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

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

1. 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.
2. 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

1. 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).
2. 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

1. 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% Na₂CO₃ solution 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% Na₂CO₃ solution 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
2. 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. 1,1-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. 2,2′-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
3. 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 FeSO₄ equivalent (Fe (II) E) per g sample = [((A₅₉₃ - c) / m)) in µg FeSO₄ 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. 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

1. Use SPSS software to carry out a statistical analysis of the experimental data.
2. Conduct the normality test using the Shapiro-Wilk test.
3. 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.
4. Express all data as mean ± SD (n = 3) and define the significance level at p < 0.05.

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...

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...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
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