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ABSTRACT
As activation of the coagulation system is both a consequence and contributor to acute
lung injury (ALI), pulmonary coagulopathy has become a potential target for therapeutic
intervention in ALI patients. We investigated the effects and possible mechanisms
of endothelial cell (EC)-anchored tissue factor pathway inhibitor (TFPI) on lipopolysaccharide
(LPS)-induced ALI in mice. To assess the effect of EC-anchored TFPI deletion on ALI
indices, TFPI knockout (cKO) mice were generated. Mice were instilled by direct intratracheal
injection LPS for the preparation of an ALI model. Evans blue dye (EBD) was injected
intravenously 2 h prior to animal sacrifice (48 h post-LPS). Lungs were fixed for
histopathology and the prepared tissue was homogenized or used to extract bronchoalveolar
lavage fluid (BALF) or detect EBD concentration. TFPI knockdown mice with ALI were
compared to wild-type (WT) mice with ALI to assess the effect of TFPI on endothelial
barrier function and inflammation. TFPI deletion markedly exacerbated LPS histopathological
changes in lung, and the LPS changes in protein, EBD extravasation, proinflammatory
cytokines TNF-α, IL-1β, and IL-6 in BALF in lung. The number and infiltration of white
blood cells (WBCs) from BALF and lung tissue of TFPI cKO mice with LPS-challenged
ALI was increased compared to WT mice with LPS-challenged ALI. We also found further
increased toll-like receptor 4 and nuclear factor kappa-light-chain-enhancer of activated
B cells activation and additional expression of vascular cell adhesion molecule 1
and reduction of angiotensin converting enzyme 2 expression in TFPI cKO+LPS mice compared
with WT+LPS mice. Endothelial-specific TFPI deficiency promoted LPS-induced pulmonary
inflammation and endothelial barrier permeability possibly via toll-like receptor
4-mediated nuclear factor kappa-light-chain-enhancer of activated B cells signaling
pathway activation.
Acute lung injury (ALI) in septic patients is characterized by increased lung vascular permeability and severe lung inflammation, which typically develop in concert and lead to progressive deterioration of lung function (Diaz et al., 2010). LPS, a cell wall component of Gram-negative bacteria, is a causative agent implicated in the pathogenesis of ALI (Andonegui et al., 2003, 2009; Everhart et al., 2006; Mehta and Malik, 2006; Bachmaier et al., 2007; Diaz et al., 2010; Karpurapu et al., 2011). Studies showed that endothelial cells (ECs) are crucial in mediating the lung’s inflammatory response by LPS (Andonegui et al., 2003, 2009). LPS binds the endothelial Toll-like receptor 4 (TLR4) via CD14, a membrane-bound glycosylphosphatidyl inositol–anchored protein (Andonegui et al., 2002; Kawagoe et al., 2008; Lloyd-Jones et al., 2008). TLR4 in turn activates signaling pathways responsible for the generation of proinflammatory cytokines via myeloid differentiation factor 88 (MyD88; Kawai et al., 1999; Medvedev et al., 2002; Bachmaier et al., 2007; Kawagoe et al., 2008). MyD88 contains the Toll-IL1-R homology (TIR) domain and death domain through which MyD88 recruits IL-1R–associated kinase 4 (IRAK4) to the Toll/IL-1 signaling domain, resulting in IRAK4 activation (Medvedev et al., 2002; Kawagoe et al., 2008). IRAK4 activates its effectors IRAK2 and IRAK1 to induce activation of NF-κB and other transcription factors required for the generation of proinflammatory cytokines and reactive oxygen species (ROS) and the activation of MAPK (Medvedev et al., 2002; Kawagoe et al., 2008; Kawai and Akira, 2010; Takeuchi and Akira, 2010). LPS induces lung neutrophil sequestration, as well as neutrophilic and macrophage generation of cytokines and ROS (Andonegui et al., 2002, 2003, 2009; Gao et al., 2002; Garrean et al., 2006; Bachmaier et al., 2007; Xu et al., 2008; Di et al., 2012), which contribute to the development of ALI (Gao et al., 2002; Bachmaier et al., 2007; Di et al., 2010, 2012). However, ECs may also have a more direct role in mediating LPS-induced loss of lung vascular barrier function and inflammation (Andonegui et al., 2002, 2003, 2009; Wang et al., 2011). A rise in intracellular Ca2+ is an essential signal required for EC contraction that precedes endothelial barrier disruption (Mehta et al., 2003; Pocock et al., 2004; Cheng et al., 2006; Mehta and Malik, 2006; Singh et al., 2007; Kini et al., 2010; Weissmann et al., 2012). It remains unknown whether Ca2+ signaling intersects with the TLR4 signaling pathway, and hence contributes to LPS-induced endothelial permeability and inflammation. Diacylglycerol (DAG), a membrane phospholipid–derived second messenger generated by LPS (Sands et al., 1994; Yamamoto et al., 1997; Monick et al., 1999; Zhang et al., 2001; Xu et al., 2005; Zhang et al., 2011), is produced upon hydrolysis of phosphatidylcholine (PC) by PC-specific phospholipase (PLC; Sands et al., 1994; Yamamoto et al., 1997; Zhang et al., 2001; Xu et al., 2005). Yamamoto et al. (1997) demonstrated that LPS induces DAG generation by binding to CD14 (Yamamoto et al., 1997), a component of LPS-binding TLR4 complex in ECs (Andonegui et al., 2002; Lloyd-Jones et al., 2008). Importantly, DAG is known to activate transient receptor potential canonical 6 (TRPC6) channels, a nonselective Ca2+ permeable ion channel (Hofmann et al., 1999; Dietrich et al., 2005a), which was shown to induce endothelial contraction (Pocock et al., 2004; Singh et al., 2007; Kini et al., 2010; Weissmann et al., 2012). In this study, we tested the hypothesis that LPS ligation of TLR4 and resulting TRPC6-dependent Ca2+ signaling intersect to mediate both the vascular leak and inflammatory features of ALI. RESULTS LPS-induced DAG generation stimulates Ca2+ entry and Ca2+ current in ECs via TRPC6 We first addressed whether LPS generates the second messenger DAG in ECs, which in turn can activate Ca2+ entry via TRPC6. We transduced FRET-based DAG (Violin et al., 2003) and Ca2+ reporters (Kim et al., 2009) in ECs isolated from WT or Trpc6−/− mouse lungs, which were stimulated with LPS. Mouse lung ECs (MLECs) were identified using VE-cadherin and PECAM (CD31) antibodies (Fig. 1 A). LPS exposure of MLECs increased DAG generation and cytosolic Ca2+ in a time-dependent manner, as evident from DAG and Ca2+ reporter activities (Fig. 1 C). TRPC6 deletion did not impair DAG generation by LPS (Fig. 1 C); rather, it prevented the increase in cytosolic Ca2+ (Fig. 1 D). Loss of TRPC6 also did not alter the expression profile of other TRPC channels (Fig. 1 B). Figure 1. LPS-generated DAG induces Ca2+ entry and activates Ca2+ current in lung ECs in a TRPC6-dependent manner. (A) Dot plot analysis of EC surface markers was performed with WT-MLECs immunostained with APC-tagged VE-cadherin and PE-tagged CD31. WT-MLECs immunostained with APC-IgG and PE-IgG were used as isotype controls. Data are representative of at least two independent experiments. (B) RNA extracted from WT and Trpc6−/− MLECs were reverse-transcribed using suitable primers, as described in the Materials and methods. GAPDH was used as internal control. C1, C4, C3, C6, C7, and GDH represent TRPC1, TRPC4, TRPC3, TRPC6, TRPC7, and GAPDH, respectively. (C and D) WT or TRPC6−/− MLECs were transduced with FRET-based DAG reporter (DAGR; C) or Ca2+ reporter (D). Cells were then stimulated with 1 µg/ml LPS and dynamic changes in CFP, FRET, and YFP were acquired. DAG and Ca2+ reporter activities were calculated by ratioing CFP/FRET intensity before and after stimulation with LPS (left). Plot shows mean ± SEM from three individual experiments (right). * indicates a significant increase in reporter activity over time zero (P 90% in TRPC6−/− MLECs (Fig. 1, G [bottom] and H), indicating the dominant role of TRPC6 channels in the response. We observed that 5 µg/ml LPS did not alter cell viability in that ECs remained fully responsive to thrombin after stimulation (Fig. 1 I). In other studies, we used 1-oleoyl-2-acetyl-sn-glycerol (OAG), a cell-permeable DAG analogue, to address whether DAG activation of TRPC6 could increase lung vascular permeability. Using isolated/perfused mouse lungs (described in Materials and methods), we determined pulmonary microvessel filtration coefficient (K f,c), a measure of vascular endothelial permeability to liquid (Vogel et al., 2000; Tauseef et al., 2008). OAG produced a fourfold increase in K f,c in WT mouse lungs, but failed to elicit a response in Trpc6−/− lungs (Fig. 2 B). Importantly, basal K f,c values did not differ in WT and Trpc6−/− lungs (Fig. 2 A). To determine whether OAG could also increase lung vascular permeability in vivo (as opposed to the aforementioned isolated/perfused organ), we injected OAG or control vehicle i.v. in mice and measured pulmonary transvascular leakage of Evans blue–labeled albumin (Tauseef et al., 2008; Knezevic et al., 2009). Infusion of OAG increased transendothelial albumin influx in WT lungs whereas the response was inhibited in Trpc6−/− lungs (Fig. 2 C). OAG increased intracellular Ca2+ in WT-MLECs but not in Trpc6−/− MLECs (Fig. 2 A). OAG did not increase intracellular Ca2+ in WT-MLECs in the absence of extracellular Ca2+ (Fig. 2 A). Together these studies show that OAG activates the TRPC6 Ca2+ entry pathway to induce lung vascular permeability. Figure 2. Direct activation of TRPC6 increases lung vascular permeability. (A) WT or Trpc6−/− MLECs were loaded with Fura 2-AM for 25 min, after which they were exposed to buffer containing 2 mM Ca2+ or Ca2+-free buffer. Changes in intracellular Ca2+ were determined in response to OAG by converting 340:380 excitation ratio intensity, as described in Materials and methods. Each representative tracing is the mean calcium response of 20–25 cells in a given field. These experiments were repeated three times. (B) Isogravimetric WT or Trpc6−/− mice lungs were perfused with 100 µM OAG for 20 min, after which intravascular pressure was raised by 10 cm of H2O. Microvascular filtration coefficient (K f,c) was determined from the slope of lung wet weight gain after normalization by lung dry weight, as described in Materials and methods. Plot shows mean ± SEM of K f,c from four individual experiments. * indicates a significant increase in K f,c over the values without OAG (0 µM; P 90% pure. HPAECs were cultured in a T-75 flask coated with gelatin and growth factor–supplemented EGM2 medium, as previously described (Knezevic et al., 2007). Transfection of cells. cDNA was transduced into MLECs using FuGENE transfection reagent according to the manufacturer’s protocol. Human pulmonary artery ECs were transfected with cDNA using a Nucleofector device obtained from Lonza (Singh et al., 2007; Kini et al., 2010). Dominant-negative (DN)-TRPC6 mutant (T. Sharma, University of Illinois, Chicago, IL) was generated by deleting LFW sequence (Δ678-680 aa) from full-length TRPC6 cDNA. LFW motif is a structural part of the pore helix of TRPC6 channel (Hofmann et al., 1999) Plasmid pEYFP-N1-TRPC6-WT containing the full-length Homo sapiens TRPC6 (available from GenBank under accession no. NM_004621) was used as a DNA template with DNA polymerase Phusion (Finnzymes) in a two-step PCR method to generate C-terminal YFP tag TRPC6 with deletion of LFW (aa 678–680). In the first step, two PCR fragments were generated using the following primer pairs: fragment A primer pairs, GFP-494-F, 5′-ATGTCGTAACAACTCCGCCCCATTGA-3′, and TRPC6-dAA678-80-R, 5′-CAAAATATAGC/TGTCTTAAAACTCTCTTCAACTG-3′; fragment B primer pairs, TRPC6-dAA678-80-F, 5′-TTTAAGACA/GCTATATTTGGACTTTCTGAAGTG-3′, and YFP-820-R, 5′-TGAACTTCAGGGTCAGCTTGC-3′. As shown in the primers TRPC6-dAA678-80-F and TRPC6-dAA678-80-R, the forward slash indicated the three amino acids being deleted (aa 678–680 or LFW residues). In the second step of PCR, a final PCR product, fragment C was generated by combining fragments A and B together as PCR DNA templates with primer pair GFP-494-F and YFP-820-R. Fragment C was digested with restriction enzymes 5′-EcoR1 and 3′-AgeI (New England Biolabs, Inc.) and ligated to the pEYFP-N1 vector (BD) also digested at the same restriction enzyme sites. The resulting plasmid was verified and analyzed by gel analysis and sequencing analysis. Immunoprecipitation. Lungs or ECs were lysed in modified RIPA buffer, and equal amounts of protein were immunoprecipitated with appropriate antibodies over night at 4°C, followed by the addition of protein A/G agarose beads for 4 h at 4°C, as described previously (Knezevic et al., 2009). For determining IRAK4 interaction with MYLK or MyD88, blots were first incubated with anti-IRAK4 antibody overnight, followed by reincubation with conformation specific mouse anti–rabbit antibody (Cell Signaling Technology) to eliminate the background from heavy chain. RT-PCR. Total RNA was isolated from mice lungs or MLECs using TRIzol agent (Invitrogen) according to the manufacturer’s instructions. RNA was quantified spectrophotometrically and reverse-transcribed using specific primers to determine the expression of various TRPC or ICAM-1 (Tiruppathi et al., 2002; Dietrich et al., 2005b; Tauseef et al., 2008). FRET-based DAG generation and Ca2+ entry assessment. WT or Trpc6−/− MLECs were transfected with FRET-based DAG (DAGR; A. Newton, University of California, San Diego, La Jolla, CA) or Ca2+ reporter (Y. Wang, University of Illinois, Champaign, IL), and DAG generation and Ca2+ mobilization were determined by direct FRET analysis in live cells. DAGR mutant was purchased from Addgene (plasmid #14865). Cells were stimulated with 1 µg/ml LPS and images were captured every 15 s. An image stack was generated with a 458-nm laser line spanning and emission wavelength ranging from 463–602 nm with 10.7-nm bandwidths. At each time point, three images were recorded: CFP, FRET, and YFP. All images were corrected for shading and background was subtracted. Threshold FRET images were used to generate a binary mask that was then divided by the corresponding CFP images to yield a ratio image reflecting DAG generation and Ca2+ mobilization. Data were analyzed by MetaMorph software. Cytosolic Ca2+ measurements. An increase in intracellular Ca2+ was measured using the Ca2+-sensitive fluorescent dye Fura 2-AM as previously described (Singh et al., 2007). The intracellular Ca2+ concentration ([Ca2+]i) was calculated using Invitrogen Fura-2 Ca2+ Imaging Calibration kit according to the manufacturer’s protocol and the equation: [Ca2+]free = K d EGTA × [(R − Rmin)/(Rmax-R)] × [F380 max/F380 min]. K d represent the dissociation constant of Fura2–Ca2+ interaction, R is fluorescence ratio, Rmin is the ratio at zero free calcium, Rmax is the ratio at saturation calcium (39 µM), F380 max is the fluorescence intensity with excitation at 380 nm, for zero free Ca2+; and F380 min is the fluorescence intensity at saturating free Ca2+. Patch-clamping of ECs. We performed the whole-cell patch-clamp technique to determine cationic currents, as previously described (Hofmann et al., 1999; Obukhov and Nowycky, 2005). The currents were acquired during 300-ms voltage ramps from −100 to +100 mV, with a 2-s inter-ramp interval using the Optopatch amplifier controlled by PCLAMP 10 software (Molecular Devices). The current amplitudes at −90 mV were plotted as a function of time. The pipette solution contained (in mM): 10 EGTA; 3.77 CaCl2; 2 MgCl2; 125 CsMeSO3; and 10 Hepes. The standard extracellular solution contained (in mM): 145 NaCl; 1.2 CaCl2; 1 MgCl2; 2.5 KCl; 10 Hepes; and 5.5 Glucose. The pH of all solutions was adjusted to 7.2. The PCLAMP 10 software package (Molecular Devices) was used for data analysis. All electrophysiological experiments were performed at room temperature (22–23°C). LPS challenge. Trpc6−/− , Mylk−/− and WT mice housed in sealed container were exposed to a nebulized solution of lyophilized E. coli LPS in sterile saline (1 mg/ml) for 1h at a driving flow rate (8 liter/min) using a small volume nebulizer (Resigard II; Marquest Medical, Englewood, CO) and sacrificed after the indicated times (Tauseef et al., 2008). Polymicrobial sepsis model. Polymicrobial sepsis was induced by CLP as previously described (Bachmaier et al., 2007; Toya et al., 2011). In brief, animals were anesthetized using 100 mg/kg ketamine and 15 mg/kg xylazine. Cecum was exposed via a midline abdominal incision, ligated below the ileo-cecal valve, and punctured using 18-gauge needle in mesenteric-antimesenteric direction. Peritoneal and skin wounds were closed using 6–0 silk sutures. Immediately after surgery, mice were resuscitated by subcutaneous injection of 1 ml of prewarmed 0.9% saline solution. Mice were monitored every 6 h for 7 d to determine their survival. Lung vascular liposomal delivery of cDNA for rescue experiments. Cationic liposomes were made using a mixture of dimethyldioctadecyl-ammonium bromide and cholesterol in chloroform, as described previously (Holinstat et al., 2006; Knezevic et al., 2009). Vector or WT-TRPC6-cDNA (50 µg) were mixed with 100 µl of liposomes. Liposomes encapsulating cDNA were injected into mouse vasculature via retroorbital injection. After 48 h, mouse lungs were used for determining lung vascular permeability or protein expression (Tauseef et al., 2008). Immunohistochemistry. Formalin-fixed, 4-µm-thick lung sections were co-immunostained with anti-TRPC6 and VE-cadherin antibodies along with appropriate Alexa Fluor–labeled secondary antibodies using manufacture protocol (Bethyl Laboratories, Inc.). Lung sections were visualized with a LSM510 confocal microscope (Carl Zeiss, Inc.; Tauseef et al., 2008). Assessment of lung vascular permeability to albumin. Evans blue–conjugated albumin (EBA; 20 mg/kg) was injected retroorbitally 30 min before sacrificing the mice to determine vascular permeability, as described previously (Peng et al., 2004; Tauseef et al., 2008; Knezevic et al., 2009). In brief, blood was obtained from the right ventricle of the heart into heparinized syringes and plasma was separated. Lungs were homogenized. Lung lysates and plasma were incubated with 2 volumes of formamide for 18 h at 55–60°C and centrifuged at 5,000 g for 30 min. The optical density of the supernatant was determined spectrophotometrically at 620 nm (Evans blue) and 740 nm (hemoglobin correction). EBA extravasation was calculated as EBA influx in lung versus that in plasma. Measurement of vessel filtration coefficient (Kf,c ). Mice were anesthetized with an i.p. injection of ketamine (100 mg/kg body weight) and xylazine (15 mg/kg body weight). Lungs were harvested and microvessel permeability was determined in isogravimetric lungs by determining microvascular filtration coefficient (K f,c; Vogel et al., 2000; Tauseef et al., 2008; Knezevic et al., 2009). In brief, outflow pressure was elevated by 10 cm H2O for 20 min in isogravimetric perfused lungs. The lung wet weight gain during this time, which is the net fluid accumulation, was recorded. At the end of each experiment, lung dry weight was determined. Kf,c (milliliters × min−1 × centimeters H2O × grams dry weight−1) was calculated from the slope of the recorded weight change normalized to the pressure change and lung dry weight. Lung edema determination. Left lungs from the same mice used for Evans blue albumin extravasation were excised and completely dried in the oven at 60°C overnight for calculation of lung wet/dry ratio (Barnard et al., 1995; Tauseef et al., 2008). BM transplantation (BMT). WT and Trpc6−/− mice were exposed to lethal irradiations with 1000 cGy at a dose rate of 100 cGy/m (Zhao et al., 2006; Weissmann et al., 2012). At 3 h after irradiation, mice were anesthetized with ketamine (100 mg/kg, i.p.) and xylazine (15 mg/kg, i.p.) and injected retroorbitally with 0.2 ml of 3 × 106 donor BM cells (TRPC6-null or WT) using a 27-gauge needle. Experiments were performed 6 wk after confirmation of BM cells transplantation by FACS analysis (see following section; Weissmann et al., 2012; Zhao et al., 2006). FACS analysis. MLECs were characterized using APC-tagged anti-VE cadherin and PE-tagged anti-CD31 antibodies. Isotype-matched APC- and PE-tagged IgG were used as negative controls. Data were acquired using Cyan-II flow Cytometry (Beckman Coulter, Inc.) and analyzed using Summit software. For determining the efficiency of BM cells transplantation in mice, BM cells were isolated from chimera mice. Cells were fixed and permeabilized with BD Cytofix/Cytoperm solution. These cells were incubated with anti-TRPC6 antibody for 30 min, followed by Alexa Fluor 488 secondary antibody. Size scatter analysis was performed using FACS analyzer to confirm the percentage of the TRPC6-expressing population (Kini et al., 2010). The analysis was done using WinMdi software. Dot blots were plotted after gating all BM cells from respective groups of mice against side scatter population. Statistical analysis. Comparisons between experimental groups were made by one-way ANOVA and post-hoc test. Differences in mean values were considered significant at P < 0.05.
To review the involvement of coagulation and fibrinolysis in the pathogenesis of acute lung injury (ALI)/acute respiratory distress syndrome (ARDS), pulmonary infection, and ventilator-induced lung injury (VILI). Published articles on experimental and clinical studies of coagulation and fibrinolysis in ALI/ARDS, pneumonia, and mechanical ventilation. Alveolar fibrin deposition is an important feature of ALI/ARDS and pulmonary infection. The mechanisms that contribute to disturbed alveolar fibrin turnover are localized tissue factor-mediated thrombin generation and depression of bronchoalveolar urokinase plasminogen activator-mediated fibrinolysis, caused by the increase of plasminogen activator inhibitors. These effects on pulmonary coagulation and fibrinolysis are regulated by various proinflammatory cytokines and are similar to those found in the intravascular spaces during severe systemic inflammation. Some studies also suggest that pulmonary coagulopathy is a feature of VILI. Recent studies have demonstrated the beneficial effect of anticoagulant therapy in sepsis. Theoretical considerations suggest that this anticoagulant therapy will benefit patients with primary lung pathology including VILI, but clinical studies are needed to examine this hypothesis before such therapy is to be advocated as a standard of care in critically ill patients.
Recognition of LPS by TLR4 on immune sentinel cells such as macrophages is thought to be key to the recruitment of neutrophils to sites of infection with Gram-negative bacteria. To explore whether endothelial TLR4 plays a role in this process, we engineered and imaged mice that expressed TLR4 exclusively on endothelium (known herein as EndotheliumTLR4 mice). Local administration of LPS into tissue induced comparable neutrophil recruitment in EndotheliumTLR4 and wild-type mice. Following systemic LPS or intraperitoneal E. coli administration, most neutrophils were sequestered in the lungs of wild-type mice and did not accumulate at primary sites of infection. In contrast, EndotheliumTLR4 mice showed reduced pulmonary capillary neutrophil sequestration over the first 24 hours; as a result, they mobilized neutrophils to primary sites of infection, cleared bacteria, and resisted a dose of E. coli that killed 50% of wild-type mice in the first 48 hours. In fact, the only defect we detected in EndotheliumTLR4 mice was a failure to accumulate neutrophils in the lungs following intratracheal administration of LPS; this response required TLR4 on bone marrow-derived immune cells. Therefore, endothelial TLR4 functions as the primary intravascular sentinel system for detection of bacteria, whereas bone marrow-derived immune cells are critical for pathogen detection at barrier sites. Nonendothelial TLR4 contributes to failure to accumulate neutrophils at primary infection sites in a disseminated systemic infection.
[∗
]Department of Pulmonary Medicine, Zhongshan Hospital, Fudan University, Shanghai,
China
[†
]Department of Cardiothoracic Surgery, Huashan Hospital, Fudan University, Shanghai,
China
[‡
]Department of Respiratory and Critical Care Medicine, Second Affiliated Hospital,
Institute of Respiratory Diseases, Zhejiang University School of Medicine, Hangzhou,
China
[§
]Shanghai Respiratory Research Institution, Shanghai, China
[||
]State Key Laboratory of Respiratory Disease, Guangzhou Medical University, Guangzhou,
China.
[¶
]Department of Nephrology, Zhongshan Hospital, Fudan University, Shanghai, China
[∗∗
]Center for Biomedical Imaging, Fudan University, Shanghai, China
[††
]Department of Pulmonary Medicine, the Central Hospital of Xuhui District, Shanghai,
China
Author notes
Address reprint requests to Chun X. Bai, MD, PhD, FCCP, Department of Pulmonary Medicine,
Zhongshan Hospital, Fudan University, Shanghai, China. E-mail:
bai.chunxue@
123456zs-hospital.sh.cn
; Jian Zhou, PhD, Department of Pulmonary Medicine, Zhongshan Hospital, Fudan University.
E-mail:
zhou.jian@
123456fudan.edu.cn
; Yuan L. Song, MD, PhD, Department of Pulmonary Medicine, Zhongshan Hospital, Fudan
University. E-mail:
ylsong70@
123456163.com
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