MIN6 is a mouse insulinoma cell line obtained from National Centre for Cell Science (NCCS), Pune, India. MIN6 display many important characteristics that are similar to pancreatic islets (Ishihara et al., 1993). For example, MIN6 cells exhibit glucose-stimulated insulin secretion (GSIS) (Cheng et al., 2012). MIN6 cells were cultured in DMEM supplemented with 10% heat-inactivated FBS, 100 U/mL penicillin, 100 μg/mL streptomycin, and 2.0 mM glutamine (Purchased from GE Healthcare, Little Chalfont, United Kingdom) in a carbon dioxide incubator maintained at 37°C with 5% CO₂. MIN6 cells with passage number between 5 and 20 were used for all the experiments (Elango et al., 2016).

The effect of naringenin on the viability of MIN6 cells was measured using an MTT assay (Mosmann, 1983). Experimentally, first, MIN6 cells (2 × 10⁴ cells/well) were plated in 96-well plates and allowed to grow for 24 h in a CO₂ incubator. Next, the growing cells were exposed to increasing concentrations (0–200 μM) of naringenin (Sigma Chemical Company, St. Louis, MO, United States) for 24 h at 37°C. After treatment, cells were replenished with 90 μL phenol-red free media containing 10 μL MTT (5 mM) and incubated for additional 3 h in the CO₂ incubator. Media was aspirated, the precipitate was dissolved in 50 μL DMSO, and the absorbance measured at 540 nm using a plate reader (Infinite 1000, Tecan, Mannedorf, Switzerland). The experiments were performed in triplicate. The relative cell viability (%) compared to control cells treated with DMSO was calculated using: Cell viability (%) = (A_sample-A _blank)/A_control-A_blank) × 100. Since an about 35% cell death was observed at 200 μM, subsequent studies were conducted with naringenin concentrations < 200 μM.

To study the protective role of naringenin on STZ-induced cytotoxicity, first, the MIN6 cells were pretreated with increasing concentration of naringenin (0–100 μM) for 24 h. Next, the naringenin-treated cells were exposed to 10 mM STZ (Primary stock of 1.0M was prepared by dissolving in 0.1M Citrate buffer pH 4.5 followed by the addition of DMSO) for 1 h and the number of viable cells estimated using MTT assay. All experiments were performed in triplicates.

2 × 10⁴ MIN6 cells/ml were transiently transfected with Nrf2-Keap1 complementation system in a 12-well plate using Lipofectamine-2000 according to the manufacturer’s protocol (Invitrogen, Carlsbad, CA, United States). Six hours after transfection, the media was replaced with a fresh batch of medium, and cells treated with naringenin (25, 50, 100 μM) for 24 h. Control and treated cells were lysed in 1X lysis buffer (pH 7.8; Promega, Madison, WI, United States), protein lysates collected, and the debris separated by centrifugation at 10,000 g at 4°C for 5 min. Total protein was estimated using the Bradford reagent (Bio-Rad Laboratories Inc, Hercules, CA, United States). Next, 100 μL luciferase substrate (prepared by mixing 10 ml of luciferase assay buffer with the lyophilized Luciferin; Promega, Madison, WI, United States) was added to the 20 μL of supernatant containing 175.0 μg of total protein and the luciferase activity measured using a luminometer (Promega, Madison, WI, United States). The developed sensor system detects the potential of naringenin to stimulate the Nrf2-Keap1 complex dissociation. A fall in luciferase signal is inversely proportional to the activation of Nrf2. The results were presented as fold change of three independent experiments.

To check the effect of naringenin on Nrf2 translocation, nuclear and cytoplasmic extracts were separated using Pierce NE-PER^® kit according to the manufacturer’s guidelines (Pierce, Rockford, IL, United States). In brief, cells (2 × 10⁴ MIN6 cells/ml) were homogenized in CER-I buffer vigorous vortexing in the pre-extraction buffer and incubated on ice for 15 min. The cellular homogenate was centrifuged at 10,000 × g for 10 min at 4°C and the supernatant containing the cytoplasmic fraction separated. Next, nuclei present in the pellet were suspended in NER buffer (provided with protease inhibitors) and the nuclear fraction generated by thorough vortex mixing for 1 min with a break for every 10 min for 40 min. The vortexed samples were centrifuged at 16,000 × g for 15 min at 4°C, to separate the supernatant containing a nuclear fraction. The separated nuclear fraction was used for western blotting.

Total protein content in the cytoplasmic and nuclear fractions was quantified with the Bradford method using BSA as a standard (Kruger, 2009). Samples (100 μg) were denatured using sample buffer at 95°C for 5 min, proteins were separated on 4–12% SDS-PAGE gel (Bio-Rad, Philadelphia, PA, United States) and electroblotted onto a nitrocellulose membrane (Schleicher & Schuell, Keene, NH, United States). Primary and secondary antibodies against Nrf2, caspase-3, β-actin and lamin-B (Santa Cruz Biotechnology, Santa Cruz, CA, United States) were used to detect corresponding proteins. The blot was developed using ECL (Bio-Rad, Philadelphia, PA, United States) and the signals captured using a gel documentation system (GBOX, Syngene, United Kingdom).

NQO1-ARE-Luc and GST-ARE-Luc reporter constructs, given by Donna D. Zhang (College of Pharmacy, University of Arizona, Tucson, AZ, United States) to Dr. Ramkumar, were used for cell-based reporter gene assay. For more details about the construction of these reporter plasmids, readers can refer to (Ramkumar et al., 2013). ARE-Luc constructs (500 ng/well) were transiently transfected into MIN6 cells in 12-well plates using Lipofectamine-2000 as described previously (Ramkumar et al., 2013). After 6 h of transfection, the media was changed and the cells exposed to 25, 50, and 100 μM naringenin for 24 h. The cell lysates were collected and the luminometric assay was carried out according to Ramkumar et al. (2013). The increase in the luciferase activity compared to control DMSO-treated cells was represented. The results were represented as the averages of at least three independent experiments.

Levels of intracellular ROS were determined by flow cytometry using an oxidation-sensitive fluorescent dye, 2,7-dichlorodihydrofluorescein diacetate (H₂DCFDA) (Eruslanov and Kusmartsev, 2010). Experimentally, first, MIN6 cells (2 × 10⁴ cells/well in 100 μL) were treated with 50 and 100 μM naringenin in complete medium for 24 h. Next, naringenin-treated cells were exposed to 10 mM STZ for 1 h and the cells were incubated with 20 μM H₂DCFDA (10 μL) for 30 min at 37°C. The reaction was terminated by washing with phosphate-buffered saline (PBS) containing 10% fetal calf serum (FCS). Cells were centrifuged at x800 g for 10 min, washed, and resuspended in 1ml PBS. The fluorescence intensity was measured using FACS (BD Biosciences, San Jose, CA, United States). Data were analyzed using BD Cell QuestTM Pro Analysis software and the shift in fluorescence intensity caused by DCF production, which is an indicator of ROS generation, represented as a histogram.

To assess the efficacy of naringenin for protecting MIN6 cells from STZ-induced cell death, the FITC Annexin V/Dead Cell Apoptosis Kit (Invitrogen), was used (Hingorani et al., 2011). Experimentally, MIN6 cells were cultured in 6-well plates and treated with naringenin (50 and 100 μM) for 24 h. Next, naringenin-treated cells were exposed to 10 mM STZ. After 1 h, the cells were trypsinized, collected and washed with cold PBS. The supernatant was discarded and the cells resuspended in 200 μL of the 1X binding buffer, containing 50 mM HEPES, 700 mM NaCl, 12.5 mM CaCl₂, pH 7.4. To 100 μL cell suspension, 5 μL annexin-V-FITC and 1 μL of PI solution (100 μg/mL) were added. Then, the cells were incubated for 15 min at room temperature in dark, and 400 μL of 1X annexin-binding buffer added. The stained cells were examined by flow cytometry using Cell QuestTM Pro Analysis software.

Animal experiments were conducted after receiving approval from the JSS Medical College Institutional Animal Ethics Committee (JSSMC/IAEC/18/5675/DEC2013), Mysuru, India. In brief, male albino mice of 4–6 weeks old, weighing about 25–30 g, were procured from the Central Animal Facility, JSS Medical College and were maintained under standard laboratory conditions: viz., temperature (25 ± 2°C) and humidity with an alternating 12 h-12 h light/dark cycles.

All the experiments were carried out with at least three replicate measurements (intra experimental) and two independent analysis (duplicate analysis) Average values were calculated and the results expressed as mean ± SEM. The statistical significance was performed using one-way ANOVA, followed by Tukey’s post hoc test using SPSS software 20 (SPSS, Cary, NC, United States); P < 0.05 was considered significant.

In order to determine whether naringenin has the potential to activate master regulator of antioxidant proteins, i.e., Nrf2, the level of activated Nrf2 was measured using a complementation system in MIN6 cells (Ramkumar et al., 2013). MIN6 cells are known to contain very low endogenous antioxidant machinery compared to many other cells and express minimal Nrf2 (Grankvist et al., 1981). In addition, MIN6 cells are very sensitive to changes in antioxidant levels in the surroundings, hence, they respond quickly to various stress signals (Cheng et al., 2012). Therefore, MIN6 cells are the best suitable cells to study the effect of ROS inducers as well as to determine the ability of antioxidant molecules to trigger Nrf2 transcription factors (Elango et al., 2016). Experimentally, the ability of naringenin (25, 50, and 100 μM for 24 h) to dissociate the Nrf2-Keap1 complex was measured using the Nrf2-Keap1 complementation system in MIN6 cells (Elango et al., 2016). After treatment with naringenin, MIN6 cells were harvested and luciferase activity measured using a luminometer. The developed sensor system detects the potential of naringenin to stimulate the Nrf2-Keap1 complex dissociation. A fall in luciferase signal is inversely proportional to the activation of Nrf2 (Ramkumar et al., 2013). The data showed a dose-dependent increase in the Nrf2 activity (Figure 2A). For example, at 50 and 100 μM concentration of naringenin an about 3.9- and 4.8- fold increase, respectively, was observed (Figure 2A). At these concentrations of naringenin showed no cytotoxic symptoms (Figure 2B).

Since naringenin exhibited the ability to activate Nrf2 in complementation assay, next, we have determined the efficacy of naringenin to promote the translocation of Nrf2 by isolating the nuclear and cytoplasmic fractions of MIN6 cells exposed to 25, 50, and 100 μM naringenin for 24 h. The amount of Nrf2 in each fraction was measured using western blotting and normalized to the respective loading controls (Elango et al., 2016). The data showed a concentration-dependent increase in the nuclear Nrf2 with a concomitant decrease in the cytosolic fraction (Figure 3A,B).

Next, to further determine whether the translocated Nrf2 is functionally active and elevates the expression of target genes NQO1 and GST, a luciferase-based reporter assay was carried out using ARE-LUC-NQO1 and ARE-LUC-GST constructs (Ramkumar et al., 2013). MIN6 cells were transfected with luciferase-expressing ARE-NQO1 and ARE-GST constructs and the transfected cells treated with 25, 50, and 100 μM naringenin for 24 h. Cell lysates were collected and analyzed for the luciferase activity. Analysis of the data revealed a dose-dependent increase in the luciferase activity in cells harboring NQO1 and GST constructs indicating that naringenin-induced Nrf2 is functionally active and promotes the expression of its target genes NQO1 and GST (Figure 3C).

Streptozotocin (STZ) is a naturally occurring toxin produced by Streptomyces achromogenes. STZ is known to induces diabetes by specifically destroying pancreatic β-cells (Schnedl et al., 1994). Mechanistically STZ inhibits DNA synthesis, induce the generation of free radicals and nitric oxide in pancreatic β-cells thereby promotes pancreatic β-cells death (Schnedl et al., 1994). As a result of pancreatic β-cells destruction, the animals exposed to STZ develop diabetes (Eleazu et al., 2013). Since naringenin potentially upregulates the expression of functionally active Nrf2 in MIN6 cells, its’ administration is predicted to reduce the cellular oxidative stress thereby protect β-cells from undergoing apoptosis (induced by STZ). To test this hypothesis, first, the levels of apoptosis induced by STZ in MIN6 cells were measured and, next, the ability of naringenin to protect MIN6 cells from STZ-induced apoptosis was quantitated.

Since naringenin showed a better cytoprotective effect in the in vitro experiments, next, its ability to decrease the complications of diabetes, especially blood glucose, was tested in preclinical animal models. Experimentally, first, diabetes was induced by MLD-STZ administration (i.p.; 50 mg/kg body weight/day) for 5 days. As shown in Figure 6A, all mice showed hyperglycemia with a serum glucose level of 357 ± 13.6 mg/dl in comparison with control mice exposed to vehicle 0.5% CMC (99.5 ± 4.9 mg/dl). Oral administration of naringenin for 45 days caused a significant (p < 0.05) dose-dependent (50 and 100 mg/kg b.w.) reduction in blood glucose levels in STZ mice. For example, blood glucose levels of STZ mice were reduced from 357 ± 13.6 mg/dl to 141.25 ± 10.6 mg/dl after administration of naringenin at the dose of 100 mg/kg b.w. These results were comparable with positive control glibenclamide at 600 μg/kg b.w (Figure 6A). However, naringenin (100 mg/kg b.w.) treated control mice did not show any statistically significant (p < 0.05) difference when compared to that of control vehicle-treated mice (Figure 6A). In summary, naringenin could reduce the STZ-induced hyperglycemia in mice.

The anti-hyperlipidemic efficacy of naringenin was investigated in STZ treated mice by measuring the levels of lipids in serum (de Almeida et al., 2012) (Table 2). STZ administered animals showed significantly (P < 0.05) increased serum (a) total cholesterol (TC) from 75.03 ± 6.2 mg/dl to 181 ± 2.0 mg/dl, (b) LDL from 16.6 ± 8.1 mg/dl to 127.2 ± 2.2 mg/dl, (c) VLDL from 10.26 ± 0.5 mg/dl to 27.06 ± 1.3 mg/dl, (d) Triglycerides (TG) from 51.33 ± 2.5 mg/dl to 135.3 ± 6.5 mg/dl, (e) Phospholipids from 82.33 ± 4.8 mg/dl to 114.5 ± 7.8 mg/dl, (f) Free fatty acids (FFA) from 2.43 ± 0.4 mg/dl to 5.56 ± 0.2 mg/dl, and significantly decreased HDL level from 48.16 ± 3.1 mg/dl to 26.7 ± 1.24mg/dl (Table 2). Oral administration of naringenin (50 and 100 mg/kg b.w.) showed a significant (P < 0.05) reduction in TC, LDL, VLDL, TG, Phospholipids, and FFA. In addition, a significant (p < 0.05) increase in the concentration of HDL compared to STZ controls was noted (Table 2). For instance, TC was decreased from 181 ± 2.0 mg/dl to 106.6 ± 4.9 mg/dl. Likewise serum LDL was reduced from 127.2 ± 2.2 mg/dl to 47.56 ± 6.1 mg/dl and VLDL from 27.06 ± 1.3 mg/dl to 17.33 ± 1.1 mg/dl. Similarly, while triglyceride content was reduced from 135.3 ± 6.5 mg/dl to 86.66 ± 5.5 mg/dl the phospholipid level decreased from 114.5 ± 7.8 mg/dl to 87.05 ± 7.2 mg/dl upon exposure to naringenin. Furthermore, whereas free fatty acids showed a significant decrease from 5.56 ± 0.2 mg/dl to 2.76 ± 0.2 mg/dl, the level of HDL content increased from 26.7 ± 1.24 mg/dl to 41.76 ± 1.2 mg/dl with naringenin administration (Table 2). Administration of the positive control glibenclamide (600 μg/kg b.w.) to STZ-induced diabetic mice has restored the lipid profile to normal levels (Table 2). The observed anti-hyperlipidemic effect of naringenin could be due to either its insulin stimulatory effect on pancreatic β-cells or due to decreased cholesterogenesis and fatty acid synthesis.

Lipid peroxidation and formation of hydroperoxides are key indicators of tissue damage, which occurs due oxidative stress (El-Aal, 2012). Since STZ is known to promote cellular oxidative stress, first, the levels of lipid peroxides were measured by boiling 0.1 ml tissue homogenate with 2.0 ml of TBA:TCA:HCl reagent for 15 min followed by cooling, and centrifuging at 10,000 rpm for 5 min to separate clear supernatant. The levels of TBARS (mM/100 g tissue) were estimated by reading the absorption at 535 nm (Niehaus and Samuelsson, 1968). Table 3 shows a significant increase in the concentration of TBARS in pancreatic tissue from about 2 mM/100 g tissue to ∼4.2 mM/100 g tissue upon STZ treatment. Administration of 50 mg/kg b.w. and 100 mg/kg b.w. naringenin significantly reduced the TBARS in pancreatic tissue to ∼3.6 mM/100 g tissue and ∼2.8 mM/100 g tissue, respectively (Table 3). The positive control 600 μg/kg b.w. glibenclamide reduced the TBARS levels to ∼2.5 mM/100 g tissue (Table 3). Treatment of control animals (exposed to vehicle 0.5% CMC) with 100 mg/kg b.w. naringenin did not change TBARS levels (Table 3).

To estimate the concentration of hydroperoxides, 0.2 ml of tissue homogenate was incubated with 1.8 ml of Fox reagent at room temperature for 30 min and the absorbance measured at 560 nm (Jiang et al., 1992). The results were expressed as mM hydroperoxides produced per 100 g tissues. The STZ-treatment significantly elevated the tissue pancreatic hydroperoxides from 16 mM/100 g tissue to ∼23 mM/100 g tissue (Table 3). Elevated pancreatic hydroperoxides were brought down to about ∼20 mM/100 g tissue and ∼17 mM/100 g tissue, respectively upon 50 mg/kg b.w. and 100 mg/kg b.w. naringenin administration (Table 3). Oral administration of glibenclamide (600 μg/kg b.w.) decreased hydroperoxides levels (to ∼16 mM/100 g) that are similar to control animals (Table 3). Treatment of control animals with 100 mg/kg b.w. naringenin did not change hydroperoxides levels (Table 3).

Since, in vitro studies have demonstrated the Nrf2 activation potential of naringenin, we have hypothesized that the cytoprotective effects of naringenin might be mediated through similar mechanisms even in mice. Therefore, protein lysates were collected from the pancreatic tissues of control and experimental animals and subjected for western blotting. Analysis of the data showed an about 2.5-fold increase in the Nrf2 expression compared to control or STZ treated mice (Figure 7). However, interestingly, a slight non-significant increase in the Nrf2 expression (compared to control mice) was observed in STZ treated mice (Figure 7). It is well known that Nrf2 regulates the cellular oxidative stress by increasing the expression of its target genes NQO1, SOD, CAT, GPX, and GSH transferase (Jung and Kwak, 2010). Therefore, we have measured the activities of these Nrf2 target genes in the pancreatic tissue (Table 4). SOD is an enzyme, which catalyzes the dis-mutation of superoxide radicals to H₂O₂ thus decreasing the possibility of superoxide anion interacting with nitric oxide that forms reactive peroxynitrite (Fukai and Ushio-Fukai, 2011). The SOD activity in the tissues was assayed according to the method developed by Kakkar et al. (1984). The reaction was initiated by addition of NADH. After incubation for 90 s, the glacial acetic acid was added to stop the reaction and the colored complex formed was extracted into butanol layer and measured at 560 nm. The SOD activity was significantly (p < 0.05) decreased in pancreatic tissue from about ∼6.4 units/min/mg protein to ∼2.1 units/min/mg protein upon STZ injection (Table 4). Administration of 50 mg/kg b.w. and 100 mg/kg b.w. naringenin significantly augmented the SOD levels in pancreatic tissue to ∼4.5 units/min/mg protein and ∼5.5 units/min/mg protein, respectively (Table 4). The positive control 600 μg/kg b.w. glibenclamide augmented the SOD levels to ∼6.1 units/min/mg protein (Table 4). Treatment of control animals with 100 mg/kg b.w. naringenin did not change SOD levels (Table 4).

Catalase is a tetrameric hemin-enzyme, which decomposes hydrogen peroxide into water and molecular oxygen (Kirkman and Gaetani, 1984). The CAT activity (μmoles of H₂O₂ consumed/min/mg protein) in the tissues was assayed by the method developed by Sinha (1972). Dichromate is converted to chromic acetate after heating in the presence of hydrogen peroxide with the formation of an intermediate perchromic acid. The produced chromic acetate was measured at 590 nm. The CAT activity significantly decreased in pancreatic tissue from about 18 units in control animals to 7.8 units; upon STZ injection (Table 4). Administration of 50 mg/kg b.w. naringenin and 100 mg/kg b.w. naringenin significantly augmented the CAT activity in pancreatic tissue to ∼13.8 and ∼18.0 units, respectively (Table 4). The positive control 600 μg/kg b.w. glibenclamide significantly elevated the CAT activity, which is similar to the activity of control animals (Table 4). No significant change was observed in mice treated with 100 mg/kg b.w. naringenin in comparison to that of control group administered with 0.5% carboxymethyl cellulose demonstrating that naringenin administration has no major impact on CAT activity (Table 4).

GPX is a selenium-containing peroxidase, which decomposes H₂O₂ and lipid peroxides through GSH, as a hydrogen donor, into the water and protects the cells from free radicals (Lubos et al., 2011). The activity of GPX was determined by the method of Rotruck et al. (1973). The GPX activity (μmoles of GSH oxidized/min/mg protein) was significantly decreased in pancreatic tissue from about from 5.4 μmoles in control animals to ∼2.8 μmoles; upon STZ treatment (Table 4). Oral administration of 50 mg/kg b.w. naringenin and 100 mg/kg b.w. naringenin significantly augmented the GPX activity in pancreatic tissue to ∼4.8 and ∼5.4 μmoles; respectively (Table 4). The positive control 600 μg/kg b.w. glibenclamide significantly augmented the GPX activity in pancreatic tissue up to 5.8 μmoles compared to STZ control animals (Table 4). No significant change was observed even in non-diabetic mice treated with naringenin in comparison to that of the control group demonstrating that naringenin administration did not affect the GPX levels (Table 4).

GST is a GSH-dependent cytosolic enzyme, which protects cells from the damage caused by ROS (Hayes and McLELLAN, 1999). In order to determine whether administration of naringenin restored the GST activity, a method developed by Habig et al. (1974) was used and the results expressed as μmoles of CDNB conjugate formed /min/mg protein. Analysis of the data represented in Table 4 showed about 75% decrease in GST activity in the pancreatic tissue, compared to vehicle-treated mice, in STZ injected animals. Oral administration of naringenin (50 mg/kg and 100 mg/kg b.w.) and glibenclamide (600 μg/kg b.w.) significantly (p < 0.05) increased the GST activity levels. For example, administration of 100 mg/kg body weight naringenin elevated the pancreatic GST activity to ∼3.3 units (Table 4). Similarly, 600 μg/kg body weight glibenclamide elevated the pancreatic GST activity to ∼3.9 units (Table 4). No significant change in GST activity was observed between vehicle control and 100 mg/kg body weight naringenin-treated non-diabetic mice (Table 4).

GSH is a potent free-radical scavenger and co-substrate for GPX activity (Sies, 1999). Several studies have shown the ability of STZ to decrease cellular GSH levels (Birben et al., 2012). Therefore, to determine whether administration of naringenin could mitigate the effect of STZ-induced GSH depletion effect, pancreatic tissues were collected and processed to estimate GSH content using Ellman’s assay (Ellman, 1959). The data in Table 4 shows a significant decrease in pancreatic tissue GSH levels upon administering mice with STZ (from ∼11.28 mg/100 g tissue to ∼6 mg/100 g tissue). However, when naringenin or glibenclamide was administered the levels had gone up to the ones present in control animals (Table 4). However, no significant changes were observed in non-diabetic mice treated with naringenin in comparison to that of a control group of mice (Table 4).

Since STZ administration is known to damage pancreas by reducing the size and number of functionally active β-cells, an attempt was made to check whether naringenin and the positive control glibenclamide restored these altered morphological features (Bâlici et al., 2015). Control and naringenin (100 mg/kg b.w.) treated non-diabetic mice showed a normal pattern with clear and well-structured pancreatic islets (Figure 8). However, the pancreatic sections of STZ control mice displayed severe necrotic changes, exclusively in the center of pancreatic islets. In addition, disappearing of the nucleus with a comparative reduction in size was observed (Figure 8). Furthermore, pancreatic islets of STZ animals treated with naringenin (50 mg/kg b.w.) exhibited moderate expansion (Figure 8). Likewise, STZ animals treated with naringenin (100 mg/kg b.w.) showed a significant improvement in the pancreatic islet with distinct cellularity changes and increase in granulation, compared to the STZ control mice (Figure 8). The positive control 600 μg/kg b.w. glibenclamide showed healthy pancreas (Figure 8).

The immunohistochemical method was used to study the distribution pattern as well as number of functional β-cells in pancreatic islets of mice (Kim et al., 2009). Control (vehicle treated) and naringenin control (100 mg/kg b.w.) groups’ showed islets with a comparatively larger region of positive immuno-reactivity, demonstrating the existence of healthy cells secreting insulin in the pancreas (Figure 9). However, the STZ control islet cells showed distorted morphology with very few insulin-positive cells compared to vehicle controls (Figure 9). While 50 mg/kg b.w. naringenin showed a moderate increase in insulin immuno-reactivity, the ones received 100 mg/kg b.w. naringenin showed the relatively large area of positive immuno-reactivity with numerous brown insulin granules in the β-cells of pancreatic islets (Figure 9). Pancreatic section of STZ mice treated with positive control compound glibenclamide showed normal histo-architecture of the pancreas with insulin cells (Figure 9). Collectively, these observations conform the role of naringenin in increasing the insulin-positive cells in the pancreas.