Biohydrogen and Biobutanol Production from Spent Coffee and Tea Waste Using Clostridium beijerinckii

Abstract

The growing advocacy for greener climates, coupled with increasing global energy demand driven by urbanization and population growth, highlights the need for sustainable solutions. Repurposing food wastes as substrates offers a promising approach to enhancing cleaner energy generation and promoting a circular economy. This study investigated the potential of spent coffee grounds (SC) and biosolids cake (BS) from tea wastes as substrates for producing valuable fuels and chemicals through acetone–ethanol–butanol (ABE) fermentation. Clostridium beijerinckii NCIMB 8052 was used to ferment 100% and 50% hydrolysates derived from Parr-treated enzyme-hydrolyzed (PEH, PEH50), Parr-treated non-hydrolyzed (PNEH, PNEH50), and non-Parr-treated hydrolyzed (NPEH) SC wastes, as well as enzyme-hydrolyzed (BSH, BSH50) and non-hydrolyzed BS wastes (NBH, NBH50). Fermentation of unmodified hydrolysates by C. beijerinckii was poor. Following CaCO₃ modification of SC and BS hydrolysates, ABE titer, yield, and productivity increased, with the highest values obtained with PEH50 and NBH. Specifically, CaCO₃ modification of SC hydrolysates led to increased butanol titer, yield, and productivity in PEH50, while the NBH exhibited higher butanol yield and productivity than the non-CaCO₃-modified hydrolysates. Additionally, H₂ gas production with PEH50 and NBH was 1.41- and 1.13-fold higher, respectively, than in other hydrolysates. These findings suggest that SC and BS hydrolysates can be valorized to butanol and hydrogen gas and, thereby, can contribute to global food wastes management, energy sustainability, and cost-effective biofuel production.

1. Introduction

The continuous depletion of fossil fuel reserves and their environmental impacts have driven the shift towards sustainable and renewable energy sources. Among the several alternatives in this transition, biobutanol and biohydrogen have gained significant recognition due to their high energy efficiency and environmental compatibility [1]. Biohydrogen generates only water as a by-product when combusted, making it a clean energy source and a key player in reducing carbon emissions [2]. Biobutanol, a four-carbon alcohol, has attracted considerable attention as adaptable biofuel due to its high energy content, lower water solubility compared to ethanol, and compatibility with gasoline engines, making it suitable for real-world applications [3]. Therefore, these biofuels could play a crucial role in addressing the challenges of energy security and environmental sustainability.

Moreover, the growing emphasis on waste valorization within the framework of a circular economy has driven the utilization of agro-industrial residues as feedstocks for biofuel production. Notably, an innovative approach to biofuel production involves using food and agricultural wastes as substrate. Spent coffee grounds (SC) and the sludge from tea leaves processing plants (biosolids) are two abundantly generated waste streams that often end up in landfills despite their rich organic constituents. Coffee is the third-most consumed beverage globally and the second most valuable commodity produced by developing countries after petroleum [4]. Coffee generates a substantial amount of waste during roasting and brewing, approximately 6 million tons annually [5]. The major coffee residue from the coffee brewing industry is the SC; although rich in carbohydrates (5000 Kcal/Kg of calorific value), proteins, lignin, cellulose, and hemicellulose [6,7], they are not utilized or potentially considered for use as feedstock for industrial processes. Furthermore, tea (Camellia sinensis L.) is the second most consumed beverage worldwide, after water [8]. Spent tea leaves sludge, the solid residues (biosolids) from the black tea brewing process, account for about 90% of total tea waste [9]. Between 1995 and 2015, global tea consumption increased 2.1-fold, with a projected 7.5% increase by 2027 [8]. The growing global demand for tea necessitates an effective waste management plan for tea industry waste to mitigate their potential environmental and climatic impacts. Biosolids cake waste (BS) is rich in proteins and amino acids (18–35%), fiber, saponins, steroids, tannins, phenols, calcium, potassium, magnesium, phosphorus, iron, sodium, chromium, nickel, and manganese [10]. Therefore, utilizing SC and BS for biofuel production presents an opportunity for value addition, transforming the waste disposal challenge into an economically and environmentally sustainable solution.

Despite the rich organic constituents of lignocellulosic food wastes, the recalcitrant nature of lignocellulose to deconstruction necessitates pretreatment processes prior to fermentation to hydrogen and butanol. The disruption of the lignin matrix in the lignocellulosic biomass (LB) and the depolymerization of its hemicellulose component to release fermentable sugars also generate lignocellulosic derived microbial inhibitory compounds (LDMICs), which can negatively impact the growth of fermenting microorganisms [11]. Researchers have utilized hydrothermal, acid, alkaline, and steam pretreatment processes to disrupt the LB structure and facilitate the release of fermentable sugars from biomass [12]. Efforts to mitigate these barriers have included modifications of pretreatment processes and fermentation media, enzymatic hydrolysis, and pH adjustment [11,13]. The choice of using Clostridium beijerinckii NCIMB 8052 (hereafter referred as C. beijerinckii) in this study was based on its ability to convert polysaccharides (starch and pectin) and lignocellulose derived sugars to high butanol to acetone ratio [11]. Additionally, C. beijerinckii contains no plasmid and appears to be more stable than some other butanol producing species that contains plasmid. Therefore, we hypothesize that the pretreatment and modification of SC- and BS-based P2 media would enhance the metabolic efficiency of C. beijerinckii during fermentation, leading to improved growth and the production of hydrogen (H₂) and acetone–butanol–ethanol (ABE). Therefore, this study investigated the production of hydrogen gas and butanol by C. beijerinckii grown on P2 media with SC and BS wastes as carbon sources. By tweaking the composition of the fermentation medium when food wastes are used as a carbon source, we aim to maximize ABE production and develop a scalable process to repurpose everyday food wastes. The goal is to advance renewable and sustainable energy systems while improving waste management by valorizing resources that are often incinerated or discarded in landfills. Findings from this study reveal enhanced bioconversion of food wastes into cleaner and renewable fuels and chemicals. Our study significantly contributes to the goal of improved waste management by transforming food wastes into value-added products.

2. Materials and Methods

2.1. Materials

Spent coffee and biosolids cake wastes obtained from J.M. Smucker Co. (Orrville, OH, USA) and Niagara Bottling LLC (Diamond Bar, CA, USA), respectively, were stored at 4 °C and were warmed to room temperature prior to analysis. C. beijerinckii NCIMB 8052 purchased from the American Type Culture Collection (Manassas, VA, USA) was stored in the refrigerator as spores at 4 °C. The spores were retrieved from the fridge where they had been stored as part of the culture collection managed by the BioEnergy Research group, Ohio State University, Wooster, OH, USA. Enzymes such as cellulase, viscozymes, and pectinase were purchased from Sigma Aldrich, (St. Louis, MO, USA).

2.2. Food Wastes Characterization

The moisture content of SC and BS was determined according to the total moisture-reference method [14]. SC was dried for 24 h in a hot air oven at 50 ± 2 °C to prevent sugar caramelization until a constant weight was achieved. Samples were pulverized using mortar and pestle, and the total solid was determined as described [15]. The elemental nitrogen and carbon content of SC and BS was determined using a Costech ECS 4010 C/N/S elemental analyzer (Costech Analytical Technologies Inc., Valencia, CA, USA), and the pH was determined using an AB15 pH Meter (Accumet Basic, Fisher Scientific, Singapore). The ash content of waste samples was determined following a modified protocol of the Test Methods for the Examination of Composting and Compost; 25 (TMECC 03.09-A) [16]. Pre-weighed samples of dried food waste were burned in a Barnstead Thermolyne 30400-Series Furnace (Model: F30428C-80) under excess air at 550 °C for up to 3 h and then cooled in a desiccator at room temperature. Afterwards, the weight of the residue (ash) was used to calculate the percentage of ash content (dry weight, w/w) in the dried food waste samples [15].

The calorific value of food waste was determined using a bomb calorimeter (Model: C 2000 Basic version 1, IKA-Werke GmbH & Co. (Staufen im Breisgau, Baden-Württemberg, Germany). Each food waste sample (200 g) was first decomposed in a decomposition unit (Model: C 5010) and was burned in the presence of oxygen gas. The higher heating value (HHV) obtained from the total heat liberated during total combustion and the heat of condensation produced from the water vapor were used to calculate the calorific value of wastes. The calorific value of samples was calculated by dividing the weight of the initial sample by the quantity of heat released during complete combustion. Samples were analyzed in triplicate to ensure reliability of data obtained. Elemental mineral composition of the food wastes was determined using Inductively Coupled Plasma Optical Emission Spectroscopy (ICP-OES) following the method described in TMECC’s 04.05, 04.06, and 04.07 protocol [15]. The ICP-OES spectrometer Prodigy High Dispersion ICP-OES (Teledyne Leeman Labs, Hudson, NH, USA) equipped with dual view, concentric nebulizer, and cyclonic spray chamber was used for the elemental mineral analysis. Argon gas (99.999% purity) was used as a carrier gas and to uphold plasma. Before injection into the ICP-OES, 7 mL of HNO₃ was added to the dried food waste samples, microwave digested, and allowed to cool at room temperature. The relationship between the distinct emission spectrum of individual elements and their concentrations in samples enabled the quantification of elements in samples on a dry weight basis, expressed in milligrams per gram of the sample.

2.3. Pretreatment and Enzymatic Hydrolysis of Food Wastes

Spent coffee grounds were dried at 50 °C for 12 h and milled using a regular kitchenware blender. SC residue was sieved to a particle size of 2 mm. The pretreatment of SC was conducted using the hydrothermal method of Zhang and Ezeji [11], with slight modification. About 95 g of SC with a moisture content of 27.86% was homogenized with 393 mL of distilled water to achieve 15% solid loading. The volume of water added to the SC waste was calculated using the formula described by Zhang and Ezeji [11]. Pretreatment of SC food waste was performed at 180 °C and a hold time of 60 min in a 1 L Parr reactor (Par 4581 HT, Moline, IL, USA). Subsequently, the pretreated SC slurry was adjusted to pH 5 using 1 M NH₄OH and was hydrolyzed using the enzyme cocktail comprising cellulase (15 FPU/g cellulose) and viscozyme (2.4 g/100 mL of SC hydrolysates), as described [11,13]. The enzyme hydrolysis of pretreated SC slurry was performed for 48 h at 80 rpm, and 50 °C in a shaker water bath (Gyrotory Waster bath shaker, New Brunswick Scientific Co., Inc., Edison, NJ, USA). Afterwards, the hydrolysate was vacuum filtered using a 0.45 µm polyethersulphone membrane filter (Nalgene^TM Rapid-flow^TM sterile filter unit).

Then, to two parts of the biosolids cake (300 g), one part of distilled water (150 mL) was added in a beaker (1 L) and stirred on moderate heat for homogenization. The resulting slurry was heated on a heating mantle at 70 °C for 10 min with constant stirring to achieve even homogenization and then was cooled to 40 °C. The pH of the biosolids slurry was adjusted to pH 6 with 1M HCL, and hydrolysis was performed through the action of pectinase enzyme (Sigma Aldrich, USA) at 40 units/g of biosolids waste. An Erlenmeyer flask (250 mL) containing BS slurry (150 mL) was secured on a G76 Gyrotory water bath shaker (New Brunswick Scientific Co., Inc., Edison, NJ, USA) and was operated for 24 h at 50 °C and 500 rpm to achieve enzyme hydrolysis.

2.4. Bacteria and Inoculum Preparation

Tryptone Glucose Yeast extract (TGY) broth containing tryptone (30 g/L), glucose (20 g/L), yeast extract (10 g/L), and L-cysteine (1.0 g/L) was used to propagate the pre-culture before use in the fermentation process. An aliquot (400 µL) of C. beijerinckii spores was heat shocked at 75 °C for 3 min (Fischer Brand Dry Bath, Waltham, MA, USA) and cooled on ice for 5 min in sterile Eppendorf tubes (1.5 mL). The heat shocked C. beijerinckii spore suspension was transferred into 10 mL of anoxic TGY medium [13] in an anaerobic chamber (Coy, Ann Arbor, MI, USA) operated with a modified atmosphere (82% N₂, 3% H₂, 15% CO₂). C. beijerinckii was incubated for 12 h at 35 ± 1 °C until an OD_600nm of 0.9–1.1 was achieved. Thereafter, the reactivated C. beijerinckii culture (2 mL) was inoculated into 18 mL of fresh anoxic and sterile TGY medium and was incubated anaerobically at 35 ± 1 °C until an OD_600nm of 0.9–1.1 was achieved in approximately 3 to 4 h.

2.5. C. beijerinckii Fermentation of Spent Coffee and Biosolids Hydrolysates

2.5.1. ABE Fermentation of Spent Coffee and Biosolids Hydrolysates

Fermentation was performed in 150 mL screw-capped Pyrex glass bottles as described [13] with minor modification. The effect of pretreatment, enzyme hydrolysis, and SC and BS concentration (50% and 100%) on the fermentation of waste hydrolysates to ABE by C. beijerinckii fermentation was investigated. Parr-treated enzyme-hydrolyzed (PEH and PEH50) and non-enzyme-hydrolyzed spent coffee hydrolysates (PNEH and PNEH50), and non-parr-treated and enzyme-hydrolyzed coffee (NPEH) were used in fermentation at 100% (PEH, PNEH, and NPEH) and 50% (PEH50 and PNEH50) concentrations. Likewise, enzyme-hydrolyzed and non-enzyme-hydrolyzed biosolids were used for fermentation at 100% (BSH and NBH) and 50% (BSH50 and NBH50) concentrations, while glucose medium (60 g/L glucose) was used as control. Food waste hydrolysates and the glucose medium (control) were similarly supplemented with 1.5 g/L yeast extract.

The pH of the SC and BS hydrolysates was adjusted to 6.0–6.5 [11,13] with 1 M NH₄OH, and 1 M HCL, respectively. The hydrolysates and 60 g/L glucose (control) were supplemented with 1.5 g/L yeast extract followed by sterilization in an autoclave at set condition (121 °C, 15 PSI for 15 min). After cooling to 45 °C, they were transferred and stored in the anaerobic chamber for 12 h to remove residual oxygen before use in fermentation. After 24 h, aliquots (1%) of filter sterilized P2 buffer (KH₂PO₄: 50 g/L; K₂HPO₄: 50 g/L; and ammonium acetate: 220 g/L), P2 mineral (MgSO₄·7H₂O: 20 g/L; MnSO₄·H₂O; FeSO₄·7H₂O, and NaCl; 1 g/L each), and P2 vitamin stock (p-aminobenzoic acid: 0.1 g/L, thiamine: 0.1 g/L, and 0.001 g/L biotin) solutions were added to the P2 fermentation medium. Notably, the ammonium acetate composition of the standard P2 buffer was adjusted to 50 g/L before use in the fermentation of SC and BS hydrolysates. The P2 medium containing glucose, SC or BS hydrolysates as carbon source (91 mL) was inoculated with C. beijerinckii at 6% v/v [11,13] of the total fermentation volume (100 mL). A 6% v/v inoculum (preculture) is typically used in ABE fermentation because this concentration provides enough fermenting microorganisms to the production medium that ensures maximum ABE production without introducing excess nutrients from the preculture. Fermentation was conducted in triplicate in the anaerobic chamber (Coy, Ann Arbor, MI, USA) operated with a modified atmosphere (82% N₂, 3% H₂, 15% CO₂) at 35 ± 1 °C. About 1.5 mL samples were collected every 12 h throughout the fermentation period.

2.5.2. Fermentation of Modified Spent Coffee and Biosolids Hydrolysates

Chemical constituents of biomass, type of biomass, concentration of inhibitory compounds in substrates, and the type of fermenting microbial strains are critical factors that influence biobutanol yield, productivity, and fermentation efficiency [17]. Notably, C. beijerinckii undergoes efficient ABE fermentation when glucose or other sugars are not limiting, which is defined as a sugar concentration above 50 g/L. Therefore, to enhance C. beijerinckii fermentation of SC and BS hydrolysates, their sugar concentration was adjusted to 60 g/L by supplementing the hydrolysates with additional glucose [13]. The pH was adjusted to 6.5 with 1 M NH₄OH or 1 M HCl as needed. The food waste hydrolysates (SC and BS) were sterilized in an autoclave operated at 121 °C, 15 psi for 10 min. After cooling to 45 °C, sterile hydrolysate was rendered anoxic by placing it in the anaerobic chamber for 6 h. Subsequently, the anoxic, sterile hydrolysate was supplemented with filter-sterilized 1% P2 vitamins, minerals, and buffer before being inoculated with 6% of C. beijerinckii pre-culture, followed by incubation as detailed in Section 2.5.1.

2.5.3. Effect of CaCO₃ Supplementation of Spent Coffee and Biosolids Hydrolysates

Supplementation of P2 medium and lignocellulosic biomass hydrolysate-based P2 medium with CaCO₃ has been shown to enhance the buffering capacity of solventogenic Clostridium, improve sugar utilization, increase butanol titer and tolerance, and enhance overall product yield and productivity during ABE fermentation [11]. To further improve the fermentation of SC and BS hydrolysates by C. beijerinckii, the hydrolysates were supplemented with 2 g/L CaCO₃ prior to fermentation. The impact of CaCO₃ (2 g/L) supplementation of 50% and 100% pre-treated and hydrolyzed SC and BS hydrolysates on microbial growth, ABE titer, productivity, yield, and biohydrogen production was evaluated. ABE fermentation of SC and BS hydrolysates supplemented with CaCO₃ was carried out following the method described in Section 2.5.1. Briefly, the sugar content of BS and SC hydrolysates was adjusted to 60 g/L glucose, followed by the addition of 1.5 g/L yeast extract and 2 g/L CaCO₃. They were sterilized in an autoclave, cooled, and made anoxic in an anerobic chamber. Prior to inoculation with C. beijerinckii pre-culture at 6% (v/v) of the total fermentation volume (100 mL), 1 mL each of P2 buffer, minerals, and vitamins stock solutions (Section 2.5.1) were added to 91 mL sterile and anoxic-modified P2 medium. The P2 medium was then inoculated with C. beijerinckii (6% v/v) and fermented in the anaerobic chamber (Coy, Ann Arbor, MI, USA) maintained under a modified atmosphere (82% N₂, 3% H₂, 15% CO₂) at 35 ± 1 °C. Fermentation samples were collected every 12 h for analyses.

2.6. Fermentation and Gas Analysis

Gases such as hydrogen (H₂) and carbon dioxide (CO₂) are byproducts of ABE fermentation [18]. To quantitatively assess H₂ and CO₂ generated during the fermentation of SC and BS hydrolysates by C. beijerinckii, fermentation was conducted as described in Section 2.5.1. Sterile and anoxic-modified SC and BS hydrolysates (70 mL aliquots) were dispensed into 150 mL serum bottles and sealed with butyl rubber stoppers and aluminum crimps. The serum bottles containing the modified substrate were sterilized in the autoclave at 121 °C for 15 min. While still hot, the pressure within the serum bottles was brought down to atmospheric pressure by inserting a sterile double-sided vacutainer needle into the rubber stopper to release the air. The degassed serum bottles were then kept in the anaerobic chamber operated with modified gases (82% N₂, 3% H₂, and 15% CO₂) for 6 h. Aliquots (0.7 mL) of filter-sterilized P2 buffer, vitamins, and mineral stock solutions were added to the fermentation medium using a sterile syringe and needle. Foil gas bags (Cel Scientific Corp. Cerritos, CA, USA) of 0.5 L were attached to the serum bottles using a two-way vacutainer needle. Fermentation was carried out in an incubator at 35 ± 1 °C in triplicate to ensure data accuracy and reliability. The foil gas bags were removed and replaced with new ones at 12 h intervals, and the gas composition of the gas was analyzed by gas chromatography (GC). The cumulative gas volume produced by C. beijerinckii grown on SC and BS hydrolysates was measured every 12 h using the water displacement method. In this method, the foil gas bag was connected to an inverted graduated cylinder, initially filled with water and placed in a water bath or trough, via a valve. When the gas foil bag was manually pressed, the applied pressure forced the gas into the cylinder, displacing an equivalent volume of water. The displaced water volume was recorded by noting the change in the water level in the graduated cylinder. The gas volume was then determined by calculating the difference between the initial and final water levels. The measurements were taken under room temperature (21 °C) and atmospheric pressure conditions.

2.7. Other Analytical Methods and Calculations

The growth of C. beijerinckii was estimated by measuring the optical density (OD_{600 nm}; pathlength, 1 cm) using a UV-visible spectrophotometer (DU^®800, Beckman Coulter Inc., Brea, CA, USA). The total reducing sugars in hydrolysates before and after fermentation was determined using the 3,5-dinitro salicylic acid (DNS) assay method [19] and High-Performance Liquid Chromatography (HPLC Waters, Milford, MA, USA). The HPLC, used for glucose quantification, was equipped with a photodiode array (PDA) detector and a 3.5 µm Xbridge C18 column of 150 mm × 4.6 mm diameter as previously reported [20]. A glucose standard calibration curve was constructed to determine the concentration of reducing sugars in samples using the UV-spectrophotometer at 546_nm.

The quantitative evaluation of fermentation products (acetone, butanol, ethanol, acetic acid, and butyric acid) was determined using the Agilent Technologies 7890A Gas Chromatography system (Agilent Technologies Inc., Wilmington, DE, USA), which was equipped with a flame ionization detector (FID) and a 30 m (length) × 320 μm (internal diameter) × 0.50 μm (HP-Innowax film) J × W 19091 N-213 capillary column (Han et al., 2013). The capillary column of the GC was calibrated and conditioned to ensure accuracy and consistent quantification. The fermentation products were analyzed as described previously [21]. The total concentration of acetone, butanol, and ethanol produced (ABE) per gram of glucose utilized was used to calculate the yield while the productivity was calculated as the rate of the ABE produced (g/L) per hour.

The gas composition (H₂ and CO₂) in the collected gas bags was quantified using the gas chromatography (GC) 7890A system (Agilent Technologies 7890, Agilent Technologies Inc., Wilmington, DE, USA). Using a gas-tight syringe and needle, 50 µL of gas was drawn from the foil gas bags, which were collected directly from the bottle headspace and injected into the GC 7890A system to quantify the H₂ gas produced. The gas chromatograph was equipped with a thermal conductivity detector (TCD) and 30 m (length) × 320 μm (internal diameter) × 0.10 μm (CP-Molsieve 5A film) J × W CP7534 capillary column with nitrogen as the carrier gas. The amount of CO₂ gas produced was analyzed using a 5890 Hewlett-Packard gas chromatograph equipped with a thermal conductivity detector (TCD) and 30 m (length) × 320 μm (internal diameter) × 3.00 μm (GS-Carbonplot film) J × W 113-3133 GC column with helium as the carrier gas. Standard gas containing 51.6% CO₂ and 48.4% H₂ was used to calibrate the GC system and serve as a reference gas for H₂ and CO₂ quantification.

2.8. Statistical Analysis

GraphPad Prism 10 (software version 10.3.1) was used for statistical analysis. Analysis of variance (ANOVA) was conducted on treatments and significant differences in treatment means were determined by multiple comparisons using Turkey’s HSD test. Significant differences between treatments in terms of cell growth, butanol, ABE concentrations, yield, productivity, sugar utilization, and H₂ gas production were determined at a 95% confidence interval.

3. Results

3.1. Enzymatic Hydrolysis and Compositional Analysis of Spent Coffee Grounds and Biosolids cake Wastes

Lignocellulosic biomass such as food wastes are rich sources of lignin, cellulose, and hemicellulose, which are recalcitrant to hydrolysis [22], so pretreatment of lignocellulosic food wastes is necessary to make available the monomeric sugars needed by microorganisms for fermentation. The compositional and elemental analysis of hydrothermal pretreated SC and BS wastes is presented in

Table 1. Compositional analysis of spent coffee grounds and biosolids cake.

Parameters	Spent Coffee Grounds	Biosolids Cake
pH	5.99 ± 0.01	11.69 ± 0.01 *
Total solids (%)	72.14 ± 0.73 *	23.04 ± 0.06
Moisture (%)	27.86 ± 0.73	76.96 ± 0.06 *
Ash (%)	1.77 ± 0.10	9.29 ± 0.27 *
Calorific value (kJ/g)	21.93 ± 0.08 *	8.27 ± 0.08
Nitrogen (%)	2.50 ± 0.10	3.92 ± 0.07 *
Organic carbon (%)	52.95 ± 0.49 *	21.72 ± 0.09
Carbon/Nitrogen ratio	21.18:1	5.5:1
Major Element (µg/g)
Aluminum (Al)	69.74 ± 2.00	49340 ± 4120 *
Boron (B)	11.10 ± 1.03 *	7.93 ± 0.01
Calcium (Ca)	1350.60 ± 15.0	97,588 ± 3620 *
Copper (Cu)	17.19 ± 2.10	18.49 ± 1.00
Iron (Fe)	65.70 ± 3.22	4543.80 ± 348 *
Potassium (K)	7486 ± 25.0 *	684.51 ± 19
Magnesium (Mg)	1591.40 ± 28.3	60,465 ± 745 *
Manganese (Mn)	30.13 ± 4.80	59.29 ± 5.0 *
Molybdenum (Mo)	<1	1.10 ± 0.10 *
Sodium (Na)	18.79 ± 1.90	803.69 ± 11 *
Phosphorus (P)	1291.60 ± 28.60 *	9483.30 ± 594
Sulphur (S)	1555.20 ± 15.04	6743.60 ± 309 *
Zinc (Zn)	7.06 ± 0.19	374.21 ± 12 *
Minor Element (µg/g)
Arsenic (As)	<1	<1
Barium (Ba)	3.32	52.58 *
Beryllium (Be)	<1	<1
Cadmium (Cd)	<1	<1
Cobalt (Co)	<1	<1
Chromium (Cr)	<1	10.16 ± 0.35 *
Lithium (Li)	1.98 ± 0.10	<1
Nickel (Ni)	1.98 ± 0.24	6.19 ± 0.20 *
Lead (Pb)	<1	0.79 ± 0.01
Antimony (Sb)	<1	3.03 ± 0.06
Selenium (Se)	2.99 ± 0.30 *	1.85 ± 0.08
Silicon (Si)	49.12 ± 2.25	1180.95 ± 324 *
Strontium (Sr)	5.22 ± 0.09	273.58 ± 18 *
Titanium (Tl)	1	<1
Vanadium (V)	<1	11.59 ± 1.00

Values are means ± standard deviations of triplicate determinations, * indicate significant difference at p ≤ 0.05. A parameter value of 100% corresponds to 100 g, 100 mg or 100 µg, depending on the unit of measurement.

. Total solids content of SC was 72.14% higher than BS (23.04%) while the calorific value of SC (21.93 kJ/g) was 2.65-fold higher than BS (8.27 kJ/g). The elemental nitrogen content of SC (2.5%) was 1.57-fold lower than BS (3.92%), while SC contained 2.44-fold more carbon (52.95%) than BS (21.72%) and represents 21.2 and 5.5 carbon to nitrogen ratios, respectively. Notably, the BS was a rich source of major and minor elements such as Al, Ca, Fe, Na, P, S, Zn, Ba, Si, Sr, Cr, and V relative to SC (Table 1). However, SC was a richer source of K (7486 µg/g) than BS.

3.2. Batch Fermentation of Non-Detoxified Spent Coffee Grounds and Biosolids Hydrolysate

The glucose concentration of the non-detoxified Parr-treated spent coffee grounds and biosolids hydrolysates (without removing the lignocellulosic-derived microbial inhibitory compounds (LDMICs) are presented in Figure 1A. Parr-treatment and enzyme hydrolysis of SC (PEH) resulted in a significant 2-fold increase in glucose concentration of PEH hydrolysate (26.18 g/L) compared to the Parr-treated non-enzyme-hydrolyzed SC waste (PNEH;13.99 g/L). The glucose concentration of non-Parr-treated SC waste (NPEH; 1.72 g/L) was 15.2-, and 8.13-fold lower than PEH and PNEH hydrolysates, respectively (Figure 1A). Likewise, the glucose concentration of biosolids hydrolysates (BSH) increased with enzyme hydrolysis (6.26 g/L) and represents a 1.60-fold increase relative to non-enzyme-hydrolyzed biosolids waste (NBH; 3.92 g/L).

The fermentation profile of C. beijerinckii grown on non-glucose-modified SC and BS hydrolysates at 50% and 100% showed poor growth, sluggish fermentation, and low ABE concentration associated with poor carbon utilization (Figure 1B–D). C. beijerinckii growth on PEH50 was 3.09-fold lower than the control (OD₆₀₀ 6.74), while C. beijerinckii growth on PNEH, NPEH, and PEH was characterized with extended adaptation growth phase and OD_{600 nm} less than one (Figure 1B,C). Likewise, C. beijerinckii growth on the BSH, NBH, BSH50, and NBH50 shows 2.82-, 2.71-, 1.84-, and 2.97-fold less growth than the control, respectively. C. beijerinckii growth with 50% BS hydrolysate was not significantly different from 100% BS hydrolysate (Figure 1E,F).

C. beijerinckii fermentation of SC and BS hydrolysate (Figure 1D) did not produce detectable acetone and butanol except in PEH50 (0.37 g/L) and control (12.25 g/L). Ethanol was the only product detected when non-modified PEH, PNEH, NPEH, and PNEH50 were used for fermentation, and these hydrolysates supported 3.63-, 1.23-, 1.29-, and 1.19-fold greater ethanol concentration, respectively, than the control (1.57 g/L. Likewise, C. beijerinckii produced only ethanol in BS hydrolysates with higher concentration in NBH (1.21 g/L) (Figure 1G).

3.3. Fermentation Profile of C. beijerinckii Grown on Glucose Adjusted SC and BS Hydrolysate

Furthermore, we hypothesized that the low glucose concentration of SC and BS hydrolysates may have resulted in poor growth of C. beijerinckii and reduced production of ABE during fermentation, hence the hydrolysates were adjusted to 60 g/L with glucose and used for fermentation. Results showed improved growth, and ABE production by C. beijerinckii grown on glucose adjusted SC and BS hydrolysates, and butanol was the highest product concentration (Figure 2). Growth of C. beijerinckii on modified SC and BS hydrolysate was enhanced and followed a typical bacterial growth pattern for C. beijerinckii. With SC hydrolysates, C. beijerinckii growth on PEH and PEH50, was 1.58- and 1.19-fold lower than the control (Figure 2B). The NPEH and PNEH achieved maximum growth at 48 h similar to PEH, PEH50, and the control that achieved maximum growth at 36 h (Figure 2A). Although C. beijerinckii had the highest cell growth on the NPEH, PEH50 was only slightly different from PNEH and NPEH (Figure 2B). The estimated growth (OD600 _nm) values of C. beijerinckii on BS hydrolysate ranged from 4.33 to 5.59 (Figure 2F). Consistent with the observation with SC hydrolysates, C. beijerinckii growth increased with fermentation time and its growth on NBH was highest at 1.19-fold more than the control. Notably, C. beijerinckii growth on BSH was 1.53, and 1.29-fold less than the control and NBH, respectively (Figure 2E,F).

Butanol and ABE concentration significantly increased with C. beijerinckii fermentation time on glucose-adjusted SC and BS hydrolysates (Figure 2C,D). Glucose adjusted PEH50 supported butanol production more than other SC hydrolysates and represents 1.47- and 1.17-fold greater butanol concentration than PEH and PNEH hydrolysates, respectively. Moreover, butanol production on the PEH50 was not significantly different from the NPEH (11.10 g/L) (Figure 2C). Notably, the least butanol production was observed with the PEH (8.10 g/L) and was 1.57-fold less than the control (12.78 g/L). Interestingly, ABE concentration was significantly higher in the PEH50 (16.81 g/L) substrate producing 1.24- and 1.11-fold more ABE than the PEH and NPEH, respectively (Figure 2D). Notably, the PEH had the lowest concentration, 1.27-fold less ABE than the control (17.19 g/L). As expected, acetone production by C. beijerinckii was highest in the control (3.21 g/L) followed by PEH50 (2.18 g/L) as shown in (Table S1). The PEH hydrolysate (3.30 g/L) supported ethanol production to a greater extent than NPEH (1.95 g/L) and the glucose control (1.57 g/L) (Table S1).

Furthermore, adjustment of BS hydrolysates with glucose significantly increased butanol and ABE production by C. beijerinckii (Figure 2G,H). Butanol concentration was highest in the C. beijerinckii-fermented NBH (11.82 g/L) and 1.07- and 1.05-fold greater than in NBH50 (11.04 g/L) and BSH50 (11.25 g/L), respectively. Butanol production by C. beijerinckii on the BSH was lowest, 1.47-fold less than the control (12.09 g/L), while on the NBH, butanol concentration was ~1.02-fold less than the control. Acetone concentration in C. beijerinckii-fermented NBH was not significantly different from BSH50, both 1.39-fold lower than the control (Table S1).

The ABE concentration of the NBH (16.28 g/L) was 1.05-fold less than the control (17.03 g/L) and was not significantly different from the NBH50 (16.27 g/L) and BSH50 (15.81 g/L) when fermented by C. beijerinckii. Although NBH (1.33-fold increase) supported more ABE production than BSH, ABE concentration of NBH was not significantly different from BSH50 (Figure 2D,H). Moreover, ABE yield and productivity increased with glucose adjustment of BS and SC hydrolysates (

Table 2. Product yield and productivity of C. beijerinckii during ABE fermentation with spent coffee grounds and biosolids hydrolysates.

Parameters	ABE Yield (g/g)		ABE Productivity (g/L/h)		Butanol Yield (g/g)		Butanol Productivity (g/L/h)		H2 Yield (mmol/g)	References
	Gluc	Glu+CaCO3	Gluc	Gluc+CaCO3	Gluc	Glu+CaCO3	Gluc	Gluc+CaCO3	Gluc+CaCO3
Spent coffee hydrolysates
PEH A	0.24 e	0.27 e	0.23 e	0.24 e	0.14 e	0.20 c	0.14 e	0.17 e	0.62 d	This study
PEH50 A	0.3 b	0.32 b	0.28 b	0.36 d	0.21 b	0.22 b	0.20 b	0.25 b	0.84 b	This study
PNEH A	0.26 d	0.29 d	0.24 d	0.26 c	0.18 d	0.20 c	0.17 d	0.18 d	0.55 e	This study
NPEH A	0.27 c	nd	0.25 c	nd	0.20 c	nd	0.19 c	nd	nd	This study
PNEH50 A	nd	0.30 c	nd	0.26 c	nd	0.22 b	nd	0.19 c	0.67 c	This study
Biosolids hydrolysates
BSH A	0.21 e	0.28 c	0.17 e	0.24 e	0.14 d	0.19 c	0.11 d	0.17 d	0.80 e	This study
BSH50 A	0.27 c	0.32 b	0.22 c	0.28 c	0.19 c	0.24 b	0.16 b	0.21 b	0.87 d	This study
NBH A	0.28 b	0.35 a	0.23 b	0.29 b	0.20 b	0.25 a	0.16 b	0.21 b	0.93 b	This study
NBH50 A	0.26 d	0.32 b	0.21 d	0.26 d	0.19 c	0.24 b	0.15 c	0.20 c	0.89 c	This study
Control (P2) A	0.39 a	0.44 a	0.29 a	0.38 a	0.29 a	0.32 a	0.21 a	0.27 a	1.06 a	This study
Grape waste B	nd	nd	nd	nd	nd	nd	nd	nd	12.21	[23]
Crude cheese whey C	nd	nd	nd	nd	nd	nd	nd	nd	0.03	[24]
Potato processing waste D	nd	nd	nd	nd	nd	nd	nd	nd	0.04	[25]
Food industry residue and manure E	nd	nd	nd	nd	nd	nd	nd	nd	0.74	[26]
Restaurants food waste F	nd	nd	nd	nd	0.17	0.06	0.17	0.17	0.08	[27]
Starch wastewater G	nd	nd	nd	nd	nd	nd	nd	nd	4.11	[28]
Molasses wastewater H	nd	nd	nd	nd	nd	nd	nd	nd	2.99	[29]
Corn starch A	0.3	nd	0.25	nd	0.22	nd	nd	nd	nd	[15]
Molasses I	0.21	nd	0.08	nd	nd	nd	nd	nd	nd	[30]
Inedible dough A	0.37	nd	0.24	nd	0.19	nd	nd	nd	nd	[15]
Batter liquid A	0.37	nd	0.31	nd	0.21	nd	nd	nd	nd	[15]
Glucose medium A	0.36	0.43	0.22	0.42	nd	nd	nd	nd	nd	[21]
Distiller dried grains with solubles (DDGS) J	0.32	nd	0.18	nd	nd	nd	nd	nd	nd	[31]
Cereal waste water K	nd	nd	nd	nd	nd	nd	nd	nd	0.79	[32]

Inoculum: ^A C. biejerinckii NCIMB 8052; ^B Sewage sludge; ^C Clostridium saccharoperbutylacetonicum; ^D Soil from tomato plant; ^E Anaerobic digester sludge; ^F C. beijerinckii BOH3; ^G Mixed liquor of Thermoanaerobacteriaceae; ^H Sludge; ^I Clostridium acetobutylicum and C. butylicum, ^J C. beijerinckii 260, ^K Dewatered sewage sludge. Values represents means of triplicate determinations and different lowercased alphabet superscript represent significant difference at p ≤ 0.05. Keys: PEH: Parr-treated and enzyme-hydrolyzed coffee (100%); PEH50: Parr-treated and enzyme-hydrolyzed coffee (50%); PNEH: Parr-treated and non-enzyme-hydrolyzed coffee (100%); PNEH50: Non-Parr-treated and enzyme-hydrolyzed coffee; BSH: Biosolids hydrolysate (100%); BSH50: Biosolids hydrolysate (50%); NBH: Non-hydrolyzed biosolids (100%); NBH50: Non-hydrolyzed biosolids (50%); Control: P2 medium (60 g/L glucose), nd (not determined).

). ABE yield (0.28 g/g) and productivity (0.23 g/L/h) of C. beijerinckii grown on the BS hydrolysate was highest in the NBH, being 1.33- and 1.07-fold more than the BSH and the NBH50, respectively (Table 2). Notably, the ABE yield and productivity of the NBH (0.28 g/g; 0.23 g/L/h) was lower than the control (0.39 g/g; 0.29 g/L/h), respectively. On PEH50 hydrolysates, ABE yield and productivity (0.3 g/g; 0.28 g/L/h) were greater than in other SC hydrolysates (Table 2). ABE yield in the PEH50 was 1.25-, 1.15-, and 1.11-fold higher than PEH, PNEH, and NPEH, respectively. The ABE productivity on the PEH50 was 1.22-fold higher than the PEH, while the ABE productivity on PEH50 was 1.05-fold less than the control.

3.4. Fermentation Profile of CaCO₃ Modified SC and BS Hydrolysate

After quantifying the potentials of SC and BS hydrolysates as substrates for C. beijerinckii-mediated butanol production, we further modified the SC and BS hydrolysates by CaCO₃ supplementation (2 g/L) to improve the fermentation efficiency. SC (PEH and PNEH) and BS (BSH and BSH50) hydrolysates at 50 and 100% were selected based on their higher butanol concentration and lower sugar supplementation needs. The choice of these hydrolysates could favor the economics of butanol production at the industrial scale by lowering the cost associated with glucose supplementation of the food waste. Supplementation of BS and SC hydrolysates with CaCO₃ (2 g/L) significantly improved C. beijerinckii metabolism and was characterized by enhanced growth, sugar utilization, butanol titer, ABE yield, and productivity (Figure 3).

CaCO₃ supplementation increased C. beijerinckii growth and ABE production in both the control and hydrolysates, with the highest levels attained in the glucose control. Notably, CaCO₃-modified PEH50 (6.39) supported C. beijerinckii growth more than other SC hydrolysates and was not significantly different from PNEH50 (6.05) (Figure 3A). CaCO₃-adjusted PEH improved C. beijerinckii growth 1.20-fold more than the non-modified PEH. C. beijerinckii growth on the PEH50 increased by 1.18-fold relative to the non-CaCO₃-modified PEH50 hydrolysate. However, the PEH50 was 1.18-fold lower than the CaCO₃-modified glucose medium (7.55). Likewise, C. beijerinckii growth on BSH, BSH50, NBH50, and NBH increased by 1.39, 1.16, 1.25, and 1.1-fold, respectively, when the hydrolysates were supplemented with CaCO₃ (Figure 3E,F). Notably, C. beijerinckii growth on NBH was not significantly different from other BS hydrolysates but was 1.23-fold less than the control.

CaCO₃ supplementation enhanced butanol concentration of both SC and BS hydrolysates. The modification of PEH hydrolysate with CaCO₃ significantly enhanced butanol titer (10.32 g/L), yield (0.20 g/g), and productivity (0.17 g/L/h) by 1.27, 1.43, and 1.21-fold relative to the non-CaCO₃-modified PEH hydrolysate (Figure 3C and Table 2). Nonetheless, butanol concentration obtained with PEH50 and PNEH50 was 1.08 and 1.17-fold, respectively, less than the glucose control (Figure 3C). Notably, ethanol concentration was less in the CaCO₃-modified hydrolysates than the non-CaCO₃-modified hydrolysates (Table S1).

Butanol production with the CaCO₃-modified BSH and BSH50 hydrolysates was 1.25 and 1.12-fold greater than the non-CaCO_3-modified hydrolysates, respectively, whereas NBH and NBH50 had 1.1 and 1.05-fold greater butanol concentration after CaCO₃ modification of the hydrolysates (Figure 3G). As expected, butanol concentration and yield in the control (13.07 g/L; 0.30 g/g) was higher than the other hydrolysates (Table 2). Consistent with the higher butanol concentration observed with glucose-adjusted NBH, CaCO₃-modified NBH produced the highest butanol (12.87 g/L) concentration, 1.25-fold more than non-CaCO₃-modified hydrolysate. Butanol yield from the NBH (0.25 g/g) was 1.25-fold greater than the non-CaCO₃-modified hydrolysate, while the butanol yield obtained with BSH50 was not significantly different from NBH50 (p ≥ 0.05). Butanol productivity with CaCO₃-modified NBH was 1.31-fold greater than the non-CaCO₃-modified hydrolysate (0.16 g/L/h) (Table 2).

CaCO₃ modification of SC hydrolysates favored ABE production, yield, and productivity, by C. beijerinckii, with higher ABE concentration, yield, and productivity obtained with PEH50 (17.37 g/L), relative to other SC hydrolysates (Figure 3D). CaCO₃-modified PEH50 supported enhanced sugar utilization and had 1.2 and 1.11-fold greater ABE yield than PEH and PNEH50, respectively (Table 2). Consequently, the maximum ABE concentration from the NBH (17.16 g/L) was 1.17 and 1.1-fold more than BSH and NBH50, respectively (Figure 3H). ABE yield from BSH, BSH50, and NBH increased by 1.33, 1.19, and 1.25, while ABE productivity went up 1.41, 1.27, and 1.26-fold, respectively, after CaCO₃ modification of the hydrolysates.

3.5. Biohydrogen Production by C. beijerinckii Grown on Spent Coffee Grounds and Biosolids Hydrolysates

Gas production by C. beijerinckii grown on the SC and BS hydrolysates ceased after 60 h of fermentation time (Figure 4). The total gas produced by C. beijerinckii on the SC hydrolysates was highest in PEH50 (2.22 L), 1.18-fold more gas than on PEH (1.88 L) but was 1.11-fold lower than on the glucose control (Figure 4A). Similarly, PNEH total gas production was 1.09-fold lower than on the PNEH50. The total gas production on BS hydrolysates ranged from 2.22 L to 2.37 L (Figure 4F). The total gas production by C. beijerinckii grown on the NBH was not significantly different from the NBH50 and BSH50 but was higher than the BSH (2.21 L). However, the total gas production by C. beijerinckii grown on the NBH and BSH was 1.04- and 1.11-fold, respectively, less than the control.

The maximum CO₂ and H₂ gas production profile when C. beijerinckii was cultured on modified SC hydrolysates revealed a significant increase in CO₂ and H₂ gas production with increasing fermentation time with the glucose control supporting the highest gas production (Figure 4B,C). Maximum H₂ gas production was obtained at 36 h after which there was a significant decline in H₂ gas production from both SC and BS-modified hydrolysates (Figure 4B,G). Cumulative H₂ gas production from SC hydrolysates ranged from 0.67 L to 0.79 L, with the largest H₂ production from the PEH50 (0.79 L), 1.14, 1.41, and 1.1-fold more H₂ production than from the PEH, PNEH, and PNEH50 (Figure 4B,C), respectively. Notably, C. beijerinckii batch fermentation produced more H₂ on 50% SC hydrolysates than 100% SC hydrolysates. H₂ yields from the SC hydrolysate ranged from 0.55 to 0.84 mmol/g glucose, while with BS hydrolysates, H₂ yield ranged from 0.80 to 0.93 mmol/g of glucose (Table 2). H₂ yield was highest with PEH50 (0.84 mmol/g glucose) and NBH (0.93 mmol/g glucose) but slightly lower than the control (1.06 mmol/g glucose). Interestingly, the H₂ gas production with NBH was 1.13-fold more than BSH. Nonetheless, NBH produced 1.14-fold less H₂ gas than the control. Furthermore, CO₂ gas produced by C. beijerinckii during the fermentation of SC hydrolysates ranged from 1.20 to 1.42 L, and the highest CO₂ gas was obtained with PEH50 (1.42 L) representing 1.18-fold more than PNEH (Figure 4D,E). However, the cumulative CO₂ gas production with the control was 1.07-fold greater than PEH50 substrate. The cumulative CO₂ gas production on BS hydrolysate was highest in NBH (1.55 L), representing 1.02-fold more CO₂ gas than the control.

4. Discussion

The carbon and nitrogen content of pretreated and hydrolyzed LB are essential nutrients that enable these feedstocks to support microbial growth and cellular metabolism [33]. Notably, the carbon–nitrogen ratio in SC hydrolysate was higher than the BS hydrolysates, while BS hydrolysates had higher concentrations of major and minor elements compared to SC hydrolysates, suggesting their potential use as nutrient supplement in microbial fermentation. Dissociated forms of metals play a crucial role in cellular metabolism of solventogenic Clostridium species. Specifically, potassium plays an important role in the regulation of pH homeostasis, protein and enzyme synthesis regulation, and transport that enables signal transduction and coenzyme metabolism [34]. Therefore, SC may serve as a suitable substrate, while BS could be an effective nutrient supplement for producing valued-added products through ABE fermentation.

The rigid cellulose–hemicellulose–lignin matrix makes LB pretreatment essential, and the choice of method and conditions for its deconstruction depends on the structural composition of the specific LB [35]. LBs (including SC food waste) contain complex carbohydrates such as cellulose and hemicellulose, and lignin that requires deconstruction to release the monomeric sugars needed by C. beijerinckii for growth and biofuel production.

The higher glucose concentration in the PNEH relative to the NPEH may be linked to the disruption of lignocellulosic matrix of SC with concomitant partial breakdown of its hemicellulose component due to Parr treatment at high temperature. Likewise, the increased sugars in PEH after enzyme hydrolysis is due to increased exposure of the disrupted lignocellulosic matrix to hydrolytic enzymes, culminating in increased release of sugars. According to Huang et al. [36], SC contains cellulose (33.10%), hemicellulose (30.03%), and lignin (24.52%). The cleavage of O-acetyl and uronic acid substitutions from hemicellulose during Parr treatment generates acetic acid and other organic acids. These acids help catalyze the hydrolysis of polysaccharides like hemicellulose into soluble oligosaccharides and subsequently into monomeric sugars during hydrothermal pretreatment of the LB [37,38]. Increased concentration of reducing sugar was reported after hydrothermal pretreatment of rice straw [39] and hydrolysis of banana peels with pectinase [40]. SC and BS food wastes contain polysaccharides (cellulose and hemicellulose) and lignin that could be degraded through pre-treatment and enzyme hydrolysis to liberate sugars that could potentially serve as nutrients for fermenting microorganisms.

C. beijerinckii exhibited greater growth on 50% SC hydrolysates than 100% SC hydrolysates. This difference was attributed to the presence of lignocellulosic-derived microbial inhibitory compounds (LDMICs) in the SC hydrolysates. Solventogenic Clostridium species are vulnerable to LDMICs that are released during various pretreatment processes of LB [12]. However, the benefits of hydrothermal pretreatments stand out over other methods based on reduced costs of procuring catalysts, alkali neutralization reagents, and sugars to support fermentation [41]. Likewise, reduced generation of LDMICs during high-pressure steam Parr treatment of LB favors butanol production more than the acidic catalyst pretreatment method [11]. The observed inhibition of growth and ABE production by C. beijerinckii when grown on non-modified SC and BS hydrolysates may be connected to the lower glucose concentration and the presence of LDMICs in hydrolysates. The SC and BS wastes contain phenolics [6,10], which could alter the cellular metabolic process of C. beijerinckii during fermentation [11,42]. Furan derivatives such as furfural and 5-hydromethylfurfural (HMF) are generated during acid and hydrothermal pretreatments of lignocellulosic feedstocks. These compounds can inhibit glycolysis and the tricarboxylic acid cycle in microorganisms by interfering with the production and activity of key enzymes, including aldehyde dehydrogenase, which are essential for cellular metabolism [43,44]. Therefore, to improve the growth of C. beijerinckii and ABE production, the sugar composition of SC and BS hydrolysate was adjusted to 60 g/L with glucose and supplemented with CaCO₃.

The relatively high butanol production with the NBH substrate may be attributed to the dense mineral content of BS hydrolysates (Figure 2G), which could have stimulated the expression and activity of key enzymes involved in the growth and ABE production pathway in C. beijerinckii. The relatively low ABE in BSH and PEH may be attributed to inhibitory compounds in the feedstock because insoluble-bound phenolics can be converted into soluble free phenolics during pretreatment and enzyme hydrolysis of biomass [45]. The increased ethanol production with undiluted PEH hydrolysates may be linked to the greater toxicity effect of LDMICs (e.g., phenolics, furfural, and HMF) on C. beijerinckii. The presence of LDMICs could stress the fermenting cells and disrupt their redox balance [11], which can cause C. beijerinckii to expend nutrient and cofactor (NADH) resources meant for solvent production to maintaining cell integrity and pH homeostasis. The production of relatively large amount of ethanol may have been a strategy by the cell to maintain energy homeostasis under stress, as ethanol production is less energy-intensive [11,42]. Moreover, the higher ABE production with PEH50 hydrolysate corroborates the assertion that LDMICs negatively impact C. beijerinckii cellular growth and metabolism.

Overliming pretreated and hydrolyzed LB can relieve the toxic effects of soluble phenolics and furan derivatives on fermenting microorganisms [46]. However, overliming incurs additional costs due to the expense of chemicals, substrate loss, and the generation and disposal of gypsum (CaSO₄·2H₂O) waste. The supplementation of SC and BS hydrolysates with CaCO₃ improved C. beijerinckii growth and butanol production. The enhanced growth, butanol and ABE titers, and productivity from the fermentation of CaCO₃-modified SC and BS hydrolysates (Figure 3A–H) may be attributed to the buffering effect of CaCO₃, increased expression and activity of key enzymes involved in growth and ABE production pathway in C. beijerinckii, which helped to ameliorate the toxic effects of LDMICs on C. beijerinckii [11,20]. While the relatively high ABE concentration obtained during the fermentation of pretreated and modified SC and BS hydrolysates substantiates our hypothesis that food wastes can be valorized into bioenergy products, it is important to note that substrate modification may be needed to enhance the robustness of the fermenting microorganisms and improve the fermentation of the feedstock to target products.

Furthermore, the higher butanol yield and productivity obtained with PEH50 and NBH hydrolysates relative to the non-CaCO₃-modified hydrolysates may be connected to the increased sugar utilization and enhanced redox balance during C. beijerinckii fermentation of food wastes supplemented with CaCO₃. Like the observations in this study, Su et al. [47] reported improved sugar utilization and butanol production with CaCO₃-supplemented corn straw hydrolysate fermented by C. acetobutylicum, while Han et al. [21] reported similar ABE yield and productivity with CaCO₃-supplemented glucose medium. As shown in Table 2, butanol and ABE yield of C. beijerinckii grown on SC and BS hydrolysates were similar to values reported when inedible dough, batter liquid, and corn starch [15] were used as substrates. ABE yield and productivity reported in fermentations with molasses and distiller dried grains with solubles (DDGS) as substrates were lower than the values obtained with PEH50 and NBH from this study [30,31]. Notably, NBH and PEH50 supported butanol yield productivity more than in restaurant food waste fermented by C. beijerinckii BOH3 [28]. Through substrate modification and improved buffering systems, the toxicity limitations of SC and BS hydrolysates can be mitigated, which will allow these substrates to be repurposed to produce value-added products through C. beijerinckii fermentation. To the best of our knowledge, this study is the first to demonstrate the potential of SC and BS food wastes as substrates for biobutanol production by C. beijerinckii.

Fermentation conditions such as pH, temperature, type of microorganisms, inoculum population, contaminants, pretreatment methods, and substrate type and concentration significantly influence H₂ production [48]. The decline in gas production after 36 h of fermentation (Figure 4B) may be due to the depletion of readily available nutrients, the accumulation of metabolic byproducts that inhibit fermentation, and the transition of the bacteria into the stationary phase [49]. As observed in our study (Figure 4B,C,G,H), Jang et al. [50] reported higher H₂ production at close to neutral fermentation medium pH than acidic pH [51,52]. Ferchichi et al. [24] reported biohydrogen yield (7.89 mmol/g lactose) from crude cheese whey fermented by C. saccharoperbutylacetonicum. The H₂ yield was lower than the yields reported for the fermentation of ripened fruits and starch wastewater, which produced 12.12 mmol/g using sewage sludge and 4.11 mmol/g using a liquor predominantly containing mixed culture from the family Thermoanaerobacteriaceae, respectively [23]. Notably, the H₂ yield reported from food industrial residue and manure (0.74 mmol/g), and cereal wastewater (0.79 mmol/g) were similar to this study [26,32]. Moreover, this study reported higher H₂ yield in the PEH50 (0.84 mmol/g) and the NBH (0.93 mmol/g) than in potato processing waste [25] and restaurant food waste fermented with C. beijerinckii BOH3 [27], suggesting the potential of SC and BS hydrolysates as viable resources for H₂ production.

Lower hydrogen yield (0.84 mmol/g) from fermentation of the PEH50 than in some previous studies [23,29] may be due to differences in substrate composition and metabolic tradeoff that resulted in the diversion of carbon flux and reducing cofactors into the production of NAD(P)H-dependent products (butanol and acetic and butyric acids). H₂ gas is produced through the activity of ferredoxin-linked hydrogenase, and the higher the solvent production, the lower the hydrogen yield [51]. H₂ production in non-CaCO₃-adjusted substrates was negligible due to poor microbial growth and acidic pH, which may have disrupted cell homeostasis and redirecting ATP towards maintaining cellular integrity rather than biohydrogen production.

Overall, the increased butanol and H₂ gas concentration, yield, and productivity obtained from NBH and PEH50 fermentation suggest that these previously underutilized and recalcitrant food wastes could serve as viable substrates for the commercial production of biobutanol and H₂ gas, and both are alternative energy sources to fossil fuels. Moreover, metabolic engineering [53] is a valuable tool that can be used to enhance biohydrogen production from food wastes like SC and BS. Additionally, using these food wastes for ABE fermentation could improve the economic viability of industrial biobutanol production, making it more profitable while aligning with the objectives of circular economy and contributing to maintaining safer climates.

5. Conclusions

This study demonstrated that lignocellulosic SC and BS hydrolysates can serve as carbon sources to produce fuels and chemicals (butanol, ABE, H₂) through C. beijerinckii fermentation. Our findings established that slight modification of the fermentation medium, in which SC and BS hydrolysates serve as carbon sources, more effectively converted these hydrolysates to fuels and chemicals. For instance, supplementation of SC and BS hydrolysates with CaCO₃ resulted in enhanced butanol and ABE titer, yield, productivity, and H₂ gas production. This approach may address challenges related to the toxicity of lignocellulosic biomass hydrolysates, low product yield, and poor productivity commonly encountered in ABE production within industrial biorefineries. Additionally, the utilization of these low-cost feedstocks (food wastes) could help reduce the high cost associated with industrial biofuel production. The study has established the possibilities of valorizing food wastes into biofuels and developed a scalable and environmentally friendly process to handle industrial food wastes in furtherance of the greener climate (reduce greenhouse gas emission) and clean energy generation goals. Future studies may focus on enhancing product yield and productivity by isolating or developing novel solventogenic microbial strains with improved tolerance to LDMICs, as well as exploring the use of multi-species fermentation for food wastes conversion to value-added products.