Mfap4-deficient (Mfap4^−/−) mice were generated in-house as previously described (16) and crossbred with C57BL/6N mice (Charles River Laboratories International) for >10 generations before they were used for experiments.

ApoE-deficient (B6.129P2-Apoe^tmlUnc/J, stock nr 002052, ApoE^−/−) mice were obtained from Jackson Laboratory and back-crossed to the C57BL/6N background. ApoE^−/− mice and double ApoE- and Mfap4-deficient (ApoE^−/−Mfap4^−/−) littermate mice were produced by ApoE^−/−Mfap4^+/− breeding pairs.

The mice were housed in separate single cages during the course of the experiment. All animal experiments were approved by the National Animal Experiments Inspectorate of Denmark (permit numbers 2012-15-2934-00047 and 2015-15-0201-00474).

Experimental AAAs were induced using a continuous infusion of Ang II as described previously (17). This model shows a strong male gender preference, recapitulating the much higher incidence of human AAA in men than in women (18). Therefore, this study only included male mice in accordance with the guidelines described in the ATVB Council Statement (19). Male ApoE^−/−Mfap4^−/− and littermate ApoE^−/− control mice were fed western diet 1 week before surgery and throughout the experiment. At the age of 10–12 weeks, subcutaneous osmotic minipumps (Alzet® Model 2004, DURECT^TM Corporation, Cupertino, CA, USA) were installed via a mid-scapular incision under mild anesthesia (2% isoflurane, IsoFlo® vet, Orion Pharma, Nivå, Denmark) supplemented with analgesia (subcutaneous injection of 5 μg/g carprofen, Rimadyl, Pfizer, Ballerup, Denmark). Adequacy of anesthesia was monitored throughout the procedure by the toe pinch reflex.

The pumps delivered saline or Ang II (Calbiochem, Merck Millipore, Darmstadt, Germany) at 1,000 ng/kg/min for 9, 21, and 28 days. The body weight and overall well-being were regularly monitored for all mice throughout the treatment period.

Mice were euthanized by CO₂/O₂ asphyxiation. Abdominal aortic tissue and/or serum was sampled and snap-frozen 9 days after surgery. Aortic diameter (AD) was measured 28 days after surgery. AAA severity was scored as previously described (20). The cardiovascular system was perfused with sterile PBS and the hearts were dissected, rinsed with sterile PBS and weighed. The aortas were carefully isolated from the heart to the iliac bifurcation, cleaned from fat and connective tissue, weighed, mounted on black wax and measured. The parts of the aortas with a maximum diameter were subsequently fixed in 4% (v/w) formaldehyde for 24 h, rinsed in PBS and paraffin-embedded.

Maximal AAA diameter in dissected aortas from ApoE^−/− and ApoE^−/−Mfap4^−/− mice was measured using a 5 mm measuring scale (Ted Pella, Inc., Redding, CA, USA) and a Canon EOS 6D camera. The measurements were performed in affected regions using Adobe® Photoshop® CC2018 (San Jose, CA, USA) in a blinded manner by two independent investigators. AAA was defined as a diameter increase >50% compared to the average aortic diameter of saline-infused mice.

In situ hybridization was performed using a modified version of the RNAScope 2.5 high-definition procedure (Advanced Cell Diagnostics, Newark, CA, USA). Mouse aortic tissues were hybridized with 20 probe pairs (421391, Advanced Cell Diagnostics) targeting nucleotides 98-1231 of mouse Mfap4 mRNA (accession number NM_029568.2) followed by branched DNA signal amplification and tyramide enhancement visualized with Liquid Permanent Red (Agilent). The sections were subsequently immunostained with anti-α-smooth muscle actin (α-SMA) antibodies (Agilent) detected with anti-mouse BrightVision horseradish peroxidase (ImmunoLogic, Duiven, the Netherlands) and visualized with Deep Space Black (BIocare Medical, Pacheco, CA, USA).

Four μm-thick serial sections were stained with hematoxylin and eosin (H&E), Verhoeff-Van Gieson, Picrosirius red and for: α-SMA, cleaved caspase-3, CD45, F4/80, Ki67, MMP-9, MMP-2, CD31, CD11b, phosphorylated (p)FAK, pSMAD2, pSMAD3, and MFAP4 (Supplementary Table 1). All immunostainings were counter-stained with hematoxylin. The stainings were performed on a Dako Autostainer Universal Staining System (Dako, Denmark A/S, Glostrup, Denmark). Stained sections were scanned at 20x magnification using NanoZoomer-XR (Hamamatsu Photonics, Hamamatsu, Japan).

The scanned images were analyzed in a blinded manner using Adobe Photoshop or ImageJ.

Ki67-positive nuclei, CD31-positive microvessels, and MMP-9-positive cells were quantified as cells per section. Collagen, α-SMA, cleaved caspase-3, CD45, F4/80, CD11b, MFAP4, MMP-2, pSMAD2, pSMAD3, and pFAK stainings were quantified as staining-positive area using automated color threshold analysis. Elastic fiber degradation was assessed as a percentage of destroyed Verhoeff van Gieson-positive tissue area. Briefly, the damaged regions showing degradation of proper elastic lamellar structure were delineated and presented as a fraction of a total intima-media area. All analyses were performed in a blinded manner. Representative images of isotype control stainings are shown in Supplementary Figure 1. Representative images of entire aortic sections are shown in Supplementary Figure 2.

Four μm-thick serial sections were deparaffinized, subjected to antigen retrieval with citrate buffer (pH 6.0), blocked with 3% BSA and subsequently double-stained for Ki-67 and CD45 or Ki-67 and α-SMA (Supplementary Table 1). The sections were counterstained with DAPI and mounted using Fluorescent Mounting Medium (Dako). Fluorescent images were visualized and acquired using Olympus BX63 microscope (Olympus) and X-cite 120LED (Lumen Dynamics) with an Olympus DP80 camera and analyzed using ImageJ.

Serum levels of mouse MFAP4 were measured using a modified AlphaLISA immunoassay (Perkin Elmer, Waltham, MA, USA) as described previously (8). The two utilized anti-MFAP4 monoclonal antibodies (HG-HYB 7-14 and HG-HYB 7-18) had been raised against human recombinant MFAP4 and cross-react with the murine MFAP4 homolog due to very high sequence similarity. Data are presented as U/ml. When measured in human serum, 1 U/ml of MFAP4 corresponds to a concentration of 38 ng/ml.

THP-1 human monocyte leukemia cell line (ATCC) was grown in RPMI-1640 medium (Gibco, TermoFisher) supplemented with 10% FBS (Sigma-Aldrich), 5,000 U/ml penicillin, 5,000 μg/ml streptomycin, and 200 mM L-glutamine (all from Gibco) at 37°C and 5% CO₂ humidity. The cells were subcultured every second-third day.

MaxiSorp 96-well plates were coated overnight at 4°C with human serum albumin (HSA) or immobilized MFAP4 (both 10 μg/ml). The cells seeded (40.000 cells/well) and differentiated with 5 nM phorbol 12-myristate 13-acetate (PMA; Sigma) for 48 h, equilibrated with complete medium for 24 h, and serum-starved for 48 h. The cells were then stimulated with 20 ng/ml TNF (R&D Systems) for 48 h. Zymography on culture supernatants was performed essentially as described above. Cell proliferation rate was assessed using WST-1 assay (Sigma) according to the manufacturer's instructions. Cell viability was assessed by CytoTox-ONE^TM Homogeneous Membrane Integrity Assay (Promega) according to the manufacturer's instructions. The zymography results were normalized to the cell proliferation index.

Fetal human primary aortic SMCs (Cell Applications) were grown in Smooth Muscle Cell Growth Medium (Cell Applications) supplemented with 5,000 U/ml penicillin and 5,000 μg/ml streptomycin. Cells from passages 3–10 were used.

The cells were seeded at HSA- and MFAP4-coated plates essentially as described above (16.800 cells/well), starved overnight in Smooth Muscle Cell Basal Medium (Cell Applications) and stimulated with indicated concentrations of TNF or Ang II for 24 h. Zymography on culture supernatants was performed essentially as described above.

Human peripheral blood mononuclear cells were isolated from buffy coats obtained from the local blood bank (permit nr DP086) using Ficoll-Paque Plus density gradient centrifugation (GE Healthcare). Briefly, the samples were layered over 15 ml of Ficoll and centrifuged brake-free for 25 min at 800 g at room temperature. The interface was removed and washed twice with PBS containing 2% FBS and 1 mM EDTA. The cells were resuspended in PBS containing 2% FBS and 1 mM EDTA. The monocytes were enriched using the EasySep™ Human CD14 Positive Selection Kit (Stemcell Technologies) according to the manufacturer's instructions. The purity of isolated monocytes was tested by flow cytometry immediately after isolation by staining with anti-CD14-FITC antibody (BD Biosciences) and was >97% in all experiments.

The lower sides of the Transwell inserts with 5.0 μm pores (Corning) were coated with 10 μg/cm² MFAP4 or HSA overnight at 4°C, washed with PBS, blocked with 10 mg/ml HSA for 1 h at room temperature and washed again. The monocytes were seeded in the upper chamber (100.000 cells/insert) in serum-free RPMI medium containing 0.5% FBS. In some experiments, the cells were pre-incubated with anti-integrin α_Vβ₃, anti-integrin α_Vβ₅ (both from Merck Millipore), or isotype control antibody (Thermo Fisher) for 30 min at room temperature before seeding. The lower chamber contained serum-free RPMI medium with 0.5% FBS ± 100 ng/ml human recombinant CCL-2 (R&D Systems). The cells were allowed to migrate for 3 h, after which the upper sides of the filters were washed with PBS and swiped with a cotton swab to remove any non-migrated cells. The lower sides of the filters were then stained with Hemacolor (Sigma) and divided into four fields. The migrated cells in each field were counted in a blinded manner by two independent investigators.

The normality of data was assessed using Shapiro-Wilk test. Levene's test was used to assess equality of variances. Non-normally distributed data were analyzed using Mann-Whitney U-test. Normally distributed data were analyzed using one-way ANOVA or Student's t-test. Comparisons of aneurysm incidence were performed using Fisher's exact test. Data are presented as means + SEM unless otherwise stated. Significance was accepted if p < 0.05. All statistical analyses were performed using GraphPad Prism (GraphPad.com).

Initially, we performed RT-qPCR analysis to assess the relative Mfap4 mRNA levels in the aorta as well as 19 additional tissues from wild-type C57BL/6N mice. The relative expression levels of Mfap4 were highest in the lung and arteries (Figure 1A). MFAP4 is an ECM molecule with the capacity to bind various ECM fibers including collagen (21). In line with this, the tissue expression profile of type I collagen (Col1a1) mRNA resembled the Mfap4 expression profile (Figure 1B).

In agreement with previous observations (16), MFAP4 was predominantly localized to the arterial elastic fibers by immunohistochemistry, and this localization was unaltered after 28 days of Ang II infusion (Figure 1C). To identify Mfap4-expressing cells, we performed in situ hybridization staining for Mfap4 transcript that revealed that Mfap4 mRNA is expressed in adventitial cells (presumably adventitial fibroblasts) and medial SMCs and that it is upregulated upon AAA infusion (Figure 1D and Supplementary Figure 3). Ang II-induced AAA development did not significantly impact MFAP4 deposition within the aortic ECM (Figure 1E) or circulating serum MFAP4 levels (Figure 1F).

Following, we assessed the changes in basic physiological parameters after Ang II infusion. We have previously reported that MAP is not affected by Mfap4 ablation in unchallenged mice when measured over periods of minutes-to-hours 5 days after placing indwelling catheters (16). In the present study, HR and MAP were measured continuously for 7 days using indwelling catheters in conscious ApoE^−/− and ApoE^−/−Mfap4^−/− mice infused with Ang II. A significantly increased HR and a tendency for increased MAP was observed in the active night period (6 p.m. to 6 a.m.) compared to the day period (6 a.m. to 6 p.m.) for both ApoE^−/− and ApoE^−/−Mfap4^−/− mice after Ang II treatment. However, we did not observe any significant differences in either HR or MAP between Ang II-infused ApoE^−/− and ApoE^−/−Mfap4^−/− mice (Supplementary Figures 4A,B). Likewise, heart weight and heart-to-body weight ratio were significantly increased after Ang II infusion but not influenced by Mfap4 genotype (Supplementary Figures 4C,D).

We next asked whether MFAP4 plays a role in the development of Ang II-induced AAA. As expected, none of the saline-infused mice developed AAAs. In contrast, 28 day-long Ang II infusion caused suprarenal AAA development, with ApoE^−/− Mfap4^−/− mice showing a significantly lower incidence rate (62%) compared to 93% incidence rate in ApoE^−/− mice (Figures 2A–C). Moreover, Ang II-infused ApoE^−/−Mfap4^−/− mice exhibited significantly lower maximal outer AAA diameter compared to ApoE^−/− mice (Figure 2D) as well as ameliorated AAA severity (Figure 2E). The dissected aorta weight was significantly reduced from 7.4 ± 3.0 mg in Ang II-treated ApoE^−/− mice to 0.4 ± 0.3 mg in ApoE^−/−Mfap4^−/− mice (Figure 2F). A similar trend was observed in the vascular luminal volume between ApoE^−/− and ApoE^−/−Mfap4^−/− mice when measured over a distance of four specific vertebrae using micro-CT in a limited number of samples (Figures 2G,H). On the other hand, there was no significant difference in survival (caused by early aneurysm rupture) between ApoE^−/− and ApoE^−/−Mfap4^−/− mice (Figure 2I). In addition, no effect of Ang II infusion or MFAP4 deficiency was observed on serum total cholesterol or triglyceride levels (Supplementary Figure 5).

Following, we evaluated the MFAP4-dependent changes in the inflammatory responses within the aortic wall. Ang II-induced inflammatory infiltration, quantified as the CD45-positive area, was significantly attenuated within the aortas of ApoE^−/−Mfap4^−/− mice compared to ApoE^−/− littermates (Figures 3A,B). Moreover, Ang II-infused ApoE^−/−Mfap4^−/− mice showed a potent reduction in CD11b-positive area in the aortas of Ang II-infused ApoE^−/−Mfap4^−/− mice compared to ApoE^−/− littermates (Figures 3C,D), suggesting monocytes/macrophages to be the affected leukocyte type. We confirmed this by staining for another macrophage marker F4/80, which yielded comparable results (data not shown). Furthermore, CD11b-positive area analyzed exclusively within the adventitial layer was also significantly lowered in Ang II-infused ApoE^−/−Mfap4^−/− mice compared to ApoE^−/− littermates (Supplementary Figure 6).

We further analyzed aortic tissue lysates (collected at day 9) by zymography to investigate MMP activity in Ang II-infused mice (Figure 3E). We observed a significant decrease in both MMP-2 (Figure 3F) and MMP-9 (Figure 3G) activity in Ang II-infused ApoE^−/−Mfap4^−/− mice compared to ApoE^−/− littermates. To confirm that, we analyzed MMP protein expression in aortic sections and found that both MMP-2 and MMP-9 expression, localized predominantly in the adventitial layer, were significantly decreased in Ang II-infused ApoE^−/−Mfap4^−/− mice compared to ApoE^−/− littermates (Figures 3H–K). These results suggest that MFAP4 promotes inflammatory responses in macrophages during AAA development.

As integrin receptors serve as main MFAP4 cellular ligands, we stained AAA sections for pFAK as a proxy for activation of integrin signaling pathways. We observed that total pFAK-positive adventitial area as well as pFAK-positive area normalized to adventitial area were significantly reduced in Ang II-infused ApoE^−/−Mfap4^−/− mice compared to ApoE^−/− littermates (Figures 3L,M and not shown).

Medial α-SMA-positive area remained unchanged between Ang II-infused ApoE^−/− and ApoE^−/−Mfap4^−/− mice (Figures 4A,B).

We then investigated the degree of cellular apoptosis and proliferation within the vessel wall. Ang II infusion resulted in an overall increase in both cleaved caspase 3-positive area and Ki-67 positive cell number. Apart from few isolated cells found throughout the tissue, the majority of cleaved caspase 3-positive staining was localized within the media at the sites of SMC degeneration. Ang II-induced apoptosis was significantly attenuated in ApoE^−/−Mfap4^−/− mice compared to ApoE^−/− littermates (Figures 4C,D).

On the other hand, Ki-67-positive cells were found throughout the vessel wall but predominantly in the adventitial layer. Ki-67 positive cell number was significantly reduced in Ang II-infused ApoE^−/−Mfap4^−/− mice compared to ApoE^−/− littermates (Figures 4E,F). Moreover, double immunofluorescent staining revealed that while single SMA-positive medial SMCs stained positive for Ki-67 (Figure 4G), the vast majority of Ki67-positive cells were CD45-positive infiltrating leukocytes (Figure 4H).

CD31-positive microvessels were essentially undetectable in aortic tissues from control mice, while numerous capillary vessels were observed in Ang II-infused aortas after 28 days. Ang II-infused ApoE^−/−Mfap4^−/− mice exhibited a 78% reduction in the observed number of microvessels compared to ApoE^−/− mice (Supplementary Figure 7).

Ang II infusion resulted in elastic membrane disruption that was significantly attenuated in ApoE^−/−Mfap4^−/− mice compared to ApoE^−/− littermates (Figures 5A,B).

We also analyzed AAA-linked fibrotic changes within the vessel wall. Ang II treatment induced adventitial collagen deposition that was significantly decreased in Ang II-infused ApoE^−/−Mfap4^−/− mice compared to ApoE^−/− littermates (Figures 5C,D). To investigate the related mechanisms, we stained AAA sections for pSMAD2 and pSMAD3, key mediators of pro-fibrotic TGF-β signaling. While the medial pSMAD-positive area was modestly influenced by MFAP4 genotype (Supplementary Figure 8), both the pSMAD2- and pSMAD3-positive area in the adventitia were highly reduced in Ang II-infused ApoE^−/−Mfap4^−/− mice compared to ApoE^−/− littermates (Figures 5E–H), showing that activation of TGF-β-dependent downstream signaling is significantly attenuated by MFAP4 deficiency.

To confirm our in vivo findings and better understand the molecular mechanisms behind MFAP4-mediated regulation of inflammatory infiltration, we evaluated the role of MFAP4 in chemotaxis of blood monocytes. We observed that MFAP4 alone was able to stimulate directional monocyte migration and that this increase could be inhibited by blocking integrin α_Vβ₃ but not integrin α_Vβ₅ (Figures 6A,B). We observed a similar tendency for monocyte chemotaxis toward CCL-2, although it did not reach statistical significance (Figures 6A,B).

Finally, we investigated if MFAP4 has a direct effect on MMP production in SMCs and macrophage-like cells. We stimulated fetal aortic SMCs and PMA-differentiated THP-1 cells with immobilized MFAP4 with or without TNF co-stimulation. MMP-2 activity in fetal aortic SMCs was independent of MFAP4 regardless of TNF or Ang II stimulation (Supplementary Figure 9A). Conversely, we observed that while TNF stimulation resulted in an overall increase in MMP-9 activity in PMA-differentiated THP-1 cells, co-stimulation with MFAP4 significantly potentiated MMP-9 activity when compared to TNF stimulation alone (Figure 6C). THP-1 cell proliferation or viability after TNF stimulation were not significantly influenced by MFAP4 (Supplementary Figures 9B,C).