocir P.
        • Curtiss L.K.
        • Levin E.G.
        • Hoover-Plow J.L.
        • Plow E.F.
        Thromb. Haemostasis. 1995; 73: 458-465
        • Grainger D.J.
        • Kirschenlohr H.L.
        • Metcalfe J.C.
        • Weissberg P.L.
        • Wade D.P.
        • Lawn R.M.
        Science. 1993; 260: 1655-1658