Natural flavonoid from Citrus reticulata Blanco; anti-inflammatory, antiviral, antioxidant

It has been demonstrated that Naringenin inhibits HepG2 cell proliferation, which is partially due to cell accumulation in the G0/G1 and G2/M phases of the cell cycle. Nuclei damage and a higher percentage of apoptotic cells are indications that naringenin induces apoptosis. Increased Bax/Bcl-2 ratios, cytochrome C release, and caspase-3 activation all indicate that naringenin initiates the mitochondrial-mediated apoptosis pathway[1]. A431 cells exposed to naringenin exhibit a dose-dependent increase in nuclear condensation and DNA fragmentation along with a significant reduction in cell viability. Naringenin-induced cell cycle arrest in the G0/G1 phase of the cell cycle is demonstrated by cell cycle studies, and caspase-3 analysis reveals a dose-dependent increase in caspase-3 activity that results in cell apoptosis[2].

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Search for new substances with antiproliferative activity and apoptosis inducing potential towards HepG2 cells is important since HCC is notoriously resistant to conventional chemotherapy. Dietary phytochemicals with significant anti-proliferative and apoptosis inducing potential are considered as agents promising for cancer therapy. Naringenin, a common dietary flavonoid abundantly present in fruits and vegetables, is believed to possess strong cytotoxic activity in numerous types of cancer cells. However, the detailed molecular mechanisms of its antiproliferative effects and apoptosis induction are still unclear. In this study, we investigated antiproliferative and apoptosis-inducing effect of naringenin in human hepatocellular carcinoma HepG2 cells. Naringenin was shown to inhibit the proliferation of HepG2 cells resulted partly from an accumulation of cells in the G0/G1 and G2/M phase of the cell cycle. Naringenin induced a rapid accumulation of p53, which might account for the naringenin-induced G0/G1 and G2/M phase arrests in Hep G2 cells. In addition, naringenin have been shown to induce apoptosis as evidenced by nuclei damage and increased proportion of apoptotic cells detected by flow cytometry analysis. Naringenin triggered the mitochondrial-mediated apoptosis pathway as shown by an increased ratio of Bax/Bcl-2, subsequent release of cytochrome C, and sequential activation of caspase-3. Our results showed that naringenin had inhibitory effect on the growth of HepG2 cell line through inhibition of cell proliferation and apoptosis induction. The elucidation of the drug targets of naringenin on inhibition of tumor cells growth should enable further development of naringenin for liver cancer therapy. [1]

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A natural predominant flavanone Naringenin, especially abundant in citrus fruits, has a wide range of pharmacological activities. The search for antiproliferative agents that reduce skin carcinoma is a task of great importance. The objective of this study was to analyze the anti-proliferative and apoptotic mechanism of naringenin using MTT assay, DNA fragmentation, nuclear condensation, change in mitochondrial membrane potential, cell cycle kinetics and caspase-3 as biomarkers and to investigate the ability to induce reactive oxygen species (ROS) initiating apoptotic cascade in human epidermoid carcinoma A431 cells. Results showed that naringenin exposure significantly reduced the cell viability of A431 cells (p<0.01) with a concomitant increase in nuclear condensation and DNA fragmentation in a dose dependent manner. The intracellular ROS generation assay showed statistically significant (p<0.001) dose-related increment in ROS production for naringenin. It also caused naringenin-mediated epidermoid carcinoma apoptosis by inducing mitochondrial depolarization. Cell cycle study showed that naringenin induced cell cycle arrest in G0/G1 phase of cell cycle and caspase-3 analysis revealed a dose dependent increment in caspase-3 activity which led to cell apoptosis. This study confirms the efficacy of naringenin that lead to cell death in epidermoid carcinoma cells via inducing ROS generation, mitochondrial depolarization, nuclear condensation, DNA fragmentation, cell cycle arrest in G0/G1 phase and caspase-3 activation. [2]

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Vascular smooth muscle cell (VSMC) proliferation and migration, which is triggered by various inflammatory stimuli, contributes importantly to the pathogenesis of atherosclerosis and restenosis. Naringenin is a citrus flavonoid with both lipid-lowering and insulin-like properties. Here, we investigated whether Naringenin affects TNF-α-induced VSMC proliferation and migration and if so, whether heme oxygenase-1 (HO-1) is involved. Rat VSMCs were treated with naringenin alone or in combination of TNF-α stimulation. We found that naringenin induced HO-1 mRNA and protein levels, as well as its activity, in VSMCs. Naringenin inhibited TNF-α-induced VSMC proliferation and migration in a dose-dependent manner. Mechanistic study demonstrated that naringenin prevented ERK/MAPK and Akt phosphorylation while left p38 MAPK and JNK unchanged. Naringenin also blocked the increase of ROS generation induced by TNF-α. More importantly, the specific HO-1 inhibitor ZnPP IX or HO-1 siRNA partially abolished the beneficial effects of naringenin on VSMCs. These results suggest that naringenin may serve as a novel drug in the treatment of these pathologies by inducing HO-1 expression/activity and subsequently decreasing VSMC proliferation and migration. [3]

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Dengue is one of the most significant health problems in tropical and sub-tropical regions throughout the world. Nearly 390 million cases are reported each year. Although a vaccine was recently approved in certain countries, an anti-dengue virus drug is still needed. Fruits and vegetables may be sources of compounds with medicinal properties, such as flavonoids. This study demonstrates the anti-dengue virus activity of the citrus flavanone naringenin, a class of flavonoid. Naringenin prevented infection with four dengue virus serotypes in Huh7.5 cells. Additionally, experiments employing subgenomic RepDV-1 and RepDV-3 replicon systems confirmed the ability of naringenin to inhibit dengue virus replication. Antiviral activity was observed even when naringenin was used to treat Huh7.5 cells 24 h after dengue virus exposure. Finally, naringenin anti-dengue virus activity was demonstrated in primary human monocytes infected with dengue virus sertoype-4, supporting the potential use of naringenin to control dengue virus replication. In conclusion, naringenin is a suitable candidate molecule for the development of specific dengue virus treatments [6].

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BACKGROUND AND PURPOSE. The aim of this study was to investigate, in vascular smooth muscle cells, the mechanical and electrophysiological effects of (+/-)-naringenin. Experimental approach: Aorta ring preparations and single tail artery myocytes were employed for functional and patch-clamp experiments, respectively. Key results: (+/-)-Naringenin induced concentration-dependent relaxation in endothelium-denuded rat aortic rings pre-contracted with either 20 mM KCl or noradrenaline (pIC(50) values of 4.74 and 4.68, respectively). Tetraethylammonium, iberiotoxin, 4-aminopyridine and 60 mM KCl antagonised (+/-)-naringenin-induced vasorelaxation, while glibenclamide did not produce any significant antagonism. Naringin [(+/-)-naringenin 7-beta-neohesperidoside] caused a concentration-dependent relaxation of rings pre-contracted with 20 mM KCl, although its potency and efficacy were significantly lower than those of (+/-)-naringenin. In rat tail artery myocytes, (+/-)-naringenin increased large conductance Ca(2+)-activated K(+) (BK(Ca)) currents in a concentration-dependent manner; this stimulation was iberiotoxin-sensitive and fully reversible upon drug wash-out. (+/-)-Naringenin accelerated the activation kinetics of BK(Ca) current, shifted, by 22 mV, the voltage dependence of the activation curve to more negative potentials, and decreased the slope of activation. (+/-)-Naringenin-induced stimulation of BK(Ca) current was insensitive either to changes in the intracellular Ca(2+) concentration or to the presence, in the pipette solution, of the fast Ca(2+) chelator BAPTA. However, such stimulation was diminished when the K(+) gradient across the membrane was reduced. Conclusions and implications: The vasorelaxant effect of the naturally-occurring flavonoid (+/-)-naringenin on endothelium-denuded vessels was due to the activation of BK(Ca) channels in myocytes [7].

The level of cholesterol and total triglycerides in plasma and the liver are significantly reduced when Naringenin supplementation is taken. Furthermore, naringenin administration reduces the levels of triglycerides and adiposity in parametrial adipose tissue. The livers of rats administered neringenin exhibit a substantial upregulation of the PPARα protein. Treatment with naringenin dramatically increases the expression of CPT-1 and UCP2, which are known to be controlled by PPARα[3]. Naringenin enhances the oxidation of hepatic fatty acids by means of a transcription program driven by PPARγ coactivator 1α/PPARα. By lowering fasting hyperinsulinemia, it inhibits the liver's and muscle's sterol regulatory element-binding protein 1c-mediated lipogenesis. Hepatic cholesterol and the production of cholesterol ester are reduced by naringenin[4]. Naringenin exhibits dose-dependent inhibition of TNF-α-induced VSMC migration and proliferation. Naringenin inhibits p38 MAPK and JNK but not ERK/MAPK and Akt phosphorylation, according to mechanistic research. Additionally, naringenin prevents TNF-α-induced increases in ROS production [5].

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Naringenin supplementation caused a significant reduction in the amount of total triglyceride and cholesterol in plasma and liver. In addition, naringenin supplementation lowered adiposity and triglyceride contents in parametrial adipose tissue. Naringenin-fed animals showed a significant increase in PPARα protein expression in the liver. Furthermore, expression of CPT-1 and UCP2, both of which are known to be regulated by PPARα, was markedly enhanced by naringenin treatment.
Conclusions: Our results indicate that the activation of PPARα transcription factor and upregulation of its fatty acid oxidation target genes by dietary Naringenin may contribute to the hypolipidemic and anti-adiposity effects in vivo. [4]

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Objective: The global epidemic of metabolic syndrome and its complications demands rapid evaluation of new and accessible interventions. Insulin resistance is the central biochemical disturbance in the metabolic syndrome. The citrus-derived flavonoid, naringenin, has lipid-lowering properties and inhibits VLDL secretion from cultured hepatocytes in a manner resembling insulin. We evaluated whether Naringenin regulates lipoprotein production and insulin sensitivity in the context of insulin resistance in vivo.

Research design and methods: LDL receptor-null (Ldlr(-/-)) mice fed a high-fat (Western) diet (42% calories from fat and 0.05% cholesterol) become dyslipidemic, insulin and glucose intolerant, and obese. Four groups of mice (standard diet, Western, and Western plus 1% or 3% wt/wt Naringenin) were fed ad libitum for 4 weeks. VLDL production and parameters of insulin and glucose tolerance were determined.

Results: We report that Naringenin treatment of Ldlr(-/-) mice fed a Western diet corrected VLDL overproduction, ameliorated hepatic steatosis, and attenuated dyslipidemia without affecting caloric intake or fat absorption. Naringenin 1) increased hepatic fatty acid oxidation through a peroxisome proliferator-activated receptor (PPAR) gamma coactivator 1alpha/PPARalpha-mediated transcription program; 2) prevented sterol regulatory element-binding protein 1c-mediated lipogenesis in both liver and muscle by reducing fasting hyperinsulinemia; 3) decreased hepatic cholesterol and cholesterol ester synthesis; 4) reduced both VLDL-derived and endogenously synthesized fatty acids, preventing muscle triglyceride accumulation; and 5) improved overall insulin sensitivity and glucose tolerance.

Conclusions: Thus, Naringenin, through its correction of many of the metabolic disturbances linked to insulin resistance, represents a promising therapeutic approach for metabolic syndrome. [5]

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Effects of Naringenin supplementation on food intake, weight gain, and organ weight [4]
Naringenin did not affect the average food intake during the 6-week experiment period (data not shown). Although the mean body weights of the naringenin-supplemented groups relative to the control group tended to increase, they were not significantly different. No significant differences were observed in liver weights across all the groups after 6 weeks of treatment (data not shown).

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Naringenin decreases adipose triglyceride (TAG) [4]
The weight of parametrial fat pad, consisting of epididymal and perirenal, was decreased in naringenin-fed rats. Compared to the control group, naringenin treatment decreased adipose tissue weight by 24.6, 14.9, and 41.1% at dietary naringenin concentrations of 0.003, 0.006, and 0.012%, respectively (Fig. 1). The difference in the parametrial adipose tissue weight at 0.012% naringenin treatment and control was significant (p < 0.05), although the differences in other groups did not reach the significance. Since triglyceride is the major components in adipose tissue, we measured adipose tissue triglyceride amounts. Consistent with the changes in parametrial fat pad weights, naringenin supplementation at highest dose significantly decreased TAG amounts in adipose tissue (Fig. 1).

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Naringenin decreases plasma triglyceride and cholesterol level [4]
Naringenin supplementation resulted in a significant lowering of the blood triglyceride levels compared to that of the control group (Table 2). Although naringenin dose-dependently lowered plasma triacylglycerol concentrations (6, 42 and 55% of control at 0.003, 0.006, and 0.012%, respectively), the difference among groups did not reach the statistical significance. The plasma total and free cholesterol concentrations were also significantly lower in the 0.006% naringenin treatment compared to the control group. However, plasma-free fatty acids concentrations were not different among groups (Table 2).

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Naringenin lowers hepatic triglyceride and cholesterol levels [4]
Hepatic triglyceride levels were decreased in rats fed a naringenin-supplemented diet relative to those of animals fed a control diet (Table 3). However, only 0.012% naringenin decreased hepatic triglyceride concentrations with a statistical significance (p < 0.05). Cholesterol concentrations in liver were also decreased in naringenin-fed rats compared to the control group, but only 0.006% naringenin resulted in a significant reduction of the hepatic cholesterol levels compared to that of the control group (p < 0.05).

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Naringenin increases hepatic PPARα, CPT-1, and UCP2 protein expression [4]
To examine whether the decrease in the plasma and hepatic triglyceride levels is due to increased hepatic fatty acid oxidation, we analyzed the expression of PPARα, a key transcriptional regulator of lipid metabolism and fatty acid β-oxidation, and 0.012% naringenin significantly induced hepatic PPARα protein expression as shown in Fig. 2. Quantitative analysis showed that PPARα protein abundance was increased by 25, 40, and 50% in 0.003, 0.006, and 0.012% naringenin-fed groups relative to the control group, respectively. We further assessed whether activation of PPARα by naringenin treatment leads to increased expression of PPARα-regulated genes such as CPT-1 and UCP2 in the liver [27–29]. Consistent with increased PPARα protein expression, naringenin dose-dependently increased CPT-1 and UCP2 protein amounts (112, 110, and 130% increase for CPT-1, and 113, 125, and 127% increase for UCP2 in 0.003, 0.006, and 0.012% naringenin group, respectively). The protein levels of PPARα, CPT1, and UCP2 in 0.012% naringenin were significantly higher than those of the control group (Fig. 2).

Antiviral Activity Assays [6]
The antiviral activity of Naringenin was assessed in two different assays. Huh7.5 cells at concentration of 2 × 104 cells/well in 96-well plates were infected with one representative strain of each of the four DENV serotypes (DENV-1/FGA/89, DENV-2/ICC265, DENV-3/5532 and DENV-4/TVP360) at a multiplicity of infection (MOI) of 10 for 90 minutes. Naringenin at its NTC was added during and after infection. Positive and negative controls for antiviral activity were included using IFN-α 2A (200 IU/mL) and mock-infected cells, respectively. After 72 h of incubation, the cell culture supernatants were stored at −86 °C for virus titration. The cells were detached and stained for flow cytometry33. A BD FACS Canto II was used to quantify cells infected by DENV. Cell culture supernatants were also employed for titration in a foci-forming immunodetection assay in C6/36 cells to confirm the FACS results44. Additionally, dose response curves were obtained with a serial dilution starting at the NTC of Naringenin. The concentration that inhibited 50% of virus infection (IC50) was obtained using nonlinear regression, followed by the construction of a sigmoidal concentration-response curve (variable slope; GraphPad) and calculation of the selectivity index (SI = CC50/IC50). All assays were performed in triplicate.

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Virucidal Assay [6]
A virucidal assay was performed as previously described with minor modifications34. Briefly, a sample of each DENV serotype (2 × 105 ffu/mL; DENV-1/FGA/89; DENV-2/ICC-265; DENV3/5532 and DENV-4/TVP360) was treated with Naringenin (250 μM) in the presence or absence of 150 μg/mL RNase A for 1 h at 37 °C. After treatment, viral RNA was extracted using a QIAamp Viral RNA Mini Kit. The RNA was reverse transcribed using 250 pmol of a random primer and Improm II Reverse Transcriptase. PCR amplification was performed as described by Lanciotti et al.45 with a D1 (5′-TCAATATGCTGAAACGCGCGAGAAACCG-3′) and D2 (5′-ATTGCACCAGCAGTCAACGTCATCTGGTTC-3′) primer pair. DENV RNA samples treated with RNase or left untreated were used as the positive and negative control, respectively.

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Time-of-Drug Addition Assay [6]
Initially, Huh7.5 cells at a density of 2 × 104 cells/well in 96-well plates were (i) treated with Naringenin for 1 h prior to DENV infection; (ii) treated with naringenin during DENV infection; (iii) treated with naringenin after DENV infection; or (iv) treated with Naringenin during and after DENV infection. An MOI of 10 was used for DENV infection, and naringenin was tested at a concentration of 250 μM. The percentage of infected cells was determined using FACS as previously described. In another experiment, cells were infected with DENV-1 strain FGA/89 (MOI: 10) as previously described and treated with Naringenin (250 μM) or with IFN-α 2A (200 IU/mL) at different time points after infection (0, 1, 2, 4, 6, 24, 30 and 48 h). The percentage of infected cells was determined using FACS. The supernatants were titrated in a foci-forming immunodetection assay 72 h post-infection.

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Naringenin Impairs DENV Replication in Huh7.5 Cells [6]
After confirming the anti-DENV effects of naringenin after viral entry into Huh7.5 cells, we determined if this flavanone impairs DENV replication. Two subgenomic replicons derived from DV1-BR/90 (RepDV1) and BR DEN3 290-02 (RepDV3) were used in these experiments27,28. DENV replicon RNAs were obtained and used to transfect Huh7.5 cells (2 μg of RNA/2 × 106 cells) in a Nucleofector 2B device according to the manufacturer’s instructions33. After 1 h of transfection, the Huh7.5 cells were treated with naringenin (250 μM). The plates were further incubated for 72 h. After incubation, the cells were recovered and subjected to FACS analysis, as described previously. As a negative control, Huh7.5 cells were transfected without RNA and were not treated. Huh7.5 cells transfected with RNA (from the DENV-1 or DENV-3 replicon) and untreated cells were used as positive controls for virus replication. The reference control consisted of treatment with recombinant IFN-α 2A (200 IU/mL) and 20 μM ribavirin.

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Antiviral Effects in Primary Human Cells [6]
PBMCs obtained from healthy donors who provided signed informed consents were purified using Ficoll-histopaque and a classical protocol46. After purification, PBMCs were plated onto 24-well plates at a density of 1 × 106 cells/well and treated with 200 μM of Naringenin for 5 days and then the exposure of annexin V/7-AAD was evaluated. Also, we quantified the number of human monocytes (CD14+) after treatment with naringenin (250, 125 and 62.5 μM) for five days. After it was established that the NTC of naringenin for human monocytes was 62.5 μM, infection with dengue was performed. Human PBMCs were plated onto 24-well plates at a density of 1 × 106 cells/well and infected with DENV-4 TVP/360 (MOI: 10) for 2 h. The inoculum was removed, and cells were treated with naringenin (62.5 μM), IFN-α 2A (200 IU/mL) or RPMI media. After incubation for 5 days at 37 °C and 5% CO2, cells were stained for DENV E protein (mAb 4G2-FITC conjugated) and with a mouse anti-human CD14 PE-Cy7 antibody47. CD14 expression was used to gate the monocytes. The percentage of DENV E-positive cells (4G2+) in this population was calculated.

Cell Viability Assay [1]
Cells were plated in 96-well plates at a density of 8×103 cells per well and incubated for 24 h with medium. The cells were rinsed with PBS and grown in a medium containing various concentrations of Naringenin (50, 100, 150, 200, 250, 300 μM). The solvent DMSO treated cells were served as control. After 24 hrs of treatment, the medium was removed and replaced by another medium containing 3-(4,5 dimethylthiazol-2yl)-2,5-diphenyl-2H-tetrazolium bromide (MTT) solution (1 mg/mL), and the cells were incubated for 2 h at 37 °C. To assess the proportion of viable cells, formazan was solubilized with 150 μL DMSO. Plates were then vortexed at room temperature for 30 mins, and the level of formazan was measured using a spectrophotometer at 575 nm.

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Observation of Morphological Changes of Cells [1]
Hep G2 cells grown on 6 cm dishes were treated with Naringenin at different dosages of 100, 150 and 200 μM for 24 h. The morphological changes were observed under an inverted microscope.

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Nuclear Staining with DAPI [1]
After treating Hep G2 cells with Naringenin (100, 150 and 200 μM) for 24 h, the cells were harvested, washed in icecold phosphate-buffered saline (PBS) and fixed with 3.7 % paraformaldehyde in PBS for 10 min at room temperature. The fixed cells were washed with PBS and stained with a 4,6-diamidino-2- phenylindile solution for 10 min at room temperature. The nuclear morphology of the cells was examined by fluorescent microscopy.

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Cell Cycle Analysis [1]
HepG2 cells were treated for 24 h with increasing concentrations of Naringenin (100, 150 and 200 μM). At the end of treatment, cells were trypsinized, and the resulting cell suspensions were centrifuged at 1000 rpm for 5 min. The cells were fixed overnight in 70 % ethanol at 4 °C and centrifuged at 1000 rpm for 5 min, and the pellets were washed twice with ice-cold PBS. Cell pellets were then resuspended in 0.5 ml of PBS containing 50 μg/ml propidium iodide and 100 μg/ml RNase A, incubated at 37 °C for 30 min, and then analyzed by flow cytometer. Cellular DNA content was analyzed by flow cytometr). At least 10 000 cells were used for each analysis, and the results were displayed as histograms. The percentage of cell distribution in G0/G1, G2/M and S phase were measured and the results were analyzed by the Modfit LT version 5.2 software for cell cycle profile.

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Western Blot Analysis [1]
Cells were treated with DMSO (control) and with different concentrations of Naringenin (100, 150 and 200 μM). After 24 h of stimulation, floating cells and adherent cells were collected and washed three times with ice-cold PBS and harvested in lysis buffer. The lysate was centrifuged at 12,000 rpm for 15 min at 4 °C, and the supernatant was collected. The protein concentration of lysates was determined by Bradford method. SDS-PAGE was performed using equivalent protein extracts (55 μg) from each sample according to Laemmli. The resolved proteins were electrophoretically transferred to polyvinylidene difluoride membranes.

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MTT assay for cell viability in HaCaT and A431 cells [2]
This assay is based on the enzymatic reduction phenomenon of 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) dye and provides a direct relationship between the viable cells and absorbance. The effect of Naringenin on cell viability was evaluated by MTT reduction assay as performed earlier [18]. Selective doses viz. 50 µM, 100 µM, 200 µM, 300 µM, 400 µM, 500 µM and 750 µM of naringenin were prepared in dimethyl sulfoxide (DMSO) in final volume of 100 µl media. After 21 h exposure, 10 µl of MTT solution (5 mg/ml stock solution) was added in each well re-incubated for 3 h at 37°C until formazan blue crystal developed. Media was discarded from each well and 100 µl of DMSO was added to dissolve formazan crystals for 10 min at 37°C. The absorbance was recorded at 540 nm by microplate reader (BIORAD-680) and relative percentage cell viability was evaluated.

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Cell morphology analysis [2]
The effect of Naringenin was analyzed for morphological changes in the cultured cells. The cells were seeded at a density of 1×104 cells/well in 96-well culture plate. After overnight incubation, the cells were treated with different concentrations of naringenin for 24 h. The cellular morphology was observed under inverted phase contrast microscope.

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Reactive oxygen species (ROS) activity assay [2]
Microscopic fluorescence imaging was used to study ROS generation in A431cells after exposure to different concentrations of Naringenin. Cells (1×104 per well) were seeded as described above for the MTT assay. Cells were then exposed to 50 µM, 100 µM, 200 µM, 300 µM, 400 µM, 500 µM and 750 µM concentrations of naringenin for 12 h. Cells were incubated with 2,7-Dichlorodihydrofluorescein diacetate (DCFH-DA) (10 mM) for 30 min at 37°C. The reaction mixture was aspirated and replaced by 200 µl of phosphate-buffered saline (PBS) in each well. The plate was kept on a shaker for 10 min at room temperature in the dark. An inverted fluorescent microscope was used to visualize intracellular fluorescence of cells and to capture images. For quantitative ROS analysis, cells (1×104 per well) were re-seeded in 96-well black bottom culture plate and allowed to adhere for 24 h in a CO2 incubator at 37°C. A431 cells were treated with different concentrations of naringenin for 12 h. After exposure, cells were incubated with DCFH-DA (10 mM) for 30 min at 37°C. Fluorescence intensity was measured by multiwell micro-plate reader at excitation wavelength of 485 nm and emission wavelength of 528 nm. Values were expressed as the percentage of fluorescence intensity relative to the control wells.

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Analysis of cellular DNA contents by flow cytometry [2]
Cell cycle phase distribution with cellular DNA contents were carried out using flow cytometry. A431 cells were seeded into 6-well plate at density 1×106 cells/ml and treated with 100 µM, 300 µM and 500 µM of Naringenin for 24 h in 5% CO2 incubator and at 37°C temperature. After 24 h incubation, the cultured cells were harvested, washed with cold PBS, fixed in 70% ethanol and treated with RNase A (10 mg/ml). Fixed cells were stained with propidium iodide (PI) dye followed by incubation for 30 min at room temperature in dark. The PI fluorescence of individual nuclei was measured using flow cytometer. Data were analyzed with the Cell Quest Pro V 3.2.1 software.

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Cell culture [3]
VSMCs were isolated from the thoracic aortas of 3- to 4-week-old male Sprague-Dawley rats as described previously (Gordon et al., 1986). Cells at passages 4–8 were used in experiments. To assess the effects of Naringenin on cultured VSMCs, we preincubated the cells with naringenin in serum-free medium and then challenged cells with 100 ng/ml TNF-α. 10−4 mM ZnPP IX was given 1 h in advance of naringenin treatment when required. An alternative approach to knock down HO-1 expression is siRNA transfection. The rat HO-1-specific siRNA (5′-CCG UGG CAG UGG GAA UUU AUG CCA U-3′) and nonspecific siRNA (5′-CCG ACG GUG AGG UUA UAU CGUGCA U-3′) (serving as a negative control) were designed by using siRNA TargetFinder software and synthesized by Invitrogen. Each siRNA was transfected into VSMCs using X-tremeGENE HP transfection reagent according to the manufacturer’s instructions. 24 h after transfection, VSMCs were treated as described previously.

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Proliferation assay [3]
Cell proliferation was analyzed by using MTT (Marshall et al., 1995) and direct cell counting assays. For the MTT assay, VSMCs were seeded in 96-well plates (1 × 104 cells per well). The cells were pretreated with naringenin (25 or 100 μM) for 1 h, and then stimulated with or without TNF-α (100 ng/ml) for 24 h in the presence of Naringenin. After that, MTT (0.2 mg/ml) was added to each well and incubated for 4 h. The supernatant was removed and the formazan crystals were dissolved in DMSO. Cell proliferation was assessed by measuring the absorbance at 550 nm using a microplate reader. MTT analysis was also confirmed by direct cell counting and BrdU incorporation assays. For the cell counting, VSMCs were seeded in 6-well plates (5 × 104 cells per well). After a similar treatment, cells were resuspended with 0.05% trypsin and 0.02% EDTA and counted by a hemocytometer. For the BrdU incorporation assay, 10 μM BrdU was added into medium for 2 h when all the treatments were done, and cells were fixed with 4% paraformaldehyde and treated with HCl. BrdU incorporation determined using anti-BrdU antibody and Alexa Fluor 555. Cells were counterstained with DAPI (Sigma). Signals were visualized by a fluorescence microscope, and the average ratios between BrdU-positive (pink) and total DAPI-stained nuclei (blue) were counted for statistic analyses.

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Migration assay [3]
TNF-α-dependent chemotaxis of VSMCs was assessed with a 24-well modified Boyden chamber containing fibronectin-coated polycarbonate membranes (8 μm pore-size) (Grotendorst et al., 1981). Briefly, the lower wells of the chamber were filled with phenol red-free DMEM supplemented with/without 100 ng/ml TNF-α in the presence or absence of Naringenin, as indicated in the figure legends. The filters were coated with 50 mg/ml fibronectin and fixed atop the bottom wells. 1 × 105 per well VSMCs were allowed to migrate for 6 h and non-migrated cells were removed from the upper side of the membrane with cotton swabs. Cells on the lower side of the membrane were stained with Hoechst 33342, and then counted in five randomly selected squares per well with a fluorescence microscope. Data was presented as numbers of migrated cells per field. Cell migration was also assessed by the scratch wound motility assay (Majack and Clowes, 1984). For this assay, VSMCs were seeded in 6-well plates (1.5 × 105 cells per well) and grew to confluence. 24 h after serum deprivation, the cells were mounted to a reusable template to create a standard wound (<3 mm). The cells were then incubated with naringenin (25 or 100 μM) for 1 h followed by stimulation with or without TNF-α (100 ng/ml) for 24 h. Wound closure rates were followed with a reference point in the field of the wound at the bottom of the plate by direct microscopic visualization. The procedure permitted photographing the identical spot each time. The remaining cell-free area was determined via microphotography and performed immediately after 24-h injury.

Animals, diets, and experimental design [4]
Male Long-Evans hooded rats obtained at 3 weeks of age (40–50 g) were housed individually in animal cages in a room with controlled temperature (23 to 24 °C) and lightning (12-h light/dark cycle) and fed a commercial non-purified diet and water prior to receiving the dietary treatments. Animals were assigned to treatments so that each group was balanced for body weight and fed one of four experimental diets (n = 6) for 6 weeks. Semi-purified, powdered diets were prepared for concentrations of Naringenin: 0, 0.003, 0.006, and 0.012% of diet. After 7 days of acclimatization, rats were assigned to one of four groups, with six animals per group, and fed semi-purified experimental diets for 6 weeks. The experimental diets contained 16% fat, 45.5% sucrose, and different Naringenin concentration (0, 0.003, 0.006, or 0.012%) (Table 1). Rats had ad libitum access to food and water during the study period. Food intake and body weight were measured throughout the experiment. At the end of the experiment, the animals were fasted for 12 h before the intraperitoneal injection of ketamine (90 mg/kg) and xylazine (10 mg/kg) and euthanized by exsanguination. Blood was immediately collected into heparinized tubes for further analysis. Livers and other tissues were perfused with physiological saline, removed, weighed, and frozen in liquid nitrogen and stored at −80 °C until further analysis. [4]

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Male C57BL/6J and Ldlr−/− mice on the C57BL/6J background were housed in pairs and maintained at 23°C on a 12-h light/dark cycle. Experiments were approved by the animal care committee of the University of Western Ontario. Eight- to 12-week-old mice were fed ad libitum a rodent standard diet (4% of calories from fat, TD8604; Harlan Teklad) or a high-fat diet containing 42% of calories from fat plus cholesterol (0.05% wt/wt). Naringenin was added to the Western diet at 1 or 3% (wt/wt). Ldlr−/− mice were fed for 4 weeks and C57BL/6J mice for 30 weeks. Food intake was measured daily, and body weight was measured biweekly. Mice were fasted for 6 h before intervention [5].

Metabolism / Metabolites
Narigenin has known human metabolites that include Narigenin 7-O-glucuronide.

Naringenin Toxicity AVIAN Red-winged blackbird Oral LD50 (Lethal Dose 50) > 100 mg/kg
Naringenin Repellency AVIAN Red-winged blackbird Oral R50 (Repellency 50) > 1 %

[1]. Naringenin (citrus flavonone) induces growth inhibition, cell cycle arrest and apoptosis in human hepatocellular carcinoma cells. Pathol Oncol Res. 2013 Oct;19(4):763-70.

[2]. Induction of apoptosis and antiproliferative activity of naringenin in human epidermoid carcinomacell through ROS generation and cell cycle arrest. PLoS One. 2014 Oct 16;9(10):e110003.

[3]. Naringenin inhibits TNF-α induced VSMC proliferation and migration via induction of HO-1. Food Chem Toxicol. 2012 Sep;50(9):3025-31.

[4]. Dietary naringenin increases hepatic peroxisome proliferators-activated receptor α proteinexpression and decreases plasma triglyceride and adiposity in rats. Eur J Nutr. 2011 Mar;50(2):81-8.

[5]. Naringenin prevents dyslipidemia, apolipoprotein B overproduction, and hyperinsulinemia in LDLreceptor-null mice with diet-induced insulin resistance. Diabetes. 2009 Oct;58(10):2198-210.

[6]. The citrus flavanone naringenin impairs dengue virus replication in human cells. Sci Rep. 2017 Feb 3;7:41864.

[7]. (+/-)-Naringenin as large conductance Ca(2+)-activated K+ (BKCa) channel opener in vascular smooth muscle cells. Br J Pharmacol. 2006 Dec;149(8):1013-21.

Naringenin is a trihydroxyflavanone that is flavanone substituted by hydroxy groups at positions 5, 6 and 4'. It is a trihydroxyflavanone and a member of 4'-hydroxyflavanones.
5,7-Dihydroxy-2-(4-hydroxyphenyl)chroman-4-one has been reported in Trichoderma virens, Cadophora gregata, and other organisms with data available.
See also: Naringenin (annotation moved to).

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(S)-naringenin is the (S)-enantiomer of naringenin. It has a role as an expectorant and a plant metabolite. It is a Naringenin and a (2S)-flavan-4-one. It is a conjugate acid of a (S)-naringenin(1-). It is an enantiomer of a (R)-naringenin.
Naringenin has been reported in Camellia sinensis, Humulus lupulus, and other organisms with data available.
See also: Naringin (has subclass); Flavanone (subclass of).

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In conclusion, we are the first to provide evidence that naringenin is highly effective in inhibiting cell proliferation and inducing apoptosis cell death in human hepatocellular carcinoma Hep G2 cells and naringenin may be a promising candidate for hepatocarcinogenesis treatment. [1]

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In conclusion, the present study provides a novel insight into the mechanism of action of Naringenin-induced apoptosis in human skin carcinoma cells without/little affecting normal skin cells and shows a link between antiproliferative and apoptotic induction, and the cell death was due to the induction of ROS mediated mitochondrial membrane depolarization, nuclear condensation and DNA fragmentation. We also observed that naringenin induced cell cycle arrest at G0/G1 phase and caspase-3 activity. Our data confirm the potential of naringenin as an agent of chemotherapeutic and cytostatic activity in human skin carcinoma cancer and therefore, may be potentially valuable for application in drug developments. For further confirmation, molecular mechanism will be carried out to elucidate the molecular pathways. [2]

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In atherosclerosis, mitogenesis is induced in VSMCs via ERK phosphorylation/activation and in turn promotes VSMC proliferation and migration (Sprague and Khalil, 2009). Such process is mediated by activation of PI3K and its downstream target Akt (Chandrasekar et al., 2004), an important regulator involved in cell metabolism, growth and vascular remodeling (Dugourd et al., 2003). In the present study, we demonstrated that TNF-α stimulation significantly increased phosphorylated ERK1/2 and Akt levels in VSMCs, which was inhibited by Naringenin treatment. More importantly, these effects of naringenin were completely abolished by pretreatment with ZnPP IX. These data indicate: (1) the blockade of mitogenesis and p-ERK/p-Akt activity may serve as the molecular basis for the functions of naringenin; (2) given that intracellular ROS can activate MAPK family members such as ERK1/2, JNK, and p38 MAPK (Adhikari et al., 2006), the importance of inhibiting upstream second messengers such as ROS by naringenin should be highlighted when evaluating the molecular mechanisms mentioned above and (3) HO-1 induction at least dominantly, if not totally, mediates the naringenin-induced deactivation of VSMCs. Taken together, our findings suggest naringenin has profound effects on various pathways involved in VSMC pathological changes in atherosclerosis and HO-1 is one of the major targets of naringenin.
In conclusion, the present study demonstrates that Naringenin inhibits TNF-α-induced VSMC proliferation and migration. We also revealed the roles of HO-1 in the anti-inflammatory and antioxidant functions of naringenin in VSMCs. Naringenin can be therefore developed as a novel drug to treat cardiovascular diseases related to VSMC proliferation and migration such as atherosclerosis. In a broader context, our findings strongly suggest that enhancement of antioxidant defense has highly therapeutic values for atherosclerosis and HO-1 ranks one of the most promising drug targets due to its dynamically modulatory property. [3]

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In summary, dietary Naringenin exerts hypolipidemic and anti-adiposity effects in vivo at physiologically relevant concentrations. These effects can be explained in part by upregulation of hepatic PPARα and its target genes and by decreasing TAG level in adipose tissue. Together, our results indicate that decreased adipose TAG amounts by Naringenin treatment may increase the release of TAG, which is in turn diverted into hepatic fatty acid oxidation, resulting in lowered plasma level of TAG. However, we could not exclude the possibility that naringenin may also upregulate fatty acid oxidation in the adipose tissue itself. Future studies should address this to clarify a potential mechanism by which dietary naringenin elicits its beneficial effects in vivo.[4]

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The protective effect of Naringenin was not restricted to mice with Ldlr deficiency. In wild-type mice, we found that naringenin significantly reduced plasma and hepatic lipids, normalized glucose tolerance and insulin sensitivity, and prevented obesity compared with Western diet–fed mice.
Collectively, these findings demonstrate that Naringenin has marked lipid- and lipoprotein-lowering potential. Naringenin normalizes hepatic VLDL production, glucose tolerance, and insulin sensitivity and prevents hepatic steatosis and obesity associated with a high-fat diet. The ability of naringenin to modulate metabolic pathways linked to the metabolic syndrome suggests that these molecules represent valuable tools in the search for regulators of insulin signaling, lipid homeostasis, and energy balance. [5]

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In conclusion, data from multiple assays (flow cytometry, viral titration and replicon system) employed to assess infection with eleven different strains representing the four DENV serotypes in two cell types (the Huh7.5 cell line and primary human monocytes) support the ability of Naringenin to target DENV replication, making naringenin a suitable candidate for the treatment of DENV infection. Our results provide novel insights for the development of specific anti-DENV drugs to treat infected patients. [6]

C₁₅H₁₂O₅

272.25

272.068

C, 66.17; H, 4.44; O, 29.38

67604-48-2

Naringenin;480-41-1

932

Off-white to light yellow solid powder

1.485g/cm3

577.5ºC at 760mmHg

247-250 °C(lit.)

2.137

3

5

1

20

363

0

FTVWIRXFELQLPI-UHFFFAOYSA-N

InChI=1S/C15H12O5/c16-9-3-1-8(2-4-9)13-7-12(19)15-11(18)5-10(17)6-14(15)20-13/h1-6,13,16-18H,7H2

5,7-dihydroxy-2-(4-hydroxyphenyl)-2,3-dihydrochromen-4-one

(±)Naringenin; 5,7-Dihydroxy-2-(4-hydroxyphenyl)chroman-4-one; CCRIS 8135; EINECS 266-769-1; DTXSID50274239; (+-)-naringenin; DTXCID50196624; 266-769-1; 67604-48-2; (±) Naringenin

2934.99.9001

Powder      -20°C    3 years

                     4°C     2 years

In solvent   -80°C    6 months

                  -20°C    1 month

Room temperature (This product is stable at ambient temperature for a few days during ordinary shipping and time spent in Customs)

DMSO : ~120 mg/mL (~440.77 mM)
H2O : < 0.1 mg/mL

Solubility in Formulation 1: ≥ 3 mg/mL (11.02 mM) (saturation unknown) in 10% DMSO + 40% PEG300 + 5% Tween80 + 45% Saline (add these co-solvents sequentially from left to right, and one by one), clear solution.
For example, if 1 mL of working solution is to be prepared, you can add 100 μL of 30.0 mg/mL clear DMSO stock solution to 400 μL PEG300 and mix evenly; then add 50 μL Tween-80 to the above solution and mix evenly; then add 450 μL normal saline to adjust the volume to 1 mL.
Preparation of saline: Dissolve 0.9 g of sodium chloride in 100 mL ddH₂ O to obtain a clear solution.

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Solubility in Formulation 2: ≥ 3 mg/mL (11.02 mM) (saturation unknown) in 10% DMSO + 90% (20% SBE-β-CD in Saline) (add these co-solvents sequentially from left to right, and one by one), clear solution.
For example, if 1 mL of working solution is to be prepared, you can add 100 μL of 30.0 mg/mL clear DMSO stock solution to 900 μL of 20% SBE-β-CD physiological saline solution and mix evenly.
Preparation of 20% SBE-β-CD in Saline (4°C，1 week): Dissolve 2 g SBE-β-CD in 10 mL saline to obtain a clear solution.

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Solubility in Formulation 3: ≥ 3 mg/mL (11.02 mM) (saturation unknown) in 10% DMSO + 90% Corn Oil (add these co-solvents sequentially from left to right, and one by one), clear solution.
For example, if 1 mL of working solution is to be prepared, you can add 100 μL of 30.0 mg/mL clear DMSO stock solution to 900 μL of corn oil and mix evenly.

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 (Please use freshly prepared in vivo formulations for optimal results.)