Role of CCR2 in the Development of Streptozotocin-Treated Diabetic Cardiomyopathy

As a result of changes to diet and lifestyle, the worldwide incidence of diabetes has been increasing rapidly. Diabetes is an independent risk factor for heart failure. Diabetic cardiomyopathy, a common complication of diabetes, demands special clinical attention because of the concealed onset, relatively rapid evolution, and very poor outcome (1). Diabetic cardiomyopathy is characterized by alterations in systolic and diastolic function as well as microangiopathy (2). Increased infiltration of monocytes and macrophages has been observed in the diabetic heart (3). Chemokines and their receptors have been shown to be involved in diabetic cardiomyopathy, and research regarding chemokines and their receptors in diabetic hearts will help to find new targets for treatment of both diabetes and cardiomyopathy (3).

As a chemokine receptor, C-C chemokine receptor type 2 (CCR2) regulates the immune response by inducing macrophage and monocyte recruitment to sites of inflammation (4,5). Indeed, CCR2 is involved in the process of diabetes. Macrophage recruitment through CCR2 into adipose tissue is believed to play a role in the development of insulin resistance and type 2 diabetes mellitus (T2DM) (6). In addition, CCR2⁺ monocyte-derived cardiac macrophages are required for adverse left ventricle remodeling (7). CCR2 is involved in the onset of a variety of diseases associated with diabetes, cardiovascular disease, hypertension, renal disease, and neurodegenerative disorders (5,8–11). These observations lead us to hypothesize that CCR2 induces recruitment of monocytes/macrophages that regulate diabetic cardiomyopathy.

In this study, we investigated the role of CCR2 in the pathogenesis of diabetic cardiomyopathy in mice with streptozotocin (STZ)–induced type 1 diabetes or db/db mice with T2DM. We proved that CCR2 knockout (KO) or treatment with a CCR2 inhibitor protects the function of diabetic hearts.

Male wild-type (WT) C57BL/6J mice (8 weeks old) and db/db mice (14 weeks old) were purchased from Beijing HFK Bioscience Co., Ltd. (Beijing, China). CCR2 KO mice with a C57BL/6 background were housed in pathogen-free animal houses at the Huazhong University of Science and Technology. All animal studies were performed in adherence with the Guide for the Care and Use of Laboratory Animals published by the National Institutes of Health and were approved by the Institutional Animal Care and Use Committee of Huazhong University of Science and Technology.

Mice were randomly divided into four groups, each containing at least 24 mice: controls (WT group), CCR2 KO (KO group), diabetes (WT-STZ group), and diabetes plus CCR2 KO (KO-STZ group). Mice were intraperitoneally injected with STZ (60 mg/kg body weight dissolved in 0.1 mol/L citrate buffer [pH 4.5]) daily for three consecutive days in order to induce type 1 diabetes. Mice in the age- and sex-matched control group were injected with citrate buffer at the same volume as that administered to mice in the WT-STZ group. STZ powder was purchased from Sigma.

The db/db mice were divided into two groups, each containing at least six mice: a control group (injected with vehicle) and a CCR2 inhibitor–treated group. Mice in the latter group were injected intraperitoneally with the CCR2 inhibitor INCB3344 (MedChemExpress, Monmouth Junction, NJ), 30 mg/kg/day for 5 weeks.

We used a Vevo 2100 high-resolution ultrasound machine equipped with a 30-MHz probe (VisualSonics, Toronto, Canada) to assess cardiac function. Mice were anesthetized with 1.5% isoflurane and examined with echocardiography. We measured hemodynamic parameters using a Millar SPR-1000 Mouse Pressure Catheter. We calculated values for cardiovascular factors using LabChart software (AD Instruments, Bella Vista, Australia). All of the measurements were obtained from at least three beats and averaged, as described previously (12,13).

Heart tissues were fixed and embedded in paraffin. We then stained the tissues with hematoxylin and eosin for preliminary assessment and with Masson trichrome to identify collagen fiber. We calculated the area of interstitial fibrosis as a percentage of the total area, and the area of perivascular fibrosis was the ratio of the fibrotic area around the vessel wall to the total vascular area.

Nine days after completing STZ treatment, we performed glucose tolerance tests using glucose infusion (1 g glucose/kg body weight) after mice were deprived of food overnight. Blood samples were taken from the mice’s tails at designated time points. We measured blood glucose using a One-Touch Ultra Glucometer (Johnson & Johnson). We used the LabAssay Triglyceride test kit (catalog no. 290-63701) for triglyceride assays and the LabAssay Cholesterol test kit (catalog no. 294-65801) for cholesterol assays (Wako Pure Chemical Co.), according to the manufacturers’ instructions, to determine the lipids extracted from serum.

Hearts were excised and then minced with fine scissors before digestion in 450 units/mL collagenase I, 125 units/mL collagenase XI, 60 units/mL DNase I, and 60 units/mL hyaluronidase (Sigma-Aldrich) for 1 hour at 37°C under agitation. Tissues were triturated and cells filtered through a 40-μm nylon mesh (BD Falcon), washed, and centrifuged (300g and 4°C for 8 min). Cells were stained at 4°C in FACS buffer. Cells were resuspended in RPMI 1640. Allophycocyanin anti-mouse CD206, FITC antimouse/human CD11b, phycoerythrin antimouse F4/80, and appropriate isotype controls were purchased from BioLegend Inc. Antimouse CD16/32 (BioLegend Inc.) was used for blocking the nonspecific binding of Ig to the Fc receptors.

Cells passed through the cytometer were gated by granularity and size, and singlets were selected in order to exclude cell clumps. CD11b⁺F4/80⁺ macrophages were selected for further analysis, and CD206⁺CD11b⁺ macrophages were selected as M2-like macrophages.

According to the instructions for the In Situ Cell Death Detection Kit (Roche, Basel, Switzerland), heart sections were treated with terminal deoxynucleotidyl TUNEL staining. All the nuclei were counterstained with DAPI. Digital images were observed under a microscope with a ×40 objective lens (Olympus, Tokyo, Japan). From each sample we obtained three random fields, and we calculated the ratio of TUNEL-positive nuclei and the total number of nuclei to define the percentage of apoptotic cells.

Total RNA was extracted from the heart tissues by using Invitrogen TRIzol reagent (Thermo Fisher Scientific, Waltham, MA). The RNA was reverse-transcribed into cDNA by using a PrimeScript RT Reagent Kit (Takara Bio Inc., Shiga, Japan). PCR confirmed CCR2 KO with the use of specific primers: CCR2_CoF: TGCTCACCAGGAAATGCCA-AG; CCR2_Mu_R: CTCGGGCGGAAAGAACCAGC; CCR2_Wt_R: TGAGCAGGAAGAGCAGGTCAGAG.

We quantified the mRNA expression of target genes after PCRs in an ABI PRISM 7900 sequence detection system and analyzed the expression by normalizing to β-actin levels and using the comparative Ct method (2^−ΔΔCt). Each reaction was performed in duplicate. Primers were as follows: β-actin forward: GCTG-TATTCCCCTCCATCG; β-actin reverse: CCAGTTGGTAA-CAATGCCATGT; CCR2 forward: ATCCACGGCATACTAT-CAACATC; CCR2 reverse: AAGGCTCACCATCATCGTAG; CD68 forward: AAAGGCCGTTACTCTCCTGC; CD68 reverse: ACTCGGGCTCTGATGTAGGT; F4/80 forward: CGT-GAGTTGCAGTTCTGCTC; F4/80 reverse: CTTGGCCCAGC-ACCGTATAA; Cd11c forward: CTGGATAGCCTTTCTTCT-GCTG; Cd11c reverse: GCACACTGTGTCCGAACTCA; tumor necrosis factor-α (TNF-α) forward: TGGAACTGGCAG-AAGAGGC; TNF-α reverse: GACAGAAGAGCGTGGTGGC; NOS2 forward: CTATGGCCGCTTTGATGTGC; NOS2 reverse: TTGGGATGCTCCATGGTCAC; interleukin (IL)-6 forward: TAGTCCTTCCTACCCCAATTTCC; IL-6 reverse: TTGGTCCTTAGCCACTCCTTC; IL-1β forward: GCAACT-GTTCCTGAACTCAACT; IL-1β reverse: ATCTTTTGGGGT-CCGTCAACT.

Total protein abstracted from the murine heart tissue was extracted in radioimmunoprecipitation buffer containing a protease inhibitor. We used a BCA Protein Assay Kit (Thermo Fisher Scientific) to determine the protein concentrations, per the manufacturer’s instructions. The same amount of denatured protein sample (60 μg) was separated by SDS-PAGE and transferred to polyvinylidene difluoride membranes (Merck-Millipore, Darmstadt, Germany). We used the following primary antibodies: cleaved caspase-3 antibody (9654S), Bcl-2 antibody (3498S), and Bax antibody (2772S) (all from Cell Signaling Technology, Danvers, MA); α–smooth muscle actin (SMA) (55135-1-AP), fibronectin (15613-1-AP), and collagen I (14695-1-AP) (all from Proteintech); and CCR2 (DF2711; ABclonal Technology). After being washed three times with Tris-buffered saline containing Tween, the membranes were incubated with horseradish peroxidase–conjugated secondary antibodies (Tianjin Sungene Biotech Co., Ltd., Tianjin, China). Bands were displayed with enhanced chemoluminescence reagents (170-5061; Bio-Rad, Hercules, CA). The expression levels of target proteins were normalized to levels of β-tubulin (Tianjin Sungene Biotech).

The mice’s hearts were kept overnight in 4% paraformaldehyde and then embedded in paraffin. To inhibit endogenous peroxidase activity, we cut the tissue into 5-μm-thick sections and treated the sections with hydrogen peroxide blocks for 15–20 min. The sections were then incubated with primary anti-F4/80 antibody (Ab6640; Abcam) at 4°C for 12 h. The sections were washed three times in PBS, incubated with fluorescent antibody at room temperature for 30 min, and then stained with DAPI to display the nuclei. We viewed at least three random fields of each sample under a fluorescent microscope and calculated the average number of positive cells.

We measured NADPH oxidase activity using an NADPH Oxidase Activity Test Kit (Genmed Scientifics, Inc., Shanghai, China), according to the manufacturer’s instructions, as previously described (14). We determined the protein concentration using a bicinchoninic acid assay. Reactive solution was mixed with buffer solution, 100 μL of the sample was added, and the culture medium was placed in an incubator for 3 min. The matrix solution was added and mixed (within 3 s), and the absorbance was then determined at 550 nm on a microplate reader (Tecan). We measured 4-hydroxynonenal (4-HNE) and 3-nitrotyrosine (3-NT) in heart tissue extracts using Cell Biolabs kits (San Diego, CA), according to the manufacturer’s protocols; these were used as markers of oxidative or nitrative stress, respectively.

Data are presented as the mean ± SEM from at least three independent experiments. We performed an unpaired Student t test to analyze comparisons between two groups, and we used one-way or two-way ANOVA to analyze comparisons among multiple groups. SPSS 13.0 software (SPSS Inc., Chicago, IL) was used for statistical analysis, and P < 0.05 represented statistical significance.

Previous studies have shown increased expression of CCR2 in patients with diabetes (15). To determine whether CCR2 is a key molecule in the development of diabetic cardiomyopathy, we examined the expression of CCR2 in the hearts of diabetic mice. We created STZ-induced diabetic C57BL/6J mice (the WT-STZ group) and compared them with the control (WT) mice (treated with saline). Mice were sacrificed 2, 4, or 6 months after being injected with STZ (60 mg/kg/day) for three consecutive days. Western blotting showed that cardiac expression of CCR2 was significantly upregulated 24 weeks after STZ treatment in WT-STZ mice (Supplementary Fig. 1A and B). PCR assays also confirmed that the mRNA level of CCR2 increases over time in the hearts of STZ-treated diabetic mice (Supplementary Fig. 1C).

To investigate the relationship between upregulation of CCR2 and STZ-induced diabetic cardiomyopathy, we created STZ-induced diabetic mice using CCR2-deficient mice (KO group) and compared them to WT-STZ mice. Animals were divided into four groups (WT, KO, WT-STZ, and KO-STZ). We assessed glucose tolerance power using an intraperitoneal glucose tolerance test (IPGTT). We measured fasting plasma glucose, serum cholesterol, triglycerides, body weight, and glucose tolerance power in each animal after saline (WT and KO groups) or STZ (WT-STZ and KO-STZ groups) treatment. The results showed that STZ injection produced marked diabetes, with increases in fasting plasma glucose, serum cholesterol, and triglycerides but decreases in body weight and glucose tolerance power (Fig. 1A–E).

We found no differences in fasting plasma glucose, serum cholesterol, triglycerides, body weight, and glucose tolerance power between WT and KO mice that were not treated with STZ, and plasma glucose levels remained below 10 mmol/L in both groups (Fig. 1A–E). We also found no significant difference in body weight or fasting glucose level between the diabetic WT and KO mice (Fig. 1A and D). Furthermore, CCR2 KO significantly decreased serum triglyceride concentrations in the diabetic mice, but it had no effect on the total serum cholesterol level (Fig. 1B and C). In addition, diabetic KO mice had higher glucose tolerance power than WT mice (Fig. 1E). Compared to WT mice, KO mice displayed obvious alterations in survival rates when injected with STZ (Fig. 1F).

To investigate whether CCR2 influenced the function of diabetic hearts, we evaluated cardiac function using transthoracic echocardiography and hemodynamic analysis. Diabetes-induced cardiac dysfunction was reflected by decreased left ventricle ejection fraction (LVEF), left ventricle fractional shortening (LVFS), dP/dt maximum rate, increased diastolic left ventricular internal diameter (LVID,d), and increased systolic left ventricular internal diameter (LVID,s) in the WT-STZ mice (Fig. 2). These changes were reversed in the KO-STZ mice: LVEF and LVFS were elevated in the KO-STZ mice, whereas LVID,d and LVID,s were decreased (Fig. 2).

Histological examination revealed that 24 weeks after the induction of diabetes, the fibrotic area progressively extended to the perivascular and interstitial areas in hearts of WT-STZ mice. This increase was significantly inhibited in hearts of KO-STZ mice (Fig. 3A). Quantitative analysis revealed that KO-STZ mice showed significantly less cardiac fibrosis than WT-STZ mice (Fig. 3B and C). We next examined the expression of fibrosis-related genes using Western blotting and RT-PCR. As shown in Fig. 4, the expression of fibrotic markers such as α-SMA, fibronectin, and collagen I was dramatically increased in diabetic hearts, but CCR2 KO retarded this expression.

These results suggest that CCR2 affects cardiac dysfunction in STZ-induced diabetic cardiomyopathy, and the increased expression of CCR2 induced by hyperglycemia might cause cardiac dysfunction and fibrosis.

Oxidative stress is closely associated with the pathogenesis of diabetes (16). We further examined the relationship between CCR2 and cardiac oxidative stress in the diabetic heart. The presence of carbonyl groups in proteins has been used as a marker of reactive oxygen species (ROS)–mediated protein oxidation (17). We thus examined NADPH oxidase activity to represent ROS levels. As shown in Fig. 5A, cardiac NADPH oxidase activity was upregulated in the myocardium in WT-STZ mice, but disruption of CCR2 in left ventricular tissues significantly inhibited this enhancement. Using ELISA, we then examined 4-HNE and 3-NT 24 weeks after inducting diabetes with STZ (Fig. 5B and C). Similarly, expression of both 4-HNE and 3-NT was also upregulated in hearts of WT-STZ mice. This increase was inhibited in hearts of KO-STZ mice (Fig. 5B and C). These results indicate an antioxidative effect of CCR2 KO in hearts of STZ-treated mice.

Hyperglycemia-induced oxidative stress increases the production of oxidative damage to DNA and affects the expression of the DNA-repairing enzyme, which leads to cell death from apoptosis (18). To further clarify the mechanism of the protective effects of CCR2 KO in STZ mice, we assessed cardiac apoptosis using the TUNEL assay. As shown in Fig. 6A and B, hearts from KO-STZ mice contained significantly fewer TUNEL-positive cells than hearts from WT-STZ mice. We also analyzed the levels of apoptotic proteins, including cleaved caspase-3, Bax, and Bcl-2. Results showed that CCR2 KO inhibited the expression of cleaved caspase-3 and the ratio of Bax to Bcl-2 in diabetic hearts (Fig. 6C–E). These data suggest that CCR2 KO protects STZ-induced cardiac dysfunction by inhibiting cell apoptosis.

Long-term exposure to oxidative stress in diabetes induces chronic inflammation (16). As immunofluorescence staining demonstrated, the number of inflammatory cells (macrophages) infiltrating the myocardium in KO-STZ mice was significantly lower than that found in WT-STZ mice (Fig. 7A and B). The mRNA expression of macrophage-specific markers (CD68 and F4/80) was lower in KO-STZ mice than in WT-STZ mice (Fig. 7C).

To further test the role of CCR2 in polarizing tissue macrophages, we used gene expression profiling to identify novel M1 and M2 macrophage gene signatures. Real-time PCR assay showed that CCR2 KO resulted in a decrease in the cardiac expression of M1 markers but also in the upregulation of the expression of genes associated with M2 polarization (Fig. 7D and E). Furthermore, using flow cytometric analysis to isolate macrophages from the heart, we found that the percentage of M2 macrophages was increased in STZ-treated CCR2 KO mice (Fig. 7F). These data demonstrate that CCR2 is required for macrophages to accumulate in the heart and for polarization to the M1 state in WT-STZ mice.

We used db/db mice treated with the CCR2 inhibitor INCB3344 to detect the role of CCR2 in the development of diabetic cardiomyopathy in T2DM. The results showed that body weight, fasting plasma glucose, IPGTT results, and serum triglyceride levels were higher in db/db mice than in WT mice (Fig. 8A–D). Moreover, treatment with the CCR2 inhibitor significantly reduced fasting plasma glucose, serum triglycerides, and IPGTT levels in the db/db mice, but it did not affect their body weight (Fig. 8A–D).

In addition, cardiac dysfunction was also reflected by lower LVEF, LVFS, and cardiac output in the db/db mice than in the WT mice; CCR2 inhibitor treatment, however, reversed these effects (Fig. 8E–G). These results indicate that CCR2 inhibition also protects cardiac function in db/db mice with T2DM.

In this study, we demonstrated that CCR2 is an important molecule not only involved in the onset and development of diabetic cardiomyopathy. CCR2 inhibition could reverse cardiac dysfunction and fibrosis and inhibit oxidative stress, myocardial apoptosis, and inflammation in the heart. Furthermore, CCR2 KO promoted the polarization of macrophages to the M2 state.

STZ treatment inhibits the enzymatic activity of cardiac contractile proteins such as myosin and actomyosin, and it gradually reduces cardiac contractile function (19). These findings provide a basis for the well-known diabetic cardiomyopathy model. In the hearts of WT mice exposed to increasing plasma glucose levels, cardiac CCR2 expression increased over time. Cardiac function deteriorated 24 weeks after STZ administration, whereas this deterioration was attenuated in the CCR2 KO mice. These results suggest that CCR2 is involved in the onset and development of STZ-induced diabetic cardiomyopathy.

Long-term hyperglycemia can promote the inflammatory response and the amount of ROS in cardiomyocytes and vascular endothelial cells. Moreover, the inflammatory response further promotes the activation of ROS through inflammatory cells such as macrophages. Oxidative stress is well known to be involved in the development of diabetic cardiomyopathy, and increased ROS production could induce various cardiovascular complications including cardiac dysfunction (16,20,21). It is important to elucidate the cellular mechanisms of ROS increases in the diabetic heart. Our results revealed that cardiac production of 4-HNE and 3-NT was increased in diabetic hearts. We also showed that the cardiac expression of NADPH oxidase is elevated in diabetic hearts. Moreover, CCR2 KO decreased the production of 4-HNE and 3-NT, and the expression of NADPH, in diabetic hearts. These results suggest that oxidative stress is increased in the diabetic heart, and CCR2 is an important molecule involved in the production of ROS and oxidative stress in the diabetic heart.

The role of monocytes/macrophages has been described as a key factor in the pathogenesis of diabetic cardiomyopathy (22). Macrophages are pleiotropic cells that can be polarized to perform a variety of critical functions related to inflammation, apoptosis, and fibrosis (23). M1 macrophages secrete inflammatory cytokines, which reduce cardiac and systemic insulin signaling and facilitate the development of diabetes (24), whereas “M2-like” macrophages may be involved in resolving inflammation and initiating repair (23) In this study, we demonstrated that STZ-induced F4/80⁺ cell infiltrates are reduced in CCR2 KO mice. Because CCR2 KO reduces M1 macrophage infiltration, we propose that proinflammatory mediators are downstream of CCR2 inhibitors in that reduced M1 macrophage infiltration leads to reduced inflammation and apoptosis.

In conclusion, we proved that inhibition of the chemokine CCR2 could inhibit oxidative stress and M1 macrophage infiltration in diabetic hearts. Our results demonstrate the mechanisms through which the expression of CCR2 in the heart, associated with persistent hyperglycemia, leads to the development of diabetic cardiomyopathy. Moreover, we suggest that CCR2 could be a useful treatment target not only for diabetic cardiomyopathy but also for diabetic complications.

Acknowledgments. The authors thank Dr. Congyi Wang, Tongji Medical College, Huazhong University of Science and Technology, for providing CCR2 knockout mice.

Funding. This work was supported by a national key research and development program (2016YFA0101100), National Natural Science Foundation of China (81400303, 81570405, 81570257, 81600616, 81700517), Major Key Technology Research Project of the Science and Technology Department of Hubei Province (2016ACA151), and key projects of Huazhong University of Science and Technology (2016JCTD107).

Duality of Interest. No potential conflicts of interest relevant to this article were reported.

Author Contributions. X.T., L.H., and Z.S. researched data and contributed to discussion. L.C., D.S., and X.M. researched data. X.L. and M.D. contributed to discussion. S.D., K.H., and F.Z. wrote, reviewed, and edited the manuscript. K.H. and F.Z. are the guarantors of this work and, as such, had full access to all the data in the study and take responsibility for the integrity of the data and the accuracy of the data analysis.