Physicochemical and Functional Characterization of Cucumis sativus L. (Poona Kheera) Mucilage and Its Application as a Coating to Inhibit Enzymatic Browning in Fresh-Cut Apples

Abstract

Enzymatic browning is a major challenge in maintaining the quality and shelf life of fresh-cut fruits, and in this context, plant-derived hydrocolloids are increasingly recognized as sustainable alternatives to synthetic additives due to their ability to retard browning while supporting quality retention. Therefore, in the present study, Cucumis sativus L. mucilage was extracted using microwave irradiation, yielding 24.56% freeze-dried irregular particles with an average size of 194.5 nm and −19.8 mV zeta potential. Various characterization techniques confirmed the amorphous structure and the presence of polysaccharides functional group. The mucilage was primarily composed of glucose (32.27%), along with arabinose, galactose, xylose, mannose, rhamnose, and minor uronic acids, reflecting a glucose-rich heteropolysaccharide. Functionally, the mucilage exhibited notable water retention (8.46 g/g), oil retention (3.21 g/g), foaming capacity (52.13%) with stability (30.46%), emulsifying capacity (90.45%) with stability (91.62%), and solubility (90.14%). Antioxidant assays revealed strong ferric reducing power (5.1 mM FeSO₄ at 10 mg/mL), DPPH scavenging (67.50%; IC₅₀ = 1.798 mg/mL), and ABTS scavenging (60.14%, IC₅₀ = 8.038 mg/mL). Anti-inflammatory evaluation indicated enhanced macrophage viability (1.38-fold at 25 mg/mL) with reduced nitric oxide production, while tyrosinase inhibition reached 60.40% (monophenolase) and 68.50% (diphenolase) at 2 mg/mL. Furthermore, when applied as an edible coating on fresh-cut apple slices, Cucumis sativus L. mucilage effectively delayed enzymatic browning in a dose-dependent manner, with 2 mg/mL maintaining apple slice brightness (L* value; 71.08) and minimizing color change (ΔE = 4.54). Overall, these findings highlight Cucumis sativus L. mucilage as a multifunctional biopolymer with promising applications in food systems and edible coatings.

1. Introduction

Enzymatic browning is a major quality defect in fresh-cut fruits, leading to undesirable changes in appearance, flavor, nutritional value, and consumer acceptability [1]. This phenomenon primarily results from the oxidation of phenolic compounds catalyzed by enzymes such as polyphenol oxidase (PPO), peroxidase (POD), and tyrosinase, producing o-quinones that subsequently polymerize into brown pigments [2]. Conventional anti-browning agents, including sulfites and ascorbic acid, are effective but are limited by safety concerns, regulatory restrictions, and poor stability during storage and processing. These drawbacks have driven growing interest in safe, natural, and multifunctional alternatives derived from plant-based materials [3]. Plant-derived mucilages have emerged as promising candidates for controlling enzymatic browning due to their dual functionality as natural hydrocolloids and bioactive polymers. Mucilage is a complex polysaccharide material widely distributed in plant tissues such as seeds, fruits, peels, and leaves, where it plays important physiological roles in water retention, protection, and nutrient regulation. From a food application perspective, mucilages offer desirable film-forming, water-holding, emulsifying, and stabilizing properties, making them suitable for edible coatings and functional food systems [4].

Cucumis sativus L. (cucumber), a member of the family Cucurbitaceae, is a widely consumed vegetable with recognized nutritional and functional value. Various parts of cucumber, including seeds, peels, and fruit tissues, are rich sources of mucilage and polysaccharides with promising physicochemical and biological properties. Although cucumber has been extensively studied for its nutritional composition and antioxidant potential, the functional application of its mucilage, particularly as a natural anti-browning agent, remains relatively underexplored [5]. Cucumber mucilage is a heteropolysaccharide matrix composed primarily of neutral sugars such as glucose, galactose, arabinose, rhamnose, and xylose, along with uronic acids including galacturonic and glucuronic acid. The presence of uronic acids imparts a negative charge, enhancing solubility, water-binding capacity, and interaction with metal ions, while branched polysaccharide structures contribute to favorable rheological behavior [6]. These structural features translate into excellent hydrocolloidal properties, including thickening, emulsification, and film formation, which are essential for edible coating applications [7]. Beyond its physicochemical functionality, cucumber mucilage has been reported to exhibit bioactive properties such as antioxidant and enzyme-inhibitory activities. The abundance of hydroxyl (–OH) and carboxyl (–COOH) functional groups enables the effective scavenging of free radicals and reactive quinones, thereby interrupting oxidative pathways involved in enzymatic browning [8]. Additionally, negatively charged uronic acid residues may chelate copper ions at the active site of tyrosinase, leading to partial suppression of its catalytic activity. These biochemical interactions are particularly relevant for controlling browning reactions in fresh-cut fruits, where tyrosinase-mediated oxidation plays a dominant role.

Recent advances in extraction technologies, especially microwave-assisted extraction, have improved the recovery of plant mucilages by reducing extraction time and preserving structural integrity and bioactivity. Microwave-assisted extraction has shown promise for producing high-quality cucumber mucilage with minimal thermal degradation, making it suitable for functional food applications [9]. Edible coatings based on natural polysaccharides act as a barrier on fruit surfaces, limiting oxygen diffusion, reducing moisture loss, and slowing enzymatic reactions. When combined with intrinsic antioxidant and enzyme-inhibitory properties, such coatings can effectively delay browning and extend the shelf life of fresh-cut produce [10]. Among fresh-cut fruits, apples are particularly susceptible to enzymatic browning and are therefore an ideal model system to evaluate the effectiveness of cucumber mucilage-based coatings. The ability of cucumber mucilage to mitigate enzymatic browning can be attributed to multiple synergistic mechanisms, including radical scavenging, reduction of o-quinones back to their phenolic forms, chelation of copper ions at the tyrosinase active site, and formation of an oxygen-limiting physical barrier on the fruit surface [11]. These effects contribute to the preservation of color, sensory attributes, and nutritional quality of fresh-cut fruits.

Despite the growing interest in plant-derived mucilages for food preservation, studies investigating cucumber (Cucumis sativus L.) mucilage as a functional anti-browning agent remain limited. Most existing reports focus either on the nutritional composition of cucumber or antioxidant activity, with little emphasis on its direct application as an edible coating for fresh-cut fruits. Moreover, the combined evaluation of microwave-assisted extraction for enhancing mucilage functionality, detailed physicochemical characterization, and systematic assessment of antioxidant and tyrosinase inhibition activities in relation to enzymatic browning control has not been comprehensively addressed. In particular, the use of cucumber mucilage as a standalone, bioactive edible coating to mitigate browning in fresh-cut apple slices represents a novel approach. By linking extraction strategy, structural functionality, and practical application in a fresh-cut fruit model, this study addresses a critical gap and advances the utilization of cucumber mucilage as a sustainable and multifunctional biopolymer for postharvest quality preservation.

Therefore, the present study aims to extract mucilage from Cucumis sativus L. using microwave-assisted extraction, characterize its physicochemical and structural properties, and evaluate its functional potential with particular emphasis on antioxidant and tyrosinase inhibition activities. Furthermore, the application of cucumber mucilage as a natural edible coating on fresh-cut apple slices was investigated to assess its effectiveness in controlling enzymatic browning and enhancing postharvest quality. This work highlights the valorization of cucumber-derived mucilage as a sustainable, bioactive, and multifunctional biopolymer for food preservation applications.

2. Materials and Methods

2.1. Chemicals and Reagents

Throughout the experiments, triple-distilled water was consistently employed to maintain reagent integrity. The following compounds were procured from Sigma-Aldrich: 2,2-diphenyl-1-picrylhydrazyl (DPPH), 2,4,6-tripyridyl-s-triazine (TPTZ), ferric chloride hexahydrate (FeCl₃·6H₂O), potassium persulfate (K₂S₂O₈), 2,2′-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) (ABTS), bovine serum albumin (BSA), L-tyrosine, L-3,4-dihydroxyphenylalanine (L-DOPA), tyrosinase enzyme, copper sulfate (CuSO₄), pyrocatechol violet, sodium hydroxide (NaOH), and ethanol (95%). All reagents were prepared fresh before use.

2.2. Raw Material and Sample Preparation

The raw material used in the present study was Indian field cucumber (Cucumis sativus L.), belonging to the Poona Kheera. The fruits were harvested at the physiological maturity stage (7 weeks after flowering), characterized by a yellow to yellow–orange skin color and fully developed. This maturity stage was selected as mucilage accumulation is highest in the seed coat and placental tissues at physiological maturity, making it suitable for mucilage extraction.

2.3. Extraction of Mucilage

Extraction of mucilage from fresh Cucumis sativus L. was performed using the microwave extraction process by following the methodology of Fernandes et al. [12]. The outer peel of the fruits was manually removed and discarded. The whole fruit pulp, including the placental tissue and seeds, was used as the extraction material, as mucilage is predominantly localized in the seed coat and surrounding parenchymatous tissues. A 50 g sample was dispersed in 1000 mL of triple distilled water (1:20 w/v) and subjected to microwave heating at different power levels (180, 360, and 540 W) for varying durations (3–5 min) with intermittent stirring. During extraction, the temperature of the extraction medium was monitored using a digital probe thermometer. The temperature increased with microwave power and exposure time and ranged between 45–65 °C at 180 W, 55–75 °C at 360 W, and 65–85 °C at 540 W. After cooling, filtration of the mixture was carried out using Whatman filter paper to eliminate particulate matter. The filtrate was then subjected to ethanol precipitation by the addition of ethanol (1:3 v/v) to facilitate mucilage separation. The precipitated mucilage was allowed to settle, collected by filtration, and subsequently freeze-dried to obtain mucilage powder. The dried mucilage powder was collected and sealed in airtight containers. The purity of the extracted Cucumis sativus L. mucilage was estimated based on its total polysaccharide content.

2.4. Characterization of Cucumis sativus L. Mucilage Structure

2.4.1. Monosaccharide Composition

The monosaccharide composition of Cucumis sativus L. mucilage was determined using HPLC following the method of Bazezew et al. [13]. In brief, 100 mg of the mucilage powder was mixed with 100 mL of 2 M trifluoroacetic acid and heated at 100 °C for 10 min to convert the polysaccharides into their constituent sugars. After cooling to room temperature, the hydrolyzed solution was passed through a 0.45 µm syringe filter and the clarified sample was analyzed using an HPLC (High Performance Liquid Chromatography) system fitted with a refractive index detector. The separation was carried out at 30 °C with a flow rate maintained between 0.5 and 1.0 mL/min, using a 20 µL injection volume and a mobile phase of acetonitrile and water in a 75:25 ratio. Identification and quantification of monosaccharides were performed by matching sample peak profiles with those obtained from calibrated sugar standards run under the same chromatographic conditions.

2.4.2. Morphological Characteristics

The surface structural analysis of the Cucumis sativus L. mucilage powder was investigating using Scanning Electron Microscopy (SEM) by following the procedure given by Geng et al. [14]. Prior to imaging, the dried sample powder (5 mg) was prepared by mounting on the aluminum stub and gold coated using a sputter coater (Quorum Q150R ES) to enhance conductivity and prevent charging under the electron beam. The samples were imaged under high vacuum with an accelerating voltage of 20.0 kV and micrographs were captured at varying magnifications (200× to 500×) to observe surface features.

2.4.3. Particle Size and Surface Charge

The measurement of the particle diameter and surface charge of the Cucumis sativus L. mucilage powder was carried out using a Zetasizer Nano ZS90 (Beckman Coulter Inc., Brea, CA, USA) equipped with dynamic light scattering based on the procedure given by Yu et al. [15]. An aqueous dispersion was prepared by sonicating the powder sample (10 mg) in deionized water (10 mL) for 10 min to ensure uniform dispersion and prevent agglomeration. The particle size was measured based on the intensity-weighted distribution using dynamic light scattering at 25 °C. Three consecutive measurements were performed for each sample.

2.4.4. Fourier Transform Infrared (FTIR) Spectroscopy

The molecular configuration and characteristic chemical groups of Cucumis sativus L. mucilage powder were analyzed using FTIR spectroscopy by following the method of Chen et al. [16]. Briefly, 2 mg of the finely Cucumis sativus L. powder mucilage was placed directly onto the diamond ATR (Attenuated Total Reflectance) crystal. Spectra were recorded in mid infra-red range (400–4000 cm⁻¹).

2.4.5. X-Ray Diffraction

The Cucumis sativus L. mucilage powder’s crystallinity was characterized through X-ray diffraction analysis by following the method of Wang et al. [17]. The scan was carried from 10° to 50° (2θ), with a scan speed of 10°/min. The sample was evenly spread on a glass sample holder to ensure a flat surface before analysis.

2.4.6. Thermal Analysis

Thermal profiling of Cucumis sativus L. mucilage powder was performed using a DSC (Differential Scanning Calorimetry) Q2000 (TA Instruments, New Castle, DE, USA) employing the procedure reported by Xu et al. [18]. A total of 10 mg of the Cucumis sativus L. mucilage powder was enclosed in an aluminum pan, with an identical pan (empty) used as a control. Thermal scanning was carried out on the sample from 15 °C to 450 °C at a rate of 10 °C/min.

2.4.7. Thermogravimetric Analysis (TGA)

Thermal stability of Cucumis sativus L. mucilage powder was evaluated using a TGA Q500 (TA Instruments, New Castle, DE, USA) by following the procedure of Bhiri et al. [19]. A total of 10 mg of Cucumis sativus L. mucilage powder was loaded into a platinum crucible and subjected to thermal analysis at temperatures from 10 °C to 950 °C at a rate of 10 °C/min. Thermal scanning above 400 °C was conducted to ensure complete thermal decomposition of the mucilage and to obtain a comprehensive thermal stability and degradation profile.

2.5. Functional Properties

2.5.1. Water and Oil Holding Capacity

The capacity of Cucumis sativus L. mucilage powder to retain water and oil was determined by the gravimetric method following the protocol described by Wang et al. [20]. A total of 1000 mg of Cucumis sativus L. mucilage powder was dispersed in 20 mL of deionized water (for water holding capacity) or 20 mL of olive oil (for oil holding capacity) utilizing centrifuge tubes that had been already weighed. Each sample was vortexed for 1 min and then left at 25 °C before being centrifuged at 5000× g for 20 min. The clear liquid layer was discarded, and the tubes were weighed again. The retained quantity of water or oil was measured and represented as grams per gram of dry matter using Equation (1).

$$\mathrm{W}\mathrm{H}\mathrm{C}/\mathrm{O}\mathrm{H}\mathrm{C}\left(\mathrm{g}/\mathrm{g}\right)=\left[\left({\displaystyle \frac{\mathrm{H}\mathrm{y}\mathrm{d}\mathrm{r}\mathrm{a}\mathrm{t}\mathrm{e}\mathrm{d} \mathrm{w}\mathrm{e}\mathrm{i}\mathrm{g}\mathrm{h}\mathrm{t}\left(\mathrm{g}\right)-\mathrm{D}\mathrm{r}\mathrm{y} \mathrm{w}\mathrm{e}\mathrm{i}\mathrm{g}\mathrm{h}\mathrm{t} \left(\mathrm{g}\right)}{\mathrm{D}\mathrm{r}\mathrm{y} \mathrm{w}\mathrm{e}\mathrm{i}\mathrm{g}\mathrm{h}\mathrm{t} \left(\mathrm{g}\right)}}\right)\right]$$

(1)

2.5.2. Foaming Capacity

To evaluate the foaming ability and retention, the methodology given by Zhang et al. [21] was employed. One g of Cucumis sativus L. powder was dispersed in 50 mL of deionized water and homogenized at for 2 min at 12,000 rpm. After homogenization, the total volume of the foam and liquid was measured. Foaming capacity was quantified based on percentage increase in volume, as outlined in Equation (2).

$$\mathrm{F}\mathrm{o}\mathrm{a}\mathrm{m}\mathrm{i}\mathrm{n}\mathrm{g} \mathrm{C}\mathrm{a}\mathrm{p}\mathrm{a}\mathrm{c}\mathrm{i}\mathrm{t}\mathrm{y} \left(\%\right)=\left[\left({\displaystyle \frac{\mathrm{I}\mathrm{n}\mathrm{c}\mathrm{r}\mathrm{e}\mathrm{a}\mathrm{s}\mathrm{e} \mathrm{i}\mathrm{n} \mathrm{v}\mathrm{o}\mathrm{l}\mathrm{u}\mathrm{m}\mathrm{e} \mathrm{a}\mathrm{f}\mathrm{t}\mathrm{e}\mathrm{r} \mathrm{w}\mathrm{h}\mathrm{i}\mathrm{p}\mathrm{p}\mathrm{i}\mathrm{n}\mathrm{g} \left(\mathrm{m}\mathrm{L}\right)-\mathrm{I}\mathrm{n}\mathrm{i}\mathrm{t}\mathrm{i}\mathrm{a}\mathrm{l} \mathrm{l}\mathrm{i}\mathrm{q}\mathrm{u}\mathrm{i}\mathrm{d} \mathrm{v}\mathrm{o}\mathrm{l}\mathrm{u}\mathrm{m}\mathrm{e} \mathrm{b}\mathrm{e}\mathrm{f}\mathrm{o}\mathrm{r}\mathrm{e} \mathrm{w}\mathrm{h}\mathrm{i}\mathrm{p}\mathrm{p}\mathrm{i}\mathrm{n}\mathrm{g} \left(\mathrm{m}\mathrm{L}\right)}{\mathrm{I}\mathrm{n}\mathrm{i}\mathrm{t}\mathrm{i}\mathrm{a}\mathrm{l} \mathrm{l}\mathrm{i}\mathrm{q}\mathrm{u}\mathrm{i}\mathrm{d} \mathrm{v}\mathrm{o}\mathrm{l}\mathrm{u}\mathrm{m}\mathrm{e} \mathrm{b}\mathrm{e}\mathrm{f}\mathrm{o}\mathrm{r}\mathrm{e} \mathrm{w}\mathrm{h}\mathrm{i}\mathrm{p}\mathrm{p}\mathrm{i}\mathrm{n}\mathrm{g} \left(\mathrm{m}\mathrm{L}\right)}}\right)\times 100\right]$$

(2)

Stability of the generated foam was assessed after maintaining the sample at 25 °C for 30 min, and the percentage stability was calculated according to Equation (3).

$$\mathrm{F}\mathrm{o}\mathrm{a}\mathrm{m}\mathrm{i}\mathrm{n}\mathrm{g} \mathrm{S}\mathrm{t}\mathrm{a}\mathrm{b}\mathrm{i}\mathrm{l}\mathrm{i}\mathrm{t}\mathrm{y} \left(\%\right)=\left[\left({\displaystyle \frac{\mathrm{F}\mathrm{o}\mathrm{a}\mathrm{m} \mathrm{v}\mathrm{o}\mathrm{l}\mathrm{u}\mathrm{m}\mathrm{e} \mathrm{m}\mathrm{e}\mathrm{a}\mathrm{s}\mathrm{u}\mathrm{r}\mathrm{e}\mathrm{d} \mathrm{a}\mathrm{f}\mathrm{t}\mathrm{e}\mathrm{r} 30 \mathrm{m}\mathrm{i}\mathrm{n} \left(\mathrm{m}\mathrm{L}\right)}{\mathrm{F}\mathrm{o}\mathrm{a}\mathrm{m} \mathrm{v}\mathrm{o}\mathrm{l}\mathrm{u}\mathrm{m}\mathrm{e} \mathrm{r}\mathrm{e}\mathrm{c}\mathrm{o}\mathrm{r}\mathrm{d}\mathrm{e}\mathrm{d} \mathrm{i}\mathrm{m}\mathrm{m}\mathrm{e}\mathrm{d}\mathrm{i}\mathrm{a}\mathrm{t}\mathrm{e}\mathrm{l}\mathrm{y} \mathrm{a}\mathrm{f}\mathrm{t}\mathrm{e}\mathrm{r} \mathrm{w}\mathrm{h}\mathrm{i}\mathrm{p}\mathrm{p}\mathrm{i}\mathrm{n}\mathrm{g} \left(\mathrm{m}\mathrm{L}\right)}}\right)\times 100\right]$$

(3)

2.5.3. Emulsifying Capacity

The evaluation of emulsification properties was carried out according to the technique presented by Lv et al. [22]. A 1 g sample of Cucumis sativus L. mucilage powder was dispersed in 20 mL of deionized water followed by homogenization for 3 min at 8500 rpm. Then, 20 mL of olive oil was gradually added while continuing homogenization for 2 min to form an emulsion. To quantify emulsifying capacity, the volume of the emulsion was measured immediately after homogenization, and the result was expressed as a percentage using Equation (4).

$$\mathrm{E}\mathrm{m}\mathrm{u}\mathrm{l}\mathrm{s}\mathrm{i}\mathrm{f}\mathrm{y}\mathrm{i}\mathrm{n}\mathrm{g} \mathrm{C}\mathrm{a}\mathrm{p}\mathrm{a}\mathrm{c}\mathrm{i}\mathrm{t}\mathrm{y}\left(\%\right)=\left[\left({\displaystyle \frac{\mathrm{H}\mathrm{e}\mathrm{i}\mathrm{g}\mathrm{h}\mathrm{t} \mathrm{o}\mathrm{f} \mathrm{s}\mathrm{t}\mathrm{a}\mathrm{b}\mathrm{l}\mathrm{e} \mathrm{e}\mathrm{m}\mathrm{u}\mathrm{l}\mathrm{s}\mathrm{i}\mathrm{o}\mathrm{n} \mathrm{l}\mathrm{a}\mathrm{y}\mathrm{e}\mathrm{r} \mathrm{f}\mathrm{o}\mathrm{r}\mathrm{m}\mathrm{e}\mathrm{d} \left(\mathrm{c}\mathrm{m}\right)}{\mathrm{T}\mathrm{o}\mathrm{t}\mathrm{a}\mathrm{l} \mathrm{c}\mathrm{o}\mathrm{l}\mathrm{u}\mathrm{m}\mathrm{n} \mathrm{h}\mathrm{e}\mathrm{i}\mathrm{g}\mathrm{h}\mathrm{t} \left(\mathrm{c}\mathrm{m}\right)}}\right)\times 100\right]$$

(4)

The emulsion was transferred into the graduated cylinders and incubated at 25 °C for 24 h. After this period, the volume of the separated oil layer was recorded. Emulsifying stability (%) was calculated using the formula given in Equation (5)

$$\mathrm{S}\mathrm{t}\mathrm{a}\mathrm{b}\mathrm{i}\mathrm{l}\mathrm{i}\mathrm{t}\mathrm{y} \mathrm{o}\mathrm{f} \mathrm{e}\mathrm{m}\mathrm{u}\mathrm{l}\mathrm{s}\mathrm{i}\mathrm{o}\mathrm{n}\left(\%\right)=\left[\left({\displaystyle \frac{\mathrm{H}\mathrm{e}\mathrm{i}\mathrm{g}\mathrm{h}\mathrm{t} \mathrm{o}\mathrm{f} \mathrm{e}\mathrm{m}\mathrm{u}\mathrm{l}\mathrm{s}\mathrm{i}\mathrm{f}\mathrm{i}\mathrm{e}\mathrm{d} \mathrm{l}\mathrm{a}\mathrm{y}\mathrm{e}\mathrm{r} \mathrm{a}\mathrm{f}\mathrm{t}\mathrm{e}\mathrm{r} \mathrm{r}\mathrm{e}\mathrm{s}\mathrm{t}\mathrm{i}\mathrm{n}\mathrm{g} \left(\mathrm{c}\mathrm{m}\right)}{\mathrm{I}\mathrm{n}\mathrm{i}\mathrm{t}\mathrm{i}\mathrm{a}\mathrm{l} \mathrm{h}\mathrm{e}\mathrm{i}\mathrm{g}\mathrm{h}\mathrm{t} \left(\mathrm{c}\mathrm{m}\right)}}\right)\times 100\right]$$

(5)

2.5.4. Solubility

The method reported by Guo et al. [23] was used to assess the solubility of Cucumis sativus L. mucilage powder by dispersing 1 g of the powder in 20 mL of deionized water in a centrifuge tube. The solution was continuously stirred for 30 min at 25 °C followed by centrifugation at 5000× g for 15 min. The upper layer was collected and dried at 105 °C in a hot air oven. Solubility (%) was determined according to Equation (6).

$$\mathrm{S}\mathrm{o}\mathrm{l}\mathrm{u}\mathrm{b}\mathrm{i}\mathrm{l}\mathrm{i}\mathrm{t}\mathrm{y}\left(\%\right)=\left[\left({\displaystyle \frac{\mathrm{D}\mathrm{r}\mathrm{i}\mathrm{e}\mathrm{d} \mathrm{w}\mathrm{e}\mathrm{i}\mathrm{g}\mathrm{h}\mathrm{t} \mathrm{o}\mathrm{f} \mathrm{s}\mathrm{u}\mathrm{p}\mathrm{e}\mathrm{r}\mathrm{n}\mathrm{a}\mathrm{t}\mathrm{e}\mathrm{n}\mathrm{t} \left(\mathrm{g}\right)}{\mathrm{W}\mathrm{e}\mathrm{i}\mathrm{g}\mathrm{h}\mathrm{t} \mathrm{o}\mathrm{f} \mathrm{m}\mathrm{u}\mathrm{c}\mathrm{i}\mathrm{l}\mathrm{a}\mathrm{g}\mathrm{e} \left(\mathrm{g}\right)}}\right)\times 100\right]$$

(6)

2.6. Biological Activity of Mucilage Powder

2.6.1. Antioxidant Activity

Ferric Ion Reducing Capacity of Cucumis sativus L. Mucilage

The FRAP (ferric reducing antioxidant power) assay was performed by following the method of Perumal et al. [24]. A freshly prepared FRAP solution was obtained by blending 300 mM sodium acetate buffer (pH 3.6), 10 mM TPTZ in 40 mM HCl, and 20 mM FeCl₃·6H₂O in a 10:1:1 (v/v/v) proportion. A volume of 8.1 µL of Cucumis sativus L. mucilage extract (2 to 10 mg/mL) was combined with 600 µL of freshly prepared FRAP working solution. The reaction mixture was incubated at 37 °C for 1 h to facilitate the reduction process. The absorbance of the Fe²⁺–TPTZ complex was recorded at 593 nm, and a standard curve was generated using 0–5 mM FeSO₄·7H₂O. Results were expressed in mM FeSO4.

Analysis of DPPH Free Radical Inhibition Potential

The antioxidant potential of Cucumis sativus L. mucilage powder was evaluated using DPPH by following the method reported by Zhang et al. [25]. DPPH was dissolved in ethanol to prepare a 0.1 mM working solution. A 1 mL volume of Cucumis sativus L. mucilage extract (2–10 mg/mL) was blended with 1 mL of DPPH reagent and homogenized. Incubation was carried out at 25 °C for 30 min. Ascorbic acid was used as the standard antioxidant reference. The absorbance was then recorded at 517 nm.

Evaluation of ABTS Radical Scavenging Potential

ABTS•⁺ scavenging potential was assessed using the methodology reported by Alias & Shafie, [26]. An equimolar mixture of 7 mM ABTS and 2.45 mM potassium persulfate was incubated in the dark at ambient temperature for 16 h to generate the ABTS radical cation. To prepare for analysis, the ABTS•⁺ solution was diluted with methanol to obtain an absorbance value of 0.700 at 734 nm. For the assay, 600 µL of the ABTS•⁺ working solution was mixed with 50 µL (2–10 mg/mL) of the mucilage sample. After a 6-minute dark incubation, absorbance was monitored at 734 nm to assess the change. The ABTS radical scavenging activity was calculated using the formula as given in Equation (7).

$$\mathrm{I}\mathrm{n}\mathrm{h}\mathrm{i}\mathrm{b}\mathrm{i}\mathrm{t}\mathrm{i}\mathrm{o}\mathrm{n} \mathrm{A}\mathrm{c}\mathrm{t}\mathrm{i}\mathrm{v}\mathrm{i}\mathrm{t}\mathrm{y}\left(\%\right)=\left[\left({\displaystyle \frac{\mathrm{A}\mathrm{b}\mathrm{s}\mathrm{o}\mathrm{r}\mathrm{b}\mathrm{a}\mathrm{n}\mathrm{c}\mathrm{e} \mathrm{o}\mathrm{f} \mathrm{c}\mathrm{o}\mathrm{n}\mathrm{t}\mathrm{r}\mathrm{o}\mathrm{l}-\mathrm{A}\mathrm{b}\mathrm{s}\mathrm{o}\mathrm{r}\mathrm{b}\mathrm{a}\mathrm{n}\mathrm{c}\mathrm{e} \mathrm{o}\mathrm{f} \mathrm{s}\mathrm{a}\mathrm{m}\mathrm{p}\mathrm{l}\mathrm{e}}{\mathrm{A}\mathrm{b}\mathrm{s}\mathrm{o}\mathrm{r}\mathrm{b}\mathrm{a}\mathrm{n}\mathrm{c}\mathrm{e} \mathrm{o}\mathrm{f} \mathrm{c}\mathrm{o}\mathrm{n}\mathrm{t}\mathrm{r}\mathrm{o}\mathrm{l}}}\right)\times 100\right]$$

(7)

2.6.2. Anti-Inflammatory Activity

Cell Culture

RAW 264.7 murine macrophage cells were maintained in Dulbecco’s modified Eagle medium (DMEM) supplemented with 10% (v/v) fetal bovine serum (FBS) and 1% (v/v) penicillin-streptomycin (containing 100 U/mL penicillin and 100 mg/mL streptomycin). The cells were cultured under standard conditions at 37 °C in a humidified incubator with 5% CO₂.

Cell Viability Test

The effect of Cucumis sativus L. mucilage on RAW 264.7 cell viability was evaluated using the MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) assay by following the method of Zheng et al. [27]. Cells were distributed in 96-well culture plates at 1 × 10⁵ cells/well and incubated for 24 h at 37 °C. After incubation, the cells were treated with 10 μL of different concentrations of the Cucumis sativus L. mucilage samples (25, 50, 100, 200, and 500 μg/mL) to make a final volume of 100 μL/well followed by incubation for 24 h. For the positive control, cells were treated with 5 μg/mL LPS (lipopolysaccharide); the blank control consisted of the same volume of DMEM. Following treatment, 20 µL of 5 mg/mL MTT in PBS (phosphate-buffered saline) was added to wells and incubated at 37 °C for 4 h for formazan synthesis. After removing the incubation medium, 100 µL of DMSO was introduced into each well to solubilize the formazan crystals formed. To ensure complete dissolution of the formazan crystals, the plate was softly shaken, followed by measuring absorbance at 570 nm with a microplate reader. Cell viability percentage was determined using Equation (8).

$$\mathrm{C}\mathrm{e}\mathrm{l}\mathrm{l} \mathrm{v}\mathrm{i}\mathrm{a}\mathrm{b}\mathrm{i}\mathrm{l}\mathrm{i}\mathrm{t}\mathrm{y}\left(\%\right)=\left[\left({\displaystyle \frac{\mathrm{A}\mathrm{b}\mathrm{s}\mathrm{o}\mathrm{r}\mathrm{b}\mathrm{a}\mathrm{n}\mathrm{c}\mathrm{e} \mathrm{o}\mathrm{f} \mathrm{c}\mathrm{e}\mathrm{l}\mathrm{l} \mathrm{c}\mathrm{u}\mathrm{l}\mathrm{t}\mathrm{u}\mathrm{r}\mathrm{e} \mathrm{c}\mathrm{o}\mathrm{n}\mathrm{t}\mathrm{a}\mathrm{i}\mathrm{n}\mathrm{i}\mathrm{n}\mathrm{g} \mathrm{m}\mathrm{u}\mathrm{c}\mathrm{i}\mathrm{l}\mathrm{a}\mathrm{g}\mathrm{e} \mathrm{s}\mathrm{o}\mathrm{l}\mathrm{u}\mathrm{t}\mathrm{i}\mathrm{o}\mathrm{n}}{\mathrm{A}\mathrm{b}\mathrm{s}\mathrm{o}\mathrm{r}\mathrm{b}\mathrm{a}\mathrm{n}\mathrm{c}\mathrm{e} \mathrm{o}\mathrm{f} \mathrm{c}\mathrm{e}\mathrm{l}\mathrm{l} \mathrm{c}\mathrm{u}\mathrm{l}\mathrm{t}\mathrm{u}\mathrm{r}\mathrm{e} \mathrm{w}\mathrm{i}\mathrm{t}\mathrm{h}\mathrm{o}\mathrm{u}\mathrm{t} \mathrm{m}\mathrm{u}\mathrm{c}\mathrm{i}\mathrm{l}\mathrm{a}\mathrm{g}\mathrm{e} \mathrm{s}\mathrm{o}\mathrm{l}\mathrm{u}\mathrm{t}\mathrm{i}\mathrm{o}\mathrm{n}}}\right)\times 100\right]$$

(8)

Determination of NO Release in Activated Macrophages

The Griess reagent method was employed to quantify NO production in RAW 264.7 cells. A total of 2 × 10⁵ RAW 264.7 cells per well were cultured in 24-well plates and left overnight to allow cell adherence. Cells were pre-incubated with Cucumis sativus L. mucilage at concentrations of 0 (control), 25, 50, 100, 200, and 500 μg/mL for 2 h, after which lipopolysaccharide (5 mg/mL) was introduced to induce stimulation to induce NO production. The cells were further incubated for 24 h at 37 °C in a humidified 5% CO₂ atmosphere. After incubation, 100 µL of the culture supernatant from each well was collected and mixed with an equal volume (100 µL) of Griess reagent in a 96-well microplate. The mixture was incubated at room temperature for 10 min in the dark, allowing for the formation of a purple azo dye. Measurement of absorbance was carried out at a wavelength of 540 nm using a plate reader.

2.6.3. Determination of Tyrosinase-Inhibition Activity

Analysis of Monophenolase Enzyme Inhibition

The ability of Cucumis sativus L. mucilage to inhibit monophenolase was determined using the protocol proposed by Al-Ajalein et al. [28] using the inhibition of L-tyrosine oxidation. A total reaction volume was prepared by combining 180 µL of 50 mM phosphate buffer (pH 6.8), 40 µL of L-tyrosine (0.5 mM), and 10 µL of mucilage sample at varying concentrations (0.5, 1, 1.5, 2.0 mg/mL) in a 96-well microplate, followed by incubation at 37 °C for 30 min. An aliquot of 1 μL tyrosinase solution (activity: 6250 U/mL) was then added to initiate the reaction. The optical density at 470 nm was continuously monitored at 20-s intervals over 15-min. The percentage inhibition was calculated relative to a control without mucilage sample.

Analysis of Diphenolase Enzyme Inhibition

Diphenolase inhibitory activity was assessed by monitoring the oxidation of L-DOPA. The reaction mixture contained 2.5 mL of 0.1 M phosphate buffer (pH 6.8), 40 µL of L-DOPA (0.5 mM), 10 µL of mucilage sample at different concentrations (0.5, 1, 1.5, 2.0 mg/mL) that was added to a 96-well plate and incubated at 37 °C for 30 min. After incubation, the dopachrome formation was measured at 470 nm at every 10 s over a period of 1 min. Inhibition percentage was calculated by comparison with the control.

2.7. Preparation of Cucumber Mucilage-Based Coating

Cucumber mucilage edible coating solutions were prepared by dissolving dried mucilage powder in distilled water to obtain concentrations of 0.5, 1.0, and 2.0 mg/mL. The solutions were stirred at ambient temperature until complete dissolution and filtered to remove any undissolved particles. The prepared coating solutions were used immediately for the treatment of fresh-cut apple slices. For coating application, fresh apples (Malus domestica cv. Red Delicious) were used in this study due to their high susceptibility to enzymatic browning. The apples were washed and cut into uniform slices. The slices were immediately dipped for 2 min in (i) distilled water (blank group), (ii) 0.01 g/mL kojic acid solution (positive control), or Cucumis sativus L. mucilage solutions at concentrations of 0.5, 1.0, and 2.0 mg/mL (treatment groups). Kojic acid was employed as a positive control because it is a well-known tyrosinase inhibitor commonly used in anti-browning studies. The selected concentration (0.01 g/mL) was used solely for comparative evaluation of anti-browning efficacy. After treatment, the slices were placed in sterile Petri dishes and stored at 4 °C for 5 days.

Anti-Browning Effect on Fresh-Cut Apple Slices

Color parameters (L*, a*, b*) were measured after 24 h intervals using a Minolta CR-400 colorimeter (Konica Minolta, Osaka, Japan). Measurements were taken specifically at the central mesocarp region of each slice to ensure consistency, and three readings per slice were recorded. The total color change (ΔE) was calculated using the CIE76 formula.

$$\Delta \mathrm{E}=\sqrt{{({\mathrm{L}}_2^{*}-{\mathrm{L}}_1^{*})}^2+{({\mathrm{a}}_2^{*}-{\mathrm{a}}_1^{*})}^2+{({\mathrm{b}}_2^{*}-{\mathrm{b}}_1^{*})}^2)}$$

(9)

where ${\mathrm{L}}_1^{*}$, ${\mathrm{a}}_1^{*}$, ${\mathrm{b}}_1^{*}$ = color values of the initial sample (Day 0), ${\mathrm{L}}_2^{*}$, ${\mathrm{a}}_2^{*}$, ${\mathrm{b}}_2^{*}$ = color values of the sample after storage, and the L* value, which is more responsive to browning while ΔE, represents the overall color change.

2.8. Statistical Analysis

All experiments were carried out in triplicate, and the statistical data were expressed as the mean average value ± standard deviation. Analysis of variance and Duncan’s multiple range tests were employed using SPSS statistical software 17.0 (SPSS, Inc., Chicago, IL, USA). A significance level of p ≤ 0.05 was used to determine statistical significance.

3. Results and Discussion

3.1. Extraction Yield of Mucilage

Microwave-assisted extraction significantly influenced the yield of mucilage from Cucumis sativus L., as shown in

Table 1. Optimization of Cucumis sativus L. mucilage yield by at different microwave power and extraction time ¹.

Microwave Power (W)	Extraction Time (min)	Yield (%)
180	3	15.51 ± 1.14 aA
180	4	18.20 ± 1.45 bA
180	5	19.81 ± 1.32 cA
360	3	19.01 ± 1.52 aB
360	4	22.32 ± 1.48 bB
360	5	23.81 ± 1.23 cB
540	3	23.42 ± 1.31 aC
540	4	25.56 ± 1.69 cC
540	5	24.01 ± 1.56 bC

¹ Data are presented as the mean ± SD (n = 3). Mean values with different lowercase (a–c) letters within a column represent significantly different values within the extraction conditions and uppercase (A–C) for extraction time, based on analysis of variance (ANOVA) and post hoc tests.

. The yield increased with both microwave power and extraction time, indicating that higher energy input and prolonged exposure enhanced mass transfer and disrupted cell walls, facilitating mucilage release. At 180 W, the extraction yield increased from 15.51% at 3 min to 19.81% at 5 min, while at 360 W, yields increased from 19.01% (3 min) to 23.81% (5 min). The highest yield (25.56%) was obtained at 540 W for 4 min, after which a slight decline (24.01%) was observed at 5 min, likely due to thermal degradation or excessive breakdown of polysaccharide chains. In addition to yield optimization, the mucilage extracted under the optimized condition (540 W, 4 min) exhibited a high polysaccharide purity of 79.56%, confirming that microwave-assisted extraction coupled with ethanol precipitation effectively recovered mucilage with minimal non-polysaccharide impurities.

Microwave-assisted extraction enhances mucilage recovery primarily through rapid volumetric heating caused by dipole rotation and ionic conduction, which disrupt hydrogen bonding within plant cell walls and promote the release of intracellular polysaccharides. Increasing the microwave power intensifies internal heating and pressure buildup, leading to improved cell wall rupture and mass transfer [29]. Similarly, longer exposure time increases solvent penetration and polysaccharide solubilization. However, excessive microwave energy or prolonged exposure, as observed at 540 W for 5 min, may result in partial depolymerization or thermal degradation of polysaccharide chains, reducing viscosity and effective mucilage recovery. Therefore, the interaction between microwave power and extraction time is critical, and the optimized condition (540 W, 4 min) represents a balance between enhanced cell disruption and the preservation of mucilage structural integrity.

3.2. Structural Characterization of Mucilage

3.2.1. Monosaccharide Composition

The monosaccharide composition of mucilage directly influences its functional properties and application potential. The sugar profile of Cucumis sativus L. mucilage is presented in

Table 2. Monosaccharide composition of Cucumis sativus L. mucilage ¹.

Monosaccharide	Composition (%)
Glucose	32.27 ± 0.09 h
Arabinose	8.09 ± 0.05 d
Galactose	13.19 ± 0.16 f
Xylose	6.89 ± 0.15 c
Galacturonic acid	4.56 ± 0.12 a
Glucuronic acid	4.60 ± 0.18 b
Mannose	20.26 ± 0.04 g
Rhamnose	9.70 ± 0.23 e

¹ Data are presented as the mean ± SD (n = 3). Mean values with different lowercase (a–h) letters within a column represent significantly different values.

, revealing a heterogeneous polysaccharide structure dominated by neutral sugars with a small amount of uronic acids. Glucose was the most abundant monosaccharide (32.27%), followed by mannose (20.26%) and galactose (13.19%), indicating that the mucilage primarily consists of glucan and mannan rich polysaccharide fractions. Arabinose (8.09%) and rhamnose (9.70%) were present, implying the occurrence of branched polysaccharide structures such as arabinogalactans or rhamnogalacturonan-type domains [30]. These sugar residues are commonly associated with side-chain branching, which enhances water retention, solubility, and the emulsifying behavior of plant-derived mucilage. Xylose (6.89%) further supports the presence of hemi cellulosic components contributing to structural complexity, and is associated with improved functional and rheological behavior, particularly shear-thinning characteristics [13]. Uronic acids, including galacturonic acid (4.56%) and glucuronic acid (4.60%), were present in lower proportions than neutral sugars, however, these acidic residues are essential for imparting anionic character to the mucilage, enabling electrostatic interactions, metal ion binding, and enhanced gel-forming ability. Their presence also contributes to the pH-dependent rheological behavior and functional versatility of the mucilage in food and pharmaceutical applications. The composition of monosaccharides, including glucose, mannose, galactose, and uronic acids, affects the viscosity, gelling ability, and water-binding properties of the mucilage, which are crucial for forming a uniform and stable coating.

3.2.2. Morphological Analysis

The surface morphology of Cucumis sativus L. mucilage powder was examined using SEM at magnifications of 200× and 500× (Figure 1A). At lower magnification (200×), the mucilage particles exhibited an irregular, heterogeneous morphology with a wide particle size distribution. The particles appeared as fragmented aggregates with rough, uneven surfaces, indicating the absence of a well-defined crystalline structure. Such irregularity is characteristic of plant-derived polysaccharides obtained through drying and grinding processes and suggests partial collapse of the native polymer matrix during extraction and dehydration. At higher magnification (500×), the surface features became more distinct, revealing angular, flaky, and fractured particles with roughness at the surface. The presence of cracks, broken edges, and porous regions indicates structural disruption of the cell wall matrix, likely caused by microwave-assisted extraction. Microwave treatment generates internal pressure, leading to cell wall rupture and the fragmentation of polysaccharide networks [31]. This structural breakdown facilitates mucilage release but also results in irregular particle morphology upon drying. The rough and porous surface topology observed in the mucilage powder is advantageous from a functional perspective, as it may enhance water absorption, swelling capacity, and the hydration rate [32]. Increased surface roughness provides a larger effective surface area, which can improve the interaction with water and other components in food and pharmaceutical formulations. Overall, the irregular, porous, and fractured surface morphology promotes rapid hydration and enhances adhesion when applied as a coating, contributing to even coverage and better moisture retention.

3.2.3. Particle Size and Charge Potential

The average particle diameter of powdered Cucumis sativus L. mucilage is presented in Figure 1B, showing a mean particle size of 194.5 nm, which indicates a relatively uniform dispersion. This small particle size suggests the efficiency of the microwave extraction in breaking down the mucilage structure into fine particles with a high surface area [33]. The submicron range of the particles suggests that the mucilage powder may possess enhanced solubility, dispersibility, and potential bioavailability when rehydrated in aqueous systems [34]. In addition, the charge potential of the mucilage particles was observed as −19.8 mV. Zeta potential serves as an essential indicator of colloidal stability, representing the electrostatic repulsion between similarly charged particles in suspension [35]. The negative charge can mainly be linked to its acidic functional moieties in the mucilage structure, such as uronic acids and hydroxyl groups commonly found in plant-derived polysaccharides [32]. In aqueous solutions, these groups may undergo dissociation, imparting a negative charge to the particle surface and promoting dispersion stability through electrostatic repulsion. Similar trends have been reported by Sharma et al. [36] in other plant mucilage. In conclusion, the small particle size and moderate negative zeta potential ensure good dispersion in coating solutions, prevent aggregation, and allow for the formation of smooth, continuous films on the fruit surface.

3.2.4. Fourier Transform Infrared Spectroscopy

The FTIR spectrum of Cucumis sativus L. mucilage, illustrated in Figure 1C, exhibited distinct absorption bands characteristic of multiple functional groups. A wide and strong absorption band appearing at 3290.10 cm⁻¹ indicates O–H stretching vibrations, reflecting the presence of abundant hydroxyl groups and strong hydrogen bonding within the mucilage matrix. The spectral feature at 2921.13 cm⁻¹ is indicative to C–H stretching of aliphatic chains, typical of carbohydrate structures [37]. A distinct absorption at 1642.16 cm⁻¹ is associated with the bending vibrations of bound water and possible C=O stretching of uronic acid groups, indicating the presence of polysaccharides [38]. The band at 1363.10 cm⁻¹ suggests C–H bending, likely from methyl or methylene groups. A strong band at 1149.03, 1074.13, 1032.16, and 1014.17 cm⁻¹ is assigned to C–O–H and C–O–C bending and stretching vibrations of glycosidic linkages, representing the polysaccharide fingerprint region [39]. The peak at 932.60 cm⁻¹ corresponds to β-glycosidic linkages, confirming the presence of β-anomeric configurations in the sugar units. Various peaks at 851.11 cm⁻¹ and 760.17 cm⁻¹ are related to the ring vibrations of pyranose structures, while the lower frequency peaks at 702.10, 571.76, 522.15, and 436.60 cm⁻¹ represent skeletal bending of the polysaccharide backbone [40]. These results collectively confirm the polysaccharide-rich nature of Cucumis sativus L. mucilage and that various functional group linkages contribute significantly to its functional properties like water retention, emulsification, and gel formation. Overall, the FTIR spectra validate that Cucumis sativus L. mucilage is a polysaccharide-rich biopolymer with functional groups that support its potential as a natural hydrocolloid in food.

3.2.5. X-Ray Diffraction

The diffraction profile obtained from the X-ray analysis of Cucumis sativus L. mucilage is shown in Figure 1D. The diffractogram displays a broad peak indicating that the mucilage is in an amorphous state. The absence of well-defined peaks confirms the lack of long-range molecular order [41]. The broad peak observed in the pattern suggests that the mucilage possesses a disordered internal structure, likely due to the random arrangement of sugar residues, branched chains, and associated functional groups. This amorphous nature is advantageous in food applications as it contributes to enhanced solubility, higher water absorption capacity, and improved swelling behavior [42]. The drying process itself contributes to the disruption of semi-crystalline order present in the mucilage. Overall, the amorphous structure facilitates solubility and swelling, which helps the coating to spread evenly and maintain flexibility.

3.2.6. Differential Scanning Calorimetry

The thermal transitions of Cucumis sativus L. mucilage are represented in Figure 1E. The DSC curve displays two major endothermic transitions, corresponding to different thermal events typically observed in mucilage. The first endothermic peak appeared at a temperature of 93.69 °C, with an onset at 26.38 °C and an endset at 137.56 °C. This transition is primarily attributed to the bound water evaporation and moisture loss from the mucilage matrix [43]. The relatively broad transition and high enthalpy value (ΔH = 233.0613 J/g) indicate a strong water retention capacity of the Cucumis sativus L. mucilage, which is characteristic of polysaccharides with multiple hydroxyl and carboxyl groups capable of forming hydrogen bonds [32]. The second thermal event occurred at a temperature of 227.04 °C, with an onset at 215.15 °C and an endset at 259.80 °C. This peak corresponds to the thermal degradation or structural rearrangement of the polysaccharide backbone, possibly due to depolymerization or the decomposition of complex carbohydrate chains [44]. The enthalpy associated with this transition (ΔH = 66.6859 J/g) and its peak height (3.4990 mW) suggest a significant energy requirement for breaking molecular interactions, indicating moderate thermal stability of the mucilage. The presence of two distinct endothermic events suggests that the mucilage has both good water retention ability and moderate thermal resistance, making it suitable for thermally processed formulations [45]. Additionally, the relatively high thermal decomposition point (>220 °C) indicates that the mucilage can retain its structure under elevated temperatures, which is desirable in food applications.

3.2.7. Thermogravimetric Analysis

The thermal degradation profile of Cucumis sativus L. mucilage was performed using TGA, and the graph is presented in Figure 1F. The TGA curve revealed a degradation pattern indicative of the complex composition of the mucilage. The overall weight loss was 80.95%, demonstrating substantial thermal decomposition upon heating. The first weight loss phase, observed between 51.20 °C and 210.75 °C, accounted for 7.79% of the total weight loss. This initial loss may be due to the surface and loosely bound water evaporation, a typical feature of hydrophilic biopolymers like mucilage that contain numerous hydroxyl groups capable of hydrogen bonding with water [46]. The second major degradation stage occurred between 210.75 °C and 293.13 °C, corresponding to the mass reduction of 31.31%. This phase likely corresponds to the depolymerization and degradation of low-molecular-weight polysaccharides and side chains, such as uronic acids and neutral sugars [47]. The breakdown of glycosidic linkages and partial volatilization of organic constituents occurs during this temperature range, reflecting the onset of structural disintegration. The third and most significant decomposition stage, between 293.13 °C and 439.77 °C, resulted in a weight loss of 40.76%. This sharp drop indicates the thermal decomposition of the primary polysaccharide backbone [48]. The steepness of this phase implies an intense thermal degradation process, which is typical for plant-derived biopolymers at elevated temperatures. After 440 °C, the TGA curve flattens out, suggesting thermal stability of the remaining carbonaceous residue with negligible weight loss beyond this point [49]. The residual mass (19.45%) likely consists of charred, thermally resistant materials or inorganic minerals present in the mucilage. These findings collectively indicate that Cucumis sativus L. mucilage possesses excellent thermal stability, suitable for applications involving heat processing, such as extrusion, baking, or encapsulation.

3.3. Functional Properties

3.3.1. Water and Oil Holding Capacity

The capacity to retain both water and oil by Cucumis sativus L. mucilage powder was 8.46 g/g and 3.21 g/g, respectively, as shown in

Table 3. Functional properties of Cucumis sativus L. mucilage ¹.

Property	Cucumis sativus L. Mucilage
Water holding capacity (g/g)	8.46 ± 0.07
Oil holding capacity (g/g)	3.21 ± 0.04
Foaming capacity (%)	52.13 ± 0.34
Foaming stability (%)	30.46 ± 0.11
Emulsifying capacity (%)	90.45 ± 0.18
Emulsifying stability (%)	91.62 ± 0.16
Solubility (%)	90.14 ± 0.17

¹ Data are presented as mean ± SD (n = 3).

. The high water holding capacity shows that the mucilage has the capability to absorb and maintain high levels of water, which may be due to the presence of hydrophilic groups such as carboxyl and hydroxyl moieties in its polysaccharide structure that facilitate hydrogen bonding with water molecules [50]. Moreover, the amorphous nature of the mucilage, as confirmed by XRD analysis, enhances its water absorption due to increased molecular mobility and open structural arrangement [51]. The high water-binding capacity means that Cucumis sativus L. mucilage can be used effectively as a thickener, stabilizer, or gelling agent in various food formulations, especially in products like bakery items, sauces, and dressings. The high oil holding capacity (3.21 g/g) of Cucumis sativus L. mucilage indicates its capacity to associate with lipophilic substances, likely through physical entrapment or hydrophobic interactions [52]. The high oil holding capacity is useful in emulsified food systems, where oil stabilization is required. The oil holding capacity may also be influenced by the porous microstructure and surface morphology of the mucilage particles, as observed in the scanning electron microscopy images, which provide capillary spaces for oil entrapment [53]. These functional properties are further supported by the presence of chemical groups identified by FTIR, such as –OH and –COOH, which play key roles in binding both water and oil. In conclusion, the high water and oil holding capacity directly enhance the coating’s ability to retain moisture and protect against enzymatic browning in fresh-cut apples.

3.3.2. Foaming Capacity and Stability

Cucumis sativus L. mucilage exhibited a foaming capacity of 52.13% and foaming stability of 30.46%, as shown in Table 3. The foaming capacity reflects the ability of the mucilage to entrap air within a liquid matrix during mechanical agitation [54]. The data demonstrate that the mucilage possesses an excellent ability to generate and retain foam, which is an important functional property in food systems that require aeration and volume enhancement, such as whipped toppings and bakery batters. The foaming capacity of the mucilage may result from its inherent surface-active molecules such as uronic acids, proteins, or amphiphilic polysaccharide fragments that can adsorb at the air–water interface, lowering interfacial tension and facilitating bubble formation [55]. The foaming stability indicates the ability of the foam to resist collapse over time, which is 30.46%. The moderate foaming stability also correlates with the mucilage’s high water holding capacity, which could delay foam breakdown by increasing the viscosity of the surrounding medium. These foaming properties are comparable to those observed for other plant-based mucilages such as X. americana seed mucilage [13], which also exhibit excellent foaming potential due to their polysaccharide-rich, hydrophilic nature. In conclusion, the foaming ability contributes to the formation of uniform, aerated films on the fruit surface, which can improve coating coverage and thickness consistency.

3.3.3. Emulsifying Capacity and Stability

The emulsifying capacity and stability of Cucumis sativus L. mucilage was 90.45% and 91.62%, respectively, as shown in Table 3. These values highlight the mucilage’s significant ability to stabilize oil–water interfaces, making it a promising natural emulsifier for food formulations. The high emulsifying capacity demonstrates the mucilage’s role in emulsion stability by strong interfacial adsorption at the oil–water boundary [56]. The observed effect can be linked to surface-active molecules including uronic acids and some proteinaceous components present in the mucilage, which reduce interfacial tension and promote droplet dispersion. The polysaccharide structure contains both hydrophilic and hydrophobic regions, enabling the formation of a stabilizing film around oil droplets and preventing their coalescence [57]. The remarkable emulsifying stability (91.62%) suggests that the emulsions formed remain intact over time, without significant phase separation or droplet aggregation. This can be attributed to the high viscosity of the mucilage solution and its ability to form a viscoelastic interfacial layer that retards droplet movement and creaming [58]. Moreover, the amorphous microstructure and small particle size of the mucilage, as observed in SEM and particle analysis, may further enhance its emulsifying properties by promoting better surface coverage and barrier formation. The high emulsifying capacity and stability values are comparable to that of cress seed mucilage [59], indicating that Cucumis sativus L. mucilage can be used as an alternative to synthetic emulsifiers like Tween or lecithin in emulsion-based products such as salad dressings, beverages, creams, and sauces. In summary, the strong emulsifying properties enable the stable dispersion of bioactive compounds in the coating solution, preventing phase separation and improving protective performance.

3.3.4. Solubility

Cucumis sativus L. mucilage exhibited a solubility of 90.14%, as shown in Table 3, indicating its excellent ability to dissolve in water. High water solubility is a critical functional property for biopolymers intended for use as hydrocolloids, thickeners, stabilizers, and delivery systems in food applications. The high solubility can be attributed to several physicochemical characteristics of the mucilage [43]. The amorphous nature, as confirmed by XRD analysis, enhances water penetration and molecular mobility, which facilitates rapid dissolution. Moreover, the presence of abundant hydrophilic functional groups (such as hydroxyl and carboxyl) identified by FTIR contributes to strong interactions with water molecules by hydrogen bonding [60]. Additionally, the small particle size and relatively high surface area, confirmed by SEM and particle size analysis, further improve the water accessibility and dispersion of mucilage particles in aqueous systems. Comparable solubility values have been reported for plant-based mucilage such as those extracted from mustard and chia seed [61,62]. Overall, the high solubility ensures rapid dissolution and uniform application of the mucilage as a coating, creating a smooth protective layer over the apple slices.

3.4. Biological Activity of Mucilage Powder

3.4.1. Antioxidant Activity

Ferric Ion Reducing Capacity of Cucumis sativus L. Mucilage

The electron-donating capacity of Cucumis sativus L. mucilage powder was determined using the ferric reducing antioxidant power assay. As shown in Figure 2A, the FRAP values increased steadily with rising concentrations of mucilage from 2 to 10 mg/mL, with reducing power ranging from 2.6 to 5.1 mM FeSO₄ at 10 mg/mL, indicating that antioxidant activity increases proportionally with concentration. The assay reflects the capacity of the antioxidants to reduce Fe³⁺ (ferric) to Fe²⁺ (ferrous), and the observed trend suggests the presence of effective reducing agents such as polyphenols, flavonoids, and uronic acids within the mucilage matrix [63]. The observed increase in reducing capacity is likely due to specific structural features of the polysaccharides, particularly hydroxyl groups that act as electron donors as well as associated phytochemicals like anthocyanins and phenolics [64]. Moreover, monosaccharide compositions and molecular weight may also influence the antioxidant activity of mucilage. The low molecular weight of mucilage is positively associated with its antioxidant activity as it provides greater exposure of hydrophilic (charged) groups on the surface [65]. These groups can readily donate hydrogen atoms and facilitate the reduction of free radicals through both electron-donating and hydrogen transfer mechanisms. Furthermore, galactose-rich mucilage has been shown to exhibit enhanced antioxidant activity. These components contribute synergistically to the antioxidant behavior by facilitating electron transfer and stabilizing reduced iron species. Overall, the results highlight the strong antioxidant potential of Cucumis sativus L. mucilage and support its application in functional foods, nutraceuticals, and packaging systems where oxidative stability is essential.

Analysis of DPPH Free Radical Inhibition Potential

The DPPH assay was carried out to evaluate the free radical scavenging capacity of Cucumis sativus L. mucilage powder across a concentration range of 2–10 mg/mL. As shown in Figure 2B, a steady and concentration-dependent increase in inhibition was observed, with values rising from 50.2% at 2 mg/mL to 67.5% at 10 mg/mL. This progressive enhancement in radical scavenging activity reflects the presence of diverse antioxidant constituents within the mucilage matrix, likely including phenolic acids, flavonoid, and uronic acid, which are known to interact with free radicals through electron transfer pathways [66]. DPPH, a nitrogen-based stable free radical, neutralizes by electron or hydrogen atom acceptance to yield a diamagnetic compound, and compounds with high reducing power can effectively neutralize it by donating electrons or hydrogen atoms [67]. The observed activity in Cucumis sativus L. mucilage indicates that such reactive functional groups are available, possibly due to the polysaccharide structure that permits molecular interactions through hydroxyl and carboxyl groups.

Moreover, the scavenging efficiency observed even at lower concentrations implies that the active molecules are not only present but also highly reactive and accessible, which may be attributed to the low molecular weight fractions or specific conformational features of the mucilage polymers [68]. The IC₅₀ value, determined to be 1.798 mg/mL, highlights the ability of the mucilage to achieve antioxidant action at minimal concentrations. Yu et al. [69] performed a study on polysaccharides derived from bamboo shoot and reported an IC₅₀ value of 7.73 mg/mL. In comparison, Cucumis sativus L. mucilage exhibited a relatively lower IC₅₀ value, indicating its suitability as a highly effective source of antioxidants. These results suggest that Cucumis sativus L. mucilage is a promising source of antioxidant agents and has potential for use as a natural additive in different food systems.

Evaluation of ABTS Radical Scavenging Potential

The ability of Cucumis sativus L. mucilage to scavenge ABTS radicals was investigated over a concentration span of 2–10 mg/mL. The scavenging activity showed a clear positive correlation with increasing concentration (Figure 2C). The inhibition activity of the mucilage increased from 25.30% at 2 mg/mL to 60.14% at 10 mg/mL, showing a steady rise across intermediate concentrations. The observed pattern demonstrates that antioxidant efficacy is dependent on the concentration of the mucilage, likely due to the increased availability of bioactives such as phenolics, flavonoids, or uronic acids at higher concentrations [70]. Moreover, the IC₅₀ parameter denotes the concentration at which 50% of ABTS radical species are effectively scavenged, which was 8.038 mg/mL. The observed results align with the existing literature on polysaccharides from plant sources. In Sukweenadhi et al. [71], extracts from Orthosiphon stamineus were reported to exhibit an IC₅₀ value of 16.91 mg/mL in the ABTS assay, which is significantly higher than that of Cucumis sativus L. mucilage. This comparison highlights the relatively stronger antioxidant activity of Cucumis sativus L. mucilage. The radical scavenging ability can be attributed to the structural features of mucilage, including hydroxyl and carboxyl groups, which play a crucial role in stabilizing ABTS⁺ radicals. Additionally, the increasing trend in ABTS inhibition confirms the functional bioactivity of mucilage, making it ideal for various food applications.

3.4.2. Anti-Inflammatory Activity

Effects of Cucumis sativus L. Mucilage on Cell Viability

The mucilage’s anti-inflammatory properties were examined in an LPS-activated RAW 264.7 macrophage cell model. Prior to assessing its anti-inflammatory effects, cell viability was measured to confirm the safety of the mucilage. As shown in Figure 3A, the mucilage exhibited no cytotoxicity at concentrations ranging from 25 to 500 mg/mL and enhanced macrophage proliferation compared to the control. Specifically, treatment with 25 mg/mL mucilage increased the proliferation index to 1.38 from the control value of 1.08, indicating a 1.28-fold increase in cell viability. However, at higher concentrations, the proliferation index gradually declined to 1.25, 1.24, 1.23, and 1.21 at 50, 100, 200, and 500 mg/mL, respectively. This decrease might be due to changes in the cell culture environment, such as osmotic pressure alterations caused by higher polysaccharide levels, which could influence cell growth [72]. However, none of the tested concentrations exhibited cytotoxic effects, highlighting the mucilage’s safety and its modulatory effect on macrophage proliferation. In conclusion, the mucilage extract demonstrated a concentration-dependent anti-inflammatory activity, with significant reduction in LPS-induced macrophage proliferation, supporting its potential as a natural anti-inflammatory agent.

Effects of Cucumis sativus L. Mucilage on Nitric Oxide (NO) Production

The inhibitory effect of the mucilage on nitric oxide production was studied in LPS-stimulated RAW 264.7 macrophages to determine its anti-inflammatory activity. In the untreated control group, the NO level was found to be 6.2 µmol/L, whereas stimulation with LPS significantly elevated NO production to 19.5 µmol/L, indicating a pronounced inflammatory response. Mucilage-treated groups showed a reduction in NO levels compared to the LPS group, with the most prominent suppression observed at lower concentrations. At 25 and 50 mg/mL, NO production was reduced to 16.7 and 16.5 µmol/L, respectively. As the concentration increased, NO levels exhibited a gradual rise, reaching 17.31, 17.90, and 18.22 µmol/L at 100, 200, and 500 mg/mL, respectively. Although a non-significant increase was noted at higher doses, the NO production remained below the level observed in the LPS-only group across all concentrations (Figure 3B). The observed decrease in NO generation suggests that the mucilage can modulate inflammatory mediator release, particularly at lower concentrations [73]. Since NO plays a critical role in inflammation and oxidative stress, its suppression reflects the anti-inflammatory potential of the mucilage. The partial downregulation of NO without completely blocking its production indicates an immunomodulatory effect, where inflammation is controlled without impairing essential cellular functions [74]. The non-linear NO response at higher mucilage concentrations may result from the saturation of iNOS (inducible nitric oxide synthase) modulating bioactive components, where inhibitory pathways are maximally engaged at lower doses. At higher concentrations, increased polysaccharide content may also induce mild osmotic or viscosity-related effects in the culture medium, influencing macrophage behavior and partially attenuating NO suppression. Such non-linear dose responses are commonly reported for plant-derived polysaccharides and reflect concentration-dependent cellular adaptation rather than loss of bioactivity. This plateau behavior is commonly reported for plant-derived polysaccharides and reflects pathway saturation rather than loss of bioactivity or assay limitations. These results highlight the mucilage’s promise as a natural agent for managing inflammation.

3.4.3. Determination of Tyrosinase-Inhibition Activity

Analysis of Monophenolase Enzyme Inhibition

The ability of Cucumis sativus L. mucilage to inhibit monophenolase activity, a function primarily associated with tyrosinase, was evaluated across a concentration range of 0.5–2 mg/mL. At 0.5 mg/mL, the mucilage exhibited 17.30% inhibition, which increased to 32.50% at 1 mg/mL, 41.30% at 1.5 mg/mL, and reached a maximum of 60.40% at 2 mg/mL. This steady increase in inhibitory activity suggests the presence of bioactive compounds within the mucilage that can effectively suppress monophenolase function (Figure 4A). The observed inhibition may be attributed to the interaction of mucilage constituents such as polyphenols, flavonoids, and uronic acid with the active site of the tyrosinase enzyme. These compounds are known to bind the copper ions present at the catalytic site, thereby blocking substrate access or altering the enzyme conformation [75]. Additionally, polysaccharides with antioxidant properties can scavenge the intermediate free radicals generated during enzymatic oxidation, further enhancing the inhibitory effect. The significant monophenolase inhibition exhibited by Cucumis sativus L. mucilage indicates its potential as a natural tyrosinase inhibitor [76]. This activity may be beneficial not only in reducing hyperpigmentation but also in mitigating oxidative stress-related inflammatory conditions, as tyrosinase activity is often linked to ROS-mediated cellular responses. These findings highlight the potential of Cucumis sativus L. mucilage as a natural inhibitor of enzymatic browning, supporting its application in cosmetic formulations and food preservation strategies.

Analysis of Diphenolase Enzyme Inhibition

Diphenolase activity inhibition was measured for the mucilage at concentrations from 0.5 to 2 mg/mL. As shown in Figure 4B, the diphenolase inhibitory activity increased progressively with increasing mucilage concentration. At 0.5 mg/mL, the inhibition percentage was 32.10%, which significantly rose to 68.50% at 2 mg/mL. This concentration-dependent increase in diphenolase inhibition suggests that the mucilage contains bioactive compounds capable of interfering with the enzyme’s activity. Diphenolase enzymes are involved in the oxidation of diphenols to quinones, which subsequently leads to browning reactions in food and contributes to melanin formation in biological systems [77]. Thus, the inhibition of diphenolase is relevant for applications in cosmetic industries, where controlling pigmentation is desirable. This effect may be linked to the presence of phenolic compounds, flavonoids, or other antioxidant constituents within the mucilage matrix that may bind to the enzyme’s active site or chelate copper ions essential for diphenolase activity [78]. These interactions hinder the enzyme’s ability to catalyze the oxidation reaction effectively. These findings suggest that the mucilage could be effectively utilized in cosmetic formulations aimed at skin whitening and in food industries to prevent enzymatic browning.

3.5. Anti-Browning Effect on the Fresh-Cut Apple Slices

The anti-browning effect of Cucumis sativus L. mucilage on fresh-cut apple slices was evaluated by monitoring L* values and total color difference (ΔE) during 5 days of storage at 4 °C, as shown in Figure 5. All treatments initially showed comparable brightness, but clear differences were observed during storage. The blank samples exhibited the most pronounced browning, with L* values declining to 67.02 and ΔE rising to 9.08 by day 5. Moreover, kojic acid effectively reduced browning, restricting the decrease in L* value to 68.21 and limiting ΔE to 6.55. This clearly indicates a visible color difference, confirming the effectiveness of the positive control and providing a benchmark for evaluating the anti-browning efficacy of the mucilage. However, Cucumis sativus L. mucilage treatments demonstrated a dose-dependent protective effect: 0.5 mg/mL moderately slowed browning with L* at 69.82 and ΔE of 5.51, while 1.0 mg/mL offered better protection (L* 70.56; ΔE 5.03). The highest concentration (2.0 mg/mL) maintained the highest brightness (L* 71.08) and the lowest degree of color change (ΔE 4.54). The observed anti-browning effect of Cucumis sativus L. mucilage is likely attributed to its polyphenolic compounds and antioxidant activity, which may scavenge reactive quinones or chelate copper at the polyphenol oxidase active site, thereby inhibiting enzymatic browning [79]. These findings indicate that Cucumis sativus L. mucilage, particularly at the highest concentration tested, effectively mitigates browning in fresh-cut apples. Similar trends have been reported in previous studies, where mucilage-based coatings, such as flaxseed mucilage incorporated into polyvinyl alcohol films, significantly reduced the total color difference (ΔE) in fresh-cut apple slices during refrigerated storage. The study demonstrated that coatings enriched with mucilage helped maintain color stability over time, slowing the progression of enzymatic browning and preserving the visual quality of the fruit [80]. These findings support the role of natural mucilage as an effective barrier and antioxidant carrier, consistent with the ΔE reduction observed in the Cucumis sativus L. mucilage-coated apple slices.

4. Conclusions

In conclusion, mucilage derived from Cucumis sativus L. using microwave-assisted extraction exhibits remarkable potential as a multifunctional biopolymer. Its favorable structural attributes, including amorphous morphology, uniform particle size, and moderate surface charge, contribute to excellent solubility, dispersibility, and colloidal stability. The presence of key polysaccharide functional groups, supported by thermal and crystallographic analyses, underscores its thermal resilience and suitability for incorporation into various thermally processed formulations. Functionally, Cucumis sativus L. mucilage demonstrates outstanding water and oil holding capacities, foaming ability, emulsifying properties, and solubility, enhancing texture and stability. Biologically, the mucilage exhibits strong antioxidant and anti-inflammatory activities along with effective tyrosinase inhibition, supporting its health-promoting applications without cytotoxic effects. Notably, its application on fresh-cut apple slices demonstrated significant anti-browning potential, highlighting its utility as a natural alternative for food preservation. Collectively, the multifunctional properties of Cucumis sativus L. mucilage underscores its suitability as a sustainable, plant-based ingredient with potential uses in food systems, particularly for extending the shelf-life and quality of fresh-cut produce. Its natural origin, functional versatility, and bioactivity offer a valuable opportunity for the development of clean-label, eco-friendly, and health-promoting products.