Nox4 and circulating free MPO synergistically promote aortic aneurysm formation

Abstract

Myeloperoxidase (MPO)-induced oxidative stress plays an important role in the pathogenesis of abdominal aortic aneurysms (AAA). MPO protein released from activated inflammatory cells can circulate and be taken up into blood vessels, but the mechanisms that regulate vascular MPO uptake, and how this might impact abdominal aortic aneurysms (AAA) formation, are poorly understood. Given that MPO utilizes hydrogen peroxide (H₂O₂) to generate the oxidant hypochlorous acid (HOCl), and that Nox4 is a major producer of vascular H₂O₂, we hypothesized that vascular Nox4 promotes MPO uptake to cooperate in AAA formation. We found that aortic MPO uptake along with Nox4 expression and H₂O₂ levels were significantly increased in angiotensin II (AngII)-induced AAA mice. Increased aortic MPO uptake was also confirmed by ex vivo incubation of AngII- or calcium chloride (CaCl₂)-treated aorta. Notably, global deletion of Nox4 prevented AngII-induced aortic MPO uptake ex vivo, while Nox4 KO mice were protected against AngII-induced AAA formation. Furthermore, overexpression of human MPO by liver-specific AAV in MPO KO mice increased circulating MPO levels and aortic MPO accumulation while promoting AAA formation. Vascular Nox4 expression contributes to aortic MPO uptake, providing a novel mechanistic linkage whereby MPO and Nox4 cooperate to promote AAA.

Supplementary Information

The online version contains supplementary material available at 10.1038/s41598-026-40599-z.

Introduction

Ruptured AAA is the 15th leading cause of death in the United States and only ~ 25% of patients with aortic rupture survive to surgery, with an additional 50% mortality following surgery^1,2. AAA is an inflammatory disease, and enhanced levels of reactive oxygen species (ROS) produced during chronic inflammation were detected locally in human AAA^3,4. Targeting oxidative stress was shown to prevent AAA formation in animal models, implying a causal role in AAA pathogenesis^5–8. However, the enzymatic sources of ROS, and interactive mechanisms whereby they promote AAA, remain to be elucidated.

Myeloperoxidase (MPO), a peroxidase enzyme abundant in polymorphonuclear leukocytes (PMNs), and to a lesser extent in monocytes and macrophages, utilizes hydrogen peroxide (H₂O₂) to promote oxidative stress through several mechanisms, including production of tyrosyl radicals and hypochlorous acid (HOCl)^9–11. MPO also amplifies inflammation and can inactivate tissue inhibitor of metalloproteinase-1 and alpha-1 antiprotease to stimulate proteolytic activity and matrix degradation^12,13, key features of AAA. PMNs are abundant in human AAA tissues and play a crucial role in elastase-induced AAA formation in mice¹⁴. Importantly, PMNs isolated from patients with AAA are activated and release more MPO compared with controls^15–17. Also, PMNs trapped within AAA tissues and the associated thrombus release MPO, plasma levels of which are elevated in patients with AAA^15,16. Smoking, a major risk factor for AAA, is likewise associated with elevated plasma MPO levels¹⁸. Interestingly, extracellular MPO can be directly taken up by blood vessels¹⁹, but the mechanisms that control uptake and accrual of vascular MPO are largely undefined.

While our previous study demonstrated increased MPO in AAA tissues and showed that pharmacologic and genetic approaches targeting MPO can prevent AAA²⁰, the source of MPO, and mechanisms whereby MPO promotes AAA, remain to be determined. The NADPH oxidase family (Nox1-Nox5) is also a major source of ROS and associated with various cardiovascular diseases, including AAA²¹. While Nox1 and Nox2 generate O₂^- in a stimulus-dependent manner, Nox4 constitutively produces H₂O₂ and is regulated by inducible expression. Previous studies showed that deletion of Nox4, or smooth muscle cell (SMC)-specific overexpression of the H₂O₂-decomposing enzyme catalase, reduces AAA formation^22,23, suggesting a crucial role of Nox4 and its product H₂O₂ in AAA pathogenesis. Given that MPO utilizes H₂O₂ to generate the powerful oxidant HOCl, we hypothesized that Nox4 promotes MPO uptake or accumulation to induce vascular oxidative stress and AAA formation.

Here, we performed ex vivo and in vivo experiments to investigate whether Nox4 contributes to aortic MPO uptake in the context of AAA. We also investigated whether overexpression of circulating human MPO protein can increase AAA formation in MPO KO mice, which we previously showed were protected against AAA formation²⁰. Moreover, using Nox4 and Nox1 deficient mice, we investigated whether Nox enzymes may participate in aortic MPO uptake.

Results

Accumulation of MPO protein was increased in the aortas of AngII-induced AAA mice independent of changes in MPO mRNA expression

Consistent with previous studies, AngII infusion significantly increased AAA formation in LDLR KO mice (Fig. 1A-C). MPO accumulation in conjunction with macrophage (Mac-3 positive) and neutrophil (Ly6G positive) infiltration were also significantly increased throughout the aortic wall in AngII-induced AAA mice (Fig. 1D), as reported previously²⁴. Increased aortic MPO accumulation in AAA mice was also confirmed by elevated MPO activity (Fig. 1E). Mature macrophages are reported to express very low levels of MPO, raising the possibility that the tissue-resident macrophages could have reactivated MPO gene expression²⁴. However, we did not detect increases in MPO mRNA levels in AngII-induced AAA, despite a tenfold increase in expression of the macrophage marker F4/80 compared with saline control (Fig. 1F), suggesting that macrophage MPO expression is not reactivated in the context of AAA. Thus, immunostaining data (Fig. 1D) raised the possibility that MPO protein is taken up into in the blood vessel wall during AAA formation.

Fig. 1.

Accumulation of MPO protein was increased in aortas of AngII-induced AAA mice independent in changes in MPO mRNA expression. AngII or saline (control) was infused for 3 weeks via osmotic minipump in LDLR KO mice. (A) Representative images of AAA from saline or AngII-infused mice. (B) AAA incidence. (C) Aortic diameter. (D) Representative histology images of MPO, Mac-3 and Ly6G (red scale bar = 20 µm, white scale bar = 100 µm). (E) MPO activity. (F) mRNA expression of MPO and F4/80 in aortic tissues. Data are represented as mean ± SEM. ns = non-significant, **p < 0.01.

Aortic MPO uptake was increased by ex vivo and in vivo AngII treatment

To investigate whether MPO uptake may be enhanced during AAA formation, abdominal aortas from LDLR KO mice were incubated with saline or AngII (100 nM) for 12 h ex vivo. Aortas were then exposed to vehicle or purified human MPO for 2 h, and MPO activity was assayed (Fig. 2A). Notably, AngII pretreatment ex vivo for 12 h significantly enhanced aortic MPO uptake (Fig. 2B). No cytotoxicity was observed as determined by lactate dehydrogenase (LDH) assay using conditioned medium (Supplemental Fig. S1).

Fig. 2.

Aortic MPO uptake was increased by ex vivo and in vivo AngII treatment. (A) Experimental scheme of aortic MPO uptake assay using AngII and purified human MPO protein. (B) Aortic MPO activity in response to AngII treatment ex vivo. (C) MPO activity in aortas harvested from AngII-infused mice for 3 days. Data are represented as mean ± SEM. ns = non-significant, *p < 0.05, **p < 0.01.

MPO taken up by aorta amplifies inflammation, inactivate tissue inhibitor of metalloproteinase-1²⁵ and alpha-1 antiprotease²⁶ to stimulate proteolytic activity and matrix degradation, key features of AAA. To investigate whether aortic MPO uptake occurs in the early phase of AAA development, we infused saline or AngII for 3 days (a time frame which precedes AAA formation) into LDLR KO mice, removed the aortas, and exposed them ex vivo to vehicle or MPO as mentioned above. Infusion of AngII in vivo likewise enhanced aortic MPO uptake ex vivo (Fig. 2C). These data suggest that MPO uptake into the aorta is stimulated early during the development of AngII-induced AAA.

Aortic Nox4 expression was induced by AngII and required for MPO uptake ex vivo

Nox4 expression was reported to be upregulated in both AngII- and CaCl₂-induced AAA tissues²⁷. Unlike Nox1 or Nox2, which primarily produce O₂^-, Nox4 primarily produces H₂O₂, which is in turn used by MPO to produce HOCl. Thus, we hypothesized that vascular Nox4-derived H₂O₂ might functionally partner with MPO by promoting both MPO vascular uptake and production of damaging oxidants. Interestingly, ex vivo treatment with AngII for 12 h resulted in upregulation of Nox4 mRNA (Fig. 3A) in conjunction with increased H₂O₂ levels (Fig. 3B) in aortas from LDLR KO mice, while Nox4 protein was strongly upregulated after AngII infusion for 14 days in vivo (Fig. 3C,D). Specificity of Nox4 antibody was verified using the aortas from Nox4 KO mice by immunohistochemical analysis (Supplemental Fig. S2). Importantly, deletion of Nox4 (a H₂O₂ producer), but not Nox1 (an O₂^- producer), blocked AngII-induced aortic MPO uptake ex vivo (Fig. 3E) despite Nox1 and Nox2 expression was similarly increased in AngII-induced AAA tissues (Supplemental Fig. S3), suggesting a specific dependence on Nox4.

Fig. 3.

Aortic Nox4 expression was induced by AngII and required for MPO uptake ex vivo. (A–C) AngII or saline (control) was infused for 3 weeks via osmotic minipump in LDLR KO mice. (A) Aortic Nox4 mRNA expression. (B) Aortic H₂O₂ levels (normalized by tissue weight). (C) Representative immunostaining images of Nox4 (white scale bar = 50 µm, red scale bar = 20 µm). (D) CaCl₂ or saline (sham) was applied to the infrarenal aorta of C57Bl/6 mice. Representative immunofluorescence images of α-smooth muscle actin and Nox4 (white scale bar = 100 µm). (E) Aortic rings harvested from WT, Nox1 KO and Nox4 KO mice were pre-incubated with saline ( −) or AngII ( +) for 12 h, and then treated with or without MPO for 2 h. After washing, MPO activity was then quantified and compared between the different groups. Data are represented as mean ± SEM. ns non-significant, ***p < 0.001.

MPO uptake was co-localized with expression of Nox4 in the CaCl₂-induced AAA model

To determine whether increased aortic MPO uptake also occurs in the context of a different AAA model, we examined aortic Nox4 expression 3 days after in vivo application of CaCl₂ to the abdominal aorta. Aortic Nox4 mRNA expression was significantly upregulated at this early time point in abdominal, but not thoracic, aorta compared to sham (saline control) (Fig. 4A). Concomitantly, CaCl₂ application was associated with increased MPO protein in the abdominal aorta as detected by immunohistochemical analysis (Fig. 4B). CaCl₂ application in abdominal aorta increased aortic diameter in abdominal, but not in thoracic aorta (Supplemental Fig. S4). We also performed ex vivo aortic MPO uptake assay using CaCl₂-treated aorta (Fig. 4C). Interestingly, uptake of MPO ex vivo was increased in abdominal, but not thoracic, aortas of CaCl₂-treated mice (Fig. 4D,E). These data suggest that MPO uptake co-localizes with induction of Nox4 and aneurysm formation in the CaCl₂ model.

Fig. 4.

MPO uptake was co-localized with expression of Nox4 in the CaCl₂-induced AAA model. (A,B) CaCl₂ or saline (sham) was applied to the infrarenal aorta of C57Bl/6 mice, and three days later, aortas were harvested and separated into thoracic and abdominal aortic portions for the indicated studies. (A) Nox4 mRNA expression in thoracic and abdominal aorta. (B) Representative MPO immunostaining images in abdominal aorta. (C) Experimental scheme of aortic MPO uptake assay in CaCl₂-treated aorta. (D) Aortic MPO activity in abdominal aorta. (E) Aortic MPO activity in thoracic aorta. Data are represented as mean ± SEM. ns non-significant, *p < 0.05, **p < 0.01, ***p < 0.001.

Aortic MPO accumulation and AAA formation in response to AngII were diminished in Nox4 deficient mice

Since Nox4 KO mice are protected against AAA formation, we next investigated the impact of Nox4 deletion on aortic MPO accumulation. MPO protein was significantly reduced in aortas of Nox4 KO mice infused with AngII (Fig. 5A & Supplemental Fig. S5). Deletion of one or both alleles of Nox4 protected against AngII-induced AAA formation (Fig. 5B,C), in conjunction with reduced macrophage infiltration, elastin degradation and MPO accumulation (Fig. 5D), while having no effect on blood pressure (data not shown). We observed mortality in several mice due to aortic rupture; two in the LDLR^-/- group, one in the LDLR^-/-Nox4^+/- group and one in the LDLR^-/-Nox4^+/- group.

Fig. 5.

Aortic MPO accumulation and AAA formation in response to AngII were diminished in Nox4 deficient mice. AngII or saline (control) was infused for 3 weeks via osmotic minipump in LDLR/Nox4 double KO or LDLR KO (control) mice. (A) Representative histology images of Nox4 and MPO (white scale bar = 50 µm, red scale bar = 20 µm). (B) AAA incidence. (C) Aortic diameter. (D) Representative histology images; H&E, Mac-3, VVG (red scale bar = 200 µm, black scale bar = 20 µm). Data are represented as mean ± SEM. *p < 0.05.

Overexpression of human MPO in MPO KO mice increased circulating MPO protein levels, aortic MPO uptake and AAA formation

MPO KO mice were previously reported to be resistant to AAA formation²⁰. MPO is produced and released by inflammatory cells and circulates systemically. Circulating MPO protein levels serve as an important inflammatory biomarker for various cardiovascular diseases, including AAA. To determine whether circulating MPO protein, in the absence of inflammatory cells, can directly promote AAA formation, human MPO (hMPO) was overexpressed in the liver of MPO KO mice. Liver-specific AAV8 containing human MPO was injected into MPO KO mice, after which AngII was infused for 3 weeks. We detected significant increases in human MPO protein levels in liver (Fig. 6A), plasma (Fig. 6B) and aorta (Fig. 6C) by Western blotting using a human MPO-specific antibody while aortic Nox4 expression was not changed by hMPO overexpression (Supplemental Fig. S6). AAA incidence (Fig. 6D) and aortic diameter (Fig. 6E,F) were increased, in conjunction with increased thrombus formation (Fig. 6G, upper panel) and elastin degradation (Fig. 6G, lower panel), in AAV8-MPO injected mice as compared to AAV8-GFP. No rupture was observed in AAV injected mice. Mitochondrial dysfunction has been implicated as a key contributor to the pathogenesis and progression of aortic aneurysm^28,29. Consistently, we observed a reduction in mitochondrial mass in the aortas of AAV-hMPO-injected mice compared with AAV-control mice (Supplemental Fig. S7). There was no significant difference in blood pressure between the groups (data not shown) or systemic inflammation as determined by cytokine profiling in plasma (e.g., IL-1β, TNFα, MCP-1, Supplemental Fig. S8). These results provide evidence that MPO detected in MPO KO mice is derived from the liver and not associated with murine inflammatory cells, demonstrating that circulating human MPO can directly contribute to AAA formation.

Fig. 6.

Overexpression of human MPO in MPO KO mice promoted circulating MPO protein levels, aortic MPO uptake and AAA formation. Liver-specific AAV8 containing human MPO or control AAV8 was administered into MPO KO mice for 2 weeks and AngII was infused by osmotic minipump for 3 weeks. MPO protein expression was measured in liver (A), plasma (B) and aorta (C). (D) AAA incidence. (E) Aortic diameter. (F) Representative in situ images of AAA. (G) Representative histology images; H&E, VVG (white scale bar = 200 µm, red scale bar = 20 µm). Data are represented as mean ± SEM. *p < 0.05, **p < 0.01.

Discussion

MPO, a leukocyte enzyme that utilizes H₂O₂ to promote oxidative stress and inflammation, plays a key role in the pathogenesis of AAA^14–16. We previously reported that MPO gene deletion prevents AAA formation in animal models of AAA²⁰. However, the sources of MPO in AAA, and the mechanisms that regulate MPO-mediated tissue damage, remain to be determined. MPO circulates in the blood and has been demonstrated to be taken up into blood vessels, though the mechanisms and consequences of vascular MPO uptake are largely unknown. Here, we found that Nox4, a unique high-output enzymatic producer of H₂O₂ that is highly expressed in vascular cells, promotes aortic MPO uptake, and that circulating MPO is capable of accumulating in the aortic wall to promote AAA formation.

MPO is highly expressed in PMNs and, to a lesser extent, in macrophages, both of which participate in AAA formation. Upon PMN activation, preformed MPO is rapidly released from storage granules, and Nox2 generates O₂^.-, which is in turn is locally dismutated to H₂O_2. MPO then converts H₂O₂ to HOCl to promote oxidative stress. We previously reported that deletion of p47phox, a key component of Nox2, attenuates AngII-induced AAA formation in ApoE-deficient mice³⁰, suggesting the crucial role of Nox2 in AAA formation. Subsequently, free MPO can then circulate in the blood and/or be taken up by tissues and cells. Notably, MPO has been detected in human AAA tissues, where it appears to be particularly enriched in the intramural thrombus³. How MPO released from inflammatory cells accumulates in AAA tissues, and the pathological consequences of aortic MPO accumulation, are unknown. In this study, we provide new data that forced expression of human MPO in MPO KO mice by liver-specific MPO delivery increased levels of liver, plasma and aortic MPO protein, thereby promoting AAA formation. This suggests that circulating MPO protein, independent of inflammatory cells, is capable of exacerbating AAA pathogenesis, although the relative contribution of cell-derived versus circulating MPO in AAA remains to be determined. MPO uptake in the aorta during AAA formation is further supported by our data from ex vivo MPO uptake assay. Aortic uptake of purified MPO protein added to the medium was enhanced by ex vivo AngII treatment or short-term in vivo infusion. Similarly, ex vivo MPO uptake was also selectively increased in CaCl₂-treated abdominal aorta. These results strongly support the hypothesis that circulating MPO can be taken up by the aorta in the context of AAA.

MPO possesses a high affinity for extracellular matrix (ECM) proteins and negatively-charged glycosaminoglycan chains due to the high abundance of positively-charged Lys and Arg residues on the protein surface^31,32. MPO protein has also been reported to be localized to the shoulder regions of atherosclerotic lesions, which are especially prone to rupture and thrombus formation³³. Cai et al. reported that MPO-generated HOCl induced ECM modification by interacting with ECM proteins in a site-specific manner and increased MPO adhesion to human coronary VSMCs in vitro, suggesting a potential mechanism linking MPO-oxidation with matrix interaction³⁴. However, how blood vessels attract and take up the circulating MPO to promote AAA formation remains unknown. Here, we provide evidence of cooperation between Nox4 expressed in the aorta and MPO-mediated AAA formation. Our data showed that AngII-induced aortic MPO ex vivo was abrogated by deletion of Nox4, but not Nox1, suggesting a specific dependence on aortic Nox4 expression. Induction of oxidant stress via the Nox family is one of the earliest signaling events induced by AngII in the aorta. Among the Nox isoforms, Nox4 is abundantly expressed in inflammatory and vascular cells, particularly VSMC³⁵ (Fig. 3C,D) and is upregulated by vascular injury³⁶ and inflammatory cytokines³⁷. Smoking, a major risk factor for AAA, is associated with both upregulated VSMC Nox4 expression³⁸ and elevated plasma MPO levels¹⁸. In VSMC, Nox4 mRNA expression level is ~ 20 times higher than that of Nox2, and up to 700 times higher than Nox1³⁹. Nox4 expression was also reported to be upregulated in both AngII- and CaCl₂-induced AAA tissues³⁰, and genetic deletion of Nox4 prevented AngII-induced AAA formation (Fig. 5). Unlike Nox1 or Nox2, which primarily produce O₂^.-, Nox4 primarily produces H₂O₂, and SMC-specific overexpression of catalase, a H₂O₂-decomposing enzyme, prevented AAA formation²³, suggesting a key role for VSMC-derived H₂O₂ in AAA. Interestingly, Nox4 expression was also reported to be upregulated by TFG-β, which plays a key role in Marfan syndrome (MFS)-associated thoracic aortic aneurysm (TAA) formation. Increased Nox4 expression and H₂O₂ levels were observed in MFS tissues and VSMCs, and MPO-derived oxidative stress contributed to TAA pathogenesis⁴⁰, further supporting a cooperative role of Nox4 with MPO in aortic aneurysm formation. In this study, Nox4 expression and concomittant MPO uptake were not increased in thoracic aorta in CaCl₂ application model. However, it remains to be determined in the AngII infusion model whether Nox4 expression in other aortic regions such as thoracic, ascending aorta and aortic root also contributes to increased MPO uptake and associated pathological changes. Further studies are required in the future to determine cell type-specific (e.g., endothelial, VSMC, inflammatory cells) source(s) of Nox4 leading to MPO uptake in the context of aortic aneurysms. Notably, mitochondrial dysfunction induced by Nox4 upregulation and subsequent astrocytic ferroptosis mediated by MPO in the hippocampus has been shown to contribute to the progression of Parkinson’s disease⁴¹, suggesting a cooperative role of Nox4 and MPO in other inflammatory disease. Given that mitochondrial dysfunction is also a critical driver of aortic aneurysm development^28,29, our observation of mitochondrial mass loss in AAV-hMPO-injected aortas suggests a potential mechanistic link between MPO activity and mitochondrial impairment, warranting further investigation.

Several limitations should be acknowledged. First, MPO activity was measured using a commercially obtained fluorescent MPO assay kit, which detects conversion of a non-fluorescent to a fluorescent compound in the presence of H₂O₂. The reagent may also react with other heme-containing proteins that catalyze oxidation of different substrates. However, we previously demonstrated that quantification of dityrosine, a specific marker of MPO activity, by LC/MS/MS correlated with data from the fluorescent MPO assay kit²⁰. Second, only commercially available KO mouse strains were used in this study, rather than in-house bred lines. Therefore, data derived from different mouse strains (e.g., mRNA expression levels) cannot be directly compared. This limitation is particularly important because the mice were not littermates or backcrossed onto a uniform genetic background, potentially introducing strain-specific variability unrelated to the targeted gene deletion. Third, AAV8-hMPO administration was tested solely in the AngII infusion model and not in the CaCl₂-induced model. Since no single animal model fully recapitulates human AAA pathology, future studies employing multiple models are warranted. Fourth, blood pressure was measured using a tail-cuff method; therefore, validation with radiotelemetry will be required. Fifth, O₂•⁻ generated by Nox2 can also be converted to hydrogen peroxide H₂O₂, a process not examined in this study. Finally, future studies using RNA sequencing and proteomic analyses will be necessary to elucidate the molecular mechanisms whereby MPO accumulation within the aortic wall contributes to AAA formation.

In conclusion, this study demonstrates that circulating MPO protein uptake by blood vessels, independent of inflammatory cell recruitment, is sufficient to promote AAA formation. We also provide evidence of a novel mechanism whereby Nox4 expressed in the aorta mediates vascular MPO uptake. Cooperation of Nox4 and MPO may contribute to oxidative stress during AAA formation, and perhaps in other conditions where the two oxidant-generating enzymes have been implicated in disease pathophysiology.

Materials and methods

Mice

MPO KO, Nox4 KO, Nox1 KO, low density lipoprotein receptor (LDLR) KO, and C57Bl/6 mice were obtained from Jackson Laboratories. For AngII infusion model, LDLR KO background mice were used because hypercholesterolemia significantly increases the incidence of AngII-induced AAA formation, independent of elevated blood pressure⁴². MPO KO and Nox4 KO mice were bred with LDLR KO mice to obtain littermate WT, heterozygous and homozygous mice in the LDLR KO background. Mice were anesthetized by isoflurane vaporizer (0.5–1.0 L/min for oxygen flowmeter, 4–5% for induction and 1–3% for maintenance, EZ Anesthesia Systems) and euthanized with intraperitoneal pentobarbital 150 mg/kg, inhaled anesthesia (isoflurane) followed by carbon dioxide (CO₂) narcosis and cervical dislocation or bilateral thoracotomy, in accordance with AVMA Panel 2007 recommendations and institutional IACUC guidelines. All mice were randomized to different treatment groups to minimize experimental variability, and surgeons were blinded to group allocation to prevent accidental or selection bias. Only male mice were used in this study because, in humans, AAA are more common in males compared with females. There is also a significant sex difference of AAA incidence in males versus females in the AngII-induced animal model of AAA (80–100% vs ~ 20%, respectively)⁴³. All animal experiments were performed at animal facilities of the Medical College of Georgia at Augusta University. Animal experimental protocols were approved by the Institutional Animal Care and Use Committee at the Medical College of Georgia at Augusta University and complied with National Institute of Health guidelines. All experiments were carried out in accordance with institutional biosafety and chemical safety guidelines and regulations.

AngII infusion in LDLR KO mice

LDLR KO mice (8- to 12-week-old male) received saline (placebo control) or AngII (1,000 ng/kg/min, Enzo Life Sciences). Ang II was infused through osmotic mini-pumps (ALZET Model 2004, Cupertino, CA) inserted into the subcutaneous tissue in the interscapular area under parenteral anesthesia as described previously^20,44–46. Mice were euthanized three weeks after minipump implantation, and abdominal aortic outer diameter was measured via microscopy. Aortic tissues, liver and blood were collected from mice for further analysis (e.g., PCR, Western blot, histology as described below). AAA was defined as local dilation of the abdominal aorta that is at least 1.5 times the size of the reference aortic diameter. In separate experiments (ex vivo aortic MPO uptake assay as described below), aortic tissues were harvested 3 days after AngII infusion (a time frame which precedes AAA formation).

CaCl₂ application in WT mice

CaCl₂ application model of AAA (using 0.5 mol/L of CaCl₂, Sigma-Aldrich) was performed as described previously^20,44–46. In brief, following laparotomy, saline (sham control) or 0.5 mol/L of CaCl₂ was applied to the infrarenal aortic adventitial surface for 15 min, followed by rinsing with 0.9% sterile saline and surgical closure. After three days (a time frame which precedes AAA formation), mice were anesthetized, and aortic tissues were collected for ex vivo MPO uptake assay as described below.

Blood pressure measurement

Blood pressure was measured using a previously validated tail cuff method (Coda 6, Kent Scientific, Torrington, CT). Mice were conditioned to the instrument and procedure for 5 consecutive days prior to pump implantation. To ensure a more robust estimation of systolic blood pressure (SBP), we used the interquartile mean of SBP measurements achieved through 30 measurement cycles every other day.

MPO activity measurement

Aortic MPO uptake levels were measured by a commercially available mouse MPO detection kit (Immunochemistry Technologies), as previously reported⁴⁷, per manufacturer’s protocol.

Immunohistochemistry

Paraffin-embedded aortic tissue sections were stained with hematoxylin and eosin (H&E), Verhoeff van Gieson (VVG), Mac-3 (BD Pharmingen), MPO (Abcam), Nox4 (Novus Biologicals), and HistoMouse-SP (Invitrogen) or DAB Substrate (Vector Labs) kits were used for visualization. To quantify the immunostaining data, total stained area in the media and adventitial region was analyzed using Image-Pro Plus Software as previously described⁴⁴.

Western blotting

Protein extraction and Western blotting were performed as described previously^20,44–46. Antibodies used in Western blotting were human MPO (Meridian Bioscience), transferrin (Abcam), and GAPDH (Invitrogen).

Quantitative PCR

Total RNA was extracted from aortic and liver tissues with QIAzol Lysis Reagent, and purified with RNeasy Lipid Tissue Mini Kit (Qiagen). Real-time quantification of mRNA levels of the genes of interest was performed using Brilliant II SYBR Green QPCR Master Mix (Agilent Technologies) per manufacturer’s instructions. Quantification of mitochondrial DNA was performed with SYBR green qRT-PCR kit (abm). Relative mitochondrial mass was determined by the ratio of mitochondrial CoxII gene to nuclear β-globin gene. Normalized Ct values were subjected to statistical analysis and fold difference was calculated by ∆∆ Ct method as described previously²⁰.

Measurement of H₂O₂ levels

Aortic H₂O₂ levels were measured by Amplex Red assay kit per manufacturer’s protocol (Invitrogen).

Ex vivo MPO uptake assay

To quantify aortic MPO uptake, we performed an ex vivo assay using isolated aorta and purified MPO derived from human PMNs (Sigma-Aldrich) as previously reported with slight modifications¹¹. Briefly, aortas were harvested from LDLR KO mice and adventitial and perivascular adipose tissues were removed. Aortas were cut into 3–4 mm rings, and pre-incubated in DMEM medium without FBS, phenol red and antibiotics. Then, aortic rings were treated with or without AngII (100 nM) for 12 h, and further incubated in the presence or absence of purified human MPO (65 nM) for 2 h. After incubation, aortic tissues were harvested to measure MPO activity and Nox4 gene expression as described above. In separate experiments, aortas were harvested after CaCl₂ application in C57Bl/6 mice or AngII infusion in LDLR KO mice for 3 days, and aortic rings were exposed to vehicle or purified human MPO for 2 h, and then washed and processed for MPO activity assay. Cytotoxicity was evaluated using LDH Cytotoxicity Assay Kit (Cayman Chemical).

Adeno-associated virus (AAV) 8-mediated human MPO delivery

AAV8 (a liver-specific serotype) containing human MPO (100 µl of viral vector at a titer of 1 × 10¹³ genome copies/ml, Vigene) was delivered into MPO KO/LDLR double KO mice via retro-orbital injection per manufacturer’s instructions. Two weeks after injection, AngII was infused to induce AAA formation as described above. Three weeks after AngII infusion, mice were sacrificed to evaluate AAA formation. Human MPO protein expression was measured in liver, plasma and aorta using human MPO specific antibody and Western blotting.

Profiling of pro-inflammatory cytokines

Mouse plasma cytokine profiling was performed at Immune Monitoring Core Facility at Medical College of Georgia. BioLegend Mouse Inflammation 13-Plex Panel FbBA228ser was used to quantify key pro-inflammatory and anti-inflammatory cytokines using a Luminex 200 instrument, and cytokine concentrations were calculated based on standard curves generated with recombinant protein standards.

Statistical analysis

All statistical analyses were performed using Graphpad Prism version 10.2.2 Software (GraphPad Software, Inc., USA). Results are expressed as mean ± SEM. Differences between two groups were analyzed by student’s t-test. Multiple group datasets were evaluated for normality, and differences were analyzed by one-way ANOVA followed by Bonferroni post-hoc analysis. p values less than 0.05 were considered to be significant.

Author contributions

H.W.K. and N.L.W. conceived and designed research, P.C., T.H., D.K., B.G., M.O., L.R., T.W.B., J.G., S.C.,M.S., R.C., N.C., P.V., H.S., H.W.K. performed experiments, P.C., T.H., D.K., B.G., H.W.K. analyzed data, P.C., T.H., D.J.F., Y.P.Z., A.G., R.L., H.W.K., N.L.W. interpreted results of experiments, P.C., T.H., D.K., B.G., M.O., H.W.K. prepared figures, H.W.K. drafted manuscript, B.G., Y.P.Z., A.G., R.L., H.W.K., N.L.W. edited and revised manuscript, H.W.K. and N.L.W. approved final version of manuscript.

Funding

This study was funded by grants AG076235 (NIH), 971459 (AHA) and 863622 (AHA).

Data availability

Datasets analyzed in this study available from the corresponding authors upon reasonable request.

Declarations

Competing interests

The authors declare no competing interests.

Ethical statement

The work was conducted in accordance with ARRIVE guidelines 2.0 (Animal Research: Reporting of In Vivo Experiments). All animal experiments were performed at animal facilities of the Medical College of Georgia at Augusta University. Animal experimental protocols were approved by the Institutional Animal Care and Use Committee at the Medical College of Georgia at Augusta University and complied with National Institute of Health guidelines. All experiments were carried out in accordance with institutional biosafety and chemical safety guidelines and regulations.