Antioxidant, antibacterial, anti-inflammatory, and anticancer properties of Cinnamomum kanehirae Hayata leaves extracts

2.3.1 Extraction of samples

The samples were mixed with solvents (50, 80, 95 % ethanol, and pure water) in a ratio of 1:50, respectively, followed by homogenization, and extracted by stirring at 25 °C for 1 h. The residue was filtered, and the same volume of extraction solvent was added for 1 h. The two extracts were combined, filtered, and concentrated at 45 °C. Then, a portion of the concentrate was freeze-dried to obtain the 4 CKHLE powders with water, 50, 80, and 95 % ethanol extraction, which was further used for cell culture and COX-2 activity determination. The concentrated extracts and the dried extract powder were stored at −20 °C until used.

2.3.2 Determination of total polyphenol content (TPC)

TPC measurements were performed by the agreement of Huang et al., (2022) and (Marsoul et al., 2020) with minor modifications. In brief, the phenolics were reduced with the Folin-Ciocalteu reagent to form a blue molybdenum-tungsten complex with the highest absorption peak at wavelength 750 nm. Each of the samples and Folin-Ciocalteu reagent was placed in a microtube with 100 μL, then shaken uniformly and standing for 2 min. Then, 2 mL of 2 % Na₂CO₃ was added with a reaction of 30 min at room temperature, followed by measured absorbance values at wavelength 750 nm. For the blank group, the Folin-Ciocalteu reagent was replaced by 2 % Na₂CO₃ and followed the same protocol. Standard curves (r² = 0.9997) were prepared as gallic acid. The above absorbance values were calculated using the following equation to calculate the samples' TPC. The results were expressed as gallic acid equivalents (mg GAE/g extract). $$\mathit{TPC}(mgGAE/mLextract)=A750nm-0.0668/0.0034$$

2.3.3 Determination of total flavonoid content (TFC)

The measurement was carried out based on the properties of strong absorbance at wavelengths of 415 nm measured by the derivatization of flavan and flavonol with aluminum chloride in the flavonoid structure. It was performed by the protocols of Huang et al., (2022) and Marsoul et al., (2020) with minor modifications. Briefly, 0.5 mL of the sample was added to 1.5 mL of 95 % ethanol, 0.1 mL of 10 % AlCl₃ 6H₂O, 0.1 mL of 1 M potassium acetate, and 2.8 mL of deionized water, then shaken until uniformly mixed. After 40 min reaction at room temperature, absorbance values were measured at wavelengths of 415 nm. The standard curve (r² = 0.9959) was prepared for the quercetin, and the obtained equation was used to calculate the TFC of the samples. The results were expressed as µg quercetin equivalent/g fresh weight (µg QE/g fresh weigh) $$\mathit{TFC}(\mathrm{\mu }\mathrm{g}\mathrm{Q}\mathrm{E}/\mathrm{g}\mathrm{f}\mathrm{r}\mathrm{e}\mathrm{s}\mathrm{h}\mathrm{w}\mathrm{e}\mathrm{i}\mathrm{g}\mathrm{h}\mathrm{t})=A415nm+0.1471/0.01209$$

2.3.4 Determination of safrole content

The method described in Reynertson et al., (2005) has been modified as appropriate. The sample mentioned above (Section 2.4.1), filtered by a 0.45 μm membrane, 20 μL, was taken and analyzed by high-performance liquid chromatography (HPLC, Hitachi, Ltd., Tokyo, Japan) with a Chromaster 5410 UV (Hitachi) detector and an RP-C18 GP (250 mm × 4.6 mm × 5 μm) column (Kanto Chemical Co., Tokyo, Japan). The wavelength was detected at 235 nm, while the retention time and peak area of the samples were analyzed using a Chromaster (Hitachi). The mobile phase was ultrapure water, acetonitrile, and methanol at a 1 mL/min flow rate. The elution gradients were as follows: 0–10 min: ultrapure water (55), acetonitrile (45); 10–30 min: ultrapure water (50), acetonitrile (45). (50), acetonitrile (50); 30–34 min methanol (1 0 0); 34–35 min: acetonitrile (15), methanol (85); 35–36 min: acetonitrile (1 0 0). Safrole standards (CAS: 94–59-7, ChemService Inc., PA., USA) were prepared in methanol at concentrations of 0.0025–1.25 μg/mL, with a standard curve (r² = 0.9985), and the obtained equation was used to calculate the concentration of safrole content in samples expressed as µg/mL.

2.4.1 Determination of DPPH radical scavenging capacity

2,2-diphenyl-1-picrylhydrazyl, also known as DPPH, has a dark purple appearance and as a stable radical with a maximum absorption peak at wavelengths 515–520 nm (Carmona-Jiménez et al., 2014, Hidayat et al., 2017), it contains the odd number of electrons and receives hydrogen protons (H⁺) from antioxidants to form more stable DPPH-H molecules (Lee et al., 2017). The method was performed with modification according to Huang et al., (2011). Various concentrations of CKHLE and gallic acid (as the control group) were taken 100 μL for each. Then, 400 μL of 100 mM Tris-HCl buffer and 500 μL of 250 μM DPPH were shaken for 20 sec. A spectrophotometer measured the absorbance values at 517 nm. For the blank group, 250 μM DPPH was replaced by 100 mM Tris-HCl buffer, and the DPPH radical scavenging rate was calculated as follows. $$\mathit{DPPH}\mathit{radical}\mathit{scavengin}\mathit{gactivity}(\%)=\left(1-\frac{A_{517nm}\mathrm{o}\mathrm{f}\mathrm{s}\mathrm{a}\mathrm{m}\mathrm{p}\mathrm{l}\mathrm{e}}{A_{517nm}\mathrm{o}\mathrm{f}\mathrm{b}\mathrm{l}\mathrm{a}\mathrm{n}\mathrm{k}}\right)\times 100$$

2.4.2 Determination of reducing power

The Fe ions were reduced to Fe²⁺ potassium hexacyanoferrate by the action of antioxidants at pH 6.6, and a Prussian blue compound formed by the action of Fe³⁺ in ferric chloride solution. Thereby, it was possible to detect yellow blood salts at a wavelength of 700 nm to indicate the reducing power of the sample. The method was performed with modification according to Huang et al., (2011). The samples and gallic acid of different concentrations were each taken at 150 μL, added 150 μL of 0.2 M sodium phosphate buffer and 1 % K₃Fe(CN)₆, mixed in a water bath (50 °C) for 20 min, cooled in an ice bath for 5 min, then added 150 μL of 10 % TCA, 0.6 mL of deionized water and 20 μL of 0.1 % Fe(CN)₆ solution in order. The reaction was incubated for 14 min, and the absorbance value was measured at 700 nm using a spectrophotometer, whereas the reducing power was calculated as follows. $$\mathrm{R}\mathrm{e}\mathrm{d}\mathrm{u}\mathrm{c}\mathrm{i}\mathrm{n}\mathrm{g}\mathrm{p}\mathrm{o}\mathrm{w}\mathrm{e}\mathrm{r}=\mathrm{A}700\mathrm{n}\mathrm{m}\mathrm{o}\mathrm{f}\mathrm{t}\mathrm{h}\mathrm{e}\mathrm{s}\mathrm{a}\mathrm{m}\mathrm{p}\mathrm{l}\mathrm{e}-\mathrm{A}700\mathrm{n}\mathrm{m}\mathrm{o}\mathrm{f}\mathrm{t}\mathrm{h}\mathrm{e}blank)$$

2.4.3 Determination of ABTS free radical scavenging ability

2,2′-azino-bis (3-ethylbenzothiazoline-6-sulfonic acid), called ABTS, has been recognized as one of the most sensitive techniques for identifying antioxidant activity due to the fast reaction kinetics involved in its reaction with antioxidants (Chanput et al., 2016). The ABTS interacted with potassium persulfate to form a blue-green chromophore with maximum absorption waves at 645, 734, and 815 nm, respectively, which was measured by spectrophotometer at the above wavelengths to calculate the antioxidant activity of the test substance (Re et al., 1999, Lee et al., 2016).

The ABTS reaction solution was prepared by mixed 5 mL of 7 mM ABTS with 89 μL of 140 mM potassium persulfate, and the reaction was carried out in a dark room for 12–16 h. The reaction solution was diluted with 95% ethanol to give an absorbance value of 0.7 ± 0.2 at 734 nm for subsequent analysis.

The sample was mixed with 990 μL of the above ABTS solution for 6 min at room temperature, and the absorbance values were measured at 734 nm. The blank group was replaced by deionized water, and the dibutyl hydroxytoluene [2,6-bis (1,1-methyl ethyl)-4-methyl phenol, butylated hydroxytoluene, BHT] standard was used as the control group. The following formula calculated the ABTS radical scavenging ability. $$\mathrm{A}\mathrm{B}\mathrm{T}\mathrm{S}\mathrm{r}\mathrm{a}\mathrm{d}\mathrm{i}\mathrm{c}\mathrm{a}\mathrm{l}\mathrm{s}\mathrm{c}\mathrm{a}\mathrm{v}\mathrm{e}\mathrm{n}\mathrm{g}\mathrm{i}\mathrm{n}\mathrm{g}\mathrm{a}\mathrm{b}\mathrm{i}\mathrm{l}\mathrm{i}\mathrm{t}\mathrm{y}(\%)=1-\frac{{\mathrm{A}}_{734nm}\mathrm{o}\mathrm{f}\mathrm{s}\mathrm{a}\mathrm{m}\mathrm{p}\mathrm{l}\mathrm{e}}{{\mathrm{A}}_{734nm}\mathrm{o}\mathrm{f}\mathrm{b}\mathrm{l}\mathrm{a}\mathrm{m}\mathrm{k}}\times 100$$

2.4.4 Determination of ferric reducing antioxidant power (FRAP)

FRAP was used to maintain the solubility of Fe at pH 3.6 and to determine the ability of the sample to reduce Fe³⁺ to Fe²⁺. As opposed to common antioxidant assays, the mechanism of FRAP involves the transfer of electrons rather than hydrogen atoms. While the Fe³⁺ reduced to Fe²⁺ in the presence of 2,4,6-try pyridyl-s-triazine (TPTZ) and formed colored compounds with strong absorbance values at 593 nm (Prior et al., 2005, Cerretani and Bendini 2010). The working reagent was prepared by mixing 1 L of 300 mM acetate buffer (pH 3.6), 10 mL of 10 mM TPTZ colorant, and 1 L of 20 mM FeCl₃ 6H₂O in the ratio of 10:1:1 and prepared in a water bath at 37 °C. A 100 μL sample was mixed with 900 μL of working reagent and protected from light for 30 min, followed by determining absorbance at 593 nm. Ascorbic acid was used as the control group. The standard curve (R² = 0.9935) was prepared with different concentrations of ferrous sulfate (FeSO₄) as the standard. The equation obtained by repeating the above steps was used to calculate the samples' ferrous sulfate equivalent (µmol FeSO₄/g).

2.4.5 Determination of oxygen radical absorbance capacity (ORAC)

ORAC was widely used to evaluate the antioxidant capacity of food products. The analysis was performed by AAPH [2,2′-azobis (2-amidinopropane) dihydrochloride] radical, which formed peroxyl radicals when heated with sufficient oxygen and accelerated fluorescence degradation. Therefore, the antioxidant capacity of the samples could be evaluated by the change in fluorescence (Skendi 2021). The 96-well plate was incubated in a water bath at 37 ℃ for 5 min, followed by the orderly addition of 50 μL of 78 nM fluorescein sodium salt for each well, 50 μL of the sample, and 25 μL of 221 mM AAPH and shaken for 10 s. The fluorescence values were recorded every 5 min at excitation of 485 nm and emission of 520 nm for 90 min. The area under a sample and blank curve were obtained by plotting the fluorescence value and time. The equation below obtained the area under the net curves (Net AUC). Trolox was used as the standard and obtained the standard curves for linear regression to calculate the equivalent of the antioxidant power of the sample. $$\mathit{Areaunderthenetcurve}(NetAUC)={\mathit{AUC}}_{\mathit{test}}-{\mathit{AUC}}_{\mathit{blank}}$$

2.5.1 Activated strains

The method was performed with modification according to Balouiri et al., (2016). The frozen Candida albicans was thawed by taking 1 mL and adding 9 mL of yeast mold broth (YM broth) liquid medium, while another 1 loop was taken and inoculated in yeast mold agar (YM agar) by scribing with a flat plate, both were incubated at 25 °C for 24 h to confirm the purity and activation of the strain. Both E. coli and B. cereus were thawed, and 1 mL of each was added to 9 mL of nutrient broth liquid medium; simultaneously, each 1 loop of bacterial broth was inoculated in nutrient agar (NA) with flat plate scribing, then incubated at 37 °C (to E. coli) and 30 ℃ (for B. cereus) for 24 h to observe the purity and activation of the strains.

2.5.2 Determination of inhibition zone

Using the Kirby-Bauer test, the concentration of the activated bacteria was prepared with sterile water, and the absorbance value at 660 nm was set to 0.1. 100 μL of each bacteria broth evenly spread on YM or NA agar plates. Then, put 8 mm diameter sterile filter paper on the plate. Add 10 μL of the sample to the paper and stand for 1 h. Cultivate the organisms at the optimal growth temperature (as described in the previous section) for 24 h and then measure the diameter of the inhibition zone. Suppose the diameter over 10 mm means it can inhibit bacteria, and the detailed definition as follows: 10 mm as slightly active, 11–15 mm as moderately active, 16–20 mm as highly active.

2.7.1 Determination of cell survival rate

The MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide) has a yellow color and acts on the respiratory chain of the mitochondrion of living cells. By the action of succinate dehydrogenase (SDH) and cytochrome C, the tetrazolium structures were changed to form formazan crystals produced was proportional to cells' survival, and cells' capacity to restore MTT (formazan contents) was known by the absorbance value, which represents the activity of mitochondrion and survival of cells. Thus MTT assay served as an indicator of cell viability (Gerlier and Thomasset 1986). The 96-well plate was incubated with 1 × 10⁵ cells/mL per well and incubated for 24 h in an incubator (5 % CO₂, 37 ℃). After removing the culture medium, 100 μL of various concentrations with CKHLE was given to each well, and DMEM served as the control group. ELISA reader measured the absorbance value at 595 nm, which was used to calculate the cell viability (%) by the equation below. $$\mathit{CellSurvivalRate}(\%)=Absorbanceat595nm(sample/controlgroup)\times 100$$

2.7.2 Determination of nitric oxide production

Nitric oxide (NO) is an osmotic gas produced by L-arginine through the action of nitric oxide synthase. As cells initiate inflammation, the activity of inducible NOS (iNOS) rapidly rises, and a significant NO radical will be produced. NO is an unstable molecule that rapidly oxidizes to the stable nitrite (NO^2–). The reaction of NO^2– with sulfanilamide and naphthyl ethylenediamine dihydrochloride in Griess-reagent was measured to quantify the NO content. The number of cells in each well of the 96-well plate was 1 × 10⁴ cells/mL and incubated for 24 h (5 % CO₂, 37 ℃). Then, remove the DMEM, give 100 μL of various concentrations of CKHLE and 0.1 μg/mL LPS, respectively, and cultivate for 24 h. Take 100 μL of supernatant in a 96-well plate, add 100 μL of Griess-reagent, protect the reaction from light for 10 min, and measure the absorbance value at 540 nm with an ELISA reader. The amount of NO production was calculated with NaNO₂ prepared as the standard curve (R² = 0.9997).

3.2.1 TPC

Polyphenols are an essential source of antioxidants in the daily diet. However, most of the polyphenols have not been absorbed in the stomach. They were accumulated in the colon and utilized by gut microorganisms, which indicates that polyphenols might be affected by intestinal health. In vitro and in vivo studies have shown that polyphenolic compounds have physiological activities such as antibacterial, anti-inflammatory, diabetes prevention, and inhibition of lipid formation (Wan et al., 2021). The TPC of the 4 CKHLEs (with water, 50, 80, and 95 % ethanol) were shown in Table 2, in which the CKHLE extracted with 50 % ethanol [a mixture of water/ethanol (50:50) v/v] had the highest TPC of 16.98 ± 0.18 mg GAE/g fresh weight). The others were 80% ethanol, water, and 95% ethanol extracted from highest to lowest, with approximately 9.34, 8.20, and 2.78 mg GAE/g fresh weight, respectively, which showed significant differences (p < 0.05). Proestos and Komaitis (2009) analyzed the TPC of 50% ethanol extracts from 5 Cinnamon species, and the results ranged from 0.69 to 2.71 mg GAE/g, with the highest TPC in Cinnamomum zeylanica species. Dvorackova et al., (2015) showed that 95 % of ethanol extracts contained the highest TPC, indicating that the extraction solvent's polarity would affect the TPC in the obtained extracts.

3.2.2 TFC

Flavonoids were one of the common active plant substances, namely quercetin, anthocyanidins, apigenin, and luteolin. Many studies have confirmed flavonoids' antioxidant, anti-inflammatory, and other physiological effects (Prasad et al., 2009, Saleem et al., 2010). This study showed that CKHLE extracted with 50% ethanol had the highest TFC content of 1.90 µg QE/g fresh weight, followed by 80% ethanol extraction with 1.16 (µg QE/g fresh weight). In comparison, 95% ethanol extract had even lower than water extraction (Table 2), all with significant differences (p < 0.05). Prasad et al., (2009) showed that 50% ethanol extracts of five species of Camphor were obtained, with the highest TFC of C. burmanni leaves (2738.4 μg QE/g), and the remaining four species ranged from 568.1 to 1564.4 μg QE/g. Przygodzka et al., (2014) reported significantly higher TFC for 50% of star anise and cinnamon ethanol extracts than absolute ethanol extracts. Still, no significant differences were found for other spices in both extract solvents.

3.2.3 Safrole content

It has been shown in animal models that safrole was reproductive, genotoxic, and carcinogenic. Therefore, governments have established regulations to reduce the risks of human exposure to safrole. The United States Food and Drug Administration (USFDA) banned safrole and safrole oil in food by allowing only edible spices with traces of natural safrole (Gad and Pham 2014); Food and Agriculture Organization of the United Nations/World Health Organization Joint Expert Committee on Food Additives (FAO/WHO JECFA) The Joint Expert Committee on Food Additives (FAO/WHO JECFA) recommends that the maximum daily dietary intake of safrole is about 14.6 μg/Kg; according to the “Scope and Limits of Use and Specification Standards for Food Additives” established by the Ministry of Health and Welfare of Taiwan, safrole classifies as aromatic alcohols in Class (II) spices. It has only used in beverages, with a limit of 1.0 mg/Kg. Therefore, CKHLE contained naturally occurring safrole. Thus, the safrole content of CKHLE was analyzed to evaluate the safety of consumption. In this study, safrole had not detected in 50% ethanol and water CKHLE, while in 80% and 95% ethanol extracts, safrole levels were 0.45 ± 0.05 ppm and 0.12 ± 0.07 ppm, respectively (Table 2), both of which were lower than the limits for food. Yet, whether CKHLE contains physiological toxicity has to be thoroughly evaluated in further animal and human trials.

However, the polarity of the extraction solvents varied the safrole content, while Saetan et al., (2017) showed that the safrole content of extracts from Cinnamomum porrectum leaves blanching at 100 °C for 60 s combined with thermal air drying decreased by 82% and 89% compared to freezing or thermal air drying, respectively; the blanching process effectively dissolved most of the safrole, while there was no significant difference in the safrole content of the other two dry-treated extracts.

3.3.1 DPPH free radical scavenging capacity

The DPPH radical scavenging assay is one of the longest-established antioxidant assays (Kumar et al., 2018). DPPH radicals dissolved in ethanol are color-stable. DPPH will change from a blue-purple compound to a yellow color when subjected to antioxidants. However, the higher ratio of absorbance value fading, the stronger the antioxidant capacity of the sample. This study investigated the effect of 4 CKHLEs (with water, 50, 80, and 95 % ethanol) on the free scavenging ability of DPPH, which were expressed as half-maximal effect concentration (EC50) (Table 3). However, low levels of EC50 imply that the sample has a higher antioxidant capacity. The EC50 of CKHLE with 80% and 50% ethanol extracts were 0.96 and 0.51 mg/mL, respectively, significantly different from the other two CKHLEs (p < 0.05). Unfortunately, there were still larger gaps when compared to gallic acid.

3.3.2 Reducing power

In this study, the EC50 of 4 CKHLEs (with water, 50, 80, and 95 % ethanol) was found that 50% ethanol extracted EC50 (1.14 ± 0.07 mg/mL) was the lowest among the 4 CKHLE, which indicated that it had that best-reducing power, followed by 80% ethanol CKHLE (1.83 ± 0.03 mg/mL) (Table 3). There was a significant difference (p < 0.05). Mathew and Abraham (2006) reported that methanol extraction of Cinnamomum verum bark was dose-dependent, and the reducing power of the extract was 2.73 at a concentration of 1 mg/mL, which indicated that it could act as an electron donor and react with free radicals to form more stable compounds and abort the oxidation reaction. Shimada et al., (1992) showed that the reduction power of Ceylon cinnamon methanol extract was due to the presence of di- and mono-hydroxy substituents in the aromatic ring, which gave it a strong ability to provide hydrogen. As known, camphor plant extracts were primarily composed of aromatic compounds. Thus, it was assumed that the reducing power of CKHLE in this study might probably arise from the same mechanism. However, identifying the active components were required to confirm the critical factor of antioxidant activity.

3.3.3 ABTS⁺ radical scavenging capacity

The ABTS⁺ radical scavenging capacity of the 4 CKHLEs in this study showed (Table 3) that 50% ethanol extraction had the lowest EC50 of about 3.80 mg/mL, followed by water and 80% ethanol extraction in that order. Still, there was no significant difference between the three. Unfortunately, the EC50 of 95% ethanol extraction was 18.00 mg/mL, about 6 folds higher than that of 50%. Hence, it was concluded that 50% ethanol extraction had the best ABTS⁺ radical scavenging capacity. Mathew and Abraham (2006) showed that the ABTS⁺ radical scavenging capacity of 50% ethanol extract of Ceylon cinnamon increased steadily with the extract concentration from 6.25 to 25 μg/mL, followed by stabilization.

3.3.4 FRAP

FRAP was a simple, rapid, and reproducible antioxidant assay initially used to determine the antioxidant capacity of human plasma but subsequently used to assess the biological activity of dietary components (Sveinsdottir et al., 2014, Kumar et al., 2018). In this study, the FRAP of the 4 types of CKHLE was investigated, and it showed that the FRAP of 50% ethanol was the highest, about 376.57 mmol FeSO₄/g, which was still significantly lower than that of ascorbic acid (p < 0.05) (Table 3). The FRAP of other extracts ranked high to low for 80 % ethanol, pure water, and 95 % ethanol, respectively. Abeysekera et al., (2013) extracted leaves and branches of Cinnamomum zeylanicum with 95% ethanol at FRAP of 125.71 ± 3.21 mmol FeSO₄/g leaves, 73.02 ± 2.81 mmol FeSO₄/g bark.

FRAP was affected by the degree of hydroxylation and confocalization of phenolic structures (flavonoid family) in antioxidants, especially when specific forms were met, e.g., the presence of 3′, 4′-dihydroxy structures in the B ring, the presence of 2, 3-double bonds connected to 4-carbonyl groups in the heterocyclic ring, and the formation of confocalization between A and B rings with 3- or 5-hydroxy groups; the A and C rings share a common carbonyl group (Hodnick et al., 1988, Bors et al., 1990, Rice-Evans et al., 1996). Hence, in this study, the FRAP of CKHLEs was correlated with TPC, which agrees with the reports of Li et al., (2017) and Yu et al., (2021); alternatively, the ortho-diphenols and tannins contents in plants were correlated with FRAP.

3.3.5 ORAC

Since different antioxidant assays were based on various sources of oxidation, their mechanisms of action were other, and the comparisons were difficult to compare directly. However some scholars believe that the mechanism of action of the ORAC method was associated with the antioxidant response in organisms; thus, the ORAC method has been considered a preferred antioxidant assay model (Chan-Blanco et al., 2007). In this study (Table 3), the ORAC values of CKHLE with 80% ethanol and water extracts were observed as 5334.51 and 5318.58 μmol TE/g sample, respectively, with no statistically significant difference between them, yet significantly higher than the other two types of CKHLE (p < 0.05). Dudonné et al., (2009) reported that the ORAC values of Eucalyptus globulus leaves and Cinnamomum zeylanicum bark extracts were 8515 and 2846 μmol Trolox/g, respectively, while the results of this study were between them. However, compared to water extracts, ORAC activity of CKHLE was superior to that of Ceylon cinnamon bark extract with the same tree genus. Hence, it indicated that CKHLE was a potential material as an antioxidant.

3.4.1 Inhibition of the bacterial activity

This study investigated the growth inhibition of C. albicans, E. coli, B. cereus by 4 CKHLEs, and the inhibition of B. cereus by 95% ethanol extract was shown as moderately active for all three strains, in which the inhibition of C. cactus was the highest, with an inhibition zone of 13 mm in diameter (Table 4). However, the 80 % and 50 % ethanol extracts were slightly active against B. cereus but failed to inhibit the growth of C. albicans and E. coli. The CKHLE extracted with water showed an inhibition zone for B. cereus. Nevertheless, the diameter of the inhibition zone did not reach the standard of inhibition activity (10 mm), while the other two strains were not inhibited. Overall, it was found that 95% ethanol extract was able to extract more antibacterial active substances from CKHL. In contrast, other CKHLE were insufficient to provide antibacterial activity, probably caused by the low concentration tested or the absence of antibacterial components. Cinnamaldehyde and eugenol from Cinnamomum burmannii have been observed to inhibit bacteria by blocking the production of β-lactamase and breaking the cell wall of bacteria (Hodnick et al., 1988). Shan et al., (2007) prepared Cinnamomum burmannii Blume with 80% methanol. The concentration of branch extract at 100 g/mL showed good inhibition ability of B. cereus, Listeria monocytogenes, S. aureus, E. coli, Salmonella anatum, especially in B. cereus and S. aureus, where the diameter of the inhibition zone reached moderately active, 15.4 ± 0.3 and 15.7 ± 0.4 mm, respectively.

3.4.2 Anti-inflammatory capacity

3.4.2.3

3.4.2.3 Nitric oxide production

Lipopolysaccharide (LPS) has been commonly used as an inducer in vitro inflammatory assays, stimulating the activation of MAPK and NFκB pathways in macrophages and promoting the production of inflammatory factors such as TNF-α, IL-6, COX-2, and iNOS (Sohn et al., 2012, Huang et al., 2014). As LPS inflamed Raw 264.7 cells, iNOS production was increased, which caused the cells to release nitric oxide (McAdam et al., 2012). However, nitric oxide (NO) was unstable and difficult to detect. In this study, the Griess-reagent was used to interact with nitrite ions (NO₂^–) to understand whether CKHLE could mitigate the inflammatory response of macrophages under the mode. In this study, the NO production in the positive control group was 20.0 μM, which was about 5 folds higher than the negative control group (4.5 μM). The 4 CKHLEs were treated with 1–25 μg/mL and compared to the positive control group, the 95% ethanol extracted CKHLE could have reduced the NO production by 12–37% (Fig. 1). Furthermore, the anti-inflammatory effect has most significantly observed in the concentration of 10 μg/mL, which reduced NO production at a maximum of 50.05% compared to the positive control group. The 50% ethanol extraction of CKHLE could reduce the NO concentration by 5–26%. Other than water-extracted CKHLE, the concentrations of the other three CKHLEs ranged from 1 to 25 μg/mL, significantly reducing NO production compared to the positive control group (p < 0.05), and the decrease in NO was more significant with higher CKHLE concentration. Lin et al. (2008) reported that the use of camphor methanol extract at a concentration of 25 μg/mL significantly inhibited NO production in Raw 264.7 cells, in agreement with the results of this study. Lee et al., (2006) found that the anti-inflammatory effect of Cinnamomum camphora n-butanol crude extract on Raw 264.7 cells significantly reduced the iNOS expression by 50–80%.

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Fig. 1

Effects of extracts from the leaves of Cinnamomum kanehirae Hayata on anti-inflammation of Raw 264.7 cells. Con.: Concentration. *There was a significant difference (p < 0.05) compared to the positive control group. The data are presented as means ± SD (n = 3). 95%: 95% ethanol extract of Cinnamomum kanehirae Hayata leaves. 80%: 80% ethanol extracts of Cinnamomum kanehirae Hayata leaves. 50%: 50% ethanol extracts of Cinnamomum kanehirae Hayata leaves. Water: water extracts of Cinnamomum kanehirae Hayata leaves.

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