Green Biorefinery Approach for Taberau (Phragmites australis) Biomass: Delignification by Coriolopsis caperata and Bioethanol Production via MIL-100(Fe)–Immobilized Saccharomyces cerevisiae

Green Biorefinery Approach for Taberau (Phragmites australis) Biomass: Delignification by Coriolopsis caperata and Bioethanol Production via MIL-100(Fe)–Immobilized Saccharomyces cerevisiae | Research Square window.SnipcartSettings = { analytics: { enabled: false } }; (function() { var accessVector = localStorage.getItem('access_vector') || ''; window.dataLayer = window.dataLayer || []; if (accessVector) { window.dataLayer.push({ user: { profile: { profileInfo: { snid: accessVector } } } }); } })(); (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0],j=d.createElement(s),dl=l!='dataLayer'?'&l='+l:'';j.async=true;j.src='https://www.googletagmanager.com/gtm.js?id='+i+dl;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-K279D39R'); Browse Preprints In Review Journals COVID-19 Preprints AJE Video Bytes Research Tools Research Promotion AJE Professional Editing AJE Rubriq About Preprint Platform In Review Editorial Policies Our Team Advisory Board Help Center Sign In Submit a Preprint Cite Share Download PDF Research Article Green Biorefinery Approach for Taberau (Phragmites australis) Biomass: Delignification by Coriolopsis caperata and Bioethanol Production via MIL-100(Fe)–Immobilized Saccharomyces cerevisiae Titin Apung Atikah, Yanetri Asi, Yuliana Yuliana, Wilson Jefriyanto, and 3 more This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-9512742/v1 This work is licensed under a CC BY 4.0 License Status: Under Review Version 1 posted 4 You are reading this latest preprint version Abstract This study explored bioethanol production from Taberau lignocellulosic biomass through fungal delignification, enzymatic hydrolysis, and fermentation using MIL-100(Fe)-immobilized yeast. The fungal pre-treatment with Coriolopsis caperata enhanced laccase activity, reaching a maximum of 520 U/L at 12 days, and significantly reduced lignin content from 20.87% to 10.92%. This reduction improved cellulose accessibility and increased the availability of fermentable sugars during hydrolysis. Enzymatic hydrolysis results showed that fungal-treated biomass produced higher reducing sugar concentrations compared to untreated and control samples, indicating effective lignin removal. Fermentation was conducted under various conditions, with optimal ethanol production achieved at a yeast dosage of 2% (w/v) and a fermentation time of 4 days, resulting in an ethanol concentration of 11.57 g/L. Characterization using FTIR, XRD, and SEM confirmed successful immobilization of yeast on MIL-100(Fe) without structural degradation. The immobilized system demonstrated improved stability and reusability compared to free yeast. The obtained ethanol concentration is within the typical range for SHF-based lignocellulosic bioethanol production. The integration of fungal pre-treatment with MOF-based immobilization provides a promising and sustainable strategy, with potential for further improvement through process optimization and alternative fermentation approaches. Bioethanol immobilization MIL-100(Fe) Phragmites australis Figures Figure 1 Figure 2 Figure 3 Figure 4 Figure 5 Figure 6 Figure 7 Figure 8 Highlights Current perspective of bioethanol production from Taberau ( Phragmites australis ) Biomass Utilizing Coriolopsis caperata , a ligninolytic fungus from Central Kalimantan, for efficient hydrolysis of lignocellulosic biomass Improved performance of fermentation due to the action of immobilized yeast ( Saccharomises cerevisiae ). MIL-100(Fe) provided a robust platform for yeast immobilization, improving stability, reusability, and catalytic efficiency. 1. Introduction Indonesia's heavy dependence on fossil fuels for energy poses a significant challenge. According to Hasan et al. (2012), fossil energy sources—petroleum (34.37%), coal (29.02%), and natural gas (23.49%)—dominate the nation's energy mix, while renewable energy contributes only about 11.95% [ 1 ]. Meanwhile, global energy demand is projected to rise by 45% by 2030, with an average annual growth rate of 1.6% [ 2 ]. However, fossil fuels are finite resources that will become increasingly scarce [ 3 – 5 ]. In addition to their finite availability, the combustion of fossil fuels contributes to environmental degradation through greenhouse gas emissions, thereby accelerating global warming [ 5 – 7 ]. These pressing challenges underscore the urgent need for sustainable, renewable energy alternatives to secure a cleaner, more resilient energy future. One viable alternative is bioethanol, a renewable energy source with significant potential to transform the energy sector [ 6 , 8 , 9 ]. In particular, lignocellulosic biomass—non-food plant material—emerges as an ideal feedstock for second-generation (G2) bioethanol production [ 9 , 10 ]. This biomass is rich in cellulose and hemicellulose, which can be converted into bioethanol through advanced processes [ 11 , 12 ]. Persson (2018) highlighted that G2 bioethanol offers lower greenhouse gas emissions than conventional fossil fuels [ 13 ], making it a more sustainable and environmentally friendly option. However, scaling up G2 bioethanol production presents several challenges. The current efficiency of feedstock selection, biomass pre-treatment, and fermentation processes still falls short of what is required for widespread commercial viability [ 14 ]. Despite these obstacles, the potential of bioethanol as a sustainable, low-emission energy source has never been more promising. Phragmites australis (locally known as Taberau ), a weed species from the Poaceae family, has emerged as a promising lignocellulosic feedstock for G2 bioethanol production [ 15 , 16 ]. Native to the peat swamp forests of Central Kalimantan, this plant is found in abundance but remains underutilized in the context of renewable energy applications. With its high cellulose and hemicellulose content [ 15 – 17 ], Taberau is considered a promising raw material for G2 bioethanol production using non-edible biomass sources. In addition to its favorable chemical composition, Taberau offers several other beneficial traits, such as rapid growth, high biomass yield, and minimal requirements for water and nutrients [ 16 – 19 ]. These characteristics position Taberau as a sustainable, low-impact biomass resource, aligning with the objectives of advancing biofuel technologies and and the transition toward more efficient and eco-friendly energy systems. However, utilizing lignocellulosic biomass effectively requires pre-treatment to remove lignin and facilitate the breakdown of cellulose and hemicellulose. Several pre-treatment methods have been developed, including base treatment [ 20 ], acid treatment [ 20 , 21 ], organic solvent-based fractionation [ 19 ], and steam explosion [ 15 , 16 ]. These methods often require the use of chemicals or extreme reaction conditions, which can lead to biomass degradation and the formation of enzymatic inhibitors that hinder the saccharification process at later stages [ 12 ]. Consequently, biological methods—particularly those utilizing white rot fungi—are increasingly being explored as an alternative. This method is advantageous because they do not rely on harsh chemicals or extreme conditions, making them more cost-effective and environmentally friendly [ 12 , 22 , 23 ]. Several studies have reported that white rot fungi are capable of lignin degradation by producing ligninolytic enzymes such as laccase, lignin peroxidase, and manganese peroxidase [ 12 , 24 – 26 ]. Roth and Spiess [ 27 ] further emphasized that laccase is particularly effective for detoxifying phenolic fragments in delignified biomass, which can otherwise inhibit cellulolytic enzyme activity during the saccharification process at later stages. Numerous white rot fungi species have been employed for biomass delignification, including Phanerochaete chrysosporium [ 28 ], Pleurotus eryngii, Irpex lacteus [ 29 ], Ganoderma lucidum, Pleurotus ostreatus, Pleurotus pulmonarius, Trametes sp. [ 30 ], and Pleurotus sajor-caju [ 31 ]. The white rot fungus Coriolopsis caperata, isolated from the previous study, has shown considerable laccase enzyme activity [ 32 , 33 ]. Therefore, biomass pre-treatment using C. caperata for the delignification process is considered a promising approach. Furthermore, the Separate Hydrolysis and Fermentation (SHF) method, which decouples the hydrolysis of cellulose from the fermentation stages, has demonstrated significant potential for second-generation (G2) bioethanol production [ 34 , 35 ]. In this process, cellulose is first converted into glucose through hydrolysis, followed by fermentation into ethanol by Saccharomyces cerevisiae (yeast) [ 34 , 35 ]. The separation of these steps in the SHF process enables independent optimization of enzymatic hydrolysis and fermentation conditions, thereby enhancing overall efficiency and reducing the inhibitory impact of ethanol on cellulolytic enzymes [ 36 ]. Moreover, S. cerevisiae can be immobilized onto various support materials, which facilitates cell separation from the fermentation medium, reduces operational costs through cell reuse across multiple fermentation cycles, enhances cell stability, and decreases contamination risks and fermentation time [ 37 , 38 ]. Common support materials for S. cerevisiae immobilization include sodium alginate [ 38 , 39 ], calcium alginate [ 40 ], and CMC-g-PVP [ 41 ]. Recently, however, metal-organic frameworks (MOFs) have recently been recognized as promising platforms for the immobilization of S. cerevisiae [ 42 , 43 ] and Escherichia coli [ 44 , 45 ]. Among these MOFs, MIL-100(Fe) has attracted considerable attention due to its regular crystal structure, excellent stability, high surface area, non-toxicity, and biodegradability, making it an ideal candidate for yeast immobilization [ 46 , 47 ]. Despite these advances, the optimization of the SHF method using Taberau biomass pre-treated with Coriolopsis caperata and MIL-100(Fe)-immobilized S. cerevisiae for bioethanol production has not yet been thoroughly investigated. The present research aims to bridge this gap by evaluating Taberau as a sustainable raw material for bioethanol production, enhanced by innovative pre-treatment and fermentation approaches. 2. Materials and Methods 2.1. Materials The materials used in this study were iron (II) sulfate heptahydrate (FeSO 4 .7H 2 O), trimesic acid (H 3 BTC), sodium hydroxide (NaOH), sodium acetate (CH 3 COONa), acetic acid (CH 3 COONa), veratryl alcohol ((CH 3 O) 2 C 6 H 3 CH 2 OH), syringaldazine ([HOC 6 H 2 (OCH 3 ) 2 CH = N-] 2 ), which were supplied by Merck. D-glucose (C 6 H 12 O 6 ), peptone, potassium dihydrogen phosphate (KH 2 PO 4 ), dipotassium hydrogen phosphate (K 2 HPO 4 ), zinc sulfate (ZnSO 4 ), iron(II) sulfate (FeSO 4 ), manganese (II) sulfate (MnSO 4 ), and magnesium sulfate (MgSO4) was used to prepare a glucose-peptone medium, while yeast extract, peptone, D-glucose (C 6 H 12 O 6 ), agar was used to prepare yeast extract peptone glucose (YPG) agar medium. 3,5-Dinitrosalicylic acid (DNS) reagent was used for estimating reducing sugar concentration. Distilled water was used as the solvent in all experiments. 2.2. Instrumentation Structural analysis of samples was conducted using XRD (Philips PW1140/90). Finely ground and dried samples were mounted on the sample stage and analyzed over a 2θ range of 5–60° employing Cu-Kα radiation to assess crystallinity. Functional group characterization was carried out using FTIR (Shimadzu FTIR-8400S) with the KBr pellet method, covering a spectral range of 400–4000 cm⁻¹. In addition, the surface morphology of the materials was examined using SEM equipped with an Advanced Microanalysis system. 2.3. Feedstocks Taberau stems was harvested from the countryside of Lais Lake, Tanjung Sangalang Village, Central Kahayan District, Pulang Pisau Regency, Central Kalimantan, Indonesia. The harvested Taberau was air-dried for 2 days on the field, and ground until they passed a 60-mesh sieve. The sieved Taberau was subsequently dried until it reached a moisture content of 10%. 2.4. Organisms and culture maintenances C. caperata was pre-cultured on potato dextrose agar (PDA, 39 g/L) plates at 27 ± 2°C for 7 days in the dark. Meanwhile, S. cerevisiae (yeast) cells maintained on YPG agar medium [ 39 ], was obtained from the available stock culture of Institut Pertanian Bogor Culture Collection (IPBCC), Bogor Indonesia. Slant media were incubated at 30°C over a period of 1–2 days to promote maximal growth. 2.5. Fungal inoculation Ten agar plugs (5 mm in diameter) from the plate culture were aseptically introduced into 100 mL of glucose–peptone medium in a 250 mL Erlenmeyer flask to obtain the liquid culture. The glucose-peptone medium was prepared as previously reported by Agnestisia et al. [ 32 ]. The cultures were maintained at 27 ± 2°C without agitation in dark conditions for 16 days, with 1 mL of veratryl alcohol (100 mM) added after 2 days to promote laccase production. Liquid samples were collected every two days and blended to obtain a homogeneous inoculation. The laccase enzyme activity was then measured. 2.6. Fungal pre-treatment Fungal pre-treatment assays were performed in triplicate using 250 mL Erlenmeyer flasks. Ten grams of Taberua biomass were conditioned with a phosphate buffer solution (pH 5) to achieve an initial moisture content of 75%. Moistened substrates underwent sterilization at 121°C for 30 minutes. After cooling, each flask received 5 mL of a homogeneous liquid culture with the highest laccase activity and was subsequently incubated without agitation at 27 ± 2°C for 35 days. The flasks were sealed with sterile, non-absorbent cotton to allow air exchange between the atmosphere and the culture. A control, consisting of 2 mL of acetate buffer solution instead of fungal inoculation, was also included for each biomass sample. The biomass was then washed after fungal pre-treatment. The contents of cellulose, hemicellulose-associated carbohydrates, and lignin in raw, control, and fungal-treated feedstocks were quantified via a gravimetric method. 2.7. Enzymatic hydrolysis Five grams of cellulase (CAS 9012-54-8, with activity of 20,000 U/g) were added to 500 mL of a 1% (w/v) substrate (raw, control, and fungal-treated feedstocks, respectively) in acetate buffer (pH = 4.8, 50 mM) and stirred at 50°C for 10, 30, 50, 70, and 90 min. The hydrolysis solution was then used to estimate reducing sugar concentration. 2.8. Preparation of yeast cell suspension Ten milliliters of sterile distilled water were added to a 24-hour-old slant culture of S. cerevisiae , after which the cells were aseptically scraped using a sterile inoculating needle and gently agitated to obtain a uniform suspension. Subsequently, 100 mL of YPG medium (agar-free) were dispensed into 250 mL Erlenmeyer flasks, plugged with cotton, sterilized, and cooled to room temperature. Each flask were inoculated with 1 mL of cell suspension under aseptic conditions and incubated at 30°C with shaking at 150 rpm for 24 h. Cells were harvested by centrifugation (6000 rpm, 15 min), washed with saline, and re-centrifuged for an additional 5 min. The supernatant was removed, and the pellet was washed with saline solution, followed by an additional centrifugation step for 5 minutes. The final pellet was subsequently washed, air-dried, and weighed, and these cells were designated as free cells. 2.9. Immobilization of yeast cells MIL-100(Fe) was synthesized in situ at room temperature by mixing an aqueous FeSO₄·7H₂O solution (3.8 g, 13.7 mmol in 120 mL) with an alkaline H₃BTC solution (1.9 g, 9.1 mmol in 30 mL of 1 M NaOH) under stirring (200 rpm). The precursor solution was added dropwise, and the mixture was stirred for 24 h. The resulting solid was washed with warm water and methanol, then dried at 60°C for 12 h. For immobilization, free yeast cells (0.25 g) were combined with MIL-100(Fe) (0.25 g) in acetate buffer (125 mL, 50 mM, pH 4.8). The immobilized cells were recovered by centrifugation (6000 rpm, 15 min), washed three times, air-dried for 24 h, and denoted as MIL-100(Fe)@yeast. 2.10. Ethanol fermentation The ethanol fermentation of the Taberau hydrolysis solution in this study was conducted using a batch fermentation method. The effects of yeast dosage and fermentation time on ethanol production were investigated. MIL-100(Fe)@yeast and yeast cell free (2% w/v) were inoculated into 500 mL Erlenmeyer flasks containing 100 mL of sterilized liquid medium (pH 5), consisting of 0.15% yeast extract, 0.25% NH₄Cl, 0.55% K₂HPO₄, and 0.025% MgSO₄·7H₂O, along with hydrolysis solution as the carbon source. The MIL-100(Fe)@yeast and yeast cell free systems were incubated at 30°C with agitation at 75 rpm for 4 hours. To assess the impact of yeast dosage and fermentation time, experiments were carried out with different MIL-100(Fe)@yeast and yeast cell free concentrations (0.5%, 1%, 1.5%, 2%, and 2.5% w/v) and fermentation durations (1 to 10 days). After the fermentation process, the MIL-100(Fe)@yeast and yeast cell free cells were separated from the fermented medium and stored for use in subsequent experiments. The fermented medium was then analyzed for ethanol and residual reducing sugar concentration. 2.11. Reusability of immobilized yeast cells The reusability of MIL-100(Fe)@yeast was tested by performing multiple fermentation cycles. After each batch, the MIL-100(Fe)@yeast were recovered from the fermentation medium by filtration or centrifugation. The recovered cells were washed with sterile water to remove residual fermentation by-products and medium components. The washed MIL-100(Fe)@yeast were then reused in subsequent fermentation cycles. The reuse experiments were carried out for up to six consecutive batches. In each cycle, the ethanol yield and glucose consumption were monitored, and the same procedure (as described above) was followed for each batch. Samples were drawn at regular intervals (every 24 hours) to measure ethanol content and sugar consumption. 2.12. Analytical methods Laccase activity was assayed via syringaldazine oxidation [ 48 ] in a 3 mL system containing sodium tartrate buffer (0.1 M, pH 5.3), syringaldazine (0.5 mM), and enzyme solution. The increase in absorbance at 525 nm (ε525 = 65,000 M⁻¹·cm⁻¹) was monitored for 2 min using a UV–Vis spectrophotometer. Furthermore, the concentration of reducing sugars was analyzed by the DNS method [ 49 ], with absorbance recorded at 540 nm using a UV–Vis spectrophotometer and quantified against a glucose standard curve. Ethanol content was determined by acid dichromate oxidation, where ethanol in a known sample mass was oxidized to acetic acid using 0.1 N potassium dichromate in the presence of sulfuric acid. In addition, the physical properties of distilled ethanol from this study consisting of density, viscosity, specific gravity, API specific gravity, and flash point, were also determined. Specific gravity was determined using the ASTM D 792 − 13 method. Viscosity was analyzed using a Saybolt viscosimeter with the ASTM D 88 test standard. Flash point were determined using a flash and fire point tester with the ASTM D 92 test standard. Specific gravity and API gravity were analyzed using the ASTM D 891-09 and ASTM D 287 test standards, respectively. 3. Results 3.1. Fungal laccase enzymes The fungal inoculation method used in this study successfully facilitated the production of laccase enzymes, with a particular focus on examining the effects of incubation period on laccase activity generated by C. caperata (Fig. 1 ). Determining the optimal incubation time is crucial for maximizing laccase production in the shortest time possible. To achieve this, incubation times were varied across a range of 2, 4, 6, 8, 10, 12, 14, and 16 days. This range was selected to assess early metabolic activity and temporal fungal performance, enabling evaluation of laccase production stability and nutrient-related effects. Similar incubation timeframes have been employed in previous studies on laccase production in other fungi, such as Ganoderma sp. [ 50 ], Trametes versicolor [ 51 ], and Trametes trogii [ 52 ]. The data obtained from these varied incubation periods will contribute to optimizing laccase production and provide a better understanding of its efficiency over time. The C. caperata fungal culture appeared to show stable growth, and the addition of 1 mL of 100 mM veratryl alcohol after two days of incubation significantly enhanced laccase production. The enzyme activity increased steadily after the second day, reaching an optimum at 12 days with an enzyme activity of 520 U/L, and then declined towards the end of the incubation period, likely due to nutrient depletion or a reduction in fungal metabolic activity. The laccase enzyme activity of C. caperata appeared to be lower than that of C. caperata BM-172 (880 U/L) [ 53 ], but is higher compared to other white rot fungi, such as Fuscoporia gilva (440 U/L) and Pleurotus tuberregium (480 U/L) [ 23 ]. Therefore, C. caperata is considered to have the potential to be used as a delignification agent for Taberua biomass in this study. 3.2. Delignification of Taberau biomass Fungal pre-treatment assays conducted on Taberua biomass showed significant delignification, as indicated by the reduction in lignin content following fungal inoculation (Table 1 ). A control group, using acetate buffer instead of the fungal inoculum, was included to isolate the effect of the fungal treatment. The most notable observation in this study was the significant reduction in lignin content in the fungal-treated feedstock (10.92% ± 1.15%), compared to the raw feedstock (20.87% ± 0.82%) and the control feedstock (19.35% ± 0.74%). This reduction in lignin content is consistent with the known role of laccase-producing fungi in delignification processes. Laccases, a group of enzymes produced by certain fungi, have been widely recognized for their ability to oxidize lignin, breaking down its complex aromatic structure into smaller fragments. The marked decrease in lignin content following fungal treatment provides strong evidence that fungal inoculation, particularly through laccase activity, was the primary mechanism responsible for the observed delignification. This delignification process also enhanced the availability of cellulose and holocellulose -associated carbohydrates, making the biomass more accessible for further biochemical conversion processes such as enzymatic hydrolysis [ 54 ]. These findings align with previous studies in which fungal treatments using laccase-producing fungi facilitated lignin degradation and improved the accessibility of polysaccharides [ 29 , 30 , 54 , 55 ]. Table 1 Chemical composition of feedstocks Composition (%) Raw feedstock Control feedstock Fungal-treated feedstock \(\:\alpha\:\) -Cellulose 39.24 \(\:\pm\:\) 0.47 38.24 \(\:\pm\:\) 1.67 45.21 \(\:\pm\:\) 1.04 Holocellulose 56.68 \(\:\pm\:\) 0.75 59.20 \(\:\pm\:\) 0.85 60.68 \(\:\pm\:\) 0.52 Lignin 20.87 \(\:\pm\:\) 0.82 19.35 \(\:\pm\:\) 0.74 10.92 \(\:\pm\:\) 1.15 Furthermore, the control group in this study, which used acetate buffer instead of fungal inoculum, did not exhibit significant changes in biomass composition, reinforcing the notion that laccase activity was the key driver behind the observed delignification. This finding is consistent with prior research, which also reported no significant changes in biomass composition in the absence of fungal inoculation, confirming the effectiveness of laccase-producing fungi in biomass pre-treatment [ 54 ]. By degrading lignin, the fungal treatment enhances the overall porosity and accessibility of the biomass, thus improving the efficiency of downstream processes like enzymatic hydrolysis, which is crucial for converting the polysaccharides into fermentable sugars for bioethanol production. 3.3. Enzymatic hydrolysis Enzymatic hydrolysis was performed to assess the sugar release potential from raw, control, and fungal-treated feedstock. The hydrolysis reaction was carried out at 50°C for various time intervals (10, 30, 50, 70, and 90 minutes). The results presented in Fig. 2 demonstrate that fungal pre-treatment significantly enhances the enzymatic hydrolysis of Taberua feedstock, as evidenced by the higher sugar release from fungal-treated feedstock compared to raw and control feedstocks. The fungal pre-treatment, which effectively degraded lignin, made the cellulose more accessible to cellulase enzymes, leading to a marked increase in sugar yield after 50 minute of hydrolysis. This finding aligns with several other studies that have shown similar improvements in sugar release following lignin removal through fungal inoculation. For instance, studies by Gupta et al. (2011) demonstrated that laccase-producing fungi can efficiently break down lignin, enhancing the accessibility of cellulose and improving the efficiency of enzymatic hydrolysis [ 56 ]. In contrast, the raw and control samples, which retained most of their lignin, exhibited slower sugar release, particularly at earlier time points, confirming the protective role of lignin in impeding cellulase action. This observation is consistent with several researcher that reported lignin removal is crucial for optimizing biomass conversion processes [ 12 , 24 , 56 ]. Overall, the results emphasize the effectiveness of fungal pre-treatment in improving the efficiency of biomass hydrolysis, offering a promising strategy for enhancing biofuel production and other biotechnological applications. 3.4. MIL-100(Fe) Immobilized yeast cells The synthesis of MIL-100(Fe) and the immobilization of yeast cells onto this material were performed. MIL-100(Fe) was prepared via an in situ method at room temperature [ 57 ], after which the obtained material was utilized as a support for yeast immobilization, yielding MIL-100(Fe)@yeast. Both MIL-100(Fe) and MIL-100(Fe)@yeast were subsequently characterized using FTIR, XRD, and SEM to confirm the successful synthesis and immobilization processes. FTIR analysis was conducted to identify functional groups and to verify the presence of characteristic MOF bonds as well as potential interactions between yeast cells and the MIL-100(Fe) framework. The immobilization of yeast cells onto the material were successfully carried out using an in situ method at room temperature. This method enables the formation of a stable MOF structure while providing a suitable surface for biological immobilization. MIL-100(Fe) is widely reported as a robust MOF with high surface area and excellent chemical stability, making it highly suitable for hosting biomolecules and microbial cells [ 42 , 43 , 58 ]. The FTIR spectra of MIL-100(Fe) and MIL-100(Fe)@yeast, recorded in the range of 400–4000 cm⁻¹, exhibit very similar spectral patterns (Fig. 3 ). This indicates that the immobilization process does not significantly affect the structural integrity of the MIL-100(Fe) framework. Characteristic absorption bands such as O–H stretching (~ 3400–3500 cm⁻¹), C = O stretching (~ 1625–1635 cm⁻¹), and Fe–O vibrations (~ 600–630 cm⁻¹) remain clearly visible after immobilization. These findings are consistent with previous reports showing that MIL-100(Fe) maintains its structural framework even after chemical modification or interaction with external species [ 59 , 60 ]. The preservation of these functional groups suggests that the coordination between Fe³⁺ ions and the organic ligands remains intact. According to Han et al. (2017), the stability of MIL-100(Fe) originates from its strong metal–ligand coordination and mesoporous architecture, which provides resistance against structural collapse during post-synthetic modification [ 60 ]. This characteristic is crucial for applications involving biological systems, where mild conditions are typically required. In addition to the characteristic peaks of MIL-100(Fe), the FTIR spectrum of MIL-100(Fe)@yeast shows additional absorption bands that can be attributed to yeast cells in Fig. 3 . The absorption peak at approximately 2926 cm⁻¹ corresponds to C–H stretching vibrations of aliphatic groups, while the band at around 1572 cm⁻¹ is associated with amide groups from proteins. These functional groups are typical components of microbial cells, including proteins and lipids, confirming the successful immobilization of yeast onto the MOF surface [ 61 ]. Furthermore, the presence of a band near 1047 cm⁻¹, corresponding to C–O stretching vibrations, indicates the presence of carbohydrates such as polysaccharides in the yeast cell wall. Yeast cell walls are primarily composed of glucans, mannoproteins, and chitin, which contribute to these characteristic FTIR signals [ 61 ]. The detection of these functional groups suggests that the yeast cells maintain their structural integrity and biochemical composition after immobilization. The coexistence of MIL-100(Fe) characteristic peaks and yeast-related bands indicates that the immobilization process occurs mainly through surface interactions, such as hydrogen bonding and electrostatic interactions, rather than through structural disruption. Similar observations have been reported in previous studies, where MOFs served as effective supports for enzyme and microbial immobilization without compromising their activity [ 42 , 43 , 58 , 62 ]. To further strengthen the identification results obtained from FTIR for MIL-100(Fe) and MIL-100(Fe)@yeast, additional supporting data in the form of XRD analysis is necessary. This data is expected to determine the crystallinity of the MIL-100(Fe) material. It is important to ensure that MIL-100(Fe) was successfully synthesized and retained its structural integrity after the yeast cell immobilization process. The XRD patterns would also reveal any changes in crystallinity due to the incorporation of yeast cells into the framework. The diffraction peaks observed in the XRD patterns of MIL-100(Fe) and MIL-100(Fe)@yeast correspond to the characteristic crystal lattice planes of the analyzed materials (Fig. 4 ). According to the JCPDS, distinct peaks at 2θ values of 6.08°, 10.22°, and 10.98° are indicative of MIL-100(Fe). These reflections confirm the successful formation of MIL-100(Fe) and the retention of its crystallinity throughout the synthesis process. A comparison of the diffractogram of MIL-100(Fe) with that of MIL-100(Fe)@yeast reveals that the immobilization of yeast cells on the MIL-100(Fe) surface does not significantly alter the overall crystalline structure of the material. The XRD pattern of MIL-100(Fe)@yeast exhibits almost identical diffraction peaks to those of MIL-100(Fe), suggesting that the framework of the MOF remains intact after the immobilization process. However, a notable difference is the appearance of a new peak at 2θ = 20.63°, which is characteristic of yeast. This additional peak is attributed to the specific diffraction pattern of the yeast cells, indicating that they have been successfully incorporated into the MIL-100(Fe) structure. The appearance of this new peak supports the conclusion that yeast cells are effectively immobilized on the MIL-100(Fe) surface, without compromising the material’s structural integrity. The persistence of the characteristic MIL-100(Fe) peaks along with the emergence of the yeast-specific peak further validates the stability and robustness of the immobilization process. These findings are consistent with the FTIR results, which also indicated that the yeast cells maintain their functional characteristics after immobilization. As the results, the XRD analysis complements the FTIR data and provides valuable confirmation that the immobilization of yeast cells on MIL-100(Fe) does not disrupt the material’s crystalline structure. This dual validation from both FTIR and XRD techniques reinforces the potential of MIL-100(Fe) as a stable and effective platform for future biocatalytic applications, where the integrity of both the support material and the immobilized cells is crucial. SEM analysis was further applied to examine the cross-sectional morphology and the distribution of synthesized MIL-100(Fe) (Fig. 5 ). The synthesized MIL-100(Fe) exhibited an octahedral particle shape, with crystallite sizes ranging from 100 nm to 2 µm, consistent with the sizes reported in the literature. However, the size and shape of the MIL-100(Fe) crystals were not uniform, which is typical for materials synthesized at room temperature. This variation in particle size is acceptable, as the synthesis was carried out under mild conditions. In contrast, when high-temperature synthesis methods are used, more homogeneous MIL-100(Fe) crystals can be produced due to better control over nucleation and crystal growth, resulting in more uniform particle size and shape. Furthermore, SEM analysis of MIL-100(Fe)@yeast provides additional insight into the immobilization process. The micrographs show a clear morphological difference between MIL-100(Fe) and MIL-100(Fe)@yeast. The pristine MIL-100(Fe) exhibits irregular agglomerated particles, which is typical for this class of MOFs due to their high surface energy and porous structure. After immobilization, the MIL-100(Fe)@yeast sample shows increased aggregation and the formation of larger clustered structures. This change in morphology indicates the interaction between the MOF particles and the biological component. However, the distinct morphology of yeast cells, which are typically oval-shaped in Saccharomyces cerevisiae , is not clearly observed in the SEM images. This may be attributed to the coverage or partial encapsulation of yeast cells by MIL-100(Fe) particles, which can obscure their natural morphology. Such phenomena have been widely reported, as microorganisms or enzymes immobilized within porous supports or MOFs often become difficult to visually distinguish due to tight coating, encapsulation within pore structures, or strong host–guest interfacial interactions that obscure clear morphological boundaries between the biological entities and the supporting material [ 43 , 63 ]. In addition, the cell wall of Saccharomyces cerevisiae , which is rich in polysaccharides and proteins, can promote adhesion to solid supports through hydrogen bonding and electrostatic interactions, further facilitating the immobilization process [ 61 , 64 ]. Importantly, although individual yeast cells are not distinctly visible, the observed increase in particle size and aggregation strongly suggests successful immobilization. These SEM results are consistent with the FTIR and XRD analyses, which confirm the presence of biological functional groups and the preservation of the MIL-100(Fe) crystalline structure. The combination of these characterization techniques provides complementary evidence that yeast cells have been successfully immobilized onto the MIL-100(Fe) framework without structural degradation. 3.5. Ethanol fermentation The ethanol fermentation of Taberau hydrolysate using both MIL-100(Fe)@yeast and free yeast systems was evaluated at different yeast dosages. The results clearly demonstrate that yeast dosage significantly affects both sugar consumption and ethanol production (Fig. 6 ). For both systems, increasing yeast dosage from 0.5% to 2.0% (w/v) resulted in a marked increase in sugar consumption and ethanol production. In the MIL-100(Fe)@yeast system, sugar consumption increased from 16.07 g/L to 27.63 g/L, accompanied by an increase in ethanol concentration from 6.90 g/L to 11.30 g/L. A similar trend was observed in the free yeast system, although at lower levels, where sugar consumption increased from 14.20 g/L to 24.80 g/L, and ethanol production from 6.00 g/L to 10.17 g/L. The maximum ethanol production in both systems was achieved at a yeast dosage of 2%, suggesting an optimal balance between cell concentration and substrate availability. This observation is consistent with previous reports indicating that appropriate yeast loading enhances substrate utilization and ethanol productivity, whereas excessive biomass does not necessarily lead to higher yields [ 65 ]. Furthermore, the MIL-100(Fe)@yeast system consistently exhibited higher sugar consumption and ethanol production compared to free yeast. This can be attributed to the advantages of cell immobilization, which include improved cell stability, higher cell density, and increased resistance to inhibitory compounds. Immobilized yeast systems have been reported to achieve higher ethanol yields and maintain stable fermentation performance [ 38 , 64 , 66 , 67 ]. However, further increasing the yeast dosage to 2.5% did not result in additional improvement. In both systems, sugar consumption decreased slightly, and ethanol production either plateaued or decreased. This indicates the existence of a threshold yeast concentration beyond which fermentation efficiency no longer improves. The decline at higher yeast loading can be attributed to cell overcrowding, competition for nutrients, and accumulation of inhibitory metabolites [ 68 ]. Fermentation time also plays a critical role in determining ethanol production efficiency. In this study, both sugar consumption and ethanol production increased significantly from day 1 to day 4 in both MIL-100(Fe)@yeast and free yeast systems (Fig. 7 ). In the MIL-100(Fe)@yeast system, sugar consumption increased from 16.53 g/L (day 1) to 27.77 g/L (day 4), accompanied by a rise in ethanol concentration from 5.60 g/L to 11.57 g/L. A similar trend was observed for free yeast, although at lower levels, where sugar consumption increased from 13.37 g/L to 24.40 g/L, and ethanol production from 3.50 g/L to 9.83 g/L over the same period. These results indicate that the active fermentation phase occurred within the first four days, during which yeast cells efficiently converted fermentable sugars into ethanol. This pattern is consistent with previous studies reporting that ethanol production typically increases during the exponential growth phase and reaches a maximum before entering the stationary phase [ 65 ]. The highest ethanol production for both systems was observed on day 4, confirming that this duration represents the optimum fermentation time. After day 4, both ethanol production and sugar consumption showed a gradual decline. In the MIL-100(Fe)@yeast system, ethanol decreased from 11.57 g/L (day 4) to 9.63 g/L (day 10), while in the free yeast system it decreased from 9.83 g/L to 5.83 g/L. This decline suggests that most fermentable sugars had been consumed and that inhibitory effects, particularly from accumulated ethanol and metabolic by-products, began to negatively affect yeast activity. Similar observations have been reported in ethanol fermentation studies, where product inhibition and substrate depletion limit further ethanol production [ 68 ]. Compared to free yeast, the MIL-100(Fe)@yeast system consistently exhibited higher sugar utilization and ethanol production across all fermentation times. Additionally, the decline after the optimum point was less pronounced in the immobilized system, indicating better stability and resistance to inhibitory conditions. This behavior is characteristic of immobilized yeast systems, where the supporting matrix enhances cell protection, maintains higher cell density, and prolongs metabolic activity [ 38 , 64 , 66 , 67 ]. Furthermore, immobilization has been widely reported to improve ethanol productivity and operational stability in lignocellulosic fermentation systems. These findings confirm that a fermentation duration of 4 days is optimal under the studied conditions. Extending fermentation beyond this point does not improve ethanol yield and may reduce efficiency due to substrate depletion and ethanol inhibition, as widely discussed in bioethanol production literature [ 38 , 64 , 66 , 67 ]. 3.6. Reusability of immobilized yeast cells The reusability of MIL-100(Fe)@yeast over six consecutive fermentation cycles showed a progressive decrease in ethanol yield with each reuse, which is a common phenomenon observed in many immobilized enzyme and cell systems (Fig. 8 ). Our results indicate that ethanol yield decreased after each cycle, likely due to a combination of factors, including the decline in yeast cell viability, the gradual breakdown of the immobilization matrix, and the accumulation of inhibitory byproducts such as ethanol. In other study, the ethanol yield dropped after four cycles, which was attributed to both cellular stress and matrix degradation [ 39 ]. Similarly, in our study, the rate of decline in ethanol yield accelerated after the third cycle, suggesting that the yeast cells' metabolic capacity and structural integrity were compromised over time. The results of this study showed a slight decrease in ethanol production after the second reuse cycle of immobilized yeast cells; however, this change was not statistically significant. This trend suggests that the immobilized system maintained relatively stable activity over repeated use, although minor variations may be associated with factors such as gradual loss of cell viability or partial nutrient depletion within the immobilization matrix. Similarly, Onaifo et al. (2016) reported no significant reduction in ethanol yield over successive reuse cycles. Minor differences between the studies may be attributed to variations in experimental conditions, particularly the immobilization strategies employed. For instance, the support material used by Onaifo et al. (2016) may have provided a more protective microenvironment for the yeast cells [ 69 ]. Additionally, differences in immobilization parameters, such as cell loading density or entrapment technique, could also contribute to the observed variations. These findings emphasize the importance of optimizing immobilization systems to enhance the stability and reusability of yeast for industrial bioethanol production. 3.7. Ethanol physical characterization The physical properties of the distilled ethanol produced in this study were analyzed, and the results indicate that the ethanol meets key specifications commonly required for industrial applications (Table 2 ). The density of 0.812 g/cm³ is in close agreement with typical values for pure ethanol, which range from 0.789 g/cm³ to 0.792 g/cm³ at 20°C. This suggests that the ethanol produced is of high purity, with minimal impurities that could affect its density. The specific gravity of 0.827 is also consistent with standard ethanol, indicating that the ethanol produced is comparable to commercially available ethanol in terms of its physical composition and suitability for biofuel applications. The viscosity of 1.65 mm²/s is within the range typically observed for ethanol (1.2–1.5 mm²/s at 20°C), which is relatively low compared to other liquids like oils or water. This moderate viscosity is advantageous for transportation and handling, as it facilitates pumping and mixing without significant resistance to flow. However, it also suggests that some dissolved impurities or fermentation by-products might slightly increase the viscosity, though it remains within acceptable limits for most industrial processes. The API specific gravity value of 42.90 further confirms the ethanol’s suitability for fuel blending, particularly in the production of bioethanol as an alternative fuel. This value, a standard measure of a liquid’s density relative to water, positions the ethanol produced in this study within the typical range expected for fuel-grade ethanol, which is essential for its use in the automotive industry and other energy applications. The flash point of 14°C, consistent with the flash point of pure ethanol (around 12–14°C), highlights the highly flammable nature of the product. This property is crucial for handling and storage, as ethanol’s volatility makes it prone to ignition at relatively low temperatures. Therefore, proper safety precautions and storage conditions are required to mitigate risks in large-scale applications Table 2 The physical properties of the distilled ethanol Parameter Value Density (g/cm³) 0.812 Viscosity ((mm 2 /s) 1.65 Specific gravity 0.827 API specific gravity 42.90 Flash point (°C) 14.00 When compared to other studies, the ethanol produced in this study shows similar physical properties to those reported in the literature. For instance, [ 69 ] reported an ethanol density of 0.79–0.81 g/cm³, viscosity between 1.4–1.7 mm²/s, and flash points in the range of 12–15°C, which are all within the values observed in our study. This consistency suggests that the fermentation process in our study was both efficient and effective, producing ethanol with standard characteristics suitable for industrial and biofuel applications. The produced ethanol conforms to established bioethanol standards; however, further optimization of fermentation conditions remains necessary. Although flash point and viscosity are within acceptable limits, fine-tuning parameters such as pH and temperature could enhance purity and reduce residual components affecting ethanol quality. In addition, research into different immobilization techniques for yeast cells or the use of various protective additives could improve the yield and purity of ethanol, ensuring that the physical characteristics remain consistent over extended production cycles. 4. Discussion The ethanol concentration obtained in this study (11.57 g/L) can be classified as moderate for lignocellulosic bioethanol production using SHF process. This value falls within the commonly reported range of 10–25 g/L for lignocellulosic substrates, indicating that the overall process performed reasonably well under the applied conditions. However, the yield did not reach higher levels typically associated with optimized or industrial-scale systems (> 40 g/L), which may be attributed to the presence of lignin-derived inhibitory compounds and the intrinsic limitations of the SHF process[ 36 , 65 ]. The moderate ethanol yield obtained in this study is largely influenced by upstream processes, especially fungal delignification and enzymatic hydrolysis.. As shown in Section 3.1 , laccase production by C. caperata reached its maximum at 12 days, indicating strong oxidative capability toward lignin degradation. This enzymatic activity played a key role in reducing lignin content from 20.87% to 10.92% (Table 1 ), as discussed in Section 3.2. The substantial lignin removal enhanced cellulose accessibility, which was further reflected in the improved sugar release during enzymatic hydrolysis (Section 3.3). The hydrolysis profile (Fig. 2 ) showed a clear increase in reducing sugar concentration up to the optimum reaction time, confirming that fungal pretreatment effectively disrupted the lignocellulosic matrix. However, despite this improvement, the remaining lignin fraction (~ 10%) and its derived compounds likely still imposed partial inhibition on both enzymatic and fermentation processes. In lignocellulosic systems, residual lignin and its degradation products, such as phenolic compounds, furfural, and hydroxymethylfurfural (HMF), are known to inhibit both enzymatic hydrolysis and microbial fermentation. These compounds can disrupt yeast cell membranes, inhibit key metabolic enzymes, and reduce ethanol productivity [ 70 ]. Thus, even though fungal delignification significantly improved biomass accessibility, it may not have completely eliminated inhibitory effects, which explains the moderate ethanol yield obtained. To better contextualize these findings, a comparison with previous studies is presented in Table 3 . As shown in Table 3 , the ethanol yield obtained in this study (11.57 g/L) falls within the range reported for lignocellulosic bioethanol production using the SHF process. Lower ethanol concentrations, such as 9.45 g/L from rice straw, have been reported for substrates treated with alkali–enzymatic methods, whereas higher values (30–31 g/L) were achieved from sweet sorghum bagasse and Douglas-fir following steam pretreatment [ 71 , 72 ]. Intermediate ethanol yields were also observed in cassava stems subjected to microwave-assisted alkali and acid pretreatments (13.66–17.84 g/L) [ 73 ]. These variations indicate that ethanol production is strongly influenced by the type and severity of pretreatment. More intensive methods, such as steam pretreatment and chemical delignification, generally enhance cellulose accessibility and lead to higher ethanol yields, but they may also increase the formation of inhibitory compounds. In contrast, the fungal delignification approach employed in this study represents a milder and more environmentally friendly strategy. This method effectively reduces lignin content while preserving carbohydrate fractions, although it may result in slightly lower ethanol production compared to more aggressive pretreatment techniques. Interestingly, the ethanol yield obtained in this study is comparable to that reported for sugarcane bagasse using Ca-alginate immobilized yeast (11.8 g/L) [ 72 ], highlighting that the performance of the MIL-100(Fe)-immobilized yeast system is within the expected range for SHF-based lignocellulosic fermentation. Table 3 Comparison of lignocellulosic bioethanol production under different conditions Biomass Pretreatment / Delignification Yeast System Method Ethanol (g/L) Rice straw [ 71 ] Alkali + enzymatic treatment Free yeast SHF 9.45 Sweet sorghum bagasse [ 72 ] Steam pretreatment Free yeast SHF 31.20 Douglas-fir [ 72 ] Steam pretreatment Free yeast SHF 30.20 Cassava stems [ 73 ] Microwave-assisted alkali (3% NaOH) Free yeast SHF 17.84 Cassava stems [ 73 ] Microwave-assisted acid (0.1 M H 2 SO) Free yeast SHF 13.66 Taberau stems (this study) Fungal delignification MIL-100(Fe)-immobilized yeast SHF 11.57 Sugarcane bagasse [ 74 ] Microwave-assisted alkali (2.75% NaOH) Ca-alginate beads-immobilized yeast SHF 11.8 The effect of yeast dosage further highlights the importance of process optimization. Ethanol production increased significantly with increasing yeast concentration up to 2% (w/v), beyond which no substantial improvement was observed. This trend reflects the balance between microbial activity and substrate availability. At optimal conditions, yeast metabolism is maximized, allowing efficient conversion of fermentable sugars produced during hydrolysis. However, excessive yeast loading can lead to mass transfer limitations and increased competition for nutrients [ 68 ]. Fermentation time also played a decisive role, with maximum ethanol production observed at 4 days. This corresponds to the active metabolic phase of yeast, during which sugar consumption and ethanol formation are at their peak. The decline observed at longer fermentation times is consistent with substrate depletion and ethanol inhibition. Importantly, this trend aligns with the hydrolysis behavior, where sugar availability is time-dependent, indicating that synchronization between hydrolysis and fermentation is critical in SHF systems. A key advantage of this study lies in the application of MIL-100(Fe)-based immobilized yeast, as discussed in Section 3.4. Characterization results (FTIR, XRD, and SEM) confirmed that yeast cells were successfully immobilized without disrupting the structural integrity of the MIL-100(Fe) framework. The presence of functional groups associated with yeast cells, along with the preservation of MOF crystallinity, indicates that immobilization occurred through surface interactions rather than structural modification. This stable immobilization likely contributed to improved fermentation performance by enhancing cell stability, protecting cells from inhibitory compounds, and maintaining a high local cell density. Furthermore, MIL-100(Fe), with its porous structure and high surface area, supports yeast metabolic activity by enabling efficient substrate diffusion and enhanced mass transfer. These characteristics explain the superior performance of immobilized yeast compared to free yeast systems, as also reported in previous studies [ 38 , 64 , 66 , 67 ]. The recycle experiments further demonstrated the practical advantages of immobilization. Ethanol production remained relatively high during the initial cycles and gradually decreased with repeated use. This decline can be attributed to cell leakage, partial deactivation, or structural changes in the immobilization matrix. Nevertheless, the ability to reuse the immobilized system highlights its potential for cost-effective and sustainable bioethanol production. Finally, the physicochemical properties of the distilled ethanol confirmed the effectiveness of downstream processing. The density (0.812 g/cm³) and flash point (14°C) are close to those of commercial ethanol fuels, indicating successful enrichment through distillation. However, slight deviations from pure ethanol values suggest the presence of residual water, which is expected in single-stage distillation systems. This highlights the importance of integrating efficient purification techniques, such as fractional distillation or dehydration, to achieve fuel-grade ethanol. In general, this study demonstrates that the integration of fungal delignification, optimized fermentation conditions, and MIL-100(Fe)-based immobilized yeast can produce bioethanol from lignocellulosic biomass with competitive performance. Although the ethanol yield remains moderate, the system offers significant advantages in terms of sustainability, reusability, and process integration, providing a promising platform for further development and scale-up. Future work should focus on improving ethanol yield through process optimization, such as enhancing enzymatic hydrolysis efficiency, reducing the formation of inhibitory compounds, and optimizing nutrient supplementation during fermentation. In addition, alternative process configurations beyond SHF are strongly recommended, particularly simultaneous saccharification and fermentation (SSF) or simultaneous saccharification and co-fermentation (SSCF), which can improve overall efficiency by reducing product inhibition and enabling better integration of enzymatic hydrolysis and fermentation steps. These approaches have been widely reported to yield higher ethanol concentrations compared to SHF systems. Furthermore, the application of advanced separation techniques, including multi-stage distillation or dehydration processes, is recommended to obtain higher-purity ethanol. Further studies on the long-term performance of immobilized yeast and the structural integrity of the MIL-100(Fe) matrix are required to enable industrial application. 5. Conclusions This study demonstrates the effective use of C. caperata for laccase production to facilitate delignification of Taberau ( Phragmites australis ) biomass, enhancing its cellulose accessibility for enzymatic hydrolysis and subsequent bioethanol production. The fungal pre-treatment reduced lignin content significantly, which in turn improved sugar release during hydrolysis. Furthermore, yeast cells immobilized on MIL-100(Fe) exhibited enhanced ethanol production compared to free yeast cells, with optimal yields achieved under specific conditions of yeast dosage and fermentation time. The immobilized yeast system retained structural integrity and effectively consumed sugars, demonstrating its potential for sustainable bioethanol production. However, ethanol yield decreased over successive reuse cycles, indicating the necessity for further optimization. The combined use of fungal pre-treatment and immobilized yeast fermentation offers a viable approach to improve lignocellulosic biomass conversion into biofuels for both eco-friendly and industrial applications. Declarations Acknowledgements The authors would like to express their sincere appreciation to colleagues and collaborators for their valuable support and contributions throughout this study. Funding This research was financially supported by the Ministry of Research, Technology, and Higher Education, Indonesia under Grant No. 1000/UN24.13/AL.O4/2024. Author Contributions TAA was responsible for the methodology, investigation, formal analysis, data curation, and drafting the original manuscript. YY and YA contributed to the investigation, methodology, formal analysis, and data curation. WJ , LT , and EJK involved to the formal analysis, data curation, and preparing visualizations. RA provided to the conceptualization, methodology, validation, supervision, and project administration, as well as contributed to the manuscript review and editing. All authors have read and approved the final manuscript. Data availability All data are included in the manuscript, and additional data can be available upon request. Declarations of Conflic of interest The authors declare that there is no conflict of interest regarding the publication of this manuscript. All authors have contributed significantly to the research and preparation of the manuscript, and no competing financial, professional, or personal interests could have influenced the work or its interpretation. References Hasan, M. H., Mahlia, T. M. I., & Nur, H. (2012). A review on energy scenario and sustainable energy in Indonesia. Renewable and Sustainable Energy Reviews , 16 (4), 2316–2328. https://doi.org/10.1016/j.rser.2011.12.007 United Nations Department of Economic and Social Affairs, P. D. (2022). W. P. P. 2022: S. of R., Raimi, D., Zhu, Y., Newell, R. G., Prest, B. C., & Bergman, A. (2022). Global Energy Outlook 2023: Sowing the Seeds of an Energy Transition . United Nation . Retrieved from http://dx.doi.org/10.17605/OSF.IO/W3B4Z%0Awww.un.org/development/ desa/pd/. Holtom, B. (2004). Fossil fuels, alternative energy and economic growth. Journal of Nursing Administration , 34 (5), 216–227. Abas, N., Kalair, A., & Khan, N. (2015). Review of fossil fuels and future energy technologies. Futures , 69 , 31–49. https://doi.org/10.1016/j.futures.2015.03.003 Wang, J., & Azam, W. (2024). Natural resource scarcity, fossil fuel energy consumption, and total greenhouse gas emissions in top emitting countries. Geoscience Frontiers , 15 (2), 2024. https://doi.org/10.1016/j.gsf.2023.101757 Merritt, H., & Barragán-Ocaña, A. (2023). The impact of market factors on the development of eco-friendly energy technologies: the case of bioethanol. Clean Technologies and Environmental Policy , 25 (2), 313–321. https://doi.org/10.1007/s10098-021-02225-6 Ripple, W. J., Wolf, C., Gregg, J. W., Rockström, J., Mann, M. E., Oreskes, N., … Crowther, T. W. (2024). The 2024 state of the climate report : Perilous times, 1–13. Rajak, R. C., & Banerjee, R. (2018). An eco-friendly process integration for second generation bioethanol production from laccase delignified Kans grass. Energy Conversion and Management , 157 , 364–371. https://doi.org/10.1016/j.enconman.2017.11.060 Lamichhane, G., Acharya, A., Poudel, D. K., Aryal, B., Gyawali, N., Niraula, P., … Parajuli, N. (2021). Recent advances in bioethanol production from Lignocellulosic biomass. International Journal of Green Energy , 18 (7), 731–744. https://doi.org/10.1080/15435075.2021.1880910 Kordala, N., Walter, M., Brzozowski, B., & Lewandowska, M. (2024). 2G-biofuel ethanol: an overview of crucial operations, advances and limitations. Biomass Conversion and Biorefinery , 14 (3), 2983–3006. https://doi.org/10.1007/s13399-022-02861-y Limayem, A., & Ricke, S. C. (2012). Lignocellulosic biomass for bioethanol production: Current perspectives, potential issues and future prospects. Progress in Energy and Combustion Science , 38 (4), 449–467. https://doi.org/10.1016/j.pecs.2012.03.002 Das, N., Jena, P. K., Padhi, D., Kumar Mohanty, M., & Sahoo, G. (2023). A comprehensive review of characterization, pretreatment and its applications on different lignocellulosic biomass for bioethanol production. Biomass Conversion and Biorefinery , 13 (2), 1503–1527. https://doi.org/10.1007/s13399-021-01294-3 Persson, T., Garcia, A. G. Y., Paz, J. O., Fraisse, C. W., & Hoogenboom, G. (2010). Reduction in greenhouse gas emissions due to the use of bio-ethanol from wheat grain and straw produced in the south-eastern USA. Journal of Agricultural Science , 148 (5), 511–527. https://doi.org/10.1017/S0021859610000316 Abdul Kareem Joyia, M., Ahmad, M., Chen, Y. F., Mustaqeem, M., Ali, A., Abbas, A., & Ashraf Gondal, M. (2024). Trends and advances in sustainable bioethanol production technologies from first to fourth generation: A critical review. Energy Conversion and Management , 321 , 2024. https://doi.org/10.1016/j.enconman.2024.119037 Cavalaglio, G., Gelosia, M., Ingles, D., Pompili, E., D’Antonio, S., & Cotana, F. (2016). Response surface methodology for the optimization of cellulosic ethanol production from Phragmites australis through pre-saccharification and simultaneous saccharification and fermentation. Industrial Crops and Products , 83 , 431–437. https://doi.org/10.1016/j.indcrop.2015.12.089 Cotana, F., Cavalaglio, G., Pisello, A. L., Gelosia, M., Ingles, D., & Pompili, E. (2015). Sustainable ethanol production from common reed (Phragmites australis) through simultaneuos saccharification and fermentation. Sustainability (Switzerland) , 7 (9), 12149–12163. https://doi.org/10.3390/su70912149 Tozluoglu, A. (2018). Bioethanol production from common reed (Phragmites australis): Enzymatic hydrolysis and fermentation. Romanian Biotechnological Letters , 23 (2), 13473–13478. Köbbing, J. F., Thevs, N., & Zerbe, S. (2013). The utilisation of reed ( Phragmites australis ): a review. Mires and Peat , 13 , 1–14. Sathitsuksanoh, N., Zhu, Z., Templeton, N., Rollin, J. A., Harvey, S. P., & Zhang, Y. H. P. (2009). Saccharification of a potential bioenergy crop, phragmites australis (Common Reed), by lignocellulose fractionation followed by enzymatic hydrolysis at decreased cellulase loadings. Industrial and Engineering Chemistry Research , 48 (13), 6441–6447. https://doi.org/10.1021/ie900291s Gao, K., Boiano, S., Marzocchella, A., & Rehmann, L. (2014). Cellulosic butanol production from alkali-pretreated switchgrass (Panicum virgatum) and phragmites (Phragmites australis). Bioresource Technology , 174 , 176–181. https://doi.org/10.1016/j.biortech.2014.09.152 Elhaak, Mohsen, Hamada, & El-Gebaly. (2015). Biofuel Production From Phragmites Australis (Cav .) and Typha Domingensis (Pers.) Plants of Burullus Lake. J. Exp. Biol. (Bot.) , 11 (2), 237–243. Retrieved from http://my.ejmanger.com/ejeb/ Suryadi, H., Judono, J. J., Putri, M. R., Eclessia, A. D., Ulhaq, J. M., Agustina, D. N., & Sumiati, T. (2022). Biodelignification of lignocellulose using ligninolytic enzymes from white-rot fungi. Heliyon , 8 (2), e08865. https://doi.org/10.1016/j.heliyon.2022.e08865 Isikhuemhen, O. S., Mikiashvili, N. A., Senwo, Z. N., & Ohimain, E. I. (2014). Biodegradation and Sugar Release from Canola Plant Biomass by Selected White Rot Fungi. Advances in Biological Chemistry , 04 (06), 395–406. https://doi.org/10.4236/abc.2014.46045 De La Torre, M., Martín-Sampedro, R., Fillat, Ú., Eugenio, M. E., Blánquez, A., Hernández, M., … Ibarra, D. (2017). Comparison of the efficiency of bacterial and fungal laccases in delignification and detoxification of steam-pretreated lignocellulosic biomass for bioethanol production. Journal of Industrial Microbiology and Biotechnology , 44 (11), 1561–1573. https://doi.org/10.1007/s10295-017-1977-1 Plácido, J., & Capareda, S. (2015). Ligninolytic enzymes: A biotechnological alternative for bioethanol production. Bioresources and Bioprocessing , 2 (1). https://doi.org/10.1186/s40643-015-0049-5 Loi, M., Glazunova, O., Fedorova, T., Logrieco, A. F., & Mulè, G. (2021). Fungal laccases: The forefront of enzymes for sustainability. Journal of Fungi , 7 (12), 1–25. https://doi.org/10.3390/jof7121048 Roth, S., & Spiess, A. C. (2015). Laccases for biorefinery applications: a critical review on challenges and perspectives. Bioprocess and Biosystems Engineering , 38 (12), 2285–2313. https://doi.org/10.1007/s00449-015-1475-7 Liong, Y. Y., Halis, R., Lai, O. M., & Mohamed, R. (2012). Conversion of lignocellulosic biomass from grass to bioethanol using materials pretreated with alkali and the white rot fungus, Phanerochaete chrysosporium. BioResources , 7 (4), 5500–5513. https://doi.org/10.15376/biores.7.4.5500-5513 López-Abelairas, M., Álvarez Pallín, M., Salvachúa, D., Lú-Chau, T., Martínez, M. J., & Lema, J. M. (2013). Optimisation of the biological pretreatment of wheat straw with white-rot fungi for ethanol production. Bioprocess and Biosystems Engineering , 36 (9), 1251–1260. https://doi.org/10.1007/s00449-012-0869-z Castoldi, R., Bracht, A., de Morais, G. R., Baesso, M. L., Correa, R. C. G., Peralta, R. A., … Peralta, R. M. (2014). Biological pretreatment of Eucalyptus grandis sawdust with white-rot fungi: Study of degradation patterns and saccharification kinetics. Chemical Engineering Journal , 258 , 240–246. https://doi.org/10.1016/j.cej.2014.07.090 Ding, C., Wang, X., & Li, M. (2019). Evaluation of six white-rot fungal pretreatments on corn stover for the production of cellulolytic and ligninolytic enzymes, reducing sugars, and ethanol. Applied Microbiology and Biotechnology , 103 (14), 5641–5652. https://doi.org/10.1007/s00253-019-09884-y Agnestisia, R., Panjaitan, D., Erdene-ochir, T., & Nion, Y. A. (2024). Decolorization of Rhodamine B by the white rot fungus Coriolopsis caperata isolated from Central Kalimantan Forest. BIO Web of Conferences , 134 , 06011. Agnestisia, R., Karelius, K., Oksal, E., & Nion, Y. A. (2025). Decolorization of textile dyes with crude ligninolytic enzymes from Coriolopsis caperata. Jurnal Bioteknologi dan Biosains Indonesia , 12 (1), 104–118. Kallel, F., Neifar, M., Kacem, I., & Ellouz Chaabouni, S. (2024). Optimization of enzymatic hydrolysis and fermentation of bleached garlic straw for bioethanol production. Journal of Material Cycles and Waste Management , 26 (5), 2880–2895. https://doi.org/10.1007/s10163-024-02016-3 Averheim, A., Stagge, S., Jönsson, L. J., Larsson, S. H., & Thyrel, M. (2024). Separate hydrolysis and fermentation of softwood bark pretreated with 2-naphthol by steam explosion. Biotechnology for Biofuels and Bioproducts , 17 (1), 1–10. https://doi.org/10.1186/s13068-024-02552-y Drissen, R. E. T., Maas, R. H. W., Tramper, J., & Beeftink, H. H. (2009). Modelling ethanol production from cellulose: Separate hydrolysis and fermentation versus simultaneous saccharification and fermentation. Biocatalysis and Biotransformation , 27 (1), 27–35. https://doi.org/10.1080/10242420802564358 Bhadana, B., & Chauhan, M. (2016). Bioethanol Production Using Saccharomyces cerevisiae with Different Perspectives: Substrates, Growth Variables, Inhibitor Reduction and Immobilization. Fermentation Technology , 5 (2), 2–5. https://doi.org/10.4172/2167-7972.1000131 Kumoro, A. C., Damayanti, A., Bahlawan, Z. A. S., Melina, M., & Puspawati, H. (2021). Bioethanol production from oil palm empty fruit bunches using saccharomyces cerevisiae immobilized on sodium alginate beads. Periodica Polytechnica Chemical Engineering , 65 (4), 493–504. https://doi.org/10.3311/PPch.16775 Mariam, I., Manzoor, K., Ali, S., & Ikram-Ul-Haq. (2009). Enhanced production of ethanol from free and immobilized Saccharomyces cerevisiae under stationary culture. Pakistan Journal of Botany , 41 (2), 821–833. Md Zain, M., Kofli, N. T., & Yahya, A. S. R. S. (2011). Bioethanol Production by Calcium Alginate-Immobilised St1 Yeast System: Effects of Size of Beads, Ratio and Concentration. IIUM Engineering Journal , 12 (4), 11–19. https://doi.org/10.31436/iiumej.v12i4.159 Gökgöz, M., & Yiğitoğlu, M. (2011). Immobilization of Saccharomyces cerevisiae on to modified carboxymethylcellulose for production of ethanol. Bioprocess and Biosystems Engineering , 34 (7), 849–857. https://doi.org/10.1007/s00449-011-0535-x Liang, K., Richardson, J. J., Doonan, C. J., Mulet, X., Ju, Y., Cui, J., … Falcaro, P. (2017). An Enzyme-Coated Metal–Organic Framework Shell for Synthetically Adaptive Cell Survival. Angewandte Chemie - International Edition , 56 (29), 8510–8515. https://doi.org/10.1002/anie.201704120 Sun, H., Li, Y., Yu, S., & Liu, J. (2020). Metal-organic frameworks (MOFs) for biopreservation: From biomacromolecules, living organisms to biological devices. Nano Today , 35 , 2024. https://doi.org/10.1016/j.nantod.2020.100985 Liang, S., Wu, X. L., Xiong, J., Zong, M. H., & Lou, W. Y. (2020). Metal-organic frameworks as novel matrices for efficient enzyme immobilization: An update review. Coordination Chemistry Reviews , 406 , 213149. https://doi.org/10.1016/j.ccr.2019.213149 Hu, J., Yuan, X., Wang, C., Shao, X., Yang, B., Abdul Razzaq, A., … Peng, Y. (2020). Self-Phosphorization of MOF-Armored Microbes for Advanced Energy Storage. Small , 16 (24), 1–9. https://doi.org/10.1002/smll.202000755 Zhong, G., Liu, D., & Zhang, J. (2018). Applications of Porous Metal-Organic Framework MIL-100(M) (M = Cr, Fe, Sc, Al, V). Crystal Growth and Design , 18 (12), 7730–7744. https://doi.org/10.1021/acs.cgd.8b01353 Zhang, F., Shi, J., Jin, Y., Fu, Y., Zhong, Y., & Zhu, W. (2015). Facile synthesis of MIL-100(Fe) under HF-free conditions and its application in the acetalization of aldehydes with diols. Chemical Engineering Journal , 259 , 183–190. https://doi.org/10.1016/j.cej.2014.07.119 Leonowicz, A., & Gzrywnowicz, K. (1981). Quantitative estimation of laccase forms in some white-rot fungi using syringaldazine as a substrate. Enzyme and Microbial Technology , 3 , 55–58. Ghose, T. K. (1987). Measurement of cellulase activities, 59 (2), 257–268. https://doi.org/doi:10.1351/pac198759020257 Sivakumar, R., Rajendran, R., Balakumar, C., Tamilvendan, M., & R. Sivakumar, R. Rajendran, C. Balakumar, M. T. (2010). Isolation , Screening and Optimization of Production Medium for Thermostable Laccase Production from Ganoderma sp. International Journal of Engineering Science , 2 (12), 7133–7141. Retrieved from http://www.ijest.info/docs/IJEST10-02-12-241.pdf Couto, S. R., Gundín, M., Lorenzo, M., & Sanromán, M. Á. (2002). Screening of supports and inducers for laccase production by Trametes versicolor in semi-solid-state conditions. Process Biochemistry , 38 (2), 249–255. https://doi.org/10.1016/S0032-9592(02)00087-0 Patrick, F., Mtui, G., Mshandete, A., & Kivaisi, A. (2010). Tanzania Journal of Science Peroxidase and Laccase in Submerged Cultures of, 36 . Hadibarata, T., & Yuniarto, A. (2020). Biodegradation of polycyclic aromatic hydrocarbons by high-laccase basidiomycetes fungi isolated from tropical forest of Borneo. Biocatalysis and Agricultural Biotechnology , 28 (July), 101717. https://doi.org/10.1016/j.bcab.2020.101717 Wan, C., & Li, Y. (2011). Effectiveness of microbial pretreatment by Ceriporiopsis subvermispora on different biomass feedstocks. Bioresource Technology , 102 (16), 7507–7512. https://doi.org/10.1016/j.biortech.2011.05.026 Bagewadi, Z. K., Mulla, S. I., & Ninnekar, H. Z. (2017). Optimization of laccase production and its application in delignification of biomass. International Journal of Recycling of Organic Waste in Agriculture , 6 (4), 351–365. https://doi.org/10.1007/s40093-017-0184-4 Gupta, R., Mehta, G., Khasa, Y. P., & Kuhad, R. C. (2011). Fungal delignification of lignocellulosic biomass improves the saccharification of cellulosics. Biodegradation , 22 (4), 797–804. https://doi.org/10.1007/s10532-010-9404-6 Pasaribu, M. H., Karelius, K., Ramdhani, E. P., Agnestisia, R., Pereiz, Z., & Toepak, E. P. (2023). Synthesis of Mil-100(Fe)@Fe3O4 Composite using Zircon Mining Magnetic Waste as an Adsorbent for Methylene Blue Dye. BIO Web of Conferences , 70 . https://doi.org/10.1051/bioconf/20237002010 Permyakova, A., Kakar, A., Bachir, J., Gkaniatsou, E., Haye, B., Menguy, N., … Sicard, C. (2022). In Situ Synthesis of a Mesoporous MIL-100 ( Fe ) Bacteria Exoskeleton. ACS Materials Letters , 5 (1), 79–84. Chávez, A. M., Rey, A., López, J., Álvarez, P. M., & Beltrán, F. J. (2021). Critical aspects of the stability and catalytic activity of MIL-100(Fe) in different advanced oxidation processes. Separation and Purification Technology , 255 (June 2020), 117660. https://doi.org/10.1016/j.seppur.2020.117660 Han, L., Qi, H., Zhang, D., Ye, G., Zhou, W., Hou, C., … Sun, Y. (2017). A facile and green synthesis of MIL-100(Fe) with high-yield and its catalytic performance. New Journal of Chemistry , 41 (22), 13504–13509. https://doi.org/10.1039/c7nj02975f Zhang, X., Zhang, X., He, W., Sun, C., Ma, J., Yuan, J., & Du, X. (2013). High-performance mesoporous LiFePO4 from Baker’s yeast. Colloids and Surfaces B: Biointerfaces , 103 (October 2012), 114–120. https://doi.org/10.1016/j.colsurfb.2012.10.002 Lyu, F., Zhang, Y., Zare, R. N., Ge, J., & Liu, Z. (2014). One-pot synthesis of protein-embedded metal-organic frameworks with enhanced biological activities. Nano Letters , 14 (10), 5761–5765. https://doi.org/10.1021/nl5026419 Yang, W., Liang, W., O’Dell, L. A., Toop, H. D., Maddigan, N., Zhang, X., … Huang, J. (2021). Insights into the Interaction between Immobilized Biocatalysts and Metal-Organic Frameworks: A Case Study of PCN-333. JACS Au , 1 (12), 2172–2181. https://doi.org/10.1021/jacsau.1c00226 Mulko, L., Rivarola, C. R., Barbero, C. A., & Acevedo, D. F. (2016). Bioethanol production by reusable Saccharomyces cerevisiae immobilized in a macroporous monolithic hydrogel matrices. Journal of Biotechnology , 233 , 56–65. https://doi.org/10.1016/j.jbiotec.2016.07.004 Zabed, H., Sahu, J. N., Suely, A., Boyce, A. N., & Faruq, G. (2017). Bioethanol production from renewable sources: Current perspectives and technological progress. Renewable and Sustainable Energy Reviews , 71 , 475–501. https://doi.org/10.1016/j.rser.2016.12.076 Kourkoutas, Y., Bekatorou, A., Banat, I. M., Marchant, R., & Koutinas, A. A. (2004). Immobilization technologies and support materials suitable in alcohol beverages production: A review. Food Microbiology , 21 (4), 377–397. https://doi.org/10.1016/j.fm.2003.10.005 Shen, H. Y., Moonjai, N., Verstrepen, K. J., & Delvaux, F. R. (2003). Impact of attachment immobilization on yeast physiology and fermentation performance. Journal of the American Society of Brewing Chemists , 61 (2), 79–87. https://doi.org/10.1094/asbcj-61-0079 Bai, F. W., Anderson, W. A., & Moo-Young, M. (2008). Ethanol fermentation technologies from sugar and starch feedstocks. Biotechnology Advances , 26 (1), 89–105. https://doi.org/10.1016/j.biotechadv.2007.09.002 Onaifo, J. O., Ikhuoria, E. U., Omorogbe, S. O., Agbonlahor, O. G., & Ifijen, I. H. (2016). Coconut Pit an Alternative Source of Producing Renewable Material (Bioethanol). Journal of Applied Chemical Science International , 6 (2), 80–84. Jönsson, L. J., & Martín, C. (2016). Pretreatment of lignocellulose: Formation of inhibitory by-products and strategies for minimizing their effects. Bioresource Technology , 199 , 103–112. https://doi.org/10.1016/j.biortech.2015.10.009 Jin, X., Song, J., & Liu, G. Q. (2020). Bioethanol production from rice straw through an enzymatic route mediated by enzymes developed in-house from Aspergillus fumigatus. Energy , 190 (xxxx), 116395. https://doi.org/10.1016/j.energy.2019.116395 Shen, F., Kumar, L., Hu, J., & Saddler, J. N. (2011). Evaluation of hemicellulose removal by xylanase and delignification on SHF and SSF for bioethanol production with steam-pretreated substrates. Bioresource Technology , 102 (19), 8945–8951. https://doi.org/10.1016/j.biortech.2011.07.028 Pooja, N. S., Sajeev, M. S., Jeeva, M. L., & Padmaja, G. (2018). Bioethanol production from microwave-assisted acid or alkali-pretreated agricultural residues of cassava using separate hydrolysis and fermentation (SHF). 3 Biotech , 8 (1), 1–12. https://doi.org/10.1007/s13205-018-1095-4 Singh, A., Sharma, P., Saran, A. K., Singh, N., & Bishnoi, N. R. (2013). Comparative study on ethanol production from pretreated sugarcane bagasse using immobilized Saccharomyces cerevisiae on various matrices. Renewable Energy , 50 , 488–493. https://doi.org/10.1016/j.renene.2012.07.003 Supplementary Files Graphicalabstract.jpg Graphical abstract Cite Share Download PDF Status: Under Review Version 1 posted Reviewers agreed at journal 06 May, 2026 Reviewers invited by journal 06 May, 2026 Editor invited by journal 27 Apr, 2026 First submitted to journal 25 Apr, 2026 You are reading this latest preprint version Research Square lets you share your work early, gain feedback from the community, and start making changes to your manuscript prior to peer review in a journal. As a division of Research Square Company, we’re committed to making research communication faster, fairer, and more useful. We do this by developing innovative software and high quality services for the global research community. Our growing team is made up of researchers and industry professionals working together to solve the most critical problems facing scientific publishing. Also discoverable on Platform About Our Team In Review Editorial Policies Advisory Board Help Center Resources Author Services Accessibility API Access RSS feed Manage Cookie Preferences © Research Square 2026 | ISSN 2693-5015 (online) Privacy Policy Terms of Service Do Not Sell My Personal Information {"props":{"pageProps":{"initialData":{"identity":"rs-9512742","acceptedTermsAndConditions":true,"allowDirectSubmit":false,"archivedVersions":[],"articleType":"Research Article","associatedPublications":[],"authors":[{"id":635186855,"identity":"3731fff8-889b-49c0-a0d6-a38525bc9ac9","order_by":0,"name":"Titin Apung Atikah","email":"","orcid":"","institution":"Universitas Palangka Raya","correspondingAuthor":false,"prefix":"","firstName":"Titin","middleName":"Apung","lastName":"Atikah","suffix":""},{"id":635186856,"identity":"d71e6719-da41-473c-9c82-5b77249ac28e","order_by":1,"name":"Yanetri Asi","email":"","orcid":"","institution":"Universitas Palangka 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05:40:45","currentVersionCode":1,"declarations":"","doi":"10.21203/rs.3.rs-9512742/v1","doiUrl":"https://doi.org/10.21203/rs.3.rs-9512742/v1","draftVersion":[],"editorialEvents":[],"editorialNote":"","failedWorkflow":false,"files":[{"id":109277924,"identity":"c29f2028-a041-452e-9646-4af82dc61838","added_by":"auto","created_at":"2026-05-14 15:34:23","extension":"jpg","order_by":1,"title":"Figure 1","display":"","copyAsset":false,"role":"figure","size":39658,"visible":true,"origin":"","legend":"\u003cp\u003eEffect of incubation time on laccase production\u003c/p\u003e","description":"","filename":"1.jpg","url":"https://assets-eu.researchsquare.com/files/rs-9512742/v1/d280ee3c5a4fc00d86746ef1.jpg"},{"id":109296220,"identity":"374892a2-3ea9-47c1-8567-76df23a07eef","added_by":"auto","created_at":"2026-05-15 08:46:16","extension":"jpg","order_by":2,"title":"Figure 2","display":"","copyAsset":false,"role":"figure","size":118447,"visible":true,"origin":"","legend":"\u003cp\u003eEffect of hydrolysis time on reducing sugar concentration\u003c/p\u003e","description":"","filename":"2.jpg","url":"https://assets-eu.researchsquare.com/files/rs-9512742/v1/09060226f5959f2c7a80f6ed.jpg"},{"id":109296045,"identity":"bb34e7de-9dfd-4012-a9c8-8230d194a981","added_by":"auto","created_at":"2026-05-15 08:44:49","extension":"jpg","order_by":3,"title":"Figure 3","display":"","copyAsset":false,"role":"figure","size":78467,"visible":true,"origin":"","legend":"\u003cp\u003eFTIR spectra of MIL-100(Fe) and MIL-100(Fe)@yeast\u003c/p\u003e","description":"","filename":"3.jpg","url":"https://assets-eu.researchsquare.com/files/rs-9512742/v1/f757017da64c8c858c28e342.jpg"},{"id":109277919,"identity":"e7de06c9-fa5e-4212-a136-ed15bc2c0999","added_by":"auto","created_at":"2026-05-14 15:34:23","extension":"jpg","order_by":4,"title":"Figure 4","display":"","copyAsset":false,"role":"figure","size":55517,"visible":true,"origin":"","legend":"\u003cp\u003eFTIR Diffractogram of MIL-100(Fe) and MIL-100(Fe)@yeast.\u003c/p\u003e\n\u003cp\u003eNote: \u0026nbsp;●, MIL-100(Fe) and \u0026nbsp;■ , yeast\u003c/p\u003e","description":"","filename":"4.jpg","url":"https://assets-eu.researchsquare.com/files/rs-9512742/v1/c03961f8b37ad9cb0da7cbef.jpg"},{"id":109296645,"identity":"1a412973-ddfc-40be-947d-09b5aa495db1","added_by":"auto","created_at":"2026-05-15 08:48:54","extension":"jpg","order_by":5,"title":"Figure 5","display":"","copyAsset":false,"role":"figure","size":172051,"visible":true,"origin":"","legend":"\u003cp\u003eSEM images of (a) MIL-100(Fe) and (b) MIL-100(Fe)@yeast\u003c/p\u003e","description":"","filename":"5.jpg","url":"https://assets-eu.researchsquare.com/files/rs-9512742/v1/937d6fcaac545054512b4bd5.jpg"},{"id":109296533,"identity":"223f5a07-bbdf-4972-a916-09e21a87d12c","added_by":"auto","created_at":"2026-05-15 08:47:59","extension":"jpg","order_by":6,"title":"Figure 6","display":"","copyAsset":false,"role":"figure","size":61780,"visible":true,"origin":"","legend":"\u003cp\u003eEffect of different free and immobilized yeast dosage on (a) sugar consumption and (b) ethanol production\u003c/p\u003e","description":"","filename":"6.jpg","url":"https://assets-eu.researchsquare.com/files/rs-9512742/v1/90a2d06f4f0f7034e100a925.jpg"},{"id":109277922,"identity":"a494b693-7936-4a93-8e2f-ee4815b84a4c","added_by":"auto","created_at":"2026-05-14 15:34:23","extension":"jpg","order_by":7,"title":"Figure 7","display":"","copyAsset":false,"role":"figure","size":63093,"visible":true,"origin":"","legend":"\u003cp\u003eEffect of different fermentation time on (a) sugar consumption and (b) ethanol production\u003c/p\u003e","description":"","filename":"7.jpg","url":"https://assets-eu.researchsquare.com/files/rs-9512742/v1/042b7f595aad2168ff29dd78.jpg"},{"id":109297838,"identity":"1788695f-2c57-4f3e-89ea-59a79399fa37","added_by":"auto","created_at":"2026-05-15 09:06:37","extension":"jpg","order_by":8,"title":"Figure 8","display":"","copyAsset":false,"role":"figure","size":98685,"visible":true,"origin":"","legend":"\u003cp\u003eReusability of MIL-100(Fe)@yeast\u003c/p\u003e","description":"","filename":"8.jpg","url":"https://assets-eu.researchsquare.com/files/rs-9512742/v1/ad371b625b37078c9d5acf90.jpg"},{"id":109296501,"identity":"c727be45-bf39-4402-b2d3-7290057ed916","added_by":"auto","created_at":"2026-05-15 08:47:34","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"manuscript-pdf","size":763725,"visible":true,"origin":"","legend":"","description":"","filename":"manuscript.pdf","url":"https://assets-eu.researchsquare.com/files/rs-9512742/v1/1ae17687-3122-4b5a-81f8-4698d2749692.pdf"},{"id":109277916,"identity":"59ea9b3f-2baa-4f83-b6c4-35ffe3ca4dde","added_by":"auto","created_at":"2026-05-14 15:34:22","extension":"jpg","order_by":1,"title":"","display":"","copyAsset":false,"role":"supplement","size":96886,"visible":true,"origin":"","legend":"\u003cp\u003eGraphical abstract\u003c/p\u003e","description":"","filename":"Graphicalabstract.jpg","url":"https://assets-eu.researchsquare.com/files/rs-9512742/v1/7b69a1b5672756e2c40a5fb8.jpg"}],"financialInterests":"","formattedTitle":"Green Biorefinery Approach for Taberau (Phragmites australis) Biomass: Delignification by Coriolopsis caperata and Bioethanol Production via MIL-100(Fe)–Immobilized Saccharomyces cerevisiae","fulltext":[{"header":"Highlights","content":"\u003cul\u003e\n \u003cli\u003eCurrent perspective of bioethanol production from Taberau (\u003cem\u003ePhragmites australis\u003c/em\u003e) Biomass\u003c/li\u003e\n \u003cli\u003eUtilizing \u003cem\u003eCoriolopsis caperata\u003c/em\u003e, a ligninolytic fungus from Central Kalimantan, for efficient hydrolysis of lignocellulosic biomass\u003c/li\u003e\n \u003cli\u003eImproved performance of fermentation due to the action of immobilized yeast (\u003cem\u003eSaccharomises cerevisiae\u003c/em\u003e).\u003c/li\u003e\n \u003cli\u003eMIL-100(Fe) provided a robust platform for yeast immobilization, improving stability, reusability, and catalytic efficiency.\u003c/li\u003e\n\u003c/ul\u003e"},{"header":"1. Introduction","content":"\u003cp\u003eIndonesia's heavy dependence on fossil fuels for energy poses a significant challenge. According to Hasan et al. (2012), fossil energy sources\u0026mdash;petroleum (34.37%), coal (29.02%), and natural gas (23.49%)\u0026mdash;dominate the nation's energy mix, while renewable energy contributes only about 11.95% [\u003cspan citationid=\"CR1\" class=\"CitationRef\"\u003e1\u003c/span\u003e]. Meanwhile, global energy demand is projected to rise by 45% by 2030, with an average annual growth rate of 1.6% [\u003cspan citationid=\"CR2\" class=\"CitationRef\"\u003e2\u003c/span\u003e]. However, fossil fuels are finite resources that will become increasingly scarce [\u003cspan additionalcitationids=\"CR4\" citationid=\"CR3\" class=\"CitationRef\"\u003e3\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e5\u003c/span\u003e]. In addition to their finite availability, the combustion of fossil fuels contributes to environmental degradation through greenhouse gas emissions, thereby accelerating global warming [\u003cspan additionalcitationids=\"CR6\" citationid=\"CR5\" class=\"CitationRef\"\u003e5\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR7\" class=\"CitationRef\"\u003e7\u003c/span\u003e]. These pressing challenges underscore the urgent need for sustainable, renewable energy alternatives to secure a cleaner, more resilient energy future.\u003c/p\u003e \u003cp\u003eOne viable alternative is bioethanol, a renewable energy source with significant potential to transform the energy sector [\u003cspan citationid=\"CR6\" class=\"CitationRef\"\u003e6\u003c/span\u003e, \u003cspan citationid=\"CR8\" class=\"CitationRef\"\u003e8\u003c/span\u003e, \u003cspan citationid=\"CR9\" class=\"CitationRef\"\u003e9\u003c/span\u003e]. In particular, lignocellulosic biomass\u0026mdash;non-food plant material\u0026mdash;emerges as an ideal feedstock for second-generation (G2) bioethanol production [\u003cspan citationid=\"CR9\" class=\"CitationRef\"\u003e9\u003c/span\u003e, \u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e]. This biomass is rich in cellulose and hemicellulose, which can be converted into bioethanol through advanced processes [\u003cspan citationid=\"CR11\" class=\"CitationRef\"\u003e11\u003c/span\u003e, \u003cspan citationid=\"CR12\" class=\"CitationRef\"\u003e12\u003c/span\u003e]. Persson (2018) highlighted that G2 bioethanol offers lower greenhouse gas emissions than conventional fossil fuels [\u003cspan citationid=\"CR13\" class=\"CitationRef\"\u003e13\u003c/span\u003e], making it a more sustainable and environmentally friendly option. However, scaling up G2 bioethanol production presents several challenges. The current efficiency of feedstock selection, biomass pre-treatment, and fermentation processes still falls short of what is required for widespread commercial viability [\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e]. Despite these obstacles, the potential of bioethanol as a sustainable, low-emission energy source has never been more promising.\u003c/p\u003e \u003cp\u003e \u003cem\u003ePhragmites australis\u003c/em\u003e (locally known as \u003cem\u003eTaberau\u003c/em\u003e), a weed species from the Poaceae family, has emerged as a promising lignocellulosic feedstock for G2 bioethanol production [\u003cspan citationid=\"CR15\" class=\"CitationRef\"\u003e15\u003c/span\u003e, \u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e16\u003c/span\u003e]. Native to the peat swamp forests of Central Kalimantan, this plant is found in abundance but remains underutilized in the context of renewable energy applications. With its high cellulose and hemicellulose content [\u003cspan additionalcitationids=\"CR16\" citationid=\"CR15\" class=\"CitationRef\"\u003e15\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR17\" class=\"CitationRef\"\u003e17\u003c/span\u003e], \u003cem\u003eTaberau\u003c/em\u003e is considered a promising raw material for G2 bioethanol production using non-edible biomass sources. In addition to its favorable chemical composition, \u003cem\u003eTaberau\u003c/em\u003e offers several other beneficial traits, such as rapid growth, high biomass yield, and minimal requirements for water and nutrients [\u003cspan additionalcitationids=\"CR17 CR18\" citationid=\"CR16\" class=\"CitationRef\"\u003e16\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR19\" class=\"CitationRef\"\u003e19\u003c/span\u003e]. These characteristics position \u003cem\u003eTaberau\u003c/em\u003e as a sustainable, low-impact biomass resource, aligning with the objectives of advancing biofuel technologies and and the transition toward more efficient and eco-friendly energy systems. However, utilizing lignocellulosic biomass effectively requires pre-treatment to remove lignin and facilitate the breakdown of cellulose and hemicellulose. Several pre-treatment methods have been developed, including base treatment [\u003cspan citationid=\"CR20\" class=\"CitationRef\"\u003e20\u003c/span\u003e], acid treatment [\u003cspan citationid=\"CR20\" class=\"CitationRef\"\u003e20\u003c/span\u003e, \u003cspan citationid=\"CR21\" class=\"CitationRef\"\u003e21\u003c/span\u003e], organic solvent-based fractionation [\u003cspan citationid=\"CR19\" class=\"CitationRef\"\u003e19\u003c/span\u003e], and steam explosion [\u003cspan citationid=\"CR15\" class=\"CitationRef\"\u003e15\u003c/span\u003e, \u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e16\u003c/span\u003e]. These methods often require the use of chemicals or extreme reaction conditions, which can lead to biomass degradation and the formation of enzymatic inhibitors that hinder the saccharification process at later stages [\u003cspan citationid=\"CR12\" class=\"CitationRef\"\u003e12\u003c/span\u003e]. Consequently, biological methods\u0026mdash;particularly those utilizing white rot fungi\u0026mdash;are increasingly being explored as an alternative. This method is advantageous because they do not rely on harsh chemicals or extreme conditions, making them more cost-effective and environmentally friendly [\u003cspan citationid=\"CR12\" class=\"CitationRef\"\u003e12\u003c/span\u003e, \u003cspan citationid=\"CR22\" class=\"CitationRef\"\u003e22\u003c/span\u003e, \u003cspan citationid=\"CR23\" class=\"CitationRef\"\u003e23\u003c/span\u003e].\u003c/p\u003e \u003cp\u003eSeveral studies have reported that white rot fungi are capable of lignin degradation by producing ligninolytic enzymes such as laccase, lignin peroxidase, and manganese peroxidase [\u003cspan citationid=\"CR12\" class=\"CitationRef\"\u003e12\u003c/span\u003e, \u003cspan additionalcitationids=\"CR25\" citationid=\"CR24\" class=\"CitationRef\"\u003e24\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR26\" class=\"CitationRef\"\u003e26\u003c/span\u003e]. Roth and Spiess [\u003cspan citationid=\"CR27\" class=\"CitationRef\"\u003e27\u003c/span\u003e] further emphasized that laccase is particularly effective for detoxifying phenolic fragments in delignified biomass, which can otherwise inhibit cellulolytic enzyme activity during the saccharification process at later stages. Numerous white rot fungi species have been employed for biomass delignification, including \u003cem\u003ePhanerochaete chrysosporium\u003c/em\u003e [\u003cspan citationid=\"CR28\" class=\"CitationRef\"\u003e28\u003c/span\u003e], \u003cem\u003ePleurotus eryngii, Irpex lacteus\u003c/em\u003e [\u003cspan citationid=\"CR29\" class=\"CitationRef\"\u003e29\u003c/span\u003e], \u003cem\u003eGanoderma lucidum, Pleurotus ostreatus, Pleurotus pulmonarius, Trametes sp.\u003c/em\u003e [\u003cspan citationid=\"CR30\" class=\"CitationRef\"\u003e30\u003c/span\u003e], \u003cem\u003eand Pleurotus sajor-caju\u003c/em\u003e [\u003cspan citationid=\"CR31\" class=\"CitationRef\"\u003e31\u003c/span\u003e]. \u003cem\u003eThe white rot fungus Coriolopsis caperata, isolated from\u003c/em\u003e the previous study, \u003cem\u003ehas shown considerable laccase enzyme activity\u003c/em\u003e [\u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e32\u003c/span\u003e, \u003cspan citationid=\"CR33\" class=\"CitationRef\"\u003e33\u003c/span\u003e]. Therefore, biomass pre-treatment using \u003cem\u003eC. caperata\u003c/em\u003e for the delignification process is considered a promising approach.\u003c/p\u003e \u003cp\u003eFurthermore, the Separate Hydrolysis and Fermentation (SHF) method, which decouples the hydrolysis of cellulose from the fermentation stages, has demonstrated significant potential for second-generation (G2) bioethanol production [\u003cspan citationid=\"CR34\" class=\"CitationRef\"\u003e34\u003c/span\u003e, \u003cspan citationid=\"CR35\" class=\"CitationRef\"\u003e35\u003c/span\u003e]. In this process, cellulose is first converted into glucose through hydrolysis, followed by fermentation into ethanol by \u003cem\u003eSaccharomyces cerevisiae\u003c/em\u003e (yeast) [\u003cspan citationid=\"CR34\" class=\"CitationRef\"\u003e34\u003c/span\u003e, \u003cspan citationid=\"CR35\" class=\"CitationRef\"\u003e35\u003c/span\u003e]. The separation of these steps in the SHF process enables independent optimization of enzymatic hydrolysis and fermentation conditions, thereby enhancing overall efficiency and reducing the inhibitory impact of ethanol on cellulolytic enzymes [\u003cspan citationid=\"CR36\" class=\"CitationRef\"\u003e36\u003c/span\u003e]. Moreover, \u003cem\u003eS. cerevisiae\u003c/em\u003e can be immobilized onto various support materials, which facilitates cell separation from the fermentation medium, reduces operational costs through cell reuse across multiple fermentation cycles, enhances cell stability, and decreases contamination risks and fermentation time [\u003cspan citationid=\"CR37\" class=\"CitationRef\"\u003e37\u003c/span\u003e, \u003cspan citationid=\"CR38\" class=\"CitationRef\"\u003e38\u003c/span\u003e]. Common support materials for \u003cem\u003eS. cerevisiae\u003c/em\u003e immobilization include sodium alginate [\u003cspan citationid=\"CR38\" class=\"CitationRef\"\u003e38\u003c/span\u003e, \u003cspan citationid=\"CR39\" class=\"CitationRef\"\u003e39\u003c/span\u003e], calcium alginate [\u003cspan citationid=\"CR40\" class=\"CitationRef\"\u003e40\u003c/span\u003e], and CMC-g-PVP [\u003cspan citationid=\"CR41\" class=\"CitationRef\"\u003e41\u003c/span\u003e]. Recently, however, metal-organic frameworks (MOFs) have recently been recognized as promising platforms for the immobilization of \u003cem\u003eS. cerevisiae\u003c/em\u003e [\u003cspan citationid=\"CR42\" class=\"CitationRef\"\u003e42\u003c/span\u003e, \u003cspan citationid=\"CR43\" class=\"CitationRef\"\u003e43\u003c/span\u003e] and \u003cem\u003eEscherichia coli\u003c/em\u003e [\u003cspan citationid=\"CR44\" class=\"CitationRef\"\u003e44\u003c/span\u003e, \u003cspan citationid=\"CR45\" class=\"CitationRef\"\u003e45\u003c/span\u003e]. Among these MOFs, MIL-100(Fe) has attracted considerable attention due to its regular crystal structure, excellent stability, high surface area, non-toxicity, and biodegradability, making it an ideal candidate for yeast immobilization [\u003cspan citationid=\"CR46\" class=\"CitationRef\"\u003e46\u003c/span\u003e, \u003cspan citationid=\"CR47\" class=\"CitationRef\"\u003e47\u003c/span\u003e]. Despite these advances, the optimization of the SHF method using \u003cem\u003eTaberau\u003c/em\u003e biomass pre-treated with \u003cem\u003eCoriolopsis caperata\u003c/em\u003e and MIL-100(Fe)-immobilized \u003cem\u003eS. cerevisiae\u003c/em\u003e for bioethanol production has not yet been thoroughly investigated. The present research aims to bridge this gap by evaluating Taberau as a sustainable raw material for bioethanol production, enhanced by innovative pre-treatment and fermentation approaches.\u003c/p\u003e"},{"header":"2. Materials and Methods","content":"\u003cdiv id=\"Sec3\" class=\"Section2\"\u003e \u003ch2\u003e2.1. Materials\u003c/h2\u003e \u003cp\u003eThe materials used in this study were iron (II) sulfate heptahydrate (FeSO\u003csub\u003e4\u003c/sub\u003e.7H\u003csub\u003e2\u003c/sub\u003eO), trimesic acid (H\u003csub\u003e3\u003c/sub\u003eBTC), sodium hydroxide (NaOH), sodium acetate (CH\u003csub\u003e3\u003c/sub\u003eCOONa), acetic acid (CH\u003csub\u003e3\u003c/sub\u003eCOONa), veratryl alcohol ((CH\u003csub\u003e3\u003c/sub\u003eO)\u003csub\u003e2\u003c/sub\u003eC\u003csub\u003e6\u003c/sub\u003eH\u003csub\u003e3\u003c/sub\u003eCH\u003csub\u003e2\u003c/sub\u003eOH), syringaldazine ([HOC\u003csub\u003e6\u003c/sub\u003eH\u003csub\u003e2\u003c/sub\u003e(OCH\u003csub\u003e3\u003c/sub\u003e)\u003csub\u003e2\u003c/sub\u003eCH\u0026thinsp;=\u0026thinsp;N-]\u003csub\u003e2\u003c/sub\u003e), which were supplied by Merck. D-glucose (C\u003csub\u003e6\u003c/sub\u003eH\u003csub\u003e12\u003c/sub\u003eO\u003csub\u003e6\u003c/sub\u003e), peptone, potassium dihydrogen phosphate (KH\u003csub\u003e2\u003c/sub\u003ePO\u003csub\u003e4\u003c/sub\u003e), dipotassium hydrogen phosphate (K\u003csub\u003e2\u003c/sub\u003eHPO\u003csub\u003e4\u003c/sub\u003e), zinc sulfate (ZnSO\u003csub\u003e4\u003c/sub\u003e), iron(II) sulfate (FeSO\u003csub\u003e4\u003c/sub\u003e), manganese (II) sulfate (MnSO\u003csub\u003e4\u003c/sub\u003e), and magnesium sulfate (MgSO4) was used to prepare a glucose-peptone medium, while yeast extract, peptone, D-glucose (C\u003csub\u003e6\u003c/sub\u003eH\u003csub\u003e12\u003c/sub\u003eO\u003csub\u003e6\u003c/sub\u003e), agar was used to prepare yeast extract peptone glucose (YPG) agar medium. 3,5-Dinitrosalicylic acid (DNS) reagent was used for estimating reducing sugar concentration. Distilled water was used as the solvent in all experiments.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec4\" class=\"Section2\"\u003e \u003ch2\u003e2.2. Instrumentation\u003c/h2\u003e \u003cp\u003eStructural analysis of samples was conducted using XRD (Philips PW1140/90). Finely ground and dried samples were mounted on the sample stage and analyzed over a 2θ range of 5\u0026ndash;60\u0026deg; employing Cu-Kα radiation to assess crystallinity. Functional group characterization was carried out using FTIR (Shimadzu FTIR-8400S) with the KBr pellet method, covering a spectral range of 400\u0026ndash;4000 cm⁻\u0026sup1;. In addition, the surface morphology of the materials was examined using SEM equipped with an Advanced Microanalysis system.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec5\" class=\"Section2\"\u003e \u003ch2\u003e2.3. Feedstocks\u003c/h2\u003e \u003cp\u003e \u003cem\u003eTaberau\u003c/em\u003e stems was harvested from the countryside of Lais Lake, Tanjung Sangalang Village, Central Kahayan District, Pulang Pisau Regency, Central Kalimantan, Indonesia. The harvested \u003cem\u003eTaberau\u003c/em\u003e was air-dried for 2 days on the field, and ground until they passed a 60-mesh sieve. The sieved \u003cem\u003eTaberau\u003c/em\u003e was subsequently dried until it reached a moisture content of 10%.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec6\" class=\"Section2\"\u003e \u003ch2\u003e2.4. Organisms and culture maintenances\u003c/h2\u003e \u003cp\u003e \u003cem\u003eC. caperata\u003c/em\u003e was pre-cultured on potato dextrose agar (PDA, 39 g/L) plates at 27\u0026thinsp;\u0026plusmn;\u0026thinsp;2\u0026deg;C for 7 days in the dark. Meanwhile, \u003cem\u003eS. cerevisiae\u003c/em\u003e (yeast) cells maintained on YPG agar medium [\u003cspan citationid=\"CR39\" class=\"CitationRef\"\u003e39\u003c/span\u003e], was obtained from the available stock culture of Institut Pertanian Bogor Culture Collection (IPBCC), Bogor Indonesia. Slant media were incubated at 30\u0026deg;C over a period of 1\u0026ndash;2 days to promote maximal growth.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec7\" class=\"Section2\"\u003e \u003ch2\u003e2.5. Fungal inoculation\u003c/h2\u003e \u003cp\u003eTen agar plugs (5 mm in diameter) from the plate culture were aseptically introduced into 100 mL of glucose\u0026ndash;peptone medium in a 250 mL Erlenmeyer flask to obtain the liquid culture. The glucose-peptone medium was prepared as previously reported by Agnestisia \u003cem\u003eet al.\u003c/em\u003e [\u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e32\u003c/span\u003e]. The cultures were maintained at 27\u0026thinsp;\u0026plusmn;\u0026thinsp;2\u0026deg;C without agitation in dark conditions for 16 days, with 1 mL of veratryl alcohol (100 mM) added after 2 days to promote laccase production. Liquid samples were collected every two days and blended to obtain a homogeneous inoculation. The laccase enzyme activity was then measured.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec8\" class=\"Section2\"\u003e \u003ch2\u003e2.6. Fungal pre-treatment\u003c/h2\u003e \u003cp\u003eFungal pre-treatment assays were performed in triplicate using 250 mL Erlenmeyer flasks. Ten grams of \u003cem\u003eTaberua\u003c/em\u003e biomass were conditioned with a phosphate buffer solution (pH 5) to achieve an initial moisture content of 75%. Moistened substrates underwent sterilization at 121\u0026deg;C for 30 minutes. After cooling, each flask received 5 mL of a homogeneous liquid culture with the highest laccase activity and was subsequently incubated without agitation at 27\u0026thinsp;\u0026plusmn;\u0026thinsp;2\u0026deg;C for 35 days. The flasks were sealed with sterile, non-absorbent cotton to allow air exchange between the atmosphere and the culture. A control, consisting of 2 mL of acetate buffer solution instead of fungal inoculation, was also included for each biomass sample. The biomass was then washed after fungal pre-treatment. The contents of cellulose, hemicellulose-associated carbohydrates, and lignin in raw, control, and fungal-treated feedstocks were quantified via a gravimetric method.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec9\" class=\"Section2\"\u003e \u003ch2\u003e2.7. Enzymatic hydrolysis\u003c/h2\u003e \u003cp\u003eFive grams of cellulase (CAS 9012-54-8, with activity of 20,000 U/g) were added to 500 mL of a 1% (w/v) substrate (raw, control, and fungal-treated feedstocks, respectively) in acetate buffer (pH\u0026thinsp;=\u0026thinsp;4.8, 50 mM) and stirred at 50\u0026deg;C for 10, 30, 50, 70, and 90 min. The hydrolysis solution was then used to estimate reducing sugar concentration.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec10\" class=\"Section2\"\u003e \u003ch2\u003e2.8. Preparation of yeast cell suspension\u003c/h2\u003e \u003cp\u003eTen milliliters of sterile distilled water were added to a 24-hour-old slant culture of \u003cem\u003eS. cerevisiae\u003c/em\u003e, after which the cells were aseptically scraped using a sterile inoculating needle and gently agitated to obtain a uniform suspension. Subsequently, 100 mL of YPG medium (agar-free) were dispensed into 250 mL Erlenmeyer flasks, plugged with cotton, sterilized, and cooled to room temperature. Each flask were inoculated with 1 mL of cell suspension under aseptic conditions and incubated at 30\u0026deg;C with shaking at 150 rpm for 24 h. Cells were harvested by centrifugation (6000 rpm, 15 min), washed with saline, and re-centrifuged for an additional 5 min. The supernatant was removed, and the pellet was washed with saline solution, followed by an additional centrifugation step for 5 minutes. The final pellet was subsequently washed, air-dried, and weighed, and these cells were designated as free cells.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec11\" class=\"Section2\"\u003e \u003ch2\u003e2.9. Immobilization of yeast cells\u003c/h2\u003e \u003cp\u003eMIL-100(Fe) was synthesized in situ at room temperature by mixing an aqueous FeSO₄\u0026middot;7H₂O solution (3.8 g, 13.7 mmol in 120 mL) with an alkaline H₃BTC solution (1.9 g, 9.1 mmol in 30 mL of 1 M NaOH) under stirring (200 rpm). The precursor solution was added dropwise, and the mixture was stirred for 24 h. The resulting solid was washed with warm water and methanol, then dried at 60\u0026deg;C for 12 h. For immobilization, free yeast cells (0.25 g) were combined with MIL-100(Fe) (0.25 g) in acetate buffer (125 mL, 50 mM, pH 4.8). The immobilized cells were recovered by centrifugation (6000 rpm, 15 min), washed three times, air-dried for 24 h, and denoted as MIL-100(Fe)@yeast.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec12\" class=\"Section2\"\u003e \u003ch2\u003e2.10. Ethanol fermentation\u003c/h2\u003e \u003cp\u003eThe ethanol fermentation of the \u003cem\u003eTaberau\u003c/em\u003e hydrolysis solution in this study was conducted using a batch fermentation method. The effects of yeast dosage and fermentation time on ethanol production were investigated. MIL-100(Fe)@yeast and yeast cell free (2% w/v) were inoculated into 500 mL Erlenmeyer flasks containing 100 mL of sterilized liquid medium (pH 5), consisting of 0.15% yeast extract, 0.25% NH₄Cl, 0.55% K₂HPO₄, and 0.025% MgSO₄\u0026middot;7H₂O, along with hydrolysis solution as the carbon source. The MIL-100(Fe)@yeast and yeast cell free systems were incubated at 30\u0026deg;C with agitation at 75 rpm for 4 hours. To assess the impact of yeast dosage and fermentation time, experiments were carried out with different MIL-100(Fe)@yeast and yeast cell free concentrations (0.5%, 1%, 1.5%, 2%, and 2.5% w/v) and fermentation durations (1 to 10 days). After the fermentation process, the MIL-100(Fe)@yeast and yeast cell free cells were separated from the fermented medium and stored for use in subsequent experiments. The fermented medium was then analyzed for ethanol and residual reducing sugar concentration.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec13\" class=\"Section2\"\u003e \u003ch2\u003e2.11. Reusability of immobilized yeast cells\u003c/h2\u003e \u003cp\u003eThe reusability of MIL-100(Fe)@yeast was tested by performing multiple fermentation cycles. After each batch, the MIL-100(Fe)@yeast were recovered from the fermentation medium by filtration or centrifugation. The recovered cells were washed with sterile water to remove residual fermentation by-products and medium components. The washed MIL-100(Fe)@yeast were then reused in subsequent fermentation cycles. The reuse experiments were carried out for up to six consecutive batches. In each cycle, the ethanol yield and glucose consumption were monitored, and the same procedure (as described above) was followed for each batch. Samples were drawn at regular intervals (every 24 hours) to measure ethanol content and sugar consumption.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec14\" class=\"Section2\"\u003e \u003ch2\u003e2.12. Analytical methods\u003c/h2\u003e \u003cp\u003eLaccase activity was assayed via syringaldazine oxidation [\u003cspan citationid=\"CR48\" class=\"CitationRef\"\u003e48\u003c/span\u003e] in a 3 mL system containing sodium tartrate buffer (0.1 M, pH 5.3), syringaldazine (0.5 mM), and enzyme solution. The increase in absorbance at 525 nm (ε525\u0026thinsp;=\u0026thinsp;65,000 M⁻\u0026sup1;\u0026middot;cm⁻\u0026sup1;) was monitored for 2 min using a UV\u0026ndash;Vis spectrophotometer. Furthermore, the concentration of reducing sugars was analyzed by the DNS method [\u003cspan citationid=\"CR49\" class=\"CitationRef\"\u003e49\u003c/span\u003e], with absorbance recorded at 540 nm using a UV\u0026ndash;Vis spectrophotometer and quantified against a glucose standard curve. Ethanol content was determined by acid dichromate oxidation, where ethanol in a known sample mass was oxidized to acetic acid using 0.1 N potassium dichromate in the presence of sulfuric acid. In addition, the physical properties of distilled ethanol from this study consisting of density, viscosity, specific gravity, API specific gravity, and flash point, were also determined. Specific gravity was determined using the ASTM D 792\u0026thinsp;\u0026minus;\u0026thinsp;13 method. Viscosity was analyzed using a Saybolt viscosimeter with the ASTM D 88 test standard. Flash point were determined using a flash and fire point tester with the ASTM D 92 test standard. Specific gravity and API gravity were analyzed using the ASTM D 891-09 and ASTM D 287 test standards, respectively.\u003c/p\u003e \u003c/div\u003e"},{"header":"3. Results","content":"\u003cdiv id=\"Sec16\" class=\"Section2\"\u003e \u003ch2\u003e3.1. Fungal laccase enzymes\u003c/h2\u003e \u003cp\u003eThe fungal inoculation method used in this study successfully facilitated the production of laccase enzymes, with a particular focus on examining the effects of incubation period on laccase activity generated by \u003cem\u003eC. caperata\u003c/em\u003e (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003e). Determining the optimal incubation time is crucial for maximizing laccase production in the shortest time possible. To achieve this, incubation times were varied across a range of 2, 4, 6, 8, 10, 12, 14, and 16 days. This range was selected to assess early metabolic activity and temporal fungal performance, enabling evaluation of laccase production stability and nutrient-related effects. Similar incubation timeframes have been employed in previous studies on laccase production in other fungi, such as \u003cem\u003eGanoderma\u003c/em\u003e sp. [\u003cspan citationid=\"CR50\" class=\"CitationRef\"\u003e50\u003c/span\u003e], \u003cem\u003eTrametes versicolor\u003c/em\u003e [\u003cspan citationid=\"CR51\" class=\"CitationRef\"\u003e51\u003c/span\u003e], and \u003cem\u003eTrametes trogii\u003c/em\u003e [\u003cspan citationid=\"CR52\" class=\"CitationRef\"\u003e52\u003c/span\u003e]. The data obtained from these varied incubation periods will contribute to optimizing laccase production and provide a better understanding of its efficiency over time.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eThe \u003cem\u003eC. caperata\u003c/em\u003e fungal culture appeared to show stable growth, and the addition of 1 mL of 100 mM veratryl alcohol after two days of incubation significantly enhanced laccase production. The enzyme activity increased steadily after the second day, reaching an optimum at 12 days with an enzyme activity of 520 U/L, and then declined towards the end of the incubation period, likely due to nutrient depletion or a reduction in fungal metabolic activity. The laccase enzyme activity of \u003cem\u003eC. caperata\u003c/em\u003e appeared to be lower than that of \u003cem\u003eC. caperata\u003c/em\u003e BM-172 (880 U/L) [\u003cspan citationid=\"CR53\" class=\"CitationRef\"\u003e53\u003c/span\u003e], but is higher compared to other white rot fungi, such as \u003cem\u003eFuscoporia gilva\u003c/em\u003e (440 U/L) and \u003cem\u003ePleurotus tuberregium\u003c/em\u003e (480 U/L) [\u003cspan citationid=\"CR23\" class=\"CitationRef\"\u003e23\u003c/span\u003e]. Therefore, \u003cem\u003eC. caperata\u003c/em\u003e is considered to have the potential to be used as a delignification agent for Taberua biomass in this study.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec17\" class=\"Section2\"\u003e \u003ch2\u003e3.2. Delignification of \u003cem\u003eTaberau\u003c/em\u003e biomass\u003c/h2\u003e \u003cp\u003eFungal pre-treatment assays conducted on \u003cem\u003eTaberua\u003c/em\u003e biomass showed significant delignification, as indicated by the reduction in lignin content following fungal inoculation (Table\u0026nbsp;\u003cspan refid=\"Tab1\" class=\"InternalRef\"\u003e1\u003c/span\u003e). A control group, using acetate buffer instead of the fungal inoculum, was included to isolate the effect of the fungal treatment. The most notable observation in this study was the significant reduction in lignin content in the fungal-treated feedstock (10.92% \u0026plusmn; 1.15%), compared to the raw feedstock (20.87% \u0026plusmn; 0.82%) and the control feedstock (19.35% \u0026plusmn; 0.74%). This reduction in lignin content is consistent with the known role of laccase-producing fungi in delignification processes. Laccases, a group of enzymes produced by certain fungi, have been widely recognized for their ability to oxidize lignin, breaking down its complex aromatic structure into smaller fragments. The marked decrease in lignin content following fungal treatment provides strong evidence that fungal inoculation, particularly through laccase activity, was the primary mechanism responsible for the observed delignification. This delignification process also enhanced the availability of cellulose and holocellulose -associated carbohydrates, making the biomass more accessible for further biochemical conversion processes such as enzymatic hydrolysis [\u003cspan citationid=\"CR54\" class=\"CitationRef\"\u003e54\u003c/span\u003e]. These findings align with previous studies in which fungal treatments using laccase-producing fungi facilitated lignin degradation and improved the accessibility of polysaccharides [\u003cspan citationid=\"CR29\" class=\"CitationRef\"\u003e29\u003c/span\u003e, \u003cspan citationid=\"CR30\" class=\"CitationRef\"\u003e30\u003c/span\u003e, \u003cspan citationid=\"CR54\" class=\"CitationRef\"\u003e54\u003c/span\u003e, \u003cspan citationid=\"CR55\" class=\"CitationRef\"\u003e55\u003c/span\u003e].\u003c/p\u003e \u003cp\u003e \u003cdiv class=\"gridtable\"\u003e\u003ctable float=\"Yes\" id=\"Tab1\" border=\"1\"\u003e \u003ccaption language=\"En\"\u003e \u003cdiv class=\"CaptionNumber\"\u003eTable 1\u003c/div\u003e \u003cdiv class=\"CaptionContent\"\u003e \u003cp\u003eChemical composition of feedstocks\u003c/p\u003e \u003c/div\u003e \u003c/caption\u003e \u003ccolgroup cols=\"4\"\u003e \u003cdiv align=\"left\" class=\"colspec\" colname=\"c1\" colnum=\"1\"\u003e\u003c/div\u003e \u003cdiv align=\"left\" class=\"colspec\" colname=\"c2\" colnum=\"2\"\u003e\u003c/div\u003e \u003cdiv align=\"left\" class=\"colspec\" colname=\"c3\" colnum=\"3\"\u003e\u003c/div\u003e \u003cdiv align=\"left\" class=\"colspec\" colname=\"c4\" colnum=\"4\"\u003e\u003c/div\u003e \u003cthead\u003e \u003ctr\u003e \u003cth align=\"left\" colname=\"c1\"\u003e \u003cp\u003eComposition (%)\u003c/p\u003e \u003c/th\u003e \u003cth align=\"left\" colname=\"c2\"\u003e \u003cp\u003eRaw feedstock\u003c/p\u003e \u003c/th\u003e \u003cth align=\"left\" colname=\"c3\"\u003e \u003cp\u003eControl feedstock\u003c/p\u003e \u003c/th\u003e \u003cth align=\"left\" colname=\"c4\"\u003e \u003cp\u003eFungal-treated feedstock\u003c/p\u003e \u003c/th\u003e \u003c/tr\u003e \u003c/thead\u003e \u003ctbody\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003e\u003cspan class=\"InlineEquation\"\u003e\u003cspan class=\"mathinline\"\u003e\\(\\:\\alpha\\:\\)\u003c/span\u003e\u003c/span\u003e-Cellulose\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c2\"\u003e \u003cp\u003e39.24 \u003cspan class=\"InlineEquation\"\u003e\u003cspan class=\"mathinline\"\u003e\\(\\:\\pm\\:\\)\u003c/span\u003e\u003c/span\u003e 0.47\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c3\"\u003e \u003cp\u003e38.24 \u003cspan class=\"InlineEquation\"\u003e\u003cspan class=\"mathinline\"\u003e\\(\\:\\pm\\:\\)\u003c/span\u003e\u003c/span\u003e 1.67\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c4\"\u003e \u003cp\u003e45.21 \u003cspan class=\"InlineEquation\"\u003e\u003cspan class=\"mathinline\"\u003e\\(\\:\\pm\\:\\)\u003c/span\u003e\u003c/span\u003e 1.04\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003eHolocellulose\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c2\"\u003e \u003cp\u003e56.68 \u003cspan class=\"InlineEquation\"\u003e\u003cspan class=\"mathinline\"\u003e\\(\\:\\pm\\:\\)\u003c/span\u003e\u003c/span\u003e 0.75\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c3\"\u003e \u003cp\u003e59.20 \u003cspan class=\"InlineEquation\"\u003e\u003cspan class=\"mathinline\"\u003e\\(\\:\\pm\\:\\)\u003c/span\u003e\u003c/span\u003e 0.85\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c4\"\u003e \u003cp\u003e60.68 \u003cspan class=\"InlineEquation\"\u003e\u003cspan class=\"mathinline\"\u003e\\(\\:\\pm\\:\\)\u003c/span\u003e\u003c/span\u003e 0.52\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003eLignin\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c2\"\u003e \u003cp\u003e20.87 \u003cspan class=\"InlineEquation\"\u003e\u003cspan class=\"mathinline\"\u003e\\(\\:\\pm\\:\\)\u003c/span\u003e\u003c/span\u003e 0.82\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c3\"\u003e \u003cp\u003e19.35 \u003cspan class=\"InlineEquation\"\u003e\u003cspan class=\"mathinline\"\u003e\\(\\:\\pm\\:\\)\u003c/span\u003e\u003c/span\u003e 0.74\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c4\"\u003e \u003cp\u003e10.92 \u003cspan class=\"InlineEquation\"\u003e\u003cspan class=\"mathinline\"\u003e\\(\\:\\pm\\:\\)\u003c/span\u003e\u003c/span\u003e 1.15\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003c/tbody\u003e \u003c/colgroup\u003e \u003c/table\u003e\u003c/div\u003e \u003c/p\u003e \u003cp\u003eFurthermore, the control group in this study, which used acetate buffer instead of fungal inoculum, did not exhibit significant changes in biomass composition, reinforcing the notion that laccase activity was the key driver behind the observed delignification. This finding is consistent with prior research, which also reported no significant changes in biomass composition in the absence of fungal inoculation, confirming the effectiveness of laccase-producing fungi in biomass pre-treatment [\u003cspan citationid=\"CR54\" class=\"CitationRef\"\u003e54\u003c/span\u003e]. By degrading lignin, the fungal treatment enhances the overall porosity and accessibility of the biomass, thus improving the efficiency of downstream processes like enzymatic hydrolysis, which is crucial for converting the polysaccharides into fermentable sugars for bioethanol production.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec18\" class=\"Section2\"\u003e \u003ch2\u003e3.3. Enzymatic hydrolysis\u003c/h2\u003e \u003cp\u003eEnzymatic hydrolysis was performed to assess the sugar release potential from raw, control, and fungal-treated feedstock. The hydrolysis reaction was carried out at 50\u0026deg;C for various time intervals (10, 30, 50, 70, and 90 minutes). The results presented in Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003e demonstrate that fungal pre-treatment significantly enhances the enzymatic hydrolysis of \u003cem\u003eTaberua\u003c/em\u003e feedstock, as evidenced by the higher sugar release from fungal-treated feedstock compared to raw and control feedstocks. The fungal pre-treatment, which effectively degraded lignin, made the cellulose more accessible to cellulase enzymes, leading to a marked increase in sugar yield after 50 minute of hydrolysis. This finding aligns with several other studies that have shown similar improvements in sugar release following lignin removal through fungal inoculation. For instance, studies by Gupta et al. (2011) demonstrated that laccase-producing fungi can efficiently break down lignin, enhancing the accessibility of cellulose and improving the efficiency of enzymatic hydrolysis [\u003cspan citationid=\"CR56\" class=\"CitationRef\"\u003e56\u003c/span\u003e]. In contrast, the raw and control samples, which retained most of their lignin, exhibited slower sugar release, particularly at earlier time points, confirming the protective role of lignin in impeding cellulase action. This observation is consistent with several researcher that reported lignin removal is crucial for optimizing biomass conversion processes [\u003cspan citationid=\"CR12\" class=\"CitationRef\"\u003e12\u003c/span\u003e, \u003cspan citationid=\"CR24\" class=\"CitationRef\"\u003e24\u003c/span\u003e, \u003cspan citationid=\"CR56\" class=\"CitationRef\"\u003e56\u003c/span\u003e]. Overall, the results emphasize the effectiveness of fungal pre-treatment in improving the efficiency of biomass hydrolysis, offering a promising strategy for enhancing biofuel production and other biotechnological applications.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec19\" class=\"Section2\"\u003e \u003ch2\u003e3.4. MIL-100(Fe) Immobilized yeast cells\u003c/h2\u003e \u003cp\u003eThe synthesis of MIL-100(Fe) and the immobilization of yeast cells onto this material were performed. MIL-100(Fe) was prepared via an in situ method at room temperature [\u003cspan citationid=\"CR57\" class=\"CitationRef\"\u003e57\u003c/span\u003e], after which the obtained material was utilized as a support for yeast immobilization, yielding MIL-100(Fe)@yeast. Both MIL-100(Fe) and MIL-100(Fe)@yeast were subsequently characterized using FTIR, XRD, and SEM to confirm the successful synthesis and immobilization processes. FTIR analysis was conducted to identify functional groups and to verify the presence of characteristic MOF bonds as well as potential interactions between yeast cells and the MIL-100(Fe) framework.\u003c/p\u003e \u003cp\u003eThe immobilization of yeast cells onto the material were successfully carried out using an in situ method at room temperature. This method enables the formation of a stable MOF structure while providing a suitable surface for biological immobilization. MIL-100(Fe) is widely reported as a robust MOF with high surface area and excellent chemical stability, making it highly suitable for hosting biomolecules and microbial cells [\u003cspan citationid=\"CR42\" class=\"CitationRef\"\u003e42\u003c/span\u003e, \u003cspan citationid=\"CR43\" class=\"CitationRef\"\u003e43\u003c/span\u003e, \u003cspan citationid=\"CR58\" class=\"CitationRef\"\u003e58\u003c/span\u003e].\u003c/p\u003e \u003cp\u003eThe FTIR spectra of MIL-100(Fe) and MIL-100(Fe)@yeast, recorded in the range of 400\u0026ndash;4000 cm⁻\u0026sup1;, exhibit very similar spectral patterns (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003e). This indicates that the immobilization process does not significantly affect the structural integrity of the MIL-100(Fe) framework. Characteristic absorption bands such as O\u0026ndash;H stretching (~\u0026thinsp;3400\u0026ndash;3500 cm⁻\u0026sup1;), C\u0026thinsp;=\u0026thinsp;O stretching (~\u0026thinsp;1625\u0026ndash;1635 cm⁻\u0026sup1;), and Fe\u0026ndash;O vibrations (~\u0026thinsp;600\u0026ndash;630 cm⁻\u0026sup1;) remain clearly visible after immobilization. These findings are consistent with previous reports showing that MIL-100(Fe) maintains its structural framework even after chemical modification or interaction with external species [\u003cspan citationid=\"CR59\" class=\"CitationRef\"\u003e59\u003c/span\u003e, \u003cspan citationid=\"CR60\" class=\"CitationRef\"\u003e60\u003c/span\u003e]. The preservation of these functional groups suggests that the coordination between Fe\u0026sup3;⁺ ions and the organic ligands remains intact. According to Han et al. (2017), the stability of MIL-100(Fe) originates from its strong metal\u0026ndash;ligand coordination and mesoporous architecture, which provides resistance against structural collapse during post-synthetic modification [\u003cspan citationid=\"CR60\" class=\"CitationRef\"\u003e60\u003c/span\u003e]. This characteristic is crucial for applications involving biological systems, where mild conditions are typically required.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eIn addition to the characteristic peaks of MIL-100(Fe), the FTIR spectrum of MIL-100(Fe)@yeast shows additional absorption bands that can be attributed to yeast cells in Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003e. The absorption peak at approximately 2926 cm⁻\u0026sup1; corresponds to C\u0026ndash;H stretching vibrations of aliphatic groups, while the band at around 1572 cm⁻\u0026sup1; is associated with amide groups from proteins. These functional groups are typical components of microbial cells, including proteins and lipids, confirming the successful immobilization of yeast onto the MOF surface [\u003cspan citationid=\"CR61\" class=\"CitationRef\"\u003e61\u003c/span\u003e]. Furthermore, the presence of a band near 1047 cm⁻\u0026sup1;, corresponding to C\u0026ndash;O stretching vibrations, indicates the presence of carbohydrates such as polysaccharides in the yeast cell wall. Yeast cell walls are primarily composed of glucans, mannoproteins, and chitin, which contribute to these characteristic FTIR signals [\u003cspan citationid=\"CR61\" class=\"CitationRef\"\u003e61\u003c/span\u003e]. The detection of these functional groups suggests that the yeast cells maintain their structural integrity and biochemical composition after immobilization.\u003c/p\u003e \u003cp\u003eThe coexistence of MIL-100(Fe) characteristic peaks and yeast-related bands indicates that the immobilization process occurs mainly through surface interactions, such as hydrogen bonding and electrostatic interactions, rather than through structural disruption. Similar observations have been reported in previous studies, where MOFs served as effective supports for enzyme and microbial immobilization without compromising their activity [\u003cspan citationid=\"CR42\" class=\"CitationRef\"\u003e42\u003c/span\u003e, \u003cspan citationid=\"CR43\" class=\"CitationRef\"\u003e43\u003c/span\u003e, \u003cspan citationid=\"CR58\" class=\"CitationRef\"\u003e58\u003c/span\u003e, \u003cspan citationid=\"CR62\" class=\"CitationRef\"\u003e62\u003c/span\u003e].\u003c/p\u003e \u003cp\u003eTo further strengthen the identification results obtained from FTIR for MIL-100(Fe) and MIL-100(Fe)@yeast, additional supporting data in the form of XRD analysis is necessary. This data is expected to determine the crystallinity of the MIL-100(Fe) material. It is important to ensure that MIL-100(Fe) was successfully synthesized and retained its structural integrity after the yeast cell immobilization process. The XRD patterns would also reveal any changes in crystallinity due to the incorporation of yeast cells into the framework. The diffraction peaks observed in the XRD patterns of MIL-100(Fe) and MIL-100(Fe)@yeast correspond to the characteristic crystal lattice planes of the analyzed materials (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003e). According to the JCPDS, distinct peaks at 2θ values of 6.08\u0026deg;, 10.22\u0026deg;, and 10.98\u0026deg; are indicative of MIL-100(Fe). These reflections confirm the successful formation of MIL-100(Fe) and the retention of its crystallinity throughout the synthesis process.\u003c/p\u003e \u003cp\u003eA comparison of the diffractogram of MIL-100(Fe) with that of MIL-100(Fe)@yeast reveals that the immobilization of yeast cells on the MIL-100(Fe) surface does not significantly alter the overall crystalline structure of the material. The XRD pattern of MIL-100(Fe)@yeast exhibits almost identical diffraction peaks to those of MIL-100(Fe), suggesting that the framework of the MOF remains intact after the immobilization process. However, a notable difference is the appearance of a new peak at 2θ\u0026thinsp;=\u0026thinsp;20.63\u0026deg;, which is characteristic of yeast. This additional peak is attributed to the specific diffraction pattern of the yeast cells, indicating that they have been successfully incorporated into the MIL-100(Fe) structure. The appearance of this new peak supports the conclusion that yeast cells are effectively immobilized on the MIL-100(Fe) surface, without compromising the material\u0026rsquo;s structural integrity. The persistence of the characteristic MIL-100(Fe) peaks along with the emergence of the yeast-specific peak further validates the stability and robustness of the immobilization process. These findings are consistent with the FTIR results, which also indicated that the yeast cells maintain their functional characteristics after immobilization. As the results, the XRD analysis complements the FTIR data and provides valuable confirmation that the immobilization of yeast cells on MIL-100(Fe) does not disrupt the material\u0026rsquo;s crystalline structure. This dual validation from both FTIR and XRD techniques reinforces the potential of MIL-100(Fe) as a stable and effective platform for future biocatalytic applications, where the integrity of both the support material and the immobilized cells is crucial.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eSEM analysis was further applied to examine the cross-sectional morphology and the distribution of synthesized MIL-100(Fe) (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003e). The synthesized MIL-100(Fe) exhibited an octahedral particle shape, with crystallite sizes ranging from 100 nm to 2 \u0026micro;m, consistent with the sizes reported in the literature. However, the size and shape of the MIL-100(Fe) crystals were not uniform, which is typical for materials synthesized at room temperature. This variation in particle size is acceptable, as the synthesis was carried out under mild conditions. In contrast, when high-temperature synthesis methods are used, more homogeneous MIL-100(Fe) crystals can be produced due to better control over nucleation and crystal growth, resulting in more uniform particle size and shape.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eFurthermore, SEM analysis of MIL-100(Fe)@yeast provides additional insight into the immobilization process. The micrographs show a clear morphological difference between MIL-100(Fe) and MIL-100(Fe)@yeast. The pristine MIL-100(Fe) exhibits irregular agglomerated particles, which is typical for this class of MOFs due to their high surface energy and porous structure. After immobilization, the MIL-100(Fe)@yeast sample shows increased aggregation and the formation of larger clustered structures. This change in morphology indicates the interaction between the MOF particles and the biological component. However, the distinct morphology of yeast cells, which are typically oval-shaped in \u003cem\u003eSaccharomyces cerevisiae\u003c/em\u003e, is not clearly observed in the SEM images. This may be attributed to the coverage or partial encapsulation of yeast cells by MIL-100(Fe) particles, which can obscure their natural morphology. Such phenomena have been widely reported, as microorganisms or enzymes immobilized within porous supports or MOFs often become difficult to visually distinguish due to tight coating, encapsulation within pore structures, or strong host\u0026ndash;guest interfacial interactions that obscure clear morphological boundaries between the biological entities and the supporting material [\u003cspan citationid=\"CR43\" class=\"CitationRef\"\u003e43\u003c/span\u003e, \u003cspan citationid=\"CR63\" class=\"CitationRef\"\u003e63\u003c/span\u003e]. In addition, the cell wall of \u003cem\u003eSaccharomyces cerevisiae\u003c/em\u003e, which is rich in polysaccharides and proteins, can promote adhesion to solid supports through hydrogen bonding and electrostatic interactions, further facilitating the immobilization process [\u003cspan citationid=\"CR61\" class=\"CitationRef\"\u003e61\u003c/span\u003e, \u003cspan citationid=\"CR64\" class=\"CitationRef\"\u003e64\u003c/span\u003e].\u003c/p\u003e \u003cp\u003eImportantly, although individual yeast cells are not distinctly visible, the observed increase in particle size and aggregation strongly suggests successful immobilization. These SEM results are consistent with the FTIR and XRD analyses, which confirm the presence of biological functional groups and the preservation of the MIL-100(Fe) crystalline structure. The combination of these characterization techniques provides complementary evidence that yeast cells have been successfully immobilized onto the MIL-100(Fe) framework without structural degradation.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec20\" class=\"Section2\"\u003e \u003ch2\u003e3.5. Ethanol fermentation\u003c/h2\u003e \u003cp\u003eThe ethanol fermentation of \u003cem\u003eTaberau\u003c/em\u003e hydrolysate using both MIL-100(Fe)@yeast and free yeast systems was evaluated at different yeast dosages. The results clearly demonstrate that yeast dosage significantly affects both sugar consumption and ethanol production (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003e). For both systems, increasing yeast dosage from 0.5% to 2.0% (w/v) resulted in a marked increase in sugar consumption and ethanol production. In the MIL-100(Fe)@yeast system, sugar consumption increased from 16.07 g/L to 27.63 g/L, accompanied by an increase in ethanol concentration from 6.90 g/L to 11.30 g/L. A similar trend was observed in the free yeast system, although at lower levels, where sugar consumption increased from 14.20 g/L to 24.80 g/L, and ethanol production from 6.00 g/L to 10.17 g/L. The maximum ethanol production in both systems was achieved at a yeast dosage of 2%, suggesting an optimal balance between cell concentration and substrate availability. This observation is consistent with previous reports indicating that appropriate yeast loading enhances substrate utilization and ethanol productivity, whereas excessive biomass does not necessarily lead to higher yields [\u003cspan citationid=\"CR65\" class=\"CitationRef\"\u003e65\u003c/span\u003e].\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eFurthermore, the MIL-100(Fe)@yeast system consistently exhibited higher sugar consumption and ethanol production compared to free yeast. This can be attributed to the advantages of cell immobilization, which include improved cell stability, higher cell density, and increased resistance to inhibitory compounds. Immobilized yeast systems have been reported to achieve higher ethanol yields and maintain stable fermentation performance [\u003cspan citationid=\"CR38\" class=\"CitationRef\"\u003e38\u003c/span\u003e, \u003cspan citationid=\"CR64\" class=\"CitationRef\"\u003e64\u003c/span\u003e, \u003cspan citationid=\"CR66\" class=\"CitationRef\"\u003e66\u003c/span\u003e, \u003cspan citationid=\"CR67\" class=\"CitationRef\"\u003e67\u003c/span\u003e]. However, further increasing the yeast dosage to 2.5% did not result in additional improvement. In both systems, sugar consumption decreased slightly, and ethanol production either plateaued or decreased. This indicates the existence of a threshold yeast concentration beyond which fermentation efficiency no longer improves. The decline at higher yeast loading can be attributed to cell overcrowding, competition for nutrients, and accumulation of inhibitory metabolites [\u003cspan citationid=\"CR68\" class=\"CitationRef\"\u003e68\u003c/span\u003e].\u003c/p\u003e \u003cp\u003eFermentation time also plays a critical role in determining ethanol production efficiency. In this study, both sugar consumption and ethanol production increased significantly from day 1 to day 4 in both MIL-100(Fe)@yeast and free yeast systems (Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003e). In the MIL-100(Fe)@yeast system, sugar consumption increased from 16.53 g/L (day 1) to 27.77 g/L (day 4), accompanied by a rise in ethanol concentration from 5.60 g/L to 11.57 g/L. A similar trend was observed for free yeast, although at lower levels, where sugar consumption increased from 13.37 g/L to 24.40 g/L, and ethanol production from 3.50 g/L to 9.83 g/L over the same period. These results indicate that the active fermentation phase occurred within the first four days, during which yeast cells efficiently converted fermentable sugars into ethanol. This pattern is consistent with previous studies reporting that ethanol production typically increases during the exponential growth phase and reaches a maximum before entering the stationary phase [\u003cspan citationid=\"CR65\" class=\"CitationRef\"\u003e65\u003c/span\u003e].\u003c/p\u003e \u003cp\u003eThe highest ethanol production for both systems was observed on day 4, confirming that this duration represents the optimum fermentation time. After day 4, both ethanol production and sugar consumption showed a gradual decline. In the MIL-100(Fe)@yeast system, ethanol decreased from 11.57 g/L (day 4) to 9.63 g/L (day 10), while in the free yeast system it decreased from 9.83 g/L to 5.83 g/L. This decline suggests that most fermentable sugars had been consumed and that inhibitory effects, particularly from accumulated ethanol and metabolic by-products, began to negatively affect yeast activity. Similar observations have been reported in ethanol fermentation studies, where product inhibition and substrate depletion limit further ethanol production [\u003cspan citationid=\"CR68\" class=\"CitationRef\"\u003e68\u003c/span\u003e].\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eCompared to free yeast, the MIL-100(Fe)@yeast system consistently exhibited higher sugar utilization and ethanol production across all fermentation times. Additionally, the decline after the optimum point was less pronounced in the immobilized system, indicating better stability and resistance to inhibitory conditions. This behavior is characteristic of immobilized yeast systems, where the supporting matrix enhances cell protection, maintains higher cell density, and prolongs metabolic activity [\u003cspan citationid=\"CR38\" class=\"CitationRef\"\u003e38\u003c/span\u003e, \u003cspan citationid=\"CR64\" class=\"CitationRef\"\u003e64\u003c/span\u003e, \u003cspan citationid=\"CR66\" class=\"CitationRef\"\u003e66\u003c/span\u003e, \u003cspan citationid=\"CR67\" class=\"CitationRef\"\u003e67\u003c/span\u003e]. Furthermore, immobilization has been widely reported to improve ethanol productivity and operational stability in lignocellulosic fermentation systems. These findings confirm that a fermentation duration of 4 days is optimal under the studied conditions. Extending fermentation beyond this point does not improve ethanol yield and may reduce efficiency due to substrate depletion and ethanol inhibition, as widely discussed in bioethanol production literature [\u003cspan citationid=\"CR38\" class=\"CitationRef\"\u003e38\u003c/span\u003e, \u003cspan citationid=\"CR64\" class=\"CitationRef\"\u003e64\u003c/span\u003e, \u003cspan citationid=\"CR66\" class=\"CitationRef\"\u003e66\u003c/span\u003e, \u003cspan citationid=\"CR67\" class=\"CitationRef\"\u003e67\u003c/span\u003e].\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec21\" class=\"Section2\"\u003e \u003ch2\u003e3.6. Reusability of immobilized yeast cells\u003c/h2\u003e \u003cp\u003eThe reusability of MIL-100(Fe)@yeast over six consecutive fermentation cycles showed a progressive decrease in ethanol yield with each reuse, which is a common phenomenon observed in many immobilized enzyme and cell systems (Fig.\u0026nbsp;\u003cspan refid=\"Fig8\" class=\"InternalRef\"\u003e8\u003c/span\u003e). Our results indicate that ethanol yield decreased after each cycle, likely due to a combination of factors, including the decline in yeast cell viability, the gradual breakdown of the immobilization matrix, and the accumulation of inhibitory byproducts such as ethanol. In other study, the ethanol yield dropped after four cycles, which was attributed to both cellular stress and matrix degradation [\u003cspan citationid=\"CR39\" class=\"CitationRef\"\u003e39\u003c/span\u003e]. Similarly, in our study, the rate of decline in ethanol yield accelerated after the third cycle, suggesting that the yeast cells' metabolic capacity and structural integrity were compromised over time.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eThe results of this study showed a slight decrease in ethanol production after the second reuse cycle of immobilized yeast cells; however, this change was not statistically significant. This trend suggests that the immobilized system maintained relatively stable activity over repeated use, although minor variations may be associated with factors such as gradual loss of cell viability or partial nutrient depletion within the immobilization matrix. Similarly, Onaifo et al. (2016) reported no significant reduction in ethanol yield over successive reuse cycles. Minor differences between the studies may be attributed to variations in experimental conditions, particularly the immobilization strategies employed. For instance, the support material used by Onaifo et al. (2016) may have provided a more protective microenvironment for the yeast cells [\u003cspan citationid=\"CR69\" class=\"CitationRef\"\u003e69\u003c/span\u003e]. Additionally, differences in immobilization parameters, such as cell loading density or entrapment technique, could also contribute to the observed variations. These findings emphasize the importance of optimizing immobilization systems to enhance the stability and reusability of yeast for industrial bioethanol production.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec22\" class=\"Section2\"\u003e \u003ch2\u003e3.7. Ethanol physical characterization\u003c/h2\u003e \u003cp\u003eThe physical properties of the distilled ethanol produced in this study were analyzed, and the results indicate that the ethanol meets key specifications commonly required for industrial applications (Table\u0026nbsp;\u003cspan refid=\"Tab2\" class=\"InternalRef\"\u003e2\u003c/span\u003e). The density of 0.812 g/cm\u0026sup3; is in close agreement with typical values for pure ethanol, which range from 0.789 g/cm\u0026sup3; to 0.792 g/cm\u0026sup3; at 20\u0026deg;C. This suggests that the ethanol produced is of high purity, with minimal impurities that could affect its density. The specific gravity of 0.827 is also consistent with standard ethanol, indicating that the ethanol produced is comparable to commercially available ethanol in terms of its physical composition and suitability for biofuel applications.\u003c/p\u003e \u003cp\u003eThe viscosity of 1.65 mm\u0026sup2;/s is within the range typically observed for ethanol (1.2\u0026ndash;1.5 mm\u0026sup2;/s at 20\u0026deg;C), which is relatively low compared to other liquids like oils or water. This moderate viscosity is advantageous for transportation and handling, as it facilitates pumping and mixing without significant resistance to flow. However, it also suggests that some dissolved impurities or fermentation by-products might slightly increase the viscosity, though it remains within acceptable limits for most industrial processes. The API specific gravity value of 42.90 further confirms the ethanol\u0026rsquo;s suitability for fuel blending, particularly in the production of bioethanol as an alternative fuel. This value, a standard measure of a liquid\u0026rsquo;s density relative to water, positions the ethanol produced in this study within the typical range expected for fuel-grade ethanol, which is essential for its use in the automotive industry and other energy applications. The flash point of 14\u0026deg;C, consistent with the flash point of pure ethanol (around 12\u0026ndash;14\u0026deg;C), highlights the highly flammable nature of the product. This property is crucial for handling and storage, as ethanol\u0026rsquo;s volatility makes it prone to ignition at relatively low temperatures. Therefore, proper safety precautions and storage conditions are required to mitigate risks in large-scale applications\u003c/p\u003e \u003cp\u003e \u003cdiv class=\"gridtable\"\u003e\u003ctable float=\"Yes\" id=\"Tab2\" border=\"1\"\u003e \u003ccaption language=\"En\"\u003e \u003cdiv class=\"CaptionNumber\"\u003eTable 2\u003c/div\u003e \u003cdiv class=\"CaptionContent\"\u003e \u003cp\u003eThe physical properties of the distilled ethanol\u003c/p\u003e \u003c/div\u003e \u003c/caption\u003e \u003ccolgroup cols=\"2\"\u003e \u003cdiv align=\"left\" class=\"colspec\" colname=\"c1\" colnum=\"1\"\u003e\u003c/div\u003e \u003cdiv align=\"char\" char=\".\" class=\"colspec\" colname=\"c2\" colnum=\"2\"\u003e\u003c/div\u003e \u003cthead\u003e \u003ctr\u003e \u003cth align=\"left\" colname=\"c1\"\u003e \u003cp\u003eParameter\u003c/p\u003e \u003c/th\u003e \u003cth align=\"left\" colname=\"c2\"\u003e \u003cp\u003eValue\u003c/p\u003e \u003c/th\u003e \u003c/tr\u003e \u003c/thead\u003e \u003ctbody\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003eDensity (g/cm\u0026sup3;)\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c2\"\u003e \u003cp\u003e0.812\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003eViscosity ((mm\u003csup\u003e2\u003c/sup\u003e/s)\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c2\"\u003e \u003cp\u003e1.65\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003eSpecific gravity\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c2\"\u003e \u003cp\u003e0.827\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003eAPI specific gravity\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c2\"\u003e \u003cp\u003e42.90\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003eFlash point (\u0026deg;C)\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c2\"\u003e \u003cp\u003e14.00\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003c/tbody\u003e \u003c/colgroup\u003e \u003c/table\u003e\u003c/div\u003e \u003c/p\u003e \u003cp\u003eWhen compared to other studies, the ethanol produced in this study shows similar physical properties to those reported in the literature. For instance, [\u003cspan citationid=\"CR69\" class=\"CitationRef\"\u003e69\u003c/span\u003e] reported an ethanol density of 0.79\u0026ndash;0.81 g/cm\u0026sup3;, viscosity between 1.4\u0026ndash;1.7 mm\u0026sup2;/s, and flash points in the range of 12\u0026ndash;15\u0026deg;C, which are all within the values observed in our study. This consistency suggests that the fermentation process in our study was both efficient and effective, producing ethanol with standard characteristics suitable for industrial and biofuel applications. The produced ethanol conforms to established bioethanol standards; however, further optimization of fermentation conditions remains necessary. Although flash point and viscosity are within acceptable limits, fine-tuning parameters such as pH and temperature could enhance purity and reduce residual components affecting ethanol quality. In addition, research into different immobilization techniques for yeast cells or the use of various protective additives could improve the yield and purity of ethanol, ensuring that the physical characteristics remain consistent over extended production cycles.\u003c/p\u003e \u003c/div\u003e"},{"header":"4. Discussion","content":"\u003cp\u003eThe ethanol concentration obtained in this study (11.57 g/L) can be classified as moderate for lignocellulosic bioethanol production using SHF process. This value falls within the commonly reported range of 10\u0026ndash;25 g/L for lignocellulosic substrates, indicating that the overall process performed reasonably well under the applied conditions. However, the yield did not reach higher levels typically associated with optimized or industrial-scale systems (\u0026gt;\u0026thinsp;40 g/L), which may be attributed to the presence of lignin-derived inhibitory compounds and the intrinsic limitations of the SHF process[\u003cspan citationid=\"CR36\" class=\"CitationRef\"\u003e36\u003c/span\u003e, \u003cspan citationid=\"CR65\" class=\"CitationRef\"\u003e65\u003c/span\u003e].\u003c/p\u003e \u003cp\u003eThe moderate ethanol yield obtained in this study is largely influenced by upstream processes, especially fungal delignification and enzymatic hydrolysis.. As shown in Section \u003cspan refid=\"Sec16\" class=\"InternalRef\"\u003e3.1\u003c/span\u003e, laccase production by C. caperata reached its maximum at 12 days, indicating strong oxidative capability toward lignin degradation. This enzymatic activity played a key role in reducing lignin content from 20.87% to 10.92% (Table\u0026nbsp;\u003cspan refid=\"Tab1\" class=\"InternalRef\"\u003e1\u003c/span\u003e), as discussed in Section 3.2. The substantial lignin removal enhanced cellulose accessibility, which was further reflected in the improved sugar release during enzymatic hydrolysis (Section 3.3). The hydrolysis profile (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003e) showed a clear increase in reducing sugar concentration up to the optimum reaction time, confirming that fungal pretreatment effectively disrupted the lignocellulosic matrix. However, despite this improvement, the remaining lignin fraction (~\u0026thinsp;10%) and its derived compounds likely still imposed partial inhibition on both enzymatic and fermentation processes.\u003c/p\u003e \u003cp\u003eIn lignocellulosic systems, residual lignin and its degradation products, such as phenolic compounds, furfural, and hydroxymethylfurfural (HMF), are known to inhibit both enzymatic hydrolysis and microbial fermentation. These compounds can disrupt yeast cell membranes, inhibit key metabolic enzymes, and reduce ethanol productivity [\u003cspan citationid=\"CR70\" class=\"CitationRef\"\u003e70\u003c/span\u003e]. Thus, even though fungal delignification significantly improved biomass accessibility, it may not have completely eliminated inhibitory effects, which explains the moderate ethanol yield obtained. To better contextualize these findings, a comparison with previous studies is presented in Table\u0026nbsp;\u003cspan refid=\"Tab3\" class=\"InternalRef\"\u003e3\u003c/span\u003e.\u003c/p\u003e \u003cp\u003eAs shown in Table\u0026nbsp;\u003cspan refid=\"Tab3\" class=\"InternalRef\"\u003e3\u003c/span\u003e, the ethanol yield obtained in this study (11.57 g/L) falls within the range reported for lignocellulosic bioethanol production using the SHF process. Lower ethanol concentrations, such as 9.45 g/L from rice straw, have been reported for substrates treated with alkali\u0026ndash;enzymatic methods, whereas higher values (30\u0026ndash;31 g/L) were achieved from sweet sorghum bagasse and Douglas-fir following steam pretreatment [\u003cspan citationid=\"CR71\" class=\"CitationRef\"\u003e71\u003c/span\u003e, \u003cspan citationid=\"CR72\" class=\"CitationRef\"\u003e72\u003c/span\u003e]. Intermediate ethanol yields were also observed in cassava stems subjected to microwave-assisted alkali and acid pretreatments (13.66\u0026ndash;17.84 g/L) [\u003cspan citationid=\"CR73\" class=\"CitationRef\"\u003e73\u003c/span\u003e]. These variations indicate that ethanol production is strongly influenced by the type and severity of pretreatment. More intensive methods, such as steam pretreatment and chemical delignification, generally enhance cellulose accessibility and lead to higher ethanol yields, but they may also increase the formation of inhibitory compounds. In contrast, the fungal delignification approach employed in this study represents a milder and more environmentally friendly strategy. This method effectively reduces lignin content while preserving carbohydrate fractions, although it may result in slightly lower ethanol production compared to more aggressive pretreatment techniques. Interestingly, the ethanol yield obtained in this study is comparable to that reported for sugarcane bagasse using Ca-alginate immobilized yeast (11.8 g/L) [\u003cspan citationid=\"CR72\" class=\"CitationRef\"\u003e72\u003c/span\u003e], highlighting that the performance of the MIL-100(Fe)-immobilized yeast system is within the expected range for SHF-based lignocellulosic fermentation.\u003c/p\u003e \u003cp\u003e \u003cdiv class=\"gridtable\"\u003e\u003ctable float=\"Yes\" id=\"Tab3\" border=\"1\"\u003e \u003ccaption language=\"En\"\u003e \u003cdiv class=\"CaptionNumber\"\u003eTable 3\u003c/div\u003e \u003cdiv class=\"CaptionContent\"\u003e \u003cp\u003eComparison of lignocellulosic bioethanol production under different conditions\u003c/p\u003e \u003c/div\u003e \u003c/caption\u003e \u003ccolgroup cols=\"5\"\u003e \u003cdiv align=\"left\" class=\"colspec\" colname=\"c1\" colnum=\"1\"\u003e\u003c/div\u003e \u003cdiv align=\"left\" class=\"colspec\" colname=\"c2\" colnum=\"2\"\u003e\u003c/div\u003e \u003cdiv align=\"left\" class=\"colspec\" colname=\"c3\" colnum=\"3\"\u003e\u003c/div\u003e \u003cdiv align=\"left\" class=\"colspec\" colname=\"c4\" colnum=\"4\"\u003e\u003c/div\u003e \u003cdiv align=\"char\" char=\".\" class=\"colspec\" colname=\"c5\" colnum=\"5\"\u003e\u003c/div\u003e \u003cthead\u003e \u003ctr\u003e \u003cth align=\"left\" colname=\"c1\"\u003e \u003cp\u003eBiomass\u003c/p\u003e \u003c/th\u003e \u003cth align=\"left\" colname=\"c2\"\u003e \u003cp\u003ePretreatment / Delignification\u003c/p\u003e \u003c/th\u003e \u003cth align=\"left\" colname=\"c3\"\u003e \u003cp\u003eYeast System\u003c/p\u003e \u003c/th\u003e \u003cth align=\"left\" colname=\"c4\"\u003e \u003cp\u003eMethod\u003c/p\u003e \u003c/th\u003e \u003cth align=\"left\" colname=\"c5\"\u003e \u003cp\u003eEthanol (g/L)\u003c/p\u003e \u003c/th\u003e \u003c/tr\u003e \u003c/thead\u003e \u003ctbody\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003eRice straw [\u003cspan citationid=\"CR71\" class=\"CitationRef\"\u003e71\u003c/span\u003e]\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c2\"\u003e \u003cp\u003eAlkali\u0026thinsp;+\u0026thinsp;enzymatic treatment\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c3\"\u003e \u003cp\u003eFree yeast\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c4\"\u003e \u003cp\u003eSHF\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c5\"\u003e \u003cp\u003e9.45\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003eSweet sorghum bagasse [\u003cspan citationid=\"CR72\" class=\"CitationRef\"\u003e72\u003c/span\u003e]\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c2\"\u003e \u003cp\u003eSteam pretreatment\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c3\"\u003e \u003cp\u003eFree yeast\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c4\"\u003e \u003cp\u003eSHF\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c5\"\u003e \u003cp\u003e31.20\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003eDouglas-fir [\u003cspan citationid=\"CR72\" class=\"CitationRef\"\u003e72\u003c/span\u003e]\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c2\"\u003e \u003cp\u003eSteam pretreatment\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c3\"\u003e \u003cp\u003eFree yeast\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c4\"\u003e \u003cp\u003eSHF\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c5\"\u003e \u003cp\u003e30.20\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003eCassava stems [\u003cspan citationid=\"CR73\" class=\"CitationRef\"\u003e73\u003c/span\u003e]\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c2\"\u003e \u003cp\u003eMicrowave-assisted alkali (3% NaOH)\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c3\"\u003e \u003cp\u003eFree yeast\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c4\"\u003e \u003cp\u003eSHF\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c5\"\u003e \u003cp\u003e17.84\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003eCassava stems [\u003cspan citationid=\"CR73\" class=\"CitationRef\"\u003e73\u003c/span\u003e]\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c2\"\u003e \u003cp\u003eMicrowave-assisted acid (0.1 M H\u003csub\u003e2\u003c/sub\u003eSO)\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c3\"\u003e \u003cp\u003eFree yeast\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c4\"\u003e \u003cp\u003eSHF\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c5\"\u003e \u003cp\u003e13.66\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003eTaberau stems (this study)\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c2\"\u003e \u003cp\u003eFungal delignification\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c3\"\u003e \u003cp\u003eMIL-100(Fe)-immobilized yeast\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c4\"\u003e \u003cp\u003eSHF\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c5\"\u003e \u003cp\u003e11.57\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003eSugarcane bagasse [\u003cspan citationid=\"CR74\" class=\"CitationRef\"\u003e74\u003c/span\u003e]\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c2\"\u003e \u003cp\u003eMicrowave-assisted alkali (2.75% NaOH)\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c3\"\u003e \u003cp\u003eCa-alginate beads-immobilized yeast\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"left\" colname=\"c4\"\u003e \u003cp\u003eSHF\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c5\"\u003e \u003cp\u003e11.8\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003c/tbody\u003e \u003c/colgroup\u003e \u003c/table\u003e\u003c/div\u003e \u003c/p\u003e \u003cp\u003eThe effect of yeast dosage further highlights the importance of process optimization. Ethanol production increased significantly with increasing yeast concentration up to 2% (w/v), beyond which no substantial improvement was observed. This trend reflects the balance between microbial activity and substrate availability. At optimal conditions, yeast metabolism is maximized, allowing efficient conversion of fermentable sugars produced during hydrolysis. However, excessive yeast loading can lead to mass transfer limitations and increased competition for nutrients [\u003cspan citationid=\"CR68\" class=\"CitationRef\"\u003e68\u003c/span\u003e].\u003c/p\u003e \u003cp\u003eFermentation time also played a decisive role, with maximum ethanol production observed at 4 days. This corresponds to the active metabolic phase of yeast, during which sugar consumption and ethanol formation are at their peak. The decline observed at longer fermentation times is consistent with substrate depletion and ethanol inhibition. Importantly, this trend aligns with the hydrolysis behavior, where sugar availability is time-dependent, indicating that synchronization between hydrolysis and fermentation is critical in SHF systems. A key advantage of this study lies in the application of MIL-100(Fe)-based immobilized yeast, as discussed in Section 3.4. Characterization results (FTIR, XRD, and SEM) confirmed that yeast cells were successfully immobilized without disrupting the structural integrity of the MIL-100(Fe) framework. The presence of functional groups associated with yeast cells, along with the preservation of MOF crystallinity, indicates that immobilization occurred through surface interactions rather than structural modification. This stable immobilization likely contributed to improved fermentation performance by enhancing cell stability, protecting cells from inhibitory compounds, and maintaining a high local cell density.\u003c/p\u003e \u003cp\u003eFurthermore, MIL-100(Fe), with its porous structure and high surface area, supports yeast metabolic activity by enabling efficient substrate diffusion and enhanced mass transfer. These characteristics explain the superior performance of immobilized yeast compared to free yeast systems, as also reported in previous studies [\u003cspan citationid=\"CR38\" class=\"CitationRef\"\u003e38\u003c/span\u003e, \u003cspan citationid=\"CR64\" class=\"CitationRef\"\u003e64\u003c/span\u003e, \u003cspan citationid=\"CR66\" class=\"CitationRef\"\u003e66\u003c/span\u003e, \u003cspan citationid=\"CR67\" class=\"CitationRef\"\u003e67\u003c/span\u003e]. The recycle experiments further demonstrated the practical advantages of immobilization. Ethanol production remained relatively high during the initial cycles and gradually decreased with repeated use. This decline can be attributed to cell leakage, partial deactivation, or structural changes in the immobilization matrix. Nevertheless, the ability to reuse the immobilized system highlights its potential for cost-effective and sustainable bioethanol production.\u003c/p\u003e \u003cp\u003eFinally, the physicochemical properties of the distilled ethanol confirmed the effectiveness of downstream processing. The density (0.812 g/cm\u0026sup3;) and flash point (14\u0026deg;C) are close to those of commercial ethanol fuels, indicating successful enrichment through distillation. However, slight deviations from pure ethanol values suggest the presence of residual water, which is expected in single-stage distillation systems. This highlights the importance of integrating efficient purification techniques, such as fractional distillation or dehydration, to achieve fuel-grade ethanol. In general, this study demonstrates that the integration of fungal delignification, optimized fermentation conditions, and MIL-100(Fe)-based immobilized yeast can produce bioethanol from lignocellulosic biomass with competitive performance. Although the ethanol yield remains moderate, the system offers significant advantages in terms of sustainability, reusability, and process integration, providing a promising platform for further development and scale-up.\u003c/p\u003e \u003cp\u003eFuture work should focus on improving ethanol yield through process optimization, such as enhancing enzymatic hydrolysis efficiency, reducing the formation of inhibitory compounds, and optimizing nutrient supplementation during fermentation. In addition, alternative process configurations beyond SHF are strongly recommended, particularly simultaneous saccharification and fermentation (SSF) or simultaneous saccharification and co-fermentation (SSCF), which can improve overall efficiency by reducing product inhibition and enabling better integration of enzymatic hydrolysis and fermentation steps. These approaches have been widely reported to yield higher ethanol concentrations compared to SHF systems. Furthermore, the application of advanced separation techniques, including multi-stage distillation or dehydration processes, is recommended to obtain higher-purity ethanol. Further studies on the long-term performance of immobilized yeast and the structural integrity of the MIL-100(Fe) matrix are required to enable industrial application.\u003c/p\u003e"},{"header":"5. Conclusions","content":"\u003cp\u003eThis study demonstrates the effective use of \u003cem\u003eC. caperata\u003c/em\u003e for laccase production to facilitate delignification of Taberau (\u003cem\u003ePhragmites australis\u003c/em\u003e) biomass, enhancing its cellulose accessibility for enzymatic hydrolysis and subsequent bioethanol production. The fungal pre-treatment reduced lignin content significantly, which in turn improved sugar release during hydrolysis. Furthermore, yeast cells immobilized on MIL-100(Fe) exhibited enhanced ethanol production compared to free yeast cells, with optimal yields achieved under specific conditions of yeast dosage and fermentation time. The immobilized yeast system retained structural integrity and effectively consumed sugars, demonstrating its potential for sustainable bioethanol production. However, ethanol yield decreased over successive reuse cycles, indicating the necessity for further optimization. The combined use of fungal pre-treatment and immobilized yeast fermentation offers a viable approach to improve lignocellulosic biomass conversion into biofuels for both eco-friendly and industrial applications.\u003c/p\u003e"},{"header":"Declarations","content":"\u003cp\u003e\u003cstrong\u003eAcknowledgements\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe authors would like to express their sincere appreciation to colleagues and collaborators for their valuable support and contributions throughout this study.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eFunding\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThis research was financially supported by the Ministry of Research, Technology, and Higher Education, Indonesia under Grant No. 1000/UN24.13/AL.O4/2024.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAuthor Contributions\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eTAA\u003c/strong\u003e was responsible for the\u0026nbsp;methodology,\u0026nbsp;investigation, formal analysis, data curation, and drafting the original manuscript. \u003cstrong\u003eYY\u0026nbsp;\u003c/strong\u003eand \u003cstrong\u003eYA\u0026nbsp;\u003c/strong\u003econtributed to the investigation, methodology, formal analysis, and data curation. \u003cstrong\u003eWJ\u003c/strong\u003e, \u003cstrong\u003eLT\u003c/strong\u003e, and \u003cstrong\u003eEJK\u003c/strong\u003e involved to the formal analysis, data curation, and preparing visualizations. \u003cstrong\u003eRA\u0026nbsp;\u003c/strong\u003eprovided to the conceptualization, methodology, validation, supervision, and project administration, as well as contributed to the manuscript review and editing. All authors have read and approved the final manuscript.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eData availability\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eAll data are included in the manuscript, and additional data can be available upon request.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eDeclarations of Conflic of interest\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe authors declare that there is no conflict of interest regarding the publication of this manuscript. All authors have contributed significantly to the research and preparation of the manuscript, and no competing financial, professional, or personal interests could have influenced the work or its interpretation.\u003c/p\u003e"},{"header":"References","content":"\u003col\u003e\n\u003cli\u003eHasan, M. H., Mahlia, T. M. I., \u0026amp; Nur, H. (2012). A review on energy scenario and sustainable energy in Indonesia. \u003cem\u003eRenewable and Sustainable Energy Reviews\u003c/em\u003e, \u003cem\u003e16\u003c/em\u003e(4), 2316\u0026ndash;2328. https://doi.org/10.1016/j.rser.2011.12.007\u003c/li\u003e\n\u003cli\u003eUnited Nations Department of Economic and Social Affairs, P. D. (2022). W. P. P. 2022: S. of R., Raimi, D., Zhu, Y., Newell, R. G., Prest, B. C., \u0026amp; Bergman, A. (2022). \u003cem\u003eGlobal Energy Outlook 2023: Sowing the Seeds of an Energy Transition\u003c/em\u003e. \u003cem\u003eUnited Nation\u003c/em\u003e. Retrieved from http://dx.doi.org/10.17605/OSF.IO/W3B4Z%0Awww.un.org/development/ desa/pd/.\u003c/li\u003e\n\u003cli\u003eHoltom, B. (2004). Fossil fuels, alternative energy and economic growth. \u003cem\u003eJournal of Nursing Administration\u003c/em\u003e, \u003cem\u003e34\u003c/em\u003e(5), 216\u0026ndash;227.\u003c/li\u003e\n\u003cli\u003eAbas, N., Kalair, A., \u0026amp; Khan, N. (2015). Review of fossil fuels and future energy technologies. \u003cem\u003eFutures\u003c/em\u003e, \u003cem\u003e69\u003c/em\u003e, 31\u0026ndash;49. https://doi.org/10.1016/j.futures.2015.03.003\u003c/li\u003e\n\u003cli\u003eWang, J., \u0026amp; Azam, W. (2024). Natural resource scarcity, fossil fuel energy consumption, and total greenhouse gas emissions in top emitting countries. \u003cem\u003eGeoscience Frontiers\u003c/em\u003e, \u003cem\u003e15\u003c/em\u003e(2), 2024. https://doi.org/10.1016/j.gsf.2023.101757\u003c/li\u003e\n\u003cli\u003eMerritt, H., \u0026amp; Barrag\u0026aacute;n-Oca\u0026ntilde;a, A. (2023). The impact of market factors on the development of eco-friendly energy technologies: the case of bioethanol. \u003cem\u003eClean Technologies and Environmental Policy\u003c/em\u003e, \u003cem\u003e25\u003c/em\u003e(2), 313\u0026ndash;321. https://doi.org/10.1007/s10098-021-02225-6\u003c/li\u003e\n\u003cli\u003eRipple, W. J., Wolf, C., Gregg, J. W., Rockstr\u0026ouml;m, J., Mann, M. E., Oreskes, N., \u0026hellip; Crowther, T. W. (2024). The 2024 state of the climate report : Perilous times, 1\u0026ndash;13.\u003c/li\u003e\n\u003cli\u003eRajak, R. C., \u0026amp; Banerjee, R. (2018). An eco-friendly process integration for second generation bioethanol production from laccase delignified Kans grass. \u003cem\u003eEnergy Conversion and Management\u003c/em\u003e, \u003cem\u003e157\u003c/em\u003e, 364\u0026ndash;371. https://doi.org/10.1016/j.enconman.2017.11.060\u003c/li\u003e\n\u003cli\u003eLamichhane, G., Acharya, A., Poudel, D. K., Aryal, B., Gyawali, N., Niraula, P., \u0026hellip; Parajuli, N. (2021). Recent advances in bioethanol production from Lignocellulosic biomass. \u003cem\u003eInternational Journal of Green Energy\u003c/em\u003e, \u003cem\u003e18\u003c/em\u003e(7), 731\u0026ndash;744. https://doi.org/10.1080/15435075.2021.1880910\u003c/li\u003e\n\u003cli\u003eKordala, N., Walter, M., Brzozowski, B., \u0026amp; Lewandowska, M. (2024). 2G-biofuel ethanol: an overview of crucial operations, advances and limitations. \u003cem\u003eBiomass Conversion and Biorefinery\u003c/em\u003e, \u003cem\u003e14\u003c/em\u003e(3), 2983\u0026ndash;3006. https://doi.org/10.1007/s13399-022-02861-y\u003c/li\u003e\n\u003cli\u003eLimayem, A., \u0026amp; Ricke, S. C. (2012). Lignocellulosic biomass for bioethanol production: Current perspectives, potential issues and future prospects. \u003cem\u003eProgress in Energy and Combustion Science\u003c/em\u003e, \u003cem\u003e38\u003c/em\u003e(4), 449\u0026ndash;467. https://doi.org/10.1016/j.pecs.2012.03.002\u003c/li\u003e\n\u003cli\u003eDas, N., Jena, P. K., Padhi, D., Kumar Mohanty, M., \u0026amp; Sahoo, G. (2023). A comprehensive review of characterization, pretreatment and its applications on different lignocellulosic biomass for bioethanol production. \u003cem\u003eBiomass Conversion and Biorefinery\u003c/em\u003e, \u003cem\u003e13\u003c/em\u003e(2), 1503\u0026ndash;1527. https://doi.org/10.1007/s13399-021-01294-3\u003c/li\u003e\n\u003cli\u003ePersson, T., Garcia, A. G. Y., Paz, J. O., Fraisse, C. W., \u0026amp; Hoogenboom, G. (2010). Reduction in greenhouse gas emissions due to the use of bio-ethanol from wheat grain and straw produced in the south-eastern USA. \u003cem\u003eJournal of Agricultural Science\u003c/em\u003e, \u003cem\u003e148\u003c/em\u003e(5), 511\u0026ndash;527. https://doi.org/10.1017/S0021859610000316\u003c/li\u003e\n\u003cli\u003eAbdul Kareem Joyia, M., Ahmad, M., Chen, Y. F., Mustaqeem, M., Ali, A., Abbas, A., \u0026amp; Ashraf Gondal, M. (2024). Trends and advances in sustainable bioethanol production technologies from first to fourth generation: A critical review. \u003cem\u003eEnergy Conversion and Management\u003c/em\u003e, \u003cem\u003e321\u003c/em\u003e, 2024. https://doi.org/10.1016/j.enconman.2024.119037\u003c/li\u003e\n\u003cli\u003eCavalaglio, G., Gelosia, M., Ingles, D., Pompili, E., D\u0026rsquo;Antonio, S., \u0026amp; Cotana, F. (2016). Response surface methodology for the optimization of cellulosic ethanol production from Phragmites australis through pre-saccharification and simultaneous saccharification and fermentation. \u003cem\u003eIndustrial Crops and Products\u003c/em\u003e, \u003cem\u003e83\u003c/em\u003e, 431\u0026ndash;437. https://doi.org/10.1016/j.indcrop.2015.12.089\u003c/li\u003e\n\u003cli\u003eCotana, F., Cavalaglio, G., Pisello, A. L., Gelosia, M., Ingles, D., \u0026amp; Pompili, E. (2015). Sustainable ethanol production from common reed (Phragmites australis) through simultaneuos saccharification and fermentation. \u003cem\u003eSustainability (Switzerland)\u003c/em\u003e, \u003cem\u003e7\u003c/em\u003e(9), 12149\u0026ndash;12163. https://doi.org/10.3390/su70912149\u003c/li\u003e\n\u003cli\u003eTozluoglu, A. (2018). Bioethanol production from common reed (Phragmites australis): Enzymatic hydrolysis and fermentation. \u003cem\u003eRomanian Biotechnological Letters\u003c/em\u003e, \u003cem\u003e23\u003c/em\u003e(2), 13473\u0026ndash;13478.\u003c/li\u003e\n\u003cli\u003eK\u0026ouml;bbing, J. F., Thevs, N., \u0026amp; Zerbe, S. (2013). The utilisation of reed ( Phragmites australis ): a review. \u003cem\u003eMires and Peat\u003c/em\u003e, \u003cem\u003e13\u003c/em\u003e, 1\u0026ndash;14.\u003c/li\u003e\n\u003cli\u003eSathitsuksanoh, N., Zhu, Z., Templeton, N., Rollin, J. A., Harvey, S. P., \u0026amp; Zhang, Y. H. P. (2009). Saccharification of a potential bioenergy crop, phragmites australis (Common Reed), by lignocellulose fractionation followed by enzymatic hydrolysis at decreased cellulase loadings. \u003cem\u003eIndustrial and Engineering Chemistry Research\u003c/em\u003e, \u003cem\u003e48\u003c/em\u003e(13), 6441\u0026ndash;6447. https://doi.org/10.1021/ie900291s\u003c/li\u003e\n\u003cli\u003eGao, K., Boiano, S., Marzocchella, A., \u0026amp; Rehmann, L. (2014). Cellulosic butanol production from alkali-pretreated switchgrass (Panicum virgatum) and phragmites (Phragmites australis). \u003cem\u003eBioresource Technology\u003c/em\u003e, \u003cem\u003e174\u003c/em\u003e, 176\u0026ndash;181. https://doi.org/10.1016/j.biortech.2014.09.152\u003c/li\u003e\n\u003cli\u003eElhaak, Mohsen, Hamada, \u0026amp; El-Gebaly. (2015). Biofuel Production From Phragmites Australis (Cav .) and Typha Domingensis (Pers.) Plants of Burullus Lake. \u003cem\u003eJ. Exp. Biol. (Bot.)\u003c/em\u003e, \u003cem\u003e11\u003c/em\u003e(2), 237\u0026ndash;243. Retrieved from http://my.ejmanger.com/ejeb/\u003c/li\u003e\n\u003cli\u003eSuryadi, H., Judono, J. J., Putri, M. R., Eclessia, A. D., Ulhaq, J. M., Agustina, D. N., \u0026amp; Sumiati, T. (2022). Biodelignification of lignocellulose using ligninolytic enzymes from white-rot fungi. \u003cem\u003eHeliyon\u003c/em\u003e, \u003cem\u003e8\u003c/em\u003e(2), e08865. https://doi.org/10.1016/j.heliyon.2022.e08865\u003c/li\u003e\n\u003cli\u003eIsikhuemhen, O. S., Mikiashvili, N. A., Senwo, Z. N., \u0026amp; Ohimain, E. I. (2014). Biodegradation and Sugar Release from Canola Plant Biomass by Selected White Rot Fungi. \u003cem\u003eAdvances in Biological Chemistry\u003c/em\u003e, \u003cem\u003e04\u003c/em\u003e(06), 395\u0026ndash;406. https://doi.org/10.4236/abc.2014.46045\u003c/li\u003e\n\u003cli\u003eDe La Torre, M., Mart\u0026iacute;n-Sampedro, R., Fillat, \u0026Uacute;., Eugenio, M. E., Bl\u0026aacute;nquez, A., Hern\u0026aacute;ndez, M., \u0026hellip; Ibarra, D. (2017). Comparison of the efficiency of bacterial and fungal laccases in delignification and detoxification of steam-pretreated lignocellulosic biomass for bioethanol production. \u003cem\u003eJournal of Industrial Microbiology and Biotechnology\u003c/em\u003e, \u003cem\u003e44\u003c/em\u003e(11), 1561\u0026ndash;1573. https://doi.org/10.1007/s10295-017-1977-1\u003c/li\u003e\n\u003cli\u003ePlácido, J., \u0026amp; Capareda, S. (2015). Ligninolytic enzymes: A biotechnological alternative for bioethanol production. \u003cem\u003eBioresources and Bioprocessing\u003c/em\u003e, \u003cem\u003e2\u003c/em\u003e(1). https://doi.org/10.1186/s40643-015-0049-5\u003c/li\u003e\n\u003cli\u003eLoi, M., Glazunova, O., Fedorova, T., Logrieco, A. F., \u0026amp; Mul\u0026egrave;, G. (2021). Fungal laccases: The forefront of enzymes for sustainability. \u003cem\u003eJournal of Fungi\u003c/em\u003e, \u003cem\u003e7\u003c/em\u003e(12), 1\u0026ndash;25. https://doi.org/10.3390/jof7121048\u003c/li\u003e\n\u003cli\u003eRoth, S., \u0026amp; Spiess, A. C. (2015). Laccases for biorefinery applications: a critical review on challenges and perspectives. \u003cem\u003eBioprocess and Biosystems Engineering\u003c/em\u003e, \u003cem\u003e38\u003c/em\u003e(12), 2285\u0026ndash;2313. https://doi.org/10.1007/s00449-015-1475-7\u003c/li\u003e\n\u003cli\u003eLiong, Y. Y., Halis, R., Lai, O. M., \u0026amp; Mohamed, R. (2012). Conversion of lignocellulosic biomass from grass to bioethanol using materials pretreated with alkali and the white rot fungus, Phanerochaete chrysosporium. \u003cem\u003eBioResources\u003c/em\u003e, \u003cem\u003e7\u003c/em\u003e(4), 5500\u0026ndash;5513. https://doi.org/10.15376/biores.7.4.5500-5513\u003c/li\u003e\n\u003cli\u003eL\u0026oacute;pez-Abelairas, M., \u0026Aacute;lvarez Pall\u0026iacute;n, M., Salvach\u0026uacute;a, D., L\u0026uacute;-Chau, T., Mart\u0026iacute;nez, M. J., \u0026amp; Lema, J. M. (2013). Optimisation of the biological pretreatment of wheat straw with white-rot fungi for ethanol production. \u003cem\u003eBioprocess and Biosystems Engineering\u003c/em\u003e, \u003cem\u003e36\u003c/em\u003e(9), 1251\u0026ndash;1260. https://doi.org/10.1007/s00449-012-0869-z\u003c/li\u003e\n\u003cli\u003eCastoldi, R., Bracht, A., de Morais, G. R., Baesso, M. L., Correa, R. C. G., Peralta, R. A., \u0026hellip; Peralta, R. M. (2014). Biological pretreatment of Eucalyptus grandis sawdust with white-rot fungi: Study of degradation patterns and saccharification kinetics. \u003cem\u003eChemical Engineering Journal\u003c/em\u003e, \u003cem\u003e258\u003c/em\u003e, 240\u0026ndash;246. https://doi.org/10.1016/j.cej.2014.07.090\u003c/li\u003e\n\u003cli\u003eDing, C., Wang, X., \u0026amp; Li, M. (2019). Evaluation of six white-rot fungal pretreatments on corn stover for the production of cellulolytic and ligninolytic enzymes, reducing sugars, and ethanol. \u003cem\u003eApplied Microbiology and Biotechnology\u003c/em\u003e, \u003cem\u003e103\u003c/em\u003e(14), 5641\u0026ndash;5652. https://doi.org/10.1007/s00253-019-09884-y\u003c/li\u003e\n\u003cli\u003eAgnestisia, R., Panjaitan, D., Erdene-ochir, T., \u0026amp; Nion, Y. A. (2024). Decolorization of Rhodamine B by the white rot fungus Coriolopsis caperata isolated from Central Kalimantan Forest. \u003cem\u003eBIO Web of Conferences\u003c/em\u003e, \u003cem\u003e134\u003c/em\u003e, 06011.\u003c/li\u003e\n\u003cli\u003eAgnestisia, R., Karelius, K., Oksal, E., \u0026amp; Nion, Y. A. (2025). Decolorization of textile dyes with crude ligninolytic enzymes from Coriolopsis caperata. \u003cem\u003eJurnal Bioteknologi dan Biosains Indonesia\u003c/em\u003e, \u003cem\u003e12\u003c/em\u003e(1), 104\u0026ndash;118.\u003c/li\u003e\n\u003cli\u003eKallel, F., Neifar, M., Kacem, I., \u0026amp; Ellouz Chaabouni, S. (2024). Optimization of enzymatic hydrolysis and fermentation of bleached garlic straw for bioethanol production. \u003cem\u003eJournal of Material Cycles and Waste Management\u003c/em\u003e, \u003cem\u003e26\u003c/em\u003e(5), 2880\u0026ndash;2895. https://doi.org/10.1007/s10163-024-02016-3\u003c/li\u003e\n\u003cli\u003eAverheim, A., Stagge, S., J\u0026ouml;nsson, L. J., Larsson, S. H., \u0026amp; Thyrel, M. (2024). Separate hydrolysis and fermentation of softwood bark pretreated with 2-naphthol by steam explosion. \u003cem\u003eBiotechnology for Biofuels and Bioproducts\u003c/em\u003e, \u003cem\u003e17\u003c/em\u003e(1), 1\u0026ndash;10. https://doi.org/10.1186/s13068-024-02552-y\u003c/li\u003e\n\u003cli\u003eDrissen, R. E. T., Maas, R. H. W., Tramper, J., \u0026amp; Beeftink, H. H. (2009). Modelling ethanol production from cellulose: Separate hydrolysis and fermentation versus simultaneous saccharification and fermentation. \u003cem\u003eBiocatalysis and Biotransformation\u003c/em\u003e, \u003cem\u003e27\u003c/em\u003e(1), 27\u0026ndash;35. https://doi.org/10.1080/10242420802564358\u003c/li\u003e\n\u003cli\u003eBhadana, B., \u0026amp; Chauhan, M. (2016). Bioethanol Production Using Saccharomyces cerevisiae with Different Perspectives: Substrates, Growth Variables, Inhibitor Reduction and Immobilization. \u003cem\u003eFermentation Technology\u003c/em\u003e, \u003cem\u003e5\u003c/em\u003e(2), 2\u0026ndash;5. https://doi.org/10.4172/2167-7972.1000131\u003c/li\u003e\n\u003cli\u003eKumoro, A. C., Damayanti, A., Bahlawan, Z. A. S., Melina, M., \u0026amp; Puspawati, H. (2021). Bioethanol production from oil palm empty fruit bunches using saccharomyces cerevisiae immobilized on sodium alginate beads. \u003cem\u003ePeriodica Polytechnica Chemical Engineering\u003c/em\u003e, \u003cem\u003e65\u003c/em\u003e(4), 493\u0026ndash;504. https://doi.org/10.3311/PPch.16775\u003c/li\u003e\n\u003cli\u003eMariam, I., Manzoor, K., Ali, S., \u0026amp; Ikram-Ul-Haq. (2009). Enhanced production of ethanol from free and immobilized Saccharomyces cerevisiae under stationary culture. \u003cem\u003ePakistan Journal of Botany\u003c/em\u003e, \u003cem\u003e41\u003c/em\u003e(2), 821\u0026ndash;833.\u003c/li\u003e\n\u003cli\u003eMd Zain, M., Kofli, N. T., \u0026amp; Yahya, A. S. R. S. (2011). Bioethanol Production by Calcium Alginate-Immobilised St1 Yeast System: Effects of Size of Beads, Ratio and Concentration. \u003cem\u003eIIUM Engineering Journal\u003c/em\u003e, \u003cem\u003e12\u003c/em\u003e(4), 11\u0026ndash;19. https://doi.org/10.31436/iiumej.v12i4.159\u003c/li\u003e\n\u003cli\u003eG\u0026ouml;kg\u0026ouml;z, M., \u0026amp; Yiğitoğlu, M. (2011). Immobilization of Saccharomyces cerevisiae on to modified carboxymethylcellulose for production of ethanol. \u003cem\u003eBioprocess and Biosystems Engineering\u003c/em\u003e, \u003cem\u003e34\u003c/em\u003e(7), 849\u0026ndash;857. https://doi.org/10.1007/s00449-011-0535-x\u003c/li\u003e\n\u003cli\u003eLiang, K., Richardson, J. J., Doonan, C. J., Mulet, X., Ju, Y., Cui, J., \u0026hellip; Falcaro, P. (2017). An Enzyme-Coated Metal\u0026ndash;Organic Framework Shell for Synthetically Adaptive Cell Survival. \u003cem\u003eAngewandte Chemie - International Edition\u003c/em\u003e, \u003cem\u003e56\u003c/em\u003e(29), 8510\u0026ndash;8515. https://doi.org/10.1002/anie.201704120\u003c/li\u003e\n\u003cli\u003eSun, H., Li, Y., Yu, S., \u0026amp; Liu, J. (2020). Metal-organic frameworks (MOFs) for biopreservation: From biomacromolecules, living organisms to biological devices. \u003cem\u003eNano Today\u003c/em\u003e, \u003cem\u003e35\u003c/em\u003e, 2024. https://doi.org/10.1016/j.nantod.2020.100985\u003c/li\u003e\n\u003cli\u003eLiang, S., Wu, X. L., Xiong, J., Zong, M. H., \u0026amp; Lou, W. Y. (2020). Metal-organic frameworks as novel matrices for efficient enzyme immobilization: An update review. \u003cem\u003eCoordination Chemistry Reviews\u003c/em\u003e, \u003cem\u003e406\u003c/em\u003e, 213149. https://doi.org/10.1016/j.ccr.2019.213149\u003c/li\u003e\n\u003cli\u003eHu, J., Yuan, X., Wang, C., Shao, X., Yang, B., Abdul Razzaq, A., \u0026hellip; Peng, Y. (2020). Self-Phosphorization of MOF-Armored Microbes for Advanced Energy Storage. \u003cem\u003eSmall\u003c/em\u003e, \u003cem\u003e16\u003c/em\u003e(24), 1\u0026ndash;9. https://doi.org/10.1002/smll.202000755\u003c/li\u003e\n\u003cli\u003eZhong, G., Liu, D., \u0026amp; Zhang, J. (2018). Applications of Porous Metal-Organic Framework MIL-100(M) (M = Cr, Fe, Sc, Al, V). \u003cem\u003eCrystal Growth and Design\u003c/em\u003e, \u003cem\u003e18\u003c/em\u003e(12), 7730\u0026ndash;7744. https://doi.org/10.1021/acs.cgd.8b01353\u003c/li\u003e\n\u003cli\u003eZhang, F., Shi, J., Jin, Y., Fu, Y., Zhong, Y., \u0026amp; Zhu, W. (2015). Facile synthesis of MIL-100(Fe) under HF-free conditions and its application in the acetalization of aldehydes with diols. \u003cem\u003eChemical Engineering Journal\u003c/em\u003e, \u003cem\u003e259\u003c/em\u003e, 183\u0026ndash;190. https://doi.org/10.1016/j.cej.2014.07.119\u003c/li\u003e\n\u003cli\u003eLeonowicz, A., \u0026amp; Gzrywnowicz, K. (1981). Quantitative estimation of laccase forms in some white-rot fungi using syringaldazine as a substrate. \u003cem\u003eEnzyme and Microbial Technology\u003c/em\u003e, \u003cem\u003e3\u003c/em\u003e, 55\u0026ndash;58.\u003c/li\u003e\n\u003cli\u003eGhose, T. K. (1987). Measurement of cellulase activities, \u003cem\u003e59\u003c/em\u003e(2), 257\u0026ndash;268. https://doi.org/doi:10.1351/pac198759020257\u003c/li\u003e\n\u003cli\u003eSivakumar, R., Rajendran, R., Balakumar, C., Tamilvendan, M., \u0026amp; R. Sivakumar, R. Rajendran, C. Balakumar, M. T. (2010). Isolation , Screening and Optimization of Production Medium for Thermostable Laccase Production from Ganoderma sp. \u003cem\u003eInternational Journal of Engineering Science\u003c/em\u003e, \u003cem\u003e2\u003c/em\u003e(12), 7133\u0026ndash;7141. Retrieved from http://www.ijest.info/docs/IJEST10-02-12-241.pdf\u003c/li\u003e\n\u003cli\u003eCouto, S. R., Gund\u0026iacute;n, M., Lorenzo, M., \u0026amp; Sanrom\u0026aacute;n, M. \u0026Aacute;. (2002). Screening of supports and inducers for laccase production by Trametes versicolor in semi-solid-state conditions. \u003cem\u003eProcess Biochemistry\u003c/em\u003e, \u003cem\u003e38\u003c/em\u003e(2), 249\u0026ndash;255. https://doi.org/10.1016/S0032-9592(02)00087-0\u003c/li\u003e\n\u003cli\u003ePatrick, F., Mtui, G., Mshandete, A., \u0026amp; Kivaisi, A. (2010). Tanzania Journal of Science Peroxidase and Laccase in Submerged Cultures of, \u003cem\u003e36\u003c/em\u003e.\u003c/li\u003e\n\u003cli\u003eHadibarata, T., \u0026amp; Yuniarto, A. (2020). Biodegradation of polycyclic aromatic hydrocarbons by high-laccase basidiomycetes fungi isolated from tropical forest of Borneo. \u003cem\u003eBiocatalysis and Agricultural Biotechnology\u003c/em\u003e, \u003cem\u003e28\u003c/em\u003e(July), 101717. https://doi.org/10.1016/j.bcab.2020.101717\u003c/li\u003e\n\u003cli\u003eWan, C., \u0026amp; Li, Y. (2011). Effectiveness of microbial pretreatment by Ceriporiopsis subvermispora on different biomass feedstocks. \u003cem\u003eBioresource Technology\u003c/em\u003e, \u003cem\u003e102\u003c/em\u003e(16), 7507\u0026ndash;7512. https://doi.org/10.1016/j.biortech.2011.05.026\u003c/li\u003e\n\u003cli\u003eBagewadi, Z. K., Mulla, S. I., \u0026amp; Ninnekar, H. Z. (2017). Optimization of laccase production and its application in delignification of biomass. \u003cem\u003eInternational Journal of Recycling of Organic Waste in Agriculture\u003c/em\u003e, \u003cem\u003e6\u003c/em\u003e(4), 351\u0026ndash;365. https://doi.org/10.1007/s40093-017-0184-4\u003c/li\u003e\n\u003cli\u003eGupta, R., Mehta, G., Khasa, Y. P., \u0026amp; Kuhad, R. C. (2011). Fungal delignification of lignocellulosic biomass improves the saccharification of cellulosics. \u003cem\u003eBiodegradation\u003c/em\u003e, \u003cem\u003e22\u003c/em\u003e(4), 797\u0026ndash;804. https://doi.org/10.1007/s10532-010-9404-6\u003c/li\u003e\n\u003cli\u003ePasaribu, M. H., Karelius, K., Ramdhani, E. P., Agnestisia, R., Pereiz, Z., \u0026amp; Toepak, E. P. (2023). Synthesis of Mil-100(Fe)@Fe3O4 Composite using Zircon Mining Magnetic Waste as an Adsorbent for Methylene Blue Dye. \u003cem\u003eBIO Web of Conferences\u003c/em\u003e, \u003cem\u003e70\u003c/em\u003e. https://doi.org/10.1051/bioconf/20237002010\u003c/li\u003e\n\u003cli\u003ePermyakova, A., Kakar, A., Bachir, J., Gkaniatsou, E., Haye, B., Menguy, N., \u0026hellip; Sicard, C. (2022). In Situ Synthesis of a Mesoporous MIL-100 ( Fe ) Bacteria Exoskeleton. \u003cem\u003eACS Materials Letters\u003c/em\u003e, \u003cem\u003e5\u003c/em\u003e(1), 79\u0026ndash;84.\u003c/li\u003e\n\u003cli\u003eCh\u0026aacute;vez, A. M., Rey, A., L\u0026oacute;pez, J., \u0026Aacute;lvarez, P. M., \u0026amp; Beltr\u0026aacute;n, F. J. (2021). Critical aspects of the stability and catalytic activity of MIL-100(Fe) in different advanced oxidation processes. \u003cem\u003eSeparation and Purification Technology\u003c/em\u003e, \u003cem\u003e255\u003c/em\u003e(June 2020), 117660. https://doi.org/10.1016/j.seppur.2020.117660\u003c/li\u003e\n\u003cli\u003eHan, L., Qi, H., Zhang, D., Ye, G., Zhou, W., Hou, C., \u0026hellip; Sun, Y. (2017). A facile and green synthesis of MIL-100(Fe) with high-yield and its catalytic performance. \u003cem\u003eNew Journal of Chemistry\u003c/em\u003e, \u003cem\u003e41\u003c/em\u003e(22), 13504\u0026ndash;13509. https://doi.org/10.1039/c7nj02975f\u003c/li\u003e\n\u003cli\u003eZhang, X., Zhang, X., He, W., Sun, C., Ma, J., Yuan, J., \u0026amp; Du, X. (2013). High-performance mesoporous LiFePO4 from Baker\u0026rsquo;s yeast. \u003cem\u003eColloids and Surfaces B: Biointerfaces\u003c/em\u003e, \u003cem\u003e103\u003c/em\u003e(October 2012), 114\u0026ndash;120. https://doi.org/10.1016/j.colsurfb.2012.10.002\u003c/li\u003e\n\u003cli\u003eLyu, F., Zhang, Y., Zare, R. N., Ge, J., \u0026amp; Liu, Z. (2014). One-pot synthesis of protein-embedded metal-organic frameworks with enhanced biological activities. \u003cem\u003eNano Letters\u003c/em\u003e, \u003cem\u003e14\u003c/em\u003e(10), 5761\u0026ndash;5765. https://doi.org/10.1021/nl5026419\u003c/li\u003e\n\u003cli\u003eYang, W., Liang, W., O\u0026rsquo;Dell, L. A., Toop, H. D., Maddigan, N., Zhang, X., \u0026hellip; Huang, J. (2021). Insights into the Interaction between Immobilized Biocatalysts and Metal-Organic Frameworks: A Case Study of PCN-333. \u003cem\u003eJACS Au\u003c/em\u003e, \u003cem\u003e1\u003c/em\u003e(12), 2172\u0026ndash;2181. https://doi.org/10.1021/jacsau.1c00226\u003c/li\u003e\n\u003cli\u003eMulko, L., Rivarola, C. R., Barbero, C. A., \u0026amp; Acevedo, D. F. (2016). Bioethanol production by reusable Saccharomyces cerevisiae immobilized in a macroporous monolithic hydrogel matrices. \u003cem\u003eJournal of Biotechnology\u003c/em\u003e, \u003cem\u003e233\u003c/em\u003e, 56\u0026ndash;65. https://doi.org/10.1016/j.jbiotec.2016.07.004\u003c/li\u003e\n\u003cli\u003eZabed, H., Sahu, J. N., Suely, A., Boyce, A. N., \u0026amp; Faruq, G. (2017). Bioethanol production from renewable sources: Current perspectives and technological progress. \u003cem\u003eRenewable and Sustainable Energy Reviews\u003c/em\u003e, \u003cem\u003e71\u003c/em\u003e, 475\u0026ndash;501. https://doi.org/10.1016/j.rser.2016.12.076\u003c/li\u003e\n\u003cli\u003eKourkoutas, Y., Bekatorou, A., Banat, I. M., Marchant, R., \u0026amp; Koutinas, A. A. (2004). Immobilization technologies and support materials suitable in alcohol beverages production: A review. \u003cem\u003eFood Microbiology\u003c/em\u003e, \u003cem\u003e21\u003c/em\u003e(4), 377\u0026ndash;397. https://doi.org/10.1016/j.fm.2003.10.005\u003c/li\u003e\n\u003cli\u003eShen, H. Y., Moonjai, N., Verstrepen, K. J., \u0026amp; Delvaux, F. R. (2003). Impact of attachment immobilization on yeast physiology and fermentation performance. \u003cem\u003eJournal of the American Society of Brewing Chemists\u003c/em\u003e, \u003cem\u003e61\u003c/em\u003e(2), 79\u0026ndash;87. https://doi.org/10.1094/asbcj-61-0079\u003c/li\u003e\n\u003cli\u003eBai, F. W., Anderson, W. A., \u0026amp; Moo-Young, M. (2008). Ethanol fermentation technologies from sugar and starch feedstocks. \u003cem\u003eBiotechnology Advances\u003c/em\u003e, \u003cem\u003e26\u003c/em\u003e(1), 89\u0026ndash;105. https://doi.org/10.1016/j.biotechadv.2007.09.002\u003c/li\u003e\n\u003cli\u003eOnaifo, J. O., Ikhuoria, E. U., Omorogbe, S. O., Agbonlahor, O. G., \u0026amp; Ifijen, I. H. (2016). Coconut Pit an Alternative Source of Producing Renewable Material (Bioethanol). \u003cem\u003eJournal of Applied Chemical Science International\u003c/em\u003e, \u003cem\u003e6\u003c/em\u003e(2), 80\u0026ndash;84.\u003c/li\u003e\n\u003cli\u003eJ\u0026ouml;nsson, L. J., \u0026amp; Mart\u0026iacute;n, C. (2016). Pretreatment of lignocellulose: Formation of inhibitory by-products and strategies for minimizing their effects. \u003cem\u003eBioresource Technology\u003c/em\u003e, \u003cem\u003e199\u003c/em\u003e, 103\u0026ndash;112. https://doi.org/10.1016/j.biortech.2015.10.009\u003c/li\u003e\n\u003cli\u003eJin, X., Song, J., \u0026amp; Liu, G. Q. (2020). Bioethanol production from rice straw through an enzymatic route mediated by enzymes developed in-house from Aspergillus fumigatus. \u003cem\u003eEnergy\u003c/em\u003e, \u003cem\u003e190\u003c/em\u003e(xxxx), 116395. https://doi.org/10.1016/j.energy.2019.116395\u003c/li\u003e\n\u003cli\u003eShen, F., Kumar, L., Hu, J., \u0026amp; Saddler, J. N. (2011). Evaluation of hemicellulose removal by xylanase and delignification on SHF and SSF for bioethanol production with steam-pretreated substrates. \u003cem\u003eBioresource Technology\u003c/em\u003e, \u003cem\u003e102\u003c/em\u003e(19), 8945\u0026ndash;8951. https://doi.org/10.1016/j.biortech.2011.07.028\u003c/li\u003e\n\u003cli\u003ePooja, N. S., Sajeev, M. S., Jeeva, M. L., \u0026amp; Padmaja, G. (2018). Bioethanol production from microwave-assisted acid or alkali-pretreated agricultural residues of cassava using separate hydrolysis and fermentation (SHF). \u003cem\u003e3 Biotech\u003c/em\u003e, \u003cem\u003e8\u003c/em\u003e(1), 1\u0026ndash;12. https://doi.org/10.1007/s13205-018-1095-4\u003c/li\u003e\n\u003cli\u003eSingh, A., Sharma, P., Saran, A. K., Singh, N., \u0026amp; Bishnoi, N. R. (2013). Comparative study on ethanol production from pretreated sugarcane bagasse using immobilized Saccharomyces cerevisiae on various matrices. \u003cem\u003eRenewable Energy\u003c/em\u003e, \u003cem\u003e50\u003c/em\u003e, 488\u0026ndash;493. https://doi.org/10.1016/j.renene.2012.07.003\u003c/li\u003e\n\u003c/ol\u003e"}],"fulltextSource":"","fullText":"","funders":[],"hasAdminPriorityOnWorkflow":false,"hasManuscriptDocX":true,"hasOptedInToPreprint":true,"hasPassedJournalQc":"","hasAnyPriority":false,"hideJournal":false,"highlight":"","institution":"","isAcceptedByJournal":false,"isAuthorSuppliedPdf":false,"isDeskRejected":"","isHiddenFromSearch":false,"isInQc":false,"isInWorkflow":false,"isPdf":false,"isPdfUpToDate":true,"isWithdrawnOrRetracted":false,"journal":{"display":true,"email":"[email protected]","identity":"applied-biochemistry-and-biotechnology","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":false,"externalIdentity":"abab","sideBox":"Learn more about [Applied Biochemistry and Biotechnology](https://www.springer.com/journal/12010)","snPcode":"12010","submissionUrl":"https://submission.nature.com/new-submission/12010/3","title":"Applied Biochemistry and Biotechnology","twitterHandle":"","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"em","reportingPortfolio":"Springer Hybrid","inReviewEnabled":true,"inReviewRevisionsEnabled":false},"keywords":"Bioethanol, immobilization, MIL-100(Fe), Phragmites australis","lastPublishedDoi":"10.21203/rs.3.rs-9512742/v1","lastPublishedDoiUrl":"https://doi.org/10.21203/rs.3.rs-9512742/v1","license":{"name":"CC BY 4.0","url":"https://creativecommons.org/licenses/by/4.0/"},"manuscriptAbstract":"\u003cp\u003eThis study explored bioethanol production from \u003cem\u003eTaberau\u003c/em\u003elignocellulosic biomass through fungal delignification, enzymatic hydrolysis, and fermentation using MIL-100(Fe)-immobilized yeast. The fungal pre-treatment with \u003cem\u003eCoriolopsis caperata\u003c/em\u003eenhanced laccase activity, reaching a maximum of 520 U/L at 12 days, and significantly reduced lignin content from 20.87% to 10.92%. This reduction improved cellulose accessibility and increased the availability of fermentable sugars during hydrolysis. Enzymatic hydrolysis results showed that fungal-treated biomass produced higher reducing sugar concentrations compared to untreated and control samples, indicating effective lignin removal. Fermentation was conducted under various conditions, with optimal ethanol production achieved at a yeast dosage of 2% (w/v) and a fermentation time of 4 days, resulting in an ethanol concentration of 11.57 g/L. Characterization using FTIR, XRD, and SEM confirmed successful immobilization of yeast on MIL-100(Fe) without structural degradation. The immobilized system demonstrated improved stability and reusability compared to free yeast. The obtained ethanol concentration is within the typical range for SHF-based lignocellulosic bioethanol production. The integration of fungal pre-treatment with MOF-based immobilization provides a promising and sustainable strategy, with potential for further improvement through process optimization and alternative fermentation approaches.\u003c/p\u003e","manuscriptTitle":"Green Biorefinery Approach for Taberau (Phragmites australis) Biomass: Delignification by Coriolopsis caperata and Bioethanol Production via MIL-100(Fe)–Immobilized Saccharomyces cerevisiae","msid":"","msnumber":"","nonDraftVersions":[{"code":1,"date":"2026-05-14 15:34:14","doi":"10.21203/rs.3.rs-9512742/v1","editorialEvents":[{"type":"communityComments","content":0},{"type":"reviewerAgreed","content":"","date":"2026-05-06T16:51:13+00:00","index":0,"fulltext":""},{"type":"reviewersInvited","content":"","date":"2026-05-06T05:04:07+00:00","index":"","fulltext":""},{"type":"editorInvited","content":"Applied Biochemistry and Biotechnology","date":"2026-04-27T20:13:52+00:00","index":"","fulltext":""},{"type":"submitted","content":"Applied Biochemistry and Biotechnology","date":"2026-04-26T00:08:59+00:00","index":"","fulltext":""}],"status":"published","journal":{"display":true,"email":"[email protected]","identity":"applied-biochemistry-and-biotechnology","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":false,"externalIdentity":"abab","sideBox":"Learn more about [Applied Biochemistry and Biotechnology](https://www.springer.com/journal/12010)","snPcode":"12010","submissionUrl":"https://submission.nature.com/new-submission/12010/3","title":"Applied Biochemistry and Biotechnology","twitterHandle":"","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"em","reportingPortfolio":"Springer Hybrid","inReviewEnabled":true,"inReviewRevisionsEnabled":false}}],"origin":"","ownerIdentity":"39b5d08e-5342-43fe-adda-74a5d7f0e4ea","owner":[],"postedDate":"May 14th, 2026","published":true,"recentEditorialEvents":[{"type":"reviewerAgreed","content":"","date":"2026-05-06T16:51:13+00:00","index":0,"fulltext":""},{"type":"reviewersInvited","content":"","date":"2026-05-06T05:04:07+00:00","index":"","fulltext":""}],"rejectedJournal":[],"revision":"","amendment":"","status":"under-review","subjectAreas":[],"tags":[],"updatedAt":"2026-05-14T15:34:14+00:00","versionOfRecord":[],"versionCreatedAt":"2026-05-14 15:34:14","video":"","vorDoi":"","vorDoiUrl":"","workflowStages":[]},"version":"v1","identity":"rs-9512742","journalConfig":"researchsquare"},"__N_SSP":true},"page":"/article/[identity]/[[...version]]","query":{"redirect":"/article/rs-9512742","identity":"rs-9512742","version":["v1"]},"buildId":"XKTyCvWXoU3ODBz1xrDgd","isFallback":false,"isExperimentalCompile":false,"dynamicIds":[84888],"gssp":true,"scriptLoader":[]}