Background
Non-insulin dependent diabetes mellitus (NIDDM, Type 2 diabetes) is a group of metabolic diseases marked by high levels of blood glucose resulting from faults in insulin resistance and insulin production [
1‐
3]. Hyperglycemia, a rapid rise in blood glucose levels in NIDDM patients occurs due to hydrolysis of starch by pancreatic α-amylase and absorption of glucose in the small intestine by α-glucosidases such as sucrase and maltase [
3]. Inhibition of these carbohydrate-hydrolyzing enzymes can significantly decrease the postprandial hyperglycemia after a mixed carbohydrate diet and can be a key strategy in the control of diabetes mellitus [
4,
5]. However, a main negative aspect of currently used therapeutic α-glucosidase inhibitors, such as the drug acarbose, is the strong α-amylase inhibitory activity causing abnormal bacterial fermentation of undigested starch in the colon that leads to abdominal distention, flatulence, meteorism and possibly diarrhea [
6,
7]. Recent studies showed that phenolic phytochemicals from plant sources are natural inhibitors of α-amylase and α-glucosidase [
8,
9] Such natural inhibitors from dietary plants could be useful since they have lower inhibitory activity against α-amylase and a stronger inhibitory activity against α-glucosidase and can be potentially used as an effective strategy for the management of postprandial hyperglycemia with minimal side effects [
9].
Zingiber mioga (Zingiberaceae family) is a herb original to eastern Asia. Its young flower buds have been used as a traditional food in Asia [
10]. Recent research has reported the antimicrobial activities of the constituents of
Zingiber mioga against several strains of bacteria, yeast, and mold [
11,
12]. The volatile ingredients of
Zingiber mioga have been studied; 2-isopropyl-3-methoxypyrazine, 2-sec-butyl-3-methoxypyrazine, and 2-isobutyl-3-methoxypyrazine were found to be the aroma compounds by GC-MS [
1]. Additionally, mioga extract was effective in inhibiting fat accumulation in 3 T3-L1 adipocytes leading to a decrease in body weight gain and a decrease in fat mass in ICR mice [
2]. However, the significance of
Zingiber mioga intake for preventing diabetes-related oxidative stress and hyperglycemia is not reported. Based on the previous obesity related findings, it is interesting to first evaluate the in vitro potential of mioga extracts again carbohydrate hydrolyzing enzyme using in vitro models and if inhibitory effect is observed, then the extracts should be evaluated using an animal model.
Therefore, the aim of this study is to examine the potential effect and mechanism of action of Zingiber mioga extract on the inhibition of postprandial hyperglycemia using both in vitro and in vivo animal models. Clear knowledge of the activity and mode of action of Zingiber mioga extract will contribute towards better understanding of the actual effect of various Zingiber mioga products towards type 2 diabetes management. To determine the above, in this study, we (i) prepared Zingiber mioga extracts (water extract of Zingiber mioga: ZMW, ethanol extract of Zingiber mioga: ZME); (ii) investigated the inhibitory activity of ZMW, ZME against α-amylase and α-glucosidase (anti-hyperglycemia potential); (iii) measured antioxidant potential using oxygen radical scavenging capacity (ORAC) assay; and (iv) evaluated the postprandial blood glucose lowering effect of ZMW, ZME after sucrose loading in a Sprague-Dawley (SD) rat and db/db mice model.
Methods
Chemicals
Zingiber mioga was obtained from a local market in Jeju, Korea. The purchased samples were identified by one of the authors (Young-In Kwon). A voucher specimen (BFC O10985) was deposited at the Bioactive Food Components Lab (BFCL) of the College of Life Science and Nano Technology, Hannam University. Rat intestinal acetone powder, porcine pancreatic α-amylase enzyme powder, starch, sucrose, and maltose were purchased from Sigma-aldrich (St. Luis, MO, USA). Unless noted, all chemicals were purchased from Sigma-aldrich (St. Luis, MO, USA).
Preparation of Zingiber mioga extracts
Zingiber mioga was crushed to a fine powder and was extracted by autoclaving the ground leaves at 121 °C for 15 min with one gram-fresh weight per 40 mL of distilled water. Zingiber mioga extract was then centrifuged at 8000 x g for 30 min, filtered through a Whatman # 1 filter, vacuum-evaporated at 60 °C, freeze dried and kept at -20 °C refrigerator until analysis.
Zingiber mioga was crushed to a fine powder and was extracted by Shaking Incubation at 40 °C for 2 h with one gram-fresh weight per 40 mL of ethanol. Zingiber mioga extract was then centrifuged at 8000 x g for 30 min, filtered through a Whatman # 1 filter, vacuum-evaporated at 60 °C, freeze dried and kept at -20 °C refrigerator until analysis.
Carbohydrate hydrolyzing enzyme inhibition
Porcine pancreatic α–amylase inhibition assay
Porcine pancreatic α-amylase inhibition was determined by the method described by Kwon et al [
3]. Sample solution (200 μL) and 0.02 M sodium phosphate buffer (pH 6.9 with 0.006 M sodium chloride, 500 μL) containing α-amylase solution (0.5 mg/mL, 5.0 MU/mL) were incubated at 25 °C for 10 min. After pre-incubation, 500 μL of a 1 % starch solution in 0.02 M sodium phosphate buffer was added. The reaction mixture was then incubated at 25 °C for 10 min. The reaction was stopped with 1.0 mL of dinitrosalicylic acid (DNS). The reaction mixture was then incubated in a boiling water bath for 5 min and cooled to room temperature. The reaction mixture was then diluted after adding distilled water, and absorbance was measured at 540 nm with ELISA microplate reader (SUNRISE; Tecan Trading AG, Saltzburg, Austria).
$$ \%\ \mathrm{inhibition} = \left(\left[\frac{\Delta {A}_{540}^{Control}-\Delta {A}_{540}^{Extract}}{\left[\Delta {A}_{540}^{Control}\right]}\right]\right)x100 $$
Sucrase and Maltase inhibition assay
The crude enzyme solution prepared from rat intestinal acetone powder Sigma-Aldrich Co. (St. Louis, MO, USA) was used as the small intestinal maltase, sucrase, and glucoamylase, showing specific activities of 0.70, 0.34 and 0.45 units/mL, respectively. Rat intestinal acetone powder (1.0 g) was suspended in 3 mL of 0.9 % saline, and the suspension was sonicated twelve times for 30 s at 4 °C. After centrifugation (10,000 × g, 30 min, 4 °C), the resulting supernatant was used for the assay. Sucrase and maltase inhibitory activity were assayed by modifying a method developed by Dahlqvist (1964) [
4]. The inhibitory activity was determined by incubating a solution of an enzyme (50 μL), 0.1 M phosphate buffer (pH 7.0, 100 μL) containing 0.4 mg/mL sucrose or maltose or 1 % soluble starch, and a solution (50 μL) with various concentrations of sample solution (between 0.05 mM and 1.0 mM) at 37 °C for 30 min. The reaction mixture was heated in a boiling water bath to stop the reaction for 10 min, and then the amount of liberated glucose was measured by the glucose oxidase method [
5]. The inhibitory activity was calculated from the formula as follows. Inhibition (%) = (C-T)/C x 100, where C is the enzyme activity without inhibitor and T is the enzyme activity with inhibitor.
Oxygen radical absorbance capacity (ORAC) assay
The peroxyl radical-scavenging capacities of
Zingiber mioga extracts were measured using ORAC [
6,
7]. The ORAC assay was carried out using a Tecan GENios multi-functional plate reader (GENios; Tecan Trading AG, Salzburg, Austria) with fluorescent filters (excitation wavelength: 485 nm, emission filter: 535 nm). In the final assay mixture, fluorescein (40 nM) was used as a target of free radical attack with either 2, 2′-azobis (2-amidinopropane) dihydrochloride (AAPH, 20 mM) as a peroxyl radical generator in peroxyl radical-scavenging capacity (ORAC
ROO·) assay or with H
2O
2 - CuSO
4 (H
2O
2, 0.75 %; CuSO
4, 5 μM) as a hydroxyl radical generator in hydroxyl radical-scavenging capacity (ORAC
HO·) assay. Trolox (1 μM) was used as a control standard and prepared fresh on a daily basis. The analyzer was programmed to record the fluorescence of fluorescein every 2 min after AAPH or H
2O
2 - CuSO
4 was added. All fluorescence measurements were expressed relative to the initial reading. Final results were calculated based on the difference in the area under the fluorescence decay curve between the blank and each sample. All data were expressed as micromoles of Trolox equivalents (TE). One ORAC unit is equivalent to the net protection area provided by 1 μM of Trolox.
Total phenolic content assay
The total phenolic content was determined by an assay modified from Velioglu et al. [
8,
9]. One milliter of sample solution was transferred into a test tube and mixed with 1 mL of 95 % ethanol and 5 mL of distilled water. To each sample 0.5 mL of 50 % (v/v) Folin-ciocalteu reagent was added and mixed. After 5 min, 1 mL of 5 % Na
2CO
3 was added to the reaction mixture and allowed to stand for 1 h. The absorbance was read at 725 nm using spectrophotometer (UV-160A; Shimadzu Inc., Kyoto, Japan). The absorbance values were converted to total phenolics and were expressed in mg equivalents of gallic acid/mL of the sample. Standard curves were established using various concentrations of gallic acid in 95 % ethanol.
Animal and study design
The inhibitory effect of Zingiber mioga extacts and acarbose on postprandial hyperglycemia after carbohydrate loads in Sprague-Dawley (SD) rats and C57BL/KsJ-db/db (db/db) mice were evaluated. The experimental protocols followed were approved by the Institutional Animal Care and Use Committee (IACUC) of the Hannam University (Approval number: HNU2015-0004). Five week-old male SD rats and C57BL/KsJ-db/db (db/db) mice were purchased from Joongang Experimental Animal Co. (Seoul, Korea) and fed a solid diet (Samyang Diet Co., Seoul, Korea) for 1 week. The rats and mice were housed in a ventilated room at 25 ± 2 °C with 50 ± 7 % relative humidity, and under an alternating 12 h light/dark cycle. After 4 groups of 5 male SD rats and db/db mice were fasted for 24 h, 2.0 g/kg of sucrose were orally administrated concurrently with 0 ~ 100 mg/kg inhibitors (Zingiber mioga extracts or Acarbose). The blood samples were then taken from the tail after administration and blood glucose levels were measured at 0, 0.5, 1, and 2 h. The glucose level in blood was determined by glucose oxidase method and compared with that of the control group, which had not taken the inhibitors.
Statistical analysis
All data are presented as mean ± Standard deviation (SD). Statistical analysis was carried out using statistical package SPSS 10 (Statistical Package for Social Science; SPSS Inc., Chicago, IL, USA) program and significance of each group was verified with the analysis of on-way analysis of variance (ANOVA) followed by the Duncan’s test of p < 0.05.
Conclusions
A sudden increase in blood glucose levels, causing hyperglycemia in NIDDM is due to insulin’s inability to effectively utilize the glucose produced by the hydrolysis of dietary carbohydrates by carbohydrate hydrolyzing enzymes. Furthermore, hyperglycemia-induced microvascular complications are likely from oxidative dysfunction from mitochondrial reactive oxygen species (ROS). Therefore, it is important to control both cellular redox status and blood glucose level for managing these diabetic complications. The ethanol extract of
Zingiber mioga has α-glucosidase inhibitory activity and potent peroxyl radical scavenging-linked antioxidant activity. Sucrose loading test showed that ZME may reduce postprandial increases of blood glucose level by acting as an intestinal α-glucosidase inhibitor. The above dual benefits (post-prandial blood glucose reduction and antioxidant activity) of ZME could support the evidence that diet rich in fruits and vegetables are associated with lower incidences of oxidation-linked diseases such as diabetes [
15‐
17]. Furthermore, the
Zingiber mioga extracts showed a strong inhibition of α-glucosidase and low inhibition of α-amylase and therefore could be potentially used as an effective complementary therapy for postprandial hyperglycemia linked to type2 diabetes with reduced side effects [
18]. It is well documented that the major side-effects of carbohydrate hydrolyzing enzyme inhibition are flatulence and diarrhea resulting from the high α-amylase inhibitory activity [
19,
20].
These in vitro and in vivo studies provide the biochemical rationale for the potential benefit of Zingiber mioga for carbohydrate hydrolyzing enzyme inhibition, with ZME appearing to be more bioactive. This observed higher efficacy of ZME can be further enhanced by identifying the bioactive components responsible for the observed activity. Further investigation is underway to identify the specific phenolic compounds in Zingiber mioga extraction that are relevant to the inhibition of carbohydrate hydrolysis enzymes.
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Competing interests
The authors declare that they have no competing interests.
Authors’ contributions
SHJ and CYC conducted the animal experiment and analyzed the data. JYL and KSH prepared the product to be tested. EMA and YIK participated in design of the study and preparation of the manuscript. All the authors read and approved the final manuscript.