Oxidative stress induces myeloperoxidase expression ... - Springer Link

Basic Res Cardiol 104:307–320 (2009) DOI 10.1007/s00395-008-0761-9

Giampiero La Rocca Antonino Di Stefano Ermanno Eleuteri Rita Anzalone Francesca Magno Simona Corrao Tiziana Loria Anna Martorana Claudio Di Gangi Marilena Colombo Fabrizio Sansone Francesco Patane` Felicia Farina Mauro Rinaldi Francesco Cappello Pantaleo Giannuzzi Giovanni Zummo Received: 3 December 2007 Returned for 1. Revision: 7 January 2008 1. Revision received: 18 June 2008 Returned for 2. Revision: 17 July 2008 2. Revision received: 17 October 2008 Accepted: 27 October 2008 Published online: 22 November 2008 G. La Rocca and A. Di Stefano contributed equally to the current work. Electronic supplementary material: The online version of this article (doi: 10.1007/s00395-008-0761-9) contains supplementary material, which is available to authorized users. Dr. G. La Rocca (&) Æ R. Anzalone F. Magno Æ S. Corrao Æ T. Loria Æ F. Farina F. Cappello Æ G. Zummo Sezione di Anatomia Umana Dipto. di Medicina Sperimentale Universita` degli Studi di Palermo Via del Vespro 129 90127 Palermo, Italy Tel.: +39-091/7655-3576 Fax: +39-091/655-3580 E-Mail: [email protected] A. Di Stefano Æ M. Colombo Laboratorio di Citoimmunopatologia Apparato Cardio-Respiratorio Fondazione S. Maugeri, IRCCS Veruno (NO), Italy

ORIGINAL CONTRIBUTION

Oxidative stress induces myeloperoxidase expression in endocardial endothelial cells from patients with chronic heart failure

j Abstract Increased oxidative stress has been implicated in the

pathogenesis of a number of cardiovascular diseases. Recent findings suggest that myeloperoxidase (MPO) may play a key role in the initiation and maintenance of chronic heart failure (CHF) by contributing to the depletion of the intracellular reservoir of nitric oxide (NO). NO consumption through MPO activity may lead to protein chlorination or nitration, leading to tissue damage. Primary cultures of human endocardial endothelial cells (EEC) obtained at heart transplantation of patients with CHF and human umbilical vein endothelial cells (HUVEC) were subjected to oxidative stress by incubation with hydrogen peroxide at non lethal (60 lM) dose for different exposure times (3 and 6 h). Treated and control cells were tested by immunohistochemistry and RTPCR for MPO and 3-chlorotyrosine expression. Both endothelial cell types expressed myeloperoxidase following oxidative stress, with higher levels in EEC. Moreover, 3-chlorotyrosine accumulation in treated cells alone indicated the presence of MPO-derived hypochlorous acid. Immunohistochemistry on sections from post-infarcted heart confirmed in vivo the endothelial positivity to MPO, 3-chlorotyrosine and, to a minor extent, nitrotyrosine. Immunohistochemical observations were confirmed by detection of MPO mRNA in both stimulated EEC and HUVEC cells. This study demonstrates for the first time that EEC can express MPO after oxidative stress, both in vitro and in vivo, followed by accumulation of 3-chlorotyrosine, an end product of oxidative stress. Deregulation of endothelial functions may contribute to the development of a number of cardiovascular diseases, including CHF. The results also highlight the notion that endothelium is not only a target but also a key player in oxidative-driven cardiovascular stress. j Key words 3-chlorotyrosine – endocardium – endothelial cells – myeloperoxidase – oxidative stress

A. Martorana Dipto. di Patologia Umana Universita` degli Studi di Palermo Palermo, Italy

C. Di Gangi Istituto di Ostetricia e Ginecologia Universita` degli Studi di Palermo Palermo, Italy

F. Sansone Æ F. Patane` Æ M. Rinaldi Divisione di Cardiochirurgia Ospedale S. Giovanni Battista Turin, Italy

BRC 761

E. Eleuteri Æ P. Giannuzzi Divisione di Cardiologia Fondazione S. Maugeri, IRCCS Veruno (NO), Italy

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Basic Research in Cardiology, Vol. 104, No. 3 (2009) Steinkopff Verlag 2008

Introduction Myeloperoxidase is a heme protein which uses hydrogen peroxide and chloride to generate the potent microbicidal hypochlorous acid (HOCl) [16]. This molecule is prominently expressed in polymorphonuclear neutrophils as well as in monocytes, but recent data show that its expression is not restricted to the myeloid cells. In fact, it is expressed even in granule-containing and pyramidal neurons of the hippocampus, as well as in several neuronal cell lines [18]. When monocytes differentiate into tissue macrophages, MPO protein is no longer present, but it has been reported that the gene can be reactivated in certain subsets of macrophages [24, 40]. Apart from its well characterised role following neutrophil activation in the oxidative burst, MPO has been recently viewed as a key player in the destabilisation of intimal homeostasis in vascular diseases. In fact, one of the better characterised processes of MPO intervention in endothelial homeostasis is the depletion of intracellular NO, to produce nitrogen dioxide, which can in turn be converted to nitrogen radical, causing nitration in tyrosine residues of proteins. Moreover, since MPO is the only enzyme capable of generating HOCl at plasma halide concentrations, it is also responsible for the formation of chlorinated aminoacids, of which 3-chlorotyrosine is a stable and detectable end-product [20, 27, 36]. It has been reported that endothelial cells are able to perform transcytosis of MPO, from the vessel lumen to the basement membrane, where the enzyme specifically triggers extracellular matrix protein nitration, as recently shown for fibronectin [1, 3]. Recent in vivo findings have also shown that the levels of circulating myeloperoxidase inversely correlate with brachial artery dilation, suggesting an active role for the molecule in interfering with NO-mediated vasodilation [46]. Endothelial dysfunction caused by an imbalance between oxidative and nitrosative stress has also been related to myocyte hypertrophy and ultimately to the development of CHF [13, 15, 17, 19, 28, 37]. Our study originated from the hypothesis that endothelial cells may intervene in myeloperoxidase metabolism not only as a target of the neutrophilderived enzyme, or through their role in the translocation of the molecule to the inner vessel wall, but also by reacting to oxidative stress with the production of a number of active molecules. Hydrogen peroxide is a reactive oxygen specie which is normally present in endothelial cells at higher concentrations than in other cell types. Moreover, at the low micromolar range (under 50 lM) it exerts a proliferative effect on human umbilical vein endothelial cells (HUVEC) [6]. Several reports in the literature suggest

that hydrogen peroxide may act as a mediator of activation of endothelial gene expression, causing for example the well known increase in vascular permeability and adhesiveness for leukocytes [5]. More generally, reactive oxygen species have been implicated in the pathophysiology of a number of cardiovascular diseases [5, 6]. The present study shows for the first time that oxidative stress induces the expression of MPO by primary human endothelial cells in culture. Since MPO is the only enzyme capable of generating HOCl at physiological halide concentrations, under the same conditions we were able to demonstrate the intracellular accumulation of 3-chlorotyrosine, one of the stable compounds derived from myeloperoxidasederived HOCl oxidation. Oxidative and nitrosative related end products were also found in post-infarct endocardial endothelial cells (EEC).

Methods j Tissue sampling For EEC isolation, soon after transplantation, tissue slices were sampled from the left ventricle, stored in cold transport buffer, and cell isolation was performed within 6 h. Samples were obtained from seven patients with post-infarct chronic heart failure (CHF). Concerning HUVEC isolation, umbilical cords were obtained from twelve subjects immediately after the partum, and stored in transport buffer [31]. Isolation of cells was performed within 2 h of tissue collection. The investigation conformed to the principles outlined in the Declaration of Helsinki. The patients gave their informed consent prior to be included in this study.

j Cells isolation and culturing: Human endocardial endothelial cells The isolation method for EEC is based on the use of collagenase to detach cells from the sub-endocardial basement membrane. Heart wall slices sampled from the left ventricle were washed extensively with Ca2+ and Mg2+ free HBSS (GIBCO, Milan, Italy) in order to remove residual blood cells. Then the slices were placed in sterile Petri dishes filled with Type II collagenase (GIBCO) diluted in Ca2+ and Mg2+ free HBSS. The dishes were placed at 37C for 15 min, without agitation, and then, after a brief rinse with collagenase solution, the reaction was stopped by adding an equal volume of Isolation Medium (same as for HUVEC, see below) and centrifuging the cells. After centrifugation, the cells were resuspended in the complete Culture Medium (same as for HUVEC, see below) and plated

G. La Rocca et al. Oxidative-driven endothelial expression of myeloperoxidase

on gelatin-coated 6-well culture plates (Corning, Milan, Italy). EEC were used for experiments at passages 2–4, as single primary cell lines.

j Cells isolation and culturing: HUVEC The method for HUVEC isolation is based on the separation of endothelial cells from vessel wall using collagenase to digest the subendothelial basement membrane, according to previously published methods [17, 24, 25] with slight modifications regarding the composition of culture and isolation media and antibiotic treatments. Briefly, for each cord (total number = 24), the vein was cannulated and perfused with a heparin solution (250 U/ml), followed by Ca2+ and Mg2+ free HBSS (GIBCO) in order to remove blood residue from the vessel. Then the vein was perfused with collagenase (Type II, GIBCO) and, after clamping of the ends, incubated for 15 min at 37C. Collagenase solution was then collected in a sterile falcon container, and the vein was thoroughly flushed with Isolation Medium (IM) (M-199 supplemented with 200 U/ml penicillin, 200 lg/ml streptomycin, 0.5 lg/ml amphotericin B and 2 Mm L-glutamine, 10% foetal bovine serum (FBS), all from GIBCO), in order to detach all endothelial cells. After centrifugation, the cells were resuspended in complete Culture Medium (CM) (M-199 with 100 U/ml penicillin, 100 lg/ml streptomycin, 0.25 lg/ml amphotericin B and 2 mM L-glutamine, 10% FBS), supplemented with the appropriate growth supplements for endothelial cells: 20 lg/ml endothelial cell growth factor (ECGF, Roche, Milan, Italy), 14 U/ml sodium heparin (SigmaAldrich, Milan, Italy) [21, 30, 31]. Cells were seeded in gelatin-coated culture flasks at a concentration of 30,000/cm2. Primary HUVEC were used for the experiments at passages 2–4. Four experimental sets were made on endothelial cells pooled from three different cords on each occasion.

j Hydrogen peroxide treatment In order to expose cells to oxidative stress, both endothelial cells were plated either in 8-well chamber slides to perform immunocytochemistry, or 6-well culture plates to extract total RNA. Cells were seeded at a density of 30,000/cm2, and, when subconfluent, subjected to a 24 h starvation (CM with 2% FBS), and then exposed to oxidative stress (60 lM H2O2 in the same culture medium) for 3 and 6 h.

j Cell cytotoxicity assay (LDH release and activity) LDH release and activity from both HUVEC and EEC was assessed by the Cytotox 96 kit (Promega),

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following manufacturer’s instructions. Briefly, cells were exposed to hydrogen peroxide in 24-well culture plates, for 3 or 6 h. Released LDH in culture supernatants of both treated and control cells was measured with a coupled enzymatic assay, which results in the conversion of a tetrazolium salt (INT) into a red formazan product. The amount of formazan was determined spectrophotometrically by reading at 490 nm. Triton X-100 (0.8%v/v, provided with the kit) was used as positive control for cytotoxicity.

j Immunocytochemistry Treated and control cells, grown in chamber slides (BD Falcon), were washed with PBS and fixed in methanol for 20 min at )20C. Dried slides were then stored at )20C until use. For the immunocytochemical procedure, cells were permeabilised with 0.1% TritonX-100 in PBS. After a subsequent rinse with PBS, slides were exposed to 0.3% H2O2 in PBS, were then blocked with 1% FBS in PBS, and incubated for 2 h with the primary antibody. The detection was performed using an avidin-biotin complex kit (LSAB2, DAKO, Milan, Italy); 3.3¢-diaminobenzidine (DAB chromogenic substrate solution, DAKO) was used as developer. Nuclear counterstaining was obtained using haematoxylin (DAKO). Myeloperoxidase was identified by the use of a polyclonal antibody (MYELOp, Novocastra, Milan, Italy) and further confirmed using a monoclonal antibody (M0748, DAKO). Endothelial cells were immunostained with a monoclonal antibody anti CD31 antigen (M0823, DAKO) and using also a monoclonal antibody for wWF antigen (M0616, DAKO). 3-nitrotyrosine and 3-chlorotyrosine were identified using a mouse monoclonal (AB7048, Abcam, Cambridge, UK) and a rabbit polyclonal antibody (HP5002, Cellsciences, Canton, MA, USA), respectively. iNOS (inducible Nitric oxide Synthase) expression was assessed using a mouse monoclonal antibody (Sc-7271, Santa Cruz Milan, Italy), while eNOS (endothelial Nitric oxide Syntase) was identified using a rabbit polyclonal antibody (Sc-654, Santa Cruz). Standard negative control was routinely performed in each experiment by omitting the first antibody. Immunopositivity on HUVEC and EEC was scored using a semiquantitative approach (0 = 0% to 3 = 100% of cells positively stained). Three independent observers evaluated the immunocytochemical results and semiquantified the percentage of positive cells for each specimen. The mean value of the three percentages was considered in this study. The conversion between direct percentage and score was made in a directly proportional fashion. Ten high-power fields were examined in each culture slide and counting of the cells was performed at ·40 magnification. As

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an additional control experiment, 3-chlorotyrosine (Sigma) and 3-nitrotyrosine (Sigma) were used in direct competition assay by excess co-incubation with the primary antibody.

j Immunohistochemistry At least three tissue blocks (1 cm · 1 cm) were cut from left ventricles of four patients who underwent heart transplantation, immediately frozen in liquid nitrogen and stored at )80C until analysis. Cryostat sections were cut and immunostained for identification of CD31 antigen, MPO, nitrotyrosine and chlorotyrosine using the same primary antibodies as previously described for immunocytochemistry. Primary antibodies were revealed as previously described [35]. Immunopositivity on EEC was scored using a semiquantitative approach (0 = 0% to 3 = 100% endothelial cells positively stained). Three independent observers evaluated the immunohistochemical results and semiquantified the percentage of positive cells for each specimen. The mean value of the three percentages was considered in this study. Ten high-power fields were examined in each tissue slide and counting of the cells was performed at ·40 magnification. Standard negative control was routinely performed in each experiment by omitting the first antibody. As an additional control experiment, 3chlorotyrosine (Sigma) and 3-nitrotyrosine (Sigma) were used in direct competition assay by excess coincubation with the primary antibody. Double immunostaining (immunofluorescence) was performed by a mouse anti NT (1:50) Abcam, ab-7048 and a rabbit anti prostacyclin synthase (1:50), Santa Cruz, sc-20933 revealed by the use of a goat anti rabbit (1:100), Sigma, F 9887 and a goat anti mouse, 1:100 (Sigma T 6528). Appropriate negative controls were performed by replacement of the primary antibody with unspecific isotype-matched control antibodies.

j Total RNA extraction Total cellular RNA was isolated using the Quick Prep Total RNA Extraction Kit (GE Healthcare, Milan, Italy) following the manufacturer’s instructions. RNA yield was evaluated spectrophotometrically (A 260/ 280) and RNA aliquots were stored at )80C until use. Total RNA fractions were used for subsequent experiments only if the A260/A280 ratio exceeded 1.6.

j RT-PCR Qualitative RT-PCR was performed using the JumpStart RED HT RT-PCR kit (Sigma-Aldrich).

RT-PCR was performed mixing 2 lg of total RNA, 0.5 lg of pd(T)23, with RNAse free water. Tubes were placed in thermal cycler at 70C for 10 minutes. The reaction comprised a reverse transcription step of 50 minutes (42C), followed by inactivation of the enzyme at 95C (5 min). Then 10 pM of specific primers were added and the reactions were cycled for 94C/ 2 min, then 35 cycles of 94C/15 s, 60C/30 s, 72C/ 60 s, with a final extension at 72C/10 min. Primers used in this study were as follows: GAPDH Forward: 5¢-AAGGTGAAGGTCGGAGTCAA-3¢; GAPDH Reverse: 5¢-AAGTGGTCGTTGAGGGCAAT3¢; product size 914 bp; Beta Actin Forward: 5¢-AAACTGGAACGGTGAAGG TG-3¢; Beta Actin Reverse: 5¢-TCAAGTTGGGGGACAAAA AG-3¢: product size 350 bp; YWHAZ Forward: 5¢-TTGGCAGCTAATGGGCTCTT-3¢; YWHAZ Reverse: 5¢-TCTGTGGGATGCAAGCAAAG3¢; product size 515 bp; MPO Forward: 5¢-TGAACATGGGGAGTGTTTCA-3¢; MPO Reverse: 5¢-CCAGCTCTGCTAACCAGGAC-3¢; product size 382 bp; vWF Forward: 5¢-GGGGTCATCTCTGGATTCAA-3¢; vWF Reverse: 5¢-CAGGTGCCTGGAATTTTCAT-3¢; product size 317 bp. Beta Actin was preferred as housekeeping gene, to GAPDH and YWHAZ, for its better linearity of expression in all the experimental conditions (data not shown). The identity of PCR products was confirmed by incubation with the appropriate restriction enzyme and subsequent visualization of the cleavage products on 2% agarose gel.

j Statistical analyses Group data were expressed as mean with indication of standard error. Differences between control and H2O2 treated groups and between different treatment regimens were evaluated by the use of a non parametric test, the Mann–Whitney-U-test; values of P < 0.05 were considered as significant.

j Western blotting Western blotting was performed as described previously [7, 25]. Briefly, MPO primary antibody was a mouse monoclonal from Santa Cruz (clone 266-6K1). Secondary antibody was a rabbit anti-mouse (Amersham Biosciences). Antibody binding was revealed by chemiluminescence (Immobilion substrate, Millipore). Positive control was a HL-60 cell lysate (Santa Cruz).


Results j Cellular extraction and culturing: immunotyping of primary endothelial cells EEC and HUVEC were isolated and cultured as described above, and subjected to morphological and immunological analyses in order to check the their conformity with the endothelial phenotype, as well as the homogeneity of the cellular populations. Freshly isolated EEC and HUVEC showed an elongated morphology (Fig. 1a, b), appearing as small Fig. 1 Morphology and immunophenotyping of endothelial cells. HUVEC (a, c) and endocardial endothelial (EEC) cells (b, d) showed the classical cobblestone morphology of endothelial cells. While at 1 day after isolation (a, b) they feature a more elongated cell shape, at subsequent passages (c, d) they become more rounded, establishing lateral cell–cell contacts and growing as a continuous monolayer. Both HUVEC (e, g) and endocardial endothelial cells (f, h) used for the 1 Day present study express the typical markers of the endothelial phenotype: CD31 (e, f) and vWF (g, h). Magnifications: 20·, bar 100 l

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islets of cells on the bottom of the culture flask 24 h after isolation. At subsequent passages (Fig. 1c, d) the cells acquired the classical ‘‘cobblestone’’ morphology, becoming more rounded and forming at confluence a continuous monolayer. Immunocytochemistry for CD31 and vWF endothelial antigens was performed on cells grown on chamber slides. Almost all cells resulted positive to both markers, showing that the isolation procedure resulted in homogeneous primary cultures, with endothelial phenotype. Figure 1e–h shows the results of this analysis. HUVEC

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j Hydrogen peroxide exposure: effects on cellular viability and endothelial markers expression Cultured endothelial cells were subjected to oxidative stress by exposure to 60 lM H2O2 for a duration of 3 and 6 h. The choice of this dose was made on the basis of the existing literature reports showing that doses between 100–200 lM are apoptotic for both HUVEC and aortic endothelial cells [5, 6, 44]. Moreover, since our main aim was to evaluate the effects of oxidative stimulation on enzyme synthesis and end-products formation of endothelial cells, we chose to perform exposure times as long as 3 and 6 h. Since different groups have used exposure times ranging from 2–6 to 18–24 h [5, 12, 41], these incubation times were in agreement with previous reports of oxidative stress evaluation on endothelial cells. We performed the LDH release and activity assay as a cytotoxicity test. For HUVEC, the 3 h treatment induced a slight and unsignificant cytotoxicity (0.53%) (P = 0.2000 as determined by the Mann–Whitney test); following 6 h treatment, the increase in cellular cytotoxicity (2.68%) reached statistical significance (P = 0.0286) (Fig. 2a, b). For EEC, the same treatment resulted in a slight and unsignificant increase of cytotoxicity at both conditions (0.82%, P = 0.1143 for 3 h treatment; 1.10%, P = 0.0571 for 6 h treatment) (Fig. 2c, d). Therefore, the concentration of H2O2 used was non-lethal. Moreover, we performed immunocytochemistry to assess variations in the expression levels of CD31 and vWF. These experiments showed that the

j Hydrogen peroxide exposure results in de novo expression of myeloperoxidase in both EEC and HUVEC Immunocytochemistry for MPO expression in primary endothelial cells exposed to 60 lM H2O2 showed that in most cases untreated cells did not express the molecule (Fig. 3a, e for HUVEC and Fig. 3b, f for EEC). In contrast, EEC and HUVEC were positively stained for MPO protein after 3 h (Fig. 3c, g) and 6 h (Fig. 3d, h) treatments. It should be noted that in two of the four cell lines of EEC a few cells positive to myeloperoxidase (score: 0.02 ± 0.01) were found also in untreated cells (not shown). On the contrary, we did not find MPO-positive cells in any of the untreated HUVEC specimens. This result is most probably related to the fact that EEC were from CHFaffected hearts, in which the altered microenvironmental conditions may severely modify endothelial phenotype. This results in the maintenance of the ectopic expression of MPO in some cells also after primary culture. In addition, as shown below, this low expression rate after three culture passages is in agreement with the relatively low expression of MPO in endocardium in vivo as assessed by immunohistochemistry. Furthermore, as depicted in supplemental figure 1, 6 h treatment showed a significant MPO increase compared to 3 h treatment for HUVEC HUVEC 3h

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Fig. 2 Effects of exposure to hydrogen peroxide on viability of endothelial cells. LDH assay was used to evaluate the cytotoxic effect of 60 lM H2O2. Data are represented as means, with the indication of standard error of three replicate experiments. Significance of differences with respect to untreated cells was calculated using the Mann–Whitney U-test. The analysis was performed for both HUVEC (a, b) and endocardial endothelial (EEC) cells (c, d), subjected to oxidative stress for 3 h (a, c) and 6 h (b, d). A slightly but significantly decreased viability was observed in 3 h H2O2 exposed HUVEC cells. On the contrary, 6 h H2O2 exposure significantly increased proliferation of EE cells

expression of the two markers was not affected by the applied stress (not shown).

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G. La Rocca et al. Oxidative-driven endothelial expression of myeloperoxidase Fig. 3 Oxidative stress causes the expression of myeloperoxidase by endothelial cells. HUVEC (a, c, e, g, on the left) and endocardial endothelial cells (b, d, f, h, on the right) were stimulated with hydrogen peroxide for 3 h (a–d) or 6 h (e–h). MPO was not detectable in untreated endothelial cells - either HUVEC (a, e) or EE cells (b, f)—at both 3 or 6 h experiments. When HUVECs were subjected to oxidative stress, MPO was detectable with a Control net cytoplasmatic positivity after 3 h (c) and 6 h (g) 3h stimulation (arrows). The same occurred for EE cells, at both 3 h (d) and 6 h (h) treatments (arrows). Magnifications: 20·, bar 100 l

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alone (score: 0.32 ± 0.06 vs. 0.16 ± 0.6 for HUVEC, P = 0.002; 0.28 ± 0.07 vs. 0.25 ± 0.1 for EEC, P = 0.588). These results showed that endothelial cells were able to produce endogenous MPO in a neutrophil-free system, following exposure to non-lethal doses of H2O2. To assess the effect of heparin on mobilisation of endothelial-derived MPO, suggested ‘‘in vivo’’ by recent studies [11], endothelial cells were incubated in a modified CM without heparin, starved and exposed to H2O2 for both 3 and 6 h. Immunocytochemistry showed that heparin removal did not

cause any variation to the number of MPO-positive cells (not shown).

j Hydrogen peroxide exposure results in the formation of end products of tyrosine oxidation: 3-chlorotyrosine in HUVECs and endocardial endothelial cells Since MPO is the only enzyme able, in the presence of hydrogen peroxide, to produce HOCl at physiological

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halide concentrations [20], we checked for the presence of 3-chlorotyrosine, one of the early and stable end products of MPO activity, which is used as a molecular fingerprint of the enzyme activity [35]. Immunocytochemistry for 3-chlorotyrosine showed that untreated cells, after three passages in culture, were negative, while a strong cytoplasmatic positivity was observed in both HUVEC (Fig. 4c, g) and EEC (Fig. 4d, h). As depicted in supplemental figure 2, there was a a five to sevenfold increase in 3-chloFig. 4 Endothelial cells exposed to hydrogen peroxide expressed 3-chlorotyrosine concomitantly to MPO expression. HUVEC (a, c, e, g, on the left) and endocardial endothelial cells (b, d, f, h, on the right) were stimulated with hydrogen peroxide for 3 h (a–d) or 6 h (e–h). 3-chlorotyrosine was not detectable in untreated endothelial cells—either HUVEC (a, e) or EE cells (b, f)—at either exposure time. When HUVECs were subjected to oxidative stress, 3-chlorotyrosine was Control recognisable with a strong cytoplasmatic positivity 3h after 3 h (c) and 6 h (g) stimulation (arrows). The same was true for endocardial endothelial cells, at both 3 h (d) and 6 h treatments (h) (arrows). Magnifications: 20·, bar 100 l

rotyrosine accumulation after 6 h compared with 3 h treatments, for both cell types (score: 1.62 ± 0.3 vs. 0.22 ± 0.07 for HUVEC, P = 0.002; 1.5 ± 0.36 vs. 0.28 ± 0.13 for EEC, P = 0.002). In the same conditions, no positive cells were detected after nitrotyrosine immunostaining following H2O2 exposure by EEC or HUVEC (not shown). In order to determine the potential contributions of NO synthases in this in vitro system, we performed immunocytochemistry to localize iNOS and eNOS HUVEC

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G. La Rocca et al. Oxidative-driven endothelial expression of myeloperoxidase Fig. 5 Endothelial cells exposed to oxidative stress express MPO mRNA after 3 h treatment. RT-PCR experiments were performed on total RNA extracted from HUVEC (a–c) and EEC (d–f). HUVEC expressed beta-actin (a) and vWF (c) both in untreated and hydrogen peroxide-exposed cells. MPO mRNA (382 bp) (b) was only detectable after exposure to oxidative stress. The results were confirmed in EEC, where expression of beta actin (d) and vWF (f) mRNAs was detected in both untreated and stressed cells. Moreover, as shown in e, also EEC expressed MPO, but only after exposure to hydrogen peroxide. M DNA ladder, C control cells; 60 lM: cells exposed to oxidative stress. The expression of MPO has been evaluated by a semiquantitative densitometric analysis of the expression in HUVEC (g) and EEC (h). Data are represented as mean, with the indication of standard deviation, of three replicate experiments. Values were normalized for beta actin expression

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expression following oxidative stress. iNOS expression was not detectable in either HUVEC or EEC following hydrogen peroxide exposure, nor it was detected in untreated cells (not shown). It is possible that this is due to the absence, in our in vitro system, of proinflammatory mediators capable of inducing iNOS expression. On the other hand, and confirming past literature results from other groups [5, 12], eNOS was amply expressed by untreated HUVEC and EEC, and its expression increased still more after oxidative stress (data not shown).

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j Detection of the mRNA for MPO in H2O2 exposed endothelial cells To demonstrate the synthesis of MPO in EEC and HUVEC following H2O2 exposure, we extracted total RNA from cells, and, following reverse transcription, end-point PCR was performed using gene-specific primers. Endothelial cells expressed the mRNAs for b-Actin and vWF both in treated and control cells (Fig. 5). The presence of MPO mRNA was observed only in treated endothelial cells (both EEC and

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HUVEC), while the 382 bp product was not detectable in control cells (Fig. 5). Further experiments were made using 40 and 45 cycles, but MPO messenger was never detectable in untreated cells. Semiquantitative determination of MPO expression levels was performed as reported previously [25]. Figure 5 shows the graph of levels of MPO messenger RNA in HUVEC and EEC after 3 h hydrogen peroxide incubation, determined after normalization with beta-actin.

j Immunohistochemistry of endocardial tissue from patients with post-infarct chronic heart failure In the endocardial heart tissue, immunopositivity of EEC was evaluated and scored for CD31 antigen, myeloperoxidase (MPO), 3-nitrotyrosine and 3-chlorotyrosine. A mean length of 4,800 ± 577 l of endothelial layer was analysed for each quantification. CD 31 antigen was uniformly expressed by all EE cells (score: 3 ± 0 (mean ± SD) (Fig. 6a). Myeloperoxidase was expressed occasionally by EE cells (score: 0.15 ± 0.1; Fig. 6b). 3-nitrotyrosine was frequently expressed by EE cells (score: 0.9 ± 0.48; Fig. 6c). 3-chlorotyrosine was amply expressed by EE cells (score: 2 ± 0.4; Fig. 6d). Endocardial capillary vessels and heart muscle were also frequently immunostained for 3-nitrotyrosine and more extensively, for 3-chlorotyrosine. Inflammatory cells around capillary vessels and infiltrating heart muscle were also occasionally immunostained for 3-nitrotyrosine and more frequently, for 3-chlorotyrosine. Fig. 6 Photomicrographs showing endocardial tissue from a patient with chronic heart failure immunostained for identification of a CD31 antigen, costitutively expressed by endothelial cells (arrows), b myeloperoxidase, randomly expressed by endothelial cells (arrows), c nitrotyrosine, moderately expressed by endothelial cells (arrows) and d 3-chlorotyrosine, more frequently expressed by endothelial cells (arrows). Results are representative of those from four patients who underwent heart transplantation after development of post-infarctual chronic heart failure. Magnification, 20·, bar 50 l

Moreover, since prostacyclin synthase is one of the most prevalent nitrated proteins [34, 50], we aimed to detect, by co-staining in immunofluorescence experiments, if this protein should be one of the potential targets of nitration in post-infarct CHF hearts. Supplemental figure 3 shows the panels of immunolocalization of prostacyclin synthase and 3-nitrotyrosine in a representative sample of human heart. The merged image (panel B) shows that, even if prostacyclin synthase should not be considered the sole nitrated protein, in almost three areas of the endocardial surface (arrows) there is a clear colocalization.

Discussion This study shows for the first time that EEC from patients with CHF, as well as HUVEC, following exposure to H2O2 express the enzyme myeloperoxidase together with 3-chlorotyrosine, an unphysiologic and specific end product of oxidative stress. These ‘‘in vitro’’ data were confirmed in ‘‘in vivo’’ experiments where MPO, 3-nitrotyrosine and 3-chlorotyrosine were identified in endocardium from patients with CHF. Myeloperoxidase is a key enzyme in the immune defense system against bacterial pathogens, on account of its capability to generate powerful oxidants such as HOCl by using hydrogen peroxide and Cl) ion at physiological levels [20, 23, 29, 35]. Moreover, protein nitration is also possible through activation of heme peroxidases, such as myeloperoxidase, by


H2O2 promoting oxidation of NO2). Other oxidants that can oxidise NO2) to nitrating intermediates include HOCl, the product of MPO-catalyzed Cl) oxidation [36, 45]. MPO has been implicated in the development of a number of cardiovascular diseases as CHF [11, 37, 40]. Moreover, several reports provide evidence of a significant association between circulating MPO levels and the risk of adverse cardiovascular events [42]. In addition, MPO has been directly linked to the NO homeostasis in the vessel wall since it is able to interact with NO to form nitrated species, thus depleting the intracellular reservoir of NO, leading to vasoconstriction or tissue damage [2, 45, 46]. Nitrosative stress due to tyrosine nitration may be exacerbated by oxidative stress. In fact, stimuli that lead to iNOS induction may also up-regulate oxidases (Xanthine Oxidase, NADPH oxidase), and concomitant elevations of NO and superoxide may lead to formation of peroxynitrite [4, 8, 14]. Nitration of tyrosine in proteins is relevant in heart failure, since this mechanism may interfere with NO redistribution in the sarcolemma altering the NO/redox balance [9, 10, 22, 38, 39]. Endothelial cells can actively interact with endogenous (e.g. neutrophil-derived) MPO, as demonstrated recently by the molecule’s binding on the surface of endothelial cells as well as by transcytosis towards the subintimal side [1, 42, 44]. In the present study, we isolated primary human endothelial cells from two different sources, e.g. EEC from the left ventricle of post infarcted CHF diseased patients and HUVEC cells from the umbilical vein. The former represent organ-specific primary cell lines, coming from a diseased organ targeted by the inflammatory processes characteristic of the disease. As reported in the Results, the isolation protocol used allowed to obtain homogeneous populations of endothelial cells. They were subjected to oxidative stress, using a non-lethal dose (60 lM) which was in the range of doses previously used in oxidative stress studies on endothelial cells. [6, 12]. Concerning the exposure times we used (3 and 6 h), this choice was made based on the rationale of measuring not simply the de novo expression of MPO in endothelial cells, but also the ‘‘long-term’’ effects of oxidative stress due, for example, to the accumulation of detectable end products as 3-chlorotyrosine. Previous studies suggest that exposure times ranging between 2–24 h should be used to investigate the pathways related to oxidative stress in endothelial cells [5, 12, 41]. Cytotoxicity tests confirmed that only in HUVEC, after 6 h exposure to hydrogen peroxide, but not in EEC, was there a significant increase of cellular cytotoxicity following H2O2 exposure. This is also in agreement

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with the recently reported evidences that endothelial cells do not show a significant increase in cell death when exposed to doses of hydrogen peroxide less than 100 lM [6, 43]. The innovative finding of this paper is that endothelial cells subjected to oxidative stress showed they are able to produce endogenous MPO in an in vitro system deprived of neutrophils and other potential sources of MPO. This result was also achieved notwithstanding starvation with lowered serum medium. In fact, since both control and treated cells were incubated (and starved) in a reduced-serum culture medium, uptake of MPO from the bovine serum can be excluded. As reported recently, untreated endothelial cells are able to bind MPO [44], but the lack of immunopositivity in untreated control cells suggests that H2O2 exerts a direct regulatory effect on the expression of MPO in the endothelium. As shown by immunocytochemistry and successively confirmed by RT-PCR, human endothelial cells from both umbilical vein and endocardium expressed MPO when exposed to 60 lM H2O2. The number of immunocytochemistry-positive cells was higher in endocardial cells than HUVEC. The sole presence of myeloperoxidase is not sufficient, per se, to provoke the important oxidative phenomena of proteins and lipids in the cardiovascular system. MPO plays its fundamental role in the metabolism of H2O2, which is used to produce more powerful oxidants such as HOCl [20]. Then HOCl is able to oxidise different substrates, leading for example to the formation of chlorinated amino acids, of which 3-chlorotyrosine constitutes a detectable and specific nonphysiologic end-product [35, 47]. Therefore, we looked for the presence of 3-chlorotyrosine in endothelial cells exposed to oxidative stress, using a novel commercial anti-chlorotyrosine antibody. Immunocytochemistry revealed the presence of a strong positivity only in treated cells, while no positivity was detectable in untreated cells, either EEC or, HUVEC demonstrating that in our ‘‘in vitro’’ model, MPO upregulation is associated to a specific end product of MPO-driven oxidative stress such as 3chlorotyrosine. Another interesting point is the absence of nitrotyrosine, in cultured endothelial cells exposed to H2O2; this molecular specie should be formed in the presence of H2O2 and MPO via a two step reaction of NO oxidation. Since macrophage-derived iNOS rather than eNOS is the main enzyme involved in NO production during the oxidative burst, it could be argued that in our in vitro system MPO action resulted in the formation of 3-chlorotyrosine rather than nitrotyrosine, also for the presence of lower levels of NO with

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respect to those achievable during inflammation [28, 33]. ‘‘In vivo’’ immunohistochemistry experiments showed that endocardium from left ventricle of postinfarct CHF patients, expressed a small amount of MPO together with moderate levels of nitrotyrosine, as already reported in the literature [1]. Interestingly, abundant levels of 3-chlorotyrosine were detected, thus confirming the ‘‘in vitro’’ observations reported for primary EEC. In order to better clarify and summarize our findings, we propose a model (depicted in Fig. 7) describing the possible pathway of coupling between endogenous MPO production and 3-chlorotyrosine accumulation in endothelial cells. In particular, the formation of HOCl due to MPO action should result also in L-arginine chlorination, therefore uncoupling eNOS and favouring the production of superoxide and the alteration of NO/ROS balance. Interestingly, eNOS uncoupling should also be a consequence of BH4 depletion, a process in which DHFR (dihydrofolate reductase) regulation plays a key role: downregulation of DHFR by hydrogen peroxide limits recycling of oxidized BH4, therefore limiting the presence of the essential cofactor in NO biosynthesis [26, 32].

Fig. 7 Proposed working model in endothelial cells of MPO action following oxidative stress. Oxidative stress (H2O2) upregulates MPO which, in the presence of hydrogen peroxide and chloride ions, catalyzes hypochlorous acid formation. HOCl-dependent oxidation may result in tyrosine chlorination of proteins, leading to 3-chlorotyrosine formation. In addition, HOCl may chlorinate L-arginine, thus ‘‘uncoupling’’ eNOS. Uncoupled eNOS may become a source of superoxide ion (O2)), which can be readily dismutated by SOD to produce again hydrogen peroxide and close the circuit. Oxidative-driven eNOS increase may further contribute to the ‘‘uncoupling’’ process. Moreover, eNOS uncoupling should also be secondary to BH4 (tetrahydrobiopterine) depletion, due to the inhibitory effect of hydrogen peroxide on DHFR activity. Finally, the levels of NO should be rapidly lowered by the interaction with superoxide, with generation of peroxynitrite (ONOO)) which can rapidly cause protein nitration at tyrosine residues. Graph proposed on the basis of our present data and references [6, 26, 27, 32, 35, 44, 48]

Finally, prostacyclin synthase appeared in vivo as one of the prominently nitrated proteins, as shown by immunofluorescence experiments. This datum should be of great clinical relevance, since tyrosine nitration of prostacyclin synthase by peroxynitrite might be an important pathophysiological event, since it not only decreases the production of prostacyclin but can further modify endothelial features, leading to increased endothelial cell apoptosis and adhesion molecule expression [34, 50]. In conclusion, our study highlights that EEC, as well as HUVEC, in the presence of non-lethal doses of hydrogen peroxide similar to those achievable in vivo, can synthesize endogenous myeloperoxidase, together with an end product of oxidative stress such as 3-chlorotyrosine. This response to oxidative stress may actively contribute to the vascular and cardiac damage in patients with various vascular and cardiac diseases including post-infarct CHF. We can only speculate on the mechanism underlying MPO induction in endothelial cells following hydrogen peroxide exposure. Further experiments are needed to establish if a direct regulatory mechanism (such as that implied in upregulation of VEGF-A) or an indirect one (such as NFkB-induced eNOS upregu-

3-Cl-tyr accumulation

MPO induction

Oxidative Stress (H2O2)

MPO Tyrosine chlorination CIDHFR

eNOS increase

H2O2 HOCI

O2BH4

“uncoupled” eNOS

SOD

L-arg chlorination

O2- + NO •

ONOO-

Tyrosine nitration

3-N-tyr accumulation


lation) are in action in endothelial cells exposed to oxidative stress [37, 49]. The uncovering of this regulatory mechanism should open new ways in clinical intervention to reduce the MPO-driven CHF burden.

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j Acknowledgments We thank Rosemary Allpress for her revision of the English language. We thank Antonella Chiara, Santina Di Gangi and Giusy Scaduto for the valuable support during the experimental work. This work was supported by Fondazione S. Maugeri, IRCCS, Ricerca corrente, and Italian Ministry of University (Ex 60% to GZ, FC, FF) grants. The authors declare no conflict of interests.

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