Background
Idiopathic pulmonary fibrosis (IPF) is a chronic, progressive, irreversible and ultimately lethal lung disease of unknown cause and unclear pathogenic mechanisms, characterized by myofibroblast accumulation and lung scarring [
1,
2]. At present, there are no reliable clinical parameters or non-invasive biomarkers predicting the clinical course of IPF [
3]. A growing body of evidence indicates that the disease can result from the abnormal behaviour of the alveolar epithelial cells, which provokes the migration, proliferation and activation of mesenchymal cells. This results in the formation of fibroblast and myofibroblast foci secreting exaggerated amounts of extracellular matrix molecules, with the subsequent destruction of the lung architecture [
4]. It has been hypothesized that an extra-pulmonary source of fibroblast/myofibroblasts exists, which likely has a bone marrow origin and can be detected in the blood [
5].
In 1994, using state-of-the-art techniques, circulating fibrocytes were identified as cells that exit the blood stream, migrate into wounds and contribute to wound repair [
6]. Fibrocytes are spindle-shaped, bone marrow-derived mesenchymal progenitor cells that co-express a variety of cell surface markers related to leukocytes, hematopoietic progenitor cells and fibroblasts. They express a variety of mesenchymal markers, including collagen I, as well as the common leukocyte marker CD45 and the hematopoietic stem cell marker CD34. They do not express T cell markers (CD3, CD4 and CD8), B cell markers (CD19) or myeloid markers (CD14) [
7]. It has been shown that, in healthy donors, they can represent up to 1 % of circulating nucleated cells [
8‐
11] and can express chemokine receptors such as CXCR4 and CCR7; they have been found in a variety of tissues under both physiological and pathological states [
9,
12]. However, scanty data exist on the fine characterization of these circulating cells, whose relative rarity in blood obviously represents an obstacle to their precise analysis.
The biological axis CXCL12/CXCR4 could be involved in mediating the contribution of fibrocytes to pulmonary fibrosis [
10]. Indeed, the high expression of CXCL12 in lung injury creates a chemokine gradient for CXCR4+ fibrocytes, which can be released from the bone marrow and recruited to the lungs [
13]. Once they extravasate and enter the target tissue, fibrocytes can differentiate into fibroblasts and myofibroblasts [
14]. So, it has been supposed that circulating fibrocytes might contribute to the intense remodelling of the pulmonary vasculature in patients with IPF, or at least represent a biomarker of disease activity [
15].
Multiple mechanisms play a role in IPF pathogenesis, including abnormal vascular repair and remodelling [
16]. During IPF, fibrogenesis is strongly associated with abnormal vascular remodelling [
17]. Indeed, there is a body of evidence suggesting that the impairment of re-endothelization mechanisms after alveolar injury may lead to the destruction of lung architecture, and consequently trigger fibrosis [
18]. Failure of re-endothelization may induce loss of the alveolar-capillary integrity, which might be the point after which fibrosis becomes inevitable [
16]. Fibrotic areas have few blood vessels, whereas adjacent non-fibrotic tissue is highly vascularized [
19]. There are almost no capillaries within the fibroblastic foci, indicating that the fibrotic process in IPF does not need neovascularization [
20]. In this regard, it has been suggested that the respective abundance of circulating endothelial cells (CEC) and endothelial progenitor cells (EPC) might reflect the balance between vascular injury/repair and potentially serve as biomarkers of the disease [
17]. Few data on CEC or EPC exist from patients with IPF.
With the aim of clarifying whether CEC and their precursors and circulating fibrocytes are altered in IPF, and to understand whether these cells may be used as biomarkers, we studied such cells in a cohort of Italian patients with IPF, some of whom were longitudinally followed. We used an innovative methodological approach, based upon sophisticated techniques that employ acoustic, multiparametric flow cytometry that allows a precise and fine analysis of these rare cell types.
Methods
Patients
All incident and prevalent patients with IPF from six Italian centres (Modena, Reggio Emilia, Bologna, Siena, Napoli and Catania) were deemed eligible for this study. All patients fulfilled 2011 American Thoracic Society/European Respiratory Society/Japanese Respiratory Society/Latin American Thoracic Association guideline diagnostic criteria [
21]. Complete medical history and lung function tests were acquired at enrolment. Six-month follow-up visits and lung function tests were scheduled for up to 2 years. Blood samples for the analysis of circulating fibrocytes and endothelial cells were obtained at enrolment and during follow-up visits.
The study has been approved by the Local Ethical Committee (Modena, Number of practice 31/12), and written informed consent was obtained from each patient.
Among the patients with IPF, 18 were treated with pirfenidone, 13 with nintedanib, and 26 were untreated. Patient characteristics are reported in Table
1.
Table 1
Patients’ characteristics
Gender | | | | |
Male | 53 | | | |
Female | 14 | | | |
Age (years) | | | 74 | 68.5–77.0 |
Time from diagnosis (years) | | | 3 | 2.0–4.5 |
Smoking history | | | | |
Non smoker | 19 | | | |
Smoker or former smoker | 41 | | | |
Forced vital capacity (% predicted) | | 75.0 | | 56.75–93.0 |
DLCO (% predicted) | | 41.0 | | 34.0–60.0 |
GAP stage (%) | | | | |
I | | 32.70 | | |
II | | 53.10 | | |
III | | 14.30 | | |
Treatment | | | | |
Pirfenidone | 18 | | | |
Nintedanib | 13 | | | |
Untreated | 26 | | | |
Blood collection and cell analysis
Thirty millilitres of blood were collected through a venous drawing in EDTA tubes. The first 3 mL of blood from the venipuncture were not used for cell analysis, because of the contaminating presence of endothelial cells derived from the vessel wall. Buffy coat was then prepared according to standard procedures, and cells were stained with different monoclonal antibodies (mAbs) for the detection of CEC, EPC and circulating fibrocytes. For the detection of CEC and EPC a minimum of 10 millions cells were stained with anti-CD45 PE (eBioscience, San Diego, CA, USA), anti-CD34 PC7 (Beckman Coulter, Hieleah, FL, USA), anti-CD133 APC (Miltenyi GmbH, Bergisch Gladbach, Germany), anti-CD14 APC-VIO770 (Miltenyi), anti-CD309 FITC (R&D Systems, Minneapolis, MN, USA) and viability probe Far-Red LIVE/DEAD.
For the detection of circulating fibrocytes a minimum of 20 million cells were stained with Red Fixable LIVE/DEAD probe (Thermo Fisher,Eugene, OR, USA ) and the following surface mAbs: anti-CD3 PE-CY 5.5 (Becton Dickinson, San José, CA, USA), anti-CD19 PE-CY 5.5 (Becton Dickinson), anti-CD45 PE (eBioscience), anti-CD34 PC7 (Beckman Coulter), anti-CD14 APC-VIO770 (Miltenyi) and anti-CXCR4 APC (Becton Dickinson). Cells were fixed and permeabilized using Cytofix/Cytoperm buffer set (Becton Dickinson) and stained with directly conjugated mAb anti-collagen I FITC (Merck Millipore, Billerica, MA, USA). Tables
2 and
3 report the mAbs used and the relative fluorochromes.
Table 2
Table summarizing the excitation sources and fluorescence emissions used for the detection of circulating endothelial cells and their precursors
Anti-CD34 | PC7 | 488/750 |
Anti-CD45 | PE | 488/530 |
Anti-CD133 | APC | 637/660 |
Anti-CD309 | FITC | 488/519 |
Anti-CD14 | APCVio770 | 637/785 |
LIVE/DEAD | Far Red | 637/>665 |
Table 3
Table summarizing the excitation sources and fluorescence emission used for the detection of circulating fibrocytes
Anti-CD34 | PC7 | 488/750 |
Anti-CD45 | PE | 488/530 |
Anti-CXCR4 | APC | 637/660 |
Anti-collagen I | FITC | 488/519 |
Anti-CD19 | PE-Cy5.5 | 488/690 |
Anti-CD3 | PE-Cy5.5 | 488/690 |
Anti-CD14 | APCVio770 | 637/785 |
LIVE/DEAD | Red Fixable | 488/615 |
Acquisition of samples
For phenotype analysis, cells were acquired using a 14-colour 4-laser high-speed Attune NxT flow cytometer (Thermo Fisher). Single staining and fluorescence minus one (FMO) controls were performed for all panels to set proper compensation and define positive signals [
22]. In order to identify rare cells like human peripheral CEC, EPC or circulating fibrocytes, it was mandatory to acquire a huge number of cells [
23], that is, of the order of several million per sample. Thus, for the phenotypic analysis, we used a novel acoustic flow cytometer able to align cells in the flow chamber using ultrasound, acquiring up to 35,000 cells per second. This was crucial to obtain the number of cells required for a correct statistical analysis, which was typically >10 million. Starting from a buffy coat, we were therefore able to clearly identify CEC, EPC or fibrocytes among peripheral blood cells.
Statistical analyses
Data were analysed by FlowJo 9.8.5 and GraphPad 6.0 software, using the Wilcoxon T test and non-parametric analysis of variance test (Kruskal–Wallis test).
Conclusions
This multicentric study is the first to provide cross-sectional and longitudinal analyses of CEC and fibrocytes amongst Italian patients with IPF. Our study was performed on blood samples—we could not analyse lung tissue from patients with IPF. Indeed, the most critical obstacle to translating information obtained from molecular or cellular in vitro or ex vivo studies into clinical applications is the scarcity of lung tissue, especially in the context of a rare disease. Although some patients undergo biopsy, in most cases either lung biopsy is not indicated, or the risk associated with the procedure precludes it from being performed. Given the fact that fibrocytes might be correlated with endothelial cells during the remodelling process of fibrotic tissue, and given that drugs used in IPF may modulate the function of CEC, the aim of this study was to understand whether more accessible cells like circulating fibrocytes and endothelial cells may be used as surrogate biomarkers of disease outcome in patients with IPF treated with different drugs.
First, we investigated the phenotype of CEC and EPC and found a significant decrease in the expression of CD309 among endothelial cell populations. Thus, it is likely that the identification of such a subpopulation could be of clinical relevance. Second, we investigated the percentage of circulating collagen I+ cells, defined as fibrocytes, in patients with IPF treated with different therapies, and we found that there was no difference compared with healthy controls. The change in the expression of CXCR4 in such cells after 6 months of therapy could be indicative of a therapeutic effect, in terms of diminished homing to the lung. However, because of the relatively small number of patients we could analyse, further data are needed to clarify this aspect.
This study had some other limitations. First, we were not able to follow up the entire IPF cohort. We also could not clarify the molecular mechanism(s) by which circulating cells expressing collagen I and endothelial cells cooperate to form fibrotic foci. However, it could be hypothesized that CEC sustain the vascularization around the fibrotic foci, and thus play a pathogenic role. In conclusion, although further studies are needed to confirm that CEC and fibrocytes may be used as surrogate biomarkers of disease presence, severity, rate of progression and treatment outcome, the change in CD309 expression in endothelial cells suggests that such receptors could likely become a new target for therapies against IPF.
Competing interests
All authors declare that they have no competing interests.
Authors’ contributions
AC and LR initiated the collaborative project. AC, LR and SC planned the project. SDB, EB, EP and LG collected the data. SC, GM, FL, CMC, MB, CV, GS, LZ and AZ recruited and followed patients and provided the clinical expertise. SDB, EB and LG were responsible for data handling and data analyses. SDB, LR and AC wrote the first draft, and all authors contributed and approved the final version of the manuscript for publication.