Introduction

Vegetables and fruits are considered to be good sources of phenolic compounds. In the past decades, phenolic compounds, especially phenolic acids, flavonoids, and anthocyanins, have been demonstrated to be important health-promoting dietary phytochemicals. Evidence has confirmed that phenolic compounds delayed the onset of some chronic diseases, such as inflammation, hypertension, arteriosclerosis and cardiovascular disease [1,2,3].

Cudrania cochinchinensis, a perennial undershrub of Moraceae family, is widely distributed in Southern China, Japan and Korea. In China, the roots of C. cochinchinensis have been used as a traditional medicine to treat humid jaundice, hepatoprotective, gonorrhea and bruising [4]. It was found that phenolic and flavonoid compounds in the extracts of C. cochinchinensis roots possessed strong tyrosinase inhibitory activities [5]. The extract of C. cochinchinensis roots exhibited strong antioxidant activities [6, 7]. While these studies offer valuable information on the chemical identity and bioactivity of the plant constituents, their stability and bioaccessibility in the gut must be taken into account. It has been reported that gastrointestinal conditions, such as temperature, pH value and related enzymes have significant effects on the stability and bioaccessibility of the phenolic compounds [8, 9]. For example, Correa-Betanzo et al. [10] found that anthocyanins have low bioavailability due to their low stability under the alkaline conditions of the small intestine. Goulas et al. [11] reported that flavonoids from carob fruit had low release efficiencies than phenolic acids during in vitro digestion.

In vitro simulated GID models have been widely used for investigating the changes in the stability, bioaccessibility, and biological activity of phytochemicals. Compared to the human clinical trials, in vitro approaches have the advantages of being more straightforward, efficient and cheaper. The study aimed to evaluate the effects of ethanol extracts from C. cochinchinensis roots under simulated GID conditions on the bioaccessibility, antioxidant activity and inhibition activities against α-glucosidase and tyrosinase, as well as cytotoxicity on HepG2 cells. Moreover, the phenolic profile changes during GID were assessed by UPLC-TOF-MS/MS.

Materials and methods

Chemicals and reagents

Gallic acid, Folin–Ciocalteu reagent (2 N), rutin, DPPH, ABTS, tripyridyltriazine (TPTZ), α-glucosidase, tyrosinase, pepsin, pancreatin, bile salts* and trolox were purchased from Sigma-Aldrich Trading Co. Ltd. (Shanghai, China). AB-8 macroporous resins were purchased from Donghong Chemical Co. Ltd. (Xian, China), and HPLC-grade formic acid and acetonitrile were obtained from Fisher Scientific (Pittsburgh, PA, USA). Cell Counting Kit-8 and HepG2 human liver cancer cells were obtained from the Beijing Solarbio Science & Technology Co., Ltd. (Beijing, China). The deionized water used in the experiment was procured using a Milli-Q system (Merck Millipore, USA). All other chemicals and reagents used in this study were of analytical grade.

Plant material and sample preparation

The C. cochinchinensis roots used in this study were collected from the Ta-pieh Mountains in LiuAn City, Anhui Province, China in September 2019. The C. cochinchinensis roots were air dried and then pulverized into a homogenate by a pulverizer (FW100, 107 Tianjin Taistite Instrument Co., Ltd, Tianjin, China) and then passed through 30–40 mesh. Samples were placed in aluminum bags, evacuated and stored at 4 °C until further used.

Preparation of the crude ethanol extracts

The flour of C. Cochinchinensis roots was extracted with 80% ethanol solution (v/v) to solvent ratio of 1:20 (w/v) with continuous stirring at room temperature for 2 h. The mixture was centrifuged at 4000 rpm, 25 °C for 15 min, and the residue was re-extracted twice with 80% ethanol as described above. After removal of ethanol with rotary evaporator under vacuum conditions at 50 °C, the crude ethanol extracts (CEE) was obtained, freeze-dried and stored at − 20 °C until use.

Purification of CEE

CEE was further purified to obtain the purified ethanol extracts (PEE) by the AB-8 macroporous adsorption resin according to the method described by Zhu et al. [12] with a slight modification. Briefly, the bioactive components of CEE were separated and enriched on a glass column (16 × 300 mm) wet-packed with 50 g AB-8 macroporous adsorption resin. The CEE was appropriately diluted with distilled water and loaded onto the column at a constant flow rate of 1.0 mL/min. After reaching equilibration, the column was washed with 100 ml distilled water to remove impurities and eluted with 70% ethanol at the flow velocity of 2 mL/min. At last, all the eluates were concentrated in a vacuum rotary evaporator at 50 °C and then freeze-dried under vacuum conditions. The dry powder of PEE was stored at – 20 °C in the dark for subsequent analysis.

Simulated in vitro gastrointestinal digestion

The in vitro digestion of PEE was carried out according to the method reported by Guimarães Drummond et al. [13], with slight modifications. Briefly, in gastric digestion (GD) step, PEE samples (500 mg) were dispersed in 5 mL of deionized water containing 35 mM NaCl at the ratio of 1:10 (w/v) to obtain 100 mg/mL PEE solution, the pH value was adjusted to 2.0 with 1 M HCl, and the mixtures were incubated in a shaker water bath at 37 °C for 5 min. Then, pepsin (25 mg) was added (E:S 1:20 w/w), and pH value was again adjusted to 2.0. The digestions were shaken at 150 rpm, 37 °C for 90 min. At the end of gastric phase, an aliquot of the supernatant was taken and freeze-dried to yield the gastric digestion-purified ethanol extracts (GD-PEE). Then, the pH value of mixtures was adjusted to 7.0 with 1 M NaHCO3. 2 mg of CaCl2, 45 mg of bile salt and 50 mg of pancreatin (E:S 1:10 w/w) were added, and the pH value of the mixtures from last step was again adjusted to 7.0 with 1 M NaHCO3 to simulate intestinal digestion. After 120 min of intestinal digestion at 37 °C, the mixture was centrifuged at 12,000 rpm for 15 min to remove impurities. The supernatants were freeze-dried to yield the intestinal digested-purified ethanol extracts (ID-PEE) and kept at − 20 °C until further used.

Analyses of total phenolic and total flavonoid

Determination of total phenolic content (TPC)

The total phenolic content was measured using the Folin–Ciocalteu spectrophotometric method [14]. Briefly, 200 μL of sample solution (0.5 mg/mL) diluted with deionized water was mixed with 100 μL of Folin–Ciocalteu reagent (2 N). The mixture was allowed to react for 6 min at ambient temperature in the dark. Then, 1 mL of 7% sodium carbonate solution (Na2CO3) was added to neutralize the reaction. The green color developed after 1 h and then absorbance value at 760 nm measured using a TU-1950 UV–vis spectrophotometer (Purkinje General Instrument Co., Beijing, China). Gallic acid was used as a reference to obtain the content of phenolic, and the results were expressed as milligram of gallic acid equivalents per gram of dry weight of sample (mg GAE/g dw).

Determination of total flavonoid content (TFC)

The content of total flavonoid in the extract was determined by using the method [15] reported previously, with minor modifications. Briefly, 2.5 mL of sample (0.25 mg/mL) diluted with deionized water and 300 μL of NaNO2 (5%) were mixed for 6 min; then 300 μL of AlCl3 (10%) was added and allowed to react at room temperature for 6 min, followed by addition of 4% sodium hydroxide (4.4 mL), and incubation for 10 min. The absorbance of the mixture was determined at 510 nm using a spectrophotometer. Rutin was used as a reference to obtain the content of total flavonoids, and the results were expressed as milligram of rutin equivalents per gram of dry weight of sample (mg RE/g dw).

Bioaccessibility of phenolic and flavonoid after simulated digestion

The bioaccessibility through an in vitro gastrointestinal digestion was calculated using the following equation:

$$ {\text{Bioaccessibility index }}\left( \% \right) = {\text{(final concentration/initial concentration) }} \times { 100,} $$

where the final concentration is the TPC/TFC at the end of gastric and intestinal digestion, and initial concentration is the TPC/TFC in the undigested purified ethanol extracts (PEE).

Identification of phenolic compounds in C. cochinchinensis roots

Individual phenolic compounds were identified by Agilent LC30 system equipped with a quaternary pump, an autosampler, thermostated column compartment and a diode array detector. Separations were carried out on Agilent C18-H reverse-phase column (150 mm × 2.1 mm × 3.5 μm) at 30 °C. The injection volume was 5 μL. The solvent system, consisting of formic acid (0.1%) in Milli-Q water (A) and acetonitrile (B), was pumped into the HPLC system at 0.6 mL/min, and wavelength of UV detector was set at 280 nm. The gradient conditions were 0–5 min, 5% B; 5–25 min, 5–20% B; 25–30 min, 20–95% B; 30–35 min, 95–5% B.

For identification, an AB SCIEX Triple TOF 5600 + system (SCIEX, Foster City, CA, USA) was used, and the MS spectrum was obtained using electrospray ionization operating in negative modes in the range of m/z 50–1200. Other major parameters were set as follows: ion spray voltage floating, − 4500 V; curtain gas, 35 psi; ion source gas1, 50 psi; ion source gas 2, 50 psi; source temperature, 550 °C; collision energy, 10 eV; declustering potential, − 80 V. Data were collected in centroid mode using MSn to obtain both protonated molecular ions and/or adducts and product ions, and MassBank online database (http://www.massbank.jp) was used for analysis.

Determination of the antioxidant activity of phenolic extractsl scavenging activity

DPPH radical scavenging activity was measured using a modified colorimetric method reported by Fraga et al. [16]. A sample solution (2 mL, 0.02 mg/mL) was mixed with 2.0 mL of 95% ethanol solution of DPPH, and the absorbance of the mixture was measured at 517 nm after incubation for 30 min in the dark at the temperature. The scavenging activity of the DPPH radicals was expressed as mmol of Trolox equivalent (TE) per gram of dry weight of sample (mmol TE/g dw).

ABTS radical scavenging activity

The ABTS assay was conducted by the method reported by Fraga et al. [16]. The stock solution of ABTS radical cation (ABTS+) was generated by mixing ABTS solution (7 mM) and potassium persulfate (2.45 mM) in the dark at room temperature for 12–16 h. Prior to analysis, the solution was diluted with ultrapure water until the absorbance of 0.70 ± 0.02 at 734 nm. A sample solution (1 mL, 0.05 mg/mL) and 2 mL of ABTS+ solution were mixed and incubated at room temperature for exactly 10 min and the absorbance measured at 734 nm. The scavenging activity of the ABTS+ radicals was expressed as mmol of Trolox equivalent (TE) per gram of dry weight of sample (mmol TE/g dw).

Ferric reducing antioxidant power assay (FRAP)

The FRAP assay of ethanol extracts was performed using the previous method [16], with some modifications. Briefly, the stock FRAP solution was prepared by mixing 300 mM acetate buffer (pH 3.6) and 10 mM tripyridyltriazine (TPTZ) solution in 40 mM HCl and 20 mM FeCl3·6H2O solution in a 10:1:1 ratio. A sample solution (400 μL, 0.1 mg/mL) was allowed to react with the FRAP solution (3 mL) at 37 °C under dark conditions. Absorbance was recorded at 593 nm after exactly 10 min of the reaction. The results of FRAP were expressed as mmol of Trolox equivalent (TE) per gram of dry weight of sample (mmol TE/g dw).

Cupric ion reducing activity (CUPRAC)

The cupric ion reducing activity was determined using the method reported by Tlili [17]. A sample solution (0.5 mL, 0.05 mg/mL) was added to a premixed reaction mixture containing 1 mL CuCl2 solution (10 mM), 1 mL ethanolic neocuproine solution (7.5 mM) and 1 mL NH4Ac buffer (1 M, pH 7.0). In addition, distilled water was used instead of CuCl2 as a blank control. Absorbance against a blank reagent was measured at 450 nm after 30 min. The results of CUPRAC were expressed as mmol of Trolox equivalent (TE) per gram of dry weight of sample (mmol TE/g dw).

Phosphomolybdenum method

The total antioxidant capacity (TAC) was measured by phosphomolybdenum method based on the protocol reported by Mocan et al. [18]. To the sample solution (0.5 mL, 0.5 mg/mL), 3 ml of the reagent (0.6 M sulfuric acid, 28 mM sodium phosphate and 4 mM ammonium molybdate) were added and the mixture was incubated at 95 °C for 90 min. Afterwards, the absorbance against a blank was read at 695 nm. The total antioxidant capacity was expressed as mmol of Trolox equivalent (TE) per gram of dry weight of sample (mmol TE/g dw).

Enzyme inhibitory activity

α-Glucosidase inhibition

The α-glucosidase inhibitory activity was analysed according to the previous method [19]. Briefly, the α-glucosidase solution was diluted to 700 U/mL with 10 mM phosphate buffer (pH 6.8). The inhibition activity was assayed by mixing 50 μL of sample solution (25 μg/mL) and 50 μL of α-glucosidase solution, and 50 μL of and incubated at 37 °C for 15 min. After that, 50 μL of 4-N-trophenyl-α-glucopyranoside (PNPG) was added, and incubate at 37 °C for 15 min. Finally, 100 μL of 0.2 M sodium carbonate solution was added to stop the reaction. The absorbance was measured at 405 nm using a microplate reader and compared with a control (buffer instead of the extracts). Acarbose was used as a positive control, and the α-glucosidase inhibitory activities were expressed as millimoles of acarbose equivalents per gram of dry weight of sample (mmol ACE/g dw).

Tyrosinase inhibition

Tyrosinase inhibitory activity was determined according to a method reported previously [20], with slight modifications. Briefly, 100 μL of phosphate buffer (pH 6.8), 100 μL of sample solution (5 μg/mL), and 50 μL of tyrosinase solution were added to a 96-well microplate and incubated at 37 °C for 15 min. Subsequently, 50 μL of l-DOPA was added and then incubated at 37 °C for 15 min. The absorbance was measured at 475 nm with a microplate reader and compared with a control (without enzyme). Kojic acid was used as a positive control, and the tyrosinase inhibitory activities were expressed as millimoles of kojic acid equivalents per gram of dry weight of sample (mmol KAE/g dw).

Cell culture and cytotoxicity

HepG2 are immortalized cell line consisting of human liver carcinoma cells, derived from the liver tissue of a 15 year-old Caucasian male who had a well-differentiated hepatocellular carcinoma. In this study, HepG2 cells were cultured in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% (v/v) fetal bovine serum and 1% (v/v) penicillin/streptomycin. Cells were cultured in cell culture CO2 incubator at 37 °C under a humidified atmosphere containing 95% air and 5% CO2.

The viability of HepG2 cells was assessed using the Cell Counting Kit-8 (CCK-8) [21]. Briefly, 100 μL of complete culture medium containing 104 HepG2 cells/well was plated in a 96-well plate. After 24 h, the complete medium was replaced by the samples and incubated for another 24 h. After reaction with 10% CCK-8 for 40 min, the optical density (OD) was read with a microplate reader at 450 nm. \({\text{Cell viability }}\left( \% \right)\, = \,\left[ {{\text{A}}_{{{\text{sample}}}} {-} {\text{A}}_{{\text{sample background}}} \left] / \right[{\text{A}}_{{{\text{control}}}} {-} {\text{A}}_{{\text{control background}}} } \right]\, \times \,{1}00.\)

Statistical analysis

Results are presented as mean ± standard deviation (n = 3). The statistical differences between the different extracts were analyzed by using one-way analysis of variance (ANOVA), followed by Duncan’s test with SPSS statistical 26 software. A p value < 0.05 was considered statistically significant.

Results and discussion

Tentative identification of phenolics from the CEE, PEE, GD-PEE, ID-PEE

UPLC-TOF-MS/MS was used to identify the phytochemicals in the ethanol extracts from C. cochinchinensis roots and its digested samples. Fifteen compounds were tentatively identified, based on their retention times, related literature, molecular mass and by comparison of their MS/MS spectra with the online databases, including five phenolic compounds (Peaks 1, 2, 3, 9 and 12) and ten flavonoid compounds (Peaks 4, 5, 6, 7, 8, 10, 11, 13, 14 and 15). MS2 spectra of tentatively identified compounds are shown in supplementary material. Total ion chromatograms (TIC) in negative mode of C. cochinchinensis samples are shown in Fig. 1. Information of all compounds tentatively identified in this study is summarized in Table 1. Structures of the tentatively identified compounds of CEE, PEE, GD-PEE, and ID-PEE samples from C. cochinchinensis roots are shown in Fig. 2. The characteristics of these compounds are discussed as follows:

Fig. 1
figure 1

Total ion chromatogram (TIC) of (A) crude ethanol extracts (CEE), (B) purified ethanol extracts (PEE), (C) gastric digestion-purified ethanol extracts (GD-PEE) and (D) intestinal digestion-purified ethanol extracts (ID-PEE) samples in negative ion mode

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Table 1 Tentative identification of bioactive compounds in C. cochinchinensis root samples (CEE, PEE, GD-PEE, ID-PEE) by UPLC-TOF-MS/MS
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Fig. 2
figure 2

Structures of the phenolic compounds identified in CEE, PEE, GD-PEE, ID-PEE samples from C. cochinchinensis roots. CEE crude ethanol extracts; PEE purified ethanol extracts; GD-PEE gastric digestion-purified ethanol extracts; ID-PEE intestinal digestion-purified ethanol extracts

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Compounds 1 and 2 exhibited the molecular ion [M–H]− at m/z 137.0301 and 153.0181, respectively, and their fragments generated highly significant peaks at m/z 93.0395 and 109.0302. The two compounds were preliminarily determined to be 4-hydroxybenzoic acid [22] and 3,5-dihydroxybenzoic acid [23], respectively. Compound 3 was proposed as 6,7-dihydroxycoumarin based on molecular ion [M–H]− at m/z 177.0231 with fragments at m/z 149.0273, 133.0318, 89.0415 and 77.0431 by matching the MS and MS/MS data with a report [24]. Compound 4, with [M–H]− ion of m/z 449.1089, produced a secondary ion peak at m/z 287.0562 [M–H–C6H10O5]− and 259.0629 [M–H–C6H10O5–CO]−, was tentatively identified as eriodictyol-7-O-glucoside, based on the literature [25]. Compounds 5 and 6, showing a molecular ion at m/z 445.9921 and 609.1472, were identified as biochanin-A8-C-glucoside and luteolin-3’,7-di-O-glucoside by matching the online databases. Specifically, compound 5 losing one glucose residue and produced fragment ion at m/z 283.0558, whereas compound 6 losing two glucose residues and produced fragment ions at m/z 447.0986 and 285.0434. Compound 7, with an [M–H]− ion of m/z 317.0657, had typical fragments of m/z 177.0213 and m/z 151.0424. These were attributed to the neutral loss of C7H7O3 and C8H5O4, respectively, resulting in the identification of the compound as myricetin [26]. Compound 8, with molecular ion [M–H]− at m/z 303.0540, produced fragments at m/z 285.0437, 217.0538, 177.0214 [M–H–C7H2O5]−, 149.0268 [M–H–C7H2O5–CO]− and 125.0268, the fragment in accordance with the mass spectrometry fragmentation pattern of taxifolin [27]. Compound 9 was tentatively identified as 9,10-dihydro-2,3,5,7-phenanthrenetetrol by matching with online database with a [M–H]− at m/z 243.0667, fragmentation ions at m/z 225.0556, 199.0761 and 175.0762. Compound 10 exhibited molecular ion [M–H]− at m/z 287.0576 and fragments at m/z 259.0620 [M–H–CO]− and 125.0249 [M–H–CO–C8H6O2]− and was tentatively identified as dihydrokaempferol by comparison with the data profile with a previous report [24]. Compound 11 was tentatively identified as kaempferol-3-O-glucoside by a related literature, followed by the fragmentation of m/z 447.0950, with a neutral loss of glucose and MS2 peak at m/z 285.0410 [28]. Compound 12, with [M–H]− ion at m/z 227.0750, was tentatively identified as resveratrol as its MS2 exhibited a product ion at m/z 185.0623 [M–H–C2H2O]− and 143.0516 [M–H–C2H2O–C2H2O]−, indicating the loss of C2H2O molecule and 2C2H2O, respectively [29]. Compound 13 exhibited the [M–H]− ion at m/z 301.0368, which further fragmented into m/z 257.0425 [M–H–CO2]−, 229.0502 [M–H–CO2–CO]−, 151.0048 [M–H–C8H6O3]−and 125.0242 [M–H–C9H4O4]−, and was identified as morin by matching the online database and literature [30]. Compound 14, with [M–H]− at m/z 285.04 and released fragments at m/z 257.0512 [M–H–CO]−, was thus tentatively identified as 3,6,3’,4’-tetrahydroxyflavone. Compound 15, with [M–H]− at m/z 287.0596 and base peak at m/z 161.0252, 135.0456 and 125.0248, was tentatively identified as 3’,4’,5,7-tetrahydroxyisoflavanone.

As shown in Fig. 1, a total of 15 compounds were also found both in the TIC of GD-PEE and ID-PEE compared with PEE. The peak intensities of most compounds were decreased with sequential gastrointestinal digestion (Table 1). This may be due to the interaction between the compounds or the bonding of the compounds to the enzyme. Cao et al. [31] reported that simulated gastrointestinal digestion changed the structures of some flavonoids. Interestingly, the intensity of 9,10-dihydro-2,3,5,7-phenanthrenetetrol (compound 9) after intestinal digestion was 1.05 times than after gastric digestion, and the intensity of morin (compound 13) after intestinal digestion was 1.16 times than gastric digestion. These findings may be explained by the release of phenolic compounds from test matrix after intestinal digestion due to the break of the bonds of these compounds to sugar residues, proteins or fibers [32]. Quatrin et al. [33] reported the same phenomenon of phenolic compounds in jaboticaba (Myrciaria trunciflora) fruit peel after in vitro GID. A similar result was observed by Zhang et al. [34] in digested Cinnamomum camphora seed kernel.

Effect of gastrointestinal digestion on TPC, TFC and bioaccessibility

The crude ethanol extracts (CEE) from C. cochinchinensis roots were purified by AB-8 macroporous adsorption resin to obtain purified ethanol extracts (PEE). As shown in Fig. 3A, the TPC (596.18 ± 1.79 mg GAE/g dw) and TFC (379.64 ± 1.83 mg RE/g dw) in PEE were the highest. Compared with CEE, the contents of total phenolic and total flavonoid increased 52.92% and 36.73%, respectively. Zhu et al. [12] studied phenolic compounds from Sanguisorba. officinalis L. and found that TPC was increased from 31.50 ± 1.31 mg GAE/g dw to 338.92 ± 1.06 mg GAE/g dw after purification by macroporous resin adsorption. A similar result in Potentilla discolor Bge was reported [35]. By purification, the TPC of the ethanol extracts was significantly increased, which is helpful for us to study the effect of in vitro gastrointestinal digestion on phenolic compounds.

Fig. 3
figure 3

Contents (A) of total phenolic and total flavonoid in CEE, PEE, GD-PEE, ID-PEE samples and bioaccessibility (B) of GD-PEE, ID-PEE from C. cochinchinensis roots. CEE crude ethanol extracts; PEE purified ethanol extracts; GD-PEE gastric digestion-purified ethanol extracts; ID-PEE intestinal digestion-purified ethanol extracts. GAE gallic acid equivalent; RE rutin equivalent; dw dry weight. Data are mean (n = 3) ± standard deviation. Different letters represent significant differences (p < 0.05)

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Simulated GID had a significant effect (P < 0.05) on TPC and TFC of PEE from C. cochinchinensis roots. As shown in Fig. 3A, the phenolic contents of GD-PEE and ID-PEE were significantly decreased by simulated in vitro digestion. The contents of total phenolic obtained in the vitro digestion were 329.85 mg GAE/g dw for gastric phase and 236.70 mg GAE/g dw for intestinal phase. Compared with the undigested PEE, the TPC was reduced by 44.67% after the gastric phase and by 60.29% at the end of the intestinal phase. The losses of phenolic compounds may be due to their interaction with other compounds and digestive enzymes or changes in molecular structure, which may influence their solubility [36]. In addition, the phenolic compounds in extracts had poor stability during GID. The decreased TPC may be caused by poor stability of phenolic compounds. Similar results have been reported by Quatrin et al. [33] and Bouayed et al. [37]. Moreover, the stability and bioaccessibility of phenolic compounds were found to mainly depend on pH value in the intestinal phase [38]. Some phenolic compounds may be degraded by non-enzymatic oxidation under near neutral conditions and in the presence of oxygen, which may be the reason for the further decrease of TPC in the intestinal stage. Similar results were previously found in the simulated GID. For example, Thomas-Valdés et al. [39] reported that the TPC in Chilean white strawberry were decreased by 18.0% after gastric phase and 55.7% after intestinal phase. Chilean red strawberry showed a high stability of TPC after the gastric phase (96.6%), followed by a significant reduction (43.0%) after the intestinal phase [40].

The total flavonoid contents were 199.09 mg RE/g dw and 126.66 mg RE/g dw for GD-PEE and ID-PEE, respectively. Compared with the PEE (undigested), the TFC was reduced by 47.55% after the gastric phase and by 66.63% at the end of the intestinal phase. As shown in Fig. 3, the loss of TFC was high than TPC. This phenomenon may be because flavonoids were less stable than phenolics in gastrointestinal digestion and may react with OH− or be degraded under high pH conditions [11].

Gastrointestinal digestion affects bioaccessibility of phenolic and flavonoid compounds. At the gastric digestion stage, the bioaccessibility of total phenolic and flavonoids were 55.66% and 52.44%, respectively. After the intestinal digestion, the bioaccessibility of total phenolic and flavonoid contents were reduced to 39.70% and 33.36%, respectively (Fig. 3B). Different types of compounds were all affected by gastrointestinal digestion. As shown in Table 1, individual compounds that had less effect on bioavailability in gastric digestion were 3,5-dihydroxybenzoic acid, 6,7-dihydroxycoumarin, myricetin, taxifolin, 9,10-dihydro-2,3,5,7-phenanthreneterol and morin. However, individual compounds that had less effect on bioavailability in intestinal digestion were 3,5-dihydroxybenzoic acid, 9,10-dihydro-2,3,5,7-phenanthreneterol and morin. The biotransformation of phenolic compounds during GID might affect their bioaccseeibility and be related to changes on their bioactivity [41], but the reasons still need further investigated.

Effect of gastrointestinal digestion on antioxidant activity

Five different antioxidant assays were used to investigate the antioxidant activity of CEE, PEE and its digested samples. As shown in Fig. 4, the antioxidant activity of PEE from C. cochinchinensis roots was significantly increased compared with CEE. Liao et al. [6] also reported that the extracts of C. cochinchinensis roots had higher DPPH radical scavenging activity and ferric ion reducing ability.

Fig. 4
figure 4

Antioxidant activities of CEE, PEE, GD-PEE and ID-PEE expressed in mmol TE/g dw, including DPPH (A), ABTS (B), FRAP (C), CUPRAC (D) and total antioxidant activity (E) assays. CEE crude ethanol extracts; PEE purified ethanol extracts; GD-PEE gastric digestion-purified ethanol extracts; ID-PEE intestinal digestion-purified ethanol extracts; TE trolox equivalent; dw dry weight. Data are mean (n = 3) ± standard deviation. Different letters represent significant differences (p < 0.05)

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A significant loss of the antioxidant activity was observed throughout the in vitro GID (Fig. 4). The scavenging activity of the samples towards the DPPH and ABTS+ radical decreased throughout the GID process (Fig. 4A, B). In DPPH assay, the values of PEE, GD-PEE and ID-PEE were 1.80 mmol TE/g dw, 1.27 mmol TE/g dw and 1.03 mmol TE/g dw. Compared with PEE, the values were 29.44% and 42.77% lower after the gastric and intestinal phase, respectively. Similarly, the ABTS+ scavenging activity of PEE was the highest (10.19 mmol TE/g dw), followed by GD-PEE (5.49 mmol TE/g dw) and ID-PEE (3.95 mmol TE/g dw). ABTS activity was reduced 46.12% after the gastric digestion, with a further loss of 15.11% after the intestinal digestion. The results were similar to that of Lucas-González et al. [42] who found that DPPH scavenging activity of persimmon fruit co-products were decreased 13.1% and 30.8%, respectively, after gastric and intestinal digestion. Tagliazucchi et al. [43] also reported that ABTS radical scavenging activity of grape polyphenols decreased during in vitro digestion.

In the FRAP and CUPRAC assays, PEE also showed the highest reducing power activity (2.39 mmol TE/g dw and 4.81 mmol TE/g dw, respectively). After gastric digestion, the values of FRAP and CUPRAC were reduced by 50.21% and 48.75% compared with PEE, respectively. At the end of intestinal digestion, the values of FRAP and CUPRAC were reduced by 59.83% and 63.12% compared with PEE, respectively. Similar results were observed in Ribes magellanicum [44] and different varieties of apple during in vitro GID [37]. On the contrary, Gullon et al. [45] found that FRAP of pomegranate peel flour was increased slightly after gastric digestion. This observation may be attributed to different types of phenolic compounds.

Total antioxidant capacity (TAC) is also an important indicator to assess the effect of in vitro digestion on antioxidant activity. A reduction of the TAC of the digested samples were observed throughout the digestion process. The TAC values at the gastric and intestinal digestion stage were reduced by 58.47% and 70.13%, respectively, with respect to the undigested PPE. Our results were in agreement with Kosti´c et al. [46] who found that TAC of phenolic from goat milk was decreased after in vitro digestion.

These results are significantly correlated with the changes in TPC during the GID process (Table 2, p < 0.01). Naeimi et al. [47] reported that myricetin, dihydrokaempferol and morin had a strong ability to scavenge DPPH free radicals, so the decrease in scavenging activity of DPPH free radicals may be closely related to the decrease in the contents of myricetin, morin and dihydrokaempferol, as shown in Table 1. In addition, the changes in ferric reducing antioxidant power may be closely related to myricetin and taxifolin contents [48]. Many previous studies have confirmed the relationship between TPC and antioxidant activity. Compounds with strong antioxidant activity, such as luteolin-3’,7-di-o-glucoside (compound 6) and resveratrol (compound 12), also decreased significantly after GID process. The change in the pH value of the food matrix from the gastric to the intestine also had a certain effect on the phenolic compounds [49]. The hydroxyl groups on the aromatic ring were partially deprotonated, and the ionization balance changed [37], resulting in a reduction in antioxidant capacity.

Table 2 Coefficients of correlation between the TPC or TFC and antioxidant activity (DPPH, ABTS, FRAP, CUPRAC, TAC), enzyme inhibition activity (α-glucosidase, tyrosinase)
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Effect of gastrointestinal digestion on enzyme inhibitory activity

The PEE from C. cochinchinensis roots showed the highest α-glucosidase inhibitory activity (0.51 mmol ACE/g dw). However, the in vitro digestion significantly reduced the inhibitory activity after the gastric (39.21%) and intestinal phases (50.98%), respectively (Table 3). The loss of inhibitory activity was correlated with the changes in the TPC throughout the GID. Similar results were reported previously. For example, Thomas-Valdés et al. [40] found that the inhibitory activity of the phenolics from the native Chilean red strawberry towards α-glucosidase was decreased gradually by the gastric and intestinal digestion. Interestingly, Kasipandi et al. [50] reported that α-glucosidase inhibitory activity of Opilia amentacea roxb fruit extracts increased slightly after gastric digestion and finally decreased significantly after the intestinal digestion. In addition, the type and structure of phenolic compounds in the samples may play an important role in enzyme inhibition. For example, resveratrol caused structural changes of the enzyme through non-competitive binding with the α-glucosidase region, thereby reducing enzyme activity [51]. Myricetin interacts with key amino acids of α-glucosidase through hydrogen bonds to inhibit α-glucosidase activity [52]. As shown in Table 1, the peak intensities of myricetin, resveratrol and most phenolic compounds were decreased gradually through in vitro digestion. This finding also explains why the inhibitory activity of α-glucosidase was reduced after GID.

Table 3 Enzyme inhibitory activities of C. cochinchinensis root extracts (CEE, PEE, GD-PEE, ID-PEE)
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Tyrosinase is a copper-containing polyphenol oxidase and is responsible for browning reactions through the phylogenetic scale. Over-activity of this enzyme was associated with human freckles, brown spots and neurodegenerative disorders [53]. As shown in Table 3, the PEE exhibited the highest inhibition activity, and the inhibitory activity was significantly decreased after the gastric digestion. It is worth noting that there were no significant difference in the inhibitory activity of tyrosinase after gastric digestion and intestinal digestion. This phenomenon may be due to the fact that the contents of phenolic compounds that inhibit tyrosinase were not significantly reduced after gastric digestion. The results suggested that phenolic compounds of C. cochinchinensis roots play an important role in the inhibitory activities for tyrosinase. It was also reported that the extracts of C. cochinchinensis roots had a strong tyrosinase inhibitory activity [5, 6].

Cytotoxic effects on HepG2 cells

As shown in Fig. 5, compared with the control, there was no significant difference in the viability of HepG2 cells when the concentration of PEE, GD-PEE and ID-PEE was lower than 75 μg/mL. In addition, there was no significant difference in the cell viability when the concentration of CEE was lower than 100 μg/mL. However, compared with PEE, the cell viability significantly changed after simulated digestion, and the highest cell viability was observed in the ID-PEE group. This result differences may be due to the decrease in the contents of phenolic compounds from the C. cochinchinensis roots after simulated GID [54].

Fig. 5
figure 5

The viability of HepG2 cells after 24 h by crude ethanol extracts (CEE), purified ethanol extracts (PEE), gastric digestion-purified ethanol extracts (GD-PEE) and intestinal digestion-purified ethanol extracts (ID-PEE). CCK-8 assay was used to assess cell viability. Data are mean (n = 6) ± standard deviation. ** indicates p < 0.05

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Correlation between the parameters

A Pearson’s correlation analysis was used to determine the correlation coefficients between the TPC or TFC and the bioactivities of ethanol extracts from C. cochinchinensis during GID. As shown in Table 2, in the simulated GID process, there were significantly positive correlations between the TPC/TFC and the antioxidant or enzyme inhibitory activity, especially the ability to scavenge ABTS+ free radicals and cupric reducing antioxidant. These results further demonstrated that the TPC/TFC of ethanol extracts from C. cochinchinensis roots during simulated GID play important roles in the bioactivity. These results were consistent with those of Lucas-Gonzalez et al. [55] and Arruda et al. [56] reporting that the phenolic and flavonoid compounds were the main contributors to the antioxidant activities from maqui berry and edible part of araticum fruit.

Conclusions

The study investigated the influence of simulated gastrointestinal digestion on phenolic bioaccessibility and bioactivity of C. cochinchinensis roots. Results showed that the total phenolic/flavonoid contents were generally decreased in the order of sequential digestions. Fifteen phenolic substances were tentatively identified by UPLC-TOF-MS/MS before and after simulated digestion, including ten flavonoids and five phenolics. These changes in the total phenolic/flavonoid contents were significantly correlated with the loss of antioxidant activity and enzyme inhibitory activity, as well as cytotoxicity on HepG2 cells. Overall, our results have helped understand the effects of gastrointestinal digestion on the bioactivity and bioaccessibility of ethanol extracts from C. cochinchinensis roots and provided a theoretical basis for potential applications in the food, cosmetics and pharmaceutical industries. However, these conclusions need to be confirmed by further studies in vivo and security evaluation.