**E**ff**ect of** *Quamoclit angulata* **Extract Supplementation on Oxidative Stress and Inflammation on Hyperglycemia-Induced Renal Damage in Type 2 Diabetic Mice**

#### **Ji Eun Park 1, Heaji Lee 1, Hyunkyung Rho 1, Seong Min Hong 2, Sun Yeou Kim <sup>2</sup> and Yunsook Lim 1,\***


Received: 19 May 2020; Accepted: 23 May 2020; Published: 27 May 2020

**Abstract:** Type 2 diabetes mellitus (T2DM) is caused by abnormalities of controlling blood glucose and insulin homeostasis. Especially, hyperglycemia causes hyper-inflammation through activation of NLRP3 inflammasome, which can lead to cell apoptosis, hypertrophy, and fibrosis. *Quamoclit angulata* (QA), one of the annual winders, has been shown ameliorative effects on diabetes. The current study investigated whether the QA extract (QAE) attenuated hyperglycemia-induced renal inflammation related to NLRP inflammasome and oxidative stress in high fat diet (HFD)-induced diabetic mice. After T2DM was induced, the mice were treated with QAE (5 or 10 mg/kg/day) by gavage for 12 weeks. The QAE supplementation reduced homeostasis model assessment insulin resistance (HOMA-IR), kidney malfunction, and glomerular hypertrophy in T2DM. Moreover, the QAE treatment significantly attenuated renal NLRP3 inflammasome dependent hyper-inflammation and consequential renal damage caused by oxidative stress, apoptosis, and fibrosis in T2DM. Furthermore, QAE normalized aberrant energy metabolism (downregulation of p-AMPK, sirtuin (SIRT)-1, and PPARγ-coactivator α (PGC-1 α)) in T2DM mice. Taken together, the results suggested that QAE as a natural product has ameliorative effects on renal damage by regulation of oxidative stress and inflammation in T2DM.

**Keywords:** *Quamoclit angulata*; type 2 diabetes; kidney damage; inflammation; oxidative stress; apoptosis; fibrosis

#### **1. Introduction**

Diabetes mellitus (DM) is considered as a metabolic disease that results in impaired glucose and insulin homeostasis [1]. Especially, insulin resistance caused by hyperglycemia, is the worldwide epidemic that is accompanied by various complications in type 2 DM (T2DM) [2]. The early stage of nephropathy (DN) is characterized by the structural changes of kidney such as damage of the glomerular basement membrane (GBM), enlargement of the mesangial cells, glomerulosclerosis, and fibrosis and renal function failure including microalbuminuria and reduced glomerular filtration rate (GFR) [3,4].

The main cause of renal damage is the hyperglycemic condition in T2DM. Hyperglycemia leads to overproduction of reactive oxygen species (ROS), which potentially causes oxidative stress and activates various cytokines, chemokines, and growth factors. Oxidative stress results from an imbalance between oxidants and antioxidants such as NAD(P)H quinone dehydrogenase-1 (NQO1), hemeoxygenase-1 (HO-1), catalase, superoxide dismutase (SOD), and glutathione peroxidase (GPx) [5–7]. The pathogenic

changes caused by hyperglycemia-induced oxidative stress modify normal cell signaling and induce hyper-inflammation, apoptosis [8]. Moreover, formation of advanced glycation end products (AGEs) and activation of the AGE receptor due to hyperglycemia directly promote inflammatory states in the kidney by increasing oxidative stress [9].

Furthermore, oxidative stress activates the nucleotide-binding oligomerization domain (NOD)-like pyrin domain containing receptor 3 (NLRP3) inflammasome [10,11]. Activated NLRP3 recruits the apoptosis-associated speck-like protein containing a caspase recruitment domain (ASC) and pro-caspase-1, and contributes to the maturation of interleukin-1β (IL-1β) by activating caspase-1. NLRP3 inflammasome and mature IL-1β activate a transcription factor, nuclear factor-κB (NF-κB), and enhance multiple proinflammatory factors such as tumor necrosis factor-α (TNF-α), interleukin-6 (IL-6), and inducible nitric oxide synthase (iNOS) [12]. Hence, the activation of NLRP3 inflammasome contributes to chronic inflammatory response as well as insulin resistance in diabetes [13,14].

Moreover, chronic hyperglycemia-induced oxidative stress and hyper-inflammation accelerate renal apoptosis [5,13,15]. Cellular apoptosis is regulated by caspases through an extrinsic and intrinsic pathway. In ongoing-diabetes, hyperglycemia induces ROS-related apoptosis by increasing the Bax/Bcl-2 ratio, which is associated with progressive activation of pro-apoptotic caspase-3 [15,16]. ROS also increases protein kinase C (PKC), which activates transcription of transforming growth factor-β (TGF-β) [17]. In general, TGF-β leads to proliferation of fibroblasts and activates α-smooth muscle actin (α-SMA), which accommodates collagen formation and cellular hypertrophy. TGF-β also causes the proliferation of mesangial cells, consequentially leading to kidney fibrosis [18]. Therefore, suppression of renal cell apoptosis and pro-fibrotic change along with NLRP3 inflammasome related hyper-inflammation would be an effective target strategy on alleviating renal damage and the progress to DN [13].

Furthermore, abnormal energy metabolism in the kidney can cause renal damage. Sirtuin1 (SIRT1) mediates inflammatory signaling and apoptosis as a molecular response to glucotoxicity via deacetylation and inhibition of transcription factor NF-κB [19]. When the AMP/ATP ratio increases, 5 adenosine monophosphate-activated protein kinase (AMPK) alleviates renal inflammation causing aberrant energy accumulation. The SIRT1/AMPK signaling pathway along with the peroxisome proliferator-activated receptor γ-coactivator α (PGC-1α) also suppresses renal hyper-inflammation and oxidative stress [20]. Hence, amelioration of SIRT1/AMPK signaling would be a possible therapeutic approach for diabetic renal damage.

In recent years, many medicinal plants have been reported as antidiabetic natural products including banaba, fenugreek, gymnema, yerba mate, etc. Among these plants, *Quamoclit angulata* (QA) is emerging as a source of therapeutic substance for diabetes and its complications. Although QA has not been fully investigated, a previously reported patent has shown that herbal agents of the *Quamoclit angulata* extract (QAE) decreased the fasting blood glucose (FBG) level and hemoglobin A1c (HbA1c) production by stimulating insulin secretion in pancreatic β cell. QAE also attenuated albuminuria, which is a major factor of DN in diabetic mice. Furthermore, QAE ameliorated angiogenesis by reducing the mRNA level of vascular endothelial growth factor (VEGF) in the ARPE 19 cell [21]. Nevertheless, little research has investigated the effects of QAE supplementation on renal damage in a hyperglycemic condition by molecular mechanisms. Hence, we examined if QAE supplementation has protective roles in renal damage via modulation of oxidative stress and inflammation in high fat diet-induced diabetic mice.

#### **2. Materials and Methods**

#### *2.1. Quamoclit Angulata Extracts (QAE)*

QA was obtained at Jeju, Korea. Aerial parts without the seed of QA (50 g) were extracted with 400 mL of water by incubation at 50 ◦C for 1 h. The sticky solid extract (50 g) was suspended in water, added to 1 kg of activated charcoal at room temperature for 1.5 h. After incubation, water, 20% ethanol

fraction, and charcoal were removed through centrifugation and filtration (0.45 μm). Fractions were mixed, concentrated in vacuo, and frozen to dry. The yields of hot water extract and activated charcoal fractions were 25% and 11%, respectively.

#### *2.2. Identification of Candidate Compounds of QAE*

The standardization of QA was analyzed by using the HPLC system (Waters Corp., Milford, MA, USA) consisting of a separation module (e2695) and a photodiode array (PDA) detector. Twenty milligrams of dried QA were dissolved in 50% methanol/water. Protocatechuic acid, chlorogenic acid, syringic acid, myricetin, and quercetin were used as standard compounds and dissolved in methanol. For the analysis of each compound or sample, a Kromasil C<sup>18</sup> column (150 <sup>×</sup> 4.6 mm, 5 <sup>μ</sup>m) was used and a column temperature was set at 30 ◦C. The mobile phase consists of 3% acetic acid/water (solvent A) and methanol (solvent B) using a gradient program of 0–10% (B) in 0–10 min, 10–70% (B) in 10–44 min, 70–100% (B) in 44–50 min. The calibration was linear in a range of 0.1–1000 μg/mL for these five compounds. The flow rate was 0.9–1.0 mL/min and the PDA detector was set at 280 nm for acquiring chromatograms.

#### *2.3. Animals Experiments*

Male C57BL/6 mice at five weeks were housed in two or three per cages and maintained in a constant environment (temperature (22 ± 1 ◦C), humidity (50 ± 5%), and 12 h light/12 h dark cycle). After seven days of adaptation, the mice were randomly allocated into two groups. The first group was a non-diabetic control group (NC), which was fed an AIN-93G diet (10% kcal fat, Research Diets, New Brunswick, NJ, USA). The second was a diabetic group (DM), which was fed a high fat diet (40% kcal fat, Research Diets, New Brunswick, NJ, USA) for four weeks.

Then, the diabetic group received an intraperitoneal administration of 30 mg/kg body weight (BW) of streptozotocin (Sigma-Aldrich, St. Louis, MO, USA) in a citric acid buffer (pH 4.4). The NC mice received an equivalent amount of solvent. After five weeks from the last injection, fasting blood glucose (FBG) levels were measured once per week during the whole period of the animal experiment. Mice with FBG >140.4 mg/dL (7.8 mmol/L) more than two times were considered as the diabetic condition. The diabetes induction protocol was referred to the previous study by Zhang et al. [22].

Mice were separated in four groups; (1) CON: Non-diabetic normal mice were gavaged with distilled water, (2) DMC: Diabetic mice were gavaged with distilled water, (3) LQ: Diabetic mice were gavaged with a low dosage of QAE (5 mg/kg/day), (4) HQ: Diabetic mice were gavaged with a high dosage of QAE (10 mg/kg/day). QAE was dissolved in distilled water. Body weight, food intake, and fasting blood glucose level were weekly monitored during the animal experiment.

The animals were sacrificed after 12 weeks of oral supplementation. Blood was collected in a heparin (Sigma-Aldrich, St. Louis, MO, USA) coated syringe from the heart, centrifuged at 850 g at 4 ◦C for 10 min to obtain plasma. The kidney was removed from mice and stored at −80 ◦C before the experiment. All experiments with mice were approved by the Institutional Animal Care and Use Committee of Kyung Hee University (KHUASP(SE)-16-005 on 14 June, 2019).

#### *2.4. Hemoglobin A1c (HbA1c) and Plasma Insulin Assay*

HbA1c levels were measured using enzyme-linked immunosorbent assay (ELISA) commercial kits (Crystal Chem., Downers Grove, Elk Grove Village, IL, USA) according to directions of the manufacturer within two weeks from the sample collection.

The plasma insulin level was measured using ELISA kits (RayBiotech, Inc., Norcross, GA, USA). The homeostasis model assessment of insulin resistance (HOMA-IR) values were calculated as follows:

HOMA-IR = fasting insulin (mmol/L) × fasting glucose (μU/mL)/22.5

#### *2.5. Oral Glucose Tolerance Test (OGTT)*

Fasted mice were administrated for 16 h with a 50% glucose solution (2 g/kg). The blood glucose level was detected at 0, 15, 30, 60, 90, and 120 min using a glucometer (OneTouch, LifeScan Inc., Malvern, PA, USA). The area under the curve (AUC) values of OGTT are calculated according to the trapezoidal rule as follows:

> AUC = (((blood glucose) <sup>i</sup> + (blood glucose) <sup>i</sup>−1) × ((time) <sup>i</sup> - (time) <sup>i</sup>−1)/2) (i = time sequence)

#### *2.6. Renal Function Test*

Urine samples were collected during three phases of the experiment (0–4 weeks; initial, 4–8 weeks; mid, and 8–12; late-points). Urinary albumin excretion was determined by the albumin assay kit (Bioassay, Hayward, CA, USA). The concentrations of urinary and plasma creatinine were calculated from interpolating the results of optical density at 515 nm into a standard curve. Concentrations of BUN were measured in accordance with the manufacturer's instructions using a commercial kit (Asan pharmaceutical, Seoul, South Korea).

#### *2.7. Histological Observation of Kidney*

Kidney tissues were fixed in 10% formaldehyde and then dehydrated through a series of alcohol. The tissues were cleared in xylene and embedded in paraffin. The sections were cut with a microtome into 5 μm, and stained with hematoxylin and eosin (H&E). Kidney morphology in stained tissue was observed using an optical microscope (Nikon ECLIPSE Ci, Nikon Instrument, Tokyo, Japan).

To calculate the glomerular area in H&E-staining, paraffin-embedded sections were measured by the Canvas 11 software (Deneba, Miami, FL, USA). The Glomerulus area was expressed as the mean of thirty glomeruli per each sample and a minimum of four samples from each group were examined. Area values are reported in <sup>μ</sup>m2 <sup>×</sup> <sup>10</sup><sup>−</sup>3.

#### *2.8. Protein Extraction and Western Blot Analysis*

The kidneys were ground and lysed on ice for 30 min. The lysate was centrifuged to remove tissue debris at 1945× *g* at 4 ◦C for 10 min. Each supernatant was centrifuged again at 9078× *g* at 4 ◦C for 30 min. Then, the final supernatant was collected for cytosolic extract. The pellet was re-crushed in a hypertonic lysis buffer for 1 h, and then the lysate was centrifuged at 9078× *g* at 4 ◦C for 20 min and the supernatant was used for nuclear extract. The protein concentration was quantified according to a BCA protein assay (ThermoFisher Scientific, Grand Island, NY, USA).

Thirty μg of each protein sample were loaded into an SDS-PAGE and transferred to poly-vinylidine fluoride (PVDF) membranes (Millipore, Marlborough, MA, USA). We used 8~12% SDS-PAGE gel according to the molecular weight (MW) of target protein(s). After the transfer, the membrane was blocked in 1~3% bovine serum albumin (BSA) in a phosphate buffed saline −0.1% Tween 20 (PBS-T), the membrane was incubated at 4 ◦C with each primary antibody. To detect primary antibodies, respective horseradish peroxide (HRP)-conjugated secondary antibodies were given to membranes. Protein bands were visualized using a chemiluminescent detector (Syngene, Cambridge, UK). Levels of targeted proteins were calculated using Syngene GeneSnap (Syngene, Cambridge, UK).

#### *2.9. Statistical Analysis*

Data were expressed as mean ± standard error of the mean (SEM). The significant differences between sample groups were determined using one-way ANOVA (significant level = 0.05).

#### **3. Results**

#### *3.1. Identification of Major Natural Compounds in QAE*

In this study, five compounds such as protocatechuic acid, chlorogenic acid, syringic acid, myricetin, and quercetin were analyzed by using the HPLC system. As shown in Table 1, the QA extract mainly involved protocatechuic acid (198.86 ± 2.26 μg/g). Meanwhile, chlorogenic acid (54.11 ± 1.81 μg/g), syringic acid (56.38 ± 0.57 μg/g), myricetin (12.42 ± 0.09 μg/g), and quercetin (13.41 ± 0.08 μg/g) were detected in QAE.

**Table 1.** Contents of protocatechuic acid, chlorogenic acid, syringic acid, myricetin, and quercetin compounds by HPLC analysis in the presence of *Quamoclit angulata* extract (QAE).


#### *3.2. E*ff*ect of QAE Supplementation on Body Weight, Food Intake, and Kidney Weight in T2DM Mice*

Body weight and food intake of all T2DM groups (the DMC group, the LQ group, and the HQ group) were significantly increased compared to the NC group. QAE supplementation for 12 weeks had no effect on body weight change in T2DM mice. With a slightly different result, kidney weight in the DMC group was significantly increased compared to the NC group, and there was no significant difference in the LQ group and the HQ group compared to the DMC group (Table 2).

**Table 2.** Effect of QAE on body weight, kidney weight, and food intake in type 2 diabetes mellitus (T2DM) mice.


Data were expressed as means ± SEM. Mean values with the same superscript letter (a,b) are not significantly different (*p* < 0.05). NC: Normal mice; DMC: Type 2 diabetic mice; LQ: Type 2 diabetic mice supplemented with a low dose (5 mg/kg/day) of QAE; HQ: Type 2 diabetic mice supplemented with a high dose (10 mg/kg/day) of QAE.

*3.3. E*ff*ect of QAE Supplementation on Fasting Blood Glucose and Plasma Insulin Levels, Homeostasis Model Assessment of Insulin Resistance (HOMA-IR), and Hemoglobin A1c (HbA1c) in T2DM Mice*

Fasting blood glucose level, plasma insulin level, HOMA-IR, and HbA1c level were as follows (Table 3). At the end of the QAE treatment period, there was no difference in the fasting blood glucose level among all groups. The plasma insulin level in the HQ group was significantly lower than that in the DMC group. HOMA-IR was significantly reduced in both QAE treated diabetic groups. The HbA1c level was significantly decreased only in the HQ group compared with the DMC group (Table 3).


**Table 3.** Effect of QAE on plasma indices related to type 2 diabetes.

Data were expressed as means ± SEM. Mean values with the same superscript letter (a,b) are not significantly different (*p* < 0.05). NC: Normal mice; DMC: Type 2 diabetic mice; LQ: Type 2 diabetic mice supplemented with a low dose (5 mg/kg/day) of QAE; HQ: Type 2 diabetic mice supplemented with a high dose (10 mg/kg/day) of QAE.

#### *3.4. E*ff*ect of QAE Supplementation on Glucose Homeostasis in T2DM Mice*

OGTT was performed to estimate insulin resistance and failure of glucose metabolism (Figure 1A), and glucose AUC was calculated as shown in Figure 1B. The figure showed that the DMC group had a high blood glucose level during 120 min after glucose administration compared to the NC group and there was no significant difference in the blood glucose level at 90 min after glucose administration among the DMC group and the QAE treatment groups. The blood glucose level of the LQ group at 120 min was remarkably reduced as compared to the DMC group and glucose AUC of the LQ group was significantly lower than that of the DMC group. On the other hand, the protein level of receptor for advanced glycation end products (RAGE) was remarkably increased in the DMC group compared to that of the NC group (Figure 1C). The HQ group showed a significant reduction of RAGE expression in comparison to the DMC group.

**Figure 1.** Effect of QAE on (**A**) glucose tolerance, (**B**) glucose area under the curve (AUC), and (**C**) renal receptor of AGE (RAGE) in T2DM mice. Data were expressed as means ± SEM. Mean values with the same superscript letter are not significantly different (*p* < 0.05). NC: Normal mice; DMC: Type 2 diabetic mice; LQ: Type 2 diabetic mice supplemented with a low dose (5 mg/kg/day) of QAE; HQ: Type 2 diabetic mice supplemented with a high dose (10 mg/kg/day) of QAE.

#### *3.5. E*ff*ect of QAE Supplementation on Kidney Function in T2DM Mice*

The ACRs of all T2DM groups were significantly higher than that in the NC group during the entire experimental period (Figure 2A). The ACRs of the QAE treatment groups decreased during the treatment period and showed a significant difference at the late stage of treatment in the diabetic mice. As shown in Figure 2B, the supplementation with a high dose of QAE significantly decreased the plasma creatinine and BUN compared with the DMC group. In representative H&E staining of the kidney (Figure 2C), the DMC group showed glomerular hypertrophy as compared to the NC group, while both the QAE treated groups ameliorated glomerular hypertrophy. The red arrow indicated mesangial expansion in the DMC group compared to the NC group. In the NC group, Bowman's space was observed as a thin white line. However, Bowman's space was broadened in the DMC group compared to that in the NC group, and was narrower in the QAE treatment groups than that in the DMC group. In addition, glomerular surface areas in histological sections of renal cortex were quantified to measure the degree of glomerular hypertrophy (Figure 2C). The glomerulus of the DMC group was significantly expanded compared with that of the NC group, while both QAE supplementation groups regardless of dose showed significantly reduced glomerular hypertrophy.

**Figure 2.** Effect of QAE on (**A**) urine albumin/creatinine ratio (ACR), (**B**) level of plasma creatinine and blood urea nitrogen (BUN), and (**C**) renal morphology and glomerular size in T2DM. Data were expressed as means ± SEM. Mean values with the same superscript letter (a,b and c) are not significantly different (*p* < 0.05). NC: Normal mice; DMC: Type 2 diabetic mice; LQ: Type 2 diabetic mice supplemented with a low dose (5 mg/kg/day) of QAE; HQ: Type 2 diabetic mice supplemented with a high dose (10 mg/kg/day) of QAE.

#### *3.6. E*ff*ect of QAE Supplementation on Oxidative Stress in T2DM Mice*

The renal 4-hydroxynonenal (4-HNE) level was examined for assessing lipid peroxidation and the level of renal protein carbonyls was used as a marker of protein oxidation caused by oxidative stress (Figure 3A). In the kidney, the 4-HNE protein level in the DMC group was significantly higher than that in the NC group. The HQ group presented a significant reduction of 4-HNE level compared to that of the DMC group. Renal levels of protein carbonyls in both QAE groups were significantly lower than that in the DMC group. In the DMC group, the protein levels of nuclear Nrf2 and its related markers such as HO-1, NQO1, catalase, MnSOD, and GPx were remarkably higher than those in the NC group. However, both QAE treatments significantly reduced the protein levels of nuclear Nrf2 and MnSOD. Moreover, a high dose of the QAE treatment significantly decreased the protein levels of HO-1, NQO1, and catalase in the diabetic mice. The protein levels of GPx and NOX4 were not significantly different among the DMC group and the QAE treatment groups (Figure 3B).

**Figure 3.** Effect of QAE on renal (**A**) 4-hydroxynonenal (4-HNE) and protein carbonyls and (**B**) antioxidant defense system in T2DM mice. Data were expressed as means ± SEM. Mean values with the same superscript letter (a,b) are not significantly different (*p* < 0.05). NC: Normal mice; DMC: Type 2 diabetic mice; LQ: Type 2 diabetic mice supplemented with a low dose (5 mg/kg/day) of QAE; HQ: Type 2 diabetic mice supplemented with a high dose (10 mg/kg/day) of QAE.

#### *3.7. E*ff*ect of QAE Supplementation on Inflammation in T2DM Mice*

The protein level of NLRP3 inflammasome was elevated in the DMC group compared to that of the NC group. However, the level of NLRP3 was significantly decreased in the QAE treatment groups compared to those in the DMC group. However, only a high dose of QAE treatment significantly lowered the protein levels of ASC, procaspase-1, caspase-1, and mature IL-1β in the diabetic mice. The protein levels of precursor IL-1β were not normalized in the QAE treatment groups compared to that in the DMC group (Figure 4A).

Furthermore, the DMC group demonstrated higher levels of inflammation related protein including monocyte chemoattractant protein (MCP)-1, CRP, nuclear NF-κB, TNF-α, IL-6, and iNOS than the NC group (Figure 4B). However, a high dose of QAE treatment in the diabetic mice reversed the protein levels of MCP-1 and nuclear NF-κB to the levels of the NC mice. In addition, the QAE treatment regardless of dose suppressed other inflammatory markers such as CRP, TNF-α, IL-6, and iNOS in the diabetic mice.

**Figure 4.** Effect of QAE on renal NLRP3 Inflammasome related hyper-inflammation in T2DM mice. Protein levels of (**A**) nucleotide-binding oligomerization domain-like pyrin domain containing receptor 3 (NLRP3) inflammasome: nucleotide-binding oligomerization domain-like pyrin domain containing receptor 3 (NLRP-3); apoptosis-associated speck-like proteins including caspase recruitment domain (ASC), caspase-1, and interleukin (IL)-1β; and (**B**) markers of pro-inflammatory response: monocyte chemoattractant protein-1 (MCP-1), C-reactive protein (CRP); and nuclear factor kappa B (NF-κB)-related inflammatory response: nuclear factor kappa B (NF-κB), tumor necrosis factor-α (TNF-α), interleukin (IL)-6, and inducible nitric oxide synthase (iNOS). Data were expressed as means ± SEM. Mean values with the same superscript letter (a,b and c) are not significantly different (*p* < 0.05). NC: Normal mice; DMC: Type 2 diabetic mice; LQ: Type 2 diabetic mice supplemented with a low dose (5 mg/kg/day) of QAE; HQ: Type 2 diabetic mice supplemented with a high dose (10 mg/kg/day) of QAE.

#### *3.8. E*ff*ect of QAE Supplementation on Energy Metabolism in T2DM Mice*

The protein levels of AMPK were not significantly different among the groups. The protein level of phosphorylated AMPK in the DMC group was significantly declined compared to that of the NC group, but those in the QAE treatment groups were increased compared to the DMC group. In addition, the QAE treatment elevated the pAMPK/AMPK ratio as much as the level of the NC group (Figure 5A). Furthermore, the protein levels of SIRT1 and PGC-1α were significantly declined in the DMC group compared to those in the NC group, but were increased in the QAE treatment groups regardless of dosage (Figure 5B).

**Figure 5.** Effect of QAE on renal AMPK/SIRT1 signaling in T2DM mice. (**A**) Phosphorylation of AMPK and (**B**) SIRT-1 and PGC-1α. Data were expressed as means ± SEM. Mean values with the same superscript letter (a,b) are not significantly different (*p* < 0.05). NC: Normal mice; DMC: Type 2 diabetic mice; LQ: Type 2 diabetic mice supplemented with a low dose (5 mg/kg/day) of QAE; HQ: Type 2 diabetic mice supplemented with a high dose (10 mg/kg/day) of QAE.

#### *3.9. E*ff*ect of QAE Supplementation on Apoptosis and Fibrosis in T2DM Mice*

The protein levels of caspase-8, caspase-3, and nuclear p53 in the DMC group were significantly higher than those in the NC group, but the high dose of QAE supplementation decreased the protein levels of caspase-8 and p53 than those in the DMC group. Furthermore, the QAE treatment regardless of dose reduced caspase-3 compared to the DMC group (Figure 6A). The protein levels of Bax in the QAE treatment groups were significantly decreased in comparison to that of the DMC group. The QAE treatment regardless of dose remarkably lowered the protein level of Bax/Bcl-2 ratio in the diabetic mice (Figure 6A). In addition, the QAE treatments reduced the protein level of ERK compared to that of the DMC group. At the same time, the protein levels of phosphorylated ERK in the QAE treatment groups were reduced compared to that in the DMC group. Moreover, the pERK/ERK ratio, an index of ERK phosphorylation, in the QAE treatment groups was also decreased compared to that in the DMC group (Figure 6A).

To examine the effect of the QAE supplementation on renal fibrosis, the protein levels of PKC-βII, TGF-β, α-SMA, and COL1A were measured (Figure 6B). The renal protein levels of PKC, TGF-β, α-SMA, and COL1A in the DMC group were significantly higher than those in the NC group. However, the QAE treatments decreased the protein levels of PKC, TGF-β, and α-SMA in comparison to the DMC group, and, in particular, a high dose of the QAE treatment declined the protein level of COL1A in the diabetic mice.

**Figure 6.** Effect of QAE on renal (**A**) apoptosis and (**B**) fibrosis in T2DM mice. Data were expressed as means ± SEM. Mean values with the same superscript letter (a,b and c) are not significantly different (*p* < 0.05). NC: Normal mice; DMC: Type 2 diabetic mice; LQ: Type 2 diabetic mice supplemented with a low dose (5 mg/kg/day) of QAE; HQ: Type 2 diabetic mice supplemented with a high dose (10 mg/kg/day) of QAE.

#### **4. Discussion**

Various studies noticed that many medicinal plants and natural products have potential biological activities. Among these plants, QA is a species of ipomoea morning glory and cultivated as an ornamental plant throughout the tropics. In this study, we aimed to investigate that dietary QAE supplementation could have beneficial effects on NLRP3 inflammasome dependent hyper-inflammation and consequential renal damage by stimulation of AMPK-SIRT1 signaling in type 2 diabetes.

The current study suggested a hypoglycemic effect of QAE presented by decreased plasma insulin, HOMA-IR, and HbA1c. HbA1c is considered as an index of average blood glucose control level, because the HbA1c level tends to increase with the averaged blood glucose levels over preceding three months. A previous study also showed a strong correlation between HbA1c and 6-h fasting glucose levels than overnight FBG levels in diabetic mice [23]. In this study, the QA supplementation ameliorated plasma insulin level, HOMA-IR, and HbA1c compared to the DMC group, although

did not significantly decrease the FBG level. In glucose tolerance test, AUC was declined in the LQ group compared to the DMC group. As shown in Table 1, QA contained five compounds such as protocatechuic acid (PCA), chlorogenic acid, syringic acid, myricetin, and quercetin. A recent study showed similar tendency that PCA significantly reduced blood glucose and plasma insulin level in the hyperglycemic condition [24]. In addition, it is known that activated ligation of AGEs to renal RAGE activated production of ROS subsequently causing oxidative stress [25]. Furthermore, chlorogenic acid (CGA) and quercetin have been shown to decrease blood glucose level by stimulating glucose uptake through the activation of AMPK in diabetic mice [26,27]. CGA has been also reported as an inhibitor of carbonic anhydrase V which has an impact on gluconeogenesis [28]. The current results showed that a high dose of QAE treatment decreased RAGE expression compared to T2DM mice. These data suggest that the QAE treatment has ameliorative effects on a hyperglycemic condition due to synergistic or additive effects of PCA, chlorogenic acid, and quercetin these active ingredients.

Moreover, there are well-known renal malfunction indicators including albuminuria, plasma creatinine, BUN, and urinary ACR level in DN. Our data showed that the QAE treatment significantly decreased urinary ACR, plasma creatinine, and BUN in diabetic mice. From these changes, it could be inferred that the QAE treatment improved renal function in a diabetic condition.

In terms of molecules, major mechanisms of hyperglycemia-induced tissue damage are as follows—the increase of intracellular AGEs formation and its receptor expression, and activation of PKC. As indicated above, the current study demonstrated that QAE supplementation reduced the protein level of RAGE in the diabetic mice. In addition, elevated protein levels of 4-HNE were decreased in the HQ group and both doses of QAE supplementation lowered protein carbonyls in the DMC group. Previous studies showed that the increased level of Nrf2 as well as 4-HNE and protein carbonyls activated Nrf2 related antioxidant defense systems [12,29]. Our results particularly demonstrated that the protein level of Nrf2 and its related antioxidant defense enzymes including NQO1, HO-1, and catalase were increased in the DMC mice but these markers were reduced in the HQ group. Especially, PCA is known to attenuate oxidative stress by decreasing the levels of ROS and malondialdehyde (MDA) in a diabetic condition [30]. Hence, it can be concluded that QAE containing PCA and CGA supplementation could alleviate cellular oxidative stress as well as activations of RAGE in diabetes.

Oxidative stress also can contribute to inflammatory response via the activation of NF-κB and downstream factors such as TNF-α, IL-6, and iNOS in DN [31]. Furthermore, oxidative stress would potentially activate NLRP3 inflammasome by initial recognition as cellular danger [32,33]. In this study, the renal protein levels of NLRP3 inflammasome, nuclear NF-κB, and subsequent inflammatory factors were higher in the DMC group compared with the NC group. A previous study demonstrated that the PCA treatment significantly reduced the secretion of pro-inflammatory cytokines in T2DM rats [34]. Furthermore, syringic acid is known to reduce oxidative stress and inflammation in diabetes [35]. Simultaneously, QAE supplementation selectively reduced the renal inflammatory factors via suppression of NLRP3 inflammasome. Therefore, the current study suggested that QAE supplementation alleviated the activation of NLRP3 inflammasome and consequential hyper-inflammation under a diabetic condition.

In consistent hyperglycemia, chronic hyper-inflammation in the kidney results in renal apoptosis via activation of caspases, proapoptotic protein Bax, p53, and mitogen activated protein kinase (MAPK) signaling [36–39]. PCA is known to reduce the protein expression levels of type IV collagen, laminin, and fibronectin in high glucose-stimulated human mesangial cells (MCs) [40]. Our results showed that pro-fibrosis related markers including PKC-βII, TGF-βI, and α-SMA as well as apoptosis related markers such as caspase-8, caspase-3, Bax/Bcl-2 ratio, and pERK/ERK ratio were declined in the QAE treated group, regardless of dose, compared with the DMC group. The present study suggested that PCA and chlorogenic acid in QAE might play major roles in the protection of renal apoptosis and fibrosis in T2DM.

How could the QAE treatment ameliorate the renal damage through suppression of oxidative stress, NLRP3 inflammasome-dependent hyper-inflammation, cell apoptosis, and pro-fibrosis in a hyperglycemic condition? There are cumulative evidences that AMPK influences intracellular signaling pathway, especially amelioration of oxidative stress via activation of antioxidant defense enzymes [41]. Metformin, which is a well-known diabetic drug, shows therapeutic mechanisms related to AMPK, which suppresses the NF-κB through activation of SIRT1 and PGC-1α [21,42,43]. PCA also increased the phosphorylation of AMPK and then activated the expression of p-Nrf2 and HO-1 in oxidative damage in HUVECs [44]. Moreover, a recent study reported that the syringic acid improved energy metabolism by regulation of mitochondrial biogenesis in diabetic rats [33]. On the other hand, SIRT1, an intracellular energy sensor, beneficially affected glucose homeostasis, cellular immunity to oxidative stress, inflammation, apoptosis, and fibrosis in the kidney [45]. In DN, one of the earliest characteristics is the loss of podocyte, which plays a crucial role in albumin processing, but SIRT1 is known to attenuate podocyte depletion and albuminuria by downregulation of claudin-1 in podocytes [46,47]. Resveratrol, a natural plant polyphenol, respectively stimulates SIRT1 and AMPK, and has a protective effect on oxidative stress and inflammatory response in the kidney [20]. A previous study showed that SIRT1 suppressed NLRP3 inflammasome activation as well as the NF-κB associated inflammatory response [48,49]. PGC-1α also regulates oxidative stress via participation in cellular signaling to mitochondrial oxidative stress and independently inhibits the NF-κB related inflammatory response [50,51]. The current studies demonstrated that the treatment of QAE containing PCA and syringic acid elevated the protein levels of SIRT1 and PGC-1α and downregulated NLRP3 inflammasome dependent inflammatory mediators in the diabetic mice. In particular, both doses of QAE supplementation has an effect on the stimulation of AMPK/SIRT pathway and a high dose of QAE supplementation decreased NLRP3 inflammation accompanied by nuclear NF-κB activation in our study. Therefore, it can be inferred that the QAE treatment has a protective effect on renal oxidative stress and hyper-inflammation under a hyperglycemic condition by involving this antagonism of SIRT1/NF-κB/NLRP3 inflammasome.

The previous study reported by our group found that the *Lespedeza bicolor* extract (LBE) containing polyphenolic compounds such as quercetin, genistein, daidzein, and naringenin has shown to exert antioxidant and anti-inflammatory effects accompanied by upregulation of the AMPK-SIRT1 pathway in the same diabetic model [52]. The current findings supported that QAE at much lower concentrations compared to LBE and other plant extracts has shown antidiabetic effects through regulation of the AMPK-SIRT related mechanism as shown in LBE treated diabetic mice [52]. Moreover, QAE supplementation attenuated pro-fibrosis as well as apoptosis in the diabetic group, which was not shown in LBE treatment groups. Therefore, it can be concluded that QAE is more effective than LBE on renal fibrosis and apoptosis in diabetes.

#### **5. Conclusions**

Taken together, we reported that QAE supplementation at a high dose had ameliorative effects on renal NLRP3 inflammasome associated hyper-inflammation and consequent renal cell apoptosis and pro-fibrosis in the HFD/STZ-induced T2DM mice. In addition, QAE supplementation regardless of the dose stimulated AMPK/SIRT1 signaling and ameliorated oxidative stress, although some molecular markers were selectively regulated at different treatment doses of QAE in diabetic renal damage. In conclusion, the current study suggested that QAE could be a potential therapeutic for improving renal damage in T2DM.

**Author Contributions:** Conceptualization, Y.L.; Data curation, J.E.P. and H.L.; Formal analysis, Y.L. and J.E.P.; Investigation, Y.L. and S.Y.K.; Funding acquisition, Y.L.; HPLC analysis, S.M.H.; Methodology, Y.L., J.E.P., H.L., and H.R.; Supervision, Y.L.; Writing—Review and editing, J.E.P., H.L., H.R., and Y.L. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was supported by iPET [115045-03] (Korea Institute of Planning and Evaluation for Technology in Food, Agriculture, Forestry and Fisheries), Ministry of Agriculture, Food and Rural Affairs.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **Abbreviations**


#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

#### *Article* **Protective E**ff**ects of Myricetin on Benzo[a]pyrene-Induced 8-Hydroxy-2**- **-Deoxyguanosine and BPDE-DNA Adduct**

#### **Seung-Cheol Jee 1, Min Kim 1, Kyeong Seok Kim 2, Hyung-Sik Kim <sup>2</sup> and Jung-Suk Sung 1,\***


Received: 30 April 2020; Accepted: 19 May 2020; Published: 21 May 2020

**Abstract:** Benzo[a]pyrene (B[a]P), a group 1 carcinogen, induces mutagenic DNA adducts. Myricetin is present in many natural foods with diverse biological activities, such as anti-oxidative and anti-cancer activities. The aim of this study was to investigate the protective effects of myricetin against B[a]P-induced toxicity. Treatment of B[a]P induced cytotoxicity on HepG2 cells, whereas co-treatment of myricetin with B[a]P reduced the formation of the B[a]P-7,8-dihydrodiol-9,10-epoxide (BPDE)-DNA adduct, which recovered cell viability. Furthermore, we found a protective effect of myricetin against B[a]P-induced genotoxicity in rats, via myricetin-induced inhibition of 8-hydroxy-2- -deoxyguanosine (8-OHdG) and BPDE-DNA adduct formation in the liver, kidney, colon, and stomach tissue. This inhibition was more prominent in the liver than in other tissues. Correspondingly, myricetin regulated the phase I and II enzymes that inhibit B[a]P metabolism and B[a]P metabolites conjugated with DNA by reducing and inducing CYP1A1 and glutathione S-transferase (GST) expression, respectively. Taken together, this showed that myricetin attenuated B[a]P-induced genotoxicity via regulation of phase I and II enzymes. Our results suggest that myricetin is anti-genotoxic, and prevents oxidative DNA damage and BPDE-DNA adduct formation via regulation of phase I and II enzymes.

**Keywords:** Benzo[a]pyrene; myricetin; oxidative stress; BPDE-DNA adduct; phase detoxifying enzyme

#### **1. Introduction**

Polycyclic aromatic hydrocarbons (PAHs) are ubiquitous environmental chemical carcinogens that lead to genetic damage and possess highly bioaccumulation characteristic [1]. Benzo[a]pyrene (B[a]P) is a well-known PAH (Figure 1A), which is listed as group 1 carcinogens by the International Agency for Research on Cancer (IARC). B[a]P is toxic and its exposure is primarily due to the food chain [2] Previous studies have shown that the average amount of people's exposure to B[a]P is 8.09–9.20 ng/day [3]. This means that humans are exposed to a low dose of B[a]P over a lifetime. The long-term exposure of B[a]P can cause angiogenesis and metastasis in the skin, lungs, liver, colorectal and stomach [4]. Other studies support that long-term exposure of low-dose B[a]P induces cell angiogenesis and metastasis [5], and a low dose of B[a]P toxicity is enhanced by interaction with PM2.5 air pollutants [6]. B[a]P is converted to B[a]P-7,8-diol-9,10-epoxide (BPDE) which is ultimate metabolite of B[a]P (Figure 1B), and it leads to genetic toxicity via covalent binding with DNA [7]. In addition, B[a]P is linked to reactive oxygen species (ROS) formation, which induces genotoxicity via the formation of 8-hydroxy-2-deoxyguanosine (8-oxo-dG) [8].

B[a]P is metabolized by phase I enzymes, such as cytochrome P450 (CYP). Furthermore, CYP1A1 is associated with the B[a]P metabolism process [9]. Previous studies have shown that B[a]P is oxidized by CYP enzymes that induce a variety of B[a]P metabolite transitions that form DNA adducts [10,11]. After formation of DNA adducts, this leads to several diseases and cancers. The xenobiotic chemicals are generally converted into water-soluble metabolites by phase II enzymes in the cells, and are easily removed by phase III enzymes [12]. Uridine 5- -diphospho (UDP)-glucuronosyltransferases (UGTs), sulfotransferases (SULTs), glutathione S-transferases (GSTs), NAD(P)H: quinine oxidoreductase type 1 (NQO1), heme oxygenase-1 (HO-1), and *N*-acetyltransferase (NAT) are known as major phase II enzymes [13]. GST is a major enzyme of phase II detoxifying enzymes. It is activated when conjugated with BPDE and reduces DNA damage by counteracting B[a]P metabolites via inhibition of BPDE-DNA adduct formation [14,15]. In addition, GSH conjugates with B[a]P-7,8-dione, which attenuates the B[a]P-induced genotoxicity [16]. Following GSH–conjugation with B[a]P metabolites via phase II detoxifying enzymes, it is excreted by transporter genes, such as ABCC1 and ABCC2 [17]. Previous studies have showed that ABCC transporters are required to detoxify B[a]P [18]. The knockdown of ABCC2 increases the concentration of B[a]P metabolites in a variety of organs [19]. ABCC2 is required for the elimination of BPDE-DNA adducts in organs by B[a]P metabolite excretion.

B[a]P is produced during food processes such as broiling, frying, and roasting. These processes increase the concentration of B[a]P, which increases the risk of disease and cancer; therefore, it is important to prevent B[a]P-induced toxicity from the natural synthesis of B[a]P [20]. Natural compounds are widely used to reduce B[a]P-induced toxicity and in cancer therapy [21,22]. Myricetin (Figure 1C), which is found in vegetables, herbs, and fresh fruits, is used as a treatment against different cancers [23]. Previously, studies have showed that myricetin has potentially therapeutic properties, such as anti-oxidant, cytoprotective, and anti-cancer properties [24]. Additionally, another study has shown that myricetin is protective after B[a]P-induced DNA damage by regulating DNA strand breaks [25]. However, cancer-causing DNA mutations occur if the BPDE-DNA adducts and ROS-induced DNA damage are not repaired by base excision repair (BER) and nucleotide excision repair (NER). Therefore, it is important to prevent DNA damage by inhibiting DNA conjugation with B[a]P metabolites. In this study, we investigated the protective effect of myricetin against B[a]P-induced genotoxicity via the inhibition of DNA adduct formation by regulating phase I and II enzymes.

**Figure 1.** The structure of chemicals. (**A**) Benzo[a]pyrene (B[a]P), (**B**) B[a]P-7,8-dihydrodiol-9,10-epoxide (BPDE), and (**C**) myricetin.

#### **2. Materials and Methods**

#### *2.1. Chemicals and Reagents*

Benzo[a]pyrene (B[a]P), myricetin, dimethyl sulfoxide (DMSO), ammonium persulfate, nuclease P1, N,N,N'N'-Tetramethyl ethylenediamine (TEMED), protease inhibitor cocktail, phosphatase inhibitor cocktail II, phosphatase inhibitor cocktail III, glycine, sodium dodecyl sulfate (SDS), tris base, sodium chloride, tween 20, and 2-mercaptoethanol were obtained from Sigma-Aldrich Chemical (St. Louis, MO, USA). Sodium pyruvate, penicillin-streptomycin, and trypsin-ethylenediaminetetraacetic acid (EDTA) were obtained from Welgene (Daegu, Korea). Alkaline phosphatase (AP) was purchased from Takara Bio Inc (Shiga, Japan). Phosphate-buffered saline (PBS) was purchased from Biosesang (Seongnam, Korea). A total of 30% acrylamide/bis solution was purchased from Bio-Rad (Hercules, CA, USA). Antibodies, such as CYP1A1, CYP1B1, GST, ABCC2, β-actin, and HRP-conjugated anti-rabbit immunoglobulin G (IgG), were purchased from Santa Cruz Biotechnology (Santa Cruz, CA, USA).

#### *2.2. Cell Culture*

Human-derived liver cancer cells (HepG2) were purchased from the American Type Culture Collection (Manassas, VA, USA). HepG2 cells were grown with Eagle's minimum essential medium (MEM, Welgene, Daegu, Korea) containing 10% fetal bovine serum (FBS, Welgene), 100 μg/mL streptomycin, 100 U/mL penicillin, and 1 mM sodium pyruvate in 100 mm<sup>2</sup> cell culture dishes. The old media were replaced with a new medium every two days. The HepG2 cells were incubated at 37 ◦C in 5% CO2 under a humidified atmosphere. The cells were used for further research when the confluency reached 80%.

#### *2.3. Animals and Housing*

Sprague-Dawley male rats (5-weeks-old; weight, 140–150 g) were purchased from Orient Bio (Seongnam-si, Korea) and were housed at a 23 ± 2 ◦C of temperature and 55 ± 1% of humidity. The room condition was maintained at specific pathogen free (SPF) with a 12 h light/12 h darkness cycle. Rodent chow and water were supplied ad libitum. The rats were randomly divided into three groups in each group (*n* = 6): (1) Control group, administration of corn oil (oral); (2) B[a]P-treated group, administration of B[a]P (2 mg/kg; daily, oral) dissolved in corn oil; (3) B[a]P co-treated with myricetin group, administration of myricetin (15 mg/kg) with B[a]P (2 mg/kg; daily, oral) dissolved in corn oil for 55 days. The concentration and period of B[a]P exposure were following other studies. To determine the B[a]P and myricetin concentration, we considered following reasons: (1) people are normally exposed to low-dose of B[a]P. Previous studies showed that people were exposed to 14 and 59.2 μg/kg/day of PAHs [26,27]. The average amount of people's exposure to B[a]P is 8.09–9.20 ng/day [3]. Additionally, previous studies showed that short-term treatment of B[a]P for in vivo used the dose of B[a]P at 25 to 200 mg/kg in rats [28]. Moreover, another study suggested that low-dose of B[a]P concentration was defined under 12 mg/kg/day [29]. On the other hand, the animals were treated with doses of 50, 100, and 200 mg/kg of myricetin to rats for a long time [30]. (2) People are unavoidably exposed to B[a]P through foods and polluted air for a lifetime. Therefore, we consider the results and determine the long-term effects of a low dose of B[a]P and myricetin. Treatments were administered at the same time daily. Behavioral tests were performed in the morning. The animal experiment protocol was approved by Sungkyunkwan University Laboratory Animal Care Service (SKKU-2013-000105, 23 March 2013) in accordance with the Ministry of Food and Drug Safety (MFDS) Animal Protection of Korea (Oh-Song, Korea).

#### *2.4. Cell Viability Assay*

To evaluate the cytotoxicity of B[a]P and myricetin on HepG2 cells, a cell viability assay was performed. A density of 1 <sup>×</sup> 10<sup>4</sup> cells/well of HepG2 cells were seeded in 96-well plates. A variety of concentrations of B[a]P (0, 1, 2.5, 5, and 10 μM) and myricetin (5, 10, 20, and 40 μM) were incubated in the wells for 48 h at 37 ◦C. After 48 h incubation, to evaluate the cell viability, 10 μL of EZ-CYTOX reagent (DOGEN, Daejeon, Korea) were treated with 100 μL MEM to each well and incubated for 2 h at 37 ◦C. Relative absorbance of each well was read at 450 nm to measure the amount of cell viability using a microplate reader (Molecular Devices, San Jose, CA, USA).

#### *2.5. BPDE-DNA Adduct Formation Analysis*

DNA extraction was performed using a QIAamp DNA Mini Kit (Qiagen, Valencia, CA, USA), according to the manufacturer's instructions. DNA was isolated and the level of BPDE-DNA adduct formation was assessed using BPDE-DNA adduct ELISA kit (Cell Biolabs, San Diego, CA, USA) and following manufacturer's instructions. Briefly, DNA samples of 2 μg/mL concentration are prepared. An amount of 100 μL of each sample is treated in 96 well plates for 2 h at 37 ◦C and rinsed two times. After washing, 100 μL of anti-BPDE antibody is treated in each well and incubated for 2 h at room temperature. Each well is rinsed 5 times using washing buffer, and then secondary antibody is added in each well for 1 h at room temperature. Each well is rinsed 3 times using washing buffer. All wells are reacted with 3,3- ,5,5- -Tetramethylbenzidine (TMB) buffer at room temperature for 20 min. Finally, the reaction is stopped by adding stop solution. The BPDE-DNA adducts formation level was evaluated by measuring the relative absorbance using a microplate reader at 450 nm.

#### *2.6. Quantification of DNA Damage via 8-Hydroxydeoxyguanosine*

We evaluated the concentration of 8-hydroxydeoxyguanosine (8-OHdG) using an 8-OHdG DNA damage ELISA kit (Cell Biolabs, San Diego, CA, USA), according to manufacturer's instructions. Briefly, isolated DNA was denatured at 95 ◦C for 5 min and immediately transferred to ice. A total of 10 units of nuclease P1 and 20 mM sodium acetate (pH 5.2) were added to total DNA, and digested DNA to nucleosides for 2 h at 37 ◦C. Next, alkaline phosphatase (Takara, Japan) added for 15 min at 37 ◦C, followed by incubation for 15 min at 50 ◦C in 100 mM Tris buffer (pH 7.5). The reaction mixtures were centrifuged for 5 min at 6000× *g* and the supernatants were extracted for assay analysis. Briefly, 50 μL of each sample is treated in wells and incubated for 10 min at room temperature. An amount of 50 μL of anti-8-OHdG antibody is treated in each well and incubated for 1 h at room temperature on orbital shaker. Each well is rinsed 3 times, 100 μL secondary antibody is added in each well and incubated for 1 h at room temperature. After incubation, 100 μl of substrate solution buffer is added in each well for 20 min at room temperature and then the reaction is stopped by adding stop solution. The absorbance of 8-OHdG was measured using a microplate reader at 450 nm (VERSA max™, Molecular Devices, San Jose, CA, USA).

#### *2.7. Western Blot Analysis*

The protein in the liver was extracted using PRO-PREP™ protein extraction solution (iNtRON, Seongnam, Korea). The protein in cells was extracted in RIPA buffer (150 mM NaCl, 1% Nonidet P-40, 50 mM Tris-HCl, and 0.25% sodium deoxycholate) (Biosolution, Seoul, Korea) containing protease inhibitor cocktail, phosphatase inhibitor cocktail II and III. Each protein concentration was quantified using the Bio-Rad protein assay (Hercules, CA, USA). After protein quantification, it was denatured at 95 ◦C for 5 min in buffer. Next, samples (50 μg) were ran on 10% SDS-polyacrylamide gel electrophoresis (SDS-PAGE) at 50 V for 60 min in running buffer. The proteins were transferred to polyvinylidene difluoride (PVDF) membranes (BioRad, Hercules, CA, USA) at 100 V for 90 min in a transfer buffer. Membranes blocked with TNT buffer containing 5% skim milk for 1 h, followed by incubation with CYP1A1, CYP1B1, and β-actin antibodies overnight at 4 ◦C. After washing for 15 min at 4 times with TNT buffer, the membrane was incubated for 45 min with secondary antibodies, and then washed for 15 min at 4 times with TNT buffer. Immunoreactivity was visualized using chemiluminescence (ECL) Plus Western Blotting reagents (Amersham Bioscience, Buckinghamshire, UK). The protein level was quantified using Quantity One Image Software (Bio-Rad, Hercules, CA, USA).

#### *2.8. Statistical Analysis*

Experimental data were evaluated in triplicates and experiments were repeated at least three times. All data were expressed as mean ± standard error of the mean (SEM). The One-way analysis of variance (ANOVA) and Tukey's multiple comparison analysis were performed to determine the significant differences between groups using GraphPad Prism 5.0 (GraphPad Software, San Diego, CA, USA). Differences were considered statistically significant when *p* < 0.05.

#### **3. Results**

#### *3.1. Protective E*ff*ect of Myricetin against B[a]P-Induced Toxicity*

HepG2 cells were treated with different concentrations of B[a]P and myricetin to evaluate their toxicity. B[a]P treatment for 48 h induced cytotoxicity in a dose-dependent manner compared with the control group (Figure 2A). The 40% inhibitory concentration of B[a]P was calculated at 10 μM for 48 h and used in subsequent experiments. Treatment of myricetin at 5, 10, 20, and 40 μM for 48 h revealed no toxicity up to 10 μM; however, cytotoxicity was shown for treatments > 20 μM when compared with the control group (Figure 2B). Then, to confirm the protective effect of myricetin on B[a]P cytotoxicity, HepG2 cells were co-treated with or without B[a]P in the presence of myricetin and the amount of the viable cells continuously analyzed compared with non-treated group. We found that myricetin co-treatment with B[a]P recovered up to 85% cell viability (Figure 2C). These results suggest that myricetin has protective effect against B[a]P-induced cytotoxicity. Based on the cell viability data, 10 μM of myricetin was used for all further in vitro experiments.

**Figure 2.** B[a]P and myricetin cytotoxicity in HepG2 cells. (**A**,**B**) HepG2 cells were treated with B[a]P (0, 1, 2.5, 5, and 10 μM) or myricetin (0, 5, 10, 20, and 40 μM) at different concentrations for 48 h. (**C**) The protective effect of myricetin against B[a]P-induced cytotoxicity was measured with B[a]P (10 μM) co-treatment with various concentrations of myricetin for 48 h. All treatment group values are significantly different when compared with controls (\* *p* < 0.05, \*\*\* *p* < 0.001) and B[a]P (# *p* < 0.05, ### *p* < 0.001). Tukey's multiple comparison test. M: myricetin.

#### *3.2. Protective E*ff*ects of Myricetin against B[a]P-Induced Oxidative DNA Damage*

To confirm the B[a]P-induced oxidative DNA damage, we calculated the concentration of 8-OHdG in the liver, kidney, colon, and stomach tissue of rats following treatment with 2 mg/kg of B[a]P and 15 mg/kg of myricetin for 55 days. B[a]P significantly increased the concentration of 8-OHdG in the liver, stomach, colon, and kidney when compared with the control group but B[a]P co-treatment with myricetin significantly reduced the 8-OHdG in the liver, stomach, and kidney respectively (Figure 3A–D). These results confirmed that myricetin protects DNA from further oxidation that contributes to cell death prevention.

**Figure 3.** Oxidative DNA damage following B[a]P (2 mg/kg) and myricetin (10 mg/kg) co-treatment. Oxidative DNA damage was measured by the concentration of 8-OHdG in liver, stomach, colon, and kidney tissues. (**A**–**D**) Myricetin was protective against B[a]P-induced 8-OHdG formation following B[a]P treatment in all tissue when compared with controls (\* *p* < 0.05, \*\* *p* < 0.01, \*\*\* *p* < 0.001) and B[a]P (# *p* < 0.05, ### *p* < 0.001). Tukey's multiple comparison test. M: myricetin.

#### *3.3. Inhibition E*ff*ect of Myricetin against BPDE-DNA Adduct Formation*

To evaluate the protective effect of myricetin against B[a]P-induced genotoxicity, the concentration of BPDE-DNA adducts was measured in HepG2 cells and the liver, stomach, colon, and kidney tissue of rats using BPDE-DNA Adduct ELISA Kit. Treatment with B[a]P, myricetin, and B[a]P + myricetin showed that B[a]P induced BPDE-DNA adduct formation when compared with the non-treatment group. However, B[a]P co-treatment with myricetin clearly decreased BPDE-DNA adduct formation when compared with B[a]P treatment alone (Figure 4A). Furthermore, B[a]P induced BPDE-DNA adduct formation in the liver, kidney, colon, and stomach of treated rats when compared with the control group. B[a]P co-treatment with myricetin inhibited the BPDE-DNA adduct formation in all rat tissue (Figure 4B–E). These results showed that the genotoxicity of B[a]P was attenuated by myricetin via inhibition of the formation of BPDE-DNA adduct.

**Figure 4.** BPDE-DNA adduct concentration in B[a]P and B[a]P + myricetin treated groups. (**A**) Effect of myricetin on BPDE-DNA adduct formation in HepG2 cells following treatment with B[a]P (10 μM) and co-treatment with myricetin (10 μM) for 48 h. (**B**–**E**) In Sprague-Dawley rats, treatment with B[a]P (2 mg/kg) and co-treatment with myricetin (15 mg/kg) for 55 days. All treatment groups are significantly different when compared with controls (\*\* *p* < 0.01, \*\*\* *p* < 0.001) and B[a]P (# *p* < 0.05, ## *p* < 0.01, ### *p* < 0.001). Tukey's multiple comparison test. M: myricetin.

#### *3.4. Regulatory E*ff*ect of Myricetin on the Expression of Phase I, II, and III Enzyme*

Our results showed that B[a]P induced CYP1A1 and CYP1B1 expression. In contrast, B[a]P + myricetin co-treatment down-regulated CYP1A1, but not CYP1B1, expression in the liver of rats (Figure 5A,B). We confirmed that myricetin co-treatment with B[a]P reduced CYP1A1 expression when compared with B[a]P treatment group (Figure 5C,D). This indicated that myricetin regulated CYP1A1 expression, thereby reducing B[a]P metabolism, which prevented DNA adduct formation.

Next, we measured GST expression in HepG2 cells following B[a]P and B[a]P + myricetin treatment. B[a]P reduced GST expression; however, co-treatment with myricetin recovered this expression when compared with B[a]P treatment alone (Figure 5C,D). In contrast, ABCC2 was not regulated by myricetin. This indicated that myricetin attenuated B[a]P-induced toxicity via reduction of oxidative DNA damage and inhibition of BPDE-DNA adduct formation by inducing GSH conjugated with B[a]P metabolites to recover the GST expression.

**Figure 5.** The expression of phase detoxifying enzymes. (**A**,**B**) The effect of myricetin (15 mg/kg) and B[a]P (2 mg/kg) treatment on CYP1A1 and CYP1B1 enzyme expression in the liver of Sprague-Dawley rats. (**C**) The effect of myricetin (10 μM) and B[a]P (10 μM) treatment on CYP1A1, GST, and ABCC2 enzyme expression in HepG2 cells. (**D**) Quantitative evaluation of relative protein expression of CYP1A1, GST, and ABCC2. All treatment groups are significantly different when compared with controls (\*\* *p* < 0.01, \*\*\* *p* < 0.001) and B[a]P (### *p* < 0.001). Tukey's multiple comparison test. M: myricetin.

#### **4. Discussion**

B[a]P is carcinogenic, and is formed during food-processing, and in tobacco smoke, waste products, and industry [31,32]. B[a]P is well-known as an ubiquitous environmental agent. Previous studies showed that people were exposed to 8.09–9.20 ng/day of B[a]P [3]. This report indicates that people are exposed to a low dose of B[a]P over their lifetime. Generally, B[a]P accumulated in humans through food; 97% of B[a]P exposure amount is associated with the food chain [2]. This indicates that the exposure of B[a]P is mainly mediated in the gastrointestinal tract such as stomach and colon. Absorbed B[a]P through the gastrointestinal tract passes through the blood vessel to the liver and undergoes the detoxification process. Then, detoxified metabolites are excreted via the urine or bile. Thus, in this study, the liver, kidney, and gastrointestinal tract (stomach and colon) were considered as target organs. After exposure to B[a]P, it is metabolized to BPDE or B[a]P radical cations, which are conjugated with DNA. Previous studies have shown that B[a]P induces genotoxicity by forming BPDE-DNA adducts or 8-OHdG [33]. Additionally, another study shows that the long-term treatment of low-dose B[a]P induces the cancer progression [5]. This means that B[a]P negatively affects the people's health causing dysfunction of hormone, cancers, and autoimmune disease [34]. Previous study shows that inhibition of DNA adduct formation reduces B[a]P mutagenesis and carcinogenesis [35]. Many studies have assessed the effect of flavonoids in reducing the B[a]P-induced toxicity [36–38].

Myricetin, which is present in many foods such as tea, vegetables, and medical plants, is a natural flavonoid that has anti-carcinogenic, -oxidant, and -inflammation effects [39,40]. Generally, the protective effects of myricetin against oxidative stress have been well-studied for a long time. Previous studies have shown that the anti-oxidant effect of myricetin is mediated by its ROS scavenging activity and activation of cellular anti-oxidative mechanisms including nuclear factor erythroid 2-related factor 2 (Nrf2)-linked pathway [41–43]. Myricetin is protective against B[a]P-induced DNA damage via oxidized pyrimidine. In contrast, oxidized purines are not reduced by myricetin [25]. This means that myricetin does not repair after receiving purine-based damage by B[a]P. Therefore, the inhibition of 8-OHdG and BPDE-DNA adducts is important for the attenuation of B[a]P-induced

DNA damage. In this study, we provide new insights into the protective effects of myricetin against B[a]P-induced toxicity.

We confirmed that myricetin recovered cell viability in HepG2 cells, indicating that myricetin could attenuate B[a]P-induced cytotoxicity (Figure 2C). Previous studies have shown that the formation of DNA adducts induces B[a]P genotoxicity via oxidative stress and BPDE. B[a]P is transited to B[a]P-7,8-dione, which enhances ROS production by inducing futile redox cycles [44]. The formation of 8-OHdG is an oxidative DNA damage marker [45]. In addition, B[a]P is sequentially transited to B[a]P-7,8-epoxide, B[a]P-7,8-dihydrodiol, and BPDE, which is a metabolite of B[a]P that causes genotoxicity via the formation of BPDE-DNA adducts [35]. On the other hand, the amount of oxidative DNA damage and the level of BPDE formation are different depending on the types of cell lines and organs [46]. Thus, to evaluate the effect of myricetin on B[a]P-induced toxicity, we analyzed 8-OHdG and BPDE-DNA adducts using HepG2 cells and various organs such as the liver, kidney and gastrointestinal tract (stomach and colon). Our results confirmed that B[a]P induced 8-OHdG and BPDE-DNA adduct formation, respectively. In contrast, co-treatment with myricetin and B[a]P reduced 8-OHdG and BPDE-DNA adduct formation in all organ tissues (Figures 3 and 4). The results indicate that reduction of B[a]P-induced cytotoxicity is induced by myricetin through reducing B[a]P metabolites-DNA adducts formation. To examine whether the side-effects affect the animals by treatment with B[a]P or myricetin, we evaluated the effect of B[a]P and myricetin on body and organ weights. We confirmed that the weights of organs including the liver, stomach, colon, and kidney were not changed in all treatment groups (supplementary information, Figure S1B–E). Moreover, the bodyweight of animals was not significantly different between the treated groups and untreated control groups (supplementary information, Figure S1A). These results indicated that there were no significant side-effects during long-term exposure to B[a]P and myricetin. Therefore, our results show that myricetin attenuates B[a]P-induced genotoxicity mainly by inhibiting oxidative DNA damage and BPDE-DNA adduct formation.

B[a]P-mediated genotoxicity is caused by the interaction of B[a]P metabolites with DNA. Our results show that myricetin reduces B[a]P-induced genotoxicity by reducing the formation of B[a]P metabolites-DNA adduct. We hypothesize that myricetin inhibits the opportunity of B[a]P metabolites conjugated with DNA. Attenuation of B[a]P-DNA adduct formation was considered to be mediated by two factors: (1) reduction in conversion of B[a]P to B[a]P metabolites, and (2) elimination of B[a]P metabolites. Previous studies have showed that the conversion of B[a]P to BPDE requires multi-enzymatic steps [47]. B[a]P is metabolized by CYP 450; its metabolites cause 8-OHdG and BPDE-DNA adduct formation. A report shows that B[a]P metabolism is regulated by CYP1A1 and CYP1B1, which are associated with B[a]P-mediated carcinogenesis [48]. CYP 450 enzymes induce B[a]P conversion to its metabolites, which conjugate with DNA to form carcinogenic DNA adducts [49]. However, our results showed that CYP1A1, but not CYP1B1, expression was decreased by myricetin treatment when compared with B[a]P alone (Figure 5A,B). A previous study has reported that the regulation of BPDE-DNA adduct formation is correlated with CYP1A1 [50]. Furthermore, the attenuation of cellular DNA damage in HepG2 cells is associated with a reduction in oxidative stress via suppression of CYP1A1 [51]. Indeed, CYP1A1 is the most efficient enzyme forming intermediate metabolite of B[a]P-derived DNA adducts in the liver [52]. Taken together with our data, this indicates that myricetin-induced attenuation of B[a]P genotoxicity is caused by reducing the intermediate metabolites of B[a]P-derived DNA adducts via regulation of CYP1A1 expression.

GST is a phase II enzyme that is conjugated with xenobiotics, including B[a]P, which is a well-known detoxification system in the body [53]. One previous study has shown that GST contributes to a reduction in BPDE-DNA adduct formation and 8-OHdG by inducing GSH-conjugation with B[a]P metabolites [54]. Our results showed that B[a]P reduced GST expression, whereas myricetin recovered this effect (Figure 5C,D). These results suggested that myricetin attenuates B[a]P-induced DNA damage by inhibiting 8-OHdG and BPDE-DNA adduct formation via GST expression induction. In contrast, phase III enzymes are not significantly regulated by myricetin (Figure 5C,D). We hypothesize that

myricetin regulates other transporter genes or does not affect phase III enzymes. Previous studies have shown that myricetin induces anticancer drug efficiency by modulating drug efflux via transporter gene regulation [55,56]. In addition, many transporter genes have been shown to localize to the basolateral or canalicular membrane, which is associated with the excretion of drugs through the bile or blood [57]. Previous studies have revealed that GSH interacting with BPDE enhances the BPDE solubility in water, which is eliminated by ABCCs [58]. Therefore, further studies are required to assess the excretion mechanism of B[a]P metabolites that are conjugated with phase II enzymes by myricetin. Taken together, myricetin attenuates B[a]P-induced 8-OHdG and BPDE-DNA adduct formation by regulating CYP1A1 and GST expression.

#### **5. Conclusions**

This study shows that myricetin reduces B[a]P-induced toxicity by inhibiting BPDE-DNA adduct and 8-OHdG formation. The inhibition of B[a]P-induced genotoxicity is associated with two factors: (1) a reduction of the B[a]P metabolism via reduced CYP1A1 expression, and (2) the elimination of B[a]P metabolites via enhanced GST expression. These results indicate that myricetin inhibits both B[a]P conversion to B[a]P metabolites and B[a]P metabolites conjugated with DNA, thereby inhibiting the formation of 8-OHdG and BPDE-DNA adducts. Our study suggests that myricetin is protective against B[a]P-induced genotoxicity by inhibiting oxidative DNA damage and BPDE-DNA adduct formation.

**Supplementary Materials:** The following are available online at http://www.mdpi.com/2076-3921/9/5/446/s1, Figure S1: Effect of B[a]P and myricetin on body and organ weights. B[a]P (2 mg/kg) alone or together with myricetin (15 mg/kg) was administered to Sprague-Dawley rats orally for 55 days. (**A**) Body weight changes of the rats during 55 days. (**B–E**) Organ weights of the rats after 55 days. M: myricetin.

**Author Contributions:** S.-C.J. designed the study, performed the experiments, analyzed the data, and wrote the manuscript. M.K. performed the experiments and analyzed the data. K.S.K. performed the experiments and analyzed the data. H.-S.K. contributed to the writing of the manuscript. J.-S.S. designed the study and contributed to the writing of the manuscript. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was supported by a grant (14162MFDS072) from the Ministry of Food and Drug Safety (MFDS, Korea, 2017) and the Dongguk University Research Fund of 2019.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

### *Article* **Nitric Oxide Modulation by Folic Acid Fortification**

**Junsei Taira 1,\* and Takayuki Ogi <sup>2</sup>**


Received: 26 March 2020; Accepted: 4 May 2020; Published: 7 May 2020

**Abstract:** Folic acid (FA) can be protected the neural tube defects (NTDs) causing nitric oxide (NO) induction, but the alleviation mechanism of the detailed FA function against NO has not yet been clarified. This study focused on elucidation of the interaction of FA and NO. FA suppressed nitrite accumulation as the NO indicator in lipopolysaccharide (LPS)-stimulated RAW264.7 cells, then the expression of the *i*NOS gene due to the LPS treatment was not inhibited by FA, suggesting that FA can modulate against NO or nitrogen radicals. NOR3 (4-ethyl-2-hydroxyamino-5-nitro-3-hexenamide) as the NO donor was used for evaluation of the NO scavenging activity of FA. FA suppressed the nitrite accumulation in a dose-dependent manner. To confirm the reaction product of FA and NO (FA-NO), liquid chromatography–mass spectrometry (LC/MS) was used to measure a similar system containing NOR3 and FA, and then detected the mass numbers of the FA-NO as *m*/*z* 470.9 (M + H)<sup>+</sup> and *m*/*z* 469.1 (M <sup>−</sup> H)−. In addition, the adducts of the FA-NO derived from 14NO and 15NO gave individual mass numbers of the isotopic ratio of nitrogen for the following products: FA-14NO, *m*/*z* 471.14 (M + H)+; *<sup>m</sup>*/*<sup>z</sup>* 469.17 (M <sup>−</sup> H)<sup>−</sup> and FA-15NO, *<sup>m</sup>*/*<sup>z</sup>* 472.16 (M <sup>+</sup> H)+; *<sup>m</sup>*/*<sup>z</sup>* 470.12 (M <sup>−</sup> H)–. To clarify the detailed NO scavenging action of FA, an electron spin resonance (ESR) study for radical detecting of the system containing carboxy-PTIO (2-(4-carboxyphenyl)-4,4,5,5-tetramethylimidazoline-1-oxyl-3-oxide) as an NO detection reagent in the presence of NOR3 and FA was performed. The carboxy-PTI (2-carboxyphenyl-4,4,5,5-tetramethylimidazoline-1-oxyl) radical produced from the reaction with NO reduced in the presence of FA showing that FA can directly scavenge NO. These results indicated that NO scavenging activity of FA reduced the accumulation of nitrite in the LPS-stimulated RAW264.7 cells. The NO modulation due to FA would be responsible for the alleviation from the failure in neural tube formation causing a high level of NO production.

**Keywords:** folic acid; nitric oxide; neural tube defects; RAW264.7 cells; NOR3; ESR; LC/MS

#### **1. Introduction**

The falling of neural tube defects (NTDs) in neural tube formation during early embryogenesis includes anencephaly, exencephaly, and spina bifida. Folic acid (FA) is known as a dietary supplement that can be prevent NTDs involving failure of the neural tube (NT) closure in the developing embryo, especially spina bifida and anencephaly in the periconceptional period [1].

Based on this information, the mandatory FA fortification has been associated with a decline in NTD prevalence in many countries [2–4]. A more recent study demonstrated that microglia activation, including the disruption of the endogenous inhibitory system (CD200-CD200R), contributes to injury in spina bifida aperta after birth [5]. Previous study indicated that a moderate level of nitric oxide (NO) and nitric oxide synthase (NOS) play a critical role in normal embryonic development [6]. Physiological concentrations of NO modulate carbon flow through the folate pathway, that is, NO

inhibits methionine synthase (MS) involving the interference transfer of the methyl group from the methyl donor, 5-methyl-tetrahydrofolate (5mTHF), to homocysteine during methionine production [7].

In previous studies, the direct effect of FA against NO produced by the NO donor, *S*-nitroso-*N*-acetyl-penicillamine (SNAP), on the process of NT closure in the chick embryo ex ovo was examined. NOS involves high NO levels due to the SNAP treatment of the inactivated MS and its activity can only be rescued by FA or vitamin B12, but not by the NOS inhibitor [8,9]. Although FA or vitamin B12 can prevent the NTDs causing the NO induction, the detailed prevention mechanism due to the FA relation to NO has not yet been clarified. Therefore, this study focused on the interaction of FA and NO, and this article describes how FA can directly scavenge NO involved in NTDs.

#### **2. Materials and Methods**

#### *2.1. Reagents*

4-Ethyl-2-hydroxyamino-5-nitro-3-hexenamide (alternate name: NOR3) and 2-(4-carboxyphenyl)- 4,4,5,5-tetramethylimidazoline-1-oxyl-3-oxide (alternate name: carboxy-PTIO) were purchased from Dojindo Molecular Technique Inc. (Kumamoto, Japan). Folic acid (FA), l-arginine, interferon-γ (IFN-γ) and lipopolysaccharide (LPS) were obtained from the FUJI Firm Wako Pure Chemical Corporation (Osaka, Japan). Dulbecco's modified Eagle's medium (DMEM) and fetal bovine serum (FBS) were obtained from Gibco BRL (Grand Island, NY, USA).

#### *2.2. Cell Culture*

Raw264.7 cells (mouse macrophages, American type culture collection) were cultured in DMEM in 10% FBS, 100 U/mL penicillin and 100 μg/mL streptomycin at 37 ◦C in a 5% CO2 atmosphere.

#### *2.3. Nitrite Assay on RAW264.7 Macrophages*

RAW264.7 macrophages in a 96-well microplate were treated with LPS (100 ng/mL), l-arginine (2 mM), and IFN-γ (100 U/mL) with or without the various concentrations of FA (25–200 μM). After culturing for 16 h, the nitrite production as an NO indicator in the medium, was determined by the Griess method as previously reported [10,11].

#### *2.4. iNOS Gene Expression*

Reverse transcriptase-polymerase chain reaction (RT-PCR) was carried out according to previously described procedures [12]. Briefly, the LPS-stimulated RAW264.7 macrophages on a 12-well microplate (2.5 <sup>×</sup> 10<sup>6</sup> cells/mL) were treated with the FA (200 <sup>μ</sup>M). The total RNA from cells was isolated from the cell lysate and the amplification of the cDNA was performed by the *i*NOS primers: 5'-CCT TGT TCA GCT ACG CCT TC-3'and 5'-CTG AGG GCT CTG TTG AGG TC-3' using PCR (GeneAmp® PCR System 9700, Applied Biosystems, Waltham, MA. USA). The PCR product of cDNA (100 ng/μL) was loaded on a DNA chip (Agilent DNA 1000 kit, Agilent Technologies, Santa Clara, CA, USA) and the electrophoresis was performed by a micro DNA analyzer (Agilent 2100 Bioanalyzer, Agilent Technologies, Santa Clara, CA, USA).

#### *2.5. Nitric Oxide (NO) Inhibitory Action*

NOR3 as the NO donor was used in the presence of various concentrations of FA (10–200 μM) as previously reported [13]. The reaction mixture containing NOR3 (200 μM), with or without FA in phosphate-buffered saline (PBS) solution, was incubated at room temperature for 60 min. The nitrite accumulation in the reaction mixture was determined using previously reported procedures [10,11]. In addition, the reaction mixture was analyzed by liquid chromatography–mass spectrometry (LC/MS) equipment. The reaction product of FA and NO was measured by LC/MS using a photodiode array detector (Quattromicro API triple-quadruple mass analyzer, Waters Corp., Milford, MA, USA) and monitored at 275 nm on a reversed-phase chromatographic column, YMC Pro C18 (i.d. 3 mm × 10 cm, YMC Co., Ltd., Kyoto, Japan) at 40 ◦C. The mobile phase consisting of 1% formic acid and acetonitrile solvent (10%) was carried out at the flow rate of 0.34 mL/min by a linear gradient to 10%, 30% and 100% for 2 min, 3 min and 7 min. The mass spectra were measured under the following conditions: electrospray ionization (ESI) at the cone voltages of 17 volts for the positive ion mode and at 21 volts for the negative ion mode.

#### *2.6. Electron Spin Resonance (ESR) Measurement*

The NO scavenging ability of FA was examined in the presence of the carboxy PTIO as the NO detection reagent [14]. The reaction mixture of FA (200 μM), NOR3 (100 μM) and carboxy-PTIO (50 μM) was prepared in PBS and incubated at room temperature for 15 min. An electron spin resonance (ESR) measurement was performed by an ESR spectrometer (JES-FR30, JEOL, Ltd., Tokyo, Japan) operating at the X-band with the modulation frequency of 100 kHz. The reaction mixture was transferred to the capillary (100 × 1.1 mm i.d. (inner diameter)., Drummond Scientific Co., Broomall, PA, USA) and placed in a quartz cell (270 mm long, 5 mm i.d., JEOL DATUM, Ltd., Tokyo, Japan). The ESR signal was measured at 9.4 GHz resonant frequency the following conditions: microwave power; 4 mW; modulation width, 0.1 mT; gain, 500; scan time, 1 min; time constant, 0.3 s.

#### *2.7. Reaction Product of Folic Acid (FA) and NO*

The reaction products of FA and NO were prepared as previously reported [11,15]. Milli-Q water (200 mL) was degassed using an ultrasonic device under reduced pressure for 30 min, subsequently a dry nitrogen gas was bubbled for 30 min in a nitrogen gas-filled glove box (762 × 450 × 478 mm, AS-600PC, AS ONE corp., Osaka, Japan). The status of deoxygenation in the aqueous solution was confirmed using an oximeter (MDS-2C, Marubishi Bioengineering Co., Ltd., Tokyo, Japan). This deoxygenated aqueous solution (1 mL) containing FA (4.4 mg), Na2S2O4 (20 mg), Na14NO2 or Na15NO2 (40 mg) was incubated in a capped vial at room temperature for 1 h under the anoxic conditions in the glove box. While the similar reaction mixture as control was prepared under the aerobic conditions. Then, the product of NO (FA-14NO and FA-15NO) was immediately analyzed by LC/MS equipment. The reaction product of FA and NO was measured by LC/MS using a photodiode array detector (Xevo-TQD triple-quadrupole mass analyzer, Waters, MA, USA) and monitored at 275 nm on a reversed-phase chromatographic column, Acquity UPLC BEH C18 (i.d. 2.1 mm × 3 cm, 1.7 μm, Waters) at 40 ◦C. The mobile phase consisting of 0.1% formic acid aqueous solution (100%) and acetonitrile solvent containing 0.1% formic acid was carried out at the flow rate of 0.40 mL/min by a linear gradient to 100%, 5%, 5% for 0.5 min, 3.5 min, 5min. These mass spectra were measured under the following conditions: ESI at the cone voltages of 30 volt in both the positive and negative ion mode.

#### **3. Results**

#### *3.1. NO Inhibitory Action*

The NO inhibitory action due to FA was evaluated for the NO production in the LPS-stimulated RAW264.7 cells. The nitrite accumulation as the NO indicator increased in the LPS treated cells. When FA was placed in the cells, the nitrite accumulation was inhibited in a dose dependent manner (Figure 1). A previous study showed a similar result that FA inhibited cytotoxicity with apoptosis due to NO [16]. This result indicated that FA has an inhibitory action on the NO production.

#### *3.2. NO Scavenging Activity*

The NO scavenging activity due to FA was evaluated using NOR3 as the NO donor. The nitrite accumulation as the NO indicator was examined with or without FA. As shown in Figure 2, FA suppressed the nitrite accumulation in a dose-dependent manner. This result suggested that FA has a potential NO or nitrogen radical scavenging activity. Therefore, the NO scavenging ability due to FA would be responsible for the suppression of the NO production in the LPS-stimulated RAW264.7 cells (Figure 1).

**Figure 1.** Inhibition of folic acid (FA) for nitric oxide (NO) production in lipopolysaccharide (LPS)-stimulated RAW264.7 macrophages. The various concentrations of FA (25, 50, 100 and 200 μM) were evaluated for the NO production in the LPS-stimulated RAW264.7 macrophages. Data are expressed as mean ± standard deviation (SD) and the significant difference was analyzed by the Student's *t*-test. \**p* < 0.01 indicates significant difference from the control.

**Figure 2.** Inhibition of nitrite accumulation due to folic acid (FA). The reaction mixture containing NOR3 (4-ethyl-2-hydroxyamino-5-nitro-3-hexenamide) (200 μM) as the NO donor with or without FA (10, 25, 50, 100 and 200 μM) in phosphate-buffered saline (PBS) solution was incubated at room temperature for 60 min. The nitrite level was used as the NO indicator. Data are expressed as mean ± SD, and the significant difference was analyzed by the Student's *t*-test. \**p* < 0.01 indicates significant difference from NOR3 without the test compounds.

#### *3.3. Suppression of iNOS Gene Expression*

The *i*NOS mRNA gene expression was induced in the LPS-stimulated RAW264.7 cells. The *i*NOS gene expression in the cells was examined with or without FA. As shown in Figure 3, the *i*NOS gene expression by the LPS treatment was not suppressed by the FA treatment. This result suggested that FA can directly scavenge NO or nitrogen radicals which would be responsible for the suppression of the NO production in the LPS-stimulated cells (Figure 1).

**Figure 3.** Inhibitory effect of folic acid (FA) for LPS stimulated *i*NOS mRNA expression in LPS-stimulated RAW264.7 macrophages. An *i*NOS mRNA gene expression was induced in the LPS-stimulated RAW264.7 cells. The *i*NOS gene expression in the cells was examined with or without FA (200 μM). Cells, cells without treatment; cells/LPS, cells treated with LPS; cells/LPS + FA, cells treated with LPS and cells + FA, cells treated with FA.

#### *3.4. Determination of FA-NO by Liquid Chromatography–Mass Spectrometry (LC*/*MS)*

To confirm the reaction product of FA and NO (FA-NO), LC/MS was used to measure in the reaction mixture containing NOR3 and FA. The high-performance liquid chromatography (HPLC) chromatogram of the FA-NO produced in the presence of FA and NOR3 indicated in Figure 4. The mass spectrum of the reaction products was detected in both the positive and negative ion mode. The mass numbers of *<sup>m</sup>*/*<sup>z</sup>* 470.9 (M <sup>+</sup> H)<sup>+</sup> and *<sup>m</sup>*/*<sup>z</sup>* 469.1 (M <sup>−</sup> H)<sup>−</sup> indicated that the FA-NO was produced by the interaction of FA and NO.

**Figure 4.** Liquid chromatography–mass spectrometry (LC/MS) chromatogram obtained from reaction mixture containing NOR3 and folic acid (FA). The LC/MS was carried out under the analytical conditions as described in the text. (**A**) Each peak of the high-performance liquid chromatography (HPLC) chromatogram indicated FA and its reaction product with NO, FA-NO. (**B**) These mass spectra indicated (**a**) FA, *m*/*z* 441.9 (M + H)<sup>+</sup> and (**b**) FA-NO, *m*/*z* 470.9 (M + H)<sup>+</sup> in the positive ion mode and (**c**) FA, *m*/*z* 440.0 (M − H)<sup>−</sup> and (**d**) FA-NO, *m*/*z* 469.1 (M − H)<sup>−</sup> in the negative ion mode of mass spectrometry.

#### *3.5. Products of FA-14NO and FA-15NO*

To confirm the ability of the FA scavenging of NO, the product of FA-NO derived from 14NO and 15NO was prepared, and then each mass spectrum was measured by LC/MS. As shown in Figures 5 and 6, each reaction product of 14NO and 15NO with FA under the conditions of aerobic and anoxia was as follows; FA-14NO, *m*/*z* 469.17 (M <sup>−</sup> H)−; *m*/*z* 471.1 4 (M + H)<sup>+</sup> for the aerobic conditions and *m*/*z* 469.12 (M <sup>−</sup> H)−; *m*/*z* 471.13 (M + H)<sup>+</sup> for the anoxic conditions (Figure 5) and also FA-15NO, *m*/*z* 470.12 (M <sup>−</sup> H)−; *m*/*z* 472.16 (M + H)<sup>+</sup> for the aerobic conditions and *m*/*z* 470.21 (M <sup>−</sup> H)−; *m*/*z* 472.14 (M + H)<sup>+</sup> for the anoxic conditions (Figure 6). The individual mass number of the FA-NO was clearly distinguished by the difference in the isotopic ratio of nitrogen. In addition, the product of FA-NO in the anoxic conditions was similar yield to that of the aerobic conditions. These results supported the assertion that FA has NO scavenging ability, resulting in the suppression of NO production in the LPS-stimulated RAW264.7 cells (Figure 1).

**Figure 5.** LC/MS chromatogram obtained from the reaction mixture containing folic acid (FA), Na2S2O4 and Na14NO2 under the conditions of (**A**) aerobic and (**B**) anoxia. The LC/MS was carried out under the analytical conditions as described in the text. Each peak of LC/MS indicated FA and its reaction product with NO, FA-NO. The HPLC chromatogram indicated FA and its reaction product with NO under the conditions of **A** (**a**) aerobic and **B** (**f**) anoxia. The TIC (total ion chromatogram) and its mass spectra of FA–14NO indicated in the negative and positive ion modes as follows. The aerobic conditions; the TIC of (**b**) and (**c**) for (**d**) *m*/*z*469.17 (M − H)− and (**e**) *m*/*z* 471.14 (M + H)+, and the anoxic conditions; the TIC of (**g**) and (**h**) for (**i**) *m*/*z* 469.12 (M − H)− and (**j**) *m*/*z* 471.13 (M + H)+.

**Figure 6.** LC/MS chromatogram obtained from the reaction mixture containing folic acid (FA), Na2S2O4 and Na15NO2 under the conditions of (**A**) aerobic and (**B**) anoxia . The LC/MS was carried out under the analytical conditions as described in the text. Each peak of LC/MS indicated FA and its reaction product with NO, FA-NO. The HPLC chromatogram indicated FA and its reaction product with NO under conditions that were **A** (**a**) aerobic and **B** (**f**) anoxia. The TIC (total ion chromatogram) and its mass spectra of FA-14NO indicated in the negative and positive ion modes as follows. the aerobic conditions; the TIC of (**b**) and (**c**) for (**d**) *m*/*z* 470.12 (M− H)− and (**e**) *m*/*z* 472.16 (M+ H)+, and the anoxic conditions; the TIC of (**g**) and (**h**) for (**i**) *m*/*z* 470.21 (M− H)− and (j) *m*/*z* 472.14 (M+ H)+.

#### *3.6. FA Scavenging NO by ESR Study*

It is known that nitronyl nitroxide carboxy-PTIO as an NO detection reagent reacts with NO, generate an imino nitroxide, carboxy-PTI radical and NO2 [17,18]. To clarify the NO scavenging action of FA, an ESR study was performed on the system containing the carboxy-PTIO in the presence of NOR3 and FA. The NO released from NOR3 was detected by carboxy-PTIO, then produced a carboxy-PTI radical as indicated by the arrows in Figure 7. The carboxy-PTI radical was reduced when FA was present in the system, indicating that FA can directly scavenge NO.

**Figure 7.** NO savenging activity of folic acid (FA). Electron spin resonance (ESR) spectrum obtained from the reaction mixture containing the carboxy-PTIO (2-phenyl-4,4,5,5-tetramethylimidazoline-1-oxyl-3-oxide, 200 μM) as the NO detection reagent in the presence of NOR3 (200 μM) with or without FA (200 μM). The NO released from NOR3 was detected by carboxy-PTIO, then produced a carboxy-PTI (2-phenyl-4,4,5,5-tetramethylimidazoline-1-oxyl) radical as indicated by the arrows.

#### **4. Discussion**

The significance of FA in the prevention of neural tube defects (NTDs) involving the failure of NT closure in the developing embryo is well recognized. NO had been shown to be able to induce NTDs in rat embryos, and biochemical studies showed that NO inhibits methionine synthase (MS) [6,7]. In a previous study, the direct effect of NO produced by the NO donor SNAP on the process of NT closure in the chick embryo ex ovo. was evaluated and the high NO levels by the SNAP treatment inhibited the MS in the methyl transfer reaction of cofactor B12 [8,9]. The alleviation mechanism due to the FA treatment was speculated to be the interference with one-carbon flow through the folate pathway [8]. Previous studies showed that 5-methyl-tetrahydrofolate (5-mTHF), the primary circulating metabolite of FA, improves the endothelial NO synthase (*e*NOS) coupling cofactor tetrahydrobiopterin (BH4), indicating that the NOS activity regulates the MS activity in the process of NT closure [8]. On the other hand, when the BH4 is limited, the uncoupled NOS produces the superoxide radical (O2 −) rather than NO, then produces peroxynitrite by the reaction between O2 - and NO which causes the endothelial dysfunction [19]. FA has the ability of O2 − scavenging, which may protect or improve the endothelial function [20–23]. However, the detailed relationship between FA and NO has not yet been clarified. In

this study, the effect of FA on the NO production in the LPS-stimulated RAW264.7 cells was examined. The nitrite accumulation in the LPS treatment cells decreased in the presence of FA (Figure 1). An inflammatory cytokine or proinflammatory cytokine including interleukin-1 (IL-1), interleukin-12 (IL-12), and interleukin-18 (IL-18), TNF-α, interferon γ (IFNγ), and granulocyte-macrophage colony stimulating factor (GM-CSF) stimulate the Janus kinase (Jak) and signal transducer and activator of transcription (STAT) pathway (JAK-STAT pathway) as a key role in signal pathways activated by growth factors and cytokines. Expression of *i*NOS and the production of NO in response to LPS/IFN γ are increased through JAK/STAT signaling [24,25]. While the expression of the *i*NOS gene due to the LPS/ IFNγ treatment was not inhibited by the FA treatment, thus the reduction NO due to FA will not be through the JAK-STAT pathway. It is suggested that FA may directly regulate against NO or producing nitrogen radicals during the NO oxidation (Figure 2). The NO-scavenging ability of FA was examined in the reaction with NOR3 as the NO donor. FA suppressed the nitrite accumulation in a dose-dependent manner suggesting that FA provides a direct scavenging ability against NO or nitrogen radicals. The result first showed another possible function of FA which can modulate the nitrogen radicals including NO. Although FA could not suppress the *i*NOS gene expression, the nitrogen radical scavenging function due to FA could suppress for the NO production in the LPS-stimulated cells (Figure 3). To better clarify the function of FA, the FA scavenging NO adduct in the reaction mixture was detected by LC/MS. The mass numbers of *m*/*z* 470.9 (M + H)<sup>+</sup> and *m*/*z* 469.1 (M <sup>−</sup> H)<sup>−</sup> were determined as the adduct of the FA scavenging NO (FA-NO). To obtain more detailed evidence, the FA-NO derived from 14NO and 15NO was measured in both the positive and negative ion mode, and each product of the 14NO or 15NO reaction with FA gave the following mass numbers: FA-14NO, *m*/*z* 471.14 (M <sup>+</sup> H)+; *<sup>m</sup>*/*<sup>z</sup>* 469.17 (M <sup>−</sup> H)<sup>−</sup> and FA-15NO, *<sup>m</sup>*/*<sup>z</sup>* 472.16 (M <sup>+</sup> H)+; *<sup>m</sup>*/*<sup>z</sup>* 470.12 (M <sup>−</sup> H)<sup>−</sup> (Figures 5A and 6A). The individual mass number of the FA-NO was clearly distinguished by the difference in the isotopic ratio of nitrogen, resulting in the product by the FA reaction with NO. NO reacts with oxygen giving rise to the formation of NO2 and potentially N2O3. To avoid the reactions, the FA reaction with NO in the anoxic conditions was examined, resulting in the product of FA-NO being of a similar yield to that of the aerobic conditions (Figures 5B and 6B). This result suggested that the reactions of FA with NO give rise to folic acid nitrosation.

To clarify the direct NO scavenging action due to FA, an ESR study was performed on the system containing carboxy-PTIO as the NO detection reagent in the presence of NOR3 and FA. The carboxy-PTI radical was then produced from the reaction with NO reduced in the presence of FA (Figure 7). This result strongly supported the assertion that FA can directly scavenge NO and the function of FA suppressed the NO production in the LPS-stimulated cells.

FA can prevent the NTDs causing the NO induction which inhibits MS involving the interference transfer of the methyl group from the 5mTHF to homocysteine in methionine production. This study proposes another alleviation mechanism whereby the NO modulation due to the FA direct scavenging of NO contributes to alleviation from the failure in the NT formation causing the high level of NO production.

#### **5. Conclusions**

This study first demonstrated that FA can directly scavenge NO, resulting in the reduction of the accumulation of nitrite in the LPS-stimulated RAW264.7 cells. In addition, the NO modulation due to FA may contribute to alleviation from the failure in neural tube formation causing the high level of NO production.

**Author Contributions:** All authors (T.O. and J.T.) participated in the experiments and research design of the study. J.T. organized this study including interpretation of the data and wrote the manuscript. All authors have read and agreed to the published version of the manuscript.

**Funding:** Works described here were supported by Grants from Okinawa Prefecture, Japan.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


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