CCDC80 May Help Prevent Aortic Tears by Keeping Artery Muscle Cells Contracting Properly

First, we examined the expression of CCDC80 in different tissues of adult mice. CCDC80 protein was expressed in several tissues including the lung, brain, and aorta; it was particularly expressed in the aorta (FigureS1A, Supporting Information), which is consistent with a recent report.^[14^] Remarkably, a high expression of CCDC80 protein was observed in VSMCs and endothelial cells but not in adventitial fibroblasts and macrophages in the aorta (Figure S1B, Supporting Information). Moreover, its expression decreased with age in male mice (Figure S1C, Supporting Information). These data indicate that CCDC80 is highly enriched in aortic VSMCs and may play an important role in maintaining vascular homeostasis.

To examine the role of CCDC80 in the aorta, we examined CCDC80 expression in the ascending aorta (ASC) of patients and mice with AD. CCDC80 mRNA and protein levels were downregulated in human AD aortas (Figure1A,B). Immunofluorescence (IF) staining showed that in human AD, CCDC80 was mainly reduced in VSMCs but not in endothelial cells (Figure 1C and Figure S2, Supporting Information). Moreover, quantitative PCR (qPCR) analysis (Figure 1D) and western blotting (WB; Figure 1E) showed that CCDC80 expression decreased in the aorta of Ang II + β‐aminopropionitrile monofumarate (BAPN)‐induced AD mice. Similarly, IF staining showed that CCDC80 was reduced in VSMCs of AD mice (Figure 1F). Collectively, these data demonstrate that CCDC80 was downregulated in both human and mice AD VSMCs, thereby indicating that CCDC80 plays a role in AD.

To investigate the role of CCDC80 in AD, we performed in vivo experiments using a mouse model of Ang II + BAPN‐induced AD. First, global CCDC80 knockout mice (CCDC80^−/−) were generated by inducing a CCDC80 genetic mutation. The global CCDC80 knockout mutation was confirmed at DNA, mRNA, and protein levels (Figure S3, Supporting Information). Then, CCDC80^−/− mice and C57BL/6J background mice (wild‐type mice, WT) were treated with Ang II + BAPN for 28 d. During the 28 d Ang II + BAPN treatment, CCDC80^−/− mice developed AD with increased frequency and severity compared with their littermate controls (55.56% (4/9) versus 100.00% (11/11), p = 0.0260) (Figure S4A,C, Supporting Information). The incidence of WT mice dying of AD and rupture was 11.11% (1/9). By contrast, 72.73% (8/11) of CCDC80^−/− mice died of aortic rupture (Figure S4B,C, Supporting Information). Blood pressure levels were similar between Ang II + BAPN‐treated CCDC80^−/− and WT mice (Table S1, Supporting Information). Vascular ultrasound imaging and maximal aortic diameter measurement at 28 days after vascular modeling demonstrated that compared with WT controls, CCDC80^−/− mice exacerbated Ang II + BAPN‐induced aortic dilation (Figure S4D–F, Supporting Information). These findings demonstrate that CCDC80 deficiency significantly increased aortic dilatation and promoted AD formation and rupture in mice.

Hypertension is the most important risk factor for AD.^[18^] Approximately 80% of patients with AD experience hypertension.^[19, 20^] Patients susceptible to the occurrence of AD primarily manifest elevated maximum systolic and mean aortic blood pressure levels.^[21^] To evaluate the effect of hypertension on CCDC80^−/− mice, we injected Ang II (1000 ng/kg/min) into male CCDC80^−/− and WT control mice for 14 d. Blood pressure was elevated in Ang II‐injected mice and was similar in Ang II‐injected CCDC80^−/− and WT mice (Table S2, Supporting Information). Remarkably, during the 14 d Ang II administration, 87.50% (21/24) of CCDC80^−/− mice experienced AD following Ang II treatment—primarily in the ASC and suprarenal abdominal aorta (AA)—compared with 8.33% (2/24) of WT mice (p < 0.0001) (Figure2A,D); 62.50% (15/24) of male CCDC80^−/− mice died of AD and rupture (Figure 2C,D) compared with 8.33% (2/24) of male WT mice. Moreover, WT and CCDC80^−/− mice treated with saline did not develop AD (Figure 2A,D). Dissected aortas in CCDC80^−/− mice exhibited a compressed true lumen as well as either an intramural hematoma or a large false lumen with thrombosed blood (Figure 2B). Vascular ultrasound imaging and maximal aortic diameter measurement performed on day 14 after modeling demonstrated that CCDC80 knockout exacerbated Ang II‐induced aortic dilation (Figure 2E–G). Hematoxylin and eosin (H&E), elastic van Gieson (EVG) staining, and Masson's staining demonstrated that AD formation, collagen deposition, elastic disarray, and elastic fiber degradation were exacerbated in Ang II‐treated CCDC80^−/− mice compared with WT mice (Figure 2H,I). Therefore, CCDC80 deficiency increases susceptibility to AD and rupture.

VSMCs are the main component cells of the vascular wall. They play an important role in maintaining vascular tone and integrity, regulating intravascular pressure, and redistributing blood volume, which are crucial for maintaining vascular homeostasis.^[22, 23^] CCDC80 was dramatically reduced in the aortas of patients and mice with AD primarily in VSMCs (Figure S1, Supporting Information). Therefore, we constructed VSMC‐specific CCDC80 knockout mice to examine the role of CCDC80 in AD.

To determine the role of VSMC‐specific CCDC80 in AD, we generated conditional CCDC80 knockout mice in VSMC (CCDC80^fl/fl SM22α Cre⁺) by crossing mice in which the exon 3 of the CCDC80 gene was flanked by two LoxP sites (CCDC80^fl/fl) with the mice containing the SM22α Cre⁺ mutation (Figure S5A, Supporting Information). The VSMC‐specific deletion of CCDC80 was confirmed at DNA, mRNA, and protein levels (Figure S5B–F, Supporting Information). There were no significant differences in expression in other tissues (heart, liver, lung, kidney, muscle, and brain) between CCDC80^fl/fl SM22α Cre⁻ and CCDC80^fl/fl SM22α Cre⁺ mice (Figure S5C, Supporting Information).

Next, we administered saline or Ang II to CCDC80^fl/fl SM22α Cre⁺ and CCDC80^fl/fl SM22α Cre⁻ mice for 14 days. Blood pressure levels were similar between Ang II‐injected CCDC80^fl/fl SM22α Cre⁺ and CCDC80^fl/fl SM22α Cre⁻ mice (Table S3, Supporting Information). Although Ang II treatment resulted in AD in both genotypes, the incidence was significantly higher in CCDC80^fl/fl SM22α Cre⁺ mice than in CCDC80^fl/fl SM22α Cre⁻ mice (80.00% versus 12.50%, p < 0.01) (Figure3A,C). AD particularly developed in the ASCs and AAs in VSMC‐specific CCDC80 knockout mice (Figure 3A). Furthermore, a higher frequency of aortic rupture was noted in CCDC80^fl/fl SM22α Cre⁺ mice (5/10, 50.00%) compared with that in CCDC80^fl/fl SM22α Cre⁻ mice (1/8, 12.50%; Figure 3B,C). After 2 weeks of Ang II treatment, vascular ultrasound imaging and maximal aortic diameter measurement demonstrated that VSMC‐specific CCDC80 knockout mice exacerbated Ang II‐induced vascular expansion compared with CCDC80^fl/fl SM22α Cre⁻ mice (Figure 3D–F).

Histopathological analysis revealed typical AD lesions and determined that AD was the main cause of aortic rupture in CCDC80^fl/fl SM22α Cre⁺ mice receiving chronic Ang II treatment (Figure 3G,H). EVG staining and Masson's staining of aortic sections obtained from Ang II‐injected CCDC80^fl/fl SM22α Cre⁺ mice showed more severe collagen deposition and media degeneration, including elastic fiber fragmentation and disorganization (Figure 3G,H). These findings suggest that the deletion of VSMC‐specific CCDC80 exacerbates Ang II‐induced AD and that VSMC‐specific CCDC80 plays a major role in regulating vascular homeostasis.

Because CCDC80^−/− mice suddenly died from aortic rupture in the early days after Ang II administration, we harvested the aortas after 3 d of Ang II administration to evaluate the initial events before aortic rupture (Figure S6A,B, Supporting Information). Compared with WT mice, elastic fibers in the ASC and AA of CCDC80^−/− mice were partially disrupted and degraded and red blood cells were present between the elastic laminas of the degraded aortic walls (Figure S6C, Supporting Information).

Next, mRNA sequencing was performed to identify differentially expressed genes (DEGs) in aortic tissues obtained from WT and CCDC80^−/− mice treated with Ang II for 3 d. In CCDC80^−/− samples, 1405 genes were upregulated (Figure4A), including those regulating ECM disassembly and SMC cell migration and proliferation (Figure 4B). By contrast, 482 genes (Figure 4A) were downregulated, particularly those enriched for SMC differentiation and muscle contraction (Figure 4B). Remarkably, the cell differentiation cluster was downregulated in CCDC80^−/− samples (Figure 4C), thereby indicating the activation of VSMC phenotype switching in CCDC80^−/− mice. In particular, the expression of VSMC contractile genes such as Myh11, SM22α, α‐SMA, and CNN1 was downregulated in CCDC80^−/− aortas, whereas the expression of VSMC synthetic genes such as Runx3, Thbs2, and Spp1 was upregulated (Figure 4C), thereby substantiating the crucial role of CCDC80 in VSMC phenotype switching in AD pathogenesis.

Synthetic VSMCs secrete MMPs and exhibit an inflammatory state via the expression of cell adhesion molecules and secretion of chemokines and inflammatory cytokines.^[24^] To further confirm the abnormal VSMC phenotype switching in CCDC80^−/− mice, we performed qPCR on an array of VSMC‐related markers in aortic tissues harvested from CCDC80^−/− mice and their littermate controls after 3 d of Ang II administration. The absence of CCDC80 significantly reduced the contractile structural markers of VSMCs, such as α‐SMA, SM22α, CNN1, and Myh11 (Figure 4D). Moreover, an increased expression of synthetic genes, such as osteopontin, proliferating cell nuclear antigen (PCNA), tumor necrosis factor‐α (TNFα), CCL2, MMP9, MMP13, and MMP14, was observed in CCDC80 knockout mice compared with their littermate controls (Figure 4E,F). Furthermore, WB, IF staining, and IHC staining analyses revealed that Ang II treatment significantly downregulated the expression of contractile markers, including α‐SMA and SM22α, while upregulating synthetic markers such as PCNA in the aorta of both genotypes. Following Ang II administration, CCDC80^−/− mice exhibited a more pronounced reduction in contractile marker levels and a greater increase in synthetic marker expression in the aorta compared with WT mice (Figure 4G—L and Figure S7A–E, Supporting Information). Moreover, CCDC80 deficiency increased the expression of inflammatory factors (e.g., TNF‐α) but displayed no effect on VSMC apoptosis in the ASC (Figures S8 and S9, Supporting Information).

MMPs are a family of proteolytic enzymes that degrade ECM proteins and are critical for cell migration and tissue remodeling under physiological and pathological conditions.^[25^] Studies have reported that MMP2 and MMP9 secreted by VSMCs play an important role in ECM degradation in aortic aneurysms and AD.^[26, 27^] Following Ang II administration, MMP2/9 activity—determined via an in situ MMP activity assay—was enhanced in the aorta of CCDC80^−/− mice compared with WT mice (Figure S10A, B, Supporting Information). The aortas harvested from CCDC80^−/− mice expressed increased Ang II‐induced MMP9 mRNA compared with WT mice (Figure 4F). Furthermore, WB revealed that MMP2 and MMP9 expression were increased in Ang II‐induced WT mice. CCDC80^−/− mice exhibited a higher expression of Ang II‐induced aortic MMP9 expression compared with WT mice; however, Ang II‐induced aortic MMP2 expression was not significantly different between CCDC80^−/− and WT mice (Figure 4M–O). IF staining revealed that CCDC80 deficiency increased Ang II‐induced MMP9 expression in VSMCs (Figure S10C,D, Supporting Information). These data indicate that CCDC80 plays a novel protective role in the context of AD by regulating VSMC phenotype switching and decreasing MMP9 expression.

To investigate the effect of CCDC80 on the phenotype switching of VSMCs, we analyzed the expression of VSMC‐specific genes including α‐SMA, SM‐22α, and MYH11 using WB and qPCR. The expression of these genes was significantly downregulated in CCDC80^−/− VSMCs under quiescent and Ang II‐induced conditions (Figure5A–D). Moreover, cell proliferation experiments showed that VSMCs isolated from CCDC80^−/− mice exhibited greater proliferation after 24 h in culture, with or without Ang II treatment, compared with those from WT mice (Figure 5E,F). Next, we investigated to study the effect of CCDC80 deletion on VSMC migration and cell contraction capability using an in vitro scratch wound assay and collagen gel contractile assay. The results showed that Ang II‐induced cell migration was enhanced in CCDC80^−/− VSMCs (Figure 5G). And, Ang II‐induced VSMCs contraction was remarkably hampered in CCDC80^−/− VSMCs (Figure S11A,B, Supporting Information). These results further confirm that CCDC80 deletion facilitates VSMC phenotype switching from a quiescent contractile phenotype to an active synthetic phenotype.

To analyze the biological role of CCDC80 in AD, we studied CCDC80‐initiated transcriptional programs. Furthermore, we performed Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway analysis of the genes differentially expressed between WT and CCDC80^−/− groups treated with Ang II. The results revealed that CCDC80 deficiency could regulate several pathways including the JAK/STAT, PI3K‐AKT, NF‐kappa B, Rap1, MAPK, and ECM‐receptor interaction signaling pathways (Figure S12A, Supporting Information). Consistently, gene‐set enrichment analysis revealed a positive correlation between CCDC80 deficiency and the JAK/STAT signaling pathway (NES = 1.4807, p < 0.0001; Figure S12B, Supporting Information).

Previous studies have reported that the JAK/STAT signaling pathway is the central node for cytokine signaling and plays an important role in regulating cell proliferation, migration, and apoptosis as well as immunity, tissue repair, inflammation, and adipose differentiation.^[28^] However, the abnormal activation of the JAK/STAT signaling pathway is significantly related to the occurrence of vascular diseases such as aortic aneurysm.^[29^] Recent studies have reported that CCDC80 interacts with JAK2 (Janus kinase 2) through the DRO1‐URB‐DRS‐Equarin‐SRPUL/sushi repeat containing protein, x‐linked (DUDES) domain to regulate the downstream signal of STAT3 (Signal transducer and activator of transcription 3).^[30^] Moreover, STAT3 can play an important role in cardiovascular diseases (such as atherosclerosis and pulmonary hypertension) through signal transduction and transcriptional activation.^[31, 32^] Therefore, we further examined the role of the JAK2/STAT3 pathway in AD. CCDC80 deletion significantly promoted Ang II‐induced expression of JAK2 (Figure6A,B). Moreover, after 3 d of Ang II administration, the levels of phosphorylated STAT3 proteins, a downstream effector of JAK2, were significantly upregulated in CCDC80‐knockout aortas (Figure 6A,C). Furthermore, CCDC80 coimmunoprecipitated with JAK2 in mouse VSMCs and this interaction significantly decreased after Ang II stimulation (Figure 6D). These results suggest that CCDC80 strongly binds to JAK2 in the absence of Ang II and that CCDC80 deletion significantly activated the JAK2/STAT3 signaling pathway.

The JAK2/STAT3 pathway‐specific inhibitor WP1066 can irreversibly inhibit JAK2 and STAT3 by degrading JAK2 proteins and subsequently blocking downstream STAT3 pathway signal transduction and activation.^[33^] To investigate the inhibitory effect of WP1066 on the JAK2/STAT3 pathway, we analyzed the expression of JAK2 and STAT3 using WB. The expression of JAK2 and p‐STAT3 was significantly downregulated in CCDC80^−/− mice aorta treated with WP1066 (Figure S13A,B, Supporting Information). Then, we examined the therapeutic effect of WP1066 on AD development in CCDC80^−/− mice with Ang II‐induced AD. Briefly, CCDC80^−/− mice were intraperitoneally injected with WP1066 daily (20 mg kg⁻¹) following Ang II administration throughout the 2 weeks of modeling.

Notably, compared with CCDC80^−/− mice that did not receive WP1066 treatment, WP1066‐treated CCDC80^−/− mice demonstrated significantly lower AD formation (90.00% vs 55.00%, p < 0.05; Figure 6E,G) and lethality (Figure 6F,G) as well as mitigated aortic dilation (Figure 6H,I). Similarly, WP1066 treatment exerted no impact on blood pressure (Table S4, Supporting Information). Histological analyses via H&E, EVG, and Masson's staining techniques revealed that WP1066 ameliorated elastin disorganization, elastic fiber degradation, and collagen deposition (Figure 6J,K), thereby further confirming the critical role of the JAK2/STAT3 pathway in AD progression.

To investigate the effect of WP1066 on phenotype switching, we performed qPCR on an array of VSMC‐related markers in aortic tissue harvested from CCDC80^−/− mice. Following Ang II administration, compared with mice that received no WP1066 treatment, WP1066‐treated CCDC80^−/− mice exhibited increased expression of VSMC contractile structural markers (including α‐SMA, SM22α, and Myh11; Figure7A), reduced expression of synthetic genes (including PCNA, MMP9, MMP13, and MMP14), and exerted no effect on CNN1, TNFα, CCL2, and MMP2 (Figure 7B,C). Similarly, WB and IF staining revealed that compared with the absence of WP1066 treatment, WP1066 administration resulted in a significant increase in contractile markers, such as α‐SMA and SM22α, in the aorta (Figure 7D,E,J–M). MMP9 expression and MMP activity were decreased in Ang II‐induced CCDC80^−/− mice treated with WP1066 (Figure 7F–I). These data suggest that JAK2/STAT3 is located downstream of CCDC80 and thus mediates VSMC differentiation and vascular degradation.

The present study showed that CCDC80 could aggravate AD by promoting VSMC phenotype switching in mice. We investigated whether VSMC phenotype switching is exacerbated in human AD. Consistent with the results observed in mice AD, a decreased expression of α‐SMA and SM22α was observed in human AD compared with human normal aorta (Figure8A,B). Similarly, TNFα expression was elevated in the aorta of patients with AD compared with the aorta of normal individuals (Figure 8C,D). We assessed the MMP activity in vivo using an in situ zymography assay. AD increased MMP2/9 activity (Figure 8E,F). Furthermore, MMP9 expression increased in the aorta of patients with AD compared with that in the aorta of normal individuals; MMP2 expression remained unchanged (Figure 8G,H). In addition, JAK2 and phosphorylation levels of STAT3 proteins were upregulated in the aorta of patients with AD compared with that in the aorta of normal individuals (Figure 8I,J). These findings suggest that VSMC phenotype switching is involved in the formation and development of human AD.

Antibodies against β‐actin (4970), GAPDH (5174T), Phospho‐Stat3 (Tyr705) (9145), STAT3 (9139T), and JAK2 (3230) for WB were purchased from Cell Signaling Technology (Boston, MA, USA). Antibodies against α‐SMA (SC32251) for WB and immunostaining were purchased from Santa Cruz Biotechnology, Inc. (Dallas, TX, USA). Antibodies against SM22α (Ab14106) for WB and immunostaining were purchased from Abcam (Cambridge, UK). Normal mouse IgG (2729) used for immunostaining and as a negative control for co‐immunoprecipitation (Co‐IP) assay was obtained from Cell Signaling Technology. Antibodies against PCNA (10205‐2‐AP) for WB and immunostaining were purchased from Proteintech Group, Inc. (Wuhan, China). Antibodies against TNFα (GB11188), MMP9 (GB11132), MMP2 (GB11130), and CD31 (GB12063) for WB and immunostaining were purchased from Servicebio Group, Inc. (Wuhan, China). Antibodies against CCDC80 (PAJ901Ra01) for immunostaining, WB and Co‐IP assay were purchased from Cloud‐Clone Corp. (Wuhan, China). Antibodies against CCDC80 (AF3410) used for immunostaining were purchased from BD Biosciences (Franklin Lakes, NJ, USA). Antibodies against JAK2 (3230) for Co‐IP assay were purchased from Cell Signaling Technology (Boston, MA, USA). BAPN (A3134) and Ang II (A9525) were purchased from Sigma (St. Louis, MO, USA). WP1066 (HY‐15312) was purchased from MCE (Monmouth, NJ, USA).

Human samples were processed according to protocols approved by the Shanghai Jiao Tong University institutional review board (IS21014). Informed consent was obtained from each participant according to the Ethics Committee of Shanghai Chest Hospital, Shanghai Jiao Tong University School of Medicine. The study was carried out according to the criteria mentioned in the Declaration of Helsinki. Human AD tissue samples were obtained from six patients undergoing aortic root and ASC replacement; five control aortic tissues were obtained from age‐matched organ donors undergoing heart transplant surgery without AD, aneurysm, coarctation, or previous aortic repair. Characteristics of patients and controls are presented in Table(Supporting Information). S5

Experimental protocols were assessed and approved by the Animal Ethics Committee of Shanghai Chest Hospital [KS(Y)22333]. Animal experimental procedures were conducted in accordance with the guidelines of the National Institutes of Health Guidelines on the Care and Use of Laboratory Animals for scientific purposes. Global CCDC80 knockout mice (CCDC80^−/−) in a C57BL/6J background were generated by the Shanghai Model Organisms Center and maintained by mating CCDC80^+/− male mice with CCDC80^+/− female mice. WT littermates were used as controls. Conditional VSMC‐specific CCDC80 knockout mice (CCDC80^fl/fl SM22α Cre) were generated by breeding CCDC80^fl/fl mice with SM22α Cre [B6.Cg‐Tg(SM22α‐Cre)1Her/J, #01 7491] mice from Jackson Laboratory. CCDC80^fl/fl littermates were used as controls. A 12 h light/dark cycle was used; the mice had ad libitum access to diet and water and were housed under a temperature of 24 °C and humidity of 40%.

CRISPR/Cas9 technology was used to induce genetic mutations using guide RNA (gRNA) and Cas9 nuclease, resulting in the coding shift of the CCDC80 reading frame or premature termination of translation. The CCDC80 transcript was obtained from the Ensemble database (ENSMUST00000 099498.9). The gRNA target sequences were designed using the website (crispor.tefor.net) and the sequences with the highest score were selected (see Table S6 for the sequence, Supporting Information). Then, CCDC80‐specific gRNAs were obtained via in vitro transcription. F0 generation mice were obtained by injecting Cas9 nuclease and gRNAs into fertilized eggs. F0 generation mice confirmed using PCR (see Table S6 for primer information, Supporting Information) were mated with WT C57BL/6J mice to obtain F1 generation heterozygous mice (CCDC80^+/−). CCDC80^+/− mice were inbred to obtain CCDC80^−/− mice. The gRNA target sequences and primer information are provided in Table S6 (Supporting Information).

The target sequence was identified according to the gene structure of CCDC80; the gRNA for this site was then designed according to the target sequence. Following activity detection, the active gRNA and Cas9 were transcribed into RNA in vitro. The homologous recombinant vector (donor vector) was constructed using In‐Fusion cloning. The vector contained a 3.1‐kb 5′ homologous arm, 0.8 kb flox region, and 3.1‐kb 3′ homologous arm. Cas9 mRNA, gRNA, and donor vector were microinjected into the fertilized eggs of C57BL/6J mice to obtain F0 mice (see Table S7 for sequence, Supporting Information). Based on the structural analysis of the CCDC80 gene, exon 3 of the CCDC80‐201 transcript was selected as the flox region. The translation of exon 3 initiated at 65.9% of the gene‐coding region. Knocking out the region would cause a frameshift mutation in the reading frame, thereby leading to the early termination of translation. F0 generation mice identified by PCR (see Table S7 for primer information, Supporting Information) were mated with WT C57BL/6J mice to obtain F1 generation heterozygous mice (CCDC80^flox/+). CCDC80^flox/+ mice were inbred to obtain CCDC80^fl/fl mice. Then, CCDC80^fl/fl mice were bred with hemizygous SM22α Cre [B6. Cg ‐Tg (SM22α‐Cre)1Her/J, #017491] mice (see Table S7 for primer information, Supporting Information) from Jackson Laboratory to obtain CCDC80^fl/fl SM22α Cre mice. SM22α‐cre transgenic hemizygous mice are viable, fertile, and normal‐sized without any significant physical or behavioral abnormalities.^[54^] These transgenic mice express Cre recombinase under the control of a mouse smooth muscle protein 22‐α (SM22α) promoter. When hybridized with strains containing loxP site‐flanked sequence of interest, CRE‐mediated recombination resulted in the deletion of flanker sequences in vascular smooth muscle cells. Thus, Mice hemizygous for the SM22α‐cre transgene were used for breeding, and female breeders also carried SM22‐Cre. The gRNA target sequences and primer information are provided in Table S7 (Supporting Information).

Ang II‐ or Ang II + BAPN‐induced AD models were established as previously reported.^[55, 56, 57^] Briefly, 5‐12 week old male knockout (CCDC80^−/− and CCDC80^fl/fl SM22α Cre) and WT mice were randomly divided into Ang II or saline treatment groups.

At 5 weeks of age, mice were anesthetized and implanted with osmotic pumps (Alzet 2004, Durect Corp., Cupertino, CA, USA) filled with Ang II (1000 ng/kg/min, Sigma) for 4 weeks. The mice were provided regular diets and BAPN (1 g/kg/day, Sigma) dissolved in water. BAPN (1 g/kg/day, Sigma) was dissolved in water and provided at the same time as Ang II administering and was maintained for 4 weeks.

At approximately 8–12 weeks of age, mice were anesthetized with isoflurane (2%) and implanted with osmotic pumps (Alzet 2002, Durect Corp.) filled with Ang II (1000 ng/kg/min, Sigma) for 2 weeks.

Male CCDC80^−/− mice (aged 8–12 weeks) were administered WP1066 (20 mg kg⁻¹ per d, MCE) or saline via an intraperitoneal injection for 2 weeks.

Blood pressure was noninvasively measured in conscious mice using the tail‐cuff plethysmography method (BP‐2010A, Softron Biotechnology, Beijing, China). Mice were trained for the measurements before measuring the pressure levels. All measurements were performed at the same time (between 14 p.m. and 17 p.m.) to ensure accurate measurements and to avoid the influence of the circadian cycle. One set of six measurements was obtained for each animal; the average blood pressure was calculated.

At 2 weeks after the experiment, the mice were euthanized under deep anesthesia for blood collection. Then, the mice were perfused with saline, and aortic tissue was obtained and photographed. The maximum diameters of the ASC and AA were measured using calipers. Subsequently, the ASC and AA were sectioned and filled with an optical cutting temperature (OCT) compound or paraffin. The sectioned specimens (5 µm) were stained with H&E (Sigma), EVG (Abcam), and Masson's trichrome (Sigma). Other segments of the aorta were stored at −80 °C for molecular analysis.

To evaluate the morphological characteristics of the thoracic aortas and AAs in vivo, ultrasound imaging was performed using a Vevo2100 cardiovascular ultrasound system (VisualSonics Inc., Ontario, Canada) with a 30 MHz transducer for mice. Following skin hair removal at ultrasonic sites, mice were anesthetized with 1.0%–2.0% isoflurane inhalation and gently fixed to a heated platform in supine position. By adjusting the direction of the ultrasonic probe, the diameters of the thoracic aorta and AA were detected in B mode.

Mouse aorta VSMCs were isolated from the thoracic aortas of WT and CCDC80^−/− mice as described previously.^[58^] Briefly, mouse aortic tissues were isolated and rinsed in phosphate‐buffered saline (PBS). Following the removal of the endothelium and adventitia, the tissues were cut into approximately 2 mm sections and treated with 0.2% collagenase type II (Worthington Biochemical Corporation, OH, USA, LS004176↗,) at 37 °C for 2 h. VSMCs were then cultured in Dulbecco's modified Eagle's medium/F12 supplemented with 10% fetal bovine serum and 1% penicillin‐streptomycin. The VSMCs were used in the experiments between passages 3 and 6. And, the VSMCs were serum‐starved for 24 h before drug treatment. Then, immunofluorescence staining was performed to detect the presence of α‐SMA to validate VSMCs (Figure S14, Supporting Information).

Protein lysates from aortic tissues or cells were lysed using lysis buffer (Roche, Mannheim, Germany, 4719964001) containing protease and phosphatase inhibitors (Roche). Protein concentrations were evaluated using the Bradford protein assay (Thermo Fisher Scientific, Waltham, MA, USA). Protein samples (20 µg per lane) were electrophoresed on sodium dodecyl sulfate–polyacrylamide gels and transferred to polyvinyl difluoride membranes. Membranes were blocked using 5% non‐fat dry milk for 1 h at room temperature and incubated with primary antibodies for 12 h at 4 °C. The membranes were washed and incubated for 1 h at room temperature with anti‐rabbit or anti‐mouse secondary antibodies conjugated to horseradish peroxidase (HRP). GAPDH or β‐actin acted as the control. The protein bands were detected via enhanced chemiluminescence (Bio‐Rad Laboratories, Inc., Hercules, CA, USA) and analyzed using the ImageJ software (National Institutes of Health, Bethesda, MD, USA).

Total RNA was extracted using TRIzol reagent (Invitrogen, Waltham, MA, USA) according to the manufacturer's protocol and reverse transcribed into cDNA using the PrimeScript RT Master Mix kit (Takara). Quantification of mRNA was performed using TB Green Premix Ex Taq (Takara); real‐time PCR experiments were conducted using the ABI PRISM 7300HT sequence detection system (Applied Biosystems, Waltham, MA, USA). The mRNA levels were normalized against GAPDH. The primer sequences used for q‐PCR are listed in Table(Supporting Information). S8

Total RNA from the aortas was extracted using TRIzol reagent (Invitrogen, Waltham, MA, USA) following the manufacturer's protocol. The concentration, quality, and integrity of RNA was verified using the NanoDrop spectrophotometer (Thermo Fisher Scientific). RNA sequencing was performed on Novaseq 6000 (Illumina, San Diego, CA, USA). After obtaining data from the machine, it was converted to the original BCL file using real‐time analysis (version v3.4.4), split using the bcl2fastq (btq) software (version v2.19), and converted into FQ. The Fastp software (version 0.19.7) was used for quality control, involving the parameter fastp‐G‐Q5‐U50‐N 15‐L150. The DESeq21 R package was used to identify DEGs and corrected via the Benjamini–Hochberg method. A DEG is defined as one with an adjusted p‐value of <0.05 and |log2 (fold change) | of ≥1. A heatmap was generated using the pheatmap R package. KEGG and Gene ontology analyses were performed to assess pathway enrichment.

VSMCs were stimulated with Ang II (1 µm) for 48 h and lysed in cold lysis buffer (87787; Thermo Fisher Scientific) with a protease inhibitor cocktail on ice for 30 min. The lysates were centrifuged at 12 000 g and 4 °C for 10 min. The cell lysates were centrifuged; protein concentrations were determined using the Pierce BCA Protein Assay kit and adjusted to ≈5 µg µL⁻¹. Approximately 300 µg of the lysates were incubated with 15 µL of Protein A/G magnetic beads (88802; Thermo Fisher Scientific) at 4 °C overnight. Next, protein‐magnetic beads were incubated with rabbit anti‐JAK2 antibody, rabbit anti‐CCDC80 antibody, or anti‐IgG (2729; Cell Signaling Technology) at 4 °C overnight. Beads containing protein–antibody complexes were mixed with 2× loading buffer, boiled for 10 min, and analyzed with anti‐JAK2 and anti‐CCDC80 antibodies.

AAs and ASCs were collected and fixed with 4% paraformaldehyde for 48 h and embedded in paraffin. Aortic tissue sections (5 µm) were stained with H&E to observe general morphology according to the manufacturer's protocol. EVG staining was performed to assess elastic fiber integrity; Masson's trichrome staining was performed to evaluate collagen content. Elastin degradation was identified as a discontinuation and widening of the elastic lamina and was evaluated by counting the number of breaks per vessel.^[59^] The grade was defined as follows: Grade 1, no degradation; Grade 2, slightly degraded elastin; Grade 3, severe degradation of elastin, grade 4, Aortic rupture. Images were viewed using a microscope (Thermo Fisher Scientific, Waltham, MA, USA) and analyzed using the Image‐Pro Plus 6.0 software (Media Cybernetics Inc., Rockville, MD, USA) to assess the collagen content.

For IF staining, the formaldehyde‐fixed sections were permeabilized with 0.3% Triton, blocked with 5% bovine serum albumin, and incubated overnight with primary antibodies against α‐SMA (1:200, Santa Cruz Biotechnology), SM22α (1:200, Abcam), TNFα (1:200, Servicebio Group, Inc.), and CCDC80 (1:100, BD Biosciences) after continuous dewaxing, rehydration, and antigen repair. Then, the sections were incubated with green fluorescent secondary antibodies (Donkey Anti‐Rabbit IgG, 488 nm, Invitrogen), red fluorescent antibodies (Donkey Anti‐Mouse IgG, 555 nm, Invitrogen), or magenta fluorescent antibodies (Goat Anti‐rat IgG, 647 nm, Invitrogen). The nuclei were stained with 4′,6‐diamidino‐2‐phenylindole (DAPI, Beyotime, Shanghai, China) for 10 min. A fluorescence microscope was used for visualization (Leica, Germany).

For IHC staining, paraffin‐embedded sections (thickness of 5 µm) were treated with 3% hydrogen peroxide after continuous dewaxing, rehydration, and antigen repair. The sections were blocked with 5% normal fetal bovine serum, incubated with primary antibodies against CCDC80 (1:100, BD Biosciences) and PCNA (1:100, Proteintech Group, Inc.) overnight at 4 °C, and incubated with HRP‐conjugated secondary antibodies for 1 h. Blotting results were visualized with diaminobenzidine‐based HRP staining. Nuclei were counterstained with hematoxylin. The sections were imaged using fluorescence microscopy.

MMP2/9 activity was assessed using a Gelatinase Assay Kit (GMS80062.1↗, GenMed Scientifics Inc., Wilmington, DE, USA) according to the manufacturer's instructions. Briefly, unfixed ASCs or AAs were embedded with OCT and cut into 8‐µm‐thick serial sections. Fluorescein‐conjugated gelatin as a fluorogenic substrate was used to analyze gelatinolytic activity. Aortic tissue sections were incubated at 4 °C for 10 min and then maintained at 37 °C for 2 h. Fluorescein‐5‐isothiocyanate fluorescence was examined by fluorescence microscopy (Leica DM3000B, Leica Microsystems, Wetzlar, Germany) and determined using the ImageJ software.

For cell counts, primary VSMCs isolated from WT and CCDC80^−/− mice were plated on 96‐well plates at a density of 1 × 10⁴ cells per well. To render the VSMCs quiescent, the VSMCs were starved in serum‐free medium for 24 h. Then, they were treated with Ang II (1 µm) for 48 h. Next, 10 µL of CCK‐8 reagent (Beyotime) was added to each well and the plates were incubated for 4 h. Absorbance values were measured at 450 nm using an enzyme standard instrument (Thermo Fisher Scientific).

The proliferation of VSMCs was detected using the BeyoClick EdU‐488 kit (Beyotime) according to the manufacturer's protocol. VSMCs were seeded at a density of 5.0 × 10⁴ cells in 24‐well plates and treated with Ang II (1 µm) for 48 h. Then, the VSMCs were incubated with 10 µmol L⁻¹ EdU solution for 2 h at 37 °C and fixed with 4% PFA at 4 °C for 15 min. Finally, 100 µL click additive solution was added to the cells, following which nucleus staining with Hoechst 33342 was performed. The fluorescence signal was visualized using a fluorescence microscope.

Primary mouse WT and CCDC80^−/− VSMCs were isolated and seeded in six‐well plates. Straight‐line scratches were made on a monolayer of VSMCs using a 200 µL sterile pipette tip. The cells were washed with PBS and treated with or without Ang II (1 µm) containing normal growth medium for an additional 18 and 40 h. The number of VSMCs that migrated into the wound was visualized using a microscope (Thermo Fisher Scientific, Waltham, MA, USA).

The contractile capabilities of VSMCs were evaluated using a collagen gel‐based assay kit (Cell Biolabs, Cell contraction assay, Cat No. CBA‐201). WT and CCDC 80↗^−/− VSMCs were cultured in serum‐free medium for 24 h before seeding them into collagen gels. VSMCs were then harvested and resuspended in complete media at 2 × 10⁶ cells mL⁻¹. Collagen gels were prepared as one part of VSMC suspension and four parts cold Collagen Gel Working Solution. A 500 µL volume of the cell–collagen mixture was added to each well of the 24‐well plate, which was then incubated for 1 h at 37 °C. Polymerized gels were added as an additional 1 mL of complete media with Ang II (1 µm) and further incubated for 2 d at 37 °C to develop contractile stress. Then, the collagen gels were gently released using a pipette tip. Digital photographs of the collagen gel lattices were taken after 12 and 24 h, and the surface area of the gel was measured using Image J.

TUNEL staining was performed to detect cell apoptosis using the In Situ Cell Death Detection kit (Beyotime). Briefly, aortic tissue sections were fixed in 4% paraformaldehyde for 15 min and permeabilized with 0.25% Triton X‐100 for 10 min. Then, tissue samples were incubated with TUNEL reagent at 37 °C for 50 min. The nuclei were stained with DAPI for 10 min. Tissue sections were visualized using a fluorescence microscope (Leica, Germany).

Continuous data were expressed as the mean ± standard error of the mean (SEM), categorical variables were shown as the percentage. The data distribution normality was determined by the Shapiro‐Wilk test, and the equality of variances by F test. For comparisons between two‐group of normally distributed data, Student's t test was used to assess similar variances, or Welch's correction was used to assess unequal variances. The Mann‐Whitney U test was performed for variables not normally distributed. For multiple comparisons (more than two groups), the homogeneity of variance was assessed by the Brown‐Forsythe test. Comparisons were performed by a 1‐way and 2‐way ANOVA analysis followed by Tukey post hoc multiple comparisons test (equal variances) or a Welch ANOVA test followed by a post hoc analysis using the Tamhane T2 method (unequal variances). For data with nonnormally distributed variables, Kruskal‐Wallis test with Dunn's multiple comparisons test was used for multigroup comparisons. The survival curves were assessed using the Kaplan–Meier method and compared using log‐rank tests. A Fisher's exact test was used to compare bivariate categorical variables. The result was statistically significant if the p‐value was <0.05. Statistical analysis was performed using GraphPad Prism version 8.0 software.

The data that support the findings of this study are available from the corresponding author upon reasonable request.