Sustainable pectin extraction from Riang husk using ultrasound-assisted extraction with deep eutectic solvents and its potential in antipollution products

Graphical abstract

Highlights

• •Ultrasound-assisted extraction combined with deep eutectic solvents (DESs) extracted highquality pectin from Riang husks.
• •Riang husk pectin expressed cytoprotective effects against hydrogen peroxide-induced oxidative stress in HaCaT cells.
• •Riang husk pectin effectively inhibited cellular damage induced by PM2.5.
• •Riang husk pectin had potential as a valuable ingredient in anti-pollution products.

Abstract

With increasing concerns about air pollution and its adverse effects on health, particularly in Thailand, the demand for antipollution products has risen significantly. Parkia timoriana (DC.) Merr., commonly known as Riang, has emerged as a promising source for developing antipollution products due to its characteristics. This study investigates the use of ultrasound-assisted extraction (UAE) combined with deep eutectic solvents (DESs) as a sustainable and efficient method for optimizing pectin extraction from Riang husks through the evaluation of a central composite design (CCD), and the structural, functional, and rheological characteristics of the extracted pectin. The antioxidant activity and protective effects against PM2.5-induced cellular damage of this method were also evaluated. The condition that exhibited the highest yield were found to be a liquid-to-solid ratio of 40 mL/g, 35 % amplitude (ultrasonic power of 28.11 W), and 60 min of extraction time. The extracted pectin was primarily composed of monosaccharides, including galacturonic acid (53.74 %), arabinose (23.97 %), galactose (12.36 %), and rhamnose (6.81 %). The degree of esterification (DE) was 73.41 %, classifying it as high methoxyl pectin. Functionally, the pectin demonstrated a solubility of 53 %, a water holding capacity of 3.88 g water/g pectin, an oil holding capacity of 3.30 g oil/g pectin, and a swelling capacity of 11.77 mL/g. Rheological analysis showed shear-thinning behavior across all pH gel forms. Furthermore, Riang husk pectin exhibited antioxidant activity, measured at 0.26 ± 0.02 mmol Trolox equivalents/g, and demonstrated cytoprotective effects against hydrogen peroxide-induced oxidative stress. It also attenuated damage caused by PM2.5 in HaCaT cells. The current study highlights UAE combined with DESs as a sustainable and effective method for obtaining high-quality pectin, contributing to the development of antipollution products and supporting sustainability goals.

1. Introduction

In recent years, antipollution products have gained significant popularity along with the growing recognition of the harmful impact on health caused by environmental pollutants. This change in consumer behavior reflects growing concerns about air quality and its impact, with research consistently finding that pollutants such as particulate matter, nitrogen dioxide, and other compounds are linked to health issues, including respiratory and heart diseases, as well as premature aging [1], [2]. These concerns are particularly relevant in Thailand, which is currently facing a significant public health crisis due to the concerning levels of particulate matter 2.5 (PM2.5) in the atmosphere, especially during the seasonal transition from winter to summer [3]. PM2.5, comprising airborne particles with diameters of less than 2.5 µm, can deeply penetrate the lungs, leading to serious long-term health risks and carrying harmful pollutants like carcinogens and heavy metals [4]. PM2.5 also poses a threat to skin health by inducing oxidative stress, causing cellular damage and irritation [5], [6], [7]. This situation underscores the urgent need for effective antipollution measures and has further increased the demand for products designed to mitigate the health impacts of pollution.

Addressing these pollution-related challenges, Parkia timoriana (DC.) Merr., known as Riang, is emerging as a promising source for cosmetic products due to its significant pectin content and potential health benefits. Native to southern Thailand and other Southeast Asian regions, Riang pods are traditionally used for their seeds, which are consumed, with the remaining pods often used as fertilizer. However, recent research has highlighted the untapped potential of Riang pod husks, which contain 15 % pectin [8]. Pectin, a polysaccharide with known antioxidative and anti-inflammatory properties, is highly valued in various industries for its capacity to create gels, stabilize products, and serves as an emulsifier, which makes it valuable in the food, cosmetic, and biomedical industries [8]. Additionally, pectin’s structural characteristics endow it with antioxidative and anti-inflammatory properties, which are crucial for developing products aimed at mitigating pollution-related skin damage [9], [10], [11]. Moreover, the ability of pectin to adsorb heavy metals makes it a promising biosorbent for combating pollution-induced toxicity [12]. However, traditional pectin extraction methods from Riang, which rely on high-temperature acid processes, are labor-intensive, time-consuming, and environmentally harmful [13].

To maximize the potential of Riang-derived pectin, it is crucial to create more efficient and sustainable extraction methods. Among advanced extraction methods, ultrasound-assisted extraction (UAE) combined with deep eutectic solvents (DESs) could potentially be a sustainable and efficient approach. Ultrasound-assisted extraction utilizes acoustic energy to enhance the extraction process through cavitation [14], reducing energy consumption, accelerating extraction times, and improving the quality of pectin [15], [16]. Meanwhile, DESs, known for their sustainability and eco-friendliness, provide a substitute for conventional solvents, being harmless while offering low volatility and excellent biodegradability [17], [18]. Although DESs may have some limitations in the scale-up process for industrial use, their application is expanding, especially in the cosmetic industry, due to the increasing preference for natural products.

Despite the growing use of DESs in cosmetic industries, previous studies have used water and acid solvents to examine pectin yield from various parts of the Riang plant [8], [13], while the application of UAE using DESs for extracting pectin from husks in the pods of Riang plant remains underexplored. Thus, this study aims to evaluate the impact of UAE with DESs, maximize the factors influencing Riang husk pectin yield by applying response surface methodology (RSM), analyze the structural characteristics of Riang husk pectin, determine its functional and rheological properties, examine the effects of pectin on cell toxicity and cellular antioxidant activity, and evaluate its protective potential on PM2.5-induced cellular damage. This research seeks to unlock the full potential of Riang husk pectin to develop antipollution products that align with sustainability goals and address the pressing need for effective solutions to mitigate pollution-related health impacts.

2. Methodology

2.1. Reagents and chemicals

Absolute ethanol, citric acid, acetic acid and 98 % sulfuric acid were purchased from RCI-Labscan (Bangkok, Thailand). Choline chloride was obtained from Loba Chemie (Tarapur, India). Betaine and glycerol were purchased from Chanjao Longevity (Bangkok, Thailand). Oxalic acid and malic acid were purchased from KemAus (New South Wales, Australia). D-(+)-Galacturonic acid monohydrate and carbazole were purchased from Merck (Darmstadt, Germany). Ascorbic acid was purchased from Loba Chemie (Tarapur, India). The other reagents used were of analytical grade.

2.2. Riang husk preparation

Ripe, dry, blackish Parkia timoriana (DC.) Merr pods were sourced from a local market in Surat Thani, Thailand. The husks of the Riang pods, which form the outer layer, were thoroughly cleaned to eliminate dust and impurities. After the cleaned husks had been dried at 60 °C for 72 h, they were finely ground, and the resulting powder was sieved through a 30-mesh sieve using a sieve shaker (AS200, USA). The powder was then kept at room temperature until further use for extraction. Prior to pectin extraction, impurities were eliminated following a modified outline adapted from Chaiyasut et al. [19]. Ethanol of 95 % (2 mL) was added to the husk powder (1 g) and boiled at 70 °C for 10 min. After centrifugation at 4,500 rpm for 15 min (min), the pellet was collected. To ensure thorough impurity removal, 30 % ethanol (20 mL) was used to treat the pellet, followed by vortexing and a second round of centrifugation. The purified pellet was subsequently collected and prepared for pectin extraction.

2.3. Preparation of PM2.5

The PM2.5 filters were supplied by the Environmental Science Research Center, Faculty of Science, Chiang Mai University, Chiang Mai. During the dry season of 2019, the researchers collected ambient PM2.5 samples following the procedure outlined by Kraisitnitikul et al. [20]. The preparation of PM2.5 followed a modified protocol based on Li et al. [21]. The particles were extracted by immersing the filters in distilled water (5 mL) and subjecting the mixture to sonication with ultrasonic frequency 37 kHz for 30 min at room temperature using an ultrasonic bath (S 70H ELMA, Germany). The sonication procedure was triplicated, and the resulting suspensions were collected. Using a vacuum, the resulting suspension was freeze-dried, and the concentrated fractions were kept at −20 °C for subsequent experiments.

2.4. Deep eutectic solvents (DESs) preparation

The DESs were formulated by combining their components in defined molar ratios of 1:2, 1:3, 1:4, 1:5, and 2: 1, followed by stirring at room temperature as described in previous studies [18]. The DESs used in this experiment consisted of combinations of betaine (Be), oxalic acid (OA), choline chloride (ChCl), malic acid (MA), acetic acid (AA), citric acid (CA), and glycerol (G).

2.5. Ultrasound-assisted extraction

Riang husk pectin was extracted using an ultrasonic probe (6 mm) processor (VCX 130, Vibra Cell, Sonics, USA) with a frequency of 20 kHz, following the protocol outlined by Chen et al. [17]. The powdered husk 0.5 g was initially mixed with different volumes (10 – 25 mL) of DES solutions in a centrifuge tube placed in an ice bath to control the high temperature occurring during the sonication process, ensuring a 2 cm gap between the bottom of the tube and ultrasound probe. Sonication was carried out using a 50 % duty cycle, alternating between 10 s of sonication and 10 s to keep the temperature constant. After sonication, the mixture underwent centrifugation at 4,500 x g at 4 °C for 15 min to collect the supernatant. The supernatant was then combined with a two-fold volume of 95 % ethanol and refrigerated at 4 °C overnight. Following refrigeration, the mixture was centrifuged again at 10,000 x g for 15 min, with 95 % ethanol used to rinse the resultant pectin precipitate to eliminate any residual solvents. Finally, the Riang husk pectin was freeze-dried and stored at −20 °C. The pectin yield was determined as follows:

$$\mathit{Pectin}yield(\%)=(driedRianghuskpectinweight/Rianghuskpowderweight)\times 100$$

2.6. Varying a single factor at a time

Five independent variables and their respective levels were systematically screened by varying a single factor at a time to determine the most appropriate factors for pectin extraction. The experimental factors included DESs, DES ratios, sonication amplitude, extraction time, and liquid–solid (L/S) ratio.

The DESs were prepared using initial ratios drawn from literature sources [17], [22], [23]. The DES combinations tested were Be: CA (1:2), Be: OA (1:2), Be: MA (1:2), Be: AA (1:3), Be: G (1:2), ChCl: CA (2:1), ChCl: AA (1:3), ChCl: MA (1:2), ChCl: OA (1:1), and ChCl: G (1:2). The extraction parameters were set to 30 % amplitude (ultrasonic power of 26.10 W), an L/S ratio of 30:1 mL/g, and 30 min of extraction time. The DES yielding the highest pectin was selected for further experimentation.

The DES ratios of Be and CA, ranging from 1:2, 1:3, 1:4, to 1:5, were tested. The extraction parameters were set to an amplitude of 30 % (ultrasonic power of 26.10 W), an L/S ratio of 30:1 mL/g, and an extraction time of 30 min. The DES ratio yielding the highest pectin was selected for further experimentation.

The various sonication amplitudes tested were 20 % (ultrasonic power of 22.79 W), 25 % (ultrasonic power of 23.85 W), 30 % (ultrasonic power of 26.10 W), 35 % (ultrasonic power of 28.11 W), and 40 % (ultrasonic power of 30.13 W). The ultrasonic power was assessed utilizing a Smart Universal Plug (HS-SUP10A, Haco Electric, Thailand) in compliance with the manufacturer's instructions. The extraction parameters were set with a DES comprising Be and CA at a 1:5 ratio, an L/S ratio of 30:1 mL/g, and an extraction time of 30 min. The amplitude exhibiting the highest pectin yield was selected for further experimentation.

The various extraction times tested were 15, 30, 45, 60, and 90 min. The extraction parameters were set with a DES comprising Be and CA at a 1:5 ratio, 35 % amplitude (ultrasonic power of 28.11 W), and an L/S ratio of 30:1 mL/g. The extraction time exhibiting the highest pectin yield was selected for further experimentation.

The L/S ratios tested were 10:1, 20:1, 30:1, 40:1, and 50:1 mL/g. The extraction parameters were set with a DES comprising Be and CA at a 1:5 ratio, 35 % of amplitude (ultrasonic power of 28.11 W), and an extraction time of 90 min.

The pectin yield from the Riang husk was assessed to identify the independent variables significantly impacting the efficiency of the extraction process. In addition, to monitor the actual temperature range during the probe sonication process, a thermometer was inserted directly into each sample solution. Temperature readings were taken at regular intervals throughout sonication, and the average temperature for each condition was calculated to represent the typical temperature during each experimental run. From these averages, the overall range of minimum and maximum temperatures observed across all variations of each independent factor was reported.

2.7. Optimization process for Riang husk pectin extraction

After completing the varying single-factor-at-a-time experiments, the key factors and their optimal levels significantly impacting pectin yield were identified and selected for further optimization using central composite design (CCD) through response surface methodology (RSM). The independent variables (L/S ratio, duration of extraction, and sonication amplitude) were coded, with their alpha values and levels presented in Table 1. The experiment consisted of non-center points (14 runs) and center points (6 runs), including the plus and minus alpha (axial) points, as shown in Table 2. A total of 20 experiments were conducted to develop mathematical models for predicting pectin yield. The association between the pectin yield (Y) and the independent variables was modeled using RSM, represented as follows:

Table 1.

The independent variables used in CCD with encoded and actual values.

Independent variables	Label	Levels
		−α	−1	0	1	+α
L/S ratio (mL/g)	X1	13.18	20	30	40	46.82
Extraction time (min)	X2	39.55	60	90	120	140.45
Amplitude (%)	X3	21.59	25	30	35	38.41

Table 2.

CCD of pectin yield.

Run	L/S ratio (mL/g), X1		Time (min), X2		Amplitude (%), X3		Yield of Pectin (%)	Percentage DE
	Experimental	Predicted	Experimental	Predicted	Experimental	Predicted	Experimental	Predicted
1	20	−1	120	1	35	1	8.52	7.17	62.89
2	46.82	1.68	90	0	30	0	11.08	10.30	68.86
3	13.18	−1.68	90	0	30	0	4.03	4.32	75.07
4	30	0	90	0	38.41	1.68	8.66	10.15	72.12
5	30	0	90	0	21.59	−1.68	3.61	4.46	73.19
6	30	0	39.55	−1.68	30	0	7.25	7.39	80.60
7	20	−1	60	−1	35	1	6.79	7.27	75.09
8	40	1	120	1	25	−1	7.60	7.34	75.53
9	30	0	90	0	30	0	6.15	7.31	71.02
10	30	0	90	0	30	0	8.34	7.31	63.35
11	30	0	90	0	30	0	6.82	7.31	84.59
12	30	0	140.45	1.68	30	0	6.10	7.22	74.64
13	40	1	120	1	35	1	10.10	10.72	75.53
14	20	−1	60	−1	25	−1	4.08	3.89	68.04
15	40	1	60	−1	35	1	12.29	10.83	76.88
16	20	−1	120	1	25	−1	4.78	3.78	74.27
17	30	0	90	0	30	0	8.66	7.31	76.95
18	30	0	90	0	30	0	7.00	7.31	77.66
19	40	1	60	−1	25	−1	6.61	7.44	80.96
20	30	0	90	0	30	0	7.65	7.31	71.97

$Y={\beta }_0+\mathrm{\Sigma }{\beta }_i{X}_i+\mathrm{\Sigma }{\beta }_{ii}{X_i}^2+\mathrm{\Sigma }{\beta }_{ij}{X}_i{X}_j$.

In this context, Xi and Xj represent the variables influencing the dependent response Y; β0, βi, βii, and βij denote the regression coefficients corresponding to the intercept, linear, quadratic, and interaction terms, respectively.

2.8. Response model validation

The response model was validated by applying optimized extraction parameters determined through CCD.

2.9. Characterization of the extracted pectin

All experimental runs were analyzed for percentage DE. After achieving the maximum yield from the optimized extraction method and conducting confirmation tests, the best treatment for characterization was identified based on percentage DE and monosaccharide composition.

2.9.1. Galacturonic acid content

The galacturonic acid content was assessed using a modified version of the carbazole-sulfate method based on Taylor [24]. In this procedure, the sample or standard solutions (200 µL) were mixed with concentrated sulfuric acid (3 mL) in test tubes. Then, 100 µL of a 0.1 % carbazole-ethanol solution was mixed in, and the samples were kept at 60 °C for 1 h. The galacturonic acid content was measured by determining the absorbance of the mixture at 525 nm.

2.9.2. Degree of esterification (DE)

The DE of low and high methoxyl pectin from citrus and Riang husk pectin was determined using a modified method from Singthong et al. [25]. Dried Riang husk pectin was analyzed through Attenuated Total Reflection Fourier Transform Infrared Spectroscopy (ATR-FTIR). The FTIR spectra were recorded between 7800 cm⁻¹ and 350 cm⁻¹ using an FTIR spectrophotometer (Thermo Scientific, Nicolet iS50, USA), with a resolution of 32 cm⁻¹ and 256 scans. The peak areas at 1730 cm⁻¹ (representing esterified carboxylic groups) and 1600 cm⁻¹ (representing carboxylic acid groups) were measured. The DE was then calculated as follows:

$$\mathit{DE}=(peakareaat1730{\mathit{cm}}^{-1}/(peakareaat1730{\mathit{cm}}^{-1}+peakareaat1600{\mathit{cm}}^{-1}))x100$$

2.9.3. Determination the composition of monosaccharide

The monosaccharide composition of Riang husk pectin was analyzed using high-performance liquid chromatography (HPLC). Prior to analysis, the pectin sample was hydrolyzed using a modified method from Wandee et al. [26]. Specifically, a 100 mg sample of Riang husk pectin was mixed with 1 mL of 0.5 M H₂SO₄ and boiled at 100 °C for 3 h. After hydrolysis, the solution’s pH was adjusted to neutral (pH 7) with 2 M NaOH. The total volume was then adjusted to 10 mL using deionized (DI) water, and the solution was filtered before HPLC analysis. The analysis was conducted using an HPLC system fitted with a Dionex CarboPac PA1 (4 × 250 mm) column and an electrochemical detector (Thermo Scientific, ICS-5000, USA). The mobile phase comprised 250 mM NaOH and deionized water in a 1:9 ratio, with a 0.8 mL/min flow rate at 30 °C. Xylose, arabinose, glucose, rhamnose, galactose, and mannose were the monosaccharide standards used for quantification.

2.10. Functional characteristics

2.10.1. Solubility

The solubility of Riang husk pectin was tested using the protocol outlined by Liew et al. [27] with minor modifications. The Riang husk pectin (0.1 g) was added to deionized water (10 mL) in a centrifuge tube and stirred thoroughly. After incubating at 40 °C for 30 min, the mixture underwent centrifugation at 4,200 rpm for 20 min. The resultant supernatant was dried at 105 °C until reaching a constant weight. The solubility was determined as follows:

$$\mathit{Solubility}(\%)=(driedsupernatantweight/pectinweight)x100$$

2.10.2. Water holding capacity (WHC)

The WHC of Riang husk pectin was assessed using the protocol outlined by Bayer et al. [28] with minor modifications. The Riang husk pectin (50 mg) was placed into distilled water (5 mL) and stirred thoroughly for 15 min. After incubation at room temperature for 60 min, the mixture underwent centrifugation at 5,000 rpm for 20 min. After centrifugation, the supernatant was carefully removed by tilting the tube onto filter paper and allowing it to drain for 30 min. The WHC was calculated as follows:

$$\mathit{WHC}=(weightoftubecontentafterdrainage)/(weightofdriedpectin)$$

2.10.3. Oil holding capacity (OHC)

The OHC of Riang husk pectin was measured using a method modified by Polanco-Lugo et al. [29]. Briefly, 1 g of Riang husk pectin was mixed with 10 mL of soybean oil and stirred for 5 h. After stirring, the mixture underwent centrifugation at 5,000 g for 15 min. The resulting supernatant was then allowed to drain for 30 min, and the retained oil was subsequently evaluated. The OHC was present as grams of oil held per gram of pectin.

2.10.4. Swelling capacity (SWC)

The swelling capacity (SWC) of Riang husk pectin was evaluated using a modified method from Wongkaew et al. [30]. For the SWC analysis, Riang husk pectin (100 mg) was combined with distilled water (5 mL) in a graduated cylinder and then kept at room temperature for 18 h. After the incubation period, the bed volume of the swollen pectin was recorded to determine its swelling capacity.

2.10.5. Color determination

An UltraScan VIS (HunterLab, USA) spectrophotometer was used to assess the color of Riang husk pectin following the CIE Lab system. The equipment was operated under the following conditions: standard D65 illumination, a 10-degree colorimetric angle, and RSIN mode. The color measurements followed the method outlined by Wongkaew et al. [30].

2.11. Gel properties

2.11.1. Preparation of gels

Gel preparation was carried out using the protocol modified by Gan et al. [31]. In this procedure, 0.025 g of Riang husk pectin, 3.25 g of sucrose, and citrate buffers (at pH levels of 2.0, 4.5, and 6.5) were combined in a beaker. The mixture was then heated up and stirred in a water bath at 100 °C until fully dissolved. The solution was then evaporated to achieve a final weight of 5 g. The prepared gels were kept for 18 h at 4 °C prior to analysis.

2.11.2. Flow analysis

The flow characteristic of Riang husk pectin were evaluated using a cone-plate rheometer (Thermo Scientific, Rheostress, Australia). Samples were placed in the rheometer and equilibrated at a measurement temperature of 25 °C. Shear rates ranging from 10 to 200 s⁻¹ were applied to evaluate the viscosity and shear stress of the gels.

2.12. Antioxidant assay

The antioxidant assay was assessed using DPPH assay, according to the method described by Myo and Khat-udomkiri [32]. The results were presented as mmol of Trolox equivalent antioxidant capacity (TEAC) per gram of pectin.

2.13. Cell culture

The HaCaT cells were cultured in Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 10 % fetal bovine serum (FBS) and 1 % penicillin/streptomycin solution. The cells were maintained in a humidified incubator (Binder, CB210, Germany) at 37 °C with 5 % CO₂ to ensure optimal growth conditions.

2.13.1. Cytotoxicity assay

The cytotoxicity assessment of HaCaT cells was conducted using the MTT assay (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide), following the method described by Supjaroenporn et al. [33]. The absorbance was measured at 570 nm. The cell viability percentage was determined as follows:

$$\mathit{Cell}viabilitypercentage=(absorbanceofthesample/absorbanceofthecontrol)\times 100$$

2.13.2. Determination of appropriate PM2.5 concentration

The assay was modified based on the procedure outlined by Zhu et al. [34]. Cells were seeded in a 96-well plate at a density of 15,000 cells per well. Following a 24 h incubation period, various concentrations of PM2.5 (ranging from 25 to 500 µg/mL) in serum- deficient medium and serum-deficient medium (control) were replaced to the cells. After treating the cells with PM2.5 for 24 h, the percentage of cell viability was assessed as follows:

$$\mathit{Cell}viabilitypercentage=(absorbanceofthecellstreatedwithPM2.5/absorbanceofthecontrol)\times 100$$

2.13.3. Protective effects of Riang husk pectin against PM2.5-induced damage in HaCaT

This assay was adapted from the methods used by Zhu et al. [34]. Cells were plated in a 96-well plate at a density of 15,000 cells per well. Following a 24-hour incubation period, various concentrations of Riang husk pectin in serum-deficient medium and serum-deficient medium (control) were added to the cells. After treating the cells with the sample for 24 h, the medium was changed to a serum-free culture medium containing 100 µg/mL of PM2.5, and the cells were then incubated for another 24 h. The percentage of cell viability was assessed using the following formula:

$$\mathit{Cell}viabilitypercentage=((absorbanceofthecellstreatedwithsample+PM2.5)/absorbanceofthecontrol)\times 100$$

2.13.4. Determination of the appropriate hydrogen peroxide (H₂O₂) concentration

The assay followed the protocol described by Myo and Khat-udomkiri [32]. After treating the cells with H₂O₂ for 24 h, the percentage of cell viability was assessed as follows:

$$\mathit{Cell}viabilitypercentage=(absorbanceof{H}_2{O}_2-treatedcell/absorbanceofthecontrol)\times 100$$

2.13.5. Cellular antioxidant assay

This assay was conducted following the method used by Myo and Khat-udomkiri [35]. After treating the cells with the sample for 24 h, the medium was changed to serum-deficient medium containing 300 uM H₂O₂, and the cells were continuously incubated for another 24 h. The percentage of cell viability was assessed using the following formula:

$$\mathit{Cell}viabilitypercentage=((absorbanceofthecellstreatedwithsample+H_2{O}_2)/absorbanceofthecontrol)\times 100$$

2.14. Statistical analysis

The statistical analysis of the CCD in RSM was conducted using R software. The pectin yield, obtained from the one factor at a time method, was evaluated using a one-way analysis of variance (ANOVA). Pairwise comparisons were made using the LSD test, with significance set at p < 0.05. The values from the characterized treatments were reported as the mean ± standard deviation (SD). A one-sample t-test was used to evaluate the discrepancy between the actual and predicted values during the validation phase. The values of cell culture were reported as mean ± standard error (SE).

3. Results and Discussion

3.1. Varying a single factor at a time

3.1.1. Influence of extraction solvents

The experiments were conducted using various DESs with the following compositions: Be: CA (1:2), Be: OA (1:2), Be: MA (1:2), Be: AA (1:3), Be: G (1:2), ChCl: CA (2:1), ChCl: OA (1:1), ChCl: MA (1:2), ChCl: AA (1:3), and ChCl: G (1:2). The extraction parameters were set as follows: amplitude 30 % (ultrasonic power of 26.10 W), L/S ratio 30:1 mL/g, and an extraction time of 30 min. The observed temperature of the treatments ranged from 17.50 to 20.00 °C. As can be observed from Fig. 1(a), the extraction solvent significantly impacted the yield of Riang husk pectin. The highest pectin yield of 3.92 ± 0.16 % was obtained using Be: CA (1:2), followed closely by ChCl: CA (2:1) with a yield of 3.87 ± 0.32 %. The lower pH of Be: CA compared to other acids, except for Be: OA, suggests its stronger acid strength, which has been shown to enhance pectin extraction yields, as noted by Yapo et al. [36] and Tan et al. [37]. These researchers found that, under consistent time and temperature conditions, pectin extraction yield significantly increased as pH decreased, indicating the effectiveness of stronger acids. Citric acid was selected for its superior performance in pectin extraction, not only for yield but also for its physicochemical properties. It is a safer and more environmentally friendly alternative to strong acids such as hydrochloric acid, making it highly suitable for food-related applications [38]. Liew et al. [27] also demonstrated that citric acid-based DESs outperformed lactic acid-based ones for pectin extraction from pomelo peels. Similarly, Gao et al. [39] emphasized that acidic conditions were more effective for polysaccharide extraction due to the influence of DES type, pH, and viscosity. Additionally, Chen et al. [17] reported that DES solvents outperformed HCl in extracting pectin from mango peels. In this study, the combination of Be and CA proved most effective, yielding the highest pectin, and was therefore selected for further experimentation.

Fig. 1.

Effect of solvents (a), DESs ratio (b), amplitude percentage (c), extraction time (d), and L/S ratio (e) on Riang husk pectin yield (distinct letters denote significantly different results between the conditions (p < 0.05)).

3.1.2. Effect of the DES ratio

The DES ratios of betaine (Be) and citric acid (CA), ranging from 1:2 to 1:5, were evaluated under the following conditions: a solvent containing Be and CA, amplitude set at 30 % (ultrasonic power of 26.10 W), L/S ratio of 30:1 mL/g, and an extraction time of 30 min. The treatment temperatures observed during these experiments ranged from 19.83 to 20.50 °C. As shown in Fig. 1(b), the maximum pectin yield of 4.10 ± 0.43 % was obtained at a DES ratio of 1:5. The pectin yield increased significantly with higher proportions of CA, particularly when compared to the 1:2 and 1:5 ratios. This finding contradicts the results of Shafie et al. [40], who reported no significant increase in Averrhoa bilimbi pectin yields when using DES mixtures of ChCl: CA at ratios of 1:1 to 1:3. The increase in pectin yield can be attributed to the acid extractant, as CA in DES aids in the acidification of the extraction medium. This facilitates the hydrolysis of bonds connecting pectin to components of the cell wall, leading to the release of pectin into the medium [15], [40]. Similarly, Cui et al. [15] found that a lower pH weakens the cell wall structure, making pectin easier to extract. In this study, the DES ratio of 1:5 resulted in a pH of 2, which was lower than that of other ratios and contributed to the highest extraction yield. Therefore, the DES ratio of 1:5 was chosen for further tests due to its optimal yield.

3.1.3. Effect of the amplitude

Ultrasound-assisted extraction (UAE) enhances the release and diffusion of target compounds by using acoustic energy to generate cavitation in the solvent [41]. In this study, the effect of ultrasonic wave amplitude ranging from 20 % to 40 % (ultrasonic power of 22.79 W to 30.13 W) was evaluated using a solvent composed of Be and CA at a 1:5 ratio, with an extraction time of 30 min and an L/S ratio of 30:1. The results showed that pectin yield increased as the amplitude percentage rose, peaking at 35 % (ultrasonic power of 28.11 W), after which it slightly decreased above 40 % (ultrasonic power of 30.13 W), as illustrated in Fig. 1(c) Significantly higher pectin yields were obtained at amplitudes of 30 % (ultrasonic power of 26.10 W), 35 %, and 40 % compared to those at 20 % (ultrasonic power of 22.79 W) and 25 % (ultrasonic power of 23.85 W) (p < 0.05). The highest yield of 4.63 ± 0.35 % was observed at an amplitude of 35 %, although this was not statistically different from the yields obtained at 30 % and 40 %. Similar findings have been observed in studies on sour orange peel pectin, where higher ultrasound power significantly increased pectin yield using the ultrasound-assisted extraction method [41]. The improvement in extraction efficiency can be attributed to the cavitation effects generated by ultrasonic waves, which facilitate penetration of the solvent into the plant matrix, facilitating better release and extraction of pectin [41], [42]. In addition, it could be attributed to the changes in the observed temperature during the sonication process. The observed average temperatures of the sonication process at 20 %, 25 %, 30 %, 35 %, and 40 % amplitude were 15.67, 16.83, 20.33, 25.67, and 28.33 °C, respectively, demonstrating an upward temperature trend with rising amplitude percentage. Spinei and Oroian [43] reported that higher temperatures during citric acid extraction notably increased the yield of pectin from grape pomace, indicating a direct correlation between temperature and extraction efficiency. Similarly, Liew et al. found that elevated temperatures contributed to a substantial increase in pectin yield from passion fruit peels, aligning with findings from other studies that emphasize the positive impact of temperature on pectin extraction [44]. Given the optimal yield achieved at 35 % amplitude (ultrasonic power of 28.11 W), this condition was selected for further experimentation.

3.1.4. Effect of extraction time

The effects of extraction time, ranging from 15 to 90 min, on the yield of Riang husk pectin were evaluated under the following conditions: a solvent composed of Be and CA at a 1:5 ratio, an amplitude of 35 % (ultrasonic power of 28.11 W), and an L/S ratio of 30:1 mL/g. As shown in Fig. 1(d), the pectin yield increased with longer extraction times. The highest yield, 8.19 ± 0.43 %, was observed at 90 min; however, no significant differences were found between pectin yields at 60 and 90 min. This suggests that the rate of dissolution of polysaccharides may plateau after a certain extraction duration. Gao et al. [39], in their study on polysaccharides extraction from Camellia oleifera Abel seed cake using ultrasound, found that prolonged extraction times reduced the yield. In contrast, Chen et al. [17] demonstrated that both extraction time and temperature positively influenced pectin yields from mango peel, with the highest yield observed at 90 min using Be and CA. They suggested that longer extraction times allowed for greater diffusion of pectin from plant material into the solvent [27]. In this study, the observed average temperatures of the sonication process for 15 to 90 min were ranging from 23.17 to 31.50 °C, demonstrating an upward temperature trend with rising extraction time. Consequently, the highest yield observed at 90 min can also be attributed to this gradual temperature increase, which appears to have remained within a safe range, preventing degradation of the compounds. The extended extraction time may have allowed the solvent to penetrate the plant tissue more effectively, breaking down cell walls and facilitating the release of additional bioactive compounds [45]. Furthermore, the moderate temperature rise seems to have improved the solvent’s efficiency, enhancing the solubility and transfer of target compounds into the solution [46]. These combined factors—effective penetration, controlled heat, and efficient compound transfer—appear to have contributed to the higher yield without compromising compound stability. In addition, Bagherian et al. [42], who studied pectin extraction from Turkish grapefruit using ultrasound as a pretreatment, identified an optimal pretreatment time of 30–40 min. They also observed that intermittent sonication produced higher yields than continuous sonication, as the pulsations created by intermittent ultrasound helped thin the boundary layer and reduce resistance [42]. Thus, 90 min was chosen as the optimal extraction time for further experiments.

3.1.5. Effect of the L/S ratio

The yield of Riang husk pectin was observed to be substantially influenced by L/S ratio. To evaluate its effect, different L/S ratios of 10:1, 20:1, 30:1, 40:1, and 50:1 mL/g were tested under the following conditions: a solvent containing Be and CA at a 1:5 ratio, an amplitude of 35 % (ultrasonic power of 28.11 W), and an extraction time of 90 min. The observed temperature of the treatments ranged from 28.83 to 29.83 °C. The highest pectin yield of 8.88 ± 0.01 % was observed at an L/S ratio of 30:1, significantly different from the other ratios, as shown in Fig. 1(e). Pectin yield increased as the L/S ratio increased, but a slight decline was observed at ratios above 30:1. This pattern aligns with the findings of Chen et al. [17]. Theoretically, a higher L/S ratio facilitates the diffusion of target compounds, improving yield [27], [47]. However, excessively high L/S ratios may lead to inefficiencies, such as unnecessary fluid use and potential negative impacts on the extraction process [39]. Given the significant result, the L/S ratio of 30:1 mL/g was chosen for subsequent studies.

3.2. Optimization process for Riang husk pectin extraction

Varying single-factor-at-a-time experiments indicated that the DES comprising Be and CA at a ratio of 1:5 yielded the highest amount of pectin from Riang husks. To further optimize the extraction process, RSM was applied, with variables including L/S ratio, extraction time, and amplitude percentage, each at three different levels. The results of the Riang pectin yield and percentage DE are depicted in Table 2, with yields ranging from 3.61 % to 12.29 % for actual values and 3.78 % to 10.83 % for predicted values. The highest pectin yield was obtained in Run 15, with an L/S ratio of 40:1 mL/g, extraction time of 60 min, and amplitude of 35 % (ultrasonic power of 28.11 W). The actual yield was 12.29 %, while the predicted yield was 10.83 %, showing close agreement between experimental and predicted results. The linear model for Riang husk pectin extraction was highly significant (p < 0.001) with a non-significant lack of fit (p = 0.3466), demonstrating that the model was suitable for predicted pectin yield, as summarized in Table 3. The coefficient of determination (R²) for pectin yield exceeded 0.84, suggesting that the model could explain over 84 % of the variability in the experimental data. The adjusted R² value of over 0.81 further supported the reliability of the model [32]. Table 4 displays the results and estimated coefficients of the linear model. The effects of the L/S ratio (X₁), extraction time (X₂), and amplitude (X₃) on the yield of Riang husk pectin, based on the 20 experimental runs, were fitted by the following linear equation:

$$\mathit{Pectin}yield\left(\%\right)=7.306+1.778\left(X_1\right)-0.052(X_2)+1.693(X_3)$$

Table 3.

ANOVA of Riang husk pectin extraction for CCD.

Source	Sum of Squares	df	Mean Square	F value	Pr (>F)
Model	87.102	9	9.678	8.584353	0.001191
Linear	82.35	3	27.4498	24.3474	6.50E-05
Interaction	2.606	3	0.8686	0.7705	0.5364
Quadratic	2.145	3	0.7151	0.6342	0.6097
Residuals	11.274	10	1.1274
Lack of fit	6.674	5	1.3347	1.4506	0.3466
Pure error	4.601	5	0.9201
Total	98.376	19
R2	0.8371
Adjusted R2	0.8066

df: Degree of freedom.

Table 4.

Regression analysis of Riang husk pectin for CCD.

Parameter	Estimated	Std. Error	t value	Pr(>|t|)
intercept	7.306398	0.223784	32.6494	4.52E-16
X1	1.778336	0.270812	6.5667	6.49E-06
X2	−0.0523	0.270812	−0.1931	0.8493
X3	1.69254	0.270812	6.2499	1.16E-05

Both the L/S ratio and amplitude had a significantly positive effect on the Riang pectin yield (p < 0.001), while the effect of extraction time had no significant effect on the Riang husk pectin yield.

The results for percentage DE are shown in Table 2, ranging from 62.89 % to 84.59 %. All extracted pectin samples indicated high-methoxyl pectin. In this case, there are no significant model terms, suggesting that the studied factors cannot predict the percentage DE of the extracted pectin. Chuenkaek et al. [48] reported that high-methoxyl pectin exhibited remarkable cell activation, enhanced skin compatibility, and excellent moisture retention. Its moisturizing, thickening, and film-forming qualities, along with its contribution to natural and sustainable solutions, make pectin highly beneficial for use in the cosmetic industry, particularly in skincare and beauty products [14].

Fig. 2 presents a response surface plot illustrating the interaction of two factors, with the third factor held constant at its central level. Fig. 2(a) depicts the correlation between the liquid-to-solid (L/S) ratio and extraction time with pectin yield. The results indicated that while longer extraction times did not lead to a significant increase in pectin yield, a higher L/S ratio resulted in higher yields. According to the regression analysis in Table 4, the extraction time had no significant effect on pectin yield (p > 0.05). These findings align with previous studies, which feature the positive impact of a higher L/S ratio on pectin yield. Similar trends have been observed in pectin extraction from durian rinds and pistachio hulls [49], [50], where increased L/S ratios led to higher yields, likely due to improved mass transfer driving forces for pectin [50]. Conversely, Perez et al. [51] found that in pectin extraction from watermelon rind, prolonged extraction times and lower L/S ratios resulted in reduced yields. Amplitude was identified as one of the critical variables affecting Riang husk pectin yield. Fig. 2(b and c) illustrates that as amplitude increases, pectin yield also improves. This result can be attributed to the cavitation effects generated by ultrasound waves, enhancing the solvent’s penetration into the plant matrix. This effect facilitates the release of pectin, increasing the efficiency and yield of the extraction process [41], [52]. Additionally, the observed average temperature during the CCD experimental runs ranged from 24.00 to 37.00 °C. The highest temperature was recorded in run 13, with an L/S ratio of 40:1 mL/g, an extraction time of 120 min, and an amplitude of 35 % (ultrasonic power of 28.11 W). The lowest temperatures were observed in runs 5 and 6, which had the lowest amplitude and extraction time, respectively. However, the treatment with the highest temperature did not yield the highest pectin. These findings suggest that the increase in temperature is primarily associated with higher amplitude settings. This upward temperature trend with increasing amplitude may contribute to the observed positive impact of amplitude on the pectin yield. However, in this optimization process for Riang husk pectin extraction, it revealed only significant linear relationships among the variables, as illustrated by the response surfaces in Fig. 2. No significant curvature was observed, suggesting that the true optimum was not within the studied range. Future studies may consider expanding the variable limits to capture potential non-linear effects and identify a true optimum for pectin yield.

Fig. 2.

Response surface plot for the effect of independent factors on Riang husk pectin yield.

3.3. Response model validation

Based on the response surface model, the highest yielded conditions for the three independent variables were identified as an L/S ratio of 40 mL/g, an extraction time of 60 min, and an amplitude of 35 % (ultrasonic power of 28.11 W). To validate the accuracy of the response model, a confirmation experiment was performed using these conditions. The model predicted a pectin yield of 10.83 %, while the actual experimental result was 12.65 ± 1.34 %. The predicted yield was not significantly different from the actual result (p ≥ 0.05), indicating that the experimental values closely aligned with the predicted ones. This confirms that the model derived from the CCD is a reliable and accurate tool for estimating pectin yield from Riang husk.

3.4. Extracted pectin characterization

3.4.1. Galacturonic acid content

The measurement of galacturonic acid is crucial for analyzing pectin structure, as it is the primary component of pectin. In this study, the content of galacturonic acid was observed to be 53.74 % mol (Table 5). Buathongjan et al. [8] reported that pectin extracted from Riang pod husks contained 70.36 % galacturonic acid, and the content in Riang pod powder was 65.38 %. Several studies have reported varying levels of galacturonic acid, including pectin from apple pomace (38.0–47.1 %) [53], grapefruit peel (44.2–60.6 %) [15], and grapefruit peel (50.0 %) [54]. The variations in galacturonic acid content across studies can be attributed to factors such as extraction methods, extraction time, and L/S ratio during the extraction process [53].

Table 5.

Monosaccharide content of pectin extracted from Riang.

Monosaccharide	mM	% mol
Rhamnose	0.19	6.81
Arabinose	0.67	23.97
Galactose	0.35	12.36
Glucose	ND	−
Xylose	0.07	2.48
Mannose	0.02	0.62
Galacturonic acid	1.51	53.74

* ND = Not detected.

3.4.2. Monosaccharide composition

The monosaccharide composition of Riang husk pectin was analyzed using ion chromatography, revealing the presence of rhamnose, xylose, galactose, arabinose, and mannose (Table 5). The predominant components were galacturonic acid (53.74 % mol), arabinose (23.97 % mol), galactose (12.36 % mol), and rhamnose (6.81 % mol), indicating that these monosaccharides may form the core structure of Riang husk pectin. These results, however, differ in percentage mol from those reported by Buathongjan et al. [8], who studied pectin from Riang pod husks. They found that the dominant monosaccharides in Riang pod husk pectin were galacturonic acid, galactose, arabinose, and rhamnose, with percentage mol values of 70.36, 11.56, 6.63, and 6.02, respectively. The discrepancies are likely due to variations in the raw material sources, extraction solvents, and methods used. Indeed, differences in pectin’s structural characteristics are often influenced by factors such as plant sources, extraction techniques, solvent choices, and extraction conditions [15]. These variables play a crucial role in determining the composition and functional properties of the extracted pectin.

3.4.3. Degree of esterification (DE)

The FTIR spectrum of Riang husk pectin, high methoxyl pectin from citrus, and low methoxyl pectin from citrus, as depicted in Fig. 3, displays a broad absorption band ranging from 3600 to 2500 cm⁻¹, corresponding to O-H stretching vibrations. This indicates the presence of free hydroxyl groups and bonded O-H bands, characteristic of carboxylic acids [55]. A prominent band around 2900 cm⁻¹ represents C-H absorption, including CH, CH₂, and CH₃ stretching and bending vibrations, associated with O-acetyl groups [29], [49], [55]. The DE is evaluated through bands found between 1800 cm⁻¹ and 1500 cm⁻¹. The stretching vibrations of the carboxylate anion (COO^–) are associated with bands around 1630 cm⁻¹ to 1600 cm⁻¹, while the stretching vibrations of the carbonyl group (C=O) in both methyl ester and carboxylic acid groups are linked to bands at 1740 cm⁻¹ to 1720 cm⁻¹ [56]. In this study, the FTIR spectrum showed bands at 1712.06 cm⁻¹ and 1610.09 cm⁻¹, consistent with these observations. Additional bands were observed at 1396.73, 1313.31, 1198.65, 1098.73, and 1015.73 cm⁻¹, falling within the range of 1300–1000 cm⁻¹. These correspond to C=O stretching, as well as CH₂ bending, OH bending, and –CH₃CO stretching [55]. It was observed that the spectra of low and high methoxyl pectin from citrus showed a more intense band around 1015 cm⁻¹, corresponding to C–O stretching vibrations. Bands at 953.38 cm⁻¹ and 889.87 cm⁻¹ were also identified, matching those reported by Wandee et al. [26], and attributed to C–C–H and C–O–H bending vibrations, as well as out-of-plane vibrations of hydroxyl groups. In terms of DE, the Riang husk pectin in this study exhibited 73.41 % esterification, categorizing it as high-methoxyl pectin. This finding aligns with the DE values of pectin extracted from Riang pod powder and Riang pod husk, which were 66.3 % and 66.2 %, respectively, as reported by Buathongjan et al. [8]. In contrast, Apirattananusorn et al. [13] found a DE of 48.3 % in water-extractable Riang pectin, classifying it as low-methoxyl pectin and suggesting that varying solvents during extraction could influence DE values.

Fig. 3.

FTIR spectral diagram of Riang husk pectin, high methoxyl pectin from citrus, and low methoxyl pectin from citrus.

3.5. Functional characterization

3.5.1. Solubility

The solubility of Riang husk pectin obtained in this study was 53.00 ± 3.46 % (Table 6). In comparison, Buathongjan et al. [8] reported that Riang Husk and Riang Pod powders contain notable amounts of soluble dietary fiber, with values of 13 % and 27 %, respectively, likely attributed to the presence of pectin and/or water-soluble hemicelluloses. The solubility observed in this study was lower than the 76.27 % reported by Liew et al. [27] for pectin from pomelo peels but comparable to the solubility of commercial citrus pectin (54 %). According to Polanco-Lugo et al. [29], the moderate solubility of pectin can affect its hydration properties, often linked to a high content of acetyl ester groups, contributing to the hydrophobic nature of pectin. This hydrophobicity can influence polysaccharide–polysaccharide and polysaccharide–water interactions [31]. Studies by Gan et al. [31] and Monsoor and Proctor [57] demonstrated that pectin extracted at varying pH levels exhibited different solubility, though the solubility remained relatively stable when dissolved in solutions at different pH values. They also noted that high-methoxyl pectin generally had lower solubility compared to low-methoxyl pectin, likely due to its higher ester group content. Pectin’s water solubility makes it well-suited for use in water-based formulations. Additionally, pectin’s solubility and appropriate viscosity enhance its value in such applications [58].

Table 6.

Functional properties and color of Riang husk pectin.

Functional properties				Color
Solubility (%)	WHC (g water/g pectin)	OHC (g oil/g pectin)	SWC (ml/g pectin)	L*	a*	b*
53.00 ± 3.46	3.88 ± 0.49	3.30 ± 0.21	11.77 ± 0.29	77.34 ± 0.01	9.02 ± 0.01	17.39 ± 0.01

Note: L* (lightness; 0 (black) to 100 (white)), a* (red (+) – green (−) axis), b* (yellow (+)- blue (−)).

3.5.2. Water holding capacity (WHC)

Riang husk pectin demonstrated WHC of 3.88 ± 0.49 g of water per gram of sample, as shown in Table 6. Ultrasound treatment enhances cavitation within the pectin structure, improving water penetration and absorption [28], [59]. According to Bayer et al. [28], pectin extracted using ultrasound with water (4.84 g/g) had a lower WHC than that obtained via chemical extraction methods (5.64 g/g). Several factors can influence the WHC, including particle size, the presence of free hydroxyl groups in the pectin structure, ionic strength, porosity, temperature, pH, applied stresses on the fibers, and the type of ions in the solution [41], [60]. The increase in WHC is often attributed to the presence of more free hydroxyl groups in the pectin structure, enhancing its ability to retain water [29]. Fibers with high WHC are valuable ingredients in modifying the viscosity and texture of products [61]. In the cosmetic industry, pectin with high WHC provides multiple advantages, including facilitating gelation, increasing viscosity, enhancing moisture absorption, supporting emulsification, enabling esterification, improving adhesion, and promoting chelation [14]. These properties make pectin a versatile ingredient for cosmetic formulations and other applications.

3.5.3. Oil holding capacity (OHC)

OHC is a key physical characteristic of plant polysaccharides, closely related to their chemical structure. Factors such as surface characteristics, thickness, charge density, and the hydrophobicity of the fiber particles can influence OHC [30], [62]. In this study, the OHC of Riang husk pectin was found to be 3.30 ± 0.21 g oil/ g sample, as shown in Table 6. The OHC values can vary widely depending on the source of the fibers. For instance, the OHC values reported for mango peel pectin, pomegranate bagasse, ripe kiwi, and passion fruit albedo were 0.81, 5.9, 6.00, and 2.03 g oil/g sample, respectively [30]. Gan et al. [31] observed that pectin extracted from the same plant at different pH levels also exhibited varying OHC values, suggesting pectin might have a larger surface area and contain more soluble dietary fiber. Ingredients with higher OHC contribute to the stability of emulsions by effectively binding oils, which is crucial for maintaining the consistency and stability of a wide range of products, including lotions, makeup formulations, shampoos, hair conditioners, and other personal care cosmetics [14]. This makes pectin a valuable component in the formulation of such products, enhancing their texture and performance.

3.5.4. Swelling capacity (SWC)

Swelling capacity (SWC) can be measured using the bed volume method, where fibers are soaked in water overnight in a volumetric cylinder, and the expansion of the fiber matrix upon water absorption is recorded [30], [60]. A higher swelling capacity typically correlates with a greater content of soluble dietary fiber. In this study, Riang husk pectin exhibited a swelling capacity of 11.77 ± 0.29 mL/g, as shown in Table 6. The SWC values can vary significantly between different plant fibers. For example, mango peel pectin has an SWC of 24.16 mL/g [30], while the SWC values for mango, pineapple, passion fruit, and guava are 4.6 mL/g, 6.6 mL/g, 7.2 mL/g, and 1.4 mL/g, respectively [63]. In the cosmetics industry, SWC is an important characteristic, as it enhances hydration, texture, and stability. When hydrated, pectin swells to form a smooth, moist particle system, contributing to the product’s overall feel and performance [14]. This makes pectin a valuable ingredient in skincare formulations, improving both functionality and sensory properties.

3.5.5. Color

The color of pectin is an important factor, as it influences the visual appeal of the final product. The color of Riang husk pectin, obtained through ultrasonic extraction, is detailed in Table 6. The visible color of the pectin can be described as milky white with a brown tint. The colorimeter readings revealed higher lightness (L*) values, with the pectin leaning toward yellow and red hues. According to Wongkaew et al. [30], the pigmentation of the biomass can significantly impact the final color of pectin, as pigments present in the plant material are often difficult to remove during the extraction process. Heating is considered a major factor influencing pectin coloration. Additionally, bound phenolic compounds or other water-soluble colorants could contribute to the high pigmentation levels in pectin. Variations in the temperature and duration of the extraction process can also affect the color of the final product [64], [65]. These factors are important to consider during the pectin extraction process, as they can influence the aesthetic qualities of pectin-based products in industries including cosmetics and food.

3.6. Gel properties

3.6.1. Flow analysis

In this study, Riang husk pectin was used to form gels at varying pH levels (2.0, 4.5, and 6.5) while maintaining a constant sucrose concentration of 65 % (w/w) for the evaluation of flow behavior. The rheological data on the Riang husk pectin gel samples are presented in Fig. 4 (a). All gel samples demonstrated shear-thinning behavior, aligning with the findings of Buathongjan et al. [8], who observed that pectin from Riang pod and Riang husk exhibited shear-thinning properties at higher concentrations (above 2 % w/v) but behaved Newtonian at lower concentrations. The higher shear stress observed relative to the shear rate at pH 4.5 and 6.5 was attributed to the increased viscosity of Riang husk pectin gel, as shown in Fig. 4(b). At pH 4.5, the viscosity of the gel rapidly declined as the shear rate increased from 10 to 30 s⁻¹ and then slightly increased up to 200 s⁻¹. In contrast, the viscosity at pH 2 and 6.5 fluctuated between shear rates of 10 and 30 s⁻¹, stabilizing at pH 2 and slightly increasing at pH 6.5. These viscosity patterns are consistent with reports on the behavior of pectin extracted from pomelo peel and Parkia speciosa pod pectin [27], [31]. This rheological behavior is influenced by the quantity and molecular weight of pectin, which is linked to factors such as protein content, as well as the presence of methyl, acetyl, and feruloyl esters in the pectin’s long chains, which facilitate crosslinking through hydrophobic interactions [29]. The shear-thinning properties of polysaccharides, such as Riang husk pectin, allow for more efficient flow, contributing to a smoother texture in formulations [27]. This makes it highly desirable for applications in food and cosmetic products where texture and viscosity play crucial roles in product performance.

Fig. 4.

Flow characteristics (a) and viscosity (b) for the Riang pectin gel at different pH.

3.7. Antioxidant property

The DPPH activity of Riang husk pectin yielded a value of 0.26 ± 0.02 mmol Trolox equivalents per gram of sample. For comparison, Buathongjan et al. [8] reported antioxidant activity values of 0.63 ± 0.01 and 0.16 ± 0.01 mmol Trolox equivalents per gram for Riang husk and pod pectin, respectively, using the same assay. Chen et al. [66] found that ultrasonic treatment can enhance the hydroxyl radical scavenging activity of pectin and observed that the antioxidant capabilities of hawthorn pectin were directly proportional to its concentration. The antioxidant activity of pectin can differ widely depending on its structure, as it primarily consists of a polymer made up of α-(1–4)-linked D-galacturonic acid. This polymer contains hydroxyl groups at the 2′ and 3′ carbon positions, which may act similarly to flavonoids and phenolic compounds. These compounds are renowned for their antioxidant properties, as they are capable of donating hydrogen or electrons to neutralize free radicals. The antioxidant action of pectin is influenced by several factors, including the quantity and type of neutral sugars, the presence of proteins and phenolic contaminants, as well as the molecular weight and degree of pectin methylation [9], [10]. These structural attributes, especially the prevalence of hydroxyl groups in polysaccharides, play a key role in the antioxidative capabilities of pectin [11].

3.8. Cell culture

3.8.1. Cytotoxicity assay

The cytotoxicity of HaCaT cells was determined using various concentrations of ascorbic acid and Riang husk pectin to determine their highest non-toxic levels for use in the cellular antioxidant assay and to assess the protective effect of Riang husk pectin against PM2.5-induced damage in HaCaT cells. The cytotoxicity assay results are shown in Fig. 5. Non-cytotoxic concentrations of ascorbic acid, ranging from 0.001 to 0.01 mg/mL, maintained over 80 % cell viability in HaCaT cells. However, higher concentrations of ascorbic acid may cause cellular damage, as although ascorbic acid is widely known for its antioxidant properties, it can exhibit pro-oxidant effects at elevated concentrations, potentially leading to cellular harm [67]. For Riang husk pectin, concentrations between 0.1 to 0.5 mg/mL resulted in cell viability of over 80 %, suggesting that these concentrations can be safely used in future cellular assays. In contrast, Abdel-Massih et al. [68] reported a decline in cell viability along with an increase in the concentration of citrus pectin, with cell viability dropping from 82 % at 1 μg/mL to 62.54 % at 750 μg/mL. The differences in concentration thresholds between this study and the current one may be due to variations in the plant source, solvents, and extraction methods used for obtaining the pectin. This suggests that Riang husk pectin exhibits low cytotoxicity and can be used at higher concentrations compared to citrus pectin for similar cellular applications.

Fig. 5.

Cytotoxicity of ascorbic acid and Riang husk pectin in different concentrations (* denote that the result significantly differs from the control (p < 0.05)).

3.8.2. Determination of appropriate PM2.5 concentration

The assessment of HaCaT cell viability after exposure to PM2.5 is depicted in Fig. 6. Cell viability significantly decreased at all concentrations compared to the control (p < 0.05). A PM2.5 concentration exceeding 100 µg/mL, resulting in 65.02 ± 0.24 % cell viability, was selected for further experiments as it provided an optimal survival rate, avoiding both insufficient stress and irreversible damage [68]. Prolonged exposure to PM2.5 induces oxidative stress and mitochondria-dependent apoptosis, potentially leading to skin irritation and damage [5]. The toxic effects of PM2.5 are due to both the particles themselves and the chemicals attached to their surfaces. Hu et al. [5] observed that PM2.5 exposure resulted in a significant, concentration-dependent increase in reactive oxygen species (ROS) and malondialdehyde (MDA) levels in HaCaT cells while also causing a dose-dependent reduction in superoxide dismutase (SOD) activity, indicating oxidative stress. Additionally, Piao et al. [6] found that PM2.5 promotes apoptotic cell death by inducing oxidative stress, leading to the degradation of critical cellular organelles such as the endoplasmic reticulum, mitochondria, and lysosomes. These results align with the research by Li et al. [70], which shows that PM2.5 exposure triggers oxidative stress and the production of pro-inflammatory substances. The PM2.5 samples used in this study were characterized by Kraisitnitikul et al. [20], who collected ambient PM2.5 during the dry season of 2019, a period with high atmospheric pollution. Their analysis revealed that daily PM2.5 concentrations exceeded 200 μg/m³ and contained organic carbon, elemental carbon, and PM2.5-bound metals such as Al, Ca, Cu, Fe, K, Mg, Mn, Pb, Sn, and Zn. These findings align with the toxicity mechanisms discussed by Hu et al. [5], Piao et al. [6], and Li et al. [70], emphasizing the role of PM2.5 in inducing oxidative stress and its harmful effects on cellular health.

Fig. 6.

Effects of PM2.5 concentrations on HaCaT cells viability (*denotes that the result differs significantly from the control (p < 0.05)).

3.8.3. Protective effects of riang pectin against PM2.5-induced damage in HaCaT

Cell viability exceeding 80 % was observed across all tested concentrations of Riang husk pectin in combination with PM2.5, significantly higher than the 67.57 % cell viability observed in cells treated with PM2.5 alone, as shown in Fig. 7. Interestingly, the increase in cell viability was not concentration-dependent on Riang husk pectin. Tang et al. [71] demonstrated that pectin can mitigate the inflammatory response caused by PM2.5 by reducing lung tissue damage, lowering levels of inflammatory cytokines, decreasing cell infiltration, and diminishing myeloperoxidase activity. Similarly, Zhu et al. [34] reported that Lycium barbarum polysaccharide effectively inhibits PM2.5-induced oxidative stress, and others, contributing to a reduction in oxidative damage, enhancing cell survival, and minimizing apoptosis. The outcomes observed in this study may be due to the ability of Riang husk pectin to reduce oxidative stress in PM2.5-treated HaCaT cells, consistent with its in vitro and cellular antioxidant activities. Additionally, pectin’s ability to bind heavy metals, depending on its structural characteristics [12], [72], could further enhance cell survival by reducing the oxidative stress caused by the metal components present in PM2.5 particles. This binding likely contributes to the protective effect of Riang husk pectin against PM2.5-induced damage in HaCaT cells.

Fig. 7.

Cell viability comparison of control, PM2.5-treated cell, and Piang husk pectin + PM2.5-treated cell (*denotes that the result differs significantly from the control (p < 0.05), (# denotes that the result differs significantly from the PM2.5-treated cell (p < 0.05)).

3.8.4. Determination of appropriate hydrogen peroxide (H₂O₂) concentration

Hydrogen peroxide (H₂O₂) is a well-known reactive oxygen species (ROS) that induces oxidative stress by generating free radicals. These free radicals can damage cellular components, leading to apoptosis. The extent of H₂O₂-induced apoptosis depends on both its concentration and the specific cellular context [73]. As shown in Fig. 8, H₂O₂ treatment leads to a significant decrease in cell viability, with the effect being directly proportional to the concentration of H₂O₂. All tested concentrations exhibited a statistically significant reduction in cell viability compared to the control group (p < 0.05). After being treated for 1 h with H₂O₂, only the cell viability at 100 μM exceeded 80 %. In contrast, at H₂O₂ concentrations of 300, 500, 700, and 900 μM, cell viabilities were 59.12 ± 3.53 %, 58.22 ± 0.57 %, 54.88 ± 1.73 %, and 47.41 ± 0.66 %, respectively. The 300 μM concentration, resulting in approximately 60 % cell viability, was selected for subsequent experiments. In establishing an oxidative stress cell model, a cell viability range of 50 % to 70 % is typically considered ideal. If cell viability is too high, the cells may not be subjected to sufficient oxidative stress, whereas excessively low survival rates could lead to irreversible cellular damage [69]. This balance ensures that the cells experience oxidative stress without causing excessive harm, potentially comprising the experimental model.

Fig. 8.

The effects of H₂O₂ on cell survival (* denotes that the result differs significantly from the control (p < 0.05)).

3.8.5. Cellular antioxidant assay

Fig. 9 demonstrates that treating HaCaT cells with 300 µM H₂O₂ for 1 h results in a survival rate of approximately 60 %, confirming the effective induction of oxidative stress. The addition of ascorbic acid at various concentrations significantly increased cell viability compared to H₂O₂-treated cells alone (p < 0.05). Specifically, ascorbic acid concentrations of 0.001, 0.005, and 0.01 mg/mL improved cell viability to 70.54 ± 3.01 %, 74.20 ± 1.95 %, and 77.53 ± 4.33 %, respectively. Ascorbic acid, a well-known antioxidant, protects HaCaT cells from oxidative stress by neutralizing ROS, such as H₂O₂, thereby preserving cell viability and preventing apoptosis or necrosis [67]. Riang husk pectin, at all tested concentrations, also improved cell viability compared to H₂O₂-treated cells, with viability percentages exceeding 67.01 %. However, the increase in cell viability at the 0.1 mg/mL concentration of Riang husk pectin was not significantly different from the cells treated with H₂O₂ alone. Wang et al. [69] demonstrated that citrus pectin exhibited strong radical scavenging effects and provided protection against oxidative damage in HepG2 cells exposed to H₂O₂. Similarly, natural polysaccharides, including pectin, enhance cellular antioxidant activity through several mechanisms: scavenging free radicals, improving the bioavailability of antioxidants, chelating metal ions, and regulating cellular antioxidant pathways. These mechanisms collectively help mitigate oxidative stress and protect cells from damage [74], [75]. This suggests that similar to citrus pectin, Riang husk pectin may offer a protective activity against oxidative stress, making it a potential candidate for reducing cellular damage caused by ROS.

Fig. 9.

Cellular antioxidant activity of Riang pectin and ascorbic acid (* denotes that the result differs significantly from the control (p < 0.05), (# denotes that the result differs significantly from H₂O₂-treated cell (p < 0.05)).

4. Conclusion

This study successfully demonstrates the efficacy of UAE combined with DESs for isolating high-quality pectin from Riang husks, utilizing a systematic approach through CCD in RSM. Among different DESs, Be: CA (1:5) was found to be the most effective deep eutectic solvent for maximizing pectin yield from Riang husks. The optimal conditions for extracting Riang husk pectin were an L/S ratio of 40 mL/g, extraction time of 60 min, and amplitude of 35 % (ultrasonic power of 28.11 W), with the L/S ratio observed to be the most influential variable on pectin yield. The extracted pectin, predominantly composed of galacturonic acid, exhibited desirable structural, functional, and rheological properties, with notable antioxidant activity and protective effects against PM2.5-induced cellular damage. These outcomes point to the potential of Riang husk pectin for use as a valuable ingredient in antipollution products. However, this study is limited to in vitro assessments, lacking in-depth mechanistic studies and in vivo validation of pectin’s efficacy. In addition, the study also lacks molecular weight (MW) analysis, a parameter for understanding pectin’s structure and bioactive properties, and control experiments to compare the UAE and DES extraction methods with conventional techniques, which would provide more insights into their effectiveness. Future research should address these gaps to provide a more thorough evaluation of the effectiveness of UAE and DES in pectin extraction and its overall functionality by exploring the detailed mechanisms of pectin’s interaction with pollutants and validating its protective efficacy in practical applications. Despite these limitations, the innovative use of UAE and DESs to extract pectin signifies a promising advancement in the development of sustainable, high-performance antipollution products, contributing to global sustainability goals and addressing the pressing need for effective pollution mitigation solutions.

CRediT authorship contribution statement

Manee Saelee: Writing – review & editing, Writing – original draft, Visualization, Methodology, Investigation, Formal analysis. Hla Myo: Writing – review & editing, Visualization, Investigation. Nuntawat Khat-udomkiri: Writing – review & editing, Visualization, Project administration, Methodology, Investigation, Funding acquisition, Formal analysis, Conceptualization.

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.