Abstract

Objective: The role of chronic inflammation in abdominal aortic aneurysm (AAA) is controversial. CD11c+ antigen-presenting cells (APCs) (dendritic cells (DCs)) have been reported in human AAA samples but their role is unclear. The effect of conditional depletion of CD11c+ cells on experimental AAA was investigated in the angiotensin II (AngII)-infused apolipoprotein E-deficient (ApoE–/–) mouse model.

Approach: CD11c-diphtheria toxin (DT or D.tox) receptor (DTR), ovalbumin (OVA) fragment aa 140–386, and enhanced green fluorescent protein (eGFP)-ApoE–/– (CD11c.DOG.ApoE–/–) mice were generated and CD11c+ cell depletion achieved with D.tox injections (8 ng/g body weight, i.p., every-other-day). AAA formation and growth were assessed by measurement of supra-renal aortic (SRA) diameter in vivo by serial ultrasound and by morphometry assessment of harvested aortas at the end of the study.

Results: Depletion of CD11c+ cells by administration of D.tox on alternative days was shown to reduce the maximum diameter of AAAs induced by 28 days AngII infusion compared with controls (D.tox, 1.58 ± 0.03 mm vs Vehicle control, 1.81 ± 0.06 mm, P<0.001). CD11c+ depletion commencing after AAA establishment by 14 days of AngII infusion, was also shown to lead to smaller AAAs than controls after a further 14 days (D.tox, 1.54 ± 0.04 mm vs Vehicle control, 1.80 ± 0.03 mm, P<0.001). Flow cytometry revealed significantly lower numbers of circulating CD44hi CD62Llo effector CD4 T cells, CD44hi CD62Llo effector CD8 T cells and B220+ B cells in CD11c+ cell-depleted mice versus controls. CD11c+ depletion attenuated SRA matrix degradation indicated by decreased neutrophil elastase activity (P=0.014), lower elastin degradation score (P=0.012) and higher collagen content (P=0.002).

Conclusion: CD11c+ cell-depletion inhibited experimental AAA development and growth associated with down-regulation of circulating effector T cells and attenuated matrix degradation. The findings suggest involvement of autoreactive immune cells in AAA pathogenesis.

Introduction

Abdominal aortic aneurysm (AAA) is an important cause of sudden death in older adults [1]. It was estimated that AAA rupture was responsible for 166000 deaths worldwide in 2016 [2]. Currently the only treatment available to avoid AAA rupture is prophylactic surgery but because of the absence of widespread availability of specialized vascular services and the perioperative complications and poor long-term durability of repair, there is substantial interest in the development of alternative therapies [3,4]. Inflammation is a consistent finding in human and experimental AAA, but whether it is a cause or consequence is controversial [3,4]. Findings from a human imaging study suggest that increased aortic inflammation predicts more rapid AAA growth and increased risk of rupture [5]. In contrast, clinical trials using antibiotics reported to reduce aortic inflammation have not shown a consistent reduction in AAA growth [6]. It seems likely though that more insights into the most appropriate inflammatory targets for an AAA drug are needed.

Examination of human AAA tissue shows classical features of chronic inflammation, including infiltration of neutrophils, macrophages, NK cells, NKT cells, T and B cells within the adventitia and media, and up-regulation of genes related to immunity including cytokine–cytokine receptor interaction, chemokine signaling pathway and antigen processing and presentation [7,8]. The ratio of CD4/CD8 cells has been reported to be up to four-folds higher in AAA tissue than in normal peripheral blood [9]. Oligoclonal T-cell infiltrates have also been identified in human AAA samples [10]. The presence of follicles containing inflammatory cells in human AAA suggests a cell-mediated antigen response [11] and IgG antibody purified from human AAA has been reported to be immune-reactive with normal aortic proteins [12]. Clonally expanded CD4+ T cells have been reported to be present in the peri-adventitial vascular-associated lymphatic tissue, suggesting that AAA may have an autoimmune component [13].

Dendritic cells (DCs) are antigen-presenting cells (APCs) that play a key role in induction of the innate immune response and are involved in maintaining immune tolerance to self-antigens [14]. Furthermore, DCs serve as a link between the innate and adaptive immune systems [15,16]. DCs are activated by endogenous or exogenous antigens and initiate the release of cytokines that activate subsequent immune responses [15,17]. In human AAA samples, DCs have been reported to be in contact with T cells along the periphery of lymphoid follicular aggregations within the adventitia, suggesting their role in antigen presentation [18]. Since DCs play a role in inducing tolerance against self-antigens, modulation of DC function could theoretically reduce aortic inflammation and remodeling.

In the present study, we hypothesized that depleting CD11c+ DCs would limit aortic remodeling and thereby reduce the severity (size) of experimental AAA. We investigated the effect of conditional depletion of CD11c+ cells on experimental AAA formation and growth in a commonly used mouse model of AAA.

Materials and methods

Mice

Approval for the animal studies was obtained from the local ethics committee and experimental work was performed in accordance with the institutional and ethical guidelines of James Cook University, Australia (AEC approval A1752), and conformed to the Guide for the Care and Use of Laboratory Animals (National Institutes of Health, U.S.A.), the Australian Code of Practice for the Care and Use of Animals for Scientific Purpose (7th Edition, 2004) and the ARRIVE criteria set by The National Centre for the Replacement, Refinement and Reduction of Animals in Research (London, U.K.). Mice were housed in an individually ventilated, temperature/humidity-controlled cage system (Aero IVC Green Line; Tecniplast) on a 12-h light/dark cycle, and maintained on normal laboratory chow and water ad libitum. All experiments took place in the Small Animal House (Building 86) at the James Cook University.

Generation of CD11c.DOG.apolipoprotein E-deficient mice

We utilized the transgenic CD11c.DTR mouse model which has previously been shown to allow effective depletion of DC over long period of time without non-specific cytotoxicity [19]. Breeding pairs of CD11c.DOG (DTR-OVA-eGFP) mice which carry the human Diphtheria toxin receptor (DTR), ovalbumin fragment aa 140–386, and enhanced green fluorescent protein (eGFP) under the control of the CD11c promoter have been previously described [19]. CD11c.DOG mice were mated with apolipoprotein E-deficient (ApoE–/–) mice [B6.129P2-Apoe tm1Unc/Arc (N10)]; sourced from the Animal Resources Centre, Canning Vale, Australia, to generate CD11c.DOG.ApoE–/–/(OVA–/–) mice. Both CD11c.DOG mice and ApoE–/– mice belonged to the same background strain C57BL/6J mice (B6). Genotyping was carried out by polymerase chain reaction (Applied Biosystems Veriti Thermal Cycler) from genomic DNA of tail biopsies using the following primers: 5′-AACCTGTGCAGATGATGTACCA-3′ and 5′-GCGATGTGCTTGATACAGAAGA-3′ for ovalbumin (OVA), and 5′-CGAAGCCAGCTTGAGTTACAGAA-3′ and 5′-AGAGCCGGAGGTGACAGATCAG-3′ for ApoE-null allele. Five microliters of the PCR products were loaded on a 1% agarose gel pre-stained with Gel-Red (Biotium, CA) and electrophoresis performed for 45 min at 100 V. PCR products were visualized by ultraviolet transillumination to confirm the genotype (Supplementary Figure S1).

Mouse model of AAA and in vivo studies

Male, 10–12 week old mice were randomized to experiments outlined below (using random number generators) after recording their baseline body weight. Transient depletion of DCs (CD11chigh MHC Class II+ cells) in experimental mice was achieved with administration of Diphtheria toxin (D.tox; 8 ng/g body weight, List Biological Laboratories Inc. CA). Proportions and absolute numbers of CD11c+ cells in the blood, spleen, thymus, lymph node and bone marrow were assessed after 24 h of D.tox injection to confirm the depletion of DC. For continuous, longer-term DC depletion, multiple D.tox i.p. injections were performed (8 ng/g body weight) on alternate days over the study period [19].

The angiotensin II (AngII)-infusion model was implemented as previously described [20]. Briefly, an osmotic micro-pump (ALZET Model 1004, Durect Corporation, U.S.A.) containing AngII (Sigma–Aldrich) dissolved in sterile water was inserted into the subcutaneous space left of the dorsal midline under anesthesia (4% isoflurane inhalation) to administer AngII at a rate of 1.0 µg/kg/min over the experimental period. Two separate animal studies were performed (Figure 1).

Schematic representation of the study design

Figure 1
Schematic representation of the study design

 

Figure 1
Schematic representation of the study design

 

AAA formation study

AngII was infused into CD11c.DOG.ApoE–/– mice (average age: 6 months) which were either administered D.tox (n=20) or vehicle control (VC, n=15) commencing at the same time as the AngII infusion which was continued for 28 days.

AAA progression study

Six-month-old male CD11c.DOG.ApoE–/– mice (n=41) were infused with AngII for 28 days. AAAs were established over 14 days during which eight mice died of aortic rupture. Thirty-three mice that remained at day 14 were allocated into two groups of similar mean suprarenal aorta (SRA) diameters based on ultrasound measurements. Group 1 (n=17) was administered D.tox and group 2 (n=16) was injected with the vehicle.

Outcome assessors were blinded to treatment groups at the time of measurement for all end point analyses. At the end of the experiments, mice were killed with carbon dioxide asphyxiation.

Assessment of AAA severity

The primary outcome of both studies was AAA severity assessed by the maximum diameter of the SRA measured from ultrasound images and morphometry using previously described reproducible protocols [21–23]. In brief, aortic diameters were measured by both ultrasound in vivo and by morphometric analysis of harvested aortas. Ultrasound measurements of the SRA were obtained prior to starting the AngII infusion and then every 1–2 weeks during the studies. Ultrasound scans were performed on anesthetized mice (i.p., 40 mg/kg ketamine, 4 mg/kg xylazine) using a MyLab™ 70 VETXV platform (Esaote, Italy) with a 40-mm linear transducer at an operating frequency of 10 MHz (LA435; Esaote, Italy) to provide a sagittal image of the aortic segment assessed. The maximum external SRA diameter was measured at peak systole using the caliper measurement feature as previously reported [24,25].

Ex vivo morphometry measurements were performed following sudden death or at day 28 after the mice were killed by CO2 asphyxiation. Aortas were perfused by PBS under constant physiological pressure. Perfused and harvested aortas (arch to iliac bifurcation) were placed on a graduated template and digitally photographed (Coolpix 4500, Nikon). Maximum diameters of the arch, thoracic, SRA and infra-renal aorta (IRA) were determined from the images using computer-aided analysis (Adobe Photoshop CS5 Extended version 12, Adobe Systems Incorporated). We have previously established that both ultrasound and ex vivo morphometric measurements can be repeated with good inter-observer reproducibility [20,22,24–26].

Sample preparation and flow cytometry

Single cell suspensions were prepared from thymus by gently grinding the thymus between frosted microscope slides. Thymocytes were released into ice-chilled MACS buffer [PBS containing 2 mM EDTA (Amresco) and 0.5% (w/v) BSA (ICN Biomedicals), adjusted to a pH of 7.4]. Splenocytes were prepared by disrupting the spleen using a 26-gauge needle and forceps in MACS buffer, and the resulting cell suspension was treated with RBC lysing buffer (BioLegend). Lymphocytes were released from the lymph nodes by carefully tearing open the capsule using two 26-gauge needles and then gently grinding the capsule between two frosted microscope slides in ice-cold MACS buffer. Suspensions were filtered through 60-μm nylon mesh. Heparinized blood samples were depleted of red cells by lysis buffer (BioLegend). Bone marrow was flushed from femoral bones using Hank’s balanced salt solution (Sigma–Aldrich) containing 2% FBS and 1 mM EDTA (Amresco) through a 23-gauge needle-fitted syringe. Cell suspensions were filtered through 60-μm nylon mesh.

Flow cytometry analysis

Flow cytometry was performed using surface markers to define target cell types as summarized in Supplementary Table S1. Antibodies were diluted in PBS containing 2 mM EDTA (Amresco) and 0.5% (w/v) BSA (ICN Biomedicals). Cells were pre-incubated for 20 min with anti-CD16/32 to prevent FcR binding before addition of surface staining antibody cocktails. Isotype-matched antibodies of irrelevant specificity were used to determine the level of non-specific staining and frequency of cells stained with test antibodies. A minimum of 50000 cells were analyzed for each sample, and a proportion of up to 1% false-positive events was accepted in the isotype control samples. Internal bead standards along with the cells of interest were simultaneously acquired directly from unmanipulated whole blood specimens. Viable cells were identified by forward/side scatter profile and in some cases by propidium iodide (PI) exclusion. A forward scatter-area against forward scatter-linear gate was used to exclude doublets from analyses. Absolute cell numbers were determined and the calculations based on mononuclear cell percentages were reported.

Plasma analysis

Blood was collected at different time points to separate platelet-poor plasma [24]. Platelet-poor plasma was separated by centrifugation of blood at 2000×g at 4°C for 10 min followed by further 10-min centrifugation at 15000×g at 4°C. The plasma samples were snap-frozen in liquid nitrogen and stored at −80°C for subsequent assessments.

Neutrophil elastase activity was assessed in plasma using a commercially available assay (Abcam) in accordance with the manufacturers’ instructions. The fluorescence was measured by using fluorescence microplate reader (PolarStar Omega, Ex/Em = 380/500 nm) in kinetic mode and then choosing two time points during the linear range and expressed as relative fluorescence unit (RFU).

Measurement of total cholesterol, high-density lipoprotein (HDL), low-density/very low-density lipoprotein (LDL/VLDL) was performed using a commercially available fluorescence assay (Abcam) in accordance with the manufacturers’ instructions.

Quantification of atherosclerotic lesion area

The severity of atherosclerosis was determined by Sudan IV staining as described previously [24]. Briefly, the left and right carotid arteries and aortic arch were dissected, excised, opened longitudinally and pinned down on a wax coated Petridish. Tissue samples were transferred to a 70% ethanol solution and stained with 0.1% Sudan IV dissolved in equal parts of acetone and 70% ethanol for 10 min in order to identify areas of atherosclerosis. Background staining was washed off by 10-min incubation in 70% ethanol. Samples were rehydrated by brief placement in H2O and digitally photographed (Coolpix 4500, Nikon). Sudan IV stained areas were quantified using Adobe Photoshop software (version CS5). The atherosclerotic lesion area was quantified and expressed as a percentage of the total luminal surface area as previously described [27].

Histological assessments

SRA segments from six mice from each group were selected using a random number generator. A total of 6 μM serial cryostat sections were cut from each SRA and processed for histopathological evaluations. Adjacent sections were stained with Hematoxylin and Eosin (H&E, Polysciences, Inc), Verhoeff-van Giesson (EVG) staining (Polysciences, Inc) and the PicroSirius Red procedure (Polysciences, Inc), as previously described [21,28]. Stained sections were photographed using a Nikon Eclipse 50i microscope fitted with a CCD Camera (DSFi1) and digital images captured to a PC supported with image analysis software (NIS Elements, version F2.30). Qualitative evaluation of elastin fiber integrity was performed on digital images by semi-quantitative grading by a blinded observer as follows: 1, no elastin degradation or mild degradation; 2, moderate; 3, moderate to severe and 4, severe elastin degradation following previously published protocols [22,29]. The percentage of the field area that stained positive for PicroSirius Red was determined from analysis of four to five fields for at least three sections per mouse for each group (n=6), and the mean value was calculated for each region. All histological assessments were performed by a blinded observer and we have previously reported that all these histological measurements can be repeated with good inter-observer reproducibility [22].

Statistical analysis

Data were analyzed using GraphPad Prism (version 7) and TIBCO Spotfire S+ (version 8.2). D’Agostino and Pearson’s tests were used to test normality of the data. Parametric or non-parametric tests were applied appropriate to data distribution. Data are reported as the median and interquartile range with maximum and minimum data points unless otherwise stated. Student’s t or Mann–Whitney test were performed for two group comparisons of parametric and non-parametric data, respectively. Data obtained as a function of time were compared within each group by two-way repeated measures analysis of variance (ANOVA) followed by Sidak’s multiple comparisons test and liner mixed effect analysis. In all cases, P-values less than 0.05 were considered significant.

Results

CD11c+ cell depletion

A single D.tox injection caused significant systemic depletion in the proportions and numbers of CD11c+ cells in the peripheral blood, spleen, thymus, lymph nodes and bone marrow of CD11c.DOG.ApoE–/– mice compared with the ApoE–/– mice after 24 h (Figure 2 and Supplementary Table S2). It has previously been described that the population of CD11c+ cells completely recover in CD11c.DOG mice 72 h after a single injection of DT [19]. Hence, for long-term depletion, D.tox injection was given on alterative days throughout the experimental period of both studies.

Proportions of CD11chi cells in the spleen, peripheral blood, thymus, lymph nodes and bone marrow samples of wild-type (clear) and CD11c.DOG.ApoE–/– mice (dark) 24 h after D.tox injection (n=15/group)

Figure 2
Proportions of CD11chi cells in the spleen, peripheral blood, thymus, lymph nodes and bone marrow samples of wild-type (clear) and CD11c.DOG.ApoE–/– mice (dark) 24 h after D.tox injection (n=15/group)

 

Figure 2
Proportions of CD11chi cells in the spleen, peripheral blood, thymus, lymph nodes and bone marrow samples of wild-type (clear) and CD11c.DOG.ApoE–/– mice (dark) 24 h after D.tox injection (n=15/group)

 

Depletion of CD11c+ cells reduced SRA expansion in response to AngII infusion

The effect of depletion of CD11c+ cells on AAA formation was assessed. AngII infusion led to an increase in maximum SRA diameter over time. SRA rupture occurred in five mice (vehicle, n=2; D.tox, n=3). Periodic ultrasound monitoring demonstrated that D.tox-mediated depletion of CD11c+ cells significantly inhibited SRA dilatation over the AngII infusion period (Figure 3A). Maximum SRA diameter of mice in the two groups were comparable prior to AngII-infusion (baseline) but significantly smaller in the D.tox group compared with vehicle controls at 14 and 28 days (P<0.001; Figure 3A). Ex vivo measurement of harvested aortas from mice surviving the 28-day AngII infusion confirmed smaller SRA diameter in CD11c+ depleted mice compared with controls (P=0.033; Figure 3B and Supplementary Figure S2A). Maximum diameter of the aortic arch was also smaller in mice with CD11c+ cell depletion compared with controls (P=0.040; Figure 3B).

CD11c+ cell depletion limited AAA induction in AngII-infused ApoE–/– mice

Figure 3
CD11c+ cell depletion limited AAA induction in AngII-infused ApoE–/– mice

(A) Effects of CD11c+ cell depletion on maximum SRA diameter in AngII-infused ApoE–/– mice. Temporal changes in the in vivo dilatation of SRA monitored fortnightly by ultrasound over a 4-week period of AngII infusion. Statistical comparisons of vehicle (n=15, clear) vs D.tox (n=20, dark) administrations were done by linear mixed effect analysis. (B) Maximum diameters of the aortic arch, thoracic aorta, SRA and IRA assessed by ex vivo morphometry. Data are expressed as median and interquartile range with maximum and minimum data points (whiskers). P-values by Mann–Whitney U test. (C) In vivo maximum diameter of the SRA was monitored weekly by ultrasound. Statistical comparisons between vehicle (n=16, clear) vs D.tox (n=17, dark) administrations were calculated with two-way repeated measures ANOVA using Dunn’s multiple comparisons test. (D) Maximum diameters of the aortic arch, thoracic aorta, SRA and IRA assessed by ex vivo morphometry. Data expressed as median and interquartile range with maximum and minimum data points (whiskers). P-values by Mann–Whitney U test. Abbreviations: SRA, supra-renal aorta; TA, thoracic aorta.

Figure 3
CD11c+ cell depletion limited AAA induction in AngII-infused ApoE–/– mice

(A) Effects of CD11c+ cell depletion on maximum SRA diameter in AngII-infused ApoE–/– mice. Temporal changes in the in vivo dilatation of SRA monitored fortnightly by ultrasound over a 4-week period of AngII infusion. Statistical comparisons of vehicle (n=15, clear) vs D.tox (n=20, dark) administrations were done by linear mixed effect analysis. (B) Maximum diameters of the aortic arch, thoracic aorta, SRA and IRA assessed by ex vivo morphometry. Data are expressed as median and interquartile range with maximum and minimum data points (whiskers). P-values by Mann–Whitney U test. (C) In vivo maximum diameter of the SRA was monitored weekly by ultrasound. Statistical comparisons between vehicle (n=16, clear) vs D.tox (n=17, dark) administrations were calculated with two-way repeated measures ANOVA using Dunn’s multiple comparisons test. (D) Maximum diameters of the aortic arch, thoracic aorta, SRA and IRA assessed by ex vivo morphometry. Data expressed as median and interquartile range with maximum and minimum data points (whiskers). P-values by Mann–Whitney U test. Abbreviations: SRA, supra-renal aorta; TA, thoracic aorta.

Depletion of CD11c+ cells limited growth of established AAAs

The effect of CD11c+ cell depletion on growth of established aortic aneurysms was assessed. AngII infusion for 2 weeks led to fatal aortic rupture in 8 of the 44 mice (18.2%) within 14 days, with all ruptures occurring at the SRA region, as previously described [30]. The remaining 33 mice were randomly allocated to vehicle control (n=16) or D.tox (n=17) administration, stratified by maximum SRA diameter so that both groups had similar mean SRA diameter at the commencement of the intervention. Continued AngII infusion resulted in aortic expansion in both groups.

Mice receiving D.tox had a significant reduction in SRA diameter as assessed by ultrasound (Figure 3C). Ex vivo measurements of harvested aortas from surviving mice confirmed this observation, with maximum diameter of the SRA (P=0.002), as well as the aortic arch (P=0.041), thoracic aorta (P=0.004) and IRA (P=0.008) significantly smaller in mice administered D.tox compared with vehicle (Figure 3D and Supplementary Figure S2B).

Depletion of CD11c+ cells reduced CD4 and CD8 T cells in the circulation

As depletion of CD11c+ cells led to reduced AAA diameter, we quantified circulating immune cells (Figure 4A). Blood collected by cardiac puncture after AngII infusion for 28 days was depleted of red cells by lysis buffer, stained with fluorescent-conjugated antibodies, and counting beads added before running on a flow cytometer. The gating strategy used for phenotyping immune cells is shown in Supplementary Figure S3. Flow cytometric analysis revealed lower proportion of T cells, especially the CD44hi CD62Llo effector CD4 and CD8 T cell subset, in the D.tox than the control group (Figure 4A,B). Circulating B-cell proportions were also found to be significantly lower in the D.tox compared with the control group (Supplementary Figure S4). Spleens were harvested and splenocytes stained with fluorescent-conjugated antibodies for flow cytometry. Lower proportion of CD4 T cells and B220+ B cells were observed in the D.tox compared with the control group (Supplementary Figure S5A,B).

CD11c+ cell depletion resulted in reduced proportions of immune cells in the circulation

Figure 4
CD11c+ cell depletion resulted in reduced proportions of immune cells in the circulation

(A,B) Lower proportions of effector CD4 and CD8 T cells were observed in CD11c+ cell depleted than control mice. Data are shown as median and interquartile ranges with maximum and minimum data points (whiskers). P-values by Mann–Whitney U test. (C) Neutrophil elastase activity in control and CD11c+ cell depleted/D.tox groups in both AAA initiation and AAA-progression studies. P-values by t test. Abbreviations: R.F.U., relative fluorescent unit; VC, vehicle control.

Figure 4
CD11c+ cell depletion resulted in reduced proportions of immune cells in the circulation

(A,B) Lower proportions of effector CD4 and CD8 T cells were observed in CD11c+ cell depleted than control mice. Data are shown as median and interquartile ranges with maximum and minimum data points (whiskers). P-values by Mann–Whitney U test. (C) Neutrophil elastase activity in control and CD11c+ cell depleted/D.tox groups in both AAA initiation and AAA-progression studies. P-values by t test. Abbreviations: R.F.U., relative fluorescent unit; VC, vehicle control.

Depletion of CD11c+ cells decreased plasma neutrophil elastase activity in mice with established AAA

Neutrophil elastase activity measured in platelet-poor plasma from mice in the AAA formation study was similar in mice receiving D.tox compared with vehicle control (Figure 4C). In contrast, in mice with established AAA, depletion of CD11c+ cells led to significantly lower plasma neutrophil elastase levels compared with vehicle controls (Figure 4C; P=0.014).

Depletion of CD11c+ cells did not affect cholesterol level or aortic atherosclerosis

Plasma cholesterol and aortic atherosclerosis were assessed in the aneurysm progression study. At the end of study period, total plasma cholesterol concentrations were comparable in mice receiving vehicle control or D.tox (P=0.460, Supplementary Figure S5A). Similarly, plasma HDL (P=0.569) concentrations and VLDL (P=0.185) were similar in mice receiving vehicle control or D.tox (Supplementary Figure S6A). Atherosclerosis plaque within the aortic arch was assessed by en face Sudan IV staining. No significant difference in mean staining area was demonstrated between the vehicle control and the D.tox groups (P=0.681; Supplementary Figure S6B).

Inhibition of AAA progression by CD11c+ cell depletion was associated with reduced aortic matrix remodeling

Histological examination of SRA sections suggested that CD11c+ depletion resulted in marked decrease in medial inflammation, adventitial atrophy and reduced expansion of luminal and outer wall diameters of the SRA (Figure 5A). EVG staining of SRA demonstrated disruption of elastic lamina in both experimental and control groups; however, the grade of elastin breaks was observed to be higher in the control group (P=0.012, Figure 5B). Assessment of PicroSirius Red staining for collagen in SRA sections demonstrated significantly lower aortic wall collagen content in the control group compared with the D.tox group (P=0.002, Figure 5C).

Depletion of CD11c+ cells limited matrix degradation in ApoE–/– mice

Figure 5
Depletion of CD11c+ cells limited matrix degradation in ApoE–/– mice

(A) Representative H&E image of AngII-infused mice showing inflammatory cell infiltration and intra-mural thrombus formation. Scale bar: 500 µm (*Lumen). (B) Comparison of medial elastin filament breaks (black structures) within EVG stained sections of suprarenal aortas. Scale bar: 500 µm (*Lumen). (C) Polarization microscopy images of PicroSirius Red staining for collagen content within the suprarenal aorta. Scale bar: 500 µm (*Lumen). (D) Quantification graph showing elastin filament degradation (n=6 aorta/group). Aortic wall elastin filament degradation was graded based on the degree of breaks in elastin filaments (graded on a scale of 1–4) as described in the ‘Materials and methods’ section. (E) Quantification of polarization images for collagen content expressed as a percentage (%) of the total suprarenal aorta section area (n=6 aorta/group). Data are shown as median and interquartile ranges. Abbreviation: VC, vehicle control.

Figure 5
Depletion of CD11c+ cells limited matrix degradation in ApoE–/– mice

(A) Representative H&E image of AngII-infused mice showing inflammatory cell infiltration and intra-mural thrombus formation. Scale bar: 500 µm (*Lumen). (B) Comparison of medial elastin filament breaks (black structures) within EVG stained sections of suprarenal aortas. Scale bar: 500 µm (*Lumen). (C) Polarization microscopy images of PicroSirius Red staining for collagen content within the suprarenal aorta. Scale bar: 500 µm (*Lumen). (D) Quantification graph showing elastin filament degradation (n=6 aorta/group). Aortic wall elastin filament degradation was graded based on the degree of breaks in elastin filaments (graded on a scale of 1–4) as described in the ‘Materials and methods’ section. (E) Quantification of polarization images for collagen content expressed as a percentage (%) of the total suprarenal aorta section area (n=6 aorta/group). Data are shown as median and interquartile ranges. Abbreviation: VC, vehicle control.

Discussion

DCs are the major APCs and play a central role in the initiation and activation of the immune response. The cytokines secreted by the DCs dictate the outcome and type of immune response [31]. In the current study, a diphtheria toxin (DT)-based system was used to induce in vivo depletion of CD11c+ cells in mice [32]. CD11c+ cells are largely DCs although CD11c+ is also expressed on intraepithelial lymphocytes and CD8+ T cells to a lesser degree [33]. The findings of this study suggest that CD11c+ cell depletion limits the severity and growth of experimental AAA in AngII-infused ApoE–/– mice. CD11c+ cell depletion was associated with significant reduction in circulating CD44hi CD62Llo effector CD4 and CD8 T-cell subsets, and a reduction in circulating B cells.

Past reports on the role of lymphocytes in AngII-induced AAA have been contradictory. AngII-infusion has been previously reported to increase the numbers of CD3+ T cells in the spleen of ApoE–/– mice [34,35]. Removal of the spleen or deficiency of lymphocytes has been reported to impair monocyte mobilization in response to AngII and protect against AAA development [35]. However, total deficiency of lymphocytes has been reported to attenuate AngII-induced atherosclerosis but not AAA in ApoE–/– mice [36]. Similarly, B cells have been reported to promote mobilization of monocytes from the spleen to the aorta in response to AngII in ApoE–/– mice [35,37]. The mobilization of the lymphocyte antigen 6C-high (Ly-6Chi) and lymphocyte antigen 6C-low (Ly-6Clo) monocytes from the spleen in response to AngII is dependent on the presence of B cells and has been reported to contribute to the development of AAA and the occurrence of aortic rupture. The role of CD11c+ cells in AngII-induced AAA formation in ApoE–/– mice has not been previously reported.

The inhibition of AAA formation and progression by CD11c+ cell depletion in the present study was associated with reduced circulating proportions of activated T and B cells, and reduced matrix degradation in the aortic wall. This finding is in line with a recent study in which depletion of plasmacytoid DCs (pDCs) abrogated CD3+ T-cell recruitment to the aorta, reduced matrix metalloproteinase (MMP) activity and inhibited AAA in the elastase mouse model [38]. Together, these data suggest an important role for DCs in promoting AAA pathogenesis. It is likely that the absence of DCs rendered T cells hyporesponsive in these mice due to defective TCR signaling. DCs act as messengers between the innate and the adaptive immune systems, and play diverse roles in T-cell immunity and tolerance such as priming of CD4 and CD8 T cells, and induction and expansion of regulatory T cells (T regs) [39]. T regs have been shown to suppress experimental AAA [40]. Importantly, patients with AAAs do not have functional circulating T regs [41]. DC may expand T regs [42], which is crucial for balancing immune responses. There are ongoing efforts to develop a vaccination against autoantigens found in the atherosclerotic plaque to induce immune tolerance and avoid tissue-damaging immune responses [43]. The state of DC presenting of antigens to lymphocytes may be crucial for inducing tolerance. A previous study reported atorvastatin administration to acutely reduce proportions of DCs and JNK expression, resulting in reduced inflammatory cell content and degradation of extracellular matrix (ECM) in the AAA wall through MMP expressions [44]. A randomized trial in AAA patients suggested that treatment with atorvastatin for 4 weeks significantly reduced inflammatory cells including DCs, macrophages and T lymphocytes along with increased interstitial collagen content and SMC proliferation in the human AAA wall [44]. Furthermore, a recent systematic review suggested that statin administration is associated with reduced AAA growth and rupture risk, although no large randomized trial has been performed [45]. It is possible that this purported benefit of statins could in part be due to favorable anti-inflammatory effects.

Neutrophil elastase is a cytotoxic serine protease stored in the azurophil granules of neutrophil granulocytes and is released following its activation. It causes degradation of a wide range of ECM proteins, including fibronectin, laminin, proteoglycans, collagens and elastin. Neutrophil elastase has shown to switch human immature DCs into Transforming Growth Factor (TGF)-β1 secreting cells [46] and promote the generation of T regs in vitro [47]. As local TGF-β1 levels and elastin filament degradation are important pathological steps in AAA formation, the neutrophil elastase activity was investigated in this study. Interestingly, in mice with established AAA, the plasma neutrophil elastase activity was reduced after CD11c+ depletion. A previous study had shown that pDC depletion inhibited AAA development in the elastase perfusion model by abrogating CD3+ T-cell recruitment and lowering MMP activity within the aorta [38]. Further, a recent study using the synthetic elastase inhibitor AZD9668 suggested that administration of this agent limited experimental AAA progression by improving intraluminal thrombus colonization by stromal cells [48]. These results align with the findings in the current study that depletion of CD11c+ cells limited inflammatory cell infiltration and elastase secretion within the aortic wall, thereby limiting AAA growth. Depletion of CD11c+ cells limited ECM degradation in our model. Elastin fibers showed increased breaks and collagen deposition was reduced in the control group compared with the CD11c+ depleted group. The findings suggest that DCs induce production of proteases that disintegrate the ECM and promote aortic wall remodeling.

The strengths of the present study include the combined assessment of the effect of CD11c+ cell depletion on both the initiation and progression of AngII-induced AAA in ApoE–/– mice. However, there are certain limitations to the study that should be acknowledged. A limitation of the current study is that blood pressure was not routinely measured. This is because past research suggests the AngII does not induce aortic aneurysm in mice through blood pressure elevation [25,48,49].

Previous studies suggest there are fundamental differences in inflammatory and immunologic responses between humans and mice [49], as well as between different mice strains [50,51]. Indeed, different strains of mice have different susceptibility to experimental AAA formation [52,53]. Thus translation of the current findings to human AAA patients is uncertain. While the present study suggests that CD11c+ depletion limits AngII induced development, there was no evidence it limited aortic rupture. The mechanisms involved in AAA development, growth and rupture have been suggested to be different. The current study was not designed to examine the effect of CD11c+ depletion on AAA rupture, which would require much larger numbers of mice.

Conclusion

The present study demonstrated that DC depletion protects mice from experimental AAA formation and progression. Mechanistically, DC depletion abrogated recruitment of T and B cells, and reduced neutrophil elastase activity resulting in reduced ECM degradation. Overall, the findings of the present study support the potential of DC vaccination or neutrophil elastase inhibitors (e.g. AZD9668) as novel approaches to treatment of AAA [54], although translation of these animal experimental findings to human disease needs to be further investigated.

Clinical perspectives

  • Role of CD11c+ APCs (DCs) in chronic inflammation AAA and the effect of conditional deletion of CD11c+ cells on experimental AAA has not been explored previously.

  • CD11c+ cell depletion was achieved with D.tox injections in transgenic mice and AAA formation and growth was assessed. CD11c+ cell depletion was shown to reduce the maximum diameter of AAAs induced by 28-days of AngII infusion. CD11c+ depletion commencing after AAA establishment by 14 days of AngII infusion was also shown to lead to smaller AAAs.

  • Further assessments showed that CD11c+ depletion led to lower proportions of CD4 and CD8 effector T cells and B cells, decreased neutrophil elastase activity and attenuated matrix degradation in experimental AAA.

Acknowledgments

We acknowledge Prof Christian Engwerda (Immunology and Infection Laboratory, QIMR Berghofer, Queensland, Australia) for providing the CD11c.DOG mice for conducting the present study.

Funding

This work was supported in part by the National Health and Medical Research Council [grant numbers 1098717, 1079369, 540403]; the Queensland Government; the Townsville Hospital Private Practice Trust; the Research Infrastructure Block Grant; the Medicine Incentive Grant; the School of Medicine, James Cook University; the Practitioner Fellowship from the National Health and Medical Research Council, Australia [grant number 1117061 (to J.G.)]; and the Senior Clinical Research Fellowship from the Queensland Government to J.G.]. The funding bodies played no role in generation of the data presented in this publication.

Competing Interests

The authors declare that there are no competing interests associated with the manuscript.

Author Contribution

S.M.K., C.S.M. and R.J.J. were involved in project administration, data curation, conceptualization, formal analysis, funding acquisition, investigation, methodology, validation, visualization, writing and editing. J.G. was involved in project administration, conceptualization, funding acquisition, investigation, methodology, supervision, writing and editing. S.L. and P.H. was involved in formal analysis and validation.

Abbreviations

     
  • AAA

    abdominal aortic aneurysm

  •  
  • AngII

    angiotensin II

  •  
  • APC

    antigen-presenting cell

  •  
  • ApoE-/-

    apolipoprotein E-deficient

  •  
  • DC

    dendritic cell

  •  
  • DOG

    DTR-OVA-eGFP

  •  
  • DTR

    diphtheria toxin receptor

  •  
  • D.tox

    diphtheria toxin

  •  
  • ECM

    extracellular matrix

  •  
  • EVG

    Verhoeff-van Giesson

  •  
  • HDL

    high-density lipoprotein

  •  
  • IRA

    infra-renal aorta

  •  
  • i.p.

    intra-peritoneal

  •  
  • LDL

    low-density lipoprotein

  •  
  • MMP

    matrix metalloproteinase

  •  
  • pDC

    plasmacytoid DC

  •  
  • SRA

    supra-renal aorta

  •  
  • TGF

    transforming growth factor

  •  
  • T reg

    regulatory T cell

  •  
  • VLDL

    very LDL

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Author notes

*

These authors contributed equally to this work.