Abstract
Introduction
Resistive breathing (RB) is the hallmark of diseases of airway obstruction, especially during exacerbations, resulting in significant mechanical stress on the lung. RB induces pulmonary inflammation and injury in previously healthy animals. Whether RB exerts additional injurious effects when superimposed on cigarette smoke (CS) exposure is unknown.
Methods
Adult mice were exposed to CS for 1 or 6 months, followed by RB for 24 h, induced through tracheal banding. Subsequently, respiratory system mechanics were assessed, bronchoalveolar lavage (BAL) was performed and cytokine levels were measured by ELISA in lung tissue samples. Surfactant protein D (SP-D) was measured in blood, and BAL and histology were performed. Emphysema was quantified by the mean linear intercept (Lm) and the destructive index (DI).
Results
CS exposure for 1 and 6 months increased BAL cellularity (∼8-fold and ∼1.9-fold to air, respectively, p<0.01). RB aggravated BAL cellularity at both time points (p=0.025 and p=0.002 to CS) and increased MCP-1 levels at 1 month (p=0.002 to control). Histology revealed augmented focal membrane thickening at 1 month following combination of RB with CS exposure (p=0.011), while at 6 months emphysema was more severe after RB in CS-exposed mice (p=0.001 to CS for both Lm and DI). CS increased Sp-D levels in BAL (p<0.01 at both time points), while RB caused increased levels in blood (p<0.01 to CS). RB decreased static compliance at 1 month of air exposure (p=0.007).
Conclusion
Combining resistive breathing with cigarette smoke exposure results in augmented inflammatory responses, increased lung injury and augmented emphysema.
Shareable abstract
Resistive breathing worsens pulmonary inflammation and emphysema caused by cigarette smoke in mice, suggesting that mechanical stress during severe airway obstruction may promote disease progression https://bit.ly/4jCjVlx
Introduction
COPD is a common chronic lung disease and one of the major causes of morbidity and mortality worldwide [1], characterised by persistent airflow limitation [2]. Although COPD has a multifactorial pathogenesis, including genetic factors, air pollution and early life events [3], cigarette smoke (CS) exposure remains the most important risk factor. Induction of inflammation and an abnormal response of the immune system in the small airways and alveoli due to CS exposure [4] results in thickening of the small airway wall and loss of alveolar attachments, with subsequent destruction of alveoli and enlargement of airspaces, the anatomical hallmarks of emphysema [5, 6]. Infiltration of lung periphery from neutrophils and macrophages is a central component of COPD pathogenesis [7, 8], a process modulated by chemokines secretion, including interleukin (IL)-8 and monocyte chemotactic protein 1 (MCP-1), for neutrophils and monocytes, respectively [9], and is reproduced in the mouse model of COPD through CS exposure both at the cellular and the cytokine level (keratinocyte-derived chemokine (KC), the mouse analogue of IL-8 and MCP-1) [10, 11].
The natural course of COPD is characterised by the presence of acute exacerbations, events that are associated with hospitalisations and increased mortality, impaired quality of life and an accelerated decline of lung function [12], whereas a prior history of exacerbation is the determining factor for future exacerbation risk. COPD exacerbations are associated with augmented inflammation of the airways, compared to stable disease [13] (including exaggerated neutrophil influx and pro-inflammatory cytokine expression [14]) and elevated circulating surfactant protein-D (Sp-D) levels [15], which are also increased in various models of epithelial lung injury in rodents [16]. During exacerbations, excessive airway narrowing due to inflammation and bronchoconstriction leads to severe airflow limitation and resistive breathing (RB) that together with marked air trapping and dynamic hyperinflation lead to significant mechanical stress on the lung [17, 18].
Increased mechanical stress imposed on the lung is known to induce pulmonary inflammation and injury through the induction of intracellular “mechanosensitive” signalling pathways [19]. Our research group has previously shown that spontaneous breathing through increased airway resistance, to mimic the mechanical consequences of severe airway obstruction, induces pulmonary inflammation and injury in previously healthy mice and rats, with upregulation of markers of innate immunity activation (e.g. neutrophil and macrophages influx) and increased expression of inflammatory cytokines [20–22]. Moreover, we have shown that RB aggravates an animal model of COPD, i.e. the model of elastase-induced emphysema [23]. The effect(s) of imposing RB on the already inflamed lungs of the mouse model of COPD (CS exposure), mimicking the clinical scenario of a stable COPD patient suffering an acute COPD exacerbation (i.e. increased RB), is not known.
We hypothesised that RB would exert a synergistic effect with CS exposure, exacerbating pulmonary inflammation and injury. To test our hypothesis, we employed the murine model of CS exposure and superimposed on it our model of RB through tracheal banding, to mimic the mechanical sequelae of bronchoconstriction during a COPD exacerbation.
Materials and methods
Animals
Adult male C57BL/6 mice (age 8–10 weeks) were purchased from the Biomedical Sciences Research Center “Alexander Fleming”. Handling of the laboratory animals was in accordance with the European Union Directive (2010/63/EU) on the protection of animals used for scientific purposes. Experimental procedures were approved by the Scientific Council and Ethics Committee of “Evangelismos” Hospital (217/4-10-2016).
CS-induced animal model of COPD
Cigarette smoke exposure was performed as previously described by our group [24] using a home-build smoking machine. Mice placed in a glass chamber received mainstream CS of five reference cigarettes (3R4F; University of Kentucky, containing 9.5 mg of tar per cigarette) four times a day with 30-min smoke-free intervals for 5 days per week. Each cigarette was burnt with 15 puffs (∼20 mL/puff) over 5 min, while the chamber was supplied with a 2 L·min−1 medical air flow, creating a smoke to air ratio of 1:6, as previously established in our laboratory [25]. The animals were exposed to CS for either 1 month (subacute exposure) or 6 months (chronic exposure). The control group were exposed to room air only.
RB through tracheal banding
Mice were subjected to RB by the method of tracheal banding, as previously described by our group [20]. After anaesthesia with an intraperitoneal injection of xylazine (5 mg·kg−1) and ketamine (90 mg·kg−1), the animals were placed under a surgical microscope. A segment of the extrathoracic trachea was exposed, its outer diameter was measured and a sterile nylon band (Ethilon) of specific length was sutured around the trachea, to provoke a reduction in surface area by half of the initial. Mice recovered from anaesthesia and returned to their cage. Although we have not measured airway resistance directly, we have previously shown that our model of tracheal banding results in more negative intrathoracic pressures (using an oesophageal catheter) (Pi/Pimax ∼22%), significantly different from quietly breathing mice (Pi/Pimax ∼9%), confirming the presence of increased work of breathing [23] and results in significant hypercapnic ventilatory failure and reduced minute ventilation, as shown in the supplementary material (supplementary tables E1 and E2).
RB was induced 24 h after the last CS exposure (of each time frame – subacute (1 month) or chronic (6 months)) and for 24 h in total to simulate the mechanical effects of an acute COPD exacerbation. Sham-operated animals served as controls. (A schematic overview of our study is presented in the supplementary material.)
Thus, the following groups of animals were created: 1 month air plus sham (n=6); 1 month air plus RB (n=11); 1 month CS plus sham (n=11); 1 month CS plus RB (n=10); 6 months air plus sham (n=8); 6 months air plus RB (n=9); 6 months CS plus sham (n=13); 6 months CS plus RB (n=11). To ensure similar exposure to CS, sham-operated mice and mice that underwent tracheal banding were assigned to each group after the final CS exposure session of the 1- or 6-month period.
Respiratory system mechanics
Respiratory system mechanics were estimated with the use of a small animal ventilator (Flexivent, Scireq), as previously described by our group [23]. By the end of the RB session, the animals were anaesthetised by an intraperitoneal injection of ketamine (90 mg·kg−1) and xylazine (10 mg·kg−1) and tracheostomised below the level of tracheal banding, after removing the nylon band, with a tracheal cannula (20 G). They were then attached to the ventilator and ventilated with a tidal volume of 7 mL·kg−1, a respiratory rate of 150 breaths·min−1 and an end-expiratory pressure of 3 cmH2O. Following 3 min of ventilation, a static volume pressure curve was performed by a stepwise inflation and deflation of the respiratory system to a total volume of 40 mL·kg−1 in seven steps in total. The static compliance of the respiratory system was estimated by the slope of the mid-linear part of the expiratory limb of the pressure–volume (PV) curve, whereas hysteresis (area between the inspiratory and expiratory limbs) was calculated by the FlexiVent software. Prior to the test (30 s), the lung volume history was standardised by a deep inspiration until the airway pressure reached 30 cmH2O, a pressure corresponding to total lung capacity in mice [26].
Blood samples
Following respiratory system mechanics measurement, the abdominal cavity was exposed, and 1 mL of blood was collected from the inferior vena cava and centrifuged (1780 ×g, 10 min, 40°C). Plasma was collected and stored at −80°C for further analysis.
Bronchoalveolar lavage fluid
After blood sampling, the animals were euthanised by exsanguination (vena cava dissection) and the thoracic cavity was exposed. Bronchoalveolar lavage fluid (BALF) was performed in the right lung with three aliquots of 0.5 mL normal saline. BALF was centrifuged (300 ×g, 10 min, 4°C); the supernatant was stored at −80°C and the cell pellet was resuspended in 1 mL of normal saline. Total cell count was measured with trypan blue stain and differential cell counts were performed on May–Grünwald stained cytospins. Percentages of macrophages, monocytes and lymphocytes were determined by counting their number in 300 cells. Eosinophil count was negligible and omitted from further analysis.
Lung histology
Following BAL, the right main bronchus was ligated, and the right lung was removed, frozen in liquid nitrogen and stored at −80°C. The left lung was fixated by gentle infusion of 4% formaldehyde under constant pressure of 20 cmH2O, removed and embedded into paraffin. Sagittal plane sections (4-μm thick) were cut and stained with haematoxylin–eosin, using classical histological techniques. The presence of pulmonary inflammation and injury was evaluated by an experienced pathologist (S. Theocharis) via a histological score originally published by Murao et al. [27], and previously also utilised by our group to evaluate RB-induced lung injury [21, 22, 28], in accordance with current standards for measurements of experimental lung injury in animals [29]. This score is also used in literature for a variety of animal models of lung injury, including lipopolysaccharide (LPS)-induced [30], CS-induced [31] and sepsis-induced [32]. Five individual key histological features of acute lung injury were evaluated: 1) capillary congestion; 2) intra-alveolar haemorrhage; 3) interstitial and 4) intra-alveolar neutrophil infiltration; and 5) focal thickening of alveolar membranes. They were scored semi-quantitatively as follows: 0, absence; or presence to a mild (1), moderate (2) or severe (3) degree. A total lung injury score was calculated. Alveolar oedema, another histological feature of lung injury, appeared rarely and thus was not further included in analysis.
Quantification of emphysema was determined only in the chronic exposure to CS (6 months) by the mean linear intercept (Lm) and the destructive index (DI), as previously described by our group [23]. For the quantification of the DI, alveolar and/or duct spaces were classified as normal, or destroyed when it met one of the following criteria: 1) two or more breaks in the alveolar wall; 2) two or more parenchymal rags in the lumen of alveolar ducts; 3) clearly abnormal morphology; and 4) classic emphysematous change [33].
Pulmonary Sp-D levels in blood and BALF
The protein levels of Sp-D were measured with ELISA, according to the standard protocol supplied by the manufacturer (DuoSet ELISA; R&D Systems), in the BALF supernatant and peripheral blood (plasma). BALF was preferred over lung tissue to quantify Sp-D levels, given that Sp-D is produced by alveolar Type II cells, secreted in the alveolar space and found mostly in the aqueous BALF [34].
Protein extraction and cytokine levels in lung tissue
Frozen lung tissue samples were homogenised, as previously described by our group [21], and protein levels of KC, the murine ortholog of IL-8 [35] and MCP-1 were determined in the lung tissue homogenates with ELISA, according to the manufacturer's protocol (DuoSet ELISA; R&D Systems) and were normalised to total protein content of lung homogenates. Protein levels of cytokines were quantified in lung tissue homogenates (over BALF only) to provide a more “holistic” image of cytokine expression (alveolar, interstitium and endothelium).
Statistical analysis
Variables were described as medians (interquartile range), and the different groups were compared with the Kruskal–Wallis test as non-parametric distributions and significance values have been adjusted by the Holm–Bonferroni correction for multiple tests [36]. Linear regression analysis was also performed to demonstrate the effect of the exposure to CS, RB and the interaction between them. To minimise the possibility of falling into Type 1 error, correlation tests and dot plots were performed, and variables with a strong correlation (Spearman rank correlation coefficient (rho) >0.6 and graphically confirmed) were not included in the linear regression analysis. A two-tailed p-value <0.05 was considered statistically significant. Statistical analyses were performed with IBM SPSS, Statistics for Windows version 26.
Comparison between the different time points (1 month versus 6 months) of CS exposure was not performed, as it was not included in the objectives of our study.
Results
Bronchoalveolar lavage fluid
CS exposure induced a significant increase in the total number of inflammatory cells measured in BALF at both time points of the experiment (∼8-fold in the subacute (p=0.002, figure 1a) and ∼1.9-fold in the chronic exposure (p<0.001, figure 2a) compared to air-exposed mice), due to increased numbers of macrophages (p<0.01 to control at both time points, figures 1b and 2b), but also due to a rise in neutrophils in the subacute and chronic exposure (p<0.001 to control, figures 1c and 2c). CS exposure increased lymphocytes in BALF (p<0.001 at both time points, figures 1d and 2d, compared to air-exposed mice). Interestingly, combining RB through tracheal banding with CS at 1 month significantly contributed to the effect of CS on total cell count (p=0.001, see supplementary table E3 and figure 1a), and neutrophil count was increased when CS was combined with RB, compared to CS alone (p=0.012, figure 1c). Moreover, lymphocytes count at the 1st month is related mostly with the coexistence of CS and RB (p=0.013, supplementary table E3). At 6 months, both macrophages and neutrophil count was further increased in CS-exposed mice after RB (p=0.027 and p=0.005, respectively, see figure 2b and c), with total cell count being related significantly with CS and RB as independent factors (p<0.001 and p=0.002, respectively, supplementary table E3).
FIGURE 1.
The effect of resistive breathing (RB) on total and differential cell counts in bronchoalveolar lavage fluid (BALF) in subacute (1 month) cigarette smoke (CS) exposure. Subacute CS exposure resulted in a) an increased total cell count, due to b) raised macrophage, c) neutrophil and d) lymphocyte count. RB further increased macrophage and neutrophil counts. Data presented as median (IQR) with overlaid datapoints. Blue box plot: air plus sham; light blue box plot: air plus RB; light red box plot: CS plus sham; red box plot: CS plus RB. #: p<0.05 to air plus sham; ¶: p<0.05 to CS plus sham.
FIGURE 2.
The effect of resistive breathing (RB) on total and differential cell counts in bronchoalveolar lavage fluid (BALF) in chronic (6 months) cigarette smoke (CS) exposure. CS exposure resulted in a) an increased total cell count, due to b) raised macrophages and d) lymphocyte count. Superimposed RB increased both macrophage and c) neutrophil count. Data presented as median (IQR) with overlaid datapoints. Blue box plot: air plus sham; light blue box plot: air plus RB; light red box plot: CS plus sham; red box plot: CS plus RB. #: p<0.05 to air plus sham, ¶: p<0.05 to CS plus sham.
Cytokine levels in lung tissue
CS exposure caused an increase in cytokine levels of KC both at 1 and at 6 months (p=0.001 for both time points, figure 3a and c, respectively). Combining RB with CS exposure did not further increase KC levels. MCP-1 levels were increased at 1 month of CS exposure only after combining with RB (p=0.002 to control, figure 3b). MCP-1 expression was not affected in the chronic CS exposure (figure 3d).
FIGURE 3.
Lung tissue protein levels of MCP-1 and keratinocyte-derived chemokine (KC) in cigarette smoke (CS)-exposed mice and the effect of resistive breathing (RB). CS exposure raised KC expression in the lung both at a) 1 month and c) 6 months. Superimposed RB did not affect KC expression. b) In contrast, RB resulted in a further increase of MCP-1 levels after 1 month of CS exposure. d) Chronic CS exposure did not affect MCP-1 protein levels in the lung. Data presented as median (IQR) with overlaid datapoints. Blue box plot: air plus sham; light blue box plot: air plus RB; light red box plot: CS plus sham; red box plot: CS plus RB. #: p<0.05 to air plus sham.
SP-D levels in blood and BALF
Exposure to CS for both 1 and 6 months induced Sp-D levels in BALF (∼2.1-fold to control, p=0.002, figure 4a and ∼1.86-fold to control, p=0.002, figure 4c, respectively). Interestingly, although RB did not cause an increase in Sp-D levels in BALF and contributed negatively to the effect of CS at 6 months (p<0.001), Sp-D levels in blood were mostly increased in groups of animals that underwent RB (either air or CS exposed, figure 4b and d).
FIGURE 4.
Bronchoalveolar lavage fluid (BALF) and blood protein levels of pulmonary surfactant protein D (Sp-D) in cigarette smoke (CS)-exposed mice and the effect of resistive breathing (RB). CS resulted in increased levels of Sp-D in BALF in both a) 1 and c) 6 months. b and d) Increased blood levels of Sp-D were noticed in groups of animals that underwent RB, denoting an increased lung permeability. Data presented as median (IQR) with overlaid datapoints. Blue box plot: air plus sham; light blue box plot: air plus RB; light red box plot: CS plus sham; red box plot: CS plus RB. #: p<0.05 to air plus sham; ¶: p<0.05 to CS plus sham.
Respiratory system mechanics
RB was associated with a reduction in static compliance at 1 month in air-exposed mice (Cst, p=0.007 to sham, figure 5a), as previously described by our group, whereas the reduction in Cst in RB and CS-exposed mice failed to reach statistical significance, due to multiple comparison correction. In contrast, at 6 months of CS exposure no difference was observed between sham-operated and RB animals (figure 5c). Hysteresis was not affected by CS or RB compared to control animals (sham and air exposed) (figure 5).
FIGURE 5.
The effect of resistive breathing (RB) on respiratory system mechanics in cigarette smoke (CS)-exposed mice. a and c) RB caused a reduction in static compliance, in accordance with the presence of lung injury only in air-exposed mice at 1 month, whereas no significant reduction was observed in mice with CS exposure at 1 or 6 months. b and d) No change in hysteresis was noticed either at 1 or at 6 months. Data presented as median (IQR) with overlaid datapoints. Blue box plot: air plus sham; light blue box plot: air plus RB; light red box plot: CS plus sham; red box plot: CS plus RB. #: p<0.05 to air plus sham.
Lung histology
An increase of total lung injury score was observed for both the subacute (1 month) and chronic (6 months) CS exposure compared to control (1 month, p=0.001 and 6 months, p=0.01, figure 6a and b and table 1), mainly due to inflammatory cell infiltration (figure 7). The addition of RB further increased focal thickening of alveolar membranes in CS-exposed mice for 1 month (p=0.011, table 1). Histological analysis following 6 months of CS exposure confirmed also the development of pulmonary emphysema (figure 7) by the increase in both Lm and DI indices, compared to air (p<0.001 for both, figure 6c and d). Adding RB significantly contributed to the variability of Lm over 6 months (p=0.025), and both Lm and DI were further increased in the combined RB and CS group, compared to CS alone (p=0.001 for both).
FIGURE 6.
Histological analysis of haematoxylin and eosin stained tissue sections of cigarette smoke (CS)-exposed mice and the effect of resistive breathing (RB). a and b) CS exposure and RB independently caused a rise in lung injury score, both at 1 and 6 months. Adding RB to 6 months of CS exposure caused a further increase in pulmonary emphysema, as measured by c) the mean linear intercept (Lm) and d) the destructive index (DI). Data presented as median (IQR) with overlaid datapoints. Blue box plot: air plus sham; light blue box plot: air plus RB; light red box plot: CS plus sham; red box plot: CS plus RB. #: p<0.05 to air plus sham; ¶: p<0.05 to CS plus sham.
TABLE 1.
Histological features of lung injury following cigarette smoke (CS) exposure and resistive breathing (RB)
| Capillary congestion | Intra-alveolar haemorrhage | Interstitial leukocyte infiltration | Ιntra-alveolar leukocyte infiltration | Focal thickening of alveolar membranes | Total score | |
|---|---|---|---|---|---|---|
| 1 month | ||||||
| Air plus sham | 0.00 (0.00–0.00) | 0.00 (0.00–0.00) | 0.00 (0.00–0.00) | 0.00 (0.00–0.00) | 0.00 (0.00–0.00) | 1.00 (0.00–1.00) |
| Air plus RB | 1.00 (0.00–1.00) | 0.00 (0.00–0.00) | 1.00 (1.00–1.00)# | 0.00 (0.00–1.00) | 1.00 (0.00–1.00) | 3.00 (2.00–3.00)# |
| CS plus sham | 1.00 (0.00–1.00) | 0.00 (0.00–0.00) | 1.00 (1.00–1.00)# | 2.00 (1.00–2.00)# | 0.00 (0.00–0.00) | 3.00 (3.00–4.00)# |
| CS plus RB | 1.00 (1.00–2.00) | 0.00 (0.00–1.00) | 1.00 (1.00–2.00)# | 1.00 (1.00–2.00) | 1.00 (1.00–2.00)#,¶ | 5.00 (3.00–9.00)# |
| 6 months | ||||||
| Air plus sham | 0.00 (0.00–0.00) | 0.00 (0.00–0.00) | 0.00 (0.00–0.00) | 0.00 (0.00–0.00) | 0.00 (0.00–0.00) | 1.00 (0.00–1.00) |
| Air plus RB | 1.00 (1.00–1.00) | 0.00 (0.00–0.00) | 1.00 (1.00–1.00)# | 1.00 (0.00–1.00) | 1.00 (0.00–1.00) | 4.00 (2.00–4.00)# |
| CS plus sham | 0.00 (0.00–0.00) | 0.00 (0.00–0.00) | 1.00 (1.00–1.00)# | 0.00 (0.00–1.00) | 1.00 (0.00–1.00) | 2.00 (1.00–3.00)# |
| CS plus RB | 0.00 (0.00–0.00) | 0.00 (0.00–0.00) | 1.00 (1.00–1.00)# | 1.00 (0.00–1.00) | 1.00 (0.00–1.00) | 3.00 (2.00–3.00)# |
Data are presented as median (IQR). #: p<0.05 to air plus sham; ¶: p<0.05 to CS plus sham.
FIGURE 7.
Representative haematoxylin and eosin stained tissue sections. Compared to control (upper left), subacute (1-month, middle left) and chronic (6-month, lower left) cigarette smoke (CS) exposure was associated with increased leukocyte infiltration. Adding resistive breathing (RB) further increased focal membrane thickening in subacute CS exposure (middle right). 6 months of CS exposure (lower left) was also marked by the presence of pulmonary emphysema that was further deteriorated by superimposed RB (lower right). Please note that for control (air plus sham) and sole RB (air plus RB) groups only one figure is depicted for both time points (1 and 6 months), to avoid complexity, since no difference was found.
Discussion
The main finding of our study is that spontaneous breathing through increased airway resistance aggravates pulmonary inflammation and emphysema in a mouse model of CS exposure.
Pathogenesis of COPD is characterised by the induction of pulmonary inflammation, including both deranged innate immunity (e.g. neutrophilic inflammation) and induction of adaptive immunity, e.g. lymphocyte infiltration, including CD8+ T-cells that are found in the lungs of COPD patients [37]. Exacerbations are acute episodes of worsening symptoms and lung function, associated with amplified pulmonary inflammation [2]. Although infections (bacterial and/or viral) are a major cause of COPD exacerbations and could explain the increased inflammatory burden, it must be acknowledged that exacerbations are heterogeneous with non-infectious causes as well [38], and the degree of inflammation (sputum neutrophils) during an exacerbation is augmented irrespective of the infectious cause and in correlation with the severity of the exacerbation [39]. Thus, further mechanisms apart from infections are needed to explain the pathogenesis of COPD exacerbations.
COPD exacerbations are associated with increased mechanical stress on the lung, through strenuous contractions of the inspiratory muscles and/or the presence of hyperinflation [40]. Although we have previously shown that in experimental animal models RB induces pulmonary inflammation and injury [20, 21], this was performed in previously healthy animals, whereas the potential synergistic effect with underlying inflammation, e.g. due to CS exposure, has not been established.
The murine model of CS exposure is a well-described model of COPD [41], leading to similar carboxyhaemoglobin levels as humans following acute exposure (10–14%) [41]. As expected, both subacute (1 month) and chronic (6 months) CS exposure induced pulmonary inflammation, as shown by the increased BALF cellularity. Exposure of lung epithelial cells to CS can cause the release of chemoattractant cytokines [42]. Accordingly, we found increased expression of KC in the lung tissue at 1 and 6 months of CS exposure.
The addition of RB resulted in a further augmentation of BALF cellularity, mainly at the subacute CS exposure. This was not associated with a further increase in KC levels, suggesting an alternative mechanism of neutrophil chemotaxis, whereas MCP-1 levels were only increased at 1 month when CS exposure was combined with RB. To our knowledge this is the first report of a “two-hit” model of CS exposure combined with the mechanical impact of breathing through increased airway resistance. Previously, increased neutrophilic inflammation was shown following a combination of CS exposure with positive-pressure mechanical ventilation [43], a study that further supports the synergy between CS and mechanical stress-induced lung injury.
Sp-D is a protein mainly synthesised by alveolar Type II cells. It has a major role in surfactant homeostasis and contributes to a wide variety of immune functions. At low levels it has an anti-inflammatory role, whereas when there is a constant and severe insult in the lung, it can assume pro-inflammatory roles to complement pulmonary innate and adaptive immunity [34]. Following injurious stimuli in the lung, Sp-D is detected in serum due to both increased expression from alveolar Type II cells and increased epithelial permeability [16]. CS exposure caused an elevation of its protein levels in BALF at both time points of the experiment but not in peripheral blood, suggesting the absence of “spill” of the protein from the alveoli to the blood. The increased levels of Sp-D in the BAL following CS exposure are in accordance with previous studies in mice at both mRNA and protein level [44]. Moreover, alveolar type II hyperplasia has been shown in a rat model of CS exposure [45]. In contrast, Sp-D levels in peripheral blood were increased after applying tracheal banding for 24 h, denoting the derangement of epithelial permeability [46] allowing its spill into the circulation. Interestingly, in COPD patients Sp-D levels in the blood are negatively associated with lung function [47] and are further increased during exacerbation [15], suggesting it can be a potential diagnostic biomarker for COPD exacerbations.
In accordance with previous studies [41], microscopic analysis of lung tissue sections revealed the presence of emphysema following 6 months of CS exposure, quantified by both the Lm and DI. Interestingly, the addition of RB aggravated the CS-induced pulmonary emphysema, in accordance with our previous findings, where RB augmented elastase-induced pulmonary emphysema [23]. Although a direct effect of increased mechanical stress is plausible, especially since protease-exposed lungs have a low threshold for mechanical disruption [48], augmented inflammation may also contribute to this effect. Indeed, increased neutrophilic infiltration is observed after the addition of RB to chronic CS exposure, which is expected to elevate neutrophil elastase activity [49], whereas RB can also increase the activity of both MMP-9 and MMP-12 in the lung, as previously shown by our group [50]. Interestingly, using novel imaging analysis, it was previously shown that “normal appearing” areas adjacent to emphysematous areas are exposed to increased mechanical deformation [51], and these areas are the sites where disease progression occurs [52].
Finally, 24 h of RB resulted in a decrease of static compliance of the respiratory system, in accordance with the presence of lung injury. Of note, this effect was only noticed at 1 month and not 6 months, probably reflecting the opposing effects of emphysema formation and the presence of lung injury on static lung compliance.
Clinical implications
Although cigarette smoking exposure remains the most recognised risk factor for COPD, it is nowadays accepted that COPD has a multifactorial pathogenesis, including genetics, early life events and air pollution [7], while exacerbations promote COPD progression and accelerate forced expiratory volume in 1 s decline [53]. COPD exacerbations are characterised by increased pulmonary inflammation due to various factors and not solely to infections. Our data raises the intriguing possibility that the increased mechanical stress imposed by increased airway resistance during a COPD exacerbation may independently exert a synergistic injurious stimulus with lung inflammation. Hitherto, animal models of COPD exacerbation usually combine a model of stable COPD, e.g. CS exposure, with an additional inflammatory stimulus, e.g. a virus/bacterial exposure or LPS [54]. However, in this case it is not feasible to discriminate between the injurious inflammatory stimulus of the additional exposure and the mechanical consequences of an exacerbation (bronchoconstriction induced by the inflammation). Finally, our data regarding the increase of Sp-D levels in the blood only after CS was combined with RB, despite an increase in BALF levels in CS only exposure, support the hypothesis of a “spill” of inflammatory markers from the lung into the circulation during exacerbation that may contribute to systemic inflammation in COPD, a phenomenon still in debate in literature [55].
We acknowledge that our study, although describing the synergistic effect of RB on CS induced inflammation and emphysema, does not provide the pathogenetic mechanisms involved. Previously, we have shown in a series of studies that mechanisms implicated in RB lung injury in healthy animals include mechanosensitive pathways, e.g. the TRPV4 channels and Src kinase signalling [28, 56] and the activation of the soluble guanylyl cyclase pathway [20]. Finally, although our model may better mimic upper airway obstruction, we believe that the pathophysiological consequences described are also applicable to peripheral airway obstruction, as in COPD [40]. This view of ours is supported by the fact that we have obtained similar findings of lung injury in the rat model of combined expiratory and inspiratory resistances, with the expiratory resistances higher than the inspiratory resistances, as is the case in intrathoracic airway obstruction, which occurs in COPD, which we have previously published [22] and the model of tracheal banding-induced lung injury, which we have also previously published [20], which was the mechanical stressor used in the current study.
A limitation of our study is that we did not include a measurement of CO2 or total particulate matter per cubic metre concentration in the exposure chamber to further standardise our experimental settings. Moreover, CS exposure per mouse was not quantified by biomarkers, such as serum carboxyhaemoglobin or cotinine levels, [41, 57] or by measuring smoke components in the lung tissue, to exclude a possible difference of CS exposure (or clearance of smoke particles from the lung) between quietly breathing and RB. On the other hand, the mice were divided into sham operation or tracheal banding 24 h after the last exposure to CS. Therefore, although differences in exposure cannot be ruled out, we believe that this mechanism does not significantly contribute to our results, especially given that we use long-term exposure periods (1 and 6 months) compared to the 24 h of RB.
In conclusion, the findings of our study suggest that RB through tracheal banding aggravates pulmonary inflammation, induces lung injury and worsens emphysema in CS-exposed mice, providing a novel preclinical animal model of COPD exacerbations.
Acknowledgements
The authors would like to thank Zoi Kollia (“Marianthi Simou” Applied Biomedical Research and Training Center, Medical School, National and Kapodistrian University of Athens, Evangelismos Hospital, Athens, Greece) for technical assistance with experimental procedures and DHL Hellas for waiving material shipping costs.
Footnotes
Provenance: Submitted article, peer reviewed.
Ethics statement: Handling of the laboratory animals was in accordance with the European Union Directive (2010/63/EU) on the protection of animals used for scientific purposes. Experimental procedures were approved by the Scientific Council and Ethics Committee of Evangelismos Hospital (217/4-10-2016).
Conflict of interest: The authors have no conflict of interest to disclose.
Support statement: This study was supported by a Grant of the Hellenic Thoracic Society. Funding information for this article has been deposited with the Open Funder Registry.
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