Combination Therapy With Clarithromycin and Montelukast Suppresses the Progression of Aortic Aneurysms in Mice With Chronic Obstructive Pulmonary Disease

Abstract

Background: Chronic obstructive pulmonary disease (COPD) is a known risk factor for aortic aneurysm (AA) enlargement and rupture. This study investigated the effects of clarithromycin (CAM) and montelukast (Mont), which are medications used to treat COPD, on AA progression in a murine model of COPD.

Methods and Results: Apolipoprotein E-deficient mice, aged 28–40 weeks, were infused with angiotensin II by osmotic pumps to induce AA formation. Some of them received COPD induction through a single dose of porcine pancreatic elastase via the trachea. Mice were divided into 3 groups: AA (n=16; AA only, treated with saline); AA-C (n=10; AA and COPD, treated with saline); and AA-Cm (n=10; AA and COPD, treated with CAM and Mont). CAM and Mont were administered orally on a daily basis. After 28 days, aortic diameter, elastin content, matrix metalloproteinase (MMP) activity, and inflammatory markers were evaluated. The AA-C group exhibited significantly larger aneurysm diameter than the AA group (2.41 vs. 1.97 mm; P<0.05). Compared with the AA-C group, the AA-Cm group had higher elastin content (46.8 vs. 32.3%; P<0.01), decreased TNF-α level (115.5 vs. 141.0 pg/mL; P<0.05), reduced MMP-9 activity (54.8 vs. 75.4 pg/mL; P<0.01), and lower M1/M2 macrophage ratio.

Conclusions: CAM and Mont attenuate AA progression in COPD by reducing inflammation, preserving elastin, and increasing infiltrated M2 macrophages, suggesting they have a therapeutic potential.

Central Figure

Aortic aneurysm (AA) is characterized by a localized weakening of the aortic wall, leading to asymptomatic enlargement and potential rupture over several years. It is estimated that approximately 170,000 people die annually worldwide due to AA-related disease.¹ The standard treatment involves surgical intervention with prosthetic graft replacement to prevent rupture; however, no therapy reliably prevents aneurysm enlargement.² Indeed, once rupture occurs, the in-hospital mortality remains as high as approximately 50% in a large registry.² Furthermore, open aortic replacement via laparotomy or thoracotomy is highly invasive and may not be suitable for elderly patients with poor surgical tolerance. Less invasive endovascular treatment may apply to these high-risk patients, but the anatomical constraints such as aneurysm location and morphology often limit their applicability. Although numerous studies, ranging from basic experimental studies to clinical observational and interventional trials, have explored the potential effects of various drugs – such as antihypertensives, anti-inflammatory agents, and even certain antibiotics – no definitive pharmacological therapy has been established to pause or reverse AA progression. Even amlodipine and candesartan, which showed promising anti-aneurysmal effects during in vivo pre-clinical studies, failed to suppress the growth of small abdominal AAs in the clinical study by Mitsui et al.³ Therefore, alternative therapies beyond current surgical approaches are needed to prevent aortic enlargement and rupture.

The pathophysiology of AA is characterized by chronic inflammation, particularly on the adventitial side of the aorta, and accumulation of inflammatory cells such as lymphocytes and macrophages.⁴ In particular, macrophages play a crucial role in AA development and progression by producing proteolytic enzymes and inflammatory mediators, leading to increased expression of cytokines, such as tumor necrosis factor-α (TNF-α), interleukin (IL)-1β, IL-6, and monocyte chemoattractant protein-1 (MCP-1) and chemokines in AA tissue.⁵ The most characteristic pathological feature of AA formation is the degradation of extracellular matrix (ECM) components, especially elastin. Degradation of ECM results from increased protease activity, including matrix metalloproteinases (MMPs) secreted by macrophages, vascular smooth muscle cells, and stromal cells.^6,7 Under normal physiological conditions, the ECM of the vascular wall undergoes continuous remodeling, with a well-balanced process of degradation of existing ECM proteins and their replacement by newly synthesized proteins. However, this balance is disrupted in the context of AA formation. Specifically, excessive ECM degradation, impaired synthesis of ECM proteins, and the apoptosis of smooth muscle cells lead to deleterious ECM remodeling that contributes to the development of AAs.⁸

Chronic obstructive pulmonary disease (COPD) is a lung disease characterized by airflow obstruction due to a combination of peripheral airway lesions and emphysema, primarily caused by long-term inhalation of harmful substances such as cigarette smoke.⁹ COPD is associated with increased local and systemic inflammation, involving elevated levels of inflammatory cells such as neutrophils, macrophages, and lymphocytes, and higher concentrations of inflammatory mediators including leukotriene B4, IL-8, and TNF-α.¹⁰ Systemic oxidative stress and increased circulating inflammatory cytokines are also observed in COPD patients, promoting pulmonary tissue damage and potentially exacerbating vascular inflammation.^10,11 Notably, COPD is a well-known independent risk factor for AA enlargement and rupture regardless of smoking status in patients.¹² However, the detailed molecular mechanisms by which COPD contributes to the progression of AA are still unclear.

In general, clarithromycin (CAM) and montelukast (Mont) are widely used therapeutic agents for COPD.^13,14 Beyond their routine application in respiratory disorders, these agents have also been investigated for their anti-aneurysmal properties.^15,16 CAM, a macrolide antibiotic, exhibits anti-aneurysmal properties by acting as a key regulator of inflammatory responses through inhibition of nuclear factor-κB (NF-κB) phosphorylation. We reported that administration of CAM suppressed the progression and rupture of AA in mice by inhibiting elastin degradation due to the suppression of macrophage infiltration and reduction of MMPs activities associated with the suppression of NF-κB phosphorylation.¹⁵ Mont operates through different mechanisms as a selective cysteinyl leukotriene receptor 1 antagonist to protect against AA formation. Our previous study showed that Mont promoted suppression of gene expression including MMP-2, MMP-9, and IL-1β, and polarized toward the anti-inflammatory M2 phenotype along with the expression of arginase-1 and IL-10 in cultured macrophages.¹⁶ Furthermore, administering Mont demonstrated decreased aortic expansion, reduced MMP-2 activity, and increased infiltration of M2 macrophages in AA mice. These reports suggest that CAM and Mont effectively prevent AA progression through their distinct molecular pathways. Additionally, because these 2 agents are already widely used in clinical practice, they have a favorable safety profile and straightforward potential for clinical application, making them particularly promising candidates for the treatment of patients with concomitant COPD and AA.

Therefore, in the present study, we aimed to clarify whether COPD accelerates AA progression and to investigate the therapeutic effects of concurrent administration of CAM and Mont in a murine model of AA with coexisting COPD. We assessed aortic diameter, elastin content, MMP activity, and inflammatory markers in ApoE-deficient mice treated with angiotensin II (Ang II) infusion to induce AA, with or without additional COPD induction by porcine pancreatic elastase (PPE). By focusing on clinically relevant, commonly prescribed medications, we sought to determine whether dual targeting of overlapping inflammatory pathways could offer a viable strategy to mitigate AA progression, especially in the context of COPD.

Methods

Experimental procedures followed the ARRIVE guideline¹⁷ and the Guide for the Care and Use of Laboratory Animals published by the National Institutes of Health,¹⁸ and were approved by the Institute for Laboratory Animal Research at the Nagoya University Graduate School of Medicine (Protocol No. M220112-001). Apolipoprotein E-deficient (ApoE^−/−) mice on a C57BL/6 background were procured from the Jackson Laboratory (Sacramento, CA, USA) and maintained on a standard diet under controlled conditions of 24℃, 12 h light/dark cycle.

Animal Preparation

Male ApoE^−/− mice aged between 28 and 40 weeks were used for the experiments. To induce AA formation, mice were infused with Ang II (Calbiochem, San Diego, CA, USA) by sustained-release osmotic pump (1,000 ng/kg/min; ALZET model 2004; DURECT, Cupertino, CA, USA) subcutaneously for 4 weeks under general anesthesia with isoflurane. To induce COPD, intratracheal instillation of PPE (0.2 units in 50 μL; E1250; Millipore Sigma, St Louis, MO, USA) was performed under general anesthesia with isoflurane.¹⁹ This model fully developed features of COPD within 4 weeks.

Experimental Protocol

As shown in Figure 1, a total of 36 mice were randomly divided into 3 groups: (1) AA-induced mice (AA group; n=16); (2) AA- and COPD-induced mice (AA-C group; n=10); and (3) AA- and COPD-induced mice with CAM and Mont treatment (AA-Cm group; n=10). In the AA group, mice received the Ang II infusion without prior COPD induction. In the AA-C group, mice underwent COPD induction via intratracheal PPE administration, followed by Ang II infusion on the same day. The AA and AA-C groups received 200 μL of saline daily via gastric gavage. In the AA-Cm group, mice underwent the same procedures as the AA-C group for COPD induction and Ang II infusion, and received daily oral administration of CAM (100 mg/kg/day; Taisho Pharmaceutical Co., Ltd, Tokyo, Japan) and Mont (10 mg/kg/day; Tokyo Chemical Industry Co., Ltd., Tokyo, Japan) dissolved in 200 μL of saline via gastric gavage. The dosages of CAM and Mont were consistent with our previous studies.^15,16 All groups were administered daily for 28 days, and then mice were euthanized. Throughout the experimental period, evaluations included weekly aortic ultrasound examinations to measure the maximum AA diameter, monitoring of aortic event rates such as rupture or sudden death, assessment of elastin content using Elastica van Gieson staining, analysis of macrophage accumulation using immunofluorescence staining, and quantification of cytokine and chemokine levels using enzyme-linked immunosorbent assay (ELISA).

Figure 1.

Experimental design. All mice were implanted with an angiotensin II-releasing pump. The group with chronic obstructive pulmonary disease (COPD) was given the porcine pancreas-derived elastase once on the operative day. The AA and AA-C groups were given saline orally, and the AA-Cm group was given clarithromycin (CAM; 100 mg/kg/day) and montelukast (Mont; 10 mg/kg/day) orally daily for 4 weeks. Ultrasound was performed once a week, and mice were euthanized on the 4th weekend. MMP, matrix metalloproteinase.

Echography

The maximum AA diameter in mice was measured using ultrasound (10 MHz; GE Healthcare, Chicago, IL, USA) before implantation and weekly thereafter. Mice were anesthetized, and ultrasound was performed in the supine position to measure the maximum AA diameter. A widely accepted clinical diagnostic standard for AA is an aortic diameter increase of >50%, so we defined AA as dilation to at least 1.5 times the preimplantation diameter.

Aortic Specimen Preparation and Measurement

The aorta was dissected from below the diaphragm to above the renal artery. The maximum minor axis of the exposed AA diameter was measured using microscopy, as previously described.²⁰ Specimen images were obtained with a DP70 digital camera (Olympus, Tokyo, Japan) attached to a stereomicroscope using DP controller software (Olympus). A 2-mm length of the suprarenal aorta was cut for histologic examination. The remaining aorta was ground and homogenized to analyze MMP enzymatic activity and perform ELISA.

Histologic Examination

Suprarenal aorta specimens were embedded in optimal cutting temperature (OCT) compound (Tissue-Tek; Sakura Finetek USA, Torrance, CA, USA) and cut into 10-μm thick sections using a cryostat microtome (CM3000; Leica Microsystems, Wetzlar, Germany). Frozen sections were fixed with 4% paraformaldehyde (FUJIFILM Wako Pure Chemical Corporation, Osaka, Japan) for 15 min and stained with Elastica van Gieson using Weigert›s resorcin-fuchsin (Muto Pure Chemicals, Tokyo, Japan), which is specific for elastic lamellae. Stained samples were photographed with an Olympus DP80 digital camera. Images were analyzed using cellSens software (Olympus) to determine the percentage area of elastin staining and medial components between the elastic lamellae (elastin gap area) relative to the total medial tissue area, as well as to identify the number of elastic lamellae and elastin breaks. For each section, the average number of elastic lamellae and the total number of breaks per elastic lamina were assessed in 3 aortic sections by circumferential counting of all laminae.

To identify macrophages in the aortic wall, immunofluorescence staining was performed. Frozen cross-sections were fixed at 4% paraformaldehyde for 15 min and blocked with 10% bovine serum albumin for 30 min. Rabbit polyclonal anti-inducible nitric oxide synthase (iNOS) antibody (1 : 50; Abcam, Cambridge, UK), rabbit polyclonal CD206 antibody (1 : 1,000; Abcam), and mouse CD68 antibody (1 : 200; Abcam) were used. Anti-mouse IgG (H+L), F(ab′)₂ fragment Alexa Fluor 488-conjugated antibody (1 : 5,000; Cell Signaling Technology, Danvers, MA, USA) and anti-rabbit IgG (H+L), F(ab′)₂ fragment Alexa Fluor 555-conjugated antibody (1 : 5,000; Cell Signaling Technology) were used as secondary antibodies for detection. Negative control experiments used mouse IgG1 and rabbit IgG isotype control antibodies at the same concentration as the primary antibodies (Cell Signaling Technology). Nucleated cells were stained with 4′, 6-diamidino-2-phenylindole Fluoromount-G (Southern Biotech, Birmingham, AL, USA). Slides were photographed with an Olympus DP80 digital camera and quantified using cellSens software ver.2.3 (Olympus).

ELISA

Infrarenal aortic tissue was homogenized using an ultrasonic disintegrator (Sonics & Materials, Inc., Newton, CT, USA) in a protein extraction radioimmunoprecipitation assay (RIPA) buffer (Fujifilm Wako Pure Chemical Corporation). The lysate protein concentration was measured with a Qubit protein assay kit and Qubit 2.0 fluorometer (Thermo Fisher Scientific, Waltham, MA, USA). Equal protein concentrations were applied to each assay kit (TNF-α, IL-1b, IL-6, MCP-1; Thermo Fisher Scientific) and detected according to the manufacturer’s protocol.

MMP-2 and MMP-9 Enzymatic Activity

Endogenous active MMP-2 and MMP-9 in aortic tissues were measured using a SensoLyte 520 MMP-2 assay kit (ANASPEC, Fremont, CA, USA) and a mouse MMP-9 activity assay kit (QuickZyme Bioscience, Leiden, The Netherlands), respectively, according to the manufacturer’s protocol. Equal total protein concentrations were applied to each assay kit and detected.

Statistical Analysis

All statistical analyses were performed with SPSS version 29 (IBM, Armonk, NY, USA). Data are presented as mean±standard error of the mean (SEM). For the comparison of aortic diameters over time between groups, 2-way repeated measures analysis of variance (ANOVA) was initially performed to assess the effects of time and group. Subsequently, repeated measures analysis was conducted within each group, followed by between-group comparisons at each time point. For comparisons among 3 groups, the normality of distribution was first assessed using the Shapiro-Wilk test. When normality was confirmed, Levene’s test was performed to evaluate the homogeneity of variances. If equal variances were assumed, 1-way ANOVA followed by Tukey’s post hoc test was conducted. In cases where equal variances were not assumed, Welch’s ANOVA followed by Games-Howell post hoc test was performed instead. When the normality assumption was violated, the Kruskal-Wallis test was used, followed by pairwise comparisons adjusted using the Bonferroni correction to control for multiple testing. Event-free survival analysis was conducted using the Log-rank test. P values <0.05 were considered statistically significant.

Results

Aortic Diameter Analysis

Representative echography images are shown in Figure 2A. The maximum aortic diameters at day 0 were 1.39±0.03 mm in the AA group, 1.43±0.02 mm in the AA-C group, and 1.43±0.02 mm in the AA-Cm group. At day 14, diameters were 1.69±0.08 mm in the AA group, 2.11±0.14 mm in the AA-C group, and 1.79±0.11 mm in the AA-Cm group, with a statistically significant difference between the AA and AA-C groups (P<0.05). At day 21, the diameters further expanded to 1.87±0.09 mm in the AA group, 2.14±0.14 mm in the AA-C group, and 2.00±0.11 mm in the AA-Cm group. At day 28, the final diameters were 1.97±0.09 mm in the AA group, 2.41±0.19 mm in the AA-C group, and 2.06±0.12 mm in the AA-Cm group (AA vs. AA-C; P<0.05; Table, Figure 2B). Although the Group×Time interaction did not reach statistical significance (P=0.102), the overall group effect and the overall time effect were significant (P<0.001). Specifically, there was a trend towards a larger aortic diameter in the AA-C group compared with the AA-Cm group (Estimate=+0.351 mm; P=0.080), suggesting a modest attenuation of aneurysm expansion in the treatment group (Table). Microscopic analysis on day 28 (Figure 3A) revealed that AA development occurred in 9 (56.3%) out of 16 cases in the AA group, 7 (70%) out of 10 cases in the AA-C group, and 6 (60%) out of 10 cases in the AA-Cm group. However, these rates did not differ significantly among the 3 groups (P>0.05). Event-free survival was assessed, with an aortic event defined as a ≥1.5-fold dilation relative to baseline diameter or death due to rupture (Figure 3B). The AA and AA-C groups each experienced 2 rupture events (12.5% in the AA group, and 20% in the AA-C group), while no ruptures were observed in the AA-Cm group. All-cause mortality was 3 (18.8%) out of 16 in the AA group, 2 (20%) out of 10 in the AA-C group, and 1 (10%) out of 10 in the AA-Cm group. Although the AA-Cm group showed fewer ruptures and lower mortality, neither difference reached statistical significance (P>0.05).

Figure 2.

Aortic diameters were measured by abdominal and thoracic ultrasound during the study period. (A) The yellow arrows indicate the outer edge of the aortic aneurysm (AA). (B) The AA-C group exhibited significantly larger aortic diameters compared with the AA group on Days 14 and 28. *P<0.05 vs. AA group on Days 14 and 28 assessed using Tukey’s test. All groups had a significant aortic dilation over the 4 weeks. Data are means±SEM. **P<0.01 vs. AA group on Day 0, ^‡P<0.05 vs. AA-C group on Day 0, ^†P<0.05 and ^††P<0.01 vs. AA-Cm group on Day 0 assessed using pairwise comparisons adjusted with the Bonferroni correction for multiple testing. AA group, AA-induced mice, treated with saline; AA-C group, AA- and COPD-induced mice, treated with saline; AA-Cm group, AA- and COPD-induced mice, treated with clarithromycin and montelukast.

Table.

Changes in Aortic Diameter Over Time in Each Group

Group	Day 0	Day 7	Day 14	Day 21	Day 28
AA	1.391±0.024	1.593±0.098	1.694±0.097	1.874±0.099	1.968±0.110
AA-C	1.428±0.030	1.868±0.120	2.106±0.116*	2.139±0.119	2.411±0.140*
AA-Cm	1.434±0.030	1.655±0.120	1.787±0.110	2.004±0.119	2.060±0.132

Aortic diameters (mean±SEM; mm) were measured by ultrasound at 5 time points (Days 0, 7, 14, 21, 28) in mice from the AA, AA-C, and AA-Cm groups. *Statistically significant differences compared with the AA group at each time point (P<0.05, 1-way ANOVA post hoc test). (1) Linear mixed-effects model: overall group effect P<0.001; time effect P<0.001; group×time interaction P=0.102. (2) Pairwise contrast (AA-C vs. AA-Cm): estimate +0.351 mm (95% confidence interval –0.05, 0.75), P=0.080. AA, aortic aneurysm; AA group, AA-induced mice, treated with saline; AA-C group, AA- and COPD-induced mice, treated with saline; AA-Cm group, AA- and COPD-induced mice, treated with clarithromycin and montelukast.

Figure 3.

Representative images and aortic event-free survival curves. Aortic events were defined as aortic diameter ≥1.5 times baseline or death from rupture. (A) The black arrows indicate aortic aneurysms (AA). (B) The AA-C group showed a higher incidence of aortic events. The AA-Cm group reduced aortic events compared with the AA-C group. The AA-Cm group had no incidence of rupture, but there was a 1.5-fold or greater dilation. AA group, AA-induced mice, treated with saline; AA-C group, AA- and COPD-induced mice, treated with saline; AA-Cm group, AA- and COPD-induced mice, treated with clarithromycin and montelukast.

Elastica van Gieson Staining

Elastica van Gieson staining in Figure 4A showed the structure of the elastic lamellae was disrupted in these groups, as shown by the arrows in the images. The aortic media in the AA-Cm group contained a higher elastin percentage than the other groups (35.3±1.2% in AA, 32.3±1.5% in AA-C, 46.8±0.96% in AA-Cm; P<0.01; Figure 4B). The average number of lamellae counted from any 5 circumferential points per section was 3.8±0.1 in AA, 3.6±0.2 in AA-C, and 4.1±0.2 in AA-Cm. Breaks, defined as areas with broken elastin fibers, were quantified circumferentially as totals of 11.0±1.5 in AA, 12.4±2.2 in AA-C, and 7.9±1.4 in AA-Cm. While there was no significant difference, the AA-Cm group tended to have more intact elastic lamellae and fewer breaks compared with other groups.

Figure 4.

Histological analysis of elastin degradation in the aortic media. (A) Representative images (scale bar=100 μm). (B) Quantification of the medial elastin area, elastin gap area, number of elastic lamellae, and breaks. The AA-C group exhibited significant elastin degradation compared with the AA-Cm group. Data are means±SEM. **P<0.01 vs. AA group assessed using Tukey tests. AA, aortic aneurysm; AA group, AA-induced mice, treated with saline; AA-C group, AA- and COPD-induced mice, treated with saline; AA-Cm group, AA- and COPD-induced mice, treated with clarithromycin and montelukast.

Expression of Cytokines and Chemokines

Protein levels in AA walls were analyzed using ELISA. No significant differences in IL-1β, IL-6, or MCP-1 expression were observed between groups. In contrast, the AA-Cm group demonstrated a significantly lower TNF-α expression than AA-C (125.2±7.3 pg/mL in AA, 141.0±4.9 pg/mL in AA-C, 115.5±4.3 pg/mL in AA-Cm; P<0.05; Figure 5).

Figure 5.

Protein expression and matrix metalloproteinase (MMP) activities in aortic tissues. The AA-Cm group showed significantly lower tumor necrosis factor-α (TNF-α) levels and decreased MMP-9 activity compared with the AA-C group. Data are means±SEM. *P<0.05 and **P<0.01 using the Kruskal-Wallis test followed by multiple comparison with Bonferroni correction. AA, aortic aneurysm; AA group, AA-induced mice, treated with saline; AA-C group, AA- and COPD-induced mice, treated with saline; AA-Cm group, AA- and COPD-induced mice, treated with clarithromycin and montelukast.

MMP-2 and MMP-9 Enzymatic Activity

Active-MMP-9 activity was significantly lower in the AA-Cm group compared with the others (76.1±3.3 pg/mL in AA, 75.4±2.4 pg/mL in AA-C, 54.8±5.8 pg/mL in AA-Cm; P<0.01; Figure 5). There was no significant difference in active-MMP-2 activity between any of the groups.

Immunostaining of Macrophages

The infiltration of macrophages in the aortic wall was detected using immunofluorescent staining (Figure 6A). In Figure 6B, the AA-Cm group exhibited significantly lower infiltration of CD68 and iNOS double-positive macrophages (M1) in the adventitia and media of the aortic wall compared with the AA-C group (33.0±4.6 vs. 64.6±5.2%; P<0.01). In contrast, the AA-Cm group had a higher percentage of CD68 and CD206 double-positive macrophages (M2) compared with the AA group (55.3±8.1 vs. 24.7±2.2%; P<0.05). The M1/M2 ratio was lowest in the AA-Cm group (2.2±0.3 in AA, 2.3±0.4 in AA-C, 0.7±0.1 in AA-Cm; P<0.01).

Figure 6.

Immunofluorescence staining of macrophage phenotypes in aortic tissues. (A) Representative images. (B) Quantification of M1 (CD68⁺iNOS⁺) and M2 (CD68⁺CD206⁺) macrophages, and the M1/M2 ratio. The AA-Cm group exhibited a lower M1/M2 ratio compared with the other groups. Data are means±SEM. *P<0.05, **P<0.01 assessed using Tukey tests and Games-Howell tests. AA, aortic aneurysm; AA group, AA-induced mice, treated with saline; AA-C group, AA- and COPD-induced mice, treated with saline; AA-Cm group, AA- and COPD-induced mice, treated with clarithromycin and montelukast; iNOS, inducible nitric oxide synthase.

Discussion

The present study investigated the progression of AA due to the co-occurrence of COPD in a murine model and the mitigating efficacy of CAM and Mont on AA progression in COPD. The primary findings of this study can be summarized as follows. First, concomitant COPD accelerated aneurysm growth: mice in the AA-C group had significantly larger maximal aortic diameters than mice in the AA group on Days 14 and 28 (P<0.05). Administration of CAM and Mont might attenuate this COPD-related expansion by 0.351 mm (95% confidence interval −0.05, 0.75) in a linear mixed-effects model, but the difference did not reach conventional statistical significance (P=0.080). Second, histological analysis showed a significantly higher elastin content in AA-Cm aortas than those in the AA or AA-C groups (P<0.01). Third, molecular assays revealed lower TNF-α levels in AA-Cm vs. AA-C (P<0.05), and reduced active MMP-9 activity in AA-Cm vs. both AA and AA-C (P<0.01), indicating suppression of inflammation and matrix degradation. Fourth, immunofluorescence demonstrated decreased infiltration of M1 macrophages (vs. AA-C), increased infiltration of M2 macrophages (vs. AA), and a significantly lower M1/M2 ratio in AA-Cm compared with both the AA and AA-C groups (P<0.01), consistent with an anti-inflammatory shift in macrophage polarization. Collectively, these data confirm that COPD augments AA progression and suggest that dual therapy with CAM and Mont may partially mitigate this process through multi-faceted anti-inflammatory and matrixstabilizing mechanisms.

No studies have been reported on inducing AA in COPD model animals. This study is the first to investigate AA progression in the context of COPD and pharmacological interventions with CAM and Mont. The expansion of the AA diameter was induced by COPD, whereas the expression levels of TNF-α and active-MMP-9 did not differ significantly. This suggests that the mechanisms by which COPD becomes a risk factor for AA may not be directly related to elevated TNF-α expression or increased MMP-9 activity within aortic tissue. COPD may exacerbate AA progression through alternative pathways that were not measured in this study. One potential mechanism involves the systemic inflammatory milieu characteristic of COPD. COPD is associated with higher levels of inflammatory cytokines such as IL-8, GM-CSF and IL-17.¹⁰ Although we did not quantify them here, these mediators could still participate in aortic inflammation and remodeling.²¹ In addition, COPD-related reduction of TIMPs may also permit excess protease activity, potentially weakening the arterial wall.²²

As the other potential mechanism, oxidative stress is a hallmark of COPD²³ and may play a significant role in AA pathogenesis. The increased production of reactive oxygen species in COPD can induce oxidative damage to vascular endothelial cells and smooth muscle cells, leading to endothelial dysfunction and apoptosis of vascular smooth muscle cells.²⁴ This cellular damage compromises the integrity of the aortic wall and disrupts normal ECM maintenance. Oxidative stress can also activate redox-sensitive transcription factors, such as NF-κB and activator protein-1,²⁵ which upregulate the expression of pro-inflammatory genes and MMP-9. Additionally, COPD induces endothelial dysfunction by decreasing nitric oxide bioavailability, resulting in vasoconstriction and increased shear stress on the aortic wall.²⁶ This hemodynamic alteration may indirectly increase mechanical stress on the aorta. COPD is an independent risk factor for AA enlargement and rupture, implying that appropriate therapeutic interventions for COPD could potentially suppress AA progression.

The therapeutic effects observed in this study are consistent with the known pharmacological properties of CAM and Mont. In our previous study, the treatment of CAM or Mont reduced MMP-2 and MMP-9 in vitro and in vivo.¹⁵ MMP-9, which is secreted by macrophages, is known to be a key protease to degrade elastin and contributes to ECM breakdown and aneurysm expansion.⁶ Macrophage infiltration into the aneurysmal wall is an important contributor to the inflammatory environment.²⁷ Although the M1/M2 ratio is dominant in AA, the simultaneous administration of CAM and Mont to AA mice reduced MMP-9 expression and reduced the M1/M2 ratio, resulting in inhibited elastin degradation. This phenomenon was reflected in AA with COPD mice. A higher elastin content was shown in treatment with CAM and Mont, indicating that this therapeutic strategy mitigated ECM degradation. In contrast, it is known that M2 macrophages are involved in tissue repair and remodeling.²⁸ Notably, Mont not only suppresses inflammatory mediators but also inhibits AA development and expansion by inducing polarization towards M2 macrophages, as evidenced by increased IL-10 and arginase-1 expression at both gene and cell surface protein levels in Mont-treated macrophages.¹⁶

Hegazy et al. reported that co-administration of CAM and Mont led to a substantial increase (approximately 144%) in the plasma concentration of Mont.²⁹ This pharmacokinetic interaction, attributed to CAM’s inhibition of cytochrome P450 3A4, the enzyme responsible for metabolizing Mont, suggests the potential for synergistic therapeutic effects when these drugs are used simultaneously. We assumed that combining CAM and Mont may offer additive or synergistic effects by targeting multiple pathways implicated in AA pathogenesis in the presence of COPD. However, the results showed less pronounced effects than anticipated. The presence of COPD as an aggravating factor likely promoted systemic inflammation, counteracting some of the beneficial effects of the treatment. It seems that the simultaneous administration of CAM and Mont remained, primarily offsetting the negative impact of COPD rather than completely halting or reversing the process of AA progression.

Study Limitations

There are several limitations. To begin with, the small sample size and the 4-week follow up may have limited statistical power, potentially obscuring differences in final aortic diameter and rupture rate. In addition, the elastase-induced COPD model in ApoE-deficient mice may not fully replicate the chronic, smoking-related pathophysiology seen in human COPD and AAs.³⁰ Moreover, the lack of monotherapy control groups for CAM and Mont precludes determination of whether the observed effects are due to synergistic interactions or attributable to 1 agent alone. Another important consideration is that treatment with CAM and Mont was initiated at the same time as the induction of both aneurysm and COPD, which does not reflect typical clinical scenarios where therapeutic interventions are started only after disease progression has occurred. A further methodological gap is the absence of COPD-specific cytokine measurements and pulmonary function testing, leaving it unclear as to whether CAM and Mont specifically mitigate COPD-related aggravation or simply exert broad anti-inflammatory, anti-aneurysmal effects. Last, drug delivery by gastric gavage and the doses used may not mirror human oral administration.

To address these limitations comprehensively, future studies should use smoking-induced COPD models with larger cohorts and extended observation periods. Treatment initiation should be delayed to better reflect clinical timing, and monotherapy arms should be included to dissect individual drug effects. Moreover, the incorporation of COPD-specific cytokine panels, pulmonary function testing, and dose-response or pharmacokinetic studies following oral administration will be essential to more accurately evaluate both efficacy and mechanistic specificity, thereby enhancing translational relevance.

Conclusions

Using a murine model of AA with coexisting COPD, we investigated the impact of combination therapy with CAM and Mont on AA progression. In this model, CAM and Mont showed a trend toward attenuation of aneurysm expansion. Although the suppression of aortic diameter expansion did not reach statistical significance, the absence of rupture events and favorable histological findings, such as preserved elastic fibers and decreased inflammatory responses, suggest potential therapeutic benefits. These observations imply that the anti-inflammatory properties of CAM and Mont may help counteract some of the detrimental processes associated with COPD-related exacerbation of aneurysmal disease. Future studies should aim to elucidate the underlying molecular mechanisms, optimize dosing strategies, and evaluate the translational potential of this therapy in clinical settings.

Acknowledgments

The authors thank the Division for Experimental Animals, Nagoya University Graduate School of Medicine for managing the animals used in this study. The authors also thank the Division for Medical Research Engineering, Nagoya University Graduate School of Medicine, for the use of a cryomicrotome (Leica Biosystems, Nußloch, Germany). During the preparation of this manuscript, the authors used DeepL translator (DeepL SE, Cologne, North Rhine-Westphalia, Germany) in order to improve language and readability.

Sources of Funding

This work was supported by the Japan Society for the Promotion of Science (JSPS) KAKENHI (Grant no. 19K09265).

Disclosures

The authors declare no conflicts of interest.

Author Contributions

S.Y. and Y.N. conceived of and designed the study. S.Y. and A.Y.-O. collected and performed the data analysis and interpretation. S.Y. wrote the article. Y.N. performed critical revisions of the article, and Y.N. and M.M. approved the article. Y.N. obtained funding. Y.N. has overall responsibility.

IRB Information

Not applicable.

Data Availability

The data will be shared on a request basis. Please contact the corresponding author directly to request data sharing.

References

• 1.   Tyrovolas S, Tyrovola D, Giné-Vázquez I, Koyanagi A, Bernabe-Ortiz A, Rodriguez-Artalejo F, et al. Global, regional, and national burden of aortic aneurysm, 1990–2017: A systematic analysis of the Global Burden of Disease Study 2017. Eur J Prev Cardiol 2022; 29: 1220–1232, doi:10.1093/eurjpc/zwab015.
• 2.   Akutsu K, Yoshino H, Shimokawa T, Ogino H, Kunihara T, Takahashi T, et al. Clinical features of 544 patients with ruptured aortic aneurysm: A report from the Tokyo Acute Aortic Super Network Database. Circ J 2024; 88: 1664–1671, doi:10.1253/circj.cj-23-0636.
• 3.   Mitsui T, Bando YK, Hirakawa A, Furusawa K, Morimoto R, Taguchi E, et al. Role of common antihypertensives in the growth of abdominal aortic aneurysm at the presurgical stage. Circ Rep 2023; 5: 405–414, doi:10.1253/circrep.cr-23-0071.
• 4.   Shimizu K, Mitchell RN, Libby P. Inflammation and cellular immune responses in abdominal aortic aneurysms. Arterioscler Thromb Vasc Biol 2006; 26: 987–994, doi:10.1161/01.ATV.0000214999.12921.4f.
• 5.   Rizas KD, Ippagunta N, Tilson MD 3rd. Immune cells and molecular mediators in the pathogenesis of the abdominal aortic aneurysm. Cardiol Rev 2009; 17: 201–210, doi:10.1097/CRD.0b013e3181b04698.
• 6.   Thompson RW, Holmes DR, Mertens RA, Liao S, Botney MD, Mecham RP, et al. Production and localization of 92-kilodalton gelatinase in abdominal aortic aneurysms: An elastolytic metalloproteinase expressed by aneurysm-infiltrating macrophages. J Clin Invest 1995; 96: 318–326, doi:10.1172/jci118037.
• 7.   Kurosawa K, Matsumura JS, Yamanouchi D. Current status of medical treatment for abdominal aortic aneurysm. Circ J 2013; 77: 2860–2866, doi:10.1253/circj.CJ-13-1252.
• 8.   Guo DC, Papke CL, He R, Milewicz DM. Pathogenesis of thoracic and abdominal aortic aneurysms. Ann N Y Acad Sci 2006; 1085: 339–352, doi:10.1196/annals.1383.013.
• 9.   Timmins SC, Diba C, Farrow CE, Schoeffel RE, Berend N, Salome CM, et al. The relationship between airflow obstruction, emphysema extent, and small airways function in COPD. Chest 2012; 142: 312–319, doi:10.1378/chest.11-2169.
• 10.   Barnes PJ. Inflammatory mechanisms in patients with chronic obstructive pulmonary disease. J Allergy Clin Immunol 2016; 138: 16–27, doi:10.1016/j.jaci.2016.05.011.
• 11.   Gan WQ, Man SF, Senthilselvan A, Sin DD. Association between chronic obstructive pulmonary disease and systemic inflammation: A systematic review and a meta-analysis. Thorax 2004; 59: 574–580, doi:10.1136/thx.2003.019588.
• 12.   Meijer CA, Kokje VBC, Van Tongeren RBM, Hamming JF, Van Bockel JH, Möller GM, et al. An association between chronic obstructive pulmonary disease and abdominal aortic aneurysm beyond smoking. Eur J Vasc and Endovasc Surg 2012; 44: 153–157, doi:10.1016/j.ejvs.2012.05.016.
• 13.   Rubinstein I, Kumar B, Schriever C. Long-term montelukast therapy in moderate to severe COPD: A preliminary observation. Respir Med 2004; 98: 134–138, doi:10.1016/j.rmed.2003.09.003.
• 14.   Simoens S, Laekeman G, Decramer M. Preventing COPD exacerbations with macrolides: A review and budget impact analysis. Respir Med 2013; 107: 637–648, doi:10.1016/j.rmed.2012.12.019.
• 15.   Uchida W, Narita Y, Yamawaki-Ogata A, Tokuda Y, Mutsuga M, Lee Fujimoto K, et al. The oral administration of clarithromycin prevents the progression and rupture of aortic aneurysm. J Vasc Surg 2018; 68: 82S–92S.e82, doi:10.1016/j.jvs.2017.12.047.
• 16.   Kawai Y, Narita Y, Yamawaki-Ogata A, Usui A, Komori K. Montelukast, a cysteinyl leukotriene receptor 1 antagonist, induces M2 macrophage polarization and inhibits murine aortic aneurysm formation. Biomed Res Int 2019; 2019: 9104680, doi:10.1155/2019/9104680.
• 17.   Percie du Sert N, Hurst V, Ahluwalia A, Alam S, Avey MT, Baker M, et al. The ARRIVE guidelines 2.0: Updated guidelines for reporting animal research. BMC Vet Res 2020; 16: 242, doi:10.1186/s12917-020-02451-y.
• 18.   National Research Council Committee for the Update of the Guide for the C, Use of Laboratory A. The National Academies Collection: Reports funded by National Institutes of Health. In: Guide for the Care and Use of Laboratory Animals. Washington (DC): National Academies Press (US).
• 19.   Vidal D, Fortunato G, Klein W, Cortizo L, Vasconcelos J, Ribeiro-Dos-Santos R, et al. Alterations in pulmonary structure by elastase administration in a model of emphysema in mice is associated with functional disturbances. Rev Port Pneumol 2012; 18: 128–136, doi:10.1016/j.rppneu.2011.12.007.
• 20.   Yamawaki-Ogata A, Fu X, Hashizume R, Fujimoto KL, Araki Y, Oshima H, et al. Therapeutic potential of bone marrow-derived mesenchymal stem cells in formed aortic aneurysms of a mouse model. Eur J Cardiothorac Surg 2014; 45: e156–e165, doi:10.1093/ejcts/ezu018.
• 21.   Miossec P, Korn T, Kuchroo VK. Interleukin-17 and type 17 helper T cells. N Engl J Med 2009; 361: 888–898, doi:10.1056/NEJMra0707449.
• 22.   Mercer PF, Shute JK, Bhowmik A, Donaldson GC, Wedzicha JA, Warner JA. MMP-9, TIMP-1 and inflammatory cells in sputum from COPD patients during exacerbation. Respir Res 2005; 6: 151, doi:10.1186/1465-9921-6-151.
• 23.   McGuinness AJ, Sapey E. Oxidative stress in COPD: Sources, markers, and potential mechanisms. J Clin Med 2017; 6: 21, doi:10.3390/jcm6020021.
• 24.   MacNee W. Pulmonary and systemic oxidant/antioxidant imbalance in chronic obstructive pulmonary disease. Proc Am Thorac Soc 2005; 2: 50–60, doi:10.1513/pats.200411-056SF.
• 25.   Meyer M, Schreck R, Baeuerle PA. H2O2 and antioxidants have opposite effects on activation of NF-kappa B and AP-1 in intact cells: AP-1 as secondary antioxidant-responsive factor. Embo J 1993; 12: 2005–2015, doi:10.1002/j.1460-2075.1993.tb05850.x.
• 26.   Maclay JD, McAllister DA, Mills NL, Paterson FP, Ludlam CA, Drost EM, et al. Vascular dysfunction in chronic obstructive pulmonary disease. Am J Respir Crit Care Med 2009; 180: 513–520, doi:10.1164/rccm.200903-0414OC.
• 27.   Raffort J, Lareyre F, Clément M, Hassen-Khodja R, Chinetti G, Mallat Z. Monocytes and macrophages in abdominal aortic aneurysm. Nat Rev Cardiol 2017; 14: 457–471, doi:10.1038/nrcardio.2017.52.
• 28.   Murray PJ, Wynn TA. Protective and pathogenic functions of macrophage subsets. Nature Reviews Immunology 2011; 11: 723–737, doi:10.1038/nri3073.
• 29.   Hegazy SK, Mabrouk MM, Elsisi AE, Mansour NO. Effect of clarithromycin and fluconazole on the pharmacokinetics of montelukast in human volunteers. Eur J Clin Pharmacol 2012; 68: 1275–1280, doi:10.1007/s00228-012-1239-0.
• 30.   Chow L, Smith D, Chokshi K, Ezegbunam W, Charoenpong P, Foley K, et al. Animal models of chronic obstructive pulmonary disease. InTech 2017, doi:10.5772/intechopen.70262.