Background
Idiopathic pulmonary fibrosis (IPF) is a chronic progressive interstitial lung disease (ILD) with few effective drug therapies [
1,
2]. An early event in IPF is recurrent epithelial injury resulting in activation of extracellular matrix (ECM)-producing fibroblasts [
3]. CXCR4 is a transmembrane protein receptor that is constitutively expressed on epithelial and myeloid cells and is involved in stem cell migration [
4]. CXCR4 is activated by the chemokine CXCL12, also known as stromal derived factor (SDF)-1 and in the bone marrow, CXCL12 is constitutively produced in order to maintain hemopoietic stem cells in the bone marrow environment [
5].
In the lung, CXCL12 is upregulated upon epithelial injury [
6]. Other CXCR4 expressing cells include fibrocytes which are bone marrow-derived, circulating progenitor cells that express the marker CD45, traffic to the lung via CXCR4/CXCL12 signalling and have been implicated in fibrotic ILD [
7]. However, an analysis of the specific cell types expressing CXCR4 in IPF has not been attempted. As a result, there is still some uncertainty as to the origin of CXCR4 expressing cells in the diseased lung. In particular, whether CXCR4
+ cells are derived from lung resident cells perhaps of epithelial origin or do they originate from the peripheral circulation remains elusive.
In a previous study, the i-body CXCR4 antagonist, AD-114, reduced experimental lung fibrosis and decreased IPF lung fibroblast invasion
in-vitro [
8], which supports the targeting of CXCR4 as a therapeutic option in IPF. CXCR4 is best known for its role in diverse disorders such as HIV-1 infection [
9], tumour development [
10], and acute myeloid leukemia [
11]. In the lung, CXCR4 expression is thought to contribute to the cellular trafficking from the vessels into the interstitium and contribute to a number of fibrotic disorders including asthma and pulmonary fibrosis [
12]. However, it is not known if circulating cells expressing only CXCR4 could be potential biomarkers of disease progression in IPF. Therefore, the first aim of this study was to investigate whether numbers of CXCR4
+ cells in the circulation of patients with IPF were elevated compared to age-matched normal controls and if there was any correlation with clinical parameters of disease.
Radiologically, honeycomb lung is defined by clusters of sub-pleural basal cystic air spaces seen on high-resolution computed tomography (HRCT) scans in patients with end-stage fibrotic ILD, and in particular those with the usual interstitial pneumonia (UIP) pattern of fibrosis. Patients with ILDs that share a radiologic and histological pattern of UIP do worse than those that have other patterns such as non-specific interstitial pneumonia (NSIP) [
13]. Histologically, honeycomb cysts are lined with cuboidal or ciliated cells that express a variety of epithelial markers such as e-cadherin and basal cell-specific keratins, supporting the hypothesis that honeycomb cysts are derived from the distal airway [
14] but further investigation into the cells of the honeycomb cyst is necessary before anti-CXCR4 therapies in IPF can be fully realised.
We have previously published that CXCR4 expression is increased in IPF lung tissue, particularly in airway and honeycomb-associated epithelial cells [
8]. However, it is not known whether these cells also express CXCL12, which would imply that an autocrine CXCR4-CXCL12 pathway could be continuously activated in IPF. Therefore, we then aimed to investigate the CXCR4/CXCL12 expression in the lung tissue of patients with IPF, non-IPF fibrotic ILD and NDC.
Methods
Patients/healthy donor/s
Patient demographics are described in Table
1 and Table
2.
Table 1
Patient demographics of patients with idiopathic pulmonary fibrosis (IPF) in the blood study
Median Age, years (SD) | 67 (11) |
Smoking History, n (%) | 14 (70%) |
Median Forced Vital Capacity (FVC), % predicted (SD) | 68 (16) |
Median Transfer Factor of Carbon Monoxide (TLCO), % predicted (SD) NDC | 43 (14) |
Table 2
Patient demographics of the donors in the tissue study
Median Age, years (SD) | 60 (16) | 61 (4) | 60 (6) |
Smoking History, n (%) | 4 (40%) | 7 (70%) | 4 (50%) |
Median Forced Vital Capacity (FVC), % predicted (SD) | n/a | 43 (16) | 48 (27) |
Median Transfer Factor of Carbon Monoxide (TLCO), % predicted (SD | n/a | 17 (8) | 43 (13) |
All patients had been diagnosed at the Alfred Hospital’s lung fibrosis multi-disciplinary team (MDT) meeting according to recommended guidelines [
15].
Blood study population
A total of 20 patients with IPF and 10 NDC volunteers (aged > 50 years) were included. Of the IPF patients, 13 were on antifibrotic medication (Pirfenidone, n = 8; Nintedanib, n = 5). Age-matched NDC donors were recruited from the Australian Red Cross Blood Donation Centre. Plasma samples from IPF and NDC donors was prepared by fractionation of whole blood.
Tissue study population
Explant lung tissue was obtained from patients with IPF (n = 10 patients) and other interstitial lung diseases (ILDs)(n = 8) at the time of lung transplantation. All patients had pathologist-verified histopathology reports of Usual Interstitial Pneumonia (UIP). None of the IPF patients undergoing lung transplantation were on antifibrotic medication. Non-IPF ILD diagnoses included non-specific interstitial pneumonia (NSIP, n = 2), chronic hypersensitivity pneumonitis (HP, n = 4), connective tissue disease associated ILD (CTD-ILD, n = 2). Tissue from non-disease controls (NDC) were obtained from deceased organ donors whose lungs had been declined for transplantation.
Because IPF typically manifests first in the lung bases and the fibrosis progresses upwards towards the lung apices, we examined tissue from multiple regions of the lungs to take in account this inherent heterogeneity. All lung tissue was obtained from the Alfred Lung Fibrosis Biobank [supported by the National Health and Medical Research Council (NHMRC) Centre for Research Excellence in Pulmonary Fibrosis].
Flow cytometric analysis
Polymorphonuclear cells were fractioned from whole blood of patients with IPF and NDC donors and frozen for later use. For cell characterization, cells were stained with an antibody cocktail containing antibodies specific for human CXCR4 (R&D systems, USA), CD4 (eBioscience, USA), CD8 (Pharmingen, USA), B-cell marker CD19 (eBioscience, USA), early myeloid marker CD33 (BD Biosciences, USA) and LIVE/DEAD™ Fixable Aqua Dead Cell Stain (ThermoFisher, USA) and analysed by flow cytometry. Only samples containing more than 5000 live cells were included for analysis.
Enzyme-linked Immunosorbent assay (ELISA)
CXCL12 in plasma was measured by ELISA according to manufacturer’s instructions (IBT Systems, Germany). Briefly, plasma samples were recovered from liquid nitrogen, thawed at room temperature and used undiluted in the assay. Antibody working solutions and CXCL12 standards were prepared on the day of the experiment in kit assay buffer. Test solutions (plasma, CXCL12 standard, or assay buffer only, 50 μl), in duplicate, were added to 100 μl of capture antibody (biotinylated anti-HuSDF1α antibody) + detection antibody (anti-HuSDF1α monoclonal antibody) and incubated for 1 h at RT. Following washing, 100 μl of conjugate working solution (goat anti-mouse IgG peroxidase conjugate) was added to each well. The plate was incubated for a further 1 h at room temperature, washed then developed using TMB-ELISA Substrate Solution (100 μl/well; supplied) for 30 min. The reaction was stopped with 2 M H2SO4 (50 μl/well; supplied) and the plate was read using a SpectraMax plate reader at 450 nm.
Immunohistochemistry (IHC)
From formalin-fixed paraffin-embedded tissue blocks, 4 μm thick paraffin sections were immunohistochemically stained for CXCR4 (dilution 1:500, clone UMB2, Cat# ab124824, Abcam, UK), CXCL12 (dilution, 1:500, clone EPR1216, Cat# ab155090, Abcam, UK), E-Cadherin (epithelial marker, dilution 1:1000, clone EP700Y, Cat# ab40772, Abcam, UK) and CD45 (myeloid marker, dilution 1:1000, clone MEM-28, Cat# ab8216, Abcam, UK). Antigen retrieval was performed in pH 6 citrate buffer (Sigma, Australia). After incubation with DakoEnVision secondary reagents (Dako, Australia), positive staining was visualized for brightfield using diaminobenzidine (Dako, Australia). Sections were counterstained in Mayer’s hemotoxylin (Sigma, Australia). All sections for brightfield analysis were stained at the same time. Sections were scanned using an Aperio Scanscope AT Turbo (Leica Biosystems, Australia) and images were captured at a resolution of 0.25 μm/pixel. The extent of overall CXCR4 expression was semi-quantified by rating the intensity and presence of CXCR4 staining across the entire section on a scale of 0 (absent), 1 (low), or 2 (medium – high) by a researcher blinded to the diagnosis of the section.
All sections for multiplex staining in both panels were stained at the same time. For multiplex staining, tyramide signal amplification was performed using Perkin-Elmer’s OPAL dye reagents (diluted 1:100 and incubated for 6 min) in 3 spectrally distinct fluorophores that have non-overlapping fluorescent spectra, allowing the individual antigens to be separated from a single image (Additional Figure
1).
Whole sections were scanned at 4× magnification using the Vectra 3 Quantitative Pathology Imaging System and the entire section was encircled to define the area to be analysed. Regions of interest (ROIs) were then automatically generated that encapsulated the area of the tissue section for subsequent analysis at 20× magnification (with a resolution 0.5 μm) using Phenochart (v 1.0, Perkin Elmer) software (Additional Figure
2). Exposure settings and emission/excitation wavelengths for each of the channels and OPAL dyes are listed in Additional Figure
2.
Sections were sequentially stained with primary antibodies in two panels and cell nuclei was stained with Hoechst in both. Panel 1 consisted of CXCR4, E-Cadherin and CXCL12. Panel 2 consisted of CXCR4, CD45 and CXCL12. The sequence of targets and dye in Panel 1 was CXCR4 (OPAL 520), e-cadherin (OPAL 570), and CXCL12 (OPAL 690). The sequence of targets and dye in Panel 2 was CXCR4 (OPAL 520), CD45 (OPAL 570) and CXCL12 (OPAL 690). Hoechst dye was applied as the last step in both panels.
Multiplex analysis was performed on tissue sections from 7 NDC donors and 7 patients with IPF using HALO imaging processing software (Indica Labs Highplex FL version 3.0) on a total of 2867 images (ROIs) (Table
3). There were 1458 ROIs captured in the e-cadherin panel and 1409 ROIs in the CD45 panel. Additional File
3 details the thresholds used for positive cell quantification. Cells in lung tissue were identified using Hoechst staining and thresholds for positivity of the panel markers were manually set by examining 2 randomly chosen ROIs from a NDC donor and IPF patient (total of 4 images). Additional File
3 shows two ROIs quantified for Panel 1 and Additional File
4 shows two ROIs quantified for Panel 2. Thresholds and corresponding cell phenotype combinations are then applied to all ROIs used in the analysis. Quantification is performed on all the ROIs from each tissue section and data is presented as percentage of Hoechst+ cells counted.
Table 3
Summary of quality control parameters for multiplex immunohistochemsitry analysis
Variable | Mean1 | SD | Mean2 | SD | p-value | |
Number of ROIs in analysis | 116 | 31.5 | 92 | 37.9 | 0.259 | |
Total number cell-sized objects | 193,341 | 43,830 | 156,428 | 59,257 | 0.209 | |
Hoechst 33258 Positive Cells | 58,854 | 28,534 | 48,439 | 34,558 | 0.209 | |
Avg Cell Area (μm2) | 126.4 | 8.0 | 125.1 | 4.8 | 0.535 | |
Avg Cytoplasm Area (μm2) | 73.0 | 11.5 | 75.2 | 7.6 | 0.902 | |
Avg Nucleus Area (μm2) | 53.4 | 5.0 | 49.9 | 5.1 | 0.259 | |
Avg Nucleus Perimeter (μm) | 34.4 | 2.1 | 33.7 | 1.8 | 0.456 | |
Avg Nucleus Roundness | 0.7 | 0.005 | 0.7 | 0.007 | 0.001** | |
Area Analyzed (μm2) | 5E+ 07 | 2E+ 07 | 4E+ 07 | 2E+ 07 | 0.383 | −2.073 |
CD45 Panel |
| Normal (N = 7) | IPF (N = 7) | Mann-Whitney U (Nonparametric) | Effect Size |
Variable | Mean1 | SD | Mean2 | SD | p-value | |
Number of ROIs in analysis | 111 | 32.4 | 91 | 33.5 | 0.318 | |
Total number cell-sized objects | 186,596 | 52,479 | 153,307 | 53,860 | 0.318 | |
Hoechst 33258 Positive Cells | 52,770 | 31,528 | 41,252 | 31,069 | 0.456 | |
Avg Cell Area (μm2) | 125.0 | 6.9 | 123.6 | 5.9 | 0.902 | |
Avg Cytoplasm Area (μm2) | 71.9 | 10.4 | 73.0 | 8.8 | 1.000 | |
Avg Nucleus Area (μm2) | 53.0 | 4.8 | 50.6 | 4.3 | 0.383 | |
Avg Nucleus Perimeter (μm) | 34.6 | 2.1 | 34.1 | 1.8 | 0.805 | |
Avg Nucleus Roundness | 0.7 | 0.005 | 0.7 | 0.006 | 0.001** | −2.309 |
Area Analyzed (μm2) | 5E+ 07 | 2E+ 07 | 5E+ 07 | 2E+ 07 | 0.646 | |
Multiplex quality control variables for each of the panels are detailed in Table
3. There were no differences in cell area, cytoplasm area, nucleus area, nucleus perimeter or total area analysed between NDC and IPF tissues. Average nucleus roundness was smaller statistically (less round) in IPF compared to NDC tissues but this parameter does not affect the number of counted cells and is likely a reflection of minor cell compression in fibrotic tissue [
16].
Statistics
Prism Software Version 7 (GraphPad Software Inc., USA) to compare IPF and NDC data. For flow cytometric analysis, between 2-group comparisons were performed using unpaired t-test with Welch’s correction. For tissue brightfield histological analysis between 3-group comparisons was made using one-way ANOVA with Dunnett’s multiple comparisons test. For tissue multiplex histological analysis between 2-group comparisons was made using Mann-Whitney U-test.
Discussion
The precise pathophysiological pathways that drive idiopathic pulmonary fibrosis (IPF) are yet to be fully understood. Earlier work suggests that the CXCR4 pathway may be a therapeutic target [
17,
18]. We have identified two distinct populations of CXCR4
+ cell, one of epithelial origin (CXCR4
+/e-cadherin
+) and one of myeloid origin (CXCR4
+/CD45
+), that were increased in lung tissue of patients with IPF compared to non-diseased control (NDC) donors. Furthermore, we now provide more evidence that dysregulation of CXCR4 expression is not unique to IPF and therefore our findings are relevant to patients with fibrotic ILDs as well.
Fibrocytes are thought to be one source of fibroblasts in IPF by differentiating into matrix-producing myofibroblasts [
19] after trafficking into the lung from the circulation. One study reported that most circulating fibrocytes express CXCR4 [
20] and importantly, fibrocytes are not found in the tissue of normal lungs [
21], therefore, lower levels of CXCR4
+ cells in IPF patient circulation compared to NDC donors was unexpected and out of line with the literature [
22]. It was not previously known if monitoring circulating cells by using only CXCR4
+ as an identifier could be a surrogate marker of active fibrosis. Our study suggests that there exists a normal circulating level of CXCR4
+ cells in healthy individuals which may be reduced as a result of active fibrosis in the lung tissue. These cells may be migrating from the circulation to the interstitial tissue through a chemical gradient produced by CXCL12-expressing cells, such as fibroblasts, or via activation in the circulation through elevated plasma CXCL12 levels.
Plasma CXCL12 concentration is elevated in chronic hypersensitivity pneumonitis, another related fibrotic ILD, and may be epithelial cell-derived [
23]. The previous studies that have observed increased numbers of CXCR4
+ cells in fibrotic lung tissue have primarily focused on the fibrocyte subtype which express haemopoietic markers like CD45 and Collagen 1 in addition to CXCR4 [
19‐
21,
24]. Relatively few studies have examined the contribution of CXCR4
+ epithelial cells to pulmonary fibrosis [
4,
25,
26]. During wound healing events, CXCR4 expression on alveolar type II (ATII) cells is important to drive epithelial cell migration and proliferation [
4,
25,
27]. Both CXCR4 and CXCL12 expression is induced in ATII cells following lung injury [
28] and both genes are regulated by a hypoxic microenvironment [
29] such as in fibrotic tissue. Our study suggests that in the fibrotic lung, chronic injury and hypoxia drives constant upregulation of CXCR4 and CXCL12 in resident epithelial cells at the epithelial-alveolar interface, leading to overproliferation and migration of neighbouring ATII cells and driving alterations in the tissue structure.
The initiating event that leads to the formation of usual interstitial pneumonia (UIP) remains unknown but hyperplastic ATII cells are often seen at areas of active fibrosis in patients with fibrotic ILD [
30] and there is some evidence to suggest that microscopic honeycomb cyst formation has a distal airway origin driven by dysregulation of the mucin gene MUC5B [
14]. Recently, Chen and colleagues demonstrated the therapeutic potential of targeting honeycomb cyst formation via MUC5B inhibition in IPF [
31]. Similarly, our study suggests that targeting epithelial CXCR4
+ cells may prevent honeycomb cyst formation in IPF and other fibrotic ILDs that share the UIP pattern. It was outside the scope of this study to determine if there were differences in CXCR4 expression between IPF and the other fibrotic ILDs but it can be concluded that the UIP phenotype includes alveolar destruction involving epithelial CXCR4
+ cells. Furthermore, these epithelial CXCR4
+ cells, unlike their lymphoid counterparts, the CXCR4
+CD45
+ cells, also expressed CXCL12, indicating that the epithelium may be the primary site of fibrotic activity driven by mechanisms involving the CXCR4/CXCL12 axis in IPF. Future studies in a larger cohort are needed to determine the precise cell type and investigate the role of CXCR4
+ cells that do not express either e-cadherin or CD45.
The limitation of our study was that the donors from whom PMBCs were obtained were not the same donors who had provided normal plasma. Furthermore, the IPF patients who provided whole blood were not the same patients from whom lung tissue was investigated. Therefore, it is difficult to determine how the quantity of CXCR4+ PBMCs relates to CXCL12 plasma levels in normal donors or to numbers of CXCR4+ cells in IPF lung tissue. Our study did not show any differences in CXCR4+/CD19+ (B-cell origin) or CXCR4+/CD33+ (early myeloid origin) cells, and therefore further investigation with a larger panel of surface markers may be necessary to delineate whether other circulating CXCR4+ cells are clinically relevant in IPF.
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