Oleic Acid Activates Mitochondrial Energy Metabolism and Reduces Oxidative Stress Resistance in the Pancreatic β-Cell Line INS-1

Abstract

Elevated concentration of saturated fatty acids in plasma adversely affects pancreatic β-cells, but the effects of unsaturated fatty acids are controversial. In this study, we examined the effects of oleic acid (OA), a monounsaturated fatty acid, on mitochondrial function, which is important for insulin secretion, using INS-1 cells, a pancreatic β-cell line derived from rats. Observations of mitochondrial membrane potential and intracellular ATP concentration showed that the electron transport chain was enhanced and ATP production increased in cells treated with OA, indicating that the response that occurs from sensing an increase in glucose concentration to the production of ATP was accelerated. Measurements of intracellular reactive oxygen species (ROS) indicated that the rate of increase in ROS after glucose stimulation was significantly higher in OA-treated cells. The mRNA expression levels of superoxide dismutase 1 and 2, which are responsive to ROS and other substances, were significantly increased in OA 1-d treated cells, but decreased in OA 7-d treated cells. It can be inferred that continued exposure to high concentrations of OA reduced ROS processing capacity and increased intracellular ROS levels. The mRNA expression of apoptosis-inducing enzyme Caspase-3 was significantly increased in OA-treated cells, although its activity was not high. However, the apoptosis induction rate after H₂O₂ stimulation was significantly higher in OA-treated cells. The high OA environment was shown to promote mitochondrial energy metabolism, leading to an increase in glucose sensitivity and a decrease in oxidative stress resistance.

INTRODUCTION

Type 2 diabetes mellitus is a disease with diverse pathological conditions, and dysfunction of insulin secretion from pancreatic β-cells is a condition seen in about half of all Japanese patients with type 2 diabetes mellitus.^1–3)

It is known that the pancreatic β-cells are adversely affected by increased free fatty acid (FFA) concentrations in the blood, due to obesity and other causes.^4,5) In particular, the adverse effects of saturated fatty acids (SFAs) on pancreatic β-cells are significant and varied, and include suppression of insulin secretion⁶⁾ and elevated nitric oxide (NO) levels.^7,8) Among others, chronic increases in palmitic acid (PA) is known to be toxic to pancreatic cells.^9,10) Induction of apoptosis,^8,11,12) decreased cell viability,¹³⁾ decreased proliferative capacity and impaired insulin secretion¹¹⁾ have been reported. It has been shown that there are multiple pathways for their toxicity, including reactive oxygen species (ROS) and mitochondrial damage,^8,12) and activation of the endoplasmic reticulum (ER) stress pathway.^14–16)

On the other hand, unsaturated fatty acid (UFA)s are not considered toxic and have been reported to mitigate cytotoxicity caused by SFAs.^17–19) OA, a monounsaturated fatty acid (MUFA), has been reported to enhance glucose-stimulated insulin secretion (GSIS),^20–22) not cause apoptosis,^17,19,23) and have beneficial effects on insulin sensitivity.²⁴⁾ In addition, protective effects such as PA-induced reduction of cell viability and ER stress^13,18) have also been reported. However, there are also reports that oleic acid (OA) induces apoptosis^8,25) and its evaluation has not been settled. Few studies have examined the effects of OA on pancreatic β-cells, focusing on mitochondrial function. Mitochondria are involved in the glucose sensitivity of GSIS intracellular mechanisms, the regulation of ER stress through Ca²⁺ flux to/from the ER, and are also a source of ROS.

In this study, we focused on the effects of OA on mitochondrial function and oxidative stress in pancreatic β-cells, which were investigated with INS-1 cells, using a rat-derived pancreatic β-cell line, observed mainly by way of imaging techniques. The effects of OA on mitochondria were investigated by fluorescence observation of mitochondrial membrane potential, intracellular ATP production, and intracellular ROS levels. Oxidative stress was evaluated by mRNA expression levels of various stress marker proteins and apoptosis-related proteins, and apoptosis was evaluated by Caspase-3 (Casp3) activity and Annexin-V binding levels.

MATERIALS AND METHODS

Materials

OA, PA, β-mercaptoethanol and bovine serum albumin (BSA) obtained from Sigma-Aldrich (St. Louis, MO, U.S.A.). RPMI 1640 (11.1 mM glucose, Ref.No.11875), fetal bovine serum (FBS, Ref.10437), Sodium pyruvate, N-(2-hydroxyethyl)piperazine-N′-2-ethanesulfonic acid (HEPES) and Penicillin Streptomycin (10000 U/mL penicillin and 10000 µg/mL streptomycin) obtained from Gibco (Carlsbad, CA, U.S.A.). GO-ATeam Cyto2 provided by Professor Hiromi Imamura, Kyoto University (GO-ATeam). ISOGEN obtained from NIPPON GENE (Tokyo, Japan). PrimeScript™ RT reagent Kit with gDNA Eraser (Perfect Real Time) and TB Green^® Premix Ex Taq™ (Til RNase Plus) obtained from TaKaRa (Shiga, Japan). Tetramethylrhodamine methyl ester (TMRE) and Lipofectamin 3000 obtained from Invitrogen (Waltham, MA, U.S.A.). Annexin-V FITC Apoptosis Staining Detection Kit obtained from Abcam (Cambridge, U.K.). Caspase Glo 3/7 Assay System obtained from Promega (Madison, WI, U.S.A.).

Fluorescence measurements of mitochondrial membrane potential, intracellular ATP concentration, and intracellular ROS were performed using research inverted system microscope IX-81 (Olympus, Tokyo, Japan) equipped with an ORCA CCD camera and W-VIEW GEMINI (Hamamatsu Photonics, Shizuoka, Japan), and controlled by image acquisition and analysis software HC Image (Hamamatsu Photonics). Fluorescence observation of apoptosis was performed using a confocal laser microscope LSM 510 META (Carl Zeiss, Baden-Wurttemberg, Germany). Unless otherwise mentioned, observations were performed in Glc(−) KRBH (Krebs–Ringer bicarbonate buffer without glucose, 135 mM NaCl, 3.6 mM KCl, 0.5 mM NaH₂PO₄, 2.0 mM CaCl₂ 2H₂O, 0.5 mM MgSO₄, 10 mM HEPES, 2.0 mM NaHCO₃).

Cell Culture

Rat INS-1 pancreatic β-cells were originally developed by Prof. Claes Wollheim (Geneva, Switzerland) and supplied by Prof. Michael R. Duchen (London, U.K.) and Prof. Tomohisa Ishikawa (Shizuoka, Japan). INS-1 cells were cultured in a humidified atmosphere of 5% CO₂ at 37 °C in RPMI-1640 medium supplemented with 10% FBS, 100 units/mL penicillin, 100 µg/mL streptomycin, 50 µM β-mercaptoethanol, 1 mM sodium pyruvate, 10 mM HEPES. Unless otherwise stated, the glucose concentration was 11 mM.

Cell Treatment with Medium Containing FFA

The FFA, OA or PA, were dissolved in absolute ethanol at a concentration of 100 mM and diluted to a ratio of 1 : 200 with culture medium containing 0.8% FA-free BSA (final conc., 200 µM), and then incubated with shaking for 2 h at 37 °C. INS-1 cells were routinely incubated for 6–7 d in the medium containing PA or OA (FFA medium), changing medium every 2 d and provided for individual experiments. Details are given in the text of each experiment.

Quantitative Real-Time PCR

The mRNA levels of INS-1 cells cultured in medium supplemented with OA or PA for 1 d or every 2 d for 7 d with medium change, were analyzed by real-time PCR. Control cells were cultured in the absence of FA. Preparation of total RNA from cells was performed with ISOGEN. The total RNA was treated with PrimeScript™ RT reagent Kit with gDNA Eraser (Perfect Real Time) to eliminate genome DNAs and to synthesize cDNA. Real-time PCR was performed on QuantStudio 3 Real Time PCR System by using TB Green^® Premix Ex Taq™ (Til RNase Plus). DNA damage-inducible transcript 3 (Ddit3, CHOP), heat shock 70 kDa protein 5 (Hspa5) and heat shock protein 90 kDa β member 1 (Hsp90b1) were measured as ER stress-related proteins, superoxide dismutase 1 (SOD1) and SOD2 as oxidative stress-related proteins, and caspase-3 (Casp3) as an apoptosis marker. Primer sequences used were as follows: 5ʹ-TGGAAGCCTGGTATGAGGATCTG-3ʹ (forward) and 5ʹ-GAGGTGCTTGTGACCTCTGCTG-3ʹ (reverse) for Ddit3, 5ʹ-TCAGCCCACCGTAACAATCAAG-3ʹ (forward) and 5ʹ-TCCAGTCAGATCAAATGTACCCAGA-3 (reverse) for Hspa5, 5ʹ-CGATGTGGATGGCACGGTAG-3ʹ (forward) and 5ʹ-GTGATGCATTTAAACCATCCAACTG-3ʹ (reverse) for Hsp90b1, 5ʹ-AGCATGGGTTCCATGTCCATC-3ʹ (forward) and 5ʹ-AGCCACATTGCCCAGGTCTC-3ʹ (reverse) SOD1, 5ʹ-TTCTGGACAAACCTGAGCCCTAA-3ʹ (forward) and 5ʹ-GAACCTTGGACTCCCACAGACAC-3ʹ (reverse) for SOD2, 5ʹ-GAGACAGACAGTGGAACTGACGATG-3ʹ (forward) and 5ʹ-GGCGCAAAGTGACTGGATGA-3ʹ (reverse) for Casp3 and GGAGATTACTGCCCTGGCTCCTA (forward) and GACTCATCGTACTCCTGCTTGCTG (reverse) for β-actin. Data were shown as the fold differences normalized to the β-actin.

Observation of Mitochondrial Membrane Potential

Mitochondrial membrane potential was evaluated by TMRE fluorescence intensity. After 4 d of culture in FFA medium with medium change every 2 d, INS-1 cells were seeded at 1.5 × 10⁵ cells/cm² on 2.5 cm diameter cover glasses and cultured in FFA medium for an additional 2 d. Fluorescence observations were performed 17–22 h after replacement with low glucose (5 mM) medium without FFA. Cell density at observation was always around 80% confluency. Cells washed in Glc(−) KRBH were stained with TMRE (25 nM) in Glc(−) KRBH for 30 min at room temperature, light-shielded, washed twice again in Glc(−) KRBH, placed on the microscope stage (30 °C), and allowed to stand for 8 min before fluorescence observation. Observations were made at excitation wavelengths of 530–550 nm and emission wavelengths of 575 nm and above every 30 s for 20 min.

Observation of Intracellular ATP Production

Intracellular ATP production was visualized by a fluorescent ATP-indicator protein, GO-ATeam.

INS-1 cells cultured in FFA medium for 3 d were seeded in culture dishes at 1.0 × 10⁵ cells/cm² and transfected with GO-ATeam Cyto2 the next day using Lipofectamine 3000. All of the above procedures were performed in FFA medium. One day after transfection, cells were seeded at 2.5 × 10⁵/cm² onto 2.5 cm diameter cover glasses and cultured in FFA medium for an additional day (6 d total in FFA medium). Cells were treated with no FFA and low glucose (5 mM) medium for 17–22 h prior to observation. Cell density at observation was always about 80% confluent. Cells were washed twice with Glc(−) KRBH and placed on the microscope stage for 8 min before fluorescence observation. GO-ATeam was excited at 487 nm, and green fluorescence (510–550 nm, the ATP-unbound form) and orange fluorescence (longer than 560 nm, the ATP-bound form) were observed using W-VIEW GEMINI. Glucose (final concentration of 20 mM) was added 1 min after the start of observation. Intracellular ATP production was evaluated by the ratio of orange to green fluorescence of GO-ATeam.

Observation of Intracellular ROS

The concentrations of intracellular ROS were evaluated by CellROX Green fluorescence intensity. After 4 d of culture in FFA medium with medium change every 2 d, INS-1 cells were seeded at 1.5 × 10⁵ cells/cm² on a 2.5 cm diameter cover glass. The cells were cultured in FFA medium for another 2 d and additional one day after a medium change. Fluorescence observations were made 17–22 h after replacement with low glucose (5 mM) medium without FFA. Cell density at the time of observation was always about 80% confluent. Cells to be observed were washed twice with Glc(−) KRBH, then light-shielded and stained with CellROX Green (5 µM) for 30 min at 37 °C. After washing twice again with Glc(−) KRBH to remove excess dye, the cells were placed on the microscope stage (30 °C) in Glc(−) KRBH for 8 min before fluorescence observation. CellROX Green was excited at 460–495 nm and observed at 510–550 nm. Glucose (final concentration of 20 µM) was added 1 min after the start of observation, and fluorescence intensity was measured every minute until 30 min.

Measurement of Caspase-3 Activity

Caspase-3 activity was measured using the Caspase Glo 3/7 Assay System. INS-1 cells cultured in FFA medium every 2 d for 7 d with medium change were seeded in 96 well plate at 1.0 × 10⁴ cells/well, stimulated with H₂O₂ (final concentration of 200 µM), and lysed by adding Caspase Glo3/7 reagent within 10 min. After an hour incubation of the cell lysates, the amount of luminescence product of caspase 3 was quantified by GloMax Multi Detection System (Promega).

Observation of Apoptosis

Apoptosis induced in cells was evaluated by interaction with Annexin-V-FITC. INS-1 cells were cultured in FFA medium for 7 d, changing medium every 2 d, and then treated for 17–22 h without FFA. KRBH (11 mM Glc) was used for observation. Cells to be observed were washed twice with KRBH (11 mM Glc) and finally placed in KRBH (11 mM Glc) including Annexin-V-FITC. After incubation for 5 min at room temperature, the cells were stimulated with H₂O₂ (final concentration 100 µM) and fluorescence intensity of Annexin-V-FITC (excitation at 460–495 nm and emission at 510–550 nm) was measured every 5 min until 40 min.

Statistical Analysis

Data from several groups were analyzed by ANOVA followed by Tukey’s post hoc test. p > 0.05 was considered statistically significant.

RESULTS

Cells Cultured in FFA Medium

Figure 1 shows INS-1 cells cultured with each FFA medium for 6 d. No significant differences were observed in the phase contrast microscopic images of both OA and PA cells compared to control cells during medium exchange every 2 d, passaging, and other treatments. PA, a saturated fatty acid, is known to cause cell death. Although there was a tendency for more cells to float during medium exchange, there was no significant difference in the number of adherent cells compared to control cells (data not shown).

Fig. 1. Phase Contrast Microscopic Image of Cells in Medium Containing FFA

INS-1 cells cultured in medium supplemented with oleic acid (OA) or palmitic acid (PA) for 6 d were observed under a phase contrast microscope. Control cells were cultured in the absence of FA. A. Control cells, B. PA-treated cells, C. OA-treated cells.

Effects of OA on mRNA Expression of Proteins Related for ER Stress, Oxygen Stress and Apoptosis

Messenger RNA expression of the proteins related to ER stress, oxygen stress and apoptosis in INS-1 cells cultured in the medium with OA or PA were analyzed (Fig. 2). In this experiment, cells were cultured with the FFA for two different periods of time: 1 d (Fig. 2A.) or 7 d (Fig. 2B.).

Fig. 2. Effects of OA and PA on mRNA Expression Levels of ER Stress Marker Proteins and Superoxide Dismutase

The mRNA levels of endoplasmic reticulum stress and oxidative stress markers in INS-1 cells cultured in medium supplemented with oleic acid (OA) or palmitic acid (PA) for 1 d (A) or 7 d (B) with medium change every 2 d were analyzed by real-time PCR. Controls were cultured in the absence of FA. Ddit3, Hspa5, and Hsp90b1 were measured as ER stress-related proteins, and SOD1 and SOD2 as oxidative stress-related proteins, and Casp3 as an apoptosis marker. Means ± standard deviation (S.D.) of 3 independent experiments are shown. There is a statistically significant difference (p < 0.05) between the different letters.

In INS-1 cells cultured in medium containing OA (OA-treated cells) for 1 d, mRNA levels of ER-stress responsive proteins, Hspa5 and Hsp90b1, were significantly higher than control cells (Fig. 2A, p < 0.0001 for both proteins) and as high as in PA-treated cells. In contrast, cells cultured with OA for 7 d (Fig. 2B) showed no ER stress response, although PA-treated cells showed higher levels of the ER-stress proteins. This suggests that ER stress may occur when the high PA environment continues, but does not in the high OA environment.

The mRNA expression levels of SOD1 and SOD2, which are elevated in response to ROS levels, were significantly higher in both PA- (SOD1: p = 0.0001, SOD2: p = 0.0034) and OA-treated cells (both of SOD1, 2: p < 0.0001) after 1 d of treatment than in control cells. The increase in expression of SOD2 indicates an increase in intramitochondrial ROS, suggesting that OA may activate mitochondrial energy metabolism. On the other hand, after 7 d of treatment, mRNA expression was significantly decreased in both PA- (SOD1: p < 0.0001, SOD2: p = 0.0002) and OA-treated cells (SOD1: p < 0.0001, SOD2: p = 0.0201) compared to control cells, suggesting that there was some change in the oxidative stress response in the cells during the 7-d period.

The mRNA expression of caspase-3, a marker of apoptosis induction, was significantly higher in OA-treated cells compared to control cells in both 1-d (p < 0.0001) and 7-d (p < 0.0001) treatments. On the other hand, PA-treated cells were significantly higher in the 1-d treatment (p < 0.0001), but returned to the level of control cells in the 7-d treatment.

Effects of OA on Mitochondrial Energy Metabolism

The effect of OA and PA on mitochondrial energy metabolism was examined in terms of its effect on changes of mitochondrial membrane potential (Fig. 3) and intracellular ATP level (Fig. 4) following glucose addition to INS-1 cells cultured in low-glucose environment (5 mM glucose) for 17–22 h.

Fig. 3. Effects of OA and PA on Mitochondrial Membrane Potential after Glucose Stimulation

Mitochondrial membrane potential of INS-1 cells cultured in medium supplemented with oleic acid (OA) or palmitic acid (PA) for 6 d with medium change every 2 d and cultured in medium with no FFA and low glucose (5 mM) for 17–22 h was evaluated by fluorescence intensity of TMRE, which binds to mitochondrial membrane in a membrane potential dependent manner. Cells were stained with 25 nM TMRE in Glc(−) KRBH for 30 min at room temperature and excess day was washed away. Fluorescence observation was started in Glc(−) KRBH, glucose (final concentration of 20 µM) was added 1 min after start, and fluorescence intensity was measured every 30 s for 20 min. For a single cell, the cytoplasmic areas (excluding the nucleus) where TMRE staining was seen were selected as broadly as possible, and changes in the average fluorescence intensity in these areas were sampled. The relative fluorescence intensities were calculated using those at the beginning of the experiment as 1. Means ± standard error (S.E.) of independent cells (Control: n = 21, PA: n = 27, OA: n = 23) are shown in the graph.

Fig. 4. Effects of OA and PA on Intracellular ATP Concentration

Intracellular ATP concentrations were measured in INS-1 cells cultured in medium supplemented with oleic acid (OA) or palmitic acid (PA) for 6 d with medium change every 2 d and cultured in medium with no FFA and low glucose (5 mM) for 17–22 h. Intracellular ATP concentration was evaluated using GO-ATeam, a fluorescent protein that changes its emission color in response to ATP concentration. The FRET system revealed that GO-ATeam emits green fluorescence (510–550 nm) when not bound to ATP and orange fluorescence (>560 nm) when bound to ATP. Fluorescence observation was started in Glc(−) KRBH and glucose was added 1 min after start (final concentration of 20 µM). Ratiometric images (A) and a graph of the ratio of fluorescence intensity (orange/green) measured every minute up to 60 min (B) are shown. The rate of increase in orange/green intensity was calculated every 10 min and shown in graph (C) as mean ± S.E. (Control: n = 21, PA: n = 22, OA: n = 30).

Mitochondrial membrane potential was expressed as a relative value with the TMRE fluorescence intensity of each cell at the beginning of the observation as 1. Figure 3 shows the change in values after the addition of glucose to cells. In control cells, values increased within 1 min after glucose addition and then gradually decreased. In OA-treated cells, this decrease was slower and tended to remain higher than in control cells for 7 to 14 min (p = 0.4319 at 12.5 min after glucose addition). These results suggest that mitochondrial membrane potential may remain relatively high in OA-treated cells after glucose addition. In contrast, in PA-treated cells, the increase after glucose addition was significantly smaller than in control cells (p = 0.0156 at 2.5 min after glucose addition), and the subsequent decrease was faster than in control cells (the p-values at 3.5 and 4.0 min after glucose addition were 0.0118 and 0.0117, respectively). It was suggested that mitochondrial membrane potential was less likely to increase after glucose addition in PA-treated cells.

To evaluate the rate of ATP synthesis, which affects mitochondrial membrane potential, we examined the change in intracellular ATP concentration after the addition of glucose. Three days after transfecting GO-ATeam, an ATP-indicator protein, into FFA-treated cells, we observed changes in fluorescence intensity of ATP-bound (green) and ATP-unbound (orange) proteins in the cells after the addition of glucose. Representative merge images of green and orange fluorescence (0 and 40 min after glucose addition) are shown in Fig. 4A, and a graph of the change in fluorescence intensity ratio of orange/green fluorescence is shown in Fig. 4B. The increase in the orange/green ratio reflects an increase in intracellular ATP concentration. To compare the ATP synthesis rate after glucose addition, the rate of increase in the orange/green ratio, was calculated every 10 min (Fig. 4C). In control cells, the increase rate from 10 to 20 min was higher than that of 0–10 min (no significant difference, p = 0.7092), and the rate decreased significantly after 20 min. This trend was generally similar for PA-treated cells. In contrast, in OA-treated cells, the 0–10 min increase rate tended to be higher than the 10–20 min increase rate (not significant) and remained relatively high until 40 min after glucose addition. Comparing the increase rate of OA-treated and control cells at the same time points, the rate of OA-treated cells tended to be greater than that of control cells at 0–10 min (not significant, p = 0.2546) and 20–30 min (not significant, p = 0.1131) after glucose addition. In OA-treated cells, the reaction from the addition of glucose to the synthesis of ATP was faster, and the higher ATP synthesis rate was maintained for longer than in control cells. The rate of increase in PA-treated cells was similar to that of control cells, and was lower than that of control cells after 40 min.

Effects of OA on Intracellular ROS Levels after Glucose Stimulation

To measure the amount of intracellular ROS derived from the mitochondrial electron transport chain (ETC), changes in intracellular ROS concentration after glucose stimulation were evaluated by changes in CellROX fluorescence intensity (Fig. 5A).

Fig. 5. Effects of OA and PA on Change in Intracellular ROS Levels after Addition of Glucose

Intracellular ROS levels were measured in INS-1 cells cultured in medium supplemented with oleic acid (OA) or palmitic acid (PA) for 7 d with medium change every 2 d and cultured in medium with no FFA and low glucose (5 mM) for 17–22 h. Cells were stained with 5 µM CellROX in Glc(−) KRBH for 30 min at 37 °C, and excess dye was washed away. Fluorescence observation was started in Glc(−) KRBH, and glucose (20 µM final of concentration) was added 1 min after the start and fluorescence intensity was measured every minute until 30 min. Intracellular fluorescence intensity was evaluated in terms of nuclear fluorescence intensity. Changes in the fluorescence intensity are shown in graph (A) as mean ± S.E. Fluorescence images (B) and mean ± S.E. of fluorescence intensity with all data points (C) at 0 and 30 min after the start are shown (Control: n = 38, PA: n = 37, OA: n = 32). There is a statistically significant difference (p < 0.05) between the different letters for each time.

CellROX fluorescence intensity of OA-treated cells cultured in fatty acid-free low glucose medium for 17–22 h before observation was significantly higher than that of control cells even before glucose addition (Fig. 5C, p < 0.0001). After glucose addition, no increase in fluorescence intensity was observed in control cells, while OA-treated cells showed a temporary decreasing trend after glucose addition, followed by a gradual increase after 5 min, reaching approximately 1.5 times the level before glucose addition after 30 min (p < 0.0001). PA-treated cells showed the same trend as OA-treated cells, but the increase after glucose addition was smaller than that of OA-treated cells, resulting that the fluorescence intensity at 30 min was significantly lower than that of OA-treated cells (p = 0.0142).

These results indicate that OA treatment increases intracellular ROS levels. The cause of the increase in ROS after glucose addition is most likely due to an increase in the amount of ROS produced in the mitochondria and/or a decrease in the ability to degrade the ROS produced.

Effect of OA on Resistance to Oxidative Stress

The increase in intracellular ROS in OA-treated cells may affect the cells’ resistance to oxidative stress. The activity of caspase-3, a marker of induced apoptosis, was measured after addition of H₂O₂ using a luminescent substrate (Fig. 6A). A large increase in activity was observed in PA-treated cells compared to control cells (p < 0.0001), while no significant difference from was observed in OA-treated cells. The induction of apoptosis after addition of H₂O₂ was evaluated by binding of FITC-labeled Annexin-V to the cells (Figs. 6B, C). The fluorescence intensity of OA-treated cells was comparable to that of control cells before the addition of H₂O₂ (0 min), but gradually increased after the addition, and was significantly higher than that of control cells after 20 min (p = 0.0001). The increase rate in fluorescence intensity of OA-treated cells over 40 min (2.5-fold, p < 0.0001) was significantly higher than that of control cells (1.8-fold). OA-treated cells likely have decreased resistance to oxidative stress. In contrast, the fluorescence intensity of PA-treated cells was significantly higher even before the addition of H₂O₂ (0 min, p < 0.0001), and their rate of increase (1.3-fold) was not higher.

Fig. 6. Apoptosis Induction after Addition of Hydrogen Peroxide

Caspase-3 activity (A) and induction of apoptosis (B, C) were measured after addition of hydrogen peroxide. Panel A; INS-1 cells cultured in medium supplemented with oleic acid (OA) or palmitic acid (PA) for 7 d, with medium changed every 2 d, were incubated for 1 h with the addition of assay reagent, and hydrogen peroxide (final conc. of 200 µM) was added within 10 min. Caspase-3 activity was quantified by the amount of luminescent product. Panel B and C; Annexin-V-FITC was added to INS-1 cells cultured in medium supplemented with oleic acid (OA) or palmitic acid (PA) for 7 d, changing medium every 2 d, and cultured in medium without FFA for 17–22 h. One minute after the start of observation, hydrogen peroxide (final concentration of 100 µM) was added, and the binding of annexin-V-FITC to the plasma membrane was evaluated by fluorescence intensity every minute until 40 min. Fluorescence images (B) and mean ± S.E. of fluorescence intensity (C) with all data points (Control: n = 34, PA: n = 39, OA: n = 42) at 0 min and 40 min after the start are shown. There is a statistically significant difference (p < 0.05) between the different letters for each time.

DISCUSSION

The cytotoxicity of saturated fatty acids such as PA has been widely reported. On the other hand, although it has been reported that OA reduces the toxicity of SFA, especially ER stress and reduced nicotinamide adenine dinucleotide phosphate (NADPH) oxidase-mediated oxidative stress,²⁶⁾ there have been few reports showing the effects of OA itself on the insulin secretory function of pancreatic β-cells. In this study, cells were cultured in OA or PA medium for 6–7 d and the effects of OA on mitochondrial energy metabolism, oxidative stress, and oxidative stress tolerance were examined in comparison to PA. The concentration of FFA added to the medium was 200 µM, which is not an extremely high blood concentration, but the effect of sustained exposure was examined by treating the cells for 6–7 d. In the phase contrast microscopic images, there were no significant differences in cell morphology or proliferation compared to control cells during treatment with OA for 6 to 7 d. It was thought that PA, a saturated fatty acid, might have adverse effects such as cell death on cells during the 6–7 d exposure, but no extreme cell death was observed.

In the mechanism by which pancreatic β-cells sense elevated glucose concentrations and secrete insulin, mitochondrial energy metabolism is involved in the initial step of producing ATP from glucose. FFA are good substrates for mitochondrial energy metabolism, producing a large amount of ATP via β-oxidation, the citric acid circuit, and the electron transfer system. The rate at which ATP is produced by ETC could control the rate at which insulin is secreted. In this study, we examined the effects of OA on mitochondrial energy metabolism and on the cell as a whole.

The cells cultured in medium containing OA for 6–7 d showed a faster increase in intracellular ATP concentration after glucose addition and higher ATP production up to 40 min later. In the OA-treated cells, mitochondrial membrane potential remained higher than in control cells until about 15 min after glucose addition, presumably because the rate of H⁺ supply from the (TCA) cycle exceeds the rate of ATP synthesis. These results likely show that GSIS is enhanced in OA-treated cells because their metabolism, which produces ATP upon sensing increased glucose concentration, has been enhanced.

Increased mitochondrial energy metabolism can simultaneously increase the production of reactive oxygen species by the electron transfer system. mRNA expression was increased in cells treated overnight with OA, SOD1 and SOD2, presumably in response to increased ROS. In contrast, SOD mRNA expression decreased in cells treated with OA for 7 d, suggesting that the cells’ ability to degrade intracellular ROS may have decreased over the previous 7 d. In the OA 7-d treated cells, the amount of intracellular ROS was actually higher and increased after glucose addition. It seems likely that ROS generated from the electron transfer system was not degraded, as was the case in control cells, but gradually accumulated. This is probably due to decreased expression of SOD enzymes.

We examined whether the continuous high ROS concentrations in OA-treated cells induces apoptosis. Although mRNA expression of caspase-3, an enzyme that induces apoptosis, was significantly higher in OA-treated cells, enzyme activity was not increased, indicating that continuous OA treatment and the resulting sustained increase in intracellular ROS concentration do not induce apoptosis. Next, the time course of apoptosis induction by the addition of H₂O₂ was examined. The induction rate of apoptosis after H₂O₂ addition was faster in OA-treated cells than in control cells, indicating that the reaction time from oxidative stress to apoptosis induction is shorter in OA-treated cells. Continuous treatment with OA was thought to cause a continuous increase in intracellular ROS concentrations, facilitating the induction of apoptosis by external oxidative stress.

Unlike OA-treated cells, PA-treated cells maintained lower mitochondrial membrane potential and produced less ATP after glucose addition than control cells, indicating that PA does not increase mitochondrial energy metabolism. In PA-treated cells, intracellular ROS has been increased and apoptosis has been induced even before the addition of H₂O₂. It has been reported that PA causes ER stress and increases intracellular ROS via NADPH oxidase.^14,27) We speculate that the high intracellular ROS levels and apoptosis induction observed in PA-treated cells in this study before the addition of H₂O₂ are probably mediated by ER stress. In this study, we show for the first time that PA (an SFA) and OA (a MUFA) have different effects on mitochondrial energy metabolism.

It has been noted that fatty acids with different hydrocarbon chain lengths and different numbers of double bonds, have different effects on cells.^26,28) In this study, we examined the effects of OA, a MUFA, on the mitochondria of pancreatic β-cells, providing a glimpse of the complex effects of OA on cells. Various types of fatty acids are present in plasma. Assuming that those structures are recognized and trigger a variety of intracellular responses, free fatty acid receptors on the plasma membrane would be a candidate for the key to elucidating the differences.^29,30) Proteins such as FABPs, fatty acid-binding proteins,³¹⁾ which intracellularly transport FFAs, are also of interest and are promising candidates for future study.

Acknowledgments

This work was supported by the Institute of Human Culture Studies (IHCS) Otsuma Women’s University Grant-in-Aid for Graduate Students (A) or (B) Grant Numbers DB1909, DB2016, DA2104, DA2203, and DA2304.

Conflict of Interest

The authors declare no conflict of interest.

REFERENCES

• 1) Mizukami H, Kudoh K. Diversity of pathophysiology in type 2 diabetes shown by islet pathology. J. Diabetes Investig., 13, 6–13 (2022).
• 2) Møller JB, Pedersen M, Tanaka H, Ohsugi M, Overgaard VR, Lynge J, Almind K, Vasconcelos N-M, Poulsen P, Keller C, Ueki K, Ingwersen HS, Pedersen KB, Kadowaki T. Body composition is the main determinant for the difference in type 2 diabetes pathophysiology between Japanese and Caucasians. Diabetes Care, 37, 796–804 (2014).
• 3) Kaneko YK. Development of therapeutic agents with a novel mechanism of action targeting pancreatic β-cells for diabetes. Biol. Pharm. Bull., 46, 640–646 (2023).
• 4) DeFronzo RA. Pathogenesis of type 2 diabetes mellitus. Med. Clin. North Am., 88, 787–835, ix (2004).
• 5) Puzziferri N, Roshek TB 3rd, Mayo HG, Gallagher R, Belle SH, Livingston EH. Long-term follow-up after bariatric surgery: a systematic review. JAMA, 312, 934–942 (2014).
• 6) Reaven GM, Hollenbeck C, Jeng C-Y, Wu MS, Ida Chen Y-D. Measurement of plasma glucose, free fatty acid, lactate, and insulin for 24 h in Patients With NIDDM. Diabetes, 37, 1020–1024 (1988).
• 7) Unger RH. How obesity causes diabetes in Zucker diabetic fatty rats. Trends Endocrinol. Metab., 8, 276–282 (1997).
• 8) Maestre I, Jordan J, Calvo S, Reig JA, Cenã V, Soria B, Prentki M, Roche E. Mitochondrial dysfunction is involved in apoptosis induced by serum withdrawal and fatty acids in the beta-cell line Ins-1. Endocrinology, 144, 335–345 (2003).
• 9) Cnop M, Hannaert JC, Hoorens A, Eizirik DL, Pipeleers DG. Inverse relationship between cytotoxicity of free fatty acids in pancreatic islet cells and cellular triglyceride accumulation. Diabetes, 50, 1771–1777 (2001).
• 10) Shimabukuro M, Zhou YT, Levi M, Unger RH. Fatty acid-induced beta cell apoptosis: a link between obesity and diabetes. Proc. Natl. Acad. Sci. U.S.A., 95, 2498–2502 (1998).
• 11) Maedler K, Spinas GA, Dyntar D, Moritz W, Kaiser N, Donath MY. Distinct effects of saturated and monounsaturated fatty acids on β-cell turnover and function. Diabetes, 50, 69–76 (2001).
• 12) Dhayal S, Welters HJ, Morgan NG. Structural requirements for the cytoprotective actions of mono-unsaturated fatty acids in the pancreatic beta-cell line, brin-bd11. Br. J. Pharmacol., 153, 1718–1727 (2008).
• 13) Acosta-Montaño P, Rodríguez-Velázquez E, Ibarra-López E, Frayde-Gómez H, Mas-Oliva J, Delgado-Coello B, Rivero IA, Alatorre-Meda M, Aguilera J, Guevara-Olaya L, García-González V. Fatty acid and lipopolysaccharide effect on beta cells proteostasis and its impact on insulin secretion. Cells, 8, 884 (2019).
• 14) Karaskov E, Scott C, Zhang L, Teodoro T, Ravazzola M, Volchuk A. Chronic palmitate but not oleate exposure induces endoplasmic reticulum stress, which may contribute to INS-1 pancreatic beta-cell apoptosis. Endocrinology, 147, 3398–3407 (2006).
• 15) Laybutt DR, Preston AM, Åkerfeldt MC, Kench JG, Busch AK, Biankin AV, Biden TJ. Endoplasmic reticulum stress contributes to beta cell apoptosis in type 2 diabetes. Diabetologia, 50, 752–763 (2007).
• 16) Cunha DA, Hekerman P, Ladriere L, Bazarra-Castro A, Ortis F, Wakeham MC, Moore F, Rasschaert J, Cardozo AK, Bellomo E, Overbergh L, Mathieu C, Lupi R, Hai T, Herchuelz A, Marchetti P, Rutter GA, Eizirik DL, Cnop M. Initiation and execution of lipotoxic ER stress in pancreatic β-cells. J. Cell Sci., 121, 2308–2318 (2008).
• 17) Moffitt JH, Fielding BA, Evershed R, Berstan R, Currie JM, Clark A. Adverse physicochemical properties of tripalmitin in beta cells lead to morphological changes and lipotoxicity in vitro. Diabetologia, 48, 1819–1829 (2005).
• 18) Sommerweiss D, Gorski T, Richter S, Garten A, Kiess W. Oleate rescues INS-1E β-cells from palmitate-induced apoptosis by preventing activation of the unfolded protein response. Biochem. Biophys. Res. Commun., 441, 770–776 (2013).
• 19) Maedler K, Oberholzer J, Bucher P, Spinas GA, Donath MY. Monounsaturated fatty acids prevent the deleterious effects of palmitate and high glucose on human pancreatic β-cell turnover and function. Diabetes, 52, 726–733 (2003).
• 20) Fujiwara K, Maekawa F, Yada T. Oleic acid interacts with GPR40 to induce Ca²⁺ signaling in rat islet β-cells: mediation by PLC and L-type Ca²⁺ channel and link to insulin release. Am. J. Physiol. Endocrinol. Metab., 289, E670–E677 (2005).
• 21) Santos LRB, Rebelato E, Graciano MFR, Abdulkader F, Curi R, Carpinelli AR. Oleic acid modulates metabolic substrate channeling during glucose-stimulated insulin secretion via NAD(P)H Oxidase. Endocrinology, 152, 3614–3621 (2011).
• 22) Gurgul E, Lortz S, Tiedge M, Jörns A, Lenzen S. Mitochondrial catalase overexpression protects insulin-producing cells against toxicity of reactive oxygen species and proinflammatory cytokines. Diabetes, 53, 2271–2280 (2004).
• 23) Eitel K, Staiger H, Rieger J, Mischak H, Brandhorst H, Brendel MD, Bretzel RG, Haring HU, Kellerer M. Protein kinase C δ activation and translocation to the nucleus are required for fatty acid–induced apoptosis of insulin-secreting cells. Diabetes, 52, 991–997 (2003).
• 24) Palomer X, Pizarro-Delgado J, Barroso E, Vázquez-Carrera M. Palmitic and oleic acid: the yin and yang of fatty acids in type 2 diabetes mellitus. Trends Endocrinol. Metab., 29, 178–190 (2018).
• 25) Wrede CE, Dickson LM, Lingohr MK, Briaud I, Rhodes CJ. Protein kinase B/akt prevents fatty acid-induced apoptosis in pancreatic β-cells (INS-1). J. Biol. Chem., 277, 49676–49684 (2002).
• 26) Lytrivi M, Castell A-L, Poitout V, Cnop M. Recent insights into mechanisms of b-cell lipo- and glucolipotoxicity in type 2 diabetes. J. Mol. Biol., 432, 1514–1534 (2020).
• 27) Inoguchi T, Li P, Umeda F, Yu HY, Kakimoto M, Imamura M, Aoki T, Etoh T, Hashimoto T, Naruse M, Sano H, Utsumi H, Nawata H. High glucose level and free fatty acid stimulate reactive oxygen species production through protein kinase C–dependent activation of NAD(P)H oxidase in cultured vascular cells. Diabetes, 49, 1939–1945 (2000).
• 28) Kiyoki Y, Kato T, Kito S, Matsuzaka T, Morioka S, Sasaki J, Makishima K, Sakamoto T, Nishikii H, Obara N, Sakata-Yanagimoto M, Sasaki T, Shimano H, Chiba S. The fatty acid elongase Elovl6 is crucial for hematopoietic stem cell engraftment and leukemia propagation. Leukemia, 37, 910–913 (2023).
• 29) Kimura I, Ichimura A, Ohue-Kitano R, Igarashi M. Free fatty acid receptors in health and disease. Physiol. Rev., 100, 171–210 (2020).
• 30) Alquier T, Peyot M-L, Latour MG, Kebede M, Sorensen CM, Gesta S, Kahn CR, Smith RD, Jetton TL, Metz TO, Prentki M, Poitout V. Deletion of GPR40 impairs glucose-induced insulin secretion in vivo in mice without affecting intracellular fuel metabolism in islets. Diabetes, 58, 2607–2615 (2009).
• 31) Prentice KJ, Saksi J, Robertson LT, Lee GY, Inouye KE, Eguchi K, Lee A, Cakici O, Otterbeck E, Cedillo P, Achenbach P, Ziegler A-G, Calay ES, Engin F, Hotamisligil GS. A hormone complex of FABP4 and nucleoside kinases regulates islet function. Nature, 600, 720–726 (2021).