Rapid microwave-assisted combined extraction and image-guided purification of Psyllium husk hydrocolloids: Structure, amino acid profiling, and functional properties

Abstract

This study reports an innovative, rapid microwave-assisted combined extraction method integrated with image-guided purification for extracting Psyllium husk mucilage (PHM). The optimized microwave-assisted water extraction (MWE) approach reduced processing time and conserved energy compared to traditional methods, including hot-water extraction (HWE) and cold-water extraction (CWE). MWE significantly improved mucilage yield (50.23 ± 0.32 % wet basis), and spray drying produced PHM yield (39.55 ± 0.63 %) with a particle size of 427.61 nm. Structural characterization revealed various chemical compounds and linkages. Sugar profiling indicated xylose (58.06 g/100 g) and arabinose (14.31 g/100 g) as major sugars, alongside diverse amino acids. Functional evaluation demonstrated PHM's superior viscosity, excellent emulsifying capability, high water holding capacity, and notable foaming properties. Overall, this sustainable and scalable extraction-purification technique enhances the quality and expands the potential applications of Psyllium husk-derived materials, providing a valuable alternative for industry-scale production.

Graphical abstract

Highlights

• •Microwave-assisted water extraction (MWE) facilitated higher purified mucilage.
• •Psyllium husk mucilage (PHM) contains xylose and arabinose as major sugars.
• •Spray-dried mucilage exhibited uniform particles and higher thermal stability.
• •PHM showed excellent anti-microbial activity against food pathogens.

1. Introduction

Mucilage is a complex, high-molecular-weight polysaccharide predominantly derived from various plant sources (Lira et al., 2023). In recent years, plant-derived mucilage has emerged as a multifunctional biopolymer with diverse industrial applications, particularly in the pharmaceutical, food, cosmetic, and biomedical sectors (Shiam et al., 2025; Tsopzong et al., 2024; Van Rooyen et al., 2024). Its unique physicochemical characteristics, including biodegradability, biocompatibility, and emulsifying potential and excellent gelling and stabilizing capabilities, make it a promising alternative to synthetic polymers (Tosif et al., 2021). Owing to its natural origin and broad spectrum of applications, mucilage is in high demand as a sustainable and safe functional ingredient. As industries shift towards eco-friendly and renewable biomaterials, mucilage has garnered substantial attention for use in controlled drug delivery systems, functional food formulations, and biodegradable packaging (Beikzadeh et al., 2020; Cakmak et al., 2023). The increasing need for clean-label and green ingredients in consumer products further underscores the importance of developing efficient and environmentally benign methods for mucilage extraction and utilization (Tosif, Bains, Chawla, Paul, et al., 2025).

Among the different botanical sources of mucilage, Plantago ovata, commonly known as Psyllium, is mainly noteworthy due to the high concentration and unique functionality of mucilage present in its husk. Psyllium husk mucilage (PHM) contains a higher amount of arabinoxylans, which are highly branched heteropolysaccharides with the capability to absorb water and swell to form viscous gels (Waleed et al., 2022a). The high mucilage yield of Psyllium husk and its excellent water-holding and gelling characteristics have become a valuable candidate in both therapeutic and industrial contexts (Neto et al., 2021). PHM is widely used in dietary fiber supplements, functional foods, and pharmaceutical formulations aimed at lowering cholesterol, managing blood glucose levels, and improving gastrointestinal health (El-Maksoud et al., 2021; Tosif et al., 2025). Furthermore, its strong film-forming and hydrogel-forming properties offer exceptional applications in biodegradable packaging and tissue engineering. As a result, PHM is in considerable demand for its multifunctional roles and natural origin, aligning well with the current trends in sustainable material science and nutraceutical innovation (Mehnath et al., 2020).

The extraction of high-purity PHM remains a substantial technical challenge. The highly viscous and gelling nature of PHM, while advantageous in end-use applications, complicates its extraction and purification processes (Neto et al., 2021). PHM produces a strong three-dimensional gelling network in the presence of water and heat. Therefore, it is difficult to remove husk particles from the gel metrics. However, conventional extraction methods, including hot water extraction (HWE) and cold-water extraction (CWE), often result in incomplete separation of mucilage from the fibrous husk due to the formation of dense gels during hydration and heating (Halász et al., 2022). These gels trapped the husk particles, leading to co-extraction and contamination, thus affecting the purity and functionality of the final product. The presence of residual husk not only alters the physicochemical properties of the mucilage but also limits its applicability in formulations requiring high structural homogeneity (Patel et al., 2019). Moreover, these traditional methods are time-consuming, labor-intensive, and inefficient in terms of solvent usage and energy consumption. The need for multiple filtration and centrifugation steps further reduces the overall scalability and economic feasibility of mucilage extraction using such conventional techniques (Strkalj et al., 2025).

A number of studies have attempted to address these challenges through chemical, enzymatic, and mechanical interventions. For instance, ultrasonication-based extraction methods, as investigated by Souza et al. (2020), offered better dispersion and release of mucilage but induced structural damage to the polymer matrix due to prolonged exposure to high-frequency waves. Similarly, enzymatic treatments, while effective in breaking down cell walls, raised concerns over enzyme residue contamination and cost-effectiveness (Van Craeyveld et al., 2009). Collectively, these findings revealed that while existing approaches show some degree of improvement, none have successfully addressed the dual challenge of rapid extraction and high-purity recovery in an environmentally sustainable manner. These challenges underline the demanding need for a novel, green, and rapid extraction method that not only enhances the yield and purity of PHM but also aligns with sustainable processing standards. Thus, microwave-assisted water extraction (MWE) is a potential mucilage extraction method because microwaves deliver volumetric dielectric heating that rapidly couples energy into the water-rich matrices, accelerating cell-wall disruption, solvent penetration, and mass transfer relative to conductive heating (Motlagh et al., 2024; Rezaei Motlagh et al., 2021). Furthermore, compared with conventional or ultrasonic approaches, MWE requires a shorter time, uses less solvent, and can effectively increase the polysaccharide yield along with purity (Bhadange et al., 2024). Mechanistically, the high-intensity radiation of microwave can build the pressure within the solvent and disrupt the cell wall of the husk to release of mucilage (Lozano Pérez et al., 2024).

Despite the increasing number of studies on mucilage extraction from various plants, to date, there is no well-established aqueous-based method that can extract PHM efficiently without significant husk residue. Furthermore, no reported technique incorporates real-time purification guidance to ensure consistency and precision during the extraction process. This lack of a scalable, selective, and eco-friendly extraction process limits the industrial utilization of PHM and restricts its broader application in high-value domains like foods, pharmaceuticals, and biomedical engineering.

To bridge this critical research gap, the present study proposes a pioneering microwave-assisted combined extraction and image-guided purification strategy for Psyllium husk mucilage. In addition to demonstrating the efficiency of this novel extraction method, the study undertakes a comprehensive characterization of the extracted PHM. Furthermore, amino acid profiling is performed to evaluate the nutritional and functional composition of the mucilage. The antimicrobial activity of the mucilage is also assessed, revealing potential biological properties.

2. Materials and methods

2.1. Materials

Psyllium husk was purchased from the Sarvoday Sat Isabgol factory, Sidhpur, Gujarat. Different media were used for microbial growth, such as nutrient agar and Mueller-Hinton Agar (MHA), and bacterial strains (Escherichia coli and Staphylococcus aureus) were procured from Hi-Media Pvt. Ltd. (Mumbai, India). All chemicals and reagents used in this study were of analytical grade and employed without any further purification.

2.2. Methods

2.2.1. Green experimental design

In this study, PHM was extracted using a two-step process to achieve the goal of extracting the purified mucilage in less time. Three different aqueous extraction methods, including hot-water extraction (HWE), cold-water extraction (CWE), and microwave-assisted water extraction (MWE), were employed to extract the mucilage from Psyllium husk (PH). In the first step, the visual husk impurities were assessed, and mucilage yield was optimized. Based on higher mucilage yield (ethanol precipitated on a wet basis) and the absence of husk impurity in mucilage solution, a suitable extraction method was selected. In the second step, PHM extracted using the selected extraction condition was further subjected to a spray-drying process to obtain uniform-sized mucilage powder.

2.2.2. Extraction of PHM using different methods

PHM was extracted using different extraction methods, including HWE, CWE, and MWE, by following the methods of (Fedeniuk & Biliaderis, 1994; Felkai-Haddache et al., 2016; Zhi et al., 2018) with minor modifications. Herein, different temperature-time combinations were used during HWE. Briefly, 2 g of Psyllium husk was dispersed into the 300 mL of distilled water and placed on a heating mantle (EM0250/CE, UK) at 60–100 °C for 30–90 min. For MWE, it was subjected to microwave (Samsung, South Korea) at 360–900 W for 2–8 min. Whereas 2 g of husk was manually added into 3000 mL of cold distilled water and placed in refrigeration (4–7 °C) for 24 h during CWE. Subsequently, after applying specific extraction conditions visual image of the solutions was captured using a mobile phone. Based on images and the visual existence of husk impurities, the most suitable method was identified. For MWE, the upper layer of mucilage was collected, and 80 % ethanol was added to precipitate the mucilage. The extraction overview of PHM is illustrated in Fig. 1. During HWE and CWE, the samples were centrifuged at 10000 ×g for 10 min, and the yield of calculated by adding 80 % ethanol to the supernatant. Based on husk impurities and ethanol-precipitated mucilage yield on a wet basis, a suitable extraction method was subjected to a spray drying process.

Fig. 1.

Schematic illustration of the extraction of PHM using green methods. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

2.3. Spray-drying of PHM

An industrial-scale spray dryer (S.M. Scientech, India) equipped with a rotary atomizer was employed to obtain uniform spray-dried PHM powder. The spray-drying condition was selected based on the research published by our group, according to Sharma et al. (2023). Spray drying was performed with an inlet temperature of 160 °C and an outlet temperature of 70 °C, using an atomizer at 2400 rpm and a feed rate of 7 mL/min. The yield of spray-dried PHM powder was calculated using the following Eq. (1).

$$\text{Yield of Mucilage}\left(\%\right)=\left[\left(\frac{\mathit{Weight\; of\; dried\; mucilage\; powder}\left(g\right)}{\mathit{Total\; weight\; of\; Psyllium\; husk}\left(g\right)}\times 100\right)\right]$$ (1)

2.4. Nutritional and chemical composition of PHM

2.4.1. Proximate analysis

Proximate parameters (moisture, ash, protein, fiber, fat) of Psyllium husk (PH) and Psyllium husk mucilage (PHM) were determined according to the standard protocol of the (AOAC, 2010) method. Total carbohydrate content was determined by the difference method.

2.4.2. Monosaccharide composition

The monosaccharide composition of PHM was determined using high-performance liquid chromatography (HPLC) (Thermo Fisher Scientific, USA) equipped with an aminex HPX-87H column and a refractive index (RI) detector. Briefly, 300 mg of spray-dried PHM was hydrolyzed with 15 mL of 4 % sulfuric acid and subjected to a water bath at boiling temperature (100 °C) for 1 h, followed by neutralization of the sample using CaCO₃. The sample was centrifuged at 8000 rpm for 15 min, and the supernatant was filtered using a 0.3 μL membrane filter. A 10 μL aliquot of the filtrate was injected into the HPLC system under the following conditions: flow rate of 0.6 mL/min, column temperature of 50 °C, and detector temperature of 35 °C. The mobile phase consisted of 0.005 M H₂SO₄ (Saikumar & Badwaik, 2025a).

2.4.3. Amino acid profiling

The amino acid composition of PHM was assessed using high-performance liquid chromatography (HPLC) following the procedure of our previously published study by Tosif, Bains, Chawla, Paul, et al. (2025). Briefly, the sample underwent hydrolysis using 10 mol L⁻¹ HCl with 6 % (w/v) phenol at 110 °C for 24 h. The essential and non-essential amino acids were derivatized using phenyl isothiocyanate (PITC) and then separated on a reverse-phase LUNA C18 column (100 Å, 5 μm, 250 × 4.6 mm). Detection was performed with a UV detector set at 254 nm. Quantification of amino acids was carried out through a multilevel internal calibration, utilizing α-aminobutyric acid as the internal standard.

2.5. Characterization of PHM

2.5.1. Proton nuclear magnetic resonance (¹H NMR)

Proton nuclear magnetic resonance (¹H NMR) spectroscopy was performed to characterize the structural characteristics and proton environment of PHM. Briefly, 15 mg of PHM was dissolved in 10 mL of deuterated water (D₂O), and the solution was transferred to an NMR tube for analysis. The ¹H NMR spectra were recorded using a Bruker Avance III 400 MHz NMR spectrometer (Bruker Avance HD III, Japan). The spectra were acquired at 25 °C with a relaxation delay of 1 s and a total of 64 scans to ensure an adequate signal-to-noise ratio. The chemical shifts (δ) were reported in parts per million (ppm).

2.5.2. Functional groups determination

Fourier transform infrared spectroscopy (FTIR) analysis of PHM was performed to identify the functional groups by following the steps of Sharma et al. (2023). Briefly, 10 mg spray-dried PHM was thoroughly blended with the potassium bromide (KBr) and pressed into a transparent, thin pellet. This pellet was analyzed under an IR beam over a spectral range of 4000 to 400 cm⁻¹, utilizing a scanning resolution of 4 cm⁻¹ by employing an FTIR spectrophotometer (PerkinElmer, Flexar, USA).

2.5.3. Surface morphology and elemental composition determination

The surface microstructure of PHM was studied using a Field Emission Scanning Electron Microscope (FE-SEM), equipped with an energy-dispersive X-ray spectroscopy (EDS) detector (Jeol-JSM-7610F plus). For analysis, 10 mg of spray-dried PHM was mounted onto a carbon-coated adhesive tape and subsequently subjected to gold sputter coating using a JEOL Smart Coater at 40 mA for 3 min. The micrographs were captured at magnifications of 1000× and 2000×, under an accelerating voltage of 15 kV. Moreover, the elemental composition of PHM was analyzed using EDS facilitated by INCA software (Oxford Instruments) integrated with the FE-SEM system.

2.5.4. Particle size and zeta potential

Particle size and zeta potential of PHM were examined for its average particle size distribution and surface charge characteristics using the dynamic light scattering (DLS) technique, conducted with a zeta sizer Nano ZS instrument (Dispersion Technology Inc., USA). Briefly, 10 mg of PHM powder was dissolved in 200 μL of deionized water, followed by centrifugation at 7000 ×g for 10 min. The supernatant was collected and subjected to ultrasonication treatment at 35 °C for 15 min. The processed sample was then analyzed for particle size and zeta potential.

2.5.5. Differential scanning calorimetry (DSC)

Differential scanning calorimetry (DSC) was employed to assess the thermal stability (occurrence of physical and chemical transformations) of PHM following the protocol described by PerkinElmer, Flexar, USA). The analysis was carried out using a Q2000 DSC instrument equipped with a single furnace, nickel‑chromium sample holder, and thermocouple-based temperature sensors. Briefly, 10 mg of PHM was placed in the calorimeter and heated from 28 °C (ambient temperature) to 450 °C at a rate of 10 °C/min under a nitrogen atmosphere (99.99 % purity) to ensure an inert environment during thermal analysis.

2.5.6. Thermogravimetric analysis (TGA)

Weight loss of PHM during the heating process was conducted using a TGA, equipped with an isothermal zone and microbalance (PerkinElmer, Tokyo, Japan). Herein, the sample was heated from 28 °C to 800 °C at a constant heating rate of 10 °C/min under a nitrogen atmosphere, ensuring an inert environment throughout the thermal decomposition.

2.6. Rheological behavior

The rheological behavior of PHM was studied by following the procedure outlined by Tosif, Bains, Chawla, Paul, et al. (2025) with minor modifications. A constant gap of 1000 μm was maintained between the rheometer plates during the measurements. The linear viscoelastic region (LVR) was established at a strain of 0.03 % which was consistently applied across all the tests. Briefly, a 2 % (w/v) suspension of PHM was prepared using double-distilled water and subjected to shear rates ranging from 1 to 100 s⁻¹ at 28 °C to assess its rheological behavior. The resulting flow curve data were analyzed using rheometer software (TRIOS v4.3.0).

2.7. Functional properties

2.7.1. Solubility

The solubility of PHM was determined by dissolving 1 % (w/v) of the sample in deionized water at room temperature (28 °C) under continuous mechanical stirring for 30 min by following the steps of Saikumar and Badwaik (2025b). The mixture was subsequently centrifuged at 7000 ×g for 15 min at 28 °C to separate the insoluble fraction. The resulting supernatant was collected and dried in a hot air oven at 110 °C until a constant weight was achieved. Solubility was then calculated using Eq. (2).

$$\text{Solubility}\left(\%\right)=\left[\left(\frac{\mathit{Weight\; of\; supernatant\; after\; drying}}{\mathit{Initial\; weight\; of\; sample}}\times 100\right)\right]$$ (2)

2.7.2. Water/oil (W/O) holding capacities

The water/oil holding capacities of the PHM sample were determined by dispersing 0.5 g of the sample in 10 mL of distilled water and 10 mL of oil, respectively, in separate tubes. Each sample was stirred to prevent clumping for 15 min. The samples were then centrifuged at 4000 rpm for 10 min, and the supernatant was decanted. The retained water or oil was quantified and expressed as grams of fluid absorbed per gram of sample, calculated using the following Eq. (3).

$$\mathrm{W}/\mathrm{O}\text{holding capacities}\left(\mathrm{g}/\mathrm{g}\right)=\left[\left(\frac{\mathit{Weight\; of}\mathit{wet}\mathit{sample}\left(g\right)-\mathit{weightof}\mathit{dry}\mathit{sample}\left(g\right)}{\mathit{Weight\; of}\mathit{dry}\mathit{sample}\left(g\right)}\times 100\right)\right]$$ (3)

2.7.3. Emulsifying capacity (EC) and emulsifying stability (ES)

EC and ES of the PHM were evaluated following the methodologies described by Sharma et al. (2023). An emulsion was prepared by homogenizing a 0.5 % (w/v) aqueous dispersion of PHM with 10 mL of flaxseed oil using a T25 ultra-Turrax high-speed homogenizer at 8000 rpm for 5 min. The resulting emulsion was then centrifuged at 6000 rpm for 10 min. EC was determined based on the volume of the emulsified layer formed, as calculated using the following Eq. (4).

$$\text{Emulsifying capacity}\left(\%\right)=\left[\left(\frac{\mathit{Initial\; emulsion\; volume}\left(\mathit{mL}\right)}{\mathit{Total\; Volume}\left(\mathit{mL}\right)}\times 100\right)\right]$$ (4)

Following homogenization, the prepared emulsion was heated in a water bath at 80 °C for 30 min and then allowed to cool to room temperature. The cooled emulsion was subsequently centrifuged at 6000 ×g for 10 min. The volume of an emulsified layer was recorded, and ES was calculated by using Eq. (5).

$$\text{Emulsifying stability}\left(\%\right)=\left[\left(\frac{\mathit{Final\; emulsion\; volume}\left(\mathit{mL}\right)}{\mathit{Initial\; emulsion\; Volume}\left(\mathit{mL}\right)}\times 100\right)\right]$$ (5)

2.7.4. Foaming capacity (FC) and foaming stability (FS)

FC and FS of PHM were assessed following the procedure described by Saikumar and Badwaik (2025a). A 0.5 % (w/v) PHM dispersion was subjected to whipping using a high-speed homogenizer at 8000 rpm for 5 min. After 30 s of settling, the volume of foam produced was measured and recorded as FC. Whereas, FS was determined by measuring the volume of foam remaining after 30 min. Both parameters were calculated using Eqs. (6), (7), respectively.

$$\mathit{Foaming\; capacity}\left(\mathrm{FC}\%\right)=\left[\left(\frac{\mathit{Whipped\; volume}\left(\mathit{mL}\right)-\mathit{Volume\; brfore\; whipping}\left(\mathit{mL}\right)}{\mathit{Total\; volume\; before\; whipping}\left(\mathit{mL}\right)}\times 100\right)\right]$$ (6)

$$\mathit{Foaming\; stability}\left(\mathrm{FS}\%\right)=\left[\left(\frac{\mathit{Volume\; after\; resting}\left(\mathit{mL}\right)-\mathit{Volume\; brfore\; whipping}\left(\mathit{mL}\right)}{\mathit{Total\; volume\; before\; whipping}\left(\mathit{mL}\right)}\times 100\right)\right]$$ (7)

2.8. Microbial killing kinetics

The time-dependent killing kinetics of food-pathogenic microbial populations were evaluated by following the protocol described by Bains et al. (2023). Sabouraud dextrose broth and Mueller-Hinton broth were used for inoculating fungal and bacterial strains, respectively. The growth of Escherichia coli and Staphylococcus aureus was monitored after 18, 24, and 48 h of incubation. Whereas Candida albicans was assessed at 48, 72, 96, and 120 h. The antimicrobial potential of PHM was quantified by calculating the logarithmic colony-forming units per milliliter (log CFU/mL) for each microorganism at a specified period.

2.9. Antimicrobial activity

The broth microdilution technique was utilized to assess the in vitro antimicrobial efficacy of the PHM, specifically to determine the minimum inhibitory concentration (MIC) and minimum bactericidal concentration (MBC) against both Gram-positive and Gram-negative bacterial strains. A series of two-fold serial dilutions of the PHM was prepared in sterile Mueller–Hinton broth. Subsequently, each dilution was inoculated with the bacterial strain under investigation. For each test, 50 μL of the diluted PHM sample was dispensed into microplate wells containing Mueller–Hinton broth, followed by the addition of 10 μL of bacterial suspension. The inoculated plates were incubated at 37 °C for 15 h. Distilled water served as the negative control, while streptomycin was used as a positive control to validate the results. The MIC was defined as the lowest concentration of the mucilage that inhibited visible bacterial growth. To determine MBC, aliquots from wells showing no visible growth were plated onto Mueller–Hinton agar, and the absence of bacterial colonies after incubation indicated bactericidal activity.

2.10. Statistical analysis

All the performed experiments were conducted in triplicate independently, and the obtained data are represented as mean values along with their respective standard deviations. Statistical analysis was performed using a one-way analysis of variance (ANOVA) with a confidence interval of 95 % (p < 0.05). The analyses were conducted using IBM SPSS Statistics software (Version 24.0, IBM Corp, Armonk, USA).

3. Results and discussion

3.1. Mucilage yield and visual husk impurity assessment

Mucilage was extracted through three distinct extraction techniques (MWE, HWE, and CWE) by applying different conditions. Among these, MWE is frequently shown to be a potential method for mucilage extraction from various sources due to its shorter time and higher yield recovery (Fernandes et al., 2024). Herein, different microwave power levels of 360 W, 520 W, 720 W, and 900 W were applied for a duration ranging from 2 to 8 min. The principal objective was to extract the purified mucilage with enhanced yield in less time. Findings indicated that when MWE was employed at varied power levels and times, the movement of Psyllium husk particles was influenced by the strong penetration ability of microwave radiation along with diploid movement (Shiehnezhad et al., 2023). After a certain period (6 min), it was observed that all the husk particles had settled at the bottom, while a distinct layer of purified mucilage formed at the top of the solution, as shown in Fig. 2a.

Fig. 2.

Extraction of PHM using different aqueous extraction methods and its visual impurities. (a) Mucilage extracted using MWE, (b) Mucilage extracted using HWE, (c) Mucilage extracted using CWE, (d) Pictorial representation of ethanol precipitated mucilage and spray-dried mucilage powder. (PHM; Psyllium husk mucilage, MWE; microwave-assisted water extraction, CWE; cold-water extraction).

At a lower microwave power of 360 W for 2 min, the husk particles migrated upward, forming a layer with an average height of 1.0 ± 0.04 cm, while the mucilage layer below measured 5.2 ± 0.02 cm (Table 1). Likewise, when the same microwave power was applied for 6 min, the results were reversed. The husk layer was observed at the bottom with a height of 0.4 ± 0.01 cm, and the purified mucilage layer above measured 0.5 ± 0.01 cm. Therefore, within 6 min of microwave treatment, all tested microwave power levels resulted in the effective settling of husk particles, allowing the upper mucilage layer to be clearly separated for yield calculation. However, at 8 min, the browning of mucilage occurred due to caramelization of sugar at a higher intensity of microwave in the presence of water. This discoloration is attributed to the microwave-induced dipole rotation and ion migration within the husk particles (Dorneles & Noreña, 2020). Moreover, the maximum mucilage yield (50.23 ± 0.32 %) was recorded at 900 W for 6 min, as shown in Fig. 3a. While increasing the extraction time from 6 to 8 min yield was significantly decreased up to 44.32 ± 0.87 % on a wet basis. These high-intensity conditions created substantial pressure and temperature within the solvent, which resulted in the breakdown of husk cell walls. The structure of the husk enables water to infiltrate the cells, promoting the release and movement of polysaccharides. Furthermore, the influence of electric and magnetic fields can interfere with hydrogen bonding, thereby improving the solubility of polysaccharides and accelerating the extraction process (Felkai-Haddache et al., 2016). In a similar study, Shiehnezhad et al. (2023) extracted the mucilage from Ocimum basilicum var. album (L.) seeds using MWE by applying different power levels of 360 W, 560 W, and 760 W for durations ranging from 2 to 7 min. The optimal yield was recorded as 17 % at 570 W for 4 min.

Table 1.

Height measurement of different layers of mucilage and husk during the extraction of PHM using MWE.

Power (W)	Time (min)	Upper layer of husk (cm)	Lower layer of husk (cm)	Mucilage layer (cm)
360 W	0	0.8 ± 0.09a	–	5.4 ± 0.02e
	2	1.0 ± 0.04b	–	5.2 ± 0.02cd
	4	–	–	5.0 ± 0.03c
	6	–	0.4 ± 0.01a	4.5 ± 0.01b
	8	–	0.5 ± 0.01b	4.3 ± 0.01a
520 W	0	0.8 ± 0.08a	–	5.5 ± 0.03de
	2	1.0 ± 0.06b	–	5.4 ± 0.02d
	4	–	0.2 ± 0.02a	5.2 ± 0.03c
	6	–	0.6 ± 0.03c	4.7 ± 0.04b
	8	–	0.4 ± 0.02b	4.5 ± 0.01a
720 W	0	0.8 ± 0.04a	–	5.5 ± 0.03e
	2	0.8 ± 0.02b	–	5.4 ± 0.04d
	4	–	0.3 ± 0.04a	5.3 ± 0.01c
	6	–	0.6 ± 0.09b	4.8 ± 0.08ab
	8	–	0.4 ± 0.05ab	4.7 ± 0.06a
900 W	0	0.7 ± 0.03a	–	5.6 ± 0.05e
	2	0.8 ± 0.01b	–	5.5 ± 0.09d
	4	–	0.2 ± 0.04a	5.3 ± 0.04c
	6	–	0.5 ± 0.02c	4.9 ± 0.03b
	8	–	0.3 ± 0.04b	4.7 ± 0.02a

Data are presented as mean ± SD (n = 3). Mean values within the same column that have different lowercase (a-d) represent significantly different values, as determined by analysis of variance (ANOVA) and post hoc tests. (Herein; empty column (−) represents the absence of husk particles).

Fig. 3.

Yield optimization of PHM. (a) MWE, (b) HWE. (PHM; Psyllium husk mucilage, MWE; microwave-assisted water extraction).

During HWE, the PHM was extracted at different temperatures (60 °C, 70 °C, 80 °C, 90 °C) and times (30, 60, 90 min). Results revealed that when PHM was extracted at 60 °C, the yield was significantly increased from 26.61 ± 0.38 % at 30 min to 36.22 ± 0.96 % at 90 min. In contrast, yield was significantly increased at 70 °C from 30.42 ± 0.75 % at 30 min to 41.83 ± 0.44 % for 90 min (Fig. 3b). This trend can be attributed to the degradation of polysaccharides at high temperatures is caused by prolonged heat exposure, which also leads to the browning of the mucilage, as shown in Fig. 2b. Similarly, Patel et al. (2019) extracted the crude polysaccharide from Psyllium seed and husks by applying high temperature (90 °C) for 3 h, and the yield was calculated to be 47.72 % and 64.55 %, respectively. However, they used anion-exchange chromatography to purify the extracted polysaccharide. Additionally, PHM forms a viscous gel when exposed to heat and water, making it difficult to extract a purified mucilage free from husk residues and discoloration. Consequently, Assi et al. (2024) optimized the PHM yield using HWE by applying the different variables. Results showed that the yield was significantly increased with increasing the temperature from 50 to 80 °C for 60 to 120 min. Higher yield (32.36 %) was found at 79.99 °C for 60.02 min. PHM extracted using CWE showed a highly viscous gel, which is attributed to the higher water holding and swelling capacity of the PHM in the presence of water due to the interaction of hydrogen bonding and other hydrophilic compounds (Fig. 2c). Thus, based on the yield on a wet basis and purity of the PHM, we employed the selected extraction condition (MWE: 900 W for 6 min) for further spray drying process. The pictorial representation of ethanol-precipitated mucilage and spray-dried mucilage powder is shown in Fig. 2d. The yield of spray-dried mucilage powder was calculated 39.55 ± 0.63 %. Overall, MWE extracted the purified mucilage in a shorter time with higher mucilage recovery compared to HWE, and CWE.

3.2. Nutritional and chemical composition of PHM

3.2.1. Proximate analysis

The proximate composition of Psyllium husk (PH) and Psyllium husk mucilage (PHM) was conducted, and the results are shown in Fig. 4a. The moisture content of PHM was found to be 5.89 ± 0.76 % and 4.05 ± 0.09 % for PH. This is attributed to the hygroscopic nature of mucilage powder and the drying process-induced retention of bound water. The Protein content of PH and PHM was calculated as 1.15 ± 0.14 % and 0.79 ± 0.36 % respectively. A significant difference was observed in fiber content for PHM (5.68 ± 0.21 %) compared to PH (19.78 ± 0.17 %), which indicated that mucilage primarily consists of soluble polysaccharides with a significant loss of insoluble fibrous material during the extraction and drying process of mucilage. A minor amount of fat content for PH and PHM was found to be (PH: 0.95 ± 0.03 %) (PHM: 0.22 ± 0.07 %), and ash content was (PH: 3.42 ± 0.12 %) (PHM: 3.44 ± 0.21 %), respectively. Total carbohydrate content was slightly reduced in PHM (89.66 ± 0.31 %) compared to PH (90.42 ± 0.17 %), which could be due to the partial loss of some soluble sugars during processing or dilution by residual moisture.

Fig. 4.

Nutritional and chemical composition of PHM. (a) Proximate composition, (b) Monosaccharides composition, (c) Amino acid chromatogram of PHM. (PHM; Psyllium husk mucilage, PH; Psyllium husk).

3.2.2. Monosaccharide composition

The HPLC analysis was conducted using standard monosaccharide references (arabinose, glucose, xylose, galactose, and rhamnose) with the standard chromatogram already reported in our previous study (Tosif et al., 2024). The monosaccharide results revealed that xylose was the predominant sugar (58.06 g/100 g), followed by arabinose (14.31 g/100 g), rhamnose (3.13 g/100 g), galactose (2.75 g/100 g), and glucose (1.33 g/100 g), as shown in Fig. 4b. The higher proportion of xylose and arabinose strongly suggested the existence of an arabinoxylan type of polysaccharide, which is a major characteristic of PHM. Arabinoxylans are composed of a β-(1 → 4)-linked xylose backbone with arabinose side chains, accounting for the dominant xylose and arabinose content observed in PHM (He et al., 2021; Kiszonas et al., 2013). Likewise, Ren et al. (2020) extracted the PH polysaccharide and fractionated it in water at different temperatures (20–100 °C). Study revealed that hetroxylan from the PH with a higher amount of arabinoxylan ratio is challenging to extract by water and requires higher temperatures. The existence of a minor amount of rhamnose, glucose, and galactose indicated the presence of branching or structural heterogeneity within mucilage, effectively due to minor pectic or galactomannan-like regions. These sugars may contribute to the techno-functional properties of mucilage, including film-forming, thickening, and emulsifying properties (de Medeiros et al., 2024).

3.2.3. Amino acid profiling

The amino acid profiling of PHM extracted using MWE revealed a diverse range of essential and non-essential amino acids with varied concentrations as determined through HPLC analysis. The standard chromatogram of amino acids is reported in our previous study by Tosif, Bains, Chawla, Paul, et al. (2025). Results showed the existence of both polar and non-polar amino acids. For instance, serine exhibited the highest relative abundance (15.31 % area), followed by isoleucine (14.70 % area), alanine (13.58 % area), and glycine (12.51 % area). Additionally, valine (11.67 % area) and leucine (10.06 % area) are both branched-chain amino acids (BCAAs), which are also found in higher amounts and may further contribute to muscle repair and metabolic health if incorporated into the food formulations (Aggarwal & Bains, 2022). The prominence of serine and glycine is consistent with strong hydration and extensive hydrogen bonding within the sugar-rich matrix, features linked to the high water-holding capacity and shear-thinning rheology reported for mucilage. A moderate amount of phenylalanine (6.22 % area) and the combined peak of arginine and threonine (4.98 % area) were present, as shown in Fig. 4c and Table 2A. On the other hand, lower amounts of lysine (1.53 % area), glutamic acid (1.32 % area), and histidine (0.51 % area) were present. The presence of essential amino acids in PHM underscores the nutritional significance of mucilage as a functional ingredient.

Table 2A.

Different amino acid compounds, retention time, and peak area of PHM.

Amino acid	Retention time (min)	Peak area	Area (%)
Asparagine	13.373	148,696	1.81
Serine	14.764	1,260,241	15.31
Glutamic acid	15.611	108,877	1.32
Glycine	16.800	1,029,464	12.51
Histidine	17.699	41,970	0.51
Arginine + Threonine	21.469	409,875	4.98
Alanine	22.911	1,117,483	13.58
Proline	25.176	260,838	3.17
Tyrosine	28.795	216,361	2.63
Valine	29.558	960,183	11.67
Lysine	32.171	125,945	1.53
Isoleucine	32.810	1,210,200	14.70
Leucine	33.255	828,034	10.06
Phenylalanine	34.187	511,863	6.22

3.3. Characterization of PHM

3.3.1. Proton nuclear magnetic resonance (¹H NMR)

The ¹H Nuclear magnetic resonance (NMR) spectrum of the PHM displayed the characteristic response of a polysaccharide structure. A broad range of signals was observed between δ 3.0–55 ppm, which is indicative of sugar ring protons (H1-H5), commonly associated with monosaccharide units (Fig. 5a). A strong, sharp peak was observed near δ 4.7 ppm, which can be attributed to the residual water signal in D₂O. The downfield region (δ 5.0–5.5 ppm) included weak anomeric proton signals (H1) of α- and β-configuration, which confirmed the presence of glycosidic linkages in the PHM polymer backbone (Patel et al., 2019). The intense peak observed around δ3.3–3.8 ppm corresponds to methine and methylene protons (H2-H6) of the sugar residues. On the other hand, minor signals between δ 1.0–1.3 ppm can be attributed to the methyl protons, such as rhamnose (a deoxy sugar which contains a methyl group at C-6). Xylose units typically resonate around δ3.2–4.2 ppm, which is in agreement with the results of the monosaccharide composition analysis.

Fig. 5.

Characterization of PHM. (a) ¹H NMR spectrum, (b) Fourier transform infrared spectroscopy (FTIR), (c) Morphological characteristics, (d) Elemental composition, (e) DSC thermogram, (f) Thermogravimetric analysis (TGA), (g) Particle size distribution, (h) Zeta potential.

3.3.2. Functional group determination

The FTIR spectrum of PHM showed characteristic peaks at 3287.68 cm⁻¹, 1362.24 cm⁻¹, 1149.02 cm⁻¹, 1077.76 cm⁻¹, 1019.22 cm⁻¹, 761.81 cm⁻¹, and 575.24 cm⁻¹. The broad peak at 3287.68 cm⁻¹ indicates O-H stretching vibrations, characteristic of hydroxyl groups, suggesting the presence of polysaccharides as shown in Fig. 5b. The peak at 1362.24 cm⁻¹ corresponds to C-H bending vibrations in CH₃ and CH₂ groups, associated with the polysaccharide backbone (Thombare et al., 2023). Peaks at 1149.02 cm⁻¹ and 1077.76 cm⁻¹ are attributed to C-O and C-O-C stretching vibrations, indicating glycosidic linkages within the polysaccharide structure. The peak at 1019.22 cm⁻¹, also associated with C-O stretching vibrations, further confirms the polysaccharide nature of the mucilage. The peak at 761.81 cm⁻¹ corresponds to C-H bending vibrations, while the peak at 575.24 cm⁻¹ may be attributed to skeletal vibrations of the polysaccharide backbone or associated minerals. These findings align with previous studies on plant-derived mucilage, confirming the successful extraction and characterization of PHM (Monge Neto et al., 2017). Subsequently, Sárossy et al. (2013) indicated that FT-IR analysis revealed a characteristic polysaccharide band at 1000–1200 cm⁻¹, in agreement with arabinoxylan references and assignable to skeletal C–O/C–O–C vibrations. Also, extraction and modification methods influenced the functional groups of PHM.

3.3.3. Scanning electron microscopy (SEM) and energy-dispersive X-ray spectroscopy (EDX)

The SEM images of the PHM are shown in Fig. 5c. At a magnification of 1000×, the images revealed that the mucilage particles exhibit a spherical morphology with a smooth surface texture. The particles appear to be relatively uniform in size, with diameters of 10 μm. At higher magnification (2000×), individual mucilage particles exhibited a more detailed surface structure. The smooth and spherical nature of these particles suggests that the spray-drying process was effective in creating a consistent and uniform powder (Tosif et al., 2024). The absence of significant surface irregularities indicates good quality and stability of the dried mucilage particles. These observations are consistent with previous studies, which have reported the formation of spherical particles during the spray drying of polysaccharide-based materials. As reported by Cervantes-Martínez et al. (2014), spray-dried aloe vera mucilage exhibited microstructural homogeneity and particle-level stability, indicating effective conversion to a robust powder form. The EDX spectrum (Fig. 5d) provided the elemental composition of the mucilage particles. The major elements detected include sodium (Na), zinc (Zn), copper (Cu), nickel (Ni), calcium (Ca), and sulfur (S). The presence of these elements is indicative of the intrinsic composition of psyllium husk mucilage, which is known to contain various minerals and trace elements. The weight percentages of the elements are as follows: Na (40 %), Zn (20 %), Cu (15 %), Ni (10 %), Ca (10 %), and S (5 %). The relatively high content of sodium suggests that it is a significant component of the mucilage, which is in agreement with the known chemical composition of Psyllium husk. Comparatively, the presence of trace elements such as Zn, Cu, and Ni is notable. These elements might originate from the natural soil environment where psyllium is grown or could be introduced during the processing stages. Previous research by Pasha et al. (2022) analyzed the elemental composition of date palm mucilage and confirmed the existence of Zn, Ca, Fe, Cu, Na, K, Mn, and Mg. However, studies addressing the elemental analysis of PHM are limited. The consistent particle size and elemental composition make the spray-dried PHM suitable for various applications, including as a dietary fiber supplement, a thickening agent in food, and a bioactive component in pharmaceuticals.

3.3.4. Differential scanning calorimetry (DSC)

Differential Scanning Calorimetry (DSC) analysis of PHM extracted using MWE, the thermogram reveals key thermal transitions indicative of its thermal stability and composition. The analysis showed an endothermic peak corresponding to the loss of bound water within the mucilage matrix, typically observed in the temperature range of 30–100 °C, as shown in Fig. 5e. This transition is attributed to the evaporation of water molecules that are physically trapped or weakly bound within the polysaccharide network of the mucilage powder (Tosif et al., 2024). Furthermore, a prominent endothermic peak observed at higher temperatures, around 200–250 °C, is likely associated with the decomposition of the mucilage's polysaccharide structure, indicating the thermal degradation of the biopolymer backbone. These thermal events are consistent with previously reported studies on plant-derived mucilage, where the initial weight loss is due to moisture evaporation followed by thermal decomposition of carbohydrate polymers at elevated temperatures (Mannai et al., 2023).

3.3.5. Thermogravimetric analysis (TGA)

The thermogravimetric analysis (TGA) data for the mucilage extracted from Psyllium husk reveal important thermal stability characteristics (Fig. 5f). Initial decomposition starts around 30 °C, indicating the presence of moisture, which is typically removed first. A significant weight loss occurs between 200 °C and 400 °C, likely attributed to the degradation of polysaccharides and other organic components, consistent with findings in other studies where plant-based mucilage exhibits similar thermal degradation patterns (Sharma et al., 2023). Furthermore, the weight loss occurring above 400 °C suggested the existence of thermally stable residues, including cellulosic, lignin, or mineral components that remain after the decomposition of organic matter. Likewise, this thermal behavior of PHM aligns with the mucilage extracted from other sources, such as flaxseed, which also revealed the major decomposition within this temperature range due to the breakdown of the polymeric structure (Rashid et al., 2019). The observed thermal stability up to 200 °C implies potential applications in food and pharmaceutical industries where moderate thermal processes are employed. This thermal profile makes CDM a suitable candidate for use as a natural thickener or stabilizer, especially in products subjected to mild heating.

3.3.6. Particle size and zeta potential

The mucilage extracted from Psyllium husk was processed using a spray dryer to obtain a fine mucilage powder. The characterization of this powder involved analyzing the particle size distribution and zeta potential to determine its physical and chemical stability. The particle size distribution analysis, as shown in Fig. 5g, revealed that the mucilage particles had an average size of 427.61 nm. This particle size is indicative of relatively small, uniform particles, which are essential for applications requiring consistent texture and stability. The uniformity in particle size can enhance the stability of dispersions, improve the bioavailability of encapsulated compounds, and ensure consistent performance in end-use applications. The zeta potential analysis, depicted in Fig. 5h, showed a value of −14.15 mV. This moderately negative zeta potential indicated that the mucilage particles possess a sufficient surface charge to provide some level of electrostatic repulsion, which can help prevent aggregation to a certain extent. In colloidal systems, zeta potential is a critical parameter as it reflects the degree of electrostatic repulsion between adjacent, similarly charged particles in dispersion (Mannai et al., 2024; Nadendla et al., 2024).

3.4. Rheological behavior

The rheological analysis of PHM reveals a distinct shear-thinning behavior, where the viscosity decreases from around 600 mPa^·s at low shear rates to below 100 mPa^·s at higher shear rates, eventually stabilizing (Fig. 6a). This characteristic is comparable to other hydrocolloids like xanthan gum and guar gum, which also exhibit similar shear-thinning profiles. Such behavior is advantageous for applications in the food industry as a thickening agent, enhancing texture and stability in products like beverages and soups. Additionally, the high initial viscosity and subsequent shear-thinning nature make PHM suitable for pharmaceutical applications, particularly in controlled drug delivery systems, as well as dietary supplements, owing to its ease of ingestion and digestion benefits. The observed rheological properties can be attributed to the alignment and disentanglement of polymer chains under shear, reducing intermolecular interactions. These findings align with the behavior of many polysaccharide solutions. For example, Ptaszek et al. (2025) studied the effect of different temperatures on the rheological behavior of the PHM. Results proved the drastic changes in the rheological behavior with varying the different temperatures.

Fig. 6.

(a). Rheological behavior of PHM and (b) Time kill kinetics to study the effect of PHM against pathogenic bacteria, (c) Time kill kinetics to study the effect of PHM against fungal strain.

3.5. Techno-functional properties

The techno-functional properties of spray-dried PHM were evaluated, and the results showed its significant potential for diverse food industrial applications. Higher water holding capacity (WHC) was observed for PHM (10.61 ± 0.19 g/g), which may be beneficial in the improvement of textural attributes, juiciness, and shelf-stability of food products (Table 2B). High water-holding capacity (WHC) arises from abundant hydroxyl groups in the polysaccharide matrix and from protein substituents capable of hydrogen bonding. For instance, the proteinaceous part of mucilage materially contributes to hydration behavior (Haruna et al., 2016). Moreover, PHM exhibited a substantial oil holding capacity (OHC) (7.05 ± 0.43 g/g), which suggested its suitability in enhancing mouthfeel, stabilizing fat-rich foods, and potentially reducing oil separation in emulsified food products. Moderate foaming capacity (38.12 ± 0.27 %) and foaming stability (45.94 ± 0.73 %) suggest PHM's possible application in products requiring aeration, though the foam stability might limit its standalone use in highly aerated products. Farahnaky et al. (2010) proved that PHM solutions exhibit weak gel-like properties and elastic behavior, with the storage modulus (G') being consistently higher than the loss modulus (G"), which suggested an ability to trap and hold liquid. Waleed et al. (2022b) indicated that the higher arabinose to xylose ratio in arabinoxylans is associated with the gel-forming capacity, which enhances both WHC and ORC, due to small micropores present on the surface. Moreover, the foaming capacity and foaming stability of PHM were found to be 38.12 ± 0.27 %, and 45.94 ± 0.73 %, respectively. Importantly, PHM demonstrated superior emulsifying properties, with an emulsifying ability of 88.09 ± 0.51 % and emulsifying stability of 93.05 ± 0.14 %, making it particularly effective for stabilizing emulsions, sauces, dressings, and dairy analogs (Tosif, Bains, Goksen, Rehman, et al., 2025). The high solubility value (91.09 ± 0.77 %) further supports its application as a versatile, easily dispersible hydrocolloid, ensuring consistent product quality and performance. Overall, PHM can be potentially useful in diverse industrial applications due to its remarkable techno-functional properties.

Table 2B.

Techno-functional properties of PHM.

Functional properties	Results
Water holding capacity (WHC) (g/g)	10.61 ± 0.19
Oil holding capacity (g/g)	7.05 ± 0.43
Foaming capacity (%)	38.12 ± 0.27
Foaming stability (%)	45.94 ± 0.73
Emulsifying ability (%)	88.09 ± 0.51
Emulsifying stability (%)	93.05 ± 0.14
Solubility (%)	91.09 ± 0.77

3.6. Microbial killing kinetics and antimicrobial activity

PHM showed promising results against both Gram-negative (Escherichia coli) and Gram-positive (Staphylococcus aureus) bacteria. The minimum inhibitory concentrations (MIC) for E. coli and S. aureus were determined to be 3.19 ± 0.04 μL/mL and 2.46 ± 0.07 μL/mL, respectively. Correspondingly, the minimum bactericidal concentrations (MBC) were found to be slightly higher, with values of 3.98 ± 0.06 μL/mL for E. coli and 3.22 ± 0.05 μL/mL for S. aureus (Table 2C). The antimicrobial activity of mucilage is due to the existence of different sugars, which contribute as antimicrobial agents (Luo et al., 2019). The microbial time-kill kinetics study provided further insight into the efficacy of PHM over extended periods. At 18 h, the bacterial population of E. coli and S. aureus decreased to 10.35 ± 0.17 and 8.43 ± 0.11 log CFU/mL, respectively. This reduction continued consistently, reaching values of 9.06 ± 0.19 and 7.91 ± 0.10 log CFU/mL at 24 h, and further decreased to 8.72 ± 0.14 and 7.39 ± 0.21 log CFU/mL at 48 h (Fig. 6b). whereas, against Candida albicans, PHM showed effective results as shown in Fig. 6c. The potential antimicrobial activity of PHM can be due to the existence of acidic sugars, which can chelate divalent cations; chelation is known to destabilize the pathogenic bacteria's outer membranes by increasing the permeability and sensitizing cells (Dantas et al., 2021). Furthermore, PHM facilitated the production of free radicals through the membrane, which increased the cell wall permeability and ultimately caused cell death in bacteria (Soleimani et al., 2023). These observations indicate a sustained antimicrobial activity of PHM over time, particularly noteworthy against S. aureus. The gradual reduction in microbial counts signifies that PHM has potential as a natural antimicrobial agent with effective and prolonged activity, suitable for food preservation or therapeutic applications.

Table 2C.

Total minimum inhibitory concentration (MIC) and minimum bactericidal concentration (MBC) of PHM.

Microorganism	MIC (μL/mL)	MBC (μL/mL)	Streptomycin (μL/mL)
Escherichia coli	3.19 ± 0.04	3.98 ± 0.06	0.83 ± 0.11
Staphylococcus aureus	2.46 ± 0.07	3.22 ± 0.05	0.79 ± 0.09

4. Conclusion

This study presents a pioneering approach to address the long-standing challenges associated with the extraction and purification of PHM, a naturally occurring biopolymer of immense industrial and therapeutic value. Traditional methods, such as hot and cold aqueous extractions, are limited by prolonged processing time, low purity, and the inevitable co-extraction of husk due to the high gelling capacity of PHM. To overcome these drawbacks, the integration of microwave-assisted extraction with image-guided purification offers a rapid, green, and highly selective alternative. The microwave technique enables efficient cellular disruption and mucilage release without compromising structural integrity, while real-time image guidance ensures consistent removal of fibrous contaminants, resulting in high-purity mucilage. The observed antimicrobial activity further broadens its utility as a natural preservative or therapeutic agent. However, the high equipment costs, potential challenges in process scalability, and the risk of localized overheating or sugar caramelization at elevated power levels may affect yield consistency and product quality. Therefore, further optimization and scale-up studies are necessary to ensure the industrial feasibility and economic viability of MWE for large-scale mucilage extraction. Overall, the extraction process aligns with green and sustainable principles, the spray-drying step contributes to notable energy consumption and may offset part of the environmental benefits achieved through eco-friendly extraction. Assessing the energy efficiency, drying yield, and potential integration of low-energy drying alternatives will be essential to establish the complete sustainability profile of the process for large-scale applications of PHM.

CRediT authorship contribution statement

Mansuri M. Tosif: Writing – original draft, Validation, Methodology, Investigation, Data curation, Conceptualization. Aarti Bains: Supervision, Methodology, Investigation, Conceptualization. Sanju Bala Dhull: Visualization, Validation, Supervision, Resources. Pardeep Kumar Sadh: Visualization, Validation, Supervision, Conceptualization. Nemat Ali: Validation, Supervision, Software, Resources. Mohammad Rashid Khan: Supervision, Software, Resources. Abdullah F. AlAsmari: Visualization, Validation, Supervision, Methodology. Nazish Muzaffar: Validation, Supervision, Software, Resources, Methodology. Prince Chawla: Writing – original draft, Visualization, Validation, Supervision, Investigation, Conceptualization.

Declaration of competing interest

Given his role as a Co-Guest Editor for the Food Chemistry: X VSI, Dr. Prince Chawla had no involvement in the peer-review of this article and had no access to information regarding its peer-review process.

Data availability

Data will be made available on request.