Telomere Dysfunction Causes Sustained Inflammation in Chronic

Obstructive Pulmonary Disease

Valerie Amsellem1, Guillaume Gary-Bobo

1, Elisabeth Marcos

1, Bernard Maitre

1, Vicky

Chaar1, Pierre Validire

2, Jean-Baptiste Stern

3, Hiba Noureddine

1, Elise Sapin

1, Dominique

Rideau1, Sophie Hue

4, Sabine Le Gouvello

4,

Jean-Luc Dubois-Randé1, Jorge Boczkowski

1, Serge Adnot

1

1INSERM U955 and Département de Physiologie-Explorations Fonctionnelles, Hôpital Henri

Mondor, AP-HP, 94010, Créteil, France

2 Institut Mutualiste Montsouris, Département anatomopathologie, Paris, France

3 Institut Mutualiste Montsouris, Département thoracique, Paris, France

4INSERM U955 and Service d’immunologie biologique, Hôpital Henri Mondor, AP-HP,

Créteil, France

Correspondence should be addressed to Serge Adnot, Hôpital Henri Mondor, Service de

Physiologie-Explorations Fonctionnelles, 94010, Créteil, France

Tel: +33 1 49 81 26 77; Fax: +33 1 49 81 26 67; E-mail: serge.adnot@inserm.fr

Funding: This study was supported by grants from the INSERM, Délégation à la Recherche

Clinique de l’AP-HP, Fondation pour la Recherche Médicale (FRM) and CARVSEN

foundation. Jorge Boczkowski was supported by the INSERM and Assistance Publique-

Hôpitaux de Paris (Contrat Hospitalier de Recherche Translationnelle).

Running head: Telomere Dysfunction and Inflammation in COPD

Descriptor number: 3.04 (endothelium)

Word count (body of text only): 3459

Page 1 of 53 AJRCCM Articles in Press. Published on September 1, 2011 as doi:10.1164/rccm.201105-0802OC

Copyright (C) 2011 by the American Thoracic Society.

Contributions of each author:

Amsellem V: design and conduct of cell culture studies, interpretation of the results

Gary-Bobo G: pulmonary vessel morphometry and immunohistochemical analyses,

mouse studies, interpretation of the results

Marcos E: technical advice, telomere length studies, interpretation of the results

Maitre B: recruitment of the patients, interpretation of the results

Chaar V: cell culture studies, interpretation of the results

Validire P: recruitment of the patients, informed consent of the patients

Noureddine H: technical advice, interpretation of the results

Stern JB: recruitment of the patients, informed consent of the patients

Sapin E: isolation and culture of endothelial cells

Rideau D: technical advice, interpretation of the results

Hue S: Luminex studies, interpretation of the results.

Le Gouvello S: quantitative PCR studies, interpretation of the results.

Dubois-Rande JL: design of the study, interpretation of the results and revision of the

manuscript

Boczkowski J: design of the study, interpretation of the results, and revision of the manuscript

Adnot S: design of the study, interpretation of the results, and writing the manuscript

This article has an online data supplement, which is accessible from this issue's table of

content online at www.atsjournals.org

Page 2 of 53

AT A GLANCE COMMENTARY

Scientific knowledge on the subject

Chronic obstructive pulmonary disease (COPD) is characterized by chronic inflammation,

which contributes to the pathogenesis of the lung disease and development of co-morbidities.

The mechanisms underlying sustained inflammation in COPD remain unknown.

What this study adds to the field

Telomere dysfunction and senescent pulmonary vascular endothelial cells having altered gene

expression profiles contribute to sustained inflammation in COPD. Telomere dysfunction in

mice leads to increased lung cytokine levels in the absence of external stimuli.

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ABSTRACT

Rationale: Chronic obstructive pulmonary disease (COPD) is associated with chronic

inflammation of unknown pathogenesis.

Objectives: To investigate whether telomere dysfunction and senescence of pulmonary

vascular endothelial cells (P-ECs) induce inflammation in COPD.

Methods: Prospective comparison of patients with COPD and age- and sex-matched control

smokers. Investigation of mice null for telomerase reverse transcriptase (Tert) or telomerase

RNA component (Terc) genes.

Measurements and Main Results: In-situ lung specimen studies showed a higher percentage

of senescent P-ECs stained for p16 and p21 in patients with COPD than in controls. Cultured

P-ECs from COPD patients exhibited early replicative senescence, with decreased cell-

population doublings, a higher percentage of beta-galactosidase-positive cells, reduced

telomerase activity, shorter telomeres, and higher p16 and p21 mRNA levels at an early cell

passage compared to controls. Senescent P-ECs released cytokines and mediators: the levels

of IL6, IL8, MCP-1, Hu-GRO, and sICAM-1 were elevated in the media of P-ECs from

patients compared to controls at an early cell passage, in proportion to the senescent P-EC

increase and telomere shortening. Upregulation of MCP-1 and sICAM-1 led to increased

monocyte adherence and migration. The elevated MCP-1, IL8, Hu-GROα, and ICAM-1 levels

measured in lungs from patients compared to controls correlated with P-EC senescence

criteria and telomere length. In Tert-/- and/or Terc-/- mouse lungs, levels of the corresponding

cytokines (MCP-1, IL8, Hu-GROα and ICAM-1) were also altered, despite the absence of

external stimuli and in proportion to telomere dysfunction.

Conclusion: Telomere dysfunction and premature P-EC senescence are major processes

perpetuating lung inflammation in COPD.

Keywords: inflammation, senescence, COPD

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INTRODUCTION

Chronic inflammation is a prominent feature of chronic obstructive pulmonary disease

(COPD) (1-3). An exaggerated inflammatory response of the airways to chronic irritants,

primarily cigarette smoke, is considered the main cause of COPD. The inflammatory process

results in remodeling of the airways and destruction of the lung parenchyma. After smoking

cessation, the inflammation persists and the levels of proinflammatory cytokines in the lungs

and bloodstream remain high in patients with COPD, even those with mild and stable forms

of the disease. This persistent inflammation may not only constitute a major driver of COPD

progression, but also contribute to the development of systemic complications of adverse

prognostic significance such as cardiovascular disease, weight loss, bone demineralization,

and muscle dysfunction (3-5). Thus, elucidating the mechanisms that underlie persistent

inflammation in COPD would probably produce major clinical benefits.

COPD is an age-related disease, and senescent cells are known to accumulate within

tissues with advancing age (6, 7). Somatic cell senescence occurs either when the replicative

potential is exhausted or in response to excessive extracellular or intracellular stress (7). Both

forms of senescence may be accelerated in COPD: premature replicative senescence may

result from increased telomere shortening (8-10) and premature stress-related senescence

from non-telomeric signals triggered by oxidative stress due primarily to cigarette smoke (11,

12). Telomeric signals are mediated chiefly via the p53-p21 pathway and non-telomeric

signals via the p16-retinoblastoma protein pathway (12). Increased numbers of p21- and p16-

stained cells have been found in the lungs of patients with emphysema compared to control

smokers, as well as at sites affected with other age-related diseases such as osteoarthritis and

atherosclerosis (6, 13). The pathogenic role played by presenescent or senescent cells in lungs

of patients with COPD remains unclear. Senescent cells that survive in vivo not only lose a

number of functions, but also acquire many changes in the expression of genes encoding

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various cytokines, proteases, and growth factors, which may affect their microenvironment

(14-16). In recent studies, we found that senescent smooth muscle cells (SMCs) were

increased in the media of remodeled vessels from patients with COPD and were located in

close proximity to actively dividing cells in the neointima. Interplay between these two cell

subsets was demonstrated in vitro by showing that senescent-SMC-conditioned media

containing cytokines caused proliferation of healthy target SMCs (17). The effects of

senescent cells on their environment may vary, however, across cell types and secreted

proteins (15). A major role of endothelial cells is to promote and maintain inflammation via

the expression of surface adhesion molecules and secreted proteins (18, 19). Our working

hypothesis here was that increased senescent pulmonary vascular endothelial cell (P-EC)

counts underlie the sustained lung inflammation in COPD. In patients and age- and sex-

matched control smokers, who were different from those investigated previously (17), we

examined lung specimens and derived cultured P-ECs to assess their susceptibility to

premature senescence and to determine whether altered P-EC functions were related to lung

inflammation. To evaluate whether telomere dysfunction per se induced lung inflammation,

we investigated mice null for the telomerase reverse transcriptase (Tert) or telomerase RNA

component (Terc) genes and not challenged by external stimuli. Part of the results of these

studies has been reported previously in abstract form (20).

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MATERIAL AND METHODS

Protocol

We prospectively recruited 31 patients undergoing lung resection surgery for localized

lung tumors at the Institut Mutualiste Montsouris (Paris, France). Among them, 16 had COPD

and 15 were control smokers matched to the COPD patients on age and sex (Table 1).

Inclusion criteria for the patients with COPD were an at least 10-pack-year smoking history

and a ratio of forced expiratory volume in 1 second (FEV1) over forced vital capacity (FVC)

<70%. The controls had to have a similar smoking history and an FEV1/FVC ratio greater

than 70%; none of the patients or controls had chronic cardiovascular, hepatic, or renal

disease or a history of cancer chemotherapy (see online data supplement).

The study was approved by the institutional review board of the Henri Mondor

Teaching Hospital. All patients and controls signed an informed consent document before

study inclusion.

Lung tissue samples collected during surgery were used for P-EC isolation, in situ

immunohistochemical studies, protein level determinations, and telomere length

measurements.

Laboratory investigations

Senescent P-ECs from peripheral pulmonary vessels were identified by double staining

with von Willebrand factor and p16 or p21 (21) (see online data supplement). Repeated

passaging of cultured P-ECs (22) from patients with COPD and controls was performed to

determine the replicative senescence threshold and cell population doubling level (PDL). P-

ECs from patients with COPD and controls were studied and compared at passage 6 and at

senescence. Subsequent analyses assessed two main criteria: PDL and the percentage of cells

with acid beta-galactosidase (β-gal) activity. Telomere length measurement using RT-qPCR

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was performed as previously described (10). Telomerase activity was quantified using the

teloTAGGG telomerase PCR Elisaplus kit (Roche Diagnostics, Meylan, France). Levels of 26

soluble factors released by P-ECs were evaluated using the Luminex®

system, and selected

factors from lung tissues were evaluated using ELISAs (see online data supplement).

Functional studies assessing monocyte migration and adhesion to P-ECs were

performed using transwell migration and monocyte adhesion assays (see online data

supplement).

Mouse studies

Tert-/- mice (Jackson laboratories, Sacramento, CA, USA) and Terc-/- mice (a gift from

Dr. Manuel Serrano, Madrid, Spain) were intercrossed to produce successive generations of

telomerase-deficient mice with decreasing telomere length. Lungs from third generation (G3)

Tert-/- and second generation (G2) Terc-/- mice were studied (23, 24).

Statistical analysis

Data are described as mean±SEM. The unpaired t-test was used to compare patients

with COPD and controls, least-square linear regression to assess correlations between

variables, and the paired t test to evaluate the effects of senescence in cells from patients with

COPD and controls. One-way ANOVA was used to compare Tert-/-, Terc-/-, and wild-type

mice. P values less than 0.05 were considered significant. Data were analyzed using

GraphPad Prism statistical software 5.0 (San Diego, CA, USA).

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RESULTS

Characteristics of patients with COPD and controls

The clinical features of the patients with COPD and controls are reported in Table 1.

The control group did not differ from the COPD group regarding the sex ratio, age, smoking

history, or body mass index. The emphysema score was low in both groups but was higher in

the patients with COPD than in the controls.

In situ analysis of p16- and p21-stained cells in lung vessels from patients with COPD

and controls

Senescent endothelial cells identified as p21- or p16-stained cells were co-stained for

von Willebrand factor (vWF) (Figure 1). The percentage of endothelial cells positive for p16

or p21 was considerably higher in large and small pulmonary vessels from patients with

COPD compared to those from controls (Figure 1A and 1B).

Replicative senescence of pulmonary vascular endothelial cells (P-ECs) from patients

with COPD and controls

More than 95% of cultured cells were endothelial cells from blood vessels (Figures E1A

and E1B). Figure 2 shows that senescence started earlier in cells from patients with COPD

than in those from controls, leading to a lower PDL in the patients with COPD (Figure 2A and

B). Patients with COPD had a higher percentage of acidic β-gal-positive cells at passage 6

than the controls; this percentage increased with subsequent passages and was similar in

patients and controls at the stage of cell senescence (Figures 2C and D). When we pooled the

patients with COPD and the controls, we found that PDL correlated positively with FEV1

(r=0.42; P<0.01) and negatively with the emphysema score (r=-0.46; P<0.01).

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Telomere length; telomerase activity; and p16, p53, and p21 mRNA levels from patients

with COPD and controls

Telomerase activity was detectable only at early cell passages and was significantly

lower in patients with COPD than in controls at passage 4 (Figure 3). Telomeres, which were

shorter in passage-4 P-ECs from patients with COPD than in those from controls, continued

to shorten during replicative senescence but had similar lengths in patients with COPD and

controls at senescence (Figure 3). Of note, telomere length at passage 4 correlated positively

with PDL (r=0.49, P<0.01). The p16, p53, and p21 mRNA showed differences in the opposite

directions: they were higher in P-ECs from patients with COPD than in those from controls at

passage 4 then increased during replicative senescence and did not differ between patients and

controls at senescence (Figure 3). Similar changes in phospho-p53(Ser15) protein levels were

observed (Figure E2).

Factors secreted by P-ECs from patients with COPD and controls during replicative

senescence

The amounts of several cytokines and growth factors released by P-ECs were measured

in conditioned media from patients with COPD and controls at passage 6 and at senescence.

Of the 26 soluble factors analyzed by the Luminex®

assay, nine were detectable in P-EC-

conditioned media: IL6, IL8, Hu-GRO, MCP-1, RANTES, sICAM-1, PAI-1, PDGF, and

FGF2. As shown in Figure 4, all these soluble factors except FGF-2 increased from passage 6

to senescence in P-ECs from controls. In P-EC-conditioned media from patients with COPD,

in contrast, no consistent increase was seen during replicative senescence, due to the already

high levels of most of these factors at passage 6. Indeed, the levels of IL6, IL8, Hu-GRO,

MCP-1, and sICAM-1 were higher in the media from patients with COPD than in those from

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controls at passage 6, whereas these levels were no longer different in the two groups at

senescence.

Effects of P-EC conditioned media on monocyte migration: role for MCP-1

We found that conditioned media from passage-6 P-ECs from patients with COPD

markedly stimulated the migration of monocytes and that this effect was not observed with

the media of passage-6 P-ECs from controls. In contrast, a similar effect was observed in

response to P-EC-conditioned media from patients with COPD and controls at replicative

senescence. We then evaluated whether the difference between patients and controls was due

to the chemoattractant activity of MCP-1. Adding the MCP-1-neutralizing antibody to the

conditioned media completely abolished the chemoattractant effect of passage-6 P-ECs from

patients with COPD, as well as that of senescent cells from patients with COPD and controls

(Figure 5A and 5B).

Monocyte adhesion to P-ECs, role for ICAM-1

Testing monocyte adhesion to P-ECs at an early cell passage revealed that P-ECs from

patients with COPD, but not those from controls, were able to recruit U937 monocytes at the

cell surface (Figure 6). In contrast, monocytes showed similar adhesion to P-ECs from

patients with COPD and controls at senescence. We then evaluated whether the difference

between patients and controls was due to the surface adhesion molecule ICAM-1. Adding

ICAM-1-neutralizing antibody completely abolished the adhesion of U937 monocytes to

passage-6 cells from patients with COPD and to senescent P-ECs from patients with COPD or

controls (Figure 6).

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Cytokine levels and telomere length in lung tissues from patients with COPD and

controls

We investigated whether the differences in IL6, IL8, MCP-1, Hu-GRO, and ICAM-1

levels in P-EC-conditioned media between patients with COPD and controls were replicated

in lung tissue extracts from the same individuals and were associated with telomere

shortening. We found that the levels of IL6, IL8, MCP-1, ICAM-1, and Hu-GROα were

higher and that telomeres were shorter in lung tissues from patients with COPD than in those

from controls (Figure 7A). In addition, the number of CD68-positive macrophages was higher

in lung sections from patients with COPD than from controls (Figure E3).

Interestingly, the levels of IL8, MCP-1, ICAM-1, and Hu-GROα, but not of IL6,

correlated negatively with telomere length measured in lung tissues (Figure 7B). Lung

cytokine levels namely IL8, MCP-1 and Hu-GROα also correlated negatively with the PDL of

cultured P-ECs (r=-0.51, P<0.05; r=-0.49, P<0.05; and r=-0.43, P<0.05, respectively).

Cytokine levels and telomere length in lung tissues from Terc-/- and Tert

-/- mice

These studies were performed to evaluate whether telomerase deficiency and

subsequent telomere shortening were associated with increased lung cytokine levels in the

absence of external stimuli. Both G2 Terc-/-

and G3 Tert-/-

mice exhibited lung telomere

dysfunction compared to wild-type mice, but G2 Terc-/-

mice had shorter lung telomeres than

G3 Tert-/-

mice, a finding reported previously in the liver and heart (25) (Figure 8A).

Compared to wild-type mice, G2 Terc-/-

mice exhibited increased lung tissue levels of MIP-2,

CXCL1 (corresponding to human IL8 and Hu-GROα, respectively), MCP-1 and ICAM-1

whereas G3 Tert-/-

mice exhibited increases only in MCP-1 and IL6. Thus, except for IL6,

lung cytokines were more consistently elevated in the G2 Terc-/-

mice, which also exhibited

greater levels of telomere dysfunction. Interestingly, individual values of lung MCP-1

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correlated inversely with lung telomere length in the overall population of mice from the three

groups (Figure 8 C).

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DISCUSSION

Our data support a major role for telomere dysfunction and P-EC senescence in the

chronic inflammation that characterizes COPD. The number of senescent P-ECs in the lungs

was higher in patients with COPD than in controls, and cultured P-ECs from patients with

COPD displayed premature replicative senescence due to decreased telomerase activity with

telomere shortening. Premature senescence of P-ECs from patients with COPD was

associated with marked overexpression of major proinflammatory cytokines and adhesion

molecules, which affected monocyte adherence and migration. Together with the elevated

cytokine levels in the lungs from patients with COPD compared to controls and the

correlation of these levels with telomere length and in vitro P-EC senescence criteria, these

findings indicate that lung inflammation in patients with COPD was directly linked to the

process of premature P-EC senescence. That increased lung cell senescence was sufficient to

cause inflammation was further supported by studies in telomerase-deficient mice showing

increased lung cytokine levels in proportion to the decrease in telomere length, even in the

absence of external stimuli.

The deleterious effect of chronic inflammation in patients with COPD is now supported

by many studies showing that inflammation contributes not only to the pathogenesis of

COPD, but also to the development of co-morbidities including cardiovascular complications,

which adversely affect the prognosis (2-5, 26). As highlighted in a recent review, in COPD

and in other chronic diseases, the problem with inflammation is not how it starts, but how it

fails to subside (27). It is well established that patients with mild COPD, even in the absence

of lung infection and long after smoking cessation, still exhibit higher circulating cytokine

levels than heavy smokers without COPD (10, 28). That damage to uninfected lungs in COPD

promotes the inflammatory process was recently suggested by studies showing that lung

volume resection surgery for emphysema significantly reduced circulating inflammatory

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mediators in COPD (29). Here, we reasoned that lung cellular mechanisms perpetuating

inflammation were related to cell senescence.

To address this hypothesis, we focused on P-ECs. We investigated lung specimens and

derived cultured P-ECs from patients with COPD and from sex- and age-matched control

smokers undergoing lung surgery. In situ studies of lung sections showed an increased

percentage of von Willebrand P-ECs stained for p21 and p16 in patients with COPD

compared to controls, in keeping with earlier data from patients with emphysema and COPD,

although these diseases were of greater severity than in our study patients (13). We then

investigated whether P-ECs derived from lung tissue extracts also showed characteristic

features of accelerated senescence when studied in vitro. P-ECs from patients with COPD

exhibited early replicative senescence compared to those of controls, with a marked decrease

in the cumulative PDL and a higher percentage of β-gal-positive cells at an early cell passage.

In the overall population of patients with COPD and controls, the relationships linking PDL to

FEV1 and to the emphysema score support a close association between cell senescence and

the severity of COPD. However, it is unlikely that cell senescence is related only to severe

emphysema, given the mild degree of emphysema in our patients. A more likely possibility is

that P-EC senescence with reduced angiogenic P-EC properties may participate in the

pathogenesis of emphysema.

A major finding from our study is that P-ECs undergoing replicative senescence

released increased amounts of several cytokines and mediators and that this process was

amplified in P-ECs from patients with COPD. Among 26 mediators investigated in P-EC-

conditioned media from patients with COPD and controls at various cell passages, nine were

detected using a Luminex®

assay. Of note, most of these secreted proteins in MP-EC-

conditioned media from controls increased with the number of passages, indicating that their

expression was linked to the normal process of replicative senescence. Several of these factors

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including IL6, IL8, Hu-GRO, MCP-1, and sICAM-1 were found in larger amounts at an early

cell passage in P-EC media from patients with COPD compared to controls, and most of them

failed to increase further during repeated cell passages. Thus, the differences between patients

with COPD and controls observed at an early cell passage but not at senescence were chiefly

due to the larger proportion of senescent cells in patients with COPD. These results are

consistent with previous studies from our laboratory showing increased susceptibility to

senescence of PA-SMCs in COPD (17). Senescent PA-SMCs were also shown to release

inflammatory mediators, although in much smaller quantities than P-ECs. That P-ECs release

20 to 10 000 times more IL8, MCP-1, Hu-GRO, or s-ICAM1 than PA-SMCs strongly

supports a prominent role for P-ECs in the general inflammatory process in COPD. Moreover,

inflammation and cell senescence influence each other, and IL8 is considered a potent inducer

of cell senescence in vitro (30). The high IL8 levels secreted by P-ECs in the present study are

consistent not only with a potential paracrine effect, but also with an endocrine effect of IL8

that may propagate cell senescence to other organs in COPD.

To evaluate whether basal lung inflammation in patients with COPD was linked to

P-EC senescence and telomere dysfunction, we measured cytokine levels and telomere length

in lung tissue extracts from our patients with COPD and controls. The levels of IL6, IL8, Hu-

GRO, MCP-1, and sICAM-1 were higher in lungs from patients with COPD than from

controls and correlated with in vitro criteria for cell senescence. Moreover, telomere length

determined from lung tissue extracts was decreased in patients with COPD compared to

controls and correlated negatively with the lung levels of these mediators (except IL6). Such a

close relationship between lung inflammation, telomere dysfunction, and P-EC senescence

criteria strongly supports a major role for premature P-EC senescence in promoting and

perpetuating the inflammatory process in COPD.

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One consequence of the increased cytokine release by P-ECs is attraction of

inflammatory cells, among which monocytes lead to alveolar macrophage accumulation (31).

In our study, P-EC-conditioned media from patients with COPD stimulated monocyte

migration to a greater extent than those from controls, and this difference was abolished by

MCP-1 antibodies. Senescent P-ECs also expressed greater amounts of the cell surface

adhesion molecule ICAM-1, which contributed substantially to monocyte adherence. These

findings constitute further evidence that cell senescence is among the COPD-associated

pulmonary vascular cell alterations that contribute to lung inflammation.

To determine whether telomere shortening responsible for increased susceptibility to

cell senescence was sufficient to cause lung inflammation, we investigated telomerase-

deficient mice, namely, Tert-/-

and Terc-/-

mice, characterized by various degrees of telomere

shortening. Interestingly, lung levels of the mouse homolog cytokines MCP-1, MIP-2,

CXCL1, and ICAM-1 were increased in the lungs from Terc-/-

mice, which had shorter lung

telomeres compared to wild-type controls and Tert-/-

mice. Moreover, MCP-1 lung levels,

which were higher in Tert-/-

and Terc-/-

mice than in wild-type controls, were increased in

proportion to the extent of telomere shortening. Taken together, these results strongly support

a causal role for cell senescence per se in inflammation, even in the absence of external

stimuli. While telomere shortening seems sufficient to increase the amounts of IL8, ICAM-1,

MCP-1, and Hu-GRO released by aging cells, combined mechanisms are probably needed to

elevate lung IL6 or other cytokines not detected in senescent P-EC-conditioned media.

The mechanisms underlying premature P-EC senescence in COPD can only be

speculated from the present study. We found that P-ECs from patients with COPD had

decreased telomerase activity and reduced telomere length, together with increased p21 and

p16 expression. These observations are consistent with a prominent role for increased cell

turnover in the occurrence of replicative cell senescence in patients with COPD. In

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accordance with this possibility, we found an inverse relationship between the PDL and

telomere length at an early passage. However, p16 expression was also higher in cells from

patients with COPD than in those from controls, suggesting a contribution of p16 in driving

premature senescence in COPD. Thus, accelerated P-EC senescence in COPD may be

attributable to a combination of both telomere shortening and oxidative stress responsible for

p16 activation.

In conclusion, our results strongly support a major contribution of telomere

dysfunction to lung inflammation and probably systemic inflammation in patients with

COPD. Slowing the cell senescence process may hold therapeutic promise for patients with

COPD.

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Acknowledgments

The authors gratefully acknowledge Manuel Serrano (Madrid, Spain) for providing the

Terc-/-

mice; Aurelie Guguin and Adeline Henry at the cytometry platform; Matthieu

Surenaud at the Luminex®

platform; Corinne Duprez et Catherine Dehoulle for Rt-qPCR

experiments; and Medhi Latiri for contributing to P-EC isolation and culture. We are indebted

to the surgeons from the chest surgery department of the Institut Mutualiste Montsouris for

providing the lung tissue samples.

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20. Amsellem V, Chaar V, Gary-Bobo G, Noureddine H, Sapin E, Maitre B, Validire P,

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Legends to figures

Figure 1: Immunolocalization and quantification of p16- and p21-stained cells in sections of

pulmonary vessels from patients with chronic obstructive pulmonary disease (COPD) and

controls. Representative photomicrographs p16 (A) and p21 (B) immunoreactivities (brown)

were located in endothelial cells identified by von Willebrand factor (vWF) staining (red).

Arrowheads show p16-positive cells (A) and p21-positive cells (B). Low magnification

bar=100 µm and high magnification bar=25 µm. The negative controls show staining with

appropriate control antibodies at the same concentration as p21 or p16 antibodies. Bar graphs

represent the percentage of endothelial cells (vWF-positive cells) expressing p16 (A) or p21

(B) in pulmonary vessels from patients with COPD and controls. Values are means±SEM.

*P<0.0001 compared with values from controls.

Figure 2: (A): Replicative senescence of vascular pulmonary endothelial cells (P-ECs) from

patients with chronic obstructive pulmonary disease (COPD) and controls. Cells were

subjected to repeated passages (P) and counted at each passage, and the population doubling

level (PDL) was calculated for patients with COPD and controls. (B): Data are means±SEM.

*P<0.01, versus controls. (C): Percentage of β-Gal-positive cells. P-ECs were stained for

senescence-associated β-Gal activity at passage 6 and at senescence when cells exhibit

proliferative arrest. Data are means±SEM. **P<0.0001 versus controls; †P<0.0001, versus

corresponding values at passage 2. (D): Representative photographs of cells stained for

senescence-associated β-Gal activity at passage 6 and at senescence.

Figure 3: Telomerase activity; telomere length; and p16, p53, and p21 mRNA levels in

vascular pulmonary endothelial cells (P-ECs) from patients with chronic obstructive

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pulmonary disease (COPD) and controls determined at passage 4 and at senescence. The T/S

ratio is the ratio of telomere repeat copy number over single-gene copy number (36B4 gene).

SF3A1 is the housekeeping gene used for p16, p53, and p21 mRNA quantification.

Each bar shows the mean±SEM. *P<0.05 and **P<0.01 vs. controls; †P<0.05 and ††P<0.01

vs. corresponding value at passage 4.

Figure 4: Levels of cytokines in the media of vascular pulmonary endothelial cells (P-ECs)

from patients with chronic obstructive pulmonary disease (COPD) and controls collected at

passage 6 and at senescence. Values are means±SEM. *P<0.05 and ***P<0.0001 vs. control

values; †P< 0.05 and ††P<0.01 vs. corresponding value at passage 6.

Figure 5: Effect of vascular pulmonary endothelial cell (P-EC)-conditioned media on

monocyte chemoattraction.

(A): Monocyte counts per field (0.6 mm2) attracted by MP-EC-conditioned media from

patients with chronic obstructive pulmonary disease (COPD) and controls at passage 6 and at

senescence in the presence of an MCP-1 neutralizing antibody (MCP-1 Ab, 2 µg/mL) or

nonspecific antibodies (IgG). Values are means±SEM. *P<0.01 vs. controls; †P<0.001 versus

MCP-1 neutralizing antibody; ‡ P≤0.001 vs. corresponding value at an early stage. Values

reflect three independent experiments.

(B): Representative photographs of filters after monocyte migration with MP-EC-conditioned

media from patients with COPD and controls at passage 6. Bar=50 µm

(C): Monocyte counts per field (0.6 mm2) in response to MCP-1 (50 ng/mL) alone and in

response to MCP-1 with a neutralizing antibody (MCP-1 Ab, 2 µg/mL) or with nonspecific

antibodies (IgG). Values are means±SEM; *P<0.001 vs. basal medium; †P<0.001 vs.

treatment with MCP-1. Values reflect three independent experiments.

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Figure 6: Adhesion of U937 monocytes to P-ECs from patients with chronic obstructive

pulmonary disease (COPD) and controls at passage 6

(A): U937 monocyte counts per field (0.6 mm2) adhering to P-ECs from controls or patients

with COPD at passage 6 and at senescence, in the presence of an anti-ICAM-1 antibody

(ICAM-1 Ab, 12 µg/mL) or nonspecific antibodies (IgG). Values are means±SEM. *P<0.05

vs. controls; †P<0.05 versus ICAM-1 neutralizing antibody; ‡ P=0.001 vs. corresponding

value at an early stage. Values reflect three independent experiments.

(B): Representative photograph of U937 monocytes adhering to the surface of P-ECs from

patients with COPD or controls at passage 6. White arrows show U937 adhering to the

endothelium surface. Bar=50 µm

Figure 7: Telomere length and cytokine levels in lung tissue extracts from patients with

COPD and controls. (A): Each bar is the mean±SEM. *P<0.05 compared with values from

controls. T/S is the ratio of the telomere repeat copy number over the single-gene copy

number (36B4 gene) (B): Correlation between telomere length and levels of IL8 (r=-0.45,

P<0.05), MCP-1 (r=-0.42, P<0.05), Hu-GROα (r=-0.44, P<0.05), and ICAM-1 (r=-0.38,

P<0.05).

Figure 8: Telomere length and cytokine levels in lungs from Tert-/-

(n=9), Terc-/-

(n=11), and

wild-type (WT; n=6) mice. (A) and (B): Each bar is the mean±SEM. *P<0.05 compared with

values from WT mice; †P≤0.001 compared with values from Tert-/-

mice. T/S is the ratio of

the telomere repeat copy number over the single-gene copy number (36B4 gene). (C):

correlation between telomere length and MCP-1 (r= -0.62, P<0.001).

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Table 1: Comparison of clinical features and pathological variables between patients with

chronic obstructive pulmonary disease and control smokers

COPD patients

(n=16)

Controls

(n=15)

P value

Females/Males 7/9 7/8 0.88

Age, years 62.8±2.1 61.3±2.6 0.65

Pack-years 45.4±5.1 31.2±5.7 0.10

Current/Former smokers 7/9 6/9 0.83

BMI kg/m2

24.3±1.1 22.5±1.3 0.33

FEV1% 72.8±3.6 92.7±3.3 <0.01

FEV1, L 2.0±0.2 2.5±0.2 0.03

FVC% 92.0±4.6 94.2±4.6 0.94

FVC, L 3.3±0.3 3.1±0.3 0.71

FEV1/FVC (%) 62.1±2.3 81.7±2.5 <0.0001

Emphysema score 17.1±1.9 4.5±1.2 <0.0001

Abbreviations: BMI, body mass index; FEV1, forced expiratory volume in 1 second; FEV1%,

percentage of the predicted FEV1 value; FVC, forced vital capacity; FVC%, percentage of the

predicted FVC value. Former smokers were defined as individuals who had not smoked

during the last year. The emphysema score was assessed on a 0-to-40 scale (32). Lung

function test values were those recorded after bronchodilators. Patients with COPD and

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controls were compared using the unpaired t-test or the Chi-square test for quantitative and

qualitative variables, respectively.

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Telomere Dysfunction Causes Sustained Inflammation in Chronic

Obstructive Pulmonary Disease

Amsellem V, Gary-Bobo G, Marcos E, Maitre B, Chaar V, Validire P, Stern JB,

Noureddine H, Sapin E, Rideau D, Hue S, Le Gouvello S,

Dubois-Rande JL, Boczkowski J, Adnot S

Online Data Supplement

Material and Methods

Study population

Thirty-one patients undergoing lung resection surgery for localized lung tumors were

recruited at the Institut Mutualiste Montsouris (Paris, France). All patients underwent

lobectomy for localized lung tumors. Among them, 16 had COPD and 15 were control

smokers matched to patients with COPD on age and sex (Table 1). Among the 16 patients

with COPD, 15 had non-small-cell lung cancer (NSCLC), including 3 with squamous cell

carcinoma and 12 with adenocarcinoma; and one had a lung localization of a breast

carcinoma. Among the 15 controls, 13 had NSCLC including 3 with squamous cell carcinoma

and 10 with adenocarcinoma; one had a lung localization of a breast adenocarcinoma and one

a lung localization of a sarcoma. The histological subtype distribution did not differ between

patients with COPD and controls.

Inclusion criteria for the patients with COPD were an at least 10-pack-year smoking

history and a ratio of expiratory volume in 1 second (FEV1) over forced vital capacity (FVC)

<70% after bronchodilator administration. All patients with COPD had mild airflow

limitation, and most of them did not have a diagnosis of COPD before lung surgery. Only 4 of

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the 16 patients with COPD were treated with inhaled beta2 agonists. None of them were

taking inhaled corticosteroids. Of these 4 patients, 3 met GOLD criteria for chronic

bronchitis. None controls received bronchodilatator therapy. Patients with a history of heart

disease, EKG abnormalities, or systolic dysfunction by echocardiography were not included

in the study; and none of the patients had hepatic disease, renal disease, or a history of cancer

chemotherapy.

The degree of emphysema was quantified using a 0-40 scale, as described by Bankier A

et al. (E1): the emphysema was rated from 0 to 4 on five computed tomography sections of

each lung, as previously reported (E2).

The study was approved by the institutional review board of the Henri Mondor

Teaching Hospital. All patients and controls signed an informed consent document before

study inclusion.

Lung tissue samples from peripheral lung at a distance from the tumor area were used

for pulmonary vascular endothelial cells (P-ECs) isolation, in situ immunohistochemical

studies, protein level determinations, and telomere length measurements.

Immunohistochemistry

Immunohistochemical analyses were performed on lung tissue samples collected from

the peripheral lung at a distance from the tumor area. Paraffin-embedded sections

deparaffinized using xylene and a graded series of ethanol dilutions were incubated in citrate

buffer (0.01 M, pH 6) at 90°C for 20 minutes. Endogenous peroxidase activity was blocked

with 3% H2O2 and 10% methanol in phosphate-buffered saline (PBS) for 10 min. Slides were

saturated for 60 minutes in 1% bovine serum albumin (BSA) and 5% goat serum in PBS 1X.

Then, immunostaining was performed in two steps. First, sections were incubated overnight

with anti-p21 mouse antibody (1:50, Cell Signaling, Boston, MA, USA), anti-p16 mouse

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antibody (1:1000, Abcam, Paris, France), or anti-CD68 mouse antibody (1/1000, Dako,

Glostrup, Denmark) followed by an anti-mouse ABC Vectastain kit horseradish peroxidase

(HRP) conjugate (Vectorlabs, Burlingame, CA, USA) or an anti-mouse HRP-conjugated

antibody (1:400 Dako, Carpinteria, CA, USA). HRP was revealed by DAB (fastDAB, Sigma,

Lyon, France) staining. Second, sections were incubated for 60 minutes with rabbit antibody

to von Willebrand factor (1:1000, Abcam, Paris, France) followed by an anti-rabbit HRP-

conjugated antibody (Vectastain ABC kit, Vectorlabs, Burlingame, CA, USA). HRP was

revealed by AEC (BD Pharmingen, San Diego, CA, USA) or Histogreen (Abcys, Paris,

France). Sections were counterstained with hematoxylin. We used IgG1, IgG2a, and IgG2b

control antibodies (R&D Systems, Minneapolis, USA) at the appropriate concentrations with

the same protocol used for staining with anti-p16, anti-p21, or anti-CD68 antibodies.

Senescent P-ECs were quantified by examining 15 random fields and counting in each field

the percentage of p16- or p21-positive cells, using the vWF-positive cells as the denominator.

Isolation of pulmonary vascular endothelial cells (P-ECs)

Lung parenchyma was collected in Hank’s Balanced Salt Solution (HBSS) with Ca2+

and Mg2+ supplemented with 50 U/mL penicillin, 50 µg/mL streptomycin, and 2.5 µg/mL

fungizone. The tissue was sectioned and digested in HBSS without Ca2+ and Mg2+ and with

dispase 2 U/mL (Invitrogen, Cergy-Pontoise, France) for 1.5 h at 37°C. Cells were separated

mechanically by flushing several times during digestion. After digestion, the tissue

homogenate suspension was filtered through a 40-µm cell strainer. The cells were pelleted at

300 g for 8 min and resuspended in endothelial cell complete media (see endothelial cell

culture section) on 0.2% gelatin. The cells were expanded to about 2 million then stained for

CD31 using the CD31-PE antibody (BD Biosciences, le Pont-de-Claix, France) and sorted

using fluorescence-activated cell sorting (FACS, MoFlo, Beckman-Coulter, Brea, CA, USA).

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After cell amplification, CD31-positive cell purity (≥97%) was confirmed by CD31 staining

and FACS (Cyan, Beckman-Coulter).

Culture of pulmonary vascular endothelial cells (P-ECs)

P-ECs were grown on 0.2% gelatin (Sigma, Lyon, France) in MCDB131 media (Gibco,

Invitrogen, Carlsbad, CA, USA) supplemented with 10% fetal calf serum, 50 U/mL penicillin,

50 µg/mL streptomycin, 2 mM L-Glutamine, 2.5 µg/mL fungizone, 25 mM HEPES, 30

µg/mL endothelial cell growth supplement, and 10 U/mL heparin. Cells were at passage 0

when the tissue homogenate was seeded for cell growth. The cells were expanded, and

endothelial cells were CD31-selected at passage 2. At each passage, when the cells reached

80%-90% confluence they were counted with Trypan blue and reseeded at 6666 cells/cm²;

viability was always greater than >95%. P-ECs were expanded until the number of cells was

sufficient for the experiments, i.e., until passage 4 or 6, depending on the ability of the cells to

proliferate. Cells from each passage were dispatched for DNA, RNA, and labeled for SA- β-

galactosidase. The onset of cell replicative senescence was determined based on cessation of

cell division, acid β-galactosidase activity, and cell morphology criteria. The cells that

reached replicative senescence were classified as being at the senescence stage. Counting cells

at each passage allowed us to compute the population-doubling level (PDL), as follows

PDL=(log10Y-log10X)/log10(2), where X is the number of cells seeded and Y is the number

of cells harvested after cell growth. We verified the origin of the endothelial cells using CD31

staining and fluorescence-activated cell sorting (FACS) at the first and last passages (Figures

E1A and E1B). Triple staining with CD31, CD34, and LYVE-1 showed no LYVE-1 staining,

establishing that the endothelial cells were from blood vessels and not from lymphatic vessels

(Figure E1B) (E3, E4).

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Immunocytochemical analyses

Cells were placed on glass cover-slips coated with gelatin 0.2%, fixed with 4%

paraformaldehyde, permeabilized with 0.2% Triton-X100, and immunostained with the

mouse antihuman CD31 primary antibody 1:50 (Santa Cruz Biotechnology, Santa Cruz, USA)

or corresponding mouse control IgG (R and D Systems, Lille, France) and the corresponding

secondary antibody labeled with AlexaFluor488 (Molecular Probes). Nuclei were stained

using DAPI at 1 µg/mL (Sigma, Lyon, France). Cover-slips were examined using a Zeiss

Axioplan 2, and images were captured with Zeiss AxioCam MRc (Zeiss, Jena, Germany).

Flow cytometry

Cells were trypsinized and washed with HBSS with 1% fetal calf serum (FCS) and

incubated with the indicated conjugated antibodies for 30 min at 4°C. Then, the cells were

washed twice with PBS 1X, fixed with PBS 1X-PFA 1%, and analyzed using a CyAnTM

ADP

analyzer (Beckman-Coulter). The antibodies used were CD31-PE (Becton Dickinson, San

Jose, CA, USA), CD34-FITC (Immunotools, Friesoythe, Germany), and LYVE-1-APC (R

and D Systems, Lille, France). All corresponding isotype controls came from Immunotools

company.

Senescence-associated β-galactosidase staining

At each passage, cells at 60% confluence were fixed with 2% formaldehyde and 0.2%

glutaraldehyde for 15 min. Then, the cells were washed with PBS and stained in a titrated

pH 6 solution containing 40 mM citric acid, 150 mM NaCl, 2 mM MgCl2, 5 mM potassium

ferrocyanide, and 1 mg/mL X-Gal.

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Assays of soluble factors using the Luminex®

system

P-ECs were grown in complete endothelial cell growth media. At the exponential stage,

cells were rinsed twice in PBS 1X and placed in reduced media containing 0.2% FCS without

heparin or endothelial cell growth supplement. After 24 h of incubation, the conditioned

media were collected and used for quantitation of 26 analytes using two milliplex plates

(Millipore, Molscheim, France) in parallel. The first milliplex plate was used to concomitantly

measure, on a single sample of conditioned media, the concentrations of the following 21

analytes: IL6, IL8, IL1ra, IL1α, IL1β, IL10, IL13, MIP1α, MIP1β, TNFα, INFγ, Fractalkine,

RANTES, Hu-GRO, MCP-1, EGF, FGF2, PDGF, VEGF, sCD40L, and CCL22. The second

milliplex plate was used for the simultaneous measurement of the concentrations of five

analytes: MMP9, PAI-1, sEselectin, sICAM-1, and sVCAM-1. Detection was performed

according to the manufacturer’s instructions. Briefly, cytokine and chemokine concentrations

were determined using antibodies for each analyte covalently immobilized to a set of

microspheres, according to a protocol developed and validated at Millipore. The analytes on

the surface of the microspheres were then detected using a cocktail of biotinylated antibodies.

Following binding of a streptavidin-phycoerythrin conjugate, the reporter fluorescent signal

was measured using the Luminex reader Bioplex 200 (Bio-Rad, Hercules, CA, USA). Data

were calculated using a calibration curve obtained in each experiment based on the respective

recombinant proteins diluted in the media used to obtain the conditioned media. Cytokine

concentrations were calculated using Bioplex Manager 6.0 (Bio-Rad) and normalized for the

number of cells used to generate the conditioned media. Levels were below the detection

threshold for IL1ra, IL1α, IL1β, IL10, IL13, MIP1 α, MIP1β, TNFα, INFγ, Fractalkine, EGF,

VEGF, sCD40L, CCL22, MMP9, sEselectin, and sVCAM-1, which are therefore not reported

in the results section. IL6, IL8, Hu-GRO, MCP-1, PAI-1, RANTES, sICAM-1, PDGF, and

FGF-2 were detectable, as reported in the results section, as graphs.

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Real-time quantitative PCR (RT-qPCR)

Total mRNA was extracted from P-ECs using RNeasy Protect Mini Kit (Qiagen, ZA

Courtaboeuf, France). First-strand cDNA was synthesized in reversed transcribed samples, as

follows: 500 ng total RNA isolated from cells, 8 U/µL M-MLV reverse transcriptase

(Invitrogen, Life Technologies, Cergy-Pontoise, France), 0,5 µg Oligo-(dT) 15 (Promega,

Charbonnières, France), and 0.8 mM mixed dNTP (GE Healthcare, Saclay, France).

Quantitative PCR was performed in a 7900HT Real-Time PCR system (Applied Biosystems,

ZA Courtaboeuf, France), using Fast SYBR Master Mix from Applied Biosystems. Table S1

reports the primer sequences designed to obtain equivalent optimal amplification efficiency

between the different assays with Fast PCR conditions. The amounts of mRNAs were

normalized for the amount of the control housekeeping gene SF3A1 (E5), which was chosen

among 3 tested housekeeping genes because of its stable expression in all samples (data not

shown). Endothelial cell mRNA expression was quantified using the ∆∆CT method. All PCR

experiments were done in triplicate in 384-well plates using a liquid handling workstation

(HAMILTON Robotics S.A.R.L., Massy, France). The sequences of primers for p16 were

Fwd 5'-GGGTCGGGTAGAGGAGGTG-3’ and Rev 5’- CATCATGACCTGGATCGGC-3’;

p21 Fwd : 5’-GAGACTCTCAGGGTCGAAAACG-3’ and Rev 5'-

GGATTAGGGCTTCCTCTTGGA-3’; p53 Fwd 5’-CCTGAGGTTGGCTCTGACTGTA-3’

and Rev 5’-TGTTCCGTCCCAGTAGATTACCA-3’ ; SF3A1 Fwd 5’-

TGCAGGATAAGACGGAATGGAAACTGA-3’ and Rev 5’-

GTAGTAAGCCAGTGAGTTGGAATCTTTG-3’.

Protein extraction and immunoblotting

Total proteins were extracted using RIPA lysis buffer (10 mM sodium phosphate pH 8,

150 mM NaCl, 1% sodium deoxycholate, 1% NP40, 0.5% SDS, 1 mM PMSF, 10 mM NaF,

1 mM sodium orthovanadate, and protease inhibitor cocktail (Roche, Basel, Switzerland).

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Immunoblots were carried out using the indicated antibodies and detected using an enhanced

chemiluminescence detection system (GE Healthcare, Munich, Germany). Densitometric

quantification was normalized for the β-actin level using Gene Tools software (Ozyme,

Montigny le Bretonneux, France). The antibodies used were anti-P-p53 Ser15 (1:1000, Cell

Signaling Technology, Boston, MA, USA) and anti-βactin (1:5000, Sigma, Saint Quentin-

Fallavier, France).

Assessment of monocyte migration

Monocytes were extracted from buffy coat of healthy blood donors provided by the

French Blood Transfusion Organization (EFS, Creteil, France). Peripheral blood mononuclear

cells (PBMCs) were isolated by Ficoll extraction. The monocytes in the PBMC population

were selected by adherence for 1 hour on a plastic culture plate in RPMI medium (Gibco)

supplemented with 10% human serum (AbCys, Paris, France), penicillin, streptomycin,

HEPES 10 mM, sodium pyruvate (1%), MEM 1%, AANE 1%, and β-mercaptoethanol

50 µM. Monocytes were collected with a cell scraper and left overnight at 4°C suspended in

RPMI complete medium.

Conditioned media tested for monocyte chemoattraction were obtained from P-ECs left

in MCDB131 medium with 0.5% BSA for 24 h. Prior to the migration assay, the monocytes

were rinsed in MCDB131 with 0.5% BSA and incubated with FcR blocking reagent for 1 h at

4°C. The frozen MCDB131 0.5% BSA-conditioned media were incubated with MCP-1

neutralizing antibody or control IgG (R and D Systems, Lille, France) at 2 µg/mL for 30 min

on a rotating wheel.

For the monocyte transwell migration assay, 5-µm pore 24-well plates (Corning,

Amsterdam, The Netherlands) were hydrated with MCDB131 0.5% BSA and placed on the

various conditioned media preincubated with MCP-1 neutralizing antibody or control IgG.

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Monocytes in MCDB131 BSA 0.5% (300 000 per well) were added on the filters and allowed

to migrate for 4 h at 37°C in 5% CO2. Recombinant human MCP-1 (R and D Systems, Lille,

France) at 50 ng/mL was used as the positive control. Monocytes on the filters were stained

with the Diff-Quick kit (Siemens Diagnostics, Saint Denis, France). Unmigrated monocytes

(top of the filter) were removed using a cotton swab, and the filters were cut out from the

transwells, mounted onto slides, and photographed under the microscope with a 20X

objective. The monocytes were then counted.

Adhesion of U937 monocytic cells to pulmonary vascular endothelial cells (P-ECs)

For adhesion assay, we used a blood-borne monocytes cell line (U937) (E6). Confluent

P-ECs in 24-well plates were starved for 4 h in 0.5% BSA media and incubated with an anti-

ICAM-1 antibody or control IgG (12.5 µg/mL, R and D Systems, Lille, France) for 1 h.

Human monocytic U937 cells grown in RPMI and preincubated with FcR blocking reagent

for 1 h (Mylteni, Paris, France) were allowed to adhere onto the endothelial cells pretreated

with ICAM-1 or IgG, for 2 h. Then, nonadhering monocytes were removed by two washes

with HBSS with calcium and magnesium. Monocytes adhering to the endothelial cells were

stained using the Diff-Quik kit (Siemens Diagnostics, Saint Denis, France). Photographs were

taken under the microscope with a 20X objective, and monocytes in each field were counted.

Telomere length analysis

Total DNA was extracted P-ECs and lung tissue using QIAamp DNA Mini kit (Qiagen,

Courtaboeuf, France) according to the manufacturer’s instructions. P-ECs were harvested,

pelleted, washed with PBS 1X, resuspended in 200 µL of PBS 1X, and homogenized in a

Tissue Lyser® homogenizer (Qiagen) before lysis in the presence of proteinase K. Lung

tissue, 20 mg, stored at -80°C was defrosted and homogenized with 80 µL of PBS 1X in a

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Tissue Lyser® homogenizer (Qiagen, Courtaboeuf, France) before lysis in the presence of

proteinase K.

The purity of the extracted DNA was checked by performing electrophoresis on 1%

agarose gel. The DNA was quantified using a spectrophotometer and the telomere length of

the extracted DNA using an RT-qPCR assay as previously described (E7).

ELISA on homogenized lung tissue

Lung tissue, 30 mg, stored at -80°C was defrosted and homogenized in a Tissue Lyser®

homogenizer (Qiagen, Courtaboeuf, France) using 300 µL of T-PER® tissue protein

extraction reagent (Thermo Scientific Pierce, Illkirch, France). The homogenate was

centrifuged at 10 000 g for 5 min to remove tissue debris, and the supernatant containing the

total lung lysate was used for the ELISA. The total protein level in the lung lysate was

quantified using the Biorad DC protein assay (Biorad, Marnes-la-Coquette, France). Before

performing each ELISA, various dilutions of protein lysates were tested to ensure that the

detected level fit within the standard linear range. All ELISA kits were from R and D Systems

(Lille, France)

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References

E1. Bankier AA, De Maertelaer V, Keyzer C, Gevenois PA. Pulmonary emphysema:

Subjective visual grading versus objective quantification with macroscopic morphometry and

thin-section ct densitometry. Radiology 1999;211:851-858.

E2. Chaouat A, Savale L, Chouaid C, Tu L, Sztrymf B, Canuet M, Maitre B, Housset B,

Brandt C, Le Corvoisier P, Weitzenblum E, Eddahibi S, Adnot S. Role for interleukin-6 in

copd-related pulmonary hypertension. Chest 2009;136:678-687.

E3. Muller AM, Hermanns MI, Skrzynski C, Nesslinger M, Muller KM, Kirkpatrick CJ.

Expression of the endothelial markers pecam-1, vwf, and cd34 in vivo and in vitro. Exp Mol

Pathol 2002;72:221-229.

E4. Podgrabinska S, Braun P, Velasco P, Kloos B, Pepper MS, Skobe M. Molecular

characterization of lymphatic endothelial cells. Proc Natl Acad Sci U S A 2002;99:16069-

16074.

E5. Mesel-Lemoine M, Cherai M, Le Gouvello S, Guillot M, Leclercq V, Klatzmann D,

Thomas-Vaslin V, Lemoine FM. Initial depletion of regulatory t cells: The missing solution to

preserve the immune functions of t lymphocytes designed for cell therapy. Blood

2006;107:381-388.

E6. DiCorleto PE, de la Motte CA. Characterization of the adhesion of the human

monocytic cell line u937 to cultured endothelial cells. J Clin Invest 1985;75:1153-1161.

E7. Savale L, Chaouat A, Bastuji-Garin S, Marcos E, Boyer L, Maitre B, Sarni M, Housset

B, Weitzenblum E, Matrat M, Le Corvoisier P, Rideau D, Boczkowski J, Dubois-Rande JL,

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Figure legends

Figure E1: Characterization of vascular pulmonary endothelial cells. (A) Localization of

CD31 by immunofluorescence. Representative picture of P-ECs stained for CD31 using a

CD31 antibody (green on merged panel). Nuclei were stained using DAPI (blue on merged

panel). (B) Representative FACS profile of P-ECs stained for CD31, CD34, and LYVE-1

using CD31-PE, CD34-FITC, and LYVE-1-APC antibodies. Left panel: FACS profile

showing the percentage of endothelial cells (CD31-positive cells). Right panel: FACS profile

showing the percentage of vascular endothelial cells (LYVE-1-/CD34+ and LYVE-1-/CD34-)

and the absence of lymphatic LYVE-1+/CD34 endothelial cells.

Figure E2: Analysis of Phospho-p53(Ser15) protein level. The amounts of

Phospho-p53(Ser15) normalized for β-actin level are shown as a bar graph. Each bar is the

mean±SEM. *<0.05 compared with MP-EC values from controls at the same stage. †<0.05

values at senescence compared with corresponding values at passage 4.

Figure E3: Immunolocalization and quantification of CD68-stained cells in sections of

pulmonary vessels from patients with chronic obstructive pulmonary disease (COPD) and

controls. Representative photomicrographs showing CD68-stained cells and vWF-stained

vessels. Arrows show CD68-positive cells, bar=100 µm. The negative control panel shows

staining using the isotype-specific control antibody at the same concentration as that of the

CD68 antibody. Bar graphs represent the number of positive cells per field in patients with

COPD and controls. Values are means±SEM. * <0.0001 compared with values from controls.

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