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Introduction

Mitohormesis refers to continuous mitochondrial remodeling in living cells to adapt to energy demands and physiological functions of different parts of the cell, including changes in dynamics of energy metabolism, biosynthesis, fusion and division, as well as autophagy (1). Abnormal mitohormesis is a key pathogenic mechanism in a variety of cardiac diseases, such as cardiac hypertrophy and heart failure, that is reflected in impaired myocardial energy metabolism and oxidative stress (2). The causes of abnormal homeostasis are associated with changes in mitohormesis-related factors, such as cytochrome c oxidase subunit 4 (COX 4), mitochondrial fission protein 1, optic atrophy 1 (OPA1), coiled-coil, Beclin1, PTEN induced putative kinase 1 (PINK1), uncoupling protein 2 (UCP 2) and dynamin-related protein 1 (DRP 1) (3,4), and, therefore, dynamic balance of mitohormesis is maintained by altering the response state of each factor. In a rat model of heart failure, 8 weeks of aerobic endurance exercise training effectively inhibited myocardial oxidative stress, increased mitochondrial biosynthesis and energy metabolism and converged mitochondrial spatial conformation from fission to fusion, which resulted in an improvement of left ventricular pumping function following acute infarction (5). It is widely accepted that skeletal muscle contraction responds directly to exercise but the pathway through which exercise regulates the myocardium is unknown.

Exercise induces skeletal muscle to secrete hormone-like substances (known as myokines) into the circulation (6), enabling modulation of distal organs. The production of the muscle factor irisin is associated with the regulation of peroxisome proliferator-activated receptor γ coactivator-1 α (PGC-1α) and the motor contraction of skeletal muscle; PGC-1α stimulates expression of fibronectin type III domain-containing protein 5 (FNDC5) gene in muscle tissue and produces a membrane protein that, after its proteolytic hydrolysis, releases into the blood a polypeptide fragment irisin (7). Currently, studies on irisin are multifaceted, particularly those associated with energy metabolism, such as cellular homeostasis and mitochondrial quality control (8).

Exercise stress is the primary mode of irisin production. Boström et al (7) reported a significant elevation in mRNA levels of PGC-1α and FNDC5, as well as an increase in plasma irisin levels, after applying an exercise stimulus to mice; however, a significant decrease in FNDC5 and irisin was observed in mice with a specific knockout of muscle PGC-1α. Although exercise has been noted to increase plasma irisin concentrations in meta-analyses (9,10), type and intensity of exercise exert different effects. In human obese female subjects, serum irisin levels were elevated following both moderate intensity intermittent and continuous exercise and post-exercise serum irisin levels were higher than intermittent exercise in continuous exercise subjects (11). By contrast, there is no significant difference in circulating irisin levels in older patients after a single session of resistance training or 21 weeks of high-intensity endurance training (12), as well as in rats subjected to moderate intensity continuous training and long-term high-intensity training (13). Among the problems that cannot be ignored are the half-life of irisin in vivo, which is <1 h (14), and experimental and subject differences.

Previous studies have confirmed that the heart secretes the most irisin (15). Many heart-associated diseases are associated with irisin, such as coronary heart disease, hypertension, cardiomyopathy and stroke (8). FNDC5/irisin improves cardiac function by activating AKT/mTOR (16) and AMP kinase α) (17). Exercise-induced elevation of circulating irisin levels improves cardiac function (18), particularly in a model of cardiac hypertrophy associated with oxidative stress and irisin modifies reactive oxygen species (ROS) levels in cardiomyocytes via protective autophagy (19) and mitochondrial biosynthesis (20); these effects are associated with the integrity of mitochondrial function. However, to the best of our knowledge, no studies have addressed the regulatory role of irisin on cardiac mitochondria undergoing oxidative stress from the perspective of mitohormesis. Here, an oxidative stress model was established using H9c2 cells to assess the potential mechanisms of exogenous irisin regulation of mitohormesis.

Materials and methods

Materials

DMEM was from HyClone. Fetal bovine serum (FBS) was purchased from Hangzhou Sijiqing Bioengineering Material Co. MTT (cat. no. G020-1-1) and malondialdehyde (MDA; cat. no. A003-2) were purchased from Nanjing Jiancheng Technology Co. RevertAid First Strand cDNA Synthesis kit (cat. no. K1622), Maxima SYBR Green/ROX qPCR Master Mix (cat. no. K0221), Lipofectamine® RNAiMAX and H2DCFDA (cat. no. D399) reagent were obtained from Thermo Fisher Scientific, Inc. Small interfering (si)RNA Interference kit (cat. no. A10001; Shanghai Gemma Genes). The primary antibodies anti-mitofusin 2 (Mfn2), anti-dynamin-1-like protein (DRP1), anti-optic atrophy 1 (OPA1), anti-cytochrome c oxidase IV (COX4), anti-uncoupling protein 2 (UCP2), β-tubulin, anti-goat and anti-rabbit IgG were purchased from Abcam. Protein extraction reagent (1 ml extraction reagent with 5 µl each protease inhibitor mixture, PMSF and protease phosphatase inhibitor mixture) was obtained from Biyun Tian Biotechnology Co. All other reagents were of analytical grade.

Cell culture

Rat cardiomyocytes (H9c2 cells; cat. no. 61761; ScienCell Research Laboratories, Inc.) were routinely cultured in DMEM containing 25 mm glucose, 10% FBS, 1% sodium pyruvate, 100 U/ml penicillin and 100 mg/ml streptomycin. Cells were maintained in a 5% CO2 atmosphere with 90% relative humidity at 37°C.

Cell sensitivity assay

Cell proliferation and viability were determined using an MTT assay. H9c2 cells (5,000–6,000 cells/well) were inoculated into 96-well plates for 12 h, then treated with different concentrations of H2O2 (0, 10, 15, 20, 40, 80 and 100 µM) for another 24 h at 37°C. Next, the culture media were discarded and 300 µl MTT solution (1 mg/ml) in PBS was added to each well for 4 h. A total of 100 µl DMSO was added to each well to dissolve purple-blue MTT formazan precipitate. After the sample was subjected to low-frequency vibration for 10 min in a shaker, absorbance was determined using an automatic microplate reader (cat. no. 1681130A; iMarK Microplate Reader; Bio-Rad Laboratories, Inc.) at an optical density of 570 nm.

Assessment of oxidative stress

H9c2 cells were incubated with H2O2 (0, 10, 15, 20 and 40 µM) at 37°C for 24 h. Lipid peroxidation was assessed using MDA assay kit. Samples underwent vortex mixing, boiling water bath >95°C for 40 min, cooling under running water, centrifugation at 8,000 g for 10 min at room temperature and supernatant collection for measurement at 532 nm using U800 (Beckman co) instrument. Finally, the MDA content was calculated. H9c2 cardiomyocytes were treated with 40 µM H2O2 for 24 h with irisin (0, 50, 75 and 100 µM) added 12 h before the end of the treatment; supernatant was then collected for MDA assay and subsequent experiments.

Mitochondrial ROS assay

H9c2 cells at 80% density were cultured in 6-well plates and incubated with DCFH-DA (10 µM) containing a ROS fluorescent probe for 20 min at 37°C and harvested by trypsinization. ROS content was determined by flow cytometry (FCM, BD Diagnostics).

Mitochondrial membrane potential (MMP) assay

H9c2 cells at 80% density were harvested by trypsinization and resuspended in 500 µl JC-1 staining buffer, incubated at 37°C for 20 min in the dark, then centrifuged at 4°C for 10 min at 600 g to remove the supernatant. H9c2 cells were washed twice with JC-1 staining buffer and resuspended in JC-1 staining buffer. MMP depolarization was detected using flow cytometry (BD Diagnostics).

Reverse transcription-quantitative (RT-q) PCR

The primers were synthesized by Invitrogen (Thermo Fisher Scientific, Inc.; Table I). Total RNA was isolated from cells using TRIzol and cDNA synthesis was conducted using a Quantscript RT kit (Tiangen Biotech Co., Ltd.). qPCR was performed using a commercial kit (SYBR premix ExTaq II, Takara Bio, Inc.). Amplification was performed in 96-well plates using a 7900 HT Fast Real Time-PCR System (Applied Biosystems; Thermo Fisher Scientific, Inc.). The thermocycling conditions were 95°C for 10 min, followed by 40 cycles at 95°C for 15 sec, 60°C for 30 sec and 72°C for 30 sec. Relative mRNA expression levels were determined using the comparative critical threshold method, as described in Applied Biosystems User Bulletin no. 2 (P/N 4303859). β-actin gene was used as the housekeeping gene.

Table I. PCR primers.

Table I.

PCR primers.

Target	Forward primer, 5′→3′	Reverse primer, 5′→3′
COX4	GCTGAGTCCTAAGTGATGCG	CTTTCCTTTGGATGTTTCCC
OPA1	CTGTAACCTTTGTCGGAGAGG	CAGCGTGTTGGCAGTGATAG
DRP1	CCTGACGCTTGTGGATTTAC	GGGATTACTGATGAACCGAAG
PGC-1α	AAGATGCCTCCTGTGACT	GATGACCGAAGTGCTTGT
siRNA 1	CCGGAGCAAUCUGAGUUAUTT	AUAACUCAGAUUGCUCCGGTT
siRNA 2	GCCAAACCAACAACUUUAUTT	AUAAAGUUGUUGGUUUGGCTT
siRNA 3	GCUCUUGAGAAUGGAUAUATT	UAUAUCCAUUCUCAAGAGCTT
Negative control	UUCUCCGAACGUGUCACGUTT	ACGUGACACGUUCGGAGAATT
β-tubulin	TGGTGGGTATGGGTCAGAAGGACTC	CATGGCTGGGGTGTTGAAGGTCTCA

[i] COX4, cytochrome c oxidase subunit 4; OPA1, optic atrophy 1; DRP1, dynamin-related protein 1; PGC-1α, peroxisome proliferator-activated receptor-γ coactivator-1 alpha.

PGC-1α siRNA transfection

The siRNA transfection was performed when the density of H9c2 cells in the culture dish reached 60–80%. siRNA and RNAiMAX transfection reagent were diluted (on ice) with Opti-MEM, then mixed at a volume ratio of 1:1 and left for 5 min at room temperature. The transfection mixture was added for 24 h, during which transfection efficiency was initially determined by fluorescence intensity. The final transfection efficiency was verified by RT-qPCR to determine the relative expression of PGC-1α mRNA. After 48 h transfection, PGC-1α protein was detected by western blotting.

Western blotting

Cellular proteins were extracted by mammalian protein extraction reagent and quantified by BCA method. Equal amounts of protein (18 µg/lane) were separated by 12% SDS-PAGE and transferred to PVDF membranes. After blocking with PBST containing 5% (w/v) non-fat dry milk powder, the membranes were incubated overnight at 4°C with anti-DPR1 (1:2,000, rabbit polyclonal), anti-Mfn2 (1:3,000, mouse monoclonal), anti-OPA1 (1:3,000, mouse monoclonal), anti-COX4 (1:3,000, mouse monoclonal), anti-UCP2 (1:1,500, rabbit polyclonal) and anti-β-tubulin (1:5,000, mouse monoclonal). The membranes were incubated with secondary HRP-conjugated goat anti-rabbit/mouse IgG (1:5,000) and then protein bands were visualized with enhanced ECL detection system (Bio-Rad Laboratories, Inc.). The protein expression was quantified by densitometry, using software AlphaEaseF, relative to β-tubulin.

Statistical analysis

All experiments were replicated three times and the experimental data were processed using GraphPad Prism 5 (Dotmatics) and expressed as mean ± standard deviation. The significance of differences between groups was analyzed by one-way ANOVA with Bonferroni post hoc test. P<0.05 was considered to indicate a statistically significant difference.

Results

Irisin improves H2O2-induced oxidation modeling

Highest intracellular MDA content was observed when experiments were performed using 40 µM H2O2 in which cardiomyocyte proliferation was not affected (Fig. 1A-C). However, when cells were treated with 75 nM irisin, the intracellular MDA content of cardiomyocytes decreased. Therefore, 40 µM H2O2 and 75 nM irisin were selected for subsequent experiments.

Figure 1. Irisin improves H2O2-induced oxidation modeling. (A) Cardiomyocyte viability was measured using MTT assay. MDA measurement in H9c2 cells treated with (B) H2O2 and (C) irisin and 40 µM H2O2. *P<0.05, **P<0.01 vs. 0. MDA, malondialdehyde; OD, optical density.

Irisin partially protects H9c2 cell mitochondria against oxidative stress

Fluorescence intensity was stronger in the H2O2 group but slightly reduced by irisin (Fig. 2A-C). Median value of ROS in the H2O2 group was more to the right, whereas the median value in the H2O2/irisin group was in between that of the control group and H2O2 group (Fig. 2E-G). MMP levels significantly decreased (Fig. 2D) and the ROS content was significantly increased (Fig. 2H) following H2O2 treatment compared with the control group, while the MMP levels significantly increased (Fig. 2D) and the ROS content significantly decreased (Fig. 2H) after the addition of irisin. Therefore, irisin was found to resist oxidative stress and maintain normal energy metabolism in H9c2 cells by partially restoring mitochondrial MMP and decreasing intracellular ROS content.

Figure 2. Detection of MMP and ROS in H9c2 cells by flow cytometry. MMP (A) control, (B) H2O2 and (C) H2O2/irisin group. (D) MMP depolarization. ROS (E) control, (F) H2O2 and (G) H2O2/irisin. (H) Median ROS content in H9c2 cardiomyocytes. **P<0.01 vs. control; #P<0.05, ##P<0.01 vs. H2O2. MMP, mitochondrial membrane potential; ROS, reactive oxygen species.

Irisin partially activates mitochondrial homeostatic indicator expression

Certain factors associated with mitohormesis were expressed at transcriptional and protein levels following the treatment of H9c2 cells with H2O2 and irisin. Following H2O2 treatment, mitochondria tended to split, and expression of DRP1 gene was significantly elevated at the transcriptional and protein levels (Figs. 3B and D), whereas the expression of the endomembrane fusion gene OPA1 was only significantly reduced at the transcriptional level (Fig. 3A) and the expression of the outer-mode fusion gene MFN2 was significantly reduced at the protein level (Fig. 3D). When irisin was added, the mitochondria exhibited a fusion trend and the DRP1 expression was significantly decreased at the transcriptional and protein levels (Fig. 3B and D). By contrast, the expression of OPA1 was significantly elevated, of which only the short type was expressed at the protein level (Fig. 3A and D) and the expression of MFN2 was significantly elevated at the protein level (Fig. 3D). This suggested that irisin partially altered the fusion and fission state of mitochondria and restored the mitochondrial homeostatic imbalance in H9c2 cells caused by excessive ROS.

Figure 3. H2O2 induced oxidative stress in H9c2 cardiomyocytes, mitohormesis-related factors were expressed at the transcriptional and protein levels after the addition of irisin. The housekeeping gene, β-tubulin, was used to normalize the expression level, and compared to the control or H2O2 group. (A) OPA1 mRNA-related expression. (B) DRP1 mRNA-related expression. (C) Western blot analysis. (D) The results of the gray-scale analysis are also shown which are expressed as a percentage (%) of β-tubulin. *P<0.05 vs. control; #P<0.05 vs. H2O2. (S-)OPA1, (short-form) optic atrophy 1; DRP1, dynamin-related protein 1; MFN2, mitofusin 2.

PGC-1α siRNA intervention

Fig. 4 shows the expression of PGC-1α mRNA following siRNA transfection. siRNA sequence of fragment 2 exhibited interference success rate >70% and was selected for subsequent experiments.

Figure 4. Small interfering RNAs interfere with PGC-1α mRNA. *P<0.05 vs. Seq.2 NC, negative control; Seq, sequence. PGC, peroxisome proliferator-activated receptor-γ coactivator.

Irisin partially protects mitochondria against oxidative stress after interfering with PGC-1α

Compared with the NC group, the MMP was significantly decreased (Fig. 5D) and ROS content was significantly increased (Fig. 5H) after interfering with PGC-1α, while the MMP was significantly increased (Fig. 5D) and the ROS content was significantly decreased (Fig. 5H) following addition of irisin.

Figure 5. Detection of MMP and ROS content in H9c2 cardiomyocytes following interference with PGC-1α. MMP (A) NC, (B) siRNA and (C) siRNA/irisin group. (D) MMP depolarization. ROS (E) NC, (F) siRNA and (G) siRNA/irisin group. (H) Median ROS content in H9c2 cells. *P<0.05, **P<0.01 vs. NC; #P<0.05, ##P<0.01 vs. siRNA. MMP, mitochondrial membrane potential; ROS, reactive oxygen species; si, small interfering; NC, negative control; PGC-1α, peroxisome proliferator-activated receptor-γ coactivator-1 alpha.

Irisin partially activates mitochondrial homeostatic indicator expression following interference with PGC-1α

Certain factors associated with mitohormesis were expressed at the transcriptional and protein levels following treatment of H9c2 cells with PGC-1α siRNA and irisin. COX4, which is associated with mitochondrial biosynthesis, was significantly decreased at the transcriptional and protein levels following interference with PGC-1α (Fig. 6B and D) and UCP2, which is associated with plasmonic leakage, was significantly increased at the protein level. The expression of OPA1 and COX4 genes associated with mitochondrial fusion was significantly elevated by irisin (Fig. 6A and B), suggesting that irisin can partially alter the imbalance in energy metabolism resulting from interference with PGC-1α by increasing mitochondrial membrane fusion and biosynthesis.

Figure 6. After interfering with peroxisome proliferator-activated receptor-γ coactivator-1α expression in H9c2 cardiomyocytes, mitohormesis-associated factors were expressed at the transcriptional and protein levels after the addition of irisin. (A) OPA1 and (B) COX4 mRNA levels. (C) Western blot analysis and (D) semi-quantification. *P<0.05, **P<0.01 vs. NC; #P<0.05 vs. siRNA. NC, negative control; siRNA, small interfering RNA; OPA1, optic atrophy 1; COX4, cytochrome c oxidase subunit 4; UCP2, uncoupling protein 2.

Discussion

ROS are produced by the mitochondrial respiratory chain and are present in the form of H2O2, which, at a certain level, leads to a change in the permeability of the mitochondrial membrane. In H9c2 cells treated with >250 µM H2O2, viability was reduced by 53.21±5.66% (21), and severe oxidative stress damage, however, this damage eventually leads to the occurrence of heart-related diseases such as heart failure, cardiac hypertrophy (22) and autophagy (23). In the present study, exogenous H2O2 treatment of H9c2 cells induced an increase in ROS content, resulting in a significant decrease in MMP. Excessive ROS cause MMP depolarization, leading to decreased efficiency of the mitochondrial electron transport chain; these changes are associated with elevated mitochondrial outer membrane permeability (24,25). The addition of 75 nM irisin effectively reduced the levels of ROS in cardiomyocytes and MMP recovery was observed. A recent study also supported that exogenous irisin decreases ROS levels in high-glucose-induced cardiomyocytes while MMP is restored (26).

In myofibroblasts, PGC1-α/FNDC5/irisin signaling is associated with the expression of mitochondrial autophagy- and fission-related PINK1, Parkin, DRP1 and microtubule-associated protein 1 light chain 3β mRNA (27). In the present study, exogenous H2O2 induced an increase in ROS levels, leading to imbalance in indicators associated with mitohormesis. Irisin resulted in a decrease in DRP1 protein expression and increase in MFN2 and OPA1 protein expression, which suggested that the mitochondria of H9c2 cells converged from damaged fission to fusion and ensured the integrity of the mitochondrial membrane, as shown by the increase in MMP. Irisin inhibits mitochondrial fission associated with DRP1 by suppressing the JNK/LATS2 signaling pathway and attenuating sepsis-mediated myocardial depression and cardiomyocyte death (28). In a hypoxia-treated cardiomyocyte model, irisin not only exerts cardioprotective effects but also decreases oxidative stress damage by increasing antioxidant enzyme activity through the activation of OPA1-induced mitochondrial autophagy (29). However, the aforementioned study did not assess differences in the expression of OPA1 isoforms (L- and S-OPA1). Although all isoforms of OPA1 promote mitochondrial ridge remodeling, the long isoform promotes mitochondrial fusion and the short isoform can better restore energetic efficiency (30). Here, increased expression of the mitochondrial inner membrane fusion protein S-OPA1 ensured the structural integrity of the inner membrane to regulate mitochondrial respiration. Previous studies have confirmed that OPA1 acts in vivo by promoting the mitochondrial cristae remodeling pathway to not only enhance mitochondrial respiratory efficiency but also alleviate mitochondrial dysfunction, mtDNA depletion, and reactive oxygen species production (31–33), which is similar to the results of the present study.

PGC-1α serves numerous physiological functions. In PGC-1α-deficient mice, organs with high mitochondrial energy requirements, such as the heart and skeletal muscle, show impaired growth and reduced mitochondrial respiratory function (34). In the present study, after transfection with PGC-1α siRNA, MMP appeared to be significantly decreased, ROS levels were elevated and the content of proton leakage-associated protein UCP2 was significantly elevated. This indicated that interference with PGC-1α increased oxidative stress state of H9c2 cells. Previous studies have confirmed that PGC-1α deficiency significantly decreases the expression of cardiac mitochondrial antioxidant enzymes superoxide dismutase 2 and thioredoxin-2 (35). Although the upregulation of UCP2 mRNA resulted in restoration of myocardial MMP and reduction of intracellular ROS levels (36), this uncoupling effect did not prevent oxidative stress in H9c2 cells at this stage. The levels of myocardial mitochondrial biosynthesis also significantly decreased after interfering with PGC-1α in the present study. This may be associated with the blockage of the pathway of mitochondrial biosynthesis. Numerous studies have confirmed that activated PGC-1α binds nuclear respiratory factor 1 (NRF1)/2 and promotes expression of mitochondrial transcription factor A (TFAM), forming the PGC-1α/NRF1/2/TFAM signaling pathway that supports mtDNA replication and protein synthesis (37–39). However, this pathway is blocked at the PGC-1α locus, decreasing myocardial mitochondrial biosynthesis. By contrast, PGC-1α overexpression increased levels of genes involved in regulating mitochondrial function and biosynthesis and enhances antioxidant and anti-apoptotic capacity of the cells (40). This suggests interference with PGC-1α increases oxidative stress in mitochondria and disrupts homeostasis.

In the present study, when PGC-1α silencing was accompanied by irisin treatment, elevated MMP and decreased ROS levels were observed. By contrast, indicators associated with mitohormesis, mRNA level of OPA1 and the protein level of COX4 were significantly elevated. Previous studies have indicated that irisin enhances mitochondrial function and respiration (41). However, restoration of mitochondrial function is associated with UCP2, whose elevated expression regulates MMP and ROS (36). Although it has been suggested that exogenous irisin (20,42) upregulates UCP2, no association was observed in the present study. Possibly due to the compensatory effect of PGC-1β following interference of PGC-1α, PGC-1β overexpression upregulates UCP2 (43). Exogenous irisin also increases mitochondrial content and enhances expression of metabolic genes associated with mitochondrial biogenesis, such as PGC-1α, TFAM, NRF1 and glucose transporter protein 4 (42,44). Although PGC-1α was disrupted, disrupting homeostasis of mitochondria, irisin promoted increased mRNA expression of OPA1 linked to mitochondrial fusion and COX4 linked to biosynthesis at the protein level.

However, the present study did not assess effects of irisin on mitohormesis indices at the vector level. Therefore, future studies should induce oxidative stress in healthy and PGC-1α-deficient rats. Moreover, exogenous irisin was injected into the two groups of rats that had already undergone oxidative stress, and after a period of time, the mitohormesis indexes of the vector cardiomyocytes were detected.

In conclusion, irisin acts partially independent of PGC-1α signaling to regulate mitohormesis imbalance by oxidative stress and maintain energy metabolism by improving mitochondrial structure. The results suggested that irisin may be a potential target for treatment of heart disease associated with abnormal mitohormesis (45) but this needs to be confirmed in vivo.

Acknowledgements

Not applicable.

Funding

The present study was supported by Shandong Provincial Natural Science Foundation, China (grant no. ZR2023MC033), National Natural Science Foundation of China (grant no. 32100006), Science and Technology Support Plan for Youth Innovation of Colleges and Universities in Shandong Province (grant no. 2022KJ088) and Shandong Provincial Natural Science Foundation, China (grant no. ZR2023MD117).

Availability of data and materials

The data generated in the present study may be requested from the corresponding author.

Authors' contributions

BW and HX conceived and designed the study and wrote the manuscript. SS, CS, WD and LL performed experiments. CS and WD analyzed data. All authors have read and approved the final manuscript. BW, HX, SS and LL confirm the authenticity of all the raw data.

Ethics approval and consent to participate

Not applicable.

Patient consent for publication

Not applicable.

Competing interests

The authors declare that they have no competing interests.

References