Alterations in Adipose Tissue and Adipokines in Heterozygous APE1/Ref-1 Deficient Mice
Article information
Abstract
Background
The role of apurinic/apyrimidinic endonuclease 1/redox factor-1 (APE1/Ref-1) in adipose tissue remains poorly understood. This study investigates adipose tissue dysfunction in heterozygous APE1/Ref-1 deficiency (APE1/Ref-1+/-) mice, focusing on changes in adipocyte physiology, oxidative stress, adipokine regulation, and adipose tissue distribution.
Methods
APE1/Ref-1 mRNA and protein levels in white adipose tissue (WAT) were measured in APE1/Ref-1+/- mice, compared to their wild-type (APE1/Ref-1+/+) controls. Oxidative stress was assessed by evaluating reactive oxygen species (ROS) levels. Histological and immunohistochemical analyses were conducted to observe adipocyte size and macrophage infiltration of WAT. Adipokine expression was measured, and micro-magnetic resonance imaging (MRI) was used to quantify abdominal fat volumes.
Results
APE1/Ref-1+/- mice exhibited significant reductions in APE1/Ref-1 mRNA and protein levels in WAT and liver tissue. These mice also showed elevated ROS levels, suggesting a regulatory role for APE1/Ref-1 in oxidative stress in WAT and liver. Histological and immunohistochemical analyses revealed hypertrophic adipocytes and macrophage infiltration in WAT, while Oil Red O staining demonstrated enhanced ectopic fat deposition in the liver of APE1/Ref-1+/- mice. These mice also displayed altered adipokine expression, with decreased adiponectin and increased leptin levels in the WAT, along with corresponding alterations in plasma levels. Despite no significant changes in overall body weight, microMRI assessments demonstrated a significant increase in visceral and subcutaneous abdominal fat volumes in APE1/Ref-1+/- mice.
Conclusion
APE1/Ref-1 is crucial in adipokine regulation and mitigating oxidative stress. These findings suggest its involvement in adipose tissue dysfunction, highlighting its potential impact on abdominal fat distribution and its implications for obesity and oxidative stress-related conditions.
INTRODUCTION
Adipose tissue is central to energy homeostasis and metabolic regulation, and it has been extensively researched to understand the molecular mechanisms underlying obesity and its complications, including the roles of adipokines, oxidative stress, and genetic factors [1]. Adequate amount and proper functioning of adipose tissue are essential for physical health. However, excessive fat accumulation due to an imbalance between energy intake and expenditure can increase the risk of serious health problems such as high blood pressure, high cholesterol, cardiovascular disease, dementia, and certain cancers [2,3]. Adipose tissue dysfunction can cause adipocyte hypertrophy, hypoxia, macrophage infiltration, cellular stress, inflammation, ectopic fat accumulation, adipokine dysregulation, and insulin resistance [4,5]. These issues can increase the risk of developing the aforementioned diseases due to excessive fat tissue accumulation, regardless of overall body weight. Recent studies suggest that the onset or severity of these diseases may not be solely correlated with obesity, highlighting the significance of adipose tissue dysfunction [4,6,7].
Oxidative stress, the imbalance between reactive oxygen species (ROS) production and detoxification, is a key factor in obesity-related metabolic dysregulation. Excessive ROS levels lead to adipose tissue dysfunction, promoting inflammation, insulin resistance, and adipose tissue expansion, which underscores the need to understand ROS regulation in adipose tissue [8]. Adipose tissue dysfunction disrupts adipokine regulation, leading to imbalances in adiponectin and leptin levels, which contribute to the development of obesity-related complications. Adiponectin has anti-inflammatory, anti-fibrotic, antioxidant, and anti-atherogenic properties, inversely correlating with body weight changes [9,10]. Conversely, elevated leptin levels in obese individuals are linked to leptin resistance and various metabolic disorders [11-13]. Adipose tissue dysfunction also causes lipid efflux from the circulatory system, leading to lipid accumulation in non-adipose tissues and subsequent ectopic fat deposition. Consequently, adipose tissue contributes to the formation of fatty liver and accumulates around the abdominal internal organs, exacerbating abdominal obesity [7].
Apurinic/apyrimidinic endonuclease 1/redox factor-1 (APE1/Ref-1) is a multifunctional protein that is crucial in DNA repair and redox regulation. Its functions include DNA base repair, redox status regulation, transcription factor activity modulation, and ROS suppression [14,15]. Emerging evidence suggests that APE1/Ref-1 plays a significant role in both adipogenesis and neurogenesis, with its redox activity being particularly crucial for these functions [16,17]. Specifically, APE1/Ref-1 has been shown to suppress the expression of key adipogenic transcription factors such as CCAAT/enhancer-binding protein (C/EBP)-α, peroxisome proliferator-activated receptor (PPAR)-γ, and adipocyte protein 2 (aP2) in vitro [16], and may be affected by adipokines. However, while the inhibitory effect of APE1/Ref-1 on adipocyte differentiation has been established in vitro, its impact on adipocyte differentiation, adipokine production, and the interplay between these factors in vivo remains unknown.
This study investigates the impact of heterozygous APE1/Ref-1 deficiency on adipose tissue function and distribution in mice. We hypothesize that APE1/Ref-1 deficiency would lead to adipose tissue dysfunction, characterized by increased oxidative stress, abnormal adipocyte morphology and distribution, and disrupted adipokine expression, ultimately contributing to increased abdominal adiposity. By elucidating the role of APE1/Ref-1 in adipose tissue function, this study aims to provide novel insights into the mechanisms underlying adipose tissue dysfunction.
METHODS
Generation of heterozygous APE1/Ref-1 deficiency mice
We generated heterozygous APE1/Ref-1 mice using Clustered regularly interspaced short palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9) technology at Macrogen (Macrogen, Seoul, Korea). APE1/Ref-1 was targeted for knockout using two single guide RNAs (sgRNAs), with in vitro validation through digestion assays and polymerase chain reaction (PCR). Mouse genotyping involved PCR amplification using specific primers. Mouse embryos, which were injected with the Cas9 protein and sgRNA molecules, were implanted into recipient mice, resulting in chimeric mice backcrossed with C57BL6/ J mice. PCR conditions included a sequence of temperature cycles and visualization on a 1% agarose gel. The mice were housed under specific pathogen-free conditions at Chungnam National University Hospital, with a 12-hour light/dark cycle at 24°C and access to distilled water and a chow diet. Male mice at 20 weeks of age were used in the experiment. Body weight was measured before euthanasia. All experimental protocols were approved by the Institutional Animal Care and Use Committee of Chungnam National University Hospital (approval numbers: CNUH-019-A0020, CNUH-020-A0022).
Measurement of ROS and superoxide production in white adipose tissue and liver
Total ROS production in white adipose tissue (WAT) and liver was quantified using a dichlorofluorescein assay kit (Abcam, Cambridge, UK) following the manufacturer’s protocol. The epididymal white adipose and liver (50 mg) tissues were homogenized in 1 mL of phosphate-buffered saline (PBS) (Welgene, Gyeongsan, Korea) using a homogenizer (Bertin Technologies, Montigny-le-Bretonneux, France), followed by centrifugation at 10,000 ×g for 5 minutes to collect the supernatant, devoid of fat. Next, 50 μL of the sample and 50 μL of the catalyst reagent were added to a 96-well black plate and incubated for 5 minutes. Subsequently, 100 μL of dichlorodihydrofluorescein solution was added, and the mixture was incubated for 30 minutes in the dark. Fluorescence was measured using a Glomax microplate reader (Promega, Madison, WI, USA) at an excitation wavelength of 475 nm and an emission range of 500 to 550 nm. Superoxide production in WAT and liver was assayed using lucigenin-enhanced chemiluminescence [18]. Briefly, a darkadapted lucigenin solution (50 μM) was prepared in Krebs– 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) buffer with the following composition: NaCl 100 mM, KCl 4.7 mM, CaCl2 1.9 mM, MgSO4 1.2 mM, K2HPO4 1.03 mM, NaHCO3 25 mM, and Na-HEPES 20 mM (pH 7.4). The solution was bubbled with a mixture of 95% O2 and 5% CO2 for 1 hour at 37°C prior to experimentation. Lucigenin solution (100 μL) was then added to each well of an opaque 96-well plate. Tissue samples (2 to 3 mm) were placed in each well, and baseline luminescence was recorded using a luminometer. Superoxide production was initiated by adding nicotinamide adenine dinucleotide phosphate (NADPH) to the wells. The relative luminescence unit was calculated by subtracting the control value from the experimental value and recording the differences. The tissue weight was determined after drying the samples at 60°C for 48 hours, and the results were expressed as the relative light unit per milligram of tissue.
Oil Red O staining
The liver samples were fixed in neutral buffered 10% formalin and then incubated in 30% sucrose (Sigma Aldrich, St. Louis, MO, USA) for cryosectioning [19]. The sections were treated with propylene glycol (Scytek, Logan, UT, USA) for 5 minutes, Oil Red O solution (Scytek) for 10 minutes, and 85% propylene glycol for 1 minute. After washing, the sections were counterstained with Mayer’s hematoxylin for 1 minute. All images were obtained using a Motic Easyscan Pro 6 (Motic, Hong Kong, China) and analyzed with ImageJ software (National Institutes of Health, Bethesda, MD, USA).
Immunohistochemistry
The epididymal WAT were fixed in neutral buffered 10% formalin and embedded in paraffin blocks for sectioning. The sections were treated with a peroxide blocking solution (Agilent Technologies, Santa Clara, CA, USA) to remove endogenous peroxidase activity. They were then incubated overnight at 4°C with anti-F4/80 (1:100; Cell Signaling, Danvers, MA, USA), anti-leptin (Abcam, 1:100), and anti-adiponectin (Abcam, 1:100). After washing twice in PBS, the sections were incubated with the Dako EnVision detection system (Agilent Technologies) for 30 minutes at room temperature. Following another wash in PBS, the sections were developed using diaminobenzidine (DAB) substrate for 3 minutes and counterstained with Mayer’s hematoxylin. All images were obtained using a Motic Easyscan Pro 6 (Motic) and analyzed with ImageJ software.
Blood chemical assay
After fasting for 12 hours, blood samples were collected from the hearts of the mice and transferred into heparin tubes. The samples were centrifuged at 2,000 ×g for 10 minutes to obtain plasma, which was then analyzed for blood chemistry. The total cholesterol, low-density lipoprotein cholesterol, high-density lipoprotein cholesterol, triglyceride (TG), and glucose levels were measured at the KPNT Analysis Service Center (KPNT, Cheongju, Korea).
Quantitative real-time reverse transcription-polymerase chain reaction
Total RNA was isolated from adipose tissue using TRIzol reagent (Invitrogen, Carlsbad, CA, USA) according to the manufacturer’s protocol. Complementary DNA was synthesized from the mRNA samples using a reverse transcription PCR kit (iNtRON Biotechnology, Seongnam, Korea). The mRNA levels were evaluated using quantitative real-time reverse transcription-polymerase chain reaction (qRT-PCR) with SYBR Green PCR Master Mix (Promega). qRT-PCR was performed in duplicate according to the manufacturer’s protocol using a QuantStudio 5 real-time PCR system (Thermo Fisher Scientific, Waltham, MA, USA). The relative expression levels of the target mRNAs were quantified using the ∆Ct method, and 18s was used as an internal control. The primer sequences for qRT-PCR were as follows: 5´-CCT CAC CCA GTG GCA AAT CTG-3´ and 5´-TCC ACA TTC CAG GAG CAT ATC T-3´ for APE1/Ref-1; 5´-TGT TCC TCT TAA TCC TGC CCA-3´ and 5´-CCA ACC TGC ACA AGT TCC CTT-3´ for adiponectin; and 5´-CTG CGT GTG TGA AAT GTC ATT G-3´ and 5´-GAG ACC CCT GTG TCG GTT C-3´ for leptin.
Western blot analysis
Tissues were homogenized using radioimmunoprecipitation lysis buffer containing a protease inhibitor, and lysates were denatured in Laemmli buffer. The protein samples (20 μg) were then loaded onto 10% sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) gels and transferred to polyvinylidene fluoride (PVDF) membranes. The membranes were blocked with non-fat dry milk in TBST for 1 hour at room temperature and immunoblotted overnight at 4°C. The primary antibodies used were APE1/Ref-1 (#MR-MA14, MediRedox Inc., Daejeon, Korea), adiponectin (#ab22554, Abcam), and leptin (#ab16227, Abcam) at a 1:1,000 dilution, as well as β-actin (#a5316, Sigma Aldrich) at a 1:5,000 dilution. After washing with the primary antibody, the membranes were incubated with the appropriate horseradish peroxidase-conjugated secondary antibody (Goat anti-Rabbit, #31460; Goat anti-Mouse, #31430, Invitrogen, Rockford, IL, USA) for 1 hour at room temperature and visualized using an enhanced chemiluminescence detection kit (Amersham Pharmacia Biotech, Piscataway, NJ, USA). All experiments were performed in triplicate. Band density was calculated using ImageJ software. In the graph, the average expression of the target proteins was normalized to β-actin expression.
Histological analysis
The isolated epididymal WAT and liver samples were fixed in 10% neutral buffered formalin solution for 24 hours. The samples were embedded in paraffin and sectioned to 5μm thickness. Hematoxylin and eosin (H&E) and immunohistochemistry staining were performed according to routine staining protocols in paraffin-embedded sections. Images were captured using a Leica S9E microscope (Leica, Wetzlar, Germany). Analysis was performed on three animals per group with site selection of three random captured images per animal. Adipocyte diameter was calculated from area measurements obtained with ImageJ software. The diameter (D) was computed using the mathematical formula: D=2×square root (area/π). Liver images were subjected to quantitative analysis of fat accumulation using ImageJ software. A standardized thresholding method was utilized to differentiate areas of fat accumulation from background staining. The extent of fat deposition within hepatocytes was quantified as a percentage of the total hepatocyte area in each image. Three independent images were analyzed per liver sample, and the average percentage of fat accumulation was calculated. The severity of fatty liver was graded according to a standardized scoring system proposed by Kleiner et al. [20]: grade 0: no steatosis (0% hepatocytes affected); grade 1: mild steatosis (1% to 33% hepatocytes affected); grade 2: moderate steatosis (34% to 66% hepatocytes affected); grade 3: severe steatosis (67% to 100% hepatocytes affected); and grade 4: massive steatosis (extensive fat accumulation with displacement of the nucleus).
Quantification of adiponectin and leptin levels in mouse plasma
Adiponectin and leptin levels in mouse plasma were determined. Whole blood samples were collected from the mice through cardiac puncture. The samples were then centrifuged at 2,000 ×g for 15 minutes to separate the plasma. To maintain the stability of adipokines, the separated plasma was stored at –80°C until analysis. For quantification of adipokine levels, plasma samples were sent to the KOMA Analytical Service Center (KOMA Biotech, Seoul, Korea). In this study, an immunology multiplex assay kit was employed, allowing for the simultaneous measurement of multiple adipokines, including adiponectin and leptin, from a single plasma sample. This method ensures the precise and concurrent assessment of these crucial adipokines in mouse plasma.
Assessing adipose tissue accumulation and volume using abdominal micro-magnetic resonance imaging
To confirm the accumulation of adipose tissue, an in vivo micromagnetic resonance imaging (MRI) study was conducted using a 7T MRI scanner (MR Solutions, Guildford, UK). During the MRI experiment, the mice were anesthetized using isoflurane United States Pharmacopeia (USP) (Troikaa Pharmaceuticals Ltd., Gujarat, India) delivered in a mixture of oxygen and air. A heating pad was used to maintain the body temperature of the animals during the examination, and respiratory gating was employed to ensure that the measurements were performed in a stable state. First, a Scout localizer scan was performed to determine the location. Subsequently, MRI scans were acquired using a fast spin-echo T1-weighted imaging sequence. Coronal sections were obtained from ventral to dorsal (TR/TE: 1,600/11 ms; field of view: 60×60 mm3; slice thickness: 1 mm; interslice gap: 1 mm), while transverse sections were acquired from the top of the testis to the bottom of the liver (TR/TE: 1,700/11 ms; field of view: 40×40 mm3; slice thickness: 1 mm; interslice gap: 1 mm).
To quantify the volume of WAT in the abdomen, the volumes of visceral fat and subcutaneous fat were measured separately, and the values were combined. For these results, we employed a thresholding technique implemented in the MicroDicom Viewer software (MicroDicom Ltd., Sofia, Bulgaria). This technique enabled the delineation of adipose tissue borders within each transverse slice. The volume of adipose tissue (mm3) was calculated by multiplying the area (mm2) of each slice by a factor of 2 (1 mm slice thickness+1 mm interslice gap) and adding the volumes of each slice. The selection of the ambiguous slide area of the transverse sections required for analysis was determined by cross-checking the positions of the testes and liver in the coronal sections. Finally, the volumes of all individual slices were summed to obtain the total visceral and subcutaneous WAT volumes [21]. The abdominal cross-sectional areas were averaged after calculating the area (mm2) of each slice.
Statistical analysis
Statistical analysis was performed using t tests to assess significance (P<0.05, P<0.01, P<0.001) between the APE1/Ref-1+/- and APE1/Ref-1+/+ groups in all experiments. Data were presented as mean±standard error of the mean, and experiments were conducted with a sample size of n=5–10 animals. All statistical analyses were performed using GraphPad Prism 8.0 software (GraphPad Software Inc., San Diego, CA, USA).
RESULTS
Characterization of heterozygous APE1/Ref-1 deficiency mice
In this study, we first described the targeting schema for the genetic modification of the APE1/Ref-1 gene, which involved the insertion of loxP sites flanking exons 2 and 3, as shown in Fig. 1A. We successfully distinguished between wild-type (APE1/Ref-1+/+) mice and heterozygous APE1/Ref-1-deficiency (APE1/Ref-1+/-) mice using PCR, confirming the genotypic modifications (Fig. 1B). The body weights of male APE1/Ref-1+/- mice at 20 weeks of age did not show significant changes (n=10) compared with those of wild-type mice (Fig. 1C). Next, we analyzed the mRNA levels of APE1/Ref-1 in the WAT of heterozygous APE1/Ref-1+/- mice. The findings revealed a significant decrease in APE1/Ref-1 mRNA expression in APE1/Ref-1+/- mice compared to wild-type mice (Fig. 1D), underscoring the functional impact of the heterozygous APE1/Ref-1 deficiency mice. Western blotting revealed a notable reduction in APE1/Ref-1 protein expression in the WAT of APE1/Ref-1+/- mice (Fig. 1E, F), and this reduction was also confirmed in the liver tissue (Fig. 1G, H). These results provide evidence for the successful creation of heterozygous APE1/Ref-1-deficiency mice.
Generation of heterozygous apurinic/apyrimidinic endonuclase1/redox factor-1 (APE1/Ref-1) deficiency mice. (A) Targeting schema of the APE1/Ref-1 gene deletion with loxP sites flanking exons 2 and 3. Knockout of the APE1/Ref-1 gene was generated via the repair of double-strand breaks using non-homologous end joining. (B) Polymerase chain reaction genotyping for detection of the APE1/Ref-1 in genomic DNA from heterozygous (APE1/Ref-1+/-) mice, wild-type (APE1/Ref-1+/+) mice. The presence of a 1,460 bp band indicates the wild-type allele, while the presence of a 568 bp band confirms the detection of the deleted allele. (C) The body weight of 20-week-old male heterozygous (APE1/Ref-1+/-) mice and wild-type (APE1/Ref-1+/+) mice (n=10). There is no significant (NS) change in body weight observed between both groups. (D) Reduced APE1/Ref-1 mRNA expression in the white adipose tissue (WAT) of APE1/Ref-1+/- mice (n=5). mRNA levels were analyzed with quantitative reverse transcription-polymerase chain reaction as described in the ‘Methods’ section. (E) Western blotting results of APE1/Ref-1 protein expression in WAT. Data presented are representative of three independent experiments. (F) Quantitative analysis for Western blotting of APE1/Ref-1 protein expression in WAT of APE1/Ref-1+/- and APE1/Ref-1+/+ mice (n=9). (G) Western blotting results of APE1/Ref-1 protein expression in liver. (H) Quantitative analysis to measure APE1/Ref-1 protein expression in liver of APE1/Ref-1+/- and APE1/Ref-1+/+ mice (n=6). The data represent the mean±standard error of the mean. Statistical significance is indicated as NS. EcoRI and BamHI means restriction endonuclease enzyme sites. aP<0.001 vs. APE1/Ref-1+/+ mice.
Increased ROS production in heterozygous APE1/Ref-1-deficient mice
Total ROS production in epididymal WAT and liver tissue was measured using the dichlorofluorescein assay, which detects the presence of a wide range of ROS species. Fig. 2A demonstrates a significant increase in total ROS production in the epididymal WAT of APE1/Ref-1+/- mice compared to wild-type controls. Consistent with this finding, Fig. 2C reveals elevated total ROS production in the liver tissue of APE1/Ref-1+/- mice. The results suggested that a partial deficiency of APE1/Ref-1 is sufficient to disturb the oxidative balance in adipose tissue and liver, leading to increased ROS production. Moreover, superoxide production in the WAT and liver tissue was analyzed using NADPH-driven superoxide production assays, which are commonly used to detect this specific type of ROS [18]. As shown in Fig. 2B, D, superoxide production was significantly elevated in both epididymal WAT and liver tissue in APE1/Ref-1+/- mice. This observation indicates a potential link between APE1/Ref-1 and the regulation of oxidative stress responses in both adipose tissue and liver, as the partial deficiency of APE1/Ref-1 leads to increased superoxide levels.
Abnormal oxidative stress state in the white adipose tissue (WAT) of heterozygous apurinic/apyrimidinic endonuclase1/redox factor- 1 (APE1/Ref-1)-deficient mice. (A) Increased total reactive oxygen species (ROS) production in WAT of APE1/Ref-1+/- mice (n=5–6). (B) Increased superoxide production in APE1/Ref-1+/- compared to APE1/Ref-1+/+ mice in WAT (n=5–6). (C) Increased total ROS production in APE1/Ref-1+/- compared to APE1/Ref-1+/+ mice (n=5). Total ROS was measured with dichlorofluorescein assay. (D) Increased superoxide production in the liver of APE1/Ref-1+/- mice (n=6). Superoxide production was measured with lucigenin-enhanced chemiluminescence. The data represent the mean±standard error of the mean. RFU, relative fluorescence unit; RLU, relative luminescence unit. aP<0.05 vs. APE1/Ref-1+/+ mice.
Adipocyte hypertrophy, inflammation and ectopic fat deposition in heterozygous APE1/Ref-1-deficient mice
Next, we investigated the effects of heterozygous APE1/Ref-1 deficiency on adipocyte morphology in mice. Histological analyses of epididymal WAT from APE1/Ref-1+/- revealed a noticeable increase in the number of hypertrophic adipocytes (Fig. 3A). Moreover, the mean adipocyte diameter in the WAT of APE1/Ref-1+/- mice was significantly larger than that of wild-type mice (71.5 μm vs. 48.8 μm, P<0.05) (Fig. 3B). Immunohistochemical staining for the macrophage marker F4/80 revealed a substantial increase in macrophage infiltration within the adipose tissue of APE1/Ref-1+/- mice (Fig. 3C, D). These findings collectively indicate that APE1/Ref-1 plays a crucial role in maintaining normal adipocyte size and regulating inflammation in adipose tissue.
Adipocyte hypertrophy in white adipose tissue (WAT) and hepatic fat accumulation in heterozygous apurinic/apyrimidinic endonuclase1/ redox factor-1 (APE1/Ref-1)-deficient mice. (A) Low power (4×) and high power (10×) image of hematoxylin and eosin (H&E) staining in WAT from the APE1/Ref-1+/+ mice and APE1/Ref-1+/- mice. Note that APE1/Ref-1+/- mice exhibit larger adipocytes compared to APE1/Ref-1+/+ mice. (B) Mean adipose diameters in WAT of APE1/Ref-1+/- mice and APE1/Ref-1+/+ mice. Mean adipose diameters in WAT of APE1/Ref-1+/- mice are significantly larger than those of APE1/Ref-1+/+ mice, indicating adipose hypertrophy (n=3). (C) Immunohistochemical detection of the macrophage-specific antigen F4/80 in WAT of APE1/Ref-1+/- mice. (D) Quantitative analysis of F4/80 expression in WAT of APE1/Ref-1+/- mice. Macrophage infiltrations are increased in adipose tissue of APE1/Ref-1+/- mice (n=3). (E) Low power (4×) and high power (40×) image H&E staining in liver tissue from APE1/Ref-1+/+ mice and APE1/Ref-1+/- mice. Fat vacuoles are observed in the liver of APE1/Ref-1+/- mice, not in APE1/Ref-1+/+ mice. (F) Quantitative analysis of hepatic steatosis in non-adipose tissue of APE1/Ref- 1+/- mice. Hepatic steatosis was analyzed by histopathological scoring as described in the ‘Methods’ section (n=3). (G) Oil Red O staining for lipid detection in frozen liver tissue sections of APE1/Ref-1+/+ and APE1/Ref-1+/- mice. (H) Quantification of the percentage of increased triglyceride content in liver tissue of APE1/Ref-1+/- mice compared to APE1/Ref-1+/+ mice. The data represent the mean±standard error of the mean (n=3–5). aP<0.05 vs. APE1/Ref-1+/+ mice.
As lipid accumulation in non-adipose tissues, especially in the liver, often indicates ectopic fat deposition, we examined whether APE1/Ref-1+/- mice exhibited any changes in fat accumulation in liver tissue. Histopathological grading of liver sections revealed a significantly higher grade of 0.67 in APE1/Ref1+/- mice compared to wild-type mice, which exhibited a grade close to zero (Fig. 3E, F). Furthermore, Oil Red O staining, used to visualize accumulated TG, showed a greater percentage of positive areas in the livers of APE1/Ref-1+/- mice (Fig. 3G, H). These findings suggest a mild fatty liver phenotype in heterozygous APE1/Ref-1 deficient mice, characterized by increased lipid accumulation and histopathological changes in the liver.
Reduced adiponectin expression in heterozygous APE1/Ref-1-deficient mice
Given the established alterations in adipose tissue in APE1/Ref-1+/- mice, we focused on changes in adipokine expression, particularly adiponectin. We investigated adiponectin expression and plasma levels in APE1/Ref-1+/- mice. qRT-PCR revealed decreased adiponectin mRNA levels in the epididymal WAT of APE1/Ref-1+/- mice compared to wild-type APE1/Ref-1+/+ mice (Fig. 4A). Quantitative analysis of adiponectin protein expression in the epididymal WAT of APE1/Ref-1+/- mice confirmed lower levels in APE1/Ref-1+/+ mice (Fig. 4B, C), which was further supported by immunohistochemical staining for adiponectin (Fig. 4D). Additionally, we analyzed adiponectin levels in the plasma of APE1/Ref-1+/- mice (Fig. 4E). Consistent with the findings in WAT, plasma levels of adiponectin were lower in APE1/Ref-1+/- mice than those in wild-type controls (2,417 pg/mL for APE1/Ref-1+/+ mice vs. 1,717 pg/mL for APE1/Ref-1+/-mice, P<0.01), collectively suggesting substantial alterations in metabolic function related to adiponectin.
Decreased adiponectin in heterozygous apurinic/apyrimidinic endonuclase1/redox factor-1 (APE1/Ref-1)-deficient mice. (A) Decreased mRNA expression of adiponectin in the white adipose tissue (WAT) of APE1/Ref-1+/- mice (n=5). mRNA levels were analyzed with quantitative reverse transcription-polymerase chain reaction as described in the ‘Methods’ section. (B) Representative Western blot analysis of adiponectin protein expression in WAT from three individual mice. (C) Quantitative analysis of adiponectin protein expression in WAT normalized to β-actin showing reduced expression in APE1/Ref-1+/- mice, compared to APE1/Ref-1+/+ mice (n=9). (D) Immunohistochemistry staining of adiponectin showing reduced expression in WAT of APE1/Ref-1+/- mice. (E) Reduced plasma adiponectin levels in APE1/ Ref-1+/- mice compared to APE1/Ref-1+/+ mice (n=6). The data represent the mean±standard error of the mean. aP<0.05; bP<0.01; cP<0.001 vs. APE1/Ref-1+/+ mice.
Elevated leptin expression in heterozygous APE1/Ref-1- deficient mice
We then directed our attention to leptin, an important adipokine in metabolic regulation. qRT-PCR analysis revealed a significant elevation in leptin mRNA levels in the epididymal WAT of APE1/ Ref-1+/- mice (Fig. 5A). Western blotting showed increased leptin protein expression levels in the epididymal WAT of APE1/Ref-1+/- mice, which were significantly higher than those in wild-type mice (Fig. 5B, C). Immunohistochemical staining visually corroborated these findings, illustrating increased leptin expression in the epididymal WAT of APE1/Ref-1+/- mice (Fig. 5D). Moreover, our investigation extended to plasma leptin levels (Fig. 5E), which were significantly elevated in APE1/Ref-1+/- mice compared to wild-type mice (889 pg/mL for APE1/Ref-1+/+ mice vs. 4,047 pg/mL for APE1/Ref-1+/- mice, P<0.001). These findings suggest a broader metabolic impact beyond the local adipose tissue. It is evident that heterozygous APE1/Ref-1 deficiency in mice leads to the upregulation of leptin expression, both at the mRNA and protein levels, in WAT and in the systemic circulation.
Elevated leptin expression in heterozygous apurinic/apyrimidinic endonuclase1/redox factor-1 (APE1/Ref-1)-deficient mice. (A) Increased leptin mRNA expression in the white adipose tissue (WAT) of APE1/Ref-1+/- mice than in APE1/Ref-1+/+ mice (n=5). mRNA levels were analyzed with quantitative reverse transcription-polymerase chain reaction as described in the ‘Methods’ section. (B) Representative Western blotting results of leptin protein expression in WAT from three individual mice. (C) Quantitative analysis of leptin protein expression in WAT normalized to β-actin showing increased expression in APE1/Ref-1+/- mice, compared to APE1/Ref-1+/+ mice (n=6). (D) Immunohistochemistry staining of leptin showing increased expression in the WAT of APE1/Ref-1+/- mice. (E) Increased leptin levels of plasma in APE1/Ref-1+/- mice compared to APE1/Ref-1+/+ mice (n=4). The data represent the mean±standard error of the mean. aP<0.001 vs. APE1/Ref-1+/+ mice.
Increases in abdominal subcutaneous and visceral adiposity in APE1/Ref-1-deficient mice
Finally, we utilized microMRI to investigate abdominal fat changes in APE1/Ref-1+/- mice. Representative MRI images of the coronal sections in Fig. 6A provide insights into the abdominal localization, with the gray box demarcating the analysis area detailed in Fig. 6C-F. The transverse section images in Fig. 6B provide a detailed view of the visceral and subcutaneous fat regions, represented by the white line in Fig. 6A. The visceral fat is identified by its location around the internal organs, while the subcutaneous fat forms a distinct layer just under the skin. MicroMRI revealed a significant increase in both the visceral or subcutaneous abdominal fat volumes in APE1/Ref-1+/- mice compared to those in APE1/Ref-1+/+ mice, as shown in Fig. 6C, D. The combined volume of abdominal white fat, comprising visceral and subcutaneous fat, was substantially higher in APE1/Ref-1+/- mice than in APE1/Ref-1+/+ mice (4,038 mm2 for APE1/Ref-1+/+ mice vs. 5,824 mm2 for APE1/Ref-1+/- mice, P<0.05) in shown in Fig. 6E. However, the average abdominal cross-sectional area in the gray box demarcating the analysis area remained unchanged, as shown in Fig. 6F, suggesting that there was no significant increase in body weight. These results provide valuable insights into the effect of heterozygous APE1/Ref1 deficiency on abdominal fat distribution and highlight its potential involvement in adiposity regulation.
Quantification of increased abdominal fat area in heterozygous apurinic/apyrimidinic endonuclase1/redox factor-1 (APE1/Ref-1) deficiency mice using micro-magnetic resonance imaging (MRI). (A) Representative microMRI images of coronal sections to determine abdominal localization. The gray box indicates the abdominal analysis area yielding the results in panels C-F. The white line indicates the position of the transverse sectional image that is shown in panel B. (B) Representative transverse sectional image of the abdomen (visceral fat [V], subcutaneous fat [S]). (C) Results of increased visceral fat volume in APE1/Ref-1+/- mice compared to APE1/Ref-1+/+ mice (n=5–6). (D) Thicker abdominal subcutaneous fat volume in APE1/Ref-1+/- mice (n=5–6). (E) APE1/Ref-1+/- mice with greater total abdominal fat, which is the sum of visceral and subcutaneous fat (n=5–6). (F) Average abdominal cross-sectional area between the two groups without difference (n=5–6). Data are presented as mean±standard error of the mean. NS, no significant. aP<0.05 vs. APE1/Ref-1+/+.
DISCUSSION
In this study, we demonstrated that heterozygous APE1/Ref-1-deficient mice exhibit increased ROS production, adipocyte hypertrophy in WAT, and ectopic fat deposition. These changes coincide with alterations in adipokine levels characterized by reduced adiponectin and elevated leptin expression. Furthermore, these observations were reinforced by microMRI analyses, which revealed increased abdominal adiposity.
Maintaining balanced level of ROS is crucial for normal WAT function. However, excessive ROS production, as observed in obesity, disrupts this delicate balance, leading to adipose tissue dysfunction [8]. Therefore, strategies aimed at restoring ROS balance in adipose tissue, such as promoting antioxidant defenses or inhibiting ROS production, hold promise for mitigating obesity-related complications and improving WAT function [22,23]. Our findings demonstrate that heterozygous APE1/Ref-1 deficiency in mice leads to a significant increase in ROS generation in adipose tissues. This finding is consistent with the established functions of APE1/Ref-1 in DNA repair and oxidative stress management. Increased ROS levels in these mice likely result from their compromised redox regulation and DNA repair abilities, given the pivotal role of APE1/Ref-1 in these processes [24]. Our study reveals that increased F4/80 staining observed in adipose tissue indicates heightened macrophage infiltration [25], suggesting a chronic inflammatory state in APE1/Ref-1+/-mice. This inflammation may be triggered by elevated ROS levels and contribute to adipocyte dysfunction.
Additionally, the increased ROS levels observed in cells lacking APE1/Ref-1 highlights the importance of this protein in maintaining cellular homeostasis under oxidative stress [26]. Studies on human gastric and colonic cell lines have demonstrated increased ROS production in the absence of APE1/Ref-1 [27]. Furthermore, recent studies have indicated that APE1/Ref-1 inhibition leads to elevated ROS levels, accumulation of lipid peroxidation, and enhanced ferroptosis [28]. Collectively, these findings emphasize the critical role of APE1/Ref-1 in mitigating oxidative stress and its potential implications for cellular functions and pathways, particularly those relevant to adipocyte physiology.
Increased ROS generation significantly impacts the expression of adipokine-related genes, which are crucial regulators of metabolic processes. Oxidative stress has been implicated in the modulation of adipokines, and our study revealed altered expression of key adipokines, such as decreased adiponectin and increased leptin levels, in heterozygous APE1/Ref-1-deficient mice. This suggests that ROS influence the regulatory pathways of these adipokines, potentially leading to metabolic imbalance. High levels of oxidative stress have been associated with the dysregulation of adipokine expression, including adiponectin, leptin, and ghrelin [29]. For instance, research has shown that epicardial adipocytes produce ROS, which correlates with metabolic changes like decreased serum adiponectin levels in severe coronary atherosclerosis [30]. Additionally, elevated leptin levels have been linked to ROS generation, primarily through NADPH oxidase activation [31]. These findings highlight the intricate interplay between oxidative stress and adipokine-related gene expression and underscore the importance of understanding these mechanisms in the context of metabolic disorders and potential therapeutic interventions.
The observed increase in fat deposition and altered fat distribution in heterozygous APE1/Ref-1-deficient mice suggest that oxidative stress may drive these alterations. Elevated oxidative stress—resulting from increased ROS levels due to APE1/Ref-1 deficiency—likely contributes to changes in adipocyte function and fat storage patterns. Previous studies have linked oxidative stress to adipocyte differentiation and lipid accumulation, indicating a potential connection between ROS generation and fat deposition.
ROS play a vital role in adipocyte differentiation, with antioxidants and ROS scavengers shown to suppress adipogenesis [32]. Studies have demonstrated that APE1/Ref-1 overexpression inhibits intracellular ROS production by blocking NADPH oxidase activity and mitochondrial ROS production [26,33,34]. However, inhibiting the redox function of APE1/Ref-1, particularly in cysteine mutants, has been reported to increase intracellular ROS levels [35,36]. Recent studies investigating the role of APE1/Ref-1 in adipocyte differentiation suggests that its redox function exerts anti-adipogenic effects by inhibiting adipocyte differentiation [16]. Silencing APE1/Ref-1 led to increased expression of key adipogenic transcription factors, such as C/EBP-alpha and PPAR-γ, which are crucial for driving adipocyte differentiation. Moreover, E3330, a redox inhibitor of APE1/Ref-1, mirrored the effects of gene silencing, resulting in the upregulation of C/EBP-alpha, PPAR-γ, and aP2, a marker of adipocyte differentiation [16].
Ectopic fat deposition in the liver was observed in heterozygous APE1/Ref-1-deficient mice. Specifically, the histopathological grade for heterozygous APE1/Ref-1 deficiency was determined to be 0.67, indicating mild fatty liver. The observed mild fatty liver phenotype in heterozygous APE1/Ref-1-deficient mice suggest a potential role of APE1/Ref-1 in regulating lipid metabolism or liver function. Oil Red O staining further confirmed increased lipid accumulation in the livers of APE1/Ref-1-deficient mice. Additionally, blood TG levels were elevated in these mice (Supplemental Fig. S1), while cholesterol, high-density lipoprotein, and low-density lipoprotein levels remained unchanged, suggests an impaired ability to metabolize TG in APE1/Ref-1-deficient mice.
The imbalance between ROS production and antioxidant defense mechanisms may facilitate lipid buildup in the liver by promoting processes like increased de novo lipogenesis and compromised lipid oxidation [37,38]. ROS and reactive nitrogen species primarily target hepatocyte proteins, lipids, and DNA, leading to structural and functional liver abnormalities [37]. Nonalcoholic fatty liver disease, a prevalent chronic liver condition globally, is closely linked with oxidative stress [38]. Consequently, heightened oxidative stress in heterozygous APE1/Ref1-deficient mice may contribute to fat deposition; however, the precise mechanism remains unclear.
A notable observation in our study was the lack of significant changes in total body weight in heterozygous APE1/Ref-1-deficient mice, despite increased fat deposition. Compensatory adjustment could result in maintaining a stable overall body weight despite changes in fat distribution [39]. One possible explanation is that the alteration of adipokines might maintain the total body weight, regardless of changes in fat distribution. Despite increased fat mass, elevated leptin levels could indicate leptin resistance, a common phenomenon in obesity where increased leptin fails to reduce appetite or increase energy expenditure [40]. In contrast, a decrease in adiponectin, an adipokine known for its beneficial metabolic effects, may further contribute to metabolic dysregulation [41]. Therefore, these changes in adipokine profiles could balance overall energy homeostasis, leading to stable body weight despite fat redistribution.
There are some limitations in this study. While it focuses on investigating the role of APE1/Ref-1 in adipose tissue dysfunction using a heterozygous APE1/Ref-1 deficiency mouse model, caution is warranted when extrapolating these findings to the human condition due to potential species differences, as the observations are based on APE1/Ref-1+/- mice. Additionally, although the study provides valuable insights into the effects of APE1/Ref-1 deficiency on adipose tissue physiology, the underlying molecular mechanisms remain incompletely understood. Furthermore, while the study suggests potential implications for metabolic disorders, future research should prioritize correlating these findings with clinical data to ascertain their relevance to human health.
In conclusion, our study elucidates the intricate relationship between heterozygous APE1/Ref-1 deficiency, ROS generation, adipose tissue dysfunction, and metabolic alterations. Overall, our study highlights the critical role of APE1/Ref-1 in maintaining cellular homeostasis, particularly in adipocyte physiology. Further research is warranted to elucidate the mechanisms underlying APE1/Ref-1 deficiency-induced adipose tissue dysfunction and its implications for metabolic health.
Supplementary Material
Supplemental Fig. S1.
Blood glucose and lipid profiles in plasma of apurinic/apyrimidinic endonuclase1/redox factor-1 (APE1/Ref-1)+/- and APE1/Ref-1+/+ mice. (A) Blood glucose, (B) triglyceride, (C) total cholesterol, (D) high-density lipoprotein (HDL) cholesterol, or (E) low-density lipoprotein (LDL) cholesterol levels in plasma of APE1/Ref-1+/- and APE1/Ref-1+/+ mice. Note that triglyceride levels were significantly elevated in APE1/Ref-1+/- mice compared to wild-type controls. Fasting blood samples were collected from APE1/Ref-1+/- and APE1/Ref-1+/+ mice as described in ‘Methods.’ Data are presented as mean±standard error of the mean (n=5). aP<0.05 vs. APE1/Ref-1+/+ mice.
Notes
CONFLICTS OF INTEREST
No potential conflict of interest relevant to this article was reported.
AUTHOR CONTRIBUTIONS
Conception or design: E.O.L., H.K.J., Y.R.L., K.H.L., B.H.J. Acquisition, analysis, or interpretation of data: E.O.L., H.J., S.K., H.K.J., Y.R.L., S.Y.A., S.P., B.H.J. Drafting the work or revising: E.O.L., B.H.J. Final approval of the manuscript: B.H.J.
Acknowledgements
This study was funded by the Basic Science Research Program of the National Research Foundation of Korea (NRF) run by the Ministry of Education (grant numbers NRF-2014R1A6A1029 617 to Byeong Hwa Jeon, RS-2023-00250674 to Hee Kyoung Joo, and NRF-2021R1I1A1A01052680 to Eun-Ok Lee).