Am J Physiol Lung Cell Mol Physiol 306: L775–L785, 2014. First published February 14, 2014; doi:10.1152/ajplung.00200.2013.

Lung endothelial barrier disruption in Lyl1-deficient mice Nelly Pirot,1 Hélène Delpech,1 Virginie Deleuze,1 Christiane Dohet,1 Monique Courtade-Saïdi,2 Céline Basset-Léobon,2 Elias Chalhoub,1 Danièle Mathieu,1 and Valérie Pinet1 1

Institut de Génétique Moléculaire de Montpellier, UMR 5535 CNRS, Montpellier University, Montpellier, France; Monique Courtade-Saïdi, Céline Basset-Léobon: Laboratoire d’Histologie-Embryologie, Faculté de Médecine Rangueil, Toulouse, France 2

Submitted 24 July 2013; accepted in final form 8 February 2014

Pirot N, Delpech H, Deleuze V, Dohet C, Courtade-Saïdi M, Basset-Léobon C, Chalhoub E, Mathieu D, Pinet V. Lung endothelial barrier disruption in Lyl1-deficient mice. Am J Physiol Lung Cell Mol Physiol 306: L775–L785, 2014. First published February 14, 2014; doi:10.1152/ajplung.00200.2013.—Maturation of newly formed vessels is a multistep phenomenon during which functional endothelial barriers are established. Disruption of vessel integrity is an important feature in many physiological and pathological processes. We previously reported that lymphoblastic leukemia-derived sequence 1 (LYL1) is required for the late stages of postnatal angiogenesis to limit the formation of new blood vessels, notably by regulating the activity of the small GTPase Rap1. In this study, we show that LYL1 is also required during the formation of the mature endothelial barrier in the lungs of adult mice. Specifically, LYL1 knockdown in human endothelial cells downregulated the expression of ARHGAP21 and ARHGAP24, which encode two Rho GTPaseactivating proteins, and this was correlated with increased RhoA activity and reorganization of the actin cytoskeleton into stress fibers. Importantly, in lungs of Lyl1-deficient mice, both vascular endothelial (VE)-cadherin and p120-catenin were poorly recruited to endothelial adherens junctions, indicative of defective cell-cell junctions. Consistent with this, higher Evans blue dye extravasation, edema, and leukocyte infiltration in the lung parenchyma of Lyl1⫺/⫺ mice than in wild-type littermates confirmed that lung vascular permeability is constitutively elevated in Lyl1⫺/⫺ adult mice. Our data show that LYL1 acts as a stabilizing signal for adherens junction formation by operating upstream of VE-cadherin and of the two GTPases Rap1 and RhoA. As increased vascular permeability is a key feature and a major mechanism of acute respiratory distress syndrome, molecules that regulate LYL1 activity could represent additional tools to modify the endothelial barrier permeability. lymphoblastic leukemia-derived sequence 1; bHLH; vascular permeability; RhoA; vascular endothelial-cadherin

that lines blood vessels forms a selective and semipermeable barrier to control the exchange of solutes, proteins, fluids, and immune cells between blood and tissues. Fine regulation of endothelial permeability is required in many physiological and pathological processes, including developmental and tumor angiogenesis, as well as immunity and inflammation (14, 25). A complex balance between cell-cell and extracellular-matrix tethering (ECM) forces and intracellular contractile forces mediated by actin and myosin maintains the integrity of the vascular barriers. Interendothelial junctions include adherens, tight, and gap junctional complexes (28). At adherens junctions (AJs), intercellular adhesion is mediated by vascular endothelial (VE)-

THE MATURE CONTINUOUS VASCULAR ENDOTHELIUM

Address for reprint requests and other correspondence: V. Pinet, Institut de Génétique Moléculaire de Montpellier, UMR 5535, CNRS, 1919 route de Mende, 34293 Montpellier cedex 5, France (e-mail:[email protected]). http://www.ajplung.org

cadherin that interacts with cytoskeletal and signaling proteins to promote junction anchoring to actin microfilaments and to transfer signals inside the cell (21). This association is required for AJ stabilization but also for the dynamic regulation of AJ opening and closing. Cell adhesion and cytoskeleton dynamics are tightly controlled by multiple members of the small guanosine triphosphatase (GTPase) superfamily (reviewed in Ref. 38). Most of the mechanisms involved in the maintenance of vascular integrity in mature vessels also play a role in stabilizing newly formed vessels during the late stages of angiogenesis. Accordingly, the transcriptional regulators acting upstream of these pathways control both the integrity of mature vessels and the stabilization of newly formed vessels (23, 27, 31, 34). The transcription factor lymphoblastic leukemia-derived sequence 1 (LYL1) is a member of the basic helix loop helix family and is closely related to TAL1 (also called SCL). LYL1 is specifically expressed in hematopoietic (19) and endothelial cells (ECs) (32) during development and adulthood. LYL1 is not essential for developmental processes because Lyl1⫺/⫺ mice show only a mild hematopoietic phenotype that affects mainly the lymphoid lineage and the long-term hematopoietic reconstitution capacity (12, 52). We recently identified angiopoietin-2, a major regulator of angiogenesis and lymphangiogenesis (5), as the first direct LYL1 target gene in human ECs (15). In adult mice, Lyl1 is expressed in both quiescent and angiogenic vessels and is required for the late stages of postnatal angiogenesis to limit the growth of newly formed vascular structures (32). Indeed, syngeneic tumors implanted in Lyl1⫺/⫺ mice develop vessels with enlarged lumens, reduced pericyte coverage, and increased permeability. We previously reported that LYL1 silencing in human ECs downregulate the expression of several molecules that contribute to endothelial junction maturation (32), namely integrin␣2, which has an important role in EC-ECM attachment (6), as well as RapGEF1 and RapGEF2, which have multiple roles through Rap1 activation in AJ maturation (30). In this study, we show that LYL1 also regulates the activity of RhoA in ECs and that LYL1 downregulation leads to impaired AJs. Moreover, the lung vascular barrier is defective in Lyl1⫺/⫺ mice. MATERIALS AND METHODS

Cell cultures. Primary ECs from human umbilical vein (HUVECs; PromoCell) and the immortalized human EC line human telomerase reverse transcriptase (hTERT-1) (51) were obtained from the United States Center for Disease Control and were cultured in complete endothelial growth medium-2 (PromoCell) supplemented with 3% fetal calf serum (FCS) (HyClone, Perbio Science). Small-interfering RNA (siRNA) transfection in HUVECs and small-hairpin RNA

1040-0605/14 Copyright © 2014 the American Physiological Society

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(shRNA) transduction in hTERT-1 cells were performed as previously described (32). The sequences of duplex RNAs were as follows: LYL1A-Forward: GAUGGAGCAAACCGCUUUG-TT; LYL1A-Reverse: CAAAGCGGUUUGCUCCAUC-TT; LYL1D-Forward: GUCAAGGAAAGGGCAGUGG-TT; LYL1D-Reverse: CCACUGCCCUUUCCUUGAC-TT. Stealth RNAi siRNA Negative Control Medium GC duplex (Life Technologies) was used as control siRNA. Gene expression analysis by real-time PCR. Total RNA was extracted from cultured cells using the High Pure RNA Isolation Kit (Roche) following the manufacturer’s instructions. RNA was primed with oligo (dT)20 or a random primer and reverse transcribed with SuperScript II (Invitrogen). The sequences of the primers used for real-time PCR analysis are listed in Table. 1. Results are shown as the means ⫾ SD. Adenoviral vector construction and viral production. Recombinant adenoviruses were generated using the Adeno-XTM Tet-Off expression system (Clontech). The Ad-TRE-LYL1 construct contains a 0.9-kb EcoRI-blunted fragment derived from pSG-FlagLYL1, in which nucleotides 407 to 1,278 of the human LYL1 cDNA (17) were cloned by PCR in phase with the FLAG Tag of the pSG5-FLAG plasmid (kindly provided by Alain Sergeant, ENS, Lyon, France). Recombinant replication-defective adenoviruses were produced by transient transfection of the constructs in human embryonic kidney 293 cells as described (26). RhoA activation assay. Cells were grown on collagen-coated 60-mm culture dishes until confluence and rinsed and lysed according to the manufacturer’s instructions (Rhotekin-RBD Protein GST Beads, Cytoskeleton). After centrifugation, lysate supernatants were incubated with agarose beads conjugated with the rhotekin Rhobinding domain that recognizes only GTP-bound active RhoA. RhoA was detected in pulled-down products and total lysates by immunoblotting using an anti-RhoA antibody (Santa Cruz Biotechnology, sc-418). Animals. Lyl1⫺/⫺ mice have been previously described (12). Mice were housed in pathogen-free conditions in the institutional animal facility. All experiments were conducted by authorized personnel (agreement numbers: 34-308 for N. Pirot and 34-368 for V. Pinet) and approved by the Institutional Review Board at the Animal Facility of the Institut de Génétique Moléculaire de Montpellier. Animals were genotyped by PCR analysis of tail DNA as described elsewhere (32). Mice were anesthetized by isofluorane inhalation and killed at 2, 4, 6, 9, and 12 mo of age; lungs were collected and fixed in neutral buffered

formalin (4% formaldehyde) overnight, dehydrated, and embedded in paraffin. Quantification of Evans blue dye extravasation within tissues and edema. Protease-activated receptor-1 (PAR-1) activating peptide (SFLLRN) or control peptide (scrambled PAR-1) (1 mg/kg; Innovagen) was injected trough the retroorbital sinus of wild-type (WT) or Lyl⫺/⫺ mice. At the same time, 20 mg/kg of Evans blue dye (EBD, Sigma) was injected in the tail vein. Thirty minutes after injection, animals were anesthetized by intraperitoneal injection of 2 mg/kg body wt xylazine (Rompun, 2% solution; Bayer Pharma) and 50 mg/kg body wt ketamine (Imalgène 500; Merial), and the chest was opened. Mice were transcardially perfused with 50 ml PBS/5 mM EDTA to remove excess circulating EBD. Organs were excised, dried, weighed, and placed in formamide at 60°C for 36 h. EBD in supernatants was quantified by spectrophotometry at 630 nm. For calculation of the lung wet-to-dry weight ratio, lungs were excised, weighed, and completely dried in an oven at 60°C overnight. Fetal liver transplantation experiments. Six-week-old C57Bl/6 (Ly5.2) Lyl1⫺/⫺ mice or WT littermates were used as recipients of fetal liver (FL) cells obtained from 14.5-day postcoitus (dpc) C57Bl/6 (Ly5.1) embryos by mashing the excised livers. Recipients were lethally irradiated with a unique 9.5-Gy dose before injection of 4 ⫻ 106 FL cells trough the retroorbital sinus. Hematopoietic reconstitution was assessed at 4 wk after FL cell grafting by peripheral blood analysis. To monitor engraftment, peripheral blood mononuclear cells were double-stained with anti-CD45.2 (Ly5.2) FITC- and antiCD45.1 (Ly5.1) phycoerythrin-conjugated antibodies and analyzed by flow cytometry. Light microscopy, immunohistochemistry, and immunofluorescence. Mouse lung sections (4 ␮m thick) were stained with hematoxylin and eosin (HE) for preliminary analysis and visualized with a NanoZoomer slide scanner controlled by the NDP.view software. For immunohistochemistry, sections were deparaffinized, rehydrated, and incubated for antigen retrieval in citrate buffer at 100°C for 30 min. Sections were incubated in 0.3% H2O2 solution at room temperature for 20 min to inhibit endogenous peroxidase and washed three times with PBS. Nonspecific antibody binding was blocked by incubation with PBS containing 20% normal horse serum for 30 min. Endogenous biotins were blocked using the Avidin-Biotin Blocking kit (Vector Laboratories, Clinisciences). Sections were then incubated with rabbit anti-Ki67 (Thermo Fisher Scientific), rat anti-MAC2 (0.5 ␮g/ml; eBioscience), goat anti-CD3⑀, or anti-PAX5 (0.4 ␮g/ml; Santa

Table 1. Sequences of the primers used in RT-qPCR Gene

Forward Primer

Reverse Primer

Sequences of the Homo sapiens primers GAPDH LYL1 ARHGAP21 ARHGAP24 ITGA2 RAP-GEF1 RAP-GEF2

ACACCCACTCCTCCACCTTT CATCTTCCCTAGCAGCCGGTTG AGCCTCGAGTACCAGTCCGC ATGGAGGGCACTGTGGTGGT CGTCTCTCAGTTTCCAAGCC ACTGATCGACAGCTCGTCCT CCCAGCTGTCAACCGATATT

F2r (PAR-1) CD31 Itga2 Lyl1 VE-cad Rap-GEF1 Rap-GEF2 ArhGAP21 ArhGAP24 18S

CCGCGTCCCTATGAGCCAGCCAGAATC CTGCCAGTCCGAAAATGGAAC ACCGCCCCTTCTGTATCTTT AGATGAGGAAACGCCCTGTA ACCGGATGACCAAGTACAGC CTGACCCAGTTCACAGAGCA CTGCACCTGAGAAAGCTGTG CACGCACATGCCTGACCAGT GCTGCCGGGCCAGGCGAACCT GTTCCGACCATAAACGATGCC

TCCACCACCCTGTTGCTGTA GTTGGTGAACACGCGCCG GGAGGGCCAACAGATTCGTGG ACTGCCCCATGGTGGCTTTC CAGTTGCCATGCTTACTGGA CCGTCGATGTAGTCTGGGTT CGATGGAGCAGAACTTGTCA

Sequences of the Mus musculus primers GGCGGCGTGTGAGGAGGGAGGC CTTCATCCACCGGGGCTATC GTGAAGGTCTTTCAGCCAGC AGCCACTGCAAGTAGCCTGT TTCTGGTTTTCTGGCAGCTT GATGAACTTGAGGAGCAGCC TAAGCGAGGAAATCCCACTG CGGATCCCCAGGAGCCCTTA CGTCGGTGTTGCTGTCGAAGGAGGG TGGTGGTGCCCTTCCGTCAAT

AJP-Lung Cell Mol Physiol • doi:10.1152/ajplung.00200.2013 • www.ajplung.org

LYL1 AND THE LUNG ENDOTHELIAL BARRIER

Cruz Biotechnology, Clinisciences) antibodies for 1 h. After being washed, lung sections were incubated with biotinylated secondary antibodies (1/500; Vector Laboratories, Clinisciences) for 45 min, washed with PBS containing 0.1% Tween, incubated with extravidin peroxidase, and revealed with 3-amino-9-ethylcarbazole (SigmaAldrich). Fixed cryostat sections were stained with fluorescein isothiocyanate-labeled tomato lectin (3 ␮g/ml, Vector Laboratories), rat anti-mouse CD31 antibody (2 ␮g/ml, BD Pharmingen), rabbit antimouse VE-cadherin antibody (1/800; kindly provided by Ph. Huber, Grenoble, France), and rabbit anti-mouse p120-catenin antibody (2 ␮g/ml; sc-13957, Santa Cruz Biotechnology, Clinisciences). Primary antibodies were detected using a chicken anti-rat IgG Alexa 488 or a donkey anti-rabbit IgG Alexa 555 anti-serum (Invitrogen), and nuclei were stained with DAPI. Perls’ Prussian blue staining. Lung sections were incubated in a 5% potassium ferrocyanide and 5% hydrochloric acid solution (1:1) for 30 min. Sections were then washed and counterstained with

A

HUVEC-siLYL1A

mRNA levels relative to controlsiRNA treated HUVECs

1 0,8

0,4

***

**

*

**

***

0,6

*** *

***

**

0,2 0

LYL1

B

HUVECs

ARH GAP21

ARH GAP24

ITGA2

GTP-bound RhoA/Total RhoA

200

100

300

200

***

100

0

sh- shCTL LYL1

Ad- AdLacZ LYL1

Active RhoA Total RhoA

DAP - F-actin

F-actin-LacZ

DAP - F-actin

DAP - F-actin

F-actin-Flag

Ad-LacZ

DAP - F-actin

*

*

Ad-LYL1

siLYL1A

siCTL

C

nuclear fast red. Hemosiderin was stained blue, and nuclei appeared red. Isolation of mouse lung ECs. Lungs were excised from 6- to 8-wk-old mice and transferred to a GentleMACS C tube containing the dissociation solution of the lung dissociation kit (Miltenyi Biotec). Tissues were dissociated with a GentleMACS dissociator (“m.lung” program) and incubated at 37°C with frequent shaking for 30 min, followed by a second dissociation with GentleMACS. Enzyme activity was stopped by addition of 2 mM EDTA and 10% FCS. The resulting suspensions were passed through a 70-␮M filter, and erythrocytes were lysed with the red blood cell lysing buffer (SigmaAldrich). Mouse lung ECs (mLECs) were then isolated from the bulk mixture by selection with anti-CD146-coated Miltenyi magnetic beads (clone: ME-9F1) and LS MACS columns. To estimate the macrophage, hematopoietic, endothelial, and smooth muscle cell content in the different fractions, cells were stained with anti-CD11bFITC, anti-CD45-PE, anti-CD31-APC, and anti-␣-smooth muscle actin (␣-SMA) antibodies before analysis by flow cytometry. For RhoA activation, mLECs were isolated, sorted by a two-step selection with anti-CD31 and anti-intercellular adhesion molecule 2 antibodies and cultured exactly as previously described (36). Western blot analysis. Whole cell extracts and immunoblotting were performed as described (15). Membranes were probed with the following antibodies: rabbit anti-integrin-␣2 (1/1,000; Millipore), goat anti-CD31 (0.4 ␮g/ml; sc-1506, Santa Cruz Biotechnology, Clinisciences), and rabbit anti-mouse VE-cadherin (1/500), followed by secondary antibodies conjugated to horseradish peroxidase (Amersham). Bands were revealed with Luminata Crescendo (Millipore). To control protein loading, membranes were reprobed with a mouse anti-␤-actin antibody (0.02 ␮g/ml; Sigma-Aldrich). Statistical analysis. Differences between experimental groups were assessed by using the unpaired Student’s t-test. Values are expressed as the means ⫾ SD. P ⬍ 0.05 was considered statistically significant.

400

300

si- siCTL LYL1

RAP GEF2

HUVECs

**

0

RAP GEF1

hTERT1 **

400 GTP-bound RhoA/Total RhoA

HUVEC -siLYL1D

1,2

L777

Fig. 1. Lymphoblastic leukemia-derived sequence 1 (LYL1) silencing reduces the expression of ARHGAP21 and ARHGAP24 genes, which encode 2 RhoGTPase-activating proteins (RhoGAPs), and increases RhoA activity and stress fiber formation. A: human umbilical vein endothelial cells (HUVECs) were transfected with anti-LYL1 (siLYL1A or siLYL1D) or control siRNAs, and total RNA was prepared 48 h after transfection. mRNA levels of the indicated genes were quantified by qRT-PCR (primers in Table 1) and normalized to GAPDH expression. Each bar is the mean ⫾ SD of the mRNA levels relative to those of control HUVECs (control siRNA) (at least 3 independent experiments), which were set at 1. *P ⬍ 0.05; **P ⬍ 0.01; ***P ⬍ 0.001 by using the Student’s t-test. B: evaluation of RhoA activity. Left: HUVECs were transfected with control siRNA (si-CTL) or LYL1AsiRNA. Human telomerase reverse transcriptase (hTERT1) cells were transduced with lentiviruses encoding either control shRNA (sh-CTL) or LYL1shRNA, and puromycin-resistant cell populations were established. Right: HUVECs were transduced with recombinant adenoviruses driving LacZ or LYL1 expression. Whole cell extracts from the different cultures were prepared and incubated with agarose beads conjugated with the rhotekin Rho-binding domain that recognizes only GTP-bound active RhoA. Because the protein concentration of the whole cell extracts was not determined, some variations in total RhoA might appear. The GTP-bound RhoA/total RhoA ratio for each extract was quantified using ImageJ. Bars show the means ⫾ SD of the GTP-bound RhoA/total RhoA ratio of at least 3 independent experiments. The ratio of control cells was arbitrarily set to 100. Images shown are representative of the 3 experiments. Vertical dashed lines have been inserted to indicate repositioned gel lanes. **P ⬍ 0.01; ***P ⬍ 0.001 by Student’s t-test. C: impact of LYL1 expression on F-actin stress fiber formation. HUVECs were transfected with control siRNA or si-LYL1A or transduced with adenoviruses expressing either LacZ or human LYL1 tagged with a flag epitope. Subconfluent HUVECs were immunostained with TRITC-Phalloidin (red) to visualize F-actin stress fibers. Ad-LacZ-transduced HUVECs were identified by staining with an anti-␤-galactosidase antibody (green), and Ad-LYL1 transduced HUVECs with an anti-flag antibody (green). LYL1-depleted cells showed stress fiber formation (arrows) that was strongly reduced in LYL1-overexpressing cells (asterisk) compared with nontransduced neighboring cells. Scale bar ⫽ 20 ␮m.

AJP-Lung Cell Mol Physiol • doi:10.1152/ajplung.00200.2013 • www.ajplung.org

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RESULTS

cifically pulled down using the GST-rhotekin-Rho binding domain. The basal levels of active RhoA were significantly higher (more than 2-fold) in LYL1-depleted HUVECs than in control (siCTL-transfected) cells (Fig. 1B, left). This result was confirmed in hTERT1 cells in which LYL1 was stably downregulated by transduction with lentiviruses encoding anti-LYL1 shRNAs (Fig. 1B, left). Reciprocally, LYL1 overexpression in HUVECs mediated by adenoviral transduction significantly reduced RhoA activity (around 2-fold) compared with cells transduced with control vector (Ad-LacZ) (Fig. 1B, right). F-actin cytoskeleton plays a major role in the maintenance of endothelial function by determining cell shape, facilitating cell adhesion to the subendothelial matrix, and participating in the regulation of junctional complexes. The dynamics and structure of F-actin filaments are mainly regulated by RhoGTPases, and RhoA is primarily involved in stress fiber formation and cell contraction. Therefore, the effect of LYL1 depletion or overexpression on F-actin structure was evaluated by immunofluorescence using TRITC-labeled phalloidin to stain F-actin (Fig. 1C). In LYL1-silenced HUVECs, stress fibers were increased (arrows in Fig. 1C) compared with siCTL-transfected cells. Specifically, 34.2% of siLYL1-transfected HUVECs showed actin stress fibers compared with 19.1% of siCTLtransfected cells. Conversely, cortical actin organization was observed only in 20.9% of LYL1-depleted HUVECs and in 33.8% of siCTL-transfected cells. Indeed, many siCTL-transfected cells were round shaped with cortical actin, whereas

B ** **

CTL PAR-1

250 200 150 100 50

CTL PAR-1 LUNG-4m

C

WT Lyl1-/1,2E-02

8,0E-03

4,0E-03

2m

250 200 150 100 50

LUNG-2m HEART-2m

D

5.50

**

5.25 5.00 4.75 4.50 4.25

0,0E+00

300

0

CTL PAR-1 HEART-4m

Wet/dry ratio in the lungs

0

350

Lyl1-/-

WT

Evans Blue extravasation ( g/g dry weight)

300

Lung PAR-1 mRNA levels normalized to -actin

Fig. 2. Basal lung vascular permeability is increased in Lyl1⫺/⫺ mice. A: control peptide ( and Œ) or protease-activated receptor-1 (PAR-1) agonist peptide (Œ and o) (1 mg/kg) was injected through the retroorbital sinus, and concomitantly Evans Blue dye was intravenously injected in 4-mo-old wildtype (WT) and Lyl1⫺/⫺ mice. After 30 min, dye extravasation was measured as described in MATERIALS AND METHODS in lung and heart. B: measurement of Evans blue dye extravasation in lung and heart of 2-mo-old WT and Lyl1⫺/⫺ mice. C: PAR-1 mRNA expression in the lungs of 2- and 4-mo-old WT and Lyl1⫺/⫺ mice. Each bar is the mean ⫾ SD of mRNA levels relative to ␤-actin (3 mice per genotype). D and E: lungs of 4- and 2-mo-old WT and Lyl1⫺/⫺ mice were excised to determine the wet/dry weight ratio as described in MATERIALS AND METHODS. **P ⬍ 0.01. Gray bars represent the mean of the corresponding set of points.

Evans Blue extravasation ( g/g dry weight)

A 350

E Wet/dry ratio in the lungs

LYL1 silencing in human ECs reduces ARHGAP21 and ARHGAP24 gene expression, increases RhoA activity, and induces the formation of F-actin stress fibers. A preliminary genome-wide array analysis of LYL1-silenced and control (negative control siRNA) HUVECs identified many genes that are potentially regulated by LYL1 (15). Among them, ARHGAP21 and ARHGAP24, which encode two Rho GTPaseactivating proteins (GAPs), were further investigated because of their involvement in the establishment of cell-cell contacts (37, 41). Using two different siRNAs against LYL1, we confirmed the downregulation of ARHGAP21 and ARHGAP24 mRNA in HUVECs by quantitative RT-PCR (48% and 30% reduction, respectively, compared with cells transfected with control siRNA) (Fig. 1A). The second siRNA (siLYL1D) had a weaker effect on ARHGAP21 and ARHGAP24 expression than siLYL1A, probably due to its less efficient downregulation of LYL1. As previously reported (32), ITGA2, RAPGEF1, and RAPGEF2 mRNA levels were also reduced in LYL1-depleted HUVECs. ARHGAP21 encodes ARHGAP21 that, in association with ␤-Arrestin1, controls RhoA activity and stress fiber formation (4). ARHGAP24 encodes ARHGAP24 that is preferentially expressed in ECs and specifically stimulates the GTPase activity of RhoA but not of Rac or CDC42 (41). Therefore, to determine whether LYL1 depletion in HUVECs affected RhoA activity, GTP-bound activated RhoA was spe-

5.50 5.25 5.00 4.75 4.50 4.25

LUNG-4m

4m

AJP-Lung Cell Mol Physiol • doi:10.1152/ajplung.00200.2013 • www.ajplung.org

LUNG-2m

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siLYL1-transfected cells were often spindle shaped with actin stress fibers (Fig. 1C, left). Reciprocally, LYL1 overexpression was associated with increased cortical actin (asterisk in Fig. 1C) compared with control cells (Ad-LacZ). These results indicate that LYL1 silencing in human ECs downregulates the expression of two ARHGAP-encoding genes and is associated with increased RhoA activity and reorganization of the actin cytoskeleton into stress fibers. Basal lung vascular permeability is elevated in adult Lyl1⫺/⫺ mice. RhoA regulates the endothelial barrier both in vitro and in vivo by controlling actin stress fiber formation (7). Stress fibers apply mechanical forces to interendothelial junctions, making them potentially permeable. Therefore, we asked whether the lack of LYL1 may affect some endothelial barriers in mice. To this aim, vascular permeability was assessed by quantifying EBD extravasation in different organs of 2- and 4-mo-old Lyl1⫺/⫺ mice. In basal conditions (Fig. 2A, Œ and ), EBD extravasation was 2.4-fold higher in lungs from 4-mo-old Lyl1⫺/⫺ mice than in WT littermates (138 ␮g/g vs.

A

2m

56.3 ␮g/g lung dry weight, respectively). Intravenous injection of rhodamin-labeled 2,000-kDa dextran was used to visualize this increased permeability in lungs of Lyl1⫺/⫺ mice (data not shown). Concomitantly, pulmonary edema, quantified by the lung wet-to-dry-weight ratio, was significantly increased in 4-mo-old Lyl1⫺/⫺ mice (5.1) compared with WT controls (4.8) (Fig. 2D). Conversely, basal lung permeability (around 200 ␮g/g lung dry weight, Fig. 2B) and lung wet-to-dry-weight ratio were comparable in 2-mo-old WT and Lyl1⫺/⫺ mice (Fig. 2E). As thrombin increases lung microvascular permeability through interaction with EC surface PAR-1 (42), we compared the effects of PAR-1 activation on endothelial permeability in Lyl1⫺/⫺ and WT mice following intravenous injection of a PAR-1-specific activating peptide (Fig. 2A, Œ and o). As expected, in WT mice, the PAR-1-activating peptide induced a 2.6-fold increase in lung permeability compared with controls (scrambled peptide). In contrast, it did not further enhance lung permeability in Lyl1⫺/⫺ mice. This lack of effect in Lyl1⫺/⫺

6m

WT

Lyl1-/-

12m

WT Lyl1-/-

120

***

**

***

80

40

0

*

4m

6m

9m

12m

*

D

***

1.6 1.2 0.8 0.4 0

2m

**

Size of the infiltrates (x104 µm2)

C Surface of infiltrates (% of total lung surface)

Number of Infiltrates/cm2

B

6

4 3 2 1 0

2m

4m

6m

9m

12m

*

5

12m

Fig. 3. Lungs from Lyl1⫺/⫺ mice show cellular infiltrates. A: paraffin-embedded lung sections from WT and Lyl1⫺/⫺ mice at 2, 6, and 12 mo of age were stained with hematoxylin-eosin (HE). Entire sections were visualized with a Nanozoomer slide scanner controlled by the NDP.view software. Scale bar ⫽ 100 ␮m. B: using the NDP.view software, we counted manually the number of infiltrates on the entire surface of each lung section and divided by the total lung area. C: surface of infiltrates was manually delineated, measured, and calculated as percentage of the total lung surface. Infiltrate quantification was performed in lungs from 2-, 4-, 6-, 9-, and 12-mo-old WT and Lyl1⫺/⫺ mice. D: taken individually, the infiltrate size was significantly larger in Lyl1⫺/⫺ mice than in WT littermates. B–D: at least 6 animals were analyzed per genotype and per age. Data are presented as the means ⫾ SE. *P ⬍ 0.05, **P ⬍ 0.01, ***P ⬍ 0.001. AJP-Lung Cell Mol Physiol • doi:10.1152/ajplung.00200.2013 • www.ajplung.org

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mice could not be attributed to reduced PAR-1 expression, as PAR-1 mRNA levels in lungs from Lyl1⫺/⫺ and WT littermates were similar (Fig. 2C). On the other hand, heart endothelial permeability was comparable in Lyl1⫺/⫺ and WT mice, both in basal conditions and after PAR-1-activating peptide injection (Fig. 2, A and B). Altogether, these experiments show that, in adult Lyl1⫺/⫺ mice, basal lung vascular permeability is constitutively elevated. In Lyl1⫺/⫺ mice, leukocyte extravasation in the lungs is independent of the hematopoietic defects. Vascular permeability is associated with the presence of cellular infiltrates, macromolecule leakage, and edema (28). To investigate LYL1 effect on cell diapedesis through the endothelial barrier, lungs of WT and Lyl1⫺/⫺ mice at different ages (2, 4, 6, 9, and 12 mo after birth) were collected and processed for histological analysis. HE staining revealed the presence of cell infiltrates already in the lungs from 2-mo-old Lyl1⫺/⫺ mice (Fig. 3A) but not from WT littermates. Quantitative analysis indicated that the number of cell infiltrates was higher in lungs from Lyl1⫺/⫺ mice than in WT littermates at all ages (Fig. 3B) as well as the lung surface occupied by the infiltrates (Fig. 3C). The cell infiltrate number and the occupied lung surface increased with age in Lyl1⫺/⫺ mice, suggesting a continuous increase of the basal lung vascular permeability in these animals. Such infiltrates are commonly seen in the lungs of mice that are continuously under mild antigen challenge, but they

A

Ki67

are usually transient and rapidly resolved thanks to successful immune surveillance. Indeed, some cellular infiltrates were observed also in 12-mo-old WT mice (Fig. 3B); however, their size was significantly smaller than in Lyl1⫺/⫺ mice (Fig. 3D). Lung section immunostaining showed that the cell infiltrates were composed of Ki67⫹ proliferating cells, including CD3⫹ T lymphocytes, PAX5⫹ B lymphocytes, and MAC2⫹ macrophages (Fig. 4A). HE staining highlighted the presence of polynuclear neutrophils, red blood cells, and large brown cells that resembled pigmented macrophages (see inset in Fig. 4A). Staining with Perls’ Prussian blue dye confirmed the presence of endocytosed red blood cells in the pigmented macrophages, as indicated by the blue coloration of hemosiderin, reflecting the degradation of hemoglobin (Fig. 4A). Altogether, these findings indicate that, in Lyl1⫺/⫺ mice, leukocyte extravasation in the pulmonary parenchyma is abnormal already at 2 mo of age. Inflammatory cell infiltrates, vascular leakiness, and edema are often associated with inflammation. However, we did not detect any difference in the mRNA expression of inflammatory cytokines (IL-1␤, monocyte chemoattractant protein-1, IL-12␤ p40, IFN-␥, TNF-␣, IL-4, IL-5, IL-6, and IL-10) between WT and Lyl1⫺/⫺ lungs (data not shown). LYL1 is expressed in several hematopoietic cell types, including B cells and immature myeloid cells that are important actors of inflammatory responses (45). Although the hematopoietic defects of Lyl1⫺/⫺ mice are not severe (12, 52), we could not exclude that they might affect directly the

Perls staining

Mac2

Pax5

CD3

* C ** 120

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6m

Number of Infiltrates/cm2

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1.2

0.8

0.4

0

6m + FL WT Ly5.1

Fig. 4. Infiltrates are composed of immune cells, and their occurrence in the lungs of Lyl1⫺/⫺ mice is independent of their hematopoietic phenotype. A: representative images of the cell composition of an infiltrate in the lung of a 1-yr-old Lyl1⫺/⫺ mouse. Paraffin-embedded lung sections were immunostained using anti-Ki67 (proliferating cells), anti-CD3⑀ (T cells), anti-PAX5 (B cells), and anti-MAC2 (macrophages) antibodies. Scale bar ⫽ 50 ␮m. Left, inset: pigmented macrophages (*). Left, middle and middle insets: confirm the cytoplasmic and nuclear staining of these 2 antibodies, respectively. Perls’ Prussian staining reveals the presence of hemosiderin (blue) in pigmented macrophages, whereas the nuclei are in red. B: fetal liver (FL) cells from Ly5.1 WT mice were grafted in irradiated Ly5.2 WT and Lyl1⫺/⫺ mice. Hematopoietic reconstitution was confirmed 1 mo later (85.8% ⫾ 4.6 and 94.4% ⫾ 1.6 of Ly5.1-positive peripheral blood mononuclear cells in the transplanted WT and Lyl1⫺/⫺ mice, respectively). At 6 mo of age, lungs were collected and HE stained. Scale bar ⫽ 100 ␮m. Quantification of the number of infiltrates/cm2 and the surface occupied by the infiltrates was performed as described in Fig. 3. 5 animals were analyzed per genotype. Data are presented as the means ⫾ SE. *P ⬍ 0.05, **P ⬍ 0.01. AJP-Lung Cell Mol Physiol • doi:10.1152/ajplung.00200.2013 • www.ajplung.org

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immune response to airways antigens. To evaluate this possibility, we examined the presence of cell infiltration in lungs of hematopoietic cell-depleted Lyl1⫺/⫺ mice after hematopoietic reconstitution with immature progenitors from WT mice. FL cells from 14.5 dpc Ly5.1 WT mice were transplanted into irradiated 6-wk-old Ly5.2 WT or Lyl1⫺/⫺ recipients. Hematopoietic reconstitution was assessed after 1 mo, and, after another 3.5 mo, lungs were collected and processed for histological analysis. After hematopoietic reconstitution, the number and the lung surface occupied by cell infiltrates in the lungs of 6-mo-old Lyl1⫺/⫺ recipients were significantly higher than in WT recipients (Fig. 4, B and C) and comparable to what was observed in Lyl1⫺/⫺ mice. These experiments demonstrate that the abnormal immune cell infiltration in the lungs of Lyl1⫺/⫺ mice is not related to their hematopoietic defects. AJ in the lungs of Lyl1⫺/⫺ mice are impaired. VE-cadherin, the major component of endothelial AJs, regulates the endothelial barrier function, and its cell surface localization is maintained through association with p120-catenin (13, 50). Therefore, we analyzed VE-cadherin and p120-catenin localization in the lung endothelium of WT and Lyl1⫺/⫺ mice. Quantification of VE-cadherin staining at cell-cell junctions revealed a significant decrease (down to 44.9% ⫾ 17.6) in the lungs of Lyl1⫺/⫺ mice compared with WT littermates (Fig. 5A). Expression of p120-catenin was also reduced in the lungs of Lyl1⫺/⫺ mice compared with controls (Fig. 5B). Twofold magnification (Fig. 5, A and B, bottom) allowed highlighting the precise cell surface expression of VE-cadherin and p120-catenin in WT lungs compared with the diffuse, cytoplasmic, or irregular staining in Lyl1⫺/⫺ mice. Concomitant staining of lung ECs with anti-CD31 antibodies or lectin (Fig. 5) did not detect any other obvious difference between WT and Lyl1⫺/⫺ mice. Therefore, the poor recruitment of

B

***

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WT

Lyl1-/-

DAPI-Lectin

Lyl1-/-

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WT

VE-cadherin at cell-cell junctions (arbitrary unit)

DAPI-VEcad

DAPI-CD31

A

both VE-cadherin and p120-catenin to ECs indicates that endothelial AJs are specifically impaired in Lyl1⫺/⫺ lungs. ECs isolated from Lyl1⫺/⫺ lung show reduced expression of integrin-␣2, RapGEF1, and RapGEF2 and increased RhoA activity. To further investigate the mechanisms by which LYL1 regulates VE-cadherin recruitment to endothelial AJs and more generally vascular permeability, we isolated ECs from lungs (mLECs) of WT and Lyl1⫺/⫺ mice. Lungs were excised and dissociated, and mLECs were positively selected with an anti-CD146 antibody. To assess mLEC enrichment, CD31⫹/ CD45⫺ cells were quantified by flow cytometry in the bulk and mLEC fractions (31.6 ⫾ 7% and 79.3 ⫾ 8.1% for WT mice and 33.9 ⫾ 4.9% and 81.6 ⫾ 4.4% for Lyl1⫺/⫺ mice, respectively) (Fig. 6A, top). Furthermore, ␣-SMA staining indicated that contamination of WT and Lyl1⫺/⫺ mLECs by smooth muscle cells/fibroblasts was lower than 5% (Fig. 6A, bottom), allowing us to ascribe the detected mRNA and protein expression levels mainly to ECs. In agreement with our previous results obtained in LYL1-depleted human ECs (32), VEcadherin mRNA and total protein levels were comparable in Lyl1⫺/⫺ and WT mLECs (Fig. 6, B and C), indicating that VE-cadherin expression is not directly controlled by LYL1. Similarly to what was observed in LYL1-depleted human ECs (see Fig. 1A), Itga2 mRNA levels and integrin-␣2 protein expression, which has an important role in EC-ECM attachment, were significantly reduced in Lyl1⫺/⫺ mLECs compared with WT cells (Fig. 6, B and C). As previously demonstrated in LYL1-depleted human ECs (32), RapGEF1 and RapGEF2 mRNA levels were also significantly reduced in Lyl1⫺/⫺ mLECs (Fig. 6B), suggesting a reduction in active Rap1. Although ArhGAP21 and ArhGAP24 mRNA levels were similar (Fig. 6B), basal level of active RhoA was significantly higher in Lyl1⫺/⫺ than in WT mLECs (Fig. 6D).

Fig. 5. Vascular endothelial (VE)-cadherin and p120-catenin recruitment at cell-cell junctions are concomitantly reduced in lungs of 6-mo-old Lyl1⫺/⫺ mice. Microscopy images illustrating the strong reduction of VE-cadherin (A) and p120-catenin (B) recruitment at cell-cell junctions in 6-mo-old Lyl1⫺/⫺ mice compared with WT littermates. Lung cryo-sections were double stained for endothelial cell visualization (green) with anti-CD31 antibody (A) or FITC-labeled lectin (B) and with specific antibodies against VE-cadherin (A) or p120-catenin (B) (red). Bottom: 2-fold magnification of the insets (dotted lines) in middle. Scale bar ⫽ 15 ␮m. VE-cadherin expression (A) was quantified in 5 random fields with the ImageJ software. Data are presented as the means ⫾ SE (4 WT and 7 Lyl1⫺/⫺ individual lungs were analyzed). VE-cadherin intensity in WT was set at 100. ***P ⬍ 0.001. AJP-Lung Cell Mol Physiol • doi:10.1152/ajplung.00200.2013 • www.ajplung.org

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INTEGRIN- 2 (150 kDa)

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CD31 (130 kDa)

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-ACTIN (45 kDa)

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100 80 60 40 20 0 VEINTEGRIN2 CADHERIN

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Arh GAP21

5

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WT Lyl1-/-

-SMA

Fig. 6. Lyl1⫺/⫺ mouse lung ECs (mLECs) show reduced expression of integrin-␣2, RapGEF1, and RapGEF2 and increased RhoA activity. A: lungs were dissociated to obtain a single cell suspension, and the bulk fraction was separated into CD146- cells and CD146⫹ mLECs, as described in MATERIALS AND METHODS. Cells were labeled with anti-CD31 and anti-CD45 antibodies to visualize ECs and hematopoietic cells, respectively, or with an anti-␣-smooth muscle actin (SMA) antibody to evaluate smooth muscle cell contamination. After dead cell exclusion, the percentages of ECs, hematopoietic cells, and smooth muscle cells were evaluated in the 3 fractions. Dot plots are representative of 6 independent preparations of mLECs. B: mRNA levels of the indicated genes in mLECs obtained from WT (solid bars) or Lyl1⫺/⫺ (open bars) mice were assessed by qRT-PCR. cDNAs were amplified with specific primers, and value was normalized to CD31 mRNA. Each bar is the mean ⫾ SD of 3 independent experiments. For each gene, the WT value was arbitrarily set to 100. *P ⬍ 0.05; **P ⬍ 0.01; ***P ⬍ 0.001 by using the Student’s t-test. C: integrin-␣2, VE-cadherin, and CD31 protein expression in whole cell lysates from CD146- cells and CD146⫹ mLECs were assessed by immunoblotting. ␤-Actin expression was used to control protein loading. Scanned autoradiographies were quantified using ImageJ to determine the protein levels relative to CD31 protein expression (a measure of EC content). Each bar is the mean ⫾ SD of 3 independent mLEC preparations. For each protein, the WT value (solid bars) was arbitrarily set to 100. **P ⬍ 0.01 by using the Student’s t-test. D: RhoA activity was assessed in whole cell extracts from WT or Lyl1⫺/⫺ mLECs. Scanned autoradiographs were quantified using ImageJ to determine the GTP-bound RhoA/total RhoA ratio for each extract. Bars show the means ⫾ SD of the GTP-bound RhoA/total RhoA ratio of 3 independent experiments. The WT mLECs ratio was arbitrarily set to 100. Images shown are representative of 3 experiments. *P ⬍ 0.05 using the Student’s t-test.

These findings extend to the adult mouse endothelium our previous results obtained in cultured human ECs showing that LYL1 depletion affects the expression and/or the localization of several important actors at endothelial junctions. DISCUSSION

We previously reported that lack of LYL1 affects the maturation of newly formed blood vessels in adult mice and

reduces active Rap1 in ECs (32). Here, we completed our analysis of LYL1 function by showing that it is required also for the formation of the endothelial barrier specifically in mature lungs by acting upstream of VE-cadherin and of RhoA. Rap1 and RhoA are among the small GTPases involved in the regulation of the endothelial barrier either by reorganizing cortical actin cytoskeleton or by controlling VE-cadherincatenin association at AJs (38). Specifically, Rap1 is involved

AJP-Lung Cell Mol Physiol • doi:10.1152/ajplung.00200.2013 • www.ajplung.org

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in the establishment and maintenance of microvascular endothelial barrier functions, whereas RhoA primarily acts antagonistically to disrupt barrier integrity. Here we show that, in addition to Rap1 (32), LYL1 also modulates RhoA activity. Different mechanisms have been reported to regulate RhoA activity through the successive action of other small GTPases. In confluent cells, active Rap1 initiates the activation of Vav2, which in turn activates Rac1 (10) that ultimately will lead to RhoA inactivation (7). In addition, Rap1 interaction with afadin drives barrier integrity restoration via a negative feedback regulation of RhoA (9). Recently, it was shown that the interaction of active Rap1 with the adaptor protein Ras-interacting protein 1 inhibits RhoA activation through ARHGAP29 modulation (33). Thus the reduced expression of the two RapGEFs induced by LYL1 depletion in human ECs (32) and mouse lung endothelium (this study) may contribute to RhoA activation as a consequence of Rap1 inhibition. LYL1 depletion in human ECs reduces the expression of ARHGAP24 and ARHGAP21 genes, which encode two RhoGAPs (4, 41), and this may also partly account for the strong increase in active RhoA. We did not observed significant change in ARHGAP24 and ARHGAP21 gene expression, consistent with the lower increase of active RhoA in Lyl1⫺/⫺ mLECs (0.5-fold) compared with LYL1-depleted HUVECs (2-fold). This difference might be due to the origin of the ECs, microvessel vs. large vessel. Whatever the mechanisms leading to RhoA activation in ECs, the primary consequence is stress fiber formation and cell contraction. Indeed, LYL1 depletion in ECs induced stress fiber formation, whereas LYL1 overexpression reduced stress fibers, leading to increased cortical actin. Association of VE-cadherin with p120-catenin regulates the formation and stabilization of AJs (24, 49, 50), leukocyte transmigration (1), and lung vascular permeability (44). We previously reported that LYL1-depleted HUVECs display discontinuous staining of VE-cadherin with obvious gaps between adjacent cells (32). Here, we show poor recruitment of both VE-cadherin and p120-catenin at AJs in the lungs of Lyl1⫺/⫺ mice, indicating loose interendothelial junctions that could cause the high basal vascular permeability observed in Lyl1⫺/⫺ lungs. The poor recruitment of both VE-cadherin and p120-catenin at AJs could be a consequence of reduced active Rap1 in Lyl1⫺/⫺ lungs. Indeed, active Rap1 stimulates AJ annealing, through VE-cadherin (18) and p120-catenin localization at AJs (8). Through distinct mechanisms, p120-catenin controls the activity of small GTPases and notably represses RhoA activity (3, 29, 48). Thus a reduction of p120 catenin at endothelial AJs in the lungs of Lyl1⫺/⫺ mice could also contribute to the increased RhoA activation observed in mLECs isolated from Lyl1⫺/⫺ lungs. RhoA is a key regulator of the endothelial barrier. Active RhoA is involved in thrombin-induced lung barrier leakiness (46), and basal endothelial lung permeability is high in RhoGDI-deficient mice (20). Conversely, inhibition of RhoA activity reduces basal permeability both in microvascular ECs in vitro and postcapillary venules in vivo (38). In Lyl1⫺/⫺ mice, lung vascular permeability is elevated and is associated with leukocyte infiltration in the pulmonary parenchyma, macromolecule extravasation, and edema. On the basis of our data in HUVECs (32) and in mLECs (this study), we propose that the concomitant increase in active RhoA and decrease in active Rap1 contribute to the lung endothelial barrier leakiness in

Lyl1⫺/⫺ mice. As RhoA activation mediates disruption of the endothelial barrier induced by PAR-1 treatment (43), constitutive RhoA activation in Lyl1⫺/⫺ lung endothelium may explain why PAR-1 activation does not further enhance lung permeability in these mice. In addition, the reduction of integrin-␣2 in Lyl1⫺/⫺ mLECs might affect cell-ECM attachment in the lung endothelium and also contribute to the loss of vascular endothelium integrity in Lyl1⫺/⫺ mice, as already shown for integrin-␣v␤5 (40). Rap1 positively regulates integrin signaling, and its inactivation impairs cell-ECM adhesions (11). Modulation of integrin-␣2 expression could also be linked to RhoA activation because phosphatidylinositol 3-AKT signaling activation downstream of integrin-␣2␤1 leads to ARHGAP26-dependent RhoA inactivation (39). In addition, deletion of the Rap1 activator RapGEF2 causes embryonic lethality at E9.5 due to hemorrhages caused by defective blood vessels (47). Similar defects have been reported also in caveolin-1-deficient mice that display alterations in lung endothelial junctions associated with increased accumulation of albumin in lung tissue (34). Likewise, endothelial-specific deletion of HIF-2␣ triggers increased dye extravasation, leukocyte infiltration in lung connective tissue, and enhanced vessel permeability (35). Four-month-old Lyl1⫺/⫺ mice show high lung permeability similar to what observed in 2-mo-old WT and Lyl1⫺/⫺ mice. LYL1

Rap1-RIAM

Rap1 translocation at the membrane

INTEGRIN 2 1

ECM-EC Contact

Rap-GEFs

Rho-GAPs

(Rap-GEF1 & 2)

(ARHGAP21 & 24)

Rap1 Activation

RhoA Activation

Rap1-Afadin

p120-catenin and VE-cadherin recruitment at AJs

VE-cadherin stabilization at AJs

Actin cytoskeleton contraction

ENDOTHELIAL BARRIER FUNCTION Fig. 7. Model illustrating how LYL1 regulates the endothelial barrier through Rap1 activation. LYL1 induces Rap1 activation by upregulating the expression of 2 RapGEFs. Then, active Rap1 associates with effector proteins like RIAM and translocates at the membrane to enhance Talin-integrin association and thereby increases integrin affinity to extracellular matrix (ECM) molecules (11). EC attachment to ECM is further increased by the direct upregulation of integrin-␣2 expression by LYL1. In addition, the interaction of active Rap1 with Afadin favors the association of p120-catenin with VE-cadherin at adherens junctions (AJs), thereby stabilizing endothelial junctions (8, 18). Finally, RhoA activity is decreased through the inhibitory action of active Rap1 (9, 33) and LYL1-mediated downregulation of 2 RhoGAPs.

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In newborn mice, alveolar formation is incomplete with highly immature and still remodeling capillary networks (2). Maturation of alveoli and capillaries is complete after 1–2 mo at the end of the alveolar stage. Thus our findings suggest that, in the absence of Lyl1, lung capillaries remain immature, similarly to the angiogenic blood vessels observed in tumors developed in Lyl1⫺/⫺ adult mice (32). In summary, we propose a model in which LYL1 acts upstream of several key players involved in endothelial barrier function, such as integrins, the small GTPases Rap1 and RhoA, and the AJ proteins VE-cadherin and p120-catenin (Fig. 7). Regardless of the upstream mechanism, both active RhoA and decreased clustering of VE-cadherin at endothelial junctions contribute to lung vascular leakiness. In humans, RhoA-mediated endothelial permeability plays a key role in the pathogenesis of pulmonary complications associated with severe acute pancreatitis (16), and reduction of VE-cadherin might explain the increased permeability and enhanced leukocyte transmigration in acute respiratory distress syndrome (22). Altogether, the knowledge of the molecular organization and function of endothelial junctions indicates that cell-cell junctions could be “molecular targets” for therapeutic intervention to counteract the disruption of vascular integrity that accompanies inflammatory pathologies. Accordingly, on the basis of our findings that LYL1 functions as a signal for inducing and maintaining vessel integrity, molecules that modulate LYL1 activity could represent additional tools to modify endothelial barrier permeability. ACKNOWLEDGMENTS We are grateful to the “Réseau des Animaleries Montpelliéraines” RAMIBiSA Facility for animal experiments. We are indebted to the MontpellierRIO Imaging Facility for microscopic image acquisition and analysis and to the “Réseau d’Histologie Experimentale de Montpellier”-RHEM Facility for expert assistance with histology. Present address for Nelly Pirot: Institut de Recherche en Cancérologie de Montpellier (U896), CRLC Val d’Aurelle, Paul Lamarque, 208 rue des Apothicaires, 34298 Montpellier cedex 5, France. Present affilation for Elias Chalhoub: FHS-University of Balamand, Beirut, Lebanon. GRANTS N. Pirot was supported by successive studentships from The French Minister of Higher Education and Research and from the “Ligue Nationale Contre le Cancer” (France). D. Mathieu and V. Pinet are supported by INSERM. This work was funded by grants from the Association pour la Recherche sur le Cancer (ARC, France) and Agence Nationale pour la Recherche (ANR-MNP, France). DISCLOSURES No conflicts of interest, financial or otherwise, are declared by the authors. AUTHOR CONTRIBUTIONS Author contributions: N.P. and V.P. conception and design of research; N.P., H.D., V.D., C.D., E.C., and V.P. performed experiments; N.P., H.D., V.D., C.D., M.C.-S., C.B.-L., E.C., and V.P. analyzed data; N.P., M.C.-S., C.B.-L., D.M., and V.P. interpreted results of experiments; N.P., D.M., and V.P. approved final version of manuscript; D.M. and V.P. edited and revised manuscript; V.P. prepared figures; V.P. drafted manuscript. REFERENCES 1. Alcaide P, Newton G, Auerbach S, Sehrawat S, Mayadas TN, Golan DE, Yacono P, Vincent P, Kowalczyk A, Luscinskas FW. p120-Catenin regulates leukocyte transmigration through an effect on VE-cadherin phosphorylation. Blood 112: 2770 –2779, 2008.

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