Microwave-Assisted Extraction of Phenolic Compounds and Antioxidants for Cosmetic Applications Using Polyol-Based Technology

Su Myat Win; Manee Saelee; Hla Myo; Nuntawat Khat-Udomkiri

doi:10.3791/67033

JoVE Journal > Chemistry

Chemistry

Microwave-Assisted Extraction of Phenolic Compounds and Antioxidants for Cosmetic Applications Using Polyol-Based Technology

Published: August 23, 2024

doi:

10.3791/67033

Su Myat Win¹, Manee Saelee¹, Hla Myo^1,2, Nuntawat Khat-Udomkiri¹

¹School of Cosmetic Science,Mae Fah Laung University, ²College of Public Health Sciences,Chulalongkorn University

Summary

This protocol details the utilization of a polyol-based microwave-assisted extraction method for extracting phenolic compounds and natural antioxidants, representing a practical and environmentally sustainable approach to the development of ready-to-use extracts.

Abstract

The utilization of polyols as green solvents for extracting bioactive compounds from plant materials has gained attention due to their safety and inert behavior with plant bioactive chemicals. This study explores the sustainable extraction of phenolic compounds and natural antioxidants from coffee silverskin using the microwave-assisted extraction (MAE) method with polyol-based solvents: glycerin, propylene glycol (PG), butylene glycol (BG), methylpropanediol (MPD), isopentyldiol (IPD), pentylene glycol, 1,2-hexanediol, and hexylene glycol (HG). A comparative analysis was conducted on conventional and non-conventional solvent extractions, focusing on their impact on the bioactive compounds of MAE, encompassing parameters such as total phenolic content (TPC), total flavonoid content (TFC), and antioxidant activities like the 1,1-diphenyl-2-picrylhydrazyl radical scavenging assay (DPPH), the 2,2′-azino-bis(-3-ethylbenzothiazoline-6-sulphonic acid) radical scavenging assay (ABTS), and the ferric reducing antioxidant power assay (FRAP). The highest values were observed for TPC with aqueous-1,2-hexanediol extraction (52.0 ± 3.0 mg GAE/g sample), TFC with aqueous-1,2-hexanediol extraction (20.0 ± 1.7 mg QE/g sample), DPPH with aqueous-HG extraction (13.6 ± 0.3 mg TE/g sample), ABTS with aqueous-pentylene glycol extraction (8.2 ± 0.1 mg TE/g sample), and FRAP with aqueous-HG extraction (21.1 ± 1.3 mg Fe (II) E/g sample). This research aims to advance eco-friendly extraction technology through natural plant components, promoting sustainability by minimizing hazardous chemical use while reducing time and energy consumption, with potential applications in cosmetics.

Introduction

Nowadays, there is a global trend towards environmental awareness in the beauty industry, leading manufacturers to focus on green technology for extracting plant components using sustainable alternatives¹. Typically, traditional solvents such as ethanol, methanol, and hexane are used to extract plant phenolic components and natural antioxidants². Nevertheless, the presence of solvent residues within plant extracts poses a potential risk to human health, inducing skin and eye irritation³, particularly concerning their intended application in cosmetics. Consequently, it is challenging to eliminate such solvent residues from the extracts, a process that demands considerable investment in time, energy, and human resources⁴. Recently, superheated water, ionic liquids, deep eutectic solvents, and bio-derived solvents have emerged as promising approaches for green solvent extraction⁵. However, their use is still limited by product separation in aqueous-based processes. To address these challenges, the development of ready-to-use extracts emerges as a viable solution⁶.

Polyols are often used in cosmetic formulations as humectants because of their good polarity and ability to retain moisture from the environment⁷. In addition, polyols such as glycerin, propylene glycol, butylene glycol, methylpropanediol, isopentyldiol, pentylene glycol, 1,2-hexanediol, and hexylene glycol can be utilized for plant extractions. They are considered non-toxic, biodegradable, environmentally friendly, non-reactive, and safe solvents for use in plant extraction⁸. Additionally, polyols can withstand the heat generated during microwave-assisted extraction (MAE) due to their elevated boiling points and polarity⁹. These polyols are generally recognized as safe (GRAS) chemicals by the United States Food and Drug Administration (FDA). Unlike conventional solvents such as ethanol or methanol, which may require rigorous removal from the extract due to their potentially harmful effects, polyols offer the advantage of minimizing the energy, time, and costs associated with solvent removal processes¹⁰. This not only streamlines the extraction process but also enhances the overall efficiency and sustainability of the extraction method. Previous investigations have employed polyols such as propylene glycol and butylene glycol as solvents in the extraction of bioactive compounds from Camellia sinensis flowers¹⁰ and coffee pulp¹¹, revealing significant potential for their role as sustainable alternative solvents in the plant extraction process. Thus, the continued development and optimization of a polyols-water solvent system holds the potential for significant advancements in green chemistry and sustainable industrial practices.

Generally, bioactive compounds found in plants are synthesized as secondary metabolites. These compounds can be categorized into three primary groups: terpenes and terpenoids, alkaloids, and phenolic compounds¹². Various extraction methods are utilized under different conditions to isolate specific bioactive compounds from plants. Bioactive compounds from plant materials can be extracted using either conventional or non-conventional techniques. Traditional methods include maceration, reflux extraction, and hydro-distillation, while non-conventional methods consist of ultrasound-assisted extraction, enzyme-assisted extraction, microwave-assisted extraction (MAE), pulsed electric field-assisted extraction, supercritical fluid extraction, and pressurized liquid extraction¹³. These non-conventional methods are designed to enhance safety by utilizing safer solvents and auxiliaries, improving energy efficiency, preventing degradation of the bioactive components, and reducing environmental pollution¹⁴.

Furthermore, MAE is among the sophisticated green technologies for extracting bioactive compounds from plants. Conventional extraction procedures require significant amounts of time, energy, and high temperatures, which over time might degrade heat-sensitive bioactive compounds¹³. In contrast to conventional thermal extractions, MAE facilitates the extraction of bioactive compounds by generating localized heating within the sample, disrupting cell structures, and enhancing mass transfer, thereby increasing the efficiency of compound extraction. Heat is transferred from inside the plant cells by microwaves, which operate on the water molecules within the plant components¹³. Moreover, MAE has advanced to improve the extraction and separation of active compounds, increasing product yield, enhancing extraction efficiency, requiring fewer chemicals, and saving time and energy while preventing the destruction of bioactive compounds¹⁵.

This research focuses on the extraction of plant phenolic compounds and natural antioxidants through microwave-assisted extraction (MAE) using different types of polyols as solvents. The total phenolic content (TPC), total flavonoid content (TFC), and antioxidant activities (DPPH, ABTS, and FRAP) of polyol-based MAE extracts are determined. Additionally, polyol-based MAE is compared with MAE using conventional solvents such as water and ethanol. This research is expected to contribute to the development of environmentally sustainable extraction technology for natural components, promoting sustainability by reducing reliance on hazardous chemicals, shortening processing times, and minimizing energy consumption in raw material production for potential applications within the cosmetics industry.

Protocol

The details of the reagents and the equipment used in this study are listed in the Table of Materials.

1. Experimental preparation

Plant sample preparation
1. Collect fresh coffee silverskin (Coffea arabica) and dry it at 60 °C in a tray dryer for 72 h¹¹.
2. Grind the dried coffee silverskin (CS) into a fine powder using a grinder and store it at room temperature for further analysis¹¹.
  NOTE: In this study, fresh CS (C. arabica) was collected from Baan Doi Chang, Mae Suai District, Chiang Rai, Thailand. CS is a byproduct obtained during the hulling process that removes the parchment layer from dried coffee beans after the cherries have been processed to remove the outer fruit layers¹⁶.
Chemicals
1. Use chemical reagents of analytical grade quality, except for the solvents in the experiment.
  NOTE: Cosmetic-grade solvents were used in the experiment.
2. Use the solvents (water, ethanol, glycerin, propylene glycol, butylene glycol, hexylene glycol, isopentyldiol, 1-2 hexanediol, pentylene glycol, and methylpropanediol) for the extraction of CS with MAE.

2. Extraction process

Sample and solvent preparation
1. Prepare the sample and solvents for the MAE procedure according to a previously reported protocol⁹ with some modifications.
2. Prepare each solvent at 60% concentration by diluting 60 mL of each solvent with distilled water and adjusting the volume to 100 mL for triplicate extractions.
  NOTE: Use 100% distilled water for water extraction.
3. Weigh 0.67 g of CS and mix with 20 mL of each extraction solvent at a 1:30 ratio in a reaction container (Figure 1A) for MAE.
  NOTE: The maximum solid-liquid amount for each vessel is 2 g of sample and 20 mL of solvent.
4. Add a magnetic stirrer bar to each vessel to ensure uniform distribution of the heat and solvent within the sample, enhancing the efficiency of the extraction process and promoting better extraction yield.
  NOTE: If nonpolar solvents are applied to the extraction process, Teflon stirrer bars can be used instead of magnetic stirrer bars to deliver effective heating through the microwave system.
5. Close each vessel firmly with a special tool (Figure 2) and place all vessels into the MAE chamber (Figure 1B).
Setting up the microwave-assisted extraction instrument and procedure
1. Perform the extraction procedure according to the reference protocol with some modifications⁹.
2. Open the monitor screen to set up the method by clicking on the toolbox icon on the top bar and selecting the SK eT rotor in the accessory section (Figure 3A,B).
3. Select the stirring rate of the stirrer bars by clicking on the stirrer section and typing 20% (Figure 4A).
  NOTE: The stirring rate can be selected from 0% to 100%.
4. Click on the door lock sector and set it to activate at temperatures exceeding 80 °C (Figure 4B).
  NOTE: This setting ensures automatic closure of the chamber door when the internal temperature surpasses 80 °C.
5. Click on the table icon on the top bar (Figure 5A) and set the temperature gradient (T1) to an extraction duration of 10 min, microwave power to 1800 W, and temperature to 120 °C.
6. Activate the stirrer by clicking on the stirrer button until the green light appears.
7. Set the blower fan speed to level 3 (maximum) (Figure 5B).
8. To hold the extraction time, select the desired extraction temperature (T2) by setting the extraction duration to 15 min, the microwave power to 1800 W, and the temperature to 120 °C.
9. Set the stirrer and fan speed as stated in section 2.2.6 and 2.2.7 (Figure 5A,B).
  NOTE: The maximum temperature and microwave power are 260 °C and 1800 W.
10. Set the cooling time by clicking on the cooling button at the lower left corner of the screen and selecting the duration of 10 min (Figure 6).
11. Save the method by clicking on the save icon at the top right corner of the screen (Figure 7A).
12. Ensure that after saving the method conditions, the extraction conditions graph will be displayed on the screen with the play button in the lower right corner (Figure 7B).
13. Start the extraction process by choosing the number of vessels used (Figure 7C).
  NOTE: Up to 15 vessels can be used in one extraction, and if the desired number of vessels is utilized, ensure the balanced placement of vessels in the chamber.
14. Following extraction, centrifuge the extracts at 4 °C, 9072 x g, for 15 min, using a refrigerated centrifuge machine.
15. Collect the supernatant with a 10 mL glass pipette (Figure 8) and store it at -20 °C in the freezer for further study.
  NOTE: Depending on the particle size and density of the plant residue, the extracts will require longer centrifugation times (20-30 min).

3. Determination of phenolic compounds

Determination of total phenolic content
1. Determine the total phenolic content of the CS extracts by referencing the protocol with some modifications¹⁷.
2. Prepare a 10-fold dilution of samples by diluting with distilled water.
3. Mix 10 µL of the diluted sample with 20 µL of undiluted Folin-Ciocalteu's reagent and allow them to react for 3 min.
4. Next, add 100 µL of 7.5% Na₂CO₃solution to the mixture in each well of a 96-well plate.
5. Prepare different concentrations for the gallic acid standard concentration range (please see Table 1 and Table 2) by diluting with distilled water.
6. Mix them with 20 µL of Folin-Ciocalteu's reagent and allow them to react for 3 min.
7. Next, add 100 µL of 7.5% Na₂CO₃solution to the mixture in each well of a 96-well plate.
8. Incubate the reaction for 30 min in the dark at room temperature.
9. Measure the absorbance of the reaction solution at 765 nm using a microplate reader (Figure 9A).
10. Plot the standard calibration curve using the concentrations of the standard and absorbance at 765 nm (Figure 10A).
11. Express the results as mg of gallic acid equivalent (GAE) per g of the sample, and calculate using the following equation¹⁸:
  NOTE: mg of gallic acid equivalent (GAE) per g of sample = [((A₇₆₅– c) / m)) in µg gallic acid equivalent × Total volume in reaction well (mL) x Dilution × weight of dry sample (1 g) x Resultant volume of extract (mL)] / [(Volume of sample added into each well (mL) x Actual weight of dry sample (g) × conversion factor from µg to mg (1000)]
  where, c = y-intercept, m = slope
Determination of the total flavonoid content
1. Determine the total flavonoid content of the CS extract according to the protocol with some modifications¹⁷.
2. Prepare a 5-fold dilution of samples by diluting with distilled water.
3. Add 50 µL of the diluted sample to 15 µL of 5% NaNO₂ and incubate in the dark for 5 min.
4. Mix 15 µL of 10% AlCl₃ solution with the reaction and keep it at room temperature for 6 min.
5. Then, add 100 µL of 1 M NaOH solution to the reaction and incubate for a further 10 min.
6. Measure the absorbance of the mixture at 510 nm (Figure 9B).
7. Prepare different concentrations of quercetin standard range (please see Table 3 and 4) by adding them to 15 µL of 5% NaNO₂ and incubating in the dark for 5 min.
8. Mix 15 µL of 10% AlCl₃ solution with the reaction and keep at room temperature for 6 min.
9. Add 100 µL of 1 M NaOH solution to the reaction and incubate further for 10 min.
10. Determine the absorbance of the standard at 510 nm (Figure 9B).
11. Plot the standard calibration curve using concentrations of the standard and absorbance at 510 nm (Figure 10B).
12. Express the results as mg of quercetin equivalent (QE) per g of the sample, calculated by equation¹⁹ as follows:
  NOTE: mg of quercetin equivalent (QE) per g of sample = [((A₅₁₀– c) / m)) in µg quercetin equivalent × Total volume in reaction well (mL) x Dilution × weight of dry sample (1 g) x Resultant volume of extract (mL)] / [(Volume of sample added into each well (mL) x Actual weight of dry sample (g) × conversion factor from µg to mg (1000)]
  where, c = y-intercept, m = slope

4. Determination of the antioxidant activities

1,1-Diphenyl-2-Picryl-Hydrazil (DPPH) radical scavenging assay
1. Determine the DPPH radical scavenging activity of CS extract according to the protocol with some modifications¹⁷.
2. Prepare a 10-fold dilution of the samples by diluting them with distilled water.
3. Mix 20 µL of the diluted samples with 135 µL of 0.1 mM DPPH solution.
4. Prepare different concentrations of the Trolox standard concentration range (please see Table 5 and 6) by mixing with 135 µL of 0.1 mM DPPH solution.
5. Incubate the mixture in the dark at room temperature for 30 min.
6. Measure the absorbance of the resultant at 517 nm (Figure 9C).
7. Plot the standard calibration curve using concentrations of the standard and % inhibition (Figure 10C).
8. Calculate the % inhibition of the DPPH assay as follows:
  % Inhibition = [(absorbance of control − absorbance of sample)/ absorbance of control] × 100
9. Express the results as mg of the Trolox equivalent antioxidant capacity per g of the sample, calculated by the following equation²⁰:
  NOTE: mg of Trolox equivalent (TE) per g of sample = [((% inhibition-c) / m) in µg Trolox equivalent × Total volume in reaction well (mL) x Dilution × weight of dry sample (1 g) x Resultant volume of extract (mL)] / [(Volume of sample added into each well (mL) x Actual weight of dry sample (g) × conversion factor from µg to mg (1000)]
  where, c = y-intercept, m = slope
2,2′-Azino-Bis-3-Ethylbenzthiazoline-6-Sulphonic acid (ABTS) radical scavenging assay
1. Determine the ABTS radical scavenging activity of CS extract using the protocol from the reference with some modifications¹⁷.
2. Prepare the ABTS·+ stock solution by mixing 7 mM ABTS and 2.45 mM potassium persulfate (1:2) and incubate in the dark at room temperature for 16 h.
3. Prepare the working solution by mixing 5 mL of ABTS·+ stock solution with 100 mL of deionized water.
4. Mix 160 µL of ABTS·+ working solution with 10 µL of the 10-fold diluted sample or Trolox standard at different concentrations (see Table 7 and Table 8).
5. Incubate the reaction in the dark at room temperature for 30 min.
6. Determine the absorbance of the mixture at 734 nm (Figure 9D).
7. Plot the standard calibration curve using concentrations of the standard and % inhibition (Figure 10D).
8. Calculate % inhibition of the ABTS assay using the following formula:
  % Inhibition = [(absorbance of control – absorbance of sample) / absorbance of control] × 100.
  1. Express the results as mg of the Trolox equivalent antioxidant capacity per g of the sample, calculated using the following equation²¹:
    NOTE: mg of trolox equivalent (TE) per g of sample = [((% inhibition – c) / m)) in µg Trolox equivalent × Total volume in reaction well (mL) x Dilution × weight of dry sample (1 g) x Resultant volume of extract (mL)] / [(Volume of sample added into each well (mL) x Actual weight of dry sample (g) × conversion factor from µg to mg (1000)]
    where, c = y-intercept, m = slope
Ferric Reducing Antioxidant Power Assay (FRAP)
1. Determine the ferric-reducing antioxidant activity of CS extract according to the protocol with some modifications¹⁷.
2. Prepare the FRAP reagent using a 30 mM acetate buffer at pH 3.6, which is a mixture of 10 mM TPTZ solution in 40 mM HCl and 20 mM FeCl₃.6H₂O solution at a ratio of 10:1:1.
3. Place the FRAP reagent into an amber bottle until required.
  NOTE: Ensure the FRAP reagent is brown. If contaminated by metal ions or other reactive compounds, the reagent will turn purple and must be discarded. Use only freshly prepared reagents.
4. Prepare a 5-fold dilution of the samples by diluting them with distilled water.
5. Add 10 µL of the diluted sample or 20 µL of FeSO₄.7H₂O at different standard concentrations (Table 9 and Table 10) to 180 µL of the FRAP solution.
6. Incubate the reaction at room temperature for 4 min.
7. Evaluate the absorbance of the mixture at 593 nm (Figure 9E).
8. Plot the standard calibration curve using concentrations of the standard and absorbance at 593 nm (Figure 10E).
9. Express the results as mg of FeSO₄ per g of the sample, calculated using the following equation²¹:
  NOTE: mg FeSO₄equivalent (Fe (II) E) per g sample = [((A₅₉₃– c) / m)) in µg FeSO₄equivalent × Total volume in reaction well (mL) x Dilution × weight of dry sample (1 g) x Resultant volume of extract (mL)] / [(Volume of sample added into each well (mL) x Actual weight of dry sample (g) × conversion factor from µg to mg (1000)]
  where, c = y-intercept, m = slope
Perform all assays (TPC, TFC, DPPH, ABTS, and FRAP) of each sample in triplicate. In this study, water was used as the blank for most assays, except DPPH, where ethanol served as the blank to address background absorbance.

5. Statistical analysis

Use SPSS software to carry out a statistical analysis of the experimental data.
Conduct the normality test using the Shapiro-Wilk test.
Compare the bioactive substances and antioxidant activities of polyols-based MAE CS extract and conventional solvents-based MAE CS extracts using one-way ANOVA with Duncan's multiple range tests.
Express all data as mean ± SD (n = 3) and define the significance level at p < 0.05.

Representative Results

Effect of polyols solvents and conventional solvents on total phenolic content, total flavonoid content, DPPH, FRAP, and ABTS antioxidant assays
Solvent polarity should be compatible with that of targeted active molecules to improve the extraction efficiency of bioactive substances from plants²². Experiments were conducted using various solvents (water, ethanol, glycerin, propylene glycol, butylene glycol, methylpropanediol, isopentyldiol, pentylene glycol, 1,2-hexanediol, and hexylene glycol) to assess their impact on the bioactive compounds and antioxidant activities of MAE coffee silverskin extract.

Effect of polyols solvents and conventional solvents on total phenolic content
The total phenolic content of each extraction with different solvents was analyzed. The highest phenolic content was yielded in samples with aqueous-1,2-hexanediol (52.0 ± 3.0 mg GAE/g sample), while the lowest TPC was revealed in samples with water extraction (31.4 ± 4.3 mg GAE/g sample), and these values were significantly different from those of all other conditions. The samples with aqueous-pentylene glycol yielded the second-highest TPC value, followed by samples with aqueous-butylene glycol, methylpropanediol, and other solvent systems (Figure 11A). When comparing samples with conventional solvents (water and aqueous-ethanol system) and samples with polyols-based solvents, significant differences in TPC values can be observed (p < 0.05).

Effect of polyols solvents and conventional solvents on total flavonoid content
The total flavonoid content of each extraction with different solvents was analyzed. The highest flavonoid content was yielded in samples with aqueous-1,2-hexanediol (20.0 ± 1.7 mg QE/g sample), demonstrating a significant difference from that of all other extracts. The samples with aqueous-isopentydiol revealed the lowest TFC value (8.8 ± 0.7 mg QE/g sample), which was not significantly different from aqueous-methyl propanediol, and aqueous-ethanol extracts. Moreover, the second highest TFC value was found in the sample with aqueous-pentylene glycol, followed by aqueous-hexylene glycol, aqueous-propylene glycol, aqueous-butylene glycol, and aqueous-glycerin (Figure 11B).

Effect of polyols solvents and conventional solvents on antioxidant assays
The antioxidant activities of the extracts with polyols and conventional solvents were evaluated using DPPH, ABTS, and FRAP assays. The highest value for the DPPH assay was measured in samples with aqueous-hexylene glycol (13.6 ± 0.3 mg TE/g sample) and the lowest in samples with aqueous-ethanol (4.5 ± 0.2 mg GAE/g sample), and these values were significantly different from other extracts (p < 0.05). The second highest DPPH values were observed in samples with aqueous-1,2-hexanediol, followed by aqueous-pentylene glycol, aqueous-methyl propanediol, and other solvent systems (Figure 11C).

The highest ABTS value was measured in samples with aqueous-pentylene glycol (8.2 ± 0.1 mg TE/g sample) and the lowest in samples with water (5.6 ± 0.04 mg GAE/g sample), and these values were significantly different from other extracts (p < 0.05). The second highest ABTS values were detected in aqueous-butylene glycol and aqueous-1,2-hexanediol, followed by samples with aqueous-glycerin, aqueous-methyl propanediol, and other solvent systems (Figure 11D).

The highest FRAP values were observed in samples with aqueous-hexylene glycol (21.1 ± 1.3 mg Fe (II) E/g sample and the lowest in water extraction (11.5 ± 0.2 Fe (II) E/g sample), with these values being significantly different (p < 0.05) for the remaining solvents. Moreover, the second highest FRAP values were found in samples with aqueous-pentylene glycol, followed by aqueous-butylene glycol, aqueous-glycerin, and other solvent systems (Figure 11E).

When comparing the antioxidant activities of samples with conventional solvents (water and aqueous ethanol), those containing polyols exhibited significantly higher antioxidant activities in all antioxidant assays (DPPH, ABTS, and FRAP) (p < 0.05).

Figure 1: Reaction in experimental containers and the MAE chamber. (A) Sample and solvent are added to the white inter-layer vessel of a Teflon container before extraction. (B) Each container is placed inside the microwave chamber before starting the extraction. Please click here to view a larger version of this figure.

Figure 2: Special tools for closing the reaction vessels. After adding the sample and solvent to the Teflon container, the lids are applied to the top of the container, placed in the vessel holder, and fastened tightly using the tools. Please click here to view a larger version of this figure.

Figure 3: Extraction method. (A) Extraction method, created by entering the method section. (B) The SK eT accessory is applied for the MAE process. Please click here to view a larger version of this figure.

Figure 4: Stirring rate and door lock function setting. (A) The magnetic stirrer bars inside each vessel can be activated by choosing the stirring rate. (B) The door lock function limits the temperature, allowing the chamber to be opened after extraction. Please click here to view a larger version of this figure.

Figure 5: Setting the extraction conditions. (A) Entering the table icon and setting the extraction conditions such as time, temperature, and microwave power. (B) Opening the stirrer button and choosing the blower speed. Please click here to view a larger version of this figure.

Figure 6: Setting the cooling time. Applying the cooling time to reduce the inside temperature in the MAE chamber. Please click here to view a larger version of this figure.

Figure 7: Starting the extraction process. (A) Saving the method created for extraction. (B) Clicking the play icon to start the extraction process. (C) Choosing the number of vessels to start the extraction. Please click here to view a larger version of this figure.

Figure 8: Picture of the final extract after extraction using MAE. Obtaining the supernatant after centrifuging. Please click here to view a larger version of this figure.

Figure 9: The 96 well-plates for determining the TPC, TFC, DPPH scavenging activity, ABTS scavenging activity and FRAP assay of the extracts. (A) Determining the TPC for the gallic acid standard plate from a concentration of 2.5-75 µg/mL and sample extracts. (B) Determining the TFC for the quercetin standard plate from concentrations of 2.5-50 µg/mL and TFC assay to measure sample extracts. (C) Determining the DPPH scavenging activity for the Trolox standard plate from concentrations of 0.25-12.5 µg/mL and the DPPH scavenging activity detection plate of sample extracts. (D) Determining the ABTS scavenging activity for the Trolox standard plate from concentrations of 0.25-5 µg/mL and the ABTS scavenging activity detection plate of sample extracts. (E) Determining the FRAP assay for the FeSO₄ standard plate from concentrations of 0.25-10 µg/mL and the FRAP assay detection plate of sample extracts. Please click here to view a larger version of this figure.

Figure 10: The standard calibration curves for the TPC, TFC, DPPH scavenging activity, ABTS scavenging activity, and FRAP assay. (A) The standard curve for determining TPC, plotted by concentrations of the gallic acid standard and absorbance at A₇₆₅. (B) The standard curve for determining TFC, plotted by concentrations of the quercetin standard and absorbance at A₅₁₀. (C) The standard curve for determining DPPH scavenging activity, plotted by concentrations of the Trolox standard and % inhibition. (D) The standard curve for determining ABTS scavenging activity, plotted by concentrations of the Trolox standard and % inhibition. (E) The standard curve for measuring the FRAP assay, plotted by concentrations of the ferrous sulfate standard and absorbance at A₅₉₃. Please click here to view a larger version of this figure.

Figure 11: The effect of solvent types on the TPC, TFC, DPPH scavenging activity, ABTS scavenging activity, and FRAP assay in the MAE of coffee silverskin. (A) The effect of solvent types on total phenolic content. (B) The effect of solvent types on total flavonoid content. (C) The effect of solvent types on DPPH scavenging activity. (D) The effect of solvent types on ABTS scavenging activity. (E) The effect of solvent types on the FRAP assay. Values are indicated as Mean ± SD (n = 3). Values with different superscript letters express a statistically significant difference (p < 0.05). Please click here to view a larger version of this figure.

Table 1: Gallic acid standard curve preparation. Preparing the standard concentration range of 2.5-75 µg/mL in the 96-well plate. B = blank, 1-7 = number of wells on the 96-well plate. Please click here to download this Table.

Table 2: Final concentration calculation of gallic acid standards. Preparing the standard concentration range of 2.5-75 µg/mL. Final concentrations (µg/mL) of gallic acid are calculated accordingly. Final concentration (µg/mL) = (Initial concentration (mg/ mL) × Initial volume (µL) / final volume (µL)) × (1000 µg/1 mg). Please click here to download this Table.

Table 3: Quercetin standard curve preparation. Preparing the standard concentration range of 2.5-50 µg/mL in the 96-well plate. B = blank, 1-7 = number of wells on the 96-well plate. Please click here to download this Table.

Table 4: Final concentration calculation table for quercetin standards. Preparing the standard concentration range of 2.5-50 µg/mL. Final concentrations (µg/mL) of quercetin are calculated accordingly. Final concentration (µg/mL) = (Initial concentration (mg/mL) × Initial volume (µL) / final volume (µL)) × (1000 µg/1 mg). Please click here to download this Table.

Table 5: Trolox standard curve preparation in the concentration range of 0.25-12.5 µg/mL. Preparing the standard concentration range of 0.25-12.5 µg/mL in the 96-well plate. B = blank, C = control, 1-7 = number of wells on the 96-well plate. Please click here to download this Table.

Table 6: Final concentration calculation of the Trolox standards for DPPH assay. Preparing the standard concentration range of 0.25-12.5 µg/mL including the final concentrations (µg/mL) of Trolox. Final concentration (µg/mL) = (Initial concentration (mg/mL) × Initial volume (µL) / final volume (µL)) × (1000 µg/1 mg). Please click here to download this Table.

Table 7: Trolox standard curve preparation in the concentration range of 0.25-5 µg/mL. Preparing the stock standard concentration range of 0.25-5 µg/mL in the 96-well plate. B = blank, C = control, 1-7 = number of wells on the 96-well plate. Please click here to download this Table.

Table 8: Final concentration calculation of the Trolox standards for ABTS assay. Preparing the standard concentration range of 0.25-5 µg/mL, including the final concentrations (µg/mL) of Trolox. Final concentration (µg/mL) = (Initial concentration (mg/mL) × Initial volume (µL) / final volume (µL)) × (1000 µg/1 mg). Please click here to download this Table.

Table 9: Final concentration calculation table for the FeSO₄ standards. Preparing the preparation of standard concentration range of 2.5-100 µg/mL, including the final concentrations (µg/mL) of FeSO₄. Final concentration (µg/mL) = (Initial concentration (mg/mL) × Initial volume (µL) / final volume (µL)) × (1000 µg/1 mg). Please click here to download this Table.

Table 10: FeSO₄ standard curve preparation. Preparing the standard concentration range of 0.25-10 µg/mL in the 96-well plate. B = blank. Please click here to download this Table.

Discussion

Various factors play a crucial role in the successful implementation of MAE, such as the phytochemical content of plant components, extraction duration, temperature, microwave power, solid-liquid ratio, and solvent concentration¹³. Plants typically exhibit varying profiles of phytochemicals; hence, the selection of natural plants rich in antioxidants and phenolic compounds is essential²³. Furthermore, distinct bioactive constituents display a variety of polarities depending on the solvent used. Likewise, solvents exhibit differing polarities. Given that the polarity of solvents plays a crucial role in determining the efficacy of bioactive compound extraction from raw materials, it is imperative that the solvent's polarity aligns with that of the targeted bioactive molecules²⁴.

In this study, various polyols were utilized to extract polyphenols and antioxidants from coffee silverskin using MAE. Polyphenols are predominantly polar, and solvents with high polarity typically enhance the yield of phenolic compounds²⁵. Compared to samples with water and ethanol, those employing different polyols demonstrated higher efficiency in all measured responses, including TPC, TFC, and antioxidant assays such as DPPH, ABTS, and FRAP. Previous studies support the findings that aqueous-polyol mixtures can enhance the extraction yield of bioactive compounds in comparison to aqueous-ethanol mixtures⁹^,¹⁰^,¹¹. When comparing different polyols, samples with a specific aqueous-polyol system failed to yield the highest value. However, it is interesting to note that samples with aqueous 1,2-hexanediol yielded the highest values in TPC and TFC assays. Meanwhile, those with aqueous-hexylene glycol yielded the highest values in DPPH and FRAP assays, and aqueous-pentylene glycol extract exhibited the highest values in the ABTS assay. The variation in values obtained from different assays such as TPC, TFC, DPPH, ABTS, and FRAP within aqueous-polyols systems can be attributed to several factors, including the distinctive characteristics of the polyols employed. Polyols exhibit differences in viscosity, polarity, and boiling points, directly influencing their efficacy in extracting bioactive compounds from plant materials²⁶. One potential explanation may be attributed to the principle of "Like dissolves like", wherein the particular solvent system is the most appropriate for facilitating the mass transfer of particular bioactive compounds²⁷. This underscores the importance of selecting a solvent with a polarity that matches that of the targeted bioactive compound.

Another possible reason might be the fact that the difference in the number of hydroxyl groups (-OH) present in the solvent significantly impacts the yield of phenolic compounds²⁸. Among these polyols, only glycerin contains three -OH groups, while the remaining polyols in this study contain two -OH groups. Solvents with a higher number of -OH groups tend to exhibit higher viscosity compared to those with fewer²⁹. Elevated viscosity can impede the efficient transfer of active compounds during the extraction process, thereby reducing the overall yield. Furthermore, the dielectric constant of a solvent, closely linked to its polarity, plays a crucial role in determining its ability to dissolve polar or nonpolar solutes. Solvents with higher dielectric constants are more adept at dissolving polar solutes, and those with lower better suited to nonpolar solutes³⁰. Among polyols, glycerin exhibits a comparatively high dielectric constant of 41.14, while hexylene glycol, pentylene glycol, and 1,2-hexanediol demonstrate lower dielectric constants of 25.86, 17.31, and 15.45, respectively³¹^,³². The findings of this study suggest that the bioactive compounds within the sample may encompass low-polar constituents.

Extraction efficiency can be enhanced by optimizing the selection and composition of solvents, and further experimentation may be required to identify the most suitable solvent system. Although the investigation exhibits potential, it is limited by its sole focus on microwave-assisted extraction with polyols and its restricted evaluation of other variables, including extraction duration, temperature, solvent concentration, solid-liquid ratio, and extraction power. In addition, a mechanistic study is necessary to understand how polyols function due to their varied dielectric constants, directly impacting their solubility of polar or nonpolar solutes. Differences in dielectric constants among polyols highlight the importance of investigating their specific mechanisms in extracting bioactive compounds. Such research would offer valuable insights into solvent-solute interactions, aiding the optimization and selection of solvent systems for efficient extraction processes.

Regarding the MAE process, there are some limitations. While MAE can provide rapid heating, precise temperature control may be challenging, potentially leading to overheating and the degradation of thermally sensitive compounds³³. However, the microwave power setting for gradient and extraction temperature can be set to the same microwave power to avoid temperature frustration during extraction. Additionally, MAE has limitations over heat-sensitive plant components. However, advanced Ethos X MAE technology can minimize the risk of deterioration by supporting effective heating at a shorter duration using dielectric heating³⁴. Each vessel of the MAE chamber has its own limited maximum solid and liquid capacity. The solid-liquid ratio over this limitation can also greatly influence the concentration of the extracts¹¹. The dissolution between solvent and solute can be promoted using stirrer bars, potentially leading to effective extraction and higher yield³⁵. Furthermore, the extracted phenolic and flavonoid compounds within the extracts can be verified through additional analysis, such as Liquid Chromatography Triple Quadrupole Mass Spectrometry (LC-QQQ) and Liquid Chromatography Quadrupole Time-of-Flight Mass Spectrometry (LC-QTOF), to establish the presence of specific bioactive compounds and their respective quantities³⁶.

The extraction of polyphenols and antioxidants from coffee silverskin using aqueous polyols through MAE demonstrated higher efficiency in comparison to water and aqueous-ethanol extracts. Based on the outcomes derived from the polyols-based MAE of CS extracts, it has been observed that the employment of the aqueous-hexylene glycol, aqueous-1,2-hexanediol, and aqueous-pentylene glycol systems resulted in significantly higher extraction yields of bioactive compounds and antioxidant capacities. Moreover, these findings underscore the potential of utilizing such extracted compounds for subsequent investigative analyses. The utilization of polyols as green solvents for bioactive compound extraction from plant materials through MAE promises environmental benefits and enhanced bioactive compound yield, offering a sustainable approach with the potential for use in cosmetic applications.

Disclosures

The authors have nothing to disclose.

Acknowledgements

This study was funded by Mae Fah Luang University. The authors would like to acknowledge the Tea and Coffee Institute of Mae Fah Luang University for facilitating the connection between the researchers and local farmers concerning the acquisition of coffee silverskin samples.

Materials

1,2-Hexanediol	Chanjao Longevity Co., Ltd.
2,2 -Azino-bis 3 ethylbenzothiazoline-6-sulfonic acid diammonium salt (ABTS)	Sigma	A1888
2,2-Diphenyl-1-picrylhydrazyl (DPPH)	Sigma	D9132
2,4,6-Tri(2-pyridyl)-s-triazine (TPTZ)	Sigma	93285
2-Digital balance	Ohaus		Pioneer
4-Digital balance	Denver	SI-234
6-hydroxy-2,5,7,8 tetramethylchroman -2-carboxylic acid (Trolox)	Sigma	238813
96-well plate	SPL Life Science
Absolute ethanol	RCI Labscan	64175
Acetic acid	RCI Labscan	64197
Aluminum chloride	Loba Chemie	898
Automatic pipette	Labnet		Biopett
Butylene glycol	Chanjao Longevity Co., Ltd.
Ethos X advanced microwave extraction	Milestone Srl, Sorisole, Italy
Ferrous sulfate	Ajex Finechem	3850
Folin-Ciocalteu's reagent	Loba Chemie	3870
Freezer SF	Sanyo	C697(GYN)
Gallic acid	Sigma	398225
Grinder	Ou Hardware Products Co.,Ltd
Hexylene glycol	Chanjao Longevity Co., Ltd.
Hydrochloric acid (37%)	RCI Labscan	AR1107
Iron (III) chloride	Loba Chemie	3820
Isopentyldiol	Chanjao Longevity Co., Ltd.
Methanol	RCI Labscan	67561
Methylpropanediol	Chanjao Longevity Co., Ltd.
Pentylene glycol	Chanjao Longevity Co., Ltd.
Potassium persulfate	Loba Chemie	5420
Propylene glycol	Chanjao Longevity Co., Ltd.
Quercetin	Sigma	Q4951
Refrigerated centrifuge	Hettich
Sodium acetate	Loba Chemie	5758
Sodium carbonate	Loba Chemie	5810
Sodium hydroxide	RCI Labscan	AR1325
Sodium nitrite	Loba Chemie	5954
SPECTROstar Nano microplate reader	BMG- LABTECH
SPSS software	IBM SPSS Statistics 20
Tray dryer	France Etuves	XUE343

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