Improvement of Thermophilic Butanol Production by Thermoanaerobacterium thermosaccharolyticum from Waste Figs Through the Gradual Addition of Butyric Acid

Abstract

This study focuses on determining the optimal fig and butyric acid concentrations for butanol production under thermophilic conditions. Waste fig is a potentially rich substrate in sugars, minerals, and vitamins, but it is insufficient for effective butanol formation when butyrate is not present in the media because butanol is produced by butyrate reduction. Therefore, butyric acid was supplemented gradually in certain concentrations to fig-containing fermentation broth. The best combination of butyric acid and fig was determined using the Box–Wilson statistical experiment design. Fig and butyric acid concentrations were set as independent variables, while butanol concentration was the objective function. When the concentrations of butyric acid and fig were near the middle of the ranges under inspection, more butanol was produced. Butanol production was the lowest as fig and butyric acid values got closer to the extremes, particularly at high concentrations. Maximum butanol of 0.32 g/L was obtained with 16 g fig/L and 1.6 g butyric acid/L. The quadratic model generated was found to be significant, and its reliability was tested with verification experiments with reproducible results. This study showed that butanol could be produced from butyrate-supplemented fig waste under thermophilic conditions with a consolidated bioprocessing approach.

1. Introduction

Ficus carica L. (fig), belonging to the Moraceae family, is well adapted to specific regions due to its unique climatic requirements and is commonly cultivated in the Middle East, Southwest Asia, and Mediterranean areas [1,2]. The world’s largest fig grower is Türkiye, which is followed by Egypt, Morocco, Iran, Algeria, and Greece [3]. In 2019, 310,000 tons of figs were produced on 52,116 hectares in Türkiye [4]. The dry fig production in Türkiye in 2019 was 90,000 tons, and 57,000 tons of dried figs were exported in 2021 [5]. Figs are rich in vitamins and minerals, fiber, and antioxidants, and their content gets more concentrated after drying. According to the USDA (United States Department of Agriculture), 100 g of dried figs contains the following: 249 kcal energy, 63.9 g carbohydrates, 9.8 g fiber, 47.9 g sugars, 0.92 g total fat, 3.3 g protein, minerals, and vitamins, with high levels of calcium, potassium, phosphorus, and magnesium [6].

Aflatoxin, a significant concern in fruit production, is a mycotoxin produced by Aspergillus species such as A. flavus and A. parasiticus [7,8]. Aflatoxin is a health hazard due to its toxic, carcinogenic, and mutagenic properties and also affects food quality [9,10]. Many food types, such as nuts, maize, rice, dried fruits, cocoa beans, and spices, are exposed to mycotoxins during the pre-harvest and post-harvest periods, especially during the drying process [11,12]. Special attention was drawn to potential aflatoxin contamination of pistachios, hazelnuts, and dried figs produced in Türkiye [11]. Aflatoxin-contaminated figs emit a bright greenish-yellow fluorescence under ultraviolet light (UV) [13,14]. Özmıhçı et al. determined the aflatoxin content in dried figs obtained from Aegean Exporters’ Associations as 69 µg/kg aflatoxin B1, 5.99 µg/kg aflatoxin B2, 84.86 µg/kg aflatoxin G1, and 5.79 µg/kg aflatoxin G2 [15].

The rapid consumption of fossil fuels has raised concerns over a potential energy crisis, while the greenhouse gas emissions resulting from their combustion contribute to global warming, which is considered the most pressing environmental issue [16,17]. This situation has led to the search for environmentally friendly fuels that may serve as an alternative to petroleum-derived fuels [18,19,20,21]. There has been an increase in interest in liquid biofuels as a result of rising oil prices, energy dependency, and environmental concerns [22,23]. In recent years, the search for alternatives to petroleum fuels has led to the development of various biofuels, including biobutanol and biomethanol, of which biodiesel and bioethanol are the most widely used [24,25]. Among biofuels, biobutanol is a more suitable biofuel than ethanol due to advantages such as higher energy content, lower vapor pressure, less corrosiveness, and less hygroscopic properties [26]. Butanol can be used as a fuel additive or directly used instead of gasoline in any gasoline engine without modification [27,28].

Butanol is a straight-chain liquid with four carbon atoms, a molecular formula of C₄H₉OH, and a molecular weight of 74.12 g/mol, and it is miscible with organic solvents. Butanol has a wide range of industrial uses, such as paint, polymer, cosmetics, and pharmaceuticals; it is also suitable to be used as a biofuel [29]. Research into butanol production by microbial fermentation was first discovered by Pasteur in 1861, then continued in 1911 with Fernbach’s isolation of a butanol-producing bacterium from potatoes. This was followed in 1912 with Weizmann’s isolation of a bacterium called Clostridium acetobutylicum, which produces acetone-butanol-ethanol [30,31]. Butanol production by microbial fermentation became increasingly important after the First World War; its use declined in the 1960s with the widespread use of chemical production processes but was revived with the energy crisis and the increase in oil prices [32].

Butanol production by microbial fermentation is based on acetone-ethanol-butanol (ABE) fermentation, which is an anaerobic fermentation process. Substrate sources can be agricultural waste, industrial by-products, or waste materials with a high sugar content [33]. ABE fermentation consists of a first acidogenic phase in which organic acids such as acetic acid and butyric acid and products such as Carbon dioxide gas (CO₂) and hydrogen gas (H₂) are produced. The second stage consists of a solventogenic phase in which the organic acids produced in the acidogenic phase are converted into alcohols, such as ethanol and butanol, and ketones, such as acetone [34,35,36].

Thermoanaerobacterium thermosaccharolyticum DSM 571 is a thermophilic bacteria reported to produce biobutanol [37]. Compared to mesophilic fermentation, thermophilic temperatures have advantages such as the ability to facilitate substrate hydrolysis and reduced risk of contamination [38,39,40].

This study aimed to investigate the effects of substrate and butyric acid concentration on butanol production from waste figs by thermophilic fermentation. Butyric acid was added gradually to fig-containing media. To assess the effect of each parameter, a Box–Wilson statistical experimental design was established. We hereby report thermophilic butanol production from waste fig by a non-genetically modified culture of Thermoanaerobacterium thermosaccharolyticum, presenting sound and novel results for the current literature.

2. Materials and Methods

2.1. Substrate and Inoculum

Waste figs obtained from the Aegean Exporter Association, Izmir, Türkiye, were used as a substrate in this study. The waste figs were chopped and then dried at 50 °C for 120 h. Then, dried figs were ground to a mesh size of less than 1 mm. The dried fig powder was homogeneous and contained all parts of the fruit, like seeds, peel, and flesh.

Thermoanaerobacterium thermosaccharolyticum DSM 571 was obtained from the German Collection of Microorganisms and Cell Cultures (DSMZ). The culture was grown at 55 °C and pH: 7.0 in a medium that contained the following, per liter [37]: 20 g of starch, 2 g of yeast extract, 1 g of NH₄Cl, 0.1 g of MgSO₄·7H₂O, potassium phosphate buffer 50 mM, 2 mL of vitamin solution [41], 0.25 g of L-cysteine·HCl, and 2 mL of trace element solution [42]. The medium was freshly prepared and autoclaved at 120 °C for 15 min before inoculation. The raised culture was transferred to the fermentation media after 24 h under anaerobic conditions at 55 °C with an inoculation percentage of 5% (v/v).

2.2. Experimental Setup and Procedure

100 mL bottles made of borosilicate glass were used for all the experiments. The bottles were sealed using rubber stoppers and screw caps. All experiments were performed in a shaker incubator at 55 °C, 50 rpm stirring speed, and 50 mL working volume for 5 days. The fermentation media contained yeast extract (2 g/L), potassium phosphate buffer (50 mM), NH₄Cl (0.5 g/L), FeSO₄·7H₂O (0.2 g/L), vitamin solution (2 mL) [41], trace element solution (2 mL) [42], MgSO₄·7H₂O (0.1 g/L), L-cysteine·HCl (0.25 g/L), and a defined concentration of fig and butyric acid according to the Box–Wilson design. Butyric acid (>99%) was obtained from Sigma Aldrich. The amounts of butyric acid determined by the Box–Wilson experiment were equally divided and added to the experimental medium a total of 10 times.

pH and Oxidation-Reduction Potential (ORP) probes of HANNA 2211 were used for pH and ORP control. All experiments were started at pH 7, and the pH was controlled at this level using 10 M NaHCO₃. Redox values generally varied from −50 mV to −250 mV. Anaerobic conditions were established by flushing the bottles with argon gas, an inert gas, for 2 min at the start of the experiments to remove dissolved oxygen from the medium and to sweep the air from the headspace.

2.3. Analytical Methods

Before the analyses, the samples were centrifuged and the supernatants were analyzed for total sugars by the acid phenol spectrophotometric method [43]. Analysis kits (Merck Spectroquant, 1.01763.0001, Darmstadt, Germany) were used to analyze total volatile fatty acids in the WTW Photolab S12 photometer. Lignin, cellulose, and hemicellulose analyses for ground-dried figs were carried out as previously described [44]. Total organic carbon, total Kjeldahl nitrogen, and total phosphorus analyses of dried figs were carried out according to the standards as described previously [45]. Standard methods were used to determine the biomass concentration [46]. The concentration of butanol was determined using a gas chromatograph (Agilent 7890N, Santa Clara, CA, USA) equipped with an HP-5 (30 m; 0.32 mm; 0.25 μm) column and a flame ionization detector (FID). Nitrogen at a flow rate of 0.5 mL/min was used as the carrier gas. In addition, H₂ (30 mL/min) and dry air (200 mL/min) flow rates were used. The maximum oven temperature was 300 °C. The temperature profile was kept at 40 °C for 1 min, increased by 6 °C/min to 130 °C, and kept at 130 °C for 1 min. The split ratio was 1:20. The sample volume was 5 µL and the total analysis time was 17 min. For analysis on the GC-FID HP5 column, the sample was passed from the water phase to the solvent phase. Samples taken at the end of thermophilic fermentation were mixed with an equal volume of dichloromethane and centrifuged at 8000 rpm for 10 min, and the lower phase was subjected to the GC for butanol analysis [47]. Fourier Transform Infrared Spectroscopy (FT-IR) analysis of the figs used in this study was performed by Perkin Elmer spectrum two ATR FT-IR. The sample spectrum was scanned 64 times within a wavenumber range of 4000–500 cm⁻¹.

2.4. Box–Wilson Design

Experimental conditions were determined using the Box–Wilson statistical experiment design based on the response surface methodology [48]. The design matrix comprised four axial, four factorial, and triplicate sets of center points totaling 11 experiments at different substrate and butyric acid concentrations. The correlation of butanol concentration with substrate and butyric acid concentrations was analyzed using the following function.

$$\mathrm{Y}\mathrm{=}{\mathrm{a}}_{\mathrm{0}}\mathrm{+}{\mathrm{a}}_{\mathrm{1}}{\mathrm{·}\mathrm{X}}_{\mathrm{1}}\mathrm{+}{\mathrm{a}}_{\mathrm{2}}{·\mathrm{X}}_{\mathrm{2}}\mathrm{+}{\mathrm{a}}_{\mathrm{12}}\mathrm{·}{\mathrm{X}}_{\mathrm{1}}{·\mathrm{X}}_{\mathrm{2}}\mathrm{+}{\mathrm{a}}_{\mathrm{11}}\mathrm{·}{\mathrm{X}}_{\mathrm{1}}\mathrm{·}{\mathrm{X}}_{\mathrm{1}}\mathrm{+}{\mathrm{a}}_{\mathrm{22}}\mathrm{·}{\mathrm{X}}_{\mathrm{2}}\mathrm{·}{\mathrm{X}}_{\mathrm{2}}$$

(1)

In this equation, Y is the predicted response, a₀ the model constant, a₁ and a₂ the linear coefficients, a₁₂ the interaction coefficient, and a₁₁ and a₂₂ the quadratic coefficients [49].

The Box–Wilson design was used to assess the effect of substrate concentration and butyric acid concentration on butanol production. The Box–Wilson design is a response surface methodology relying on evaluating the relationship between observed outcomes and experimental variables. The main steps of this design consist of carrying out statistically designed experiments, determining the model coefficients as a result of the experiment, obtaining prediction results, and finally verifying the adequacy of the model [50,51]. The inspected ranges for the independent variables were substrate concentrations (X₁) = 0–30 g/L and butyric acid concentration (X₂) = 0–4 g/L, respectively. Butanol concentration (Y) was the objective function. The concentration ranges of butyric acid to be added were determined based on preliminary experiments with Thermoanaerobacterium thermosaccharolyticum. To prevent a shock loading on the inoculum, each butyric acid level was added gradually in equal volumes a total of 10 times during 5-day fermentation periods. Butyric acid was added every 12 h, with the first time being at the beginning of the experiment.

3. Results and Discussion

The content of fig waste used in this study is presented in

Table 1. Composition analysis results of figs used in this study.

Parameter	Content (%, w/w)
Total sugar	57
Total organic carbon	30.80
Total Kjeldahl nitrogen	3.55
Total phosphorus	1.24
Lignin (Acid detergent lignin)	7.68
Cellulose	0.45
Hemicellulose	5.49

. As can be seen, the carbon content of fig is 30.8%, and the nitrogen and phosphorus contents are 3.55% and 1.24%, respectively. On the other hand, the lignin ratio (7.68%) is higher compared to the cellulose (0.45%) and hemicellulose (5.49%) content. The total sugar content of the fig used is 57%. When all these are evaluated, it can be said that the fig is rich in carbohydrates and that polysaccharides are found in lesser amounts in the carbohydrate profile. Fourier Transform Infrared Spectroscopy (FT-IR) analysis was performed to get the chemical makeup of dried figs, and the results are shown in Figure 1.

The peaks obtained in the FT-IR spectrum support that the dried figs may contain carbohydrates, proteins, fats, and phenolic compounds. A wide band was observed at 3600–3000 cm⁻¹, indicating the presence of –OH groups, water molecules, and phenolic compounds in the sample [52]. A sharp peak was observed at 2916 cm⁻¹, referring to C-H bonds that are present in carbohydrates and fats [53]. The sharp peak around 1730 cm⁻¹ can be attributed to C=O stretching vibrations in ester, aldehyde, and keton structures found in organic acids, as well as ester compounds [54]. The moderate intensity peak between 1600 and 1500 cm⁻¹ indicates the presence of C=C double-bond stretching vibrations which might be present in aromatic cycles of phenolic compounds [55]. Many peaks of C-H bending and C-O stretching vibrations in carbohydrates and other organic compounds in fig have been monitored between 1500 and 1000 cm⁻¹. This region can also be named as the fingerprint region that helps to determine specific compounds [54]. The most prominent and sharp peak in the spectrum was recorded at 1021 cm⁻¹, where C-O stretching vibrations are usually present [55]. This peak may be an indicator of eter, ester, and alcohol compounds within sugars and organic acids in figs. For N-H stretching vibrations, amid-I band (C=O stretching vibrations), amid-II band (N-H and C-N stretching vibrations), and C-N stretching vibration peak regions around 1650 cm⁻¹, 1550 cm⁻¹, 1175 cm⁻¹, respectively, are indicators [56].

Figure 2 illustrates the time-varying biomass concentration of T. thermosaccharolyticum DSM 571 in the growth media. It took roughly 10 h for the culture to acclimate to the growth medium and surroundings, as shown in Figure 2. Following the 8–10 h lag phase, the culture grew exponentially for about 12 h. The biomass concentration remained slightly stable between the 22nd and 24th h, then increased to about 0.35 g/L, where it reached the stationary growth phase. The culture was harvested for inoculation after the exponential phase corresponding to the 36th h of the growth period. Experimentally measured biomass concentrations and time data were correlated with the logistic growth equation shown in Equation (2) [57] using Minitab (trial version) software. The curve shown as a straight line in Figure 2 was drawn using the Logistic equation, and it was compatible with the curve obtained using experimental points with an accuracy of 88%. X_∞ and k were calculated as 0.356 g/L and 0.283 1/h, respectively.

$$\mathrm{X}\mathrm{=}\frac{{\mathrm{X}}_{\mathrm{0}}{\mathrm{e}}^{\mathrm{k}\mathrm{t}}}{\mathrm{1}\mathrm{-}\frac{{\mathrm{X}}_{\mathrm{0}}}{{\mathrm{X}}_{\mathrm{\infty }}}\left(\mathrm{1}\mathrm{-}{\mathrm{e}}^{\mathrm{k}\mathrm{t}}\right)}$$

(2)

where X was the biomass concentration at any time (g/L), X₀ was the initial biomass concentration (g/L), X_∞ was the carrying capacity (g/L), k was the carrying capacity coefficient (1/h), and t was time (h).

During butanol fermentation by C. acetobutylicum, butyric acid production takes place in the acidogenic phase. The main products are acetate, butyrate, CO_2, and H₂. The ratio of acetate to butyrate is approximately 1:2. The production of acetoin and lactate as by-products can also be observed in this fermentation [30]. The organic acids formed lower the pH of the medium. Depending on the pH drop, the acids formed and excreted by the cell are taken back into the cell and converted into solvents like acetone, ethanol, and butanol [32]. A similar situation is observed in Thermoanaerobacterium, a thermophilic bacterium. The pH drops, and acetone and butanol are formed, depending on the amount of acetic acid and butyric acid produced [39]. For butanol production, butyric acid must be present in the medium. In this study, since we were not able to determine the organic acid profile and other solvents like acetone and ethanol, butyric acid was externally added by neglecting any microbial butyric acid contribution.

Table 2 shows the experimental results and model predictions in terms of butanol concentration, obtained from the Box–Wilson experimental design, where fig concentration (X₁) and butyric acid concentration (X₂) are independent variables. X₁ was varied in the range of 0–30 g/L and X₂ in the range of 0–4 g/L, respectively. The axial and factorial points were conducted in duplicate, and since the results were consistent, their averages were used in the evaluation. To enhance the experimental reliability, the center point was conducted in triplicate. Table 2 shows that the lowest butanol production was 0.19 g/L and the highest butanol production was 0.36 g/L at the central point with 15 g fig/L and 2 g/L butyric acid concentrations. Measurements performed in triplicate at the center point exhibited a standard deviation of 0.0058, demonstrating minimal variability and thereby confirming the high consistency and reliability of the experimental data. The model predictions and experimental butanol concentrations were found to be fairly consistent.

Table 2. The Box–Wilson design matrix, along with the experimental results and model predictions.

	X1	X2	Y
	Fig Concentration (g/L)	Butyric Acid Concentration (g/L)	Butanol Concentration (g/L)
			Experimental	Prediction
Axial points
A1	0	2	0.27	0.29
A2	30	2	0.2	0.23
A3	15	0	0.19	0.22
A4	15	4	0.14	0.15
Factorial points
F1	25.6	3.42	0.24	0.22
F2	25.6	0.58	0.22	0.19
F3	4.39	3.42	0.19	0.18
F4	4.39	0.58	0.33	0.31
Center point
C1	15	2	0.35	0.36
C2	15	2	0.36	0.36
C3	15	2	0.36	0.36

Table 2. The Box–Wilson design matrix, along with the experimental results and model predictions.

	X1	X2	Y
	Fig Concentration (g/L)	Butyric Acid Concentration (g/L)	Butanol Concentration (g/L)
			Experimental	Prediction
Axial points
A1	0	2	0.27	0.29
A2	30	2	0.2	0.23
A3	15	0	0.19	0.22
A4	15	4	0.14	0.15
Factorial points
F1	25.6	3.42	0.24	0.22
F2	25.6	0.58	0.22	0.19
F3	4.39	3.42	0.19	0.18
F4	4.39	0.58	0.33	0.31
Center point
C1	15	2	0.35	0.36
C2	15	2	0.36	0.36
C3	15	2	0.36	0.36

Table 3. Model summary statistics, Analysis of variance (ANOVA), and coefficients.

MODEL SUMMARY
R	R2		R2adj		Std. Error	F	df1		df2		p
0.962	0.925		0.850		0.03074	−0.0004	−0.043		0.90		0.008
ANOVA
Model		Sum of Squares		df		Mean Square		F		p
Regression		0.058		5		0.012		12.314		0.008
Residual		0.005		5		0.001
Total		0.063		10
COEFFICIENTS
Model		Unstandardized Coefficients		Std. Error		Standardized Coefficients Beta		t		p
(Constant)		0.230 (ao)		0.047				4.929		0.004
X1		0.006 (a1)		0.004		0.715		1.446		0.208
X2		0.112 (a2)		0.031		1.789		3.612		0.015
X1×X1		−0.0004 (a11)		0.000		−1.649		−3.820		0.012
X2×X2		−0.042 (a22)		0.006		−2.821		−6.530		0.001
X1×X2		0.003 (a12)		0.001		0.955		2.603		0.048

summarizes the results of the statistical analysis performed using IBM SPSS Statistics 31 software. The p-value of 0.008 (<0.05) and a high F value of 12.31 show that the model is significant. A strong positive correlation was observed, with an R value of 0.962 and a corresponding R² of 0.925, indicating that approximately 92.5% of the variance in the dependent variable is explained by the model. The adjusted R² value of 0.850 indicates that the model explains approximately 85% of the variance in the dependent variable, taking into account the number of independent variables, thereby demonstrating good model fit and strong explanatory power. Model function coefficients are shown at the bottom in Table 3. In addition, standardized coefficients, t, and p values for each independent variable are also presented. Accordingly, when the p-values are examined, the fact that the p-value is below 0.05 in all conditions except when the fig concentration is alone shows that it has a significant effect on butanol production. The impact of the linear, interaction, and quadratic concepts in the model on butanol concentration can be seen better when the standardized coefficients are examined. When these coefficients are examined together with their p values, it is seen that the highest significant effect was X₂×X₂, the butyric acid quadratic effect (p = 0.001, Beta: −2.821), followed by X₂, the butyric acid linear component (p = 0.015, Beta = 1.789), and X₁×X₁, the fig quadratic component (p = 0.012, Beta = −1.649). Although X₁×X₂, the contribution of the interaction component (p = 0.048, Beta = 0.955), is also significant, it is understood that X₁, the fig linear component, did not have a significant contribution to butanol production (p = 0.208, Beta = 0.715). From this analysis, it can be said that the components of butyric acid concentration in the function make a more significant and effective contribution to butanol production than the components of fig concentration. The effect of the quadratic component of fig concentration was non-negligible, and the interaction component, in which both fig and butyric acid concentrations were evaluated together, also made a significant contribution. Figure 3a,b was drawn to better understand the effects of the independent variables on the objective function.

In butanol production via thermophilic fermentation, the process occurs through a two-stage pathway. In the first stage, sugars are converted to carboxylic acids, predominantly butyric acid. In the second stage, the produced butyric acid is converted to butanol via the following pathway: butyric acid → (via ctfAB) butyryl-CoA → (via adhE) butyraldehyde → (via bdhAB/adhE) butanol [58,59].

For this study, the following steps are used to explain how butanol is formed from waste figs during thermophilic fermentation:

waste fig → enzymatic hydrolysis → mono and oligosaccharides → butyric acid within fatty acids → butanol

The polysaccharides in figs are converted into monosaccharides and oligosaccharides by enzymatic hydrolysis in the first step. When the fermentable sugars are transformed into mixed fatty acids, butyric acid is microbially reduced with the reducing power, which yields butanol. Thus, for effective butanol formation, butyrate and fermentable sugars must be present in sufficient and balanced amounts in the fermentation environment. In a typical fermentation, it may not be possible for microorganisms to selectively follow such a path and focus on butanol formation, since fermentation conditions, organic acid profile, reducing power, and thermodynamic conditions can push the organism to form other solvents and ketones, such as ethanol or acetone. Butyric acid has an intermediate precursor role in butanol formation. Since butanol production proceeds through a two-step fermentation process in which butyric acid is first formed and subsequently reduced to butanol, exogenous butyric acid was added to the medium in this study to simulate the natural pathway and assess its contribution to butanol formation. The butyric acid concentration to be added was given to the fermentation medium gradually rather than with a single injection to prevent microorganisms from being exposed to shock butyric acid loading, to prevent a sharp pH drop, and to understand the appropriate butyric acid feeding regime. Since butyric acid is not formed suddenly during butanol fermentation but is released over time and reduced to butanol, we aimed to try to mimic this natural process and see the possibility of directing it. So far, studies on thermophilic butanol fermentation at different ratios of glucose (G) to butyrate (B) [35,60] have been reported in the literature, and a butyrate-to-glucose (B/G) ratio of 0.5 has been addressed to be the ideal ratio [35].

To examine the selected parameters over the full range, a model equation was created using the coefficients obtained from the Box–Wilson analysis, and the graphs in Figure 3 were plotted. Figure 3a shows the variation of butanol concentration with fig concentration at different fixed butyric acid concentrations. The highest butanol production occurred at 2 g/L butyric acid. In general, butanol concentration increased to about 10–20 g/L, fig concentration range, except at 0 g/L butyric acid concentration because of substrate limitation, and it decreased at fig concentrations higher than 20 g/L due to substrate inhibition. At substrate concentrations higher than 20 g/L, the bacteria may switch to another microbial activity instead of butanol production. On the other hand, at 0 g/L butyric acid concentration, butanol peaked at about 7 g/L fig concentration. It can be said that while the highest butanol production occurred at 1.6 g/L butyric acid and 16 g/L fig concentration, the lowest butanol production was obtained when fig and butyric acid concentrations were 30 and 0 g/L, respectively. The second lowest butanol concentration was found at a concentration of 4 g/L butyric acid and 30 g figs/L. A better view of the effect of butyric concentration at fixed fig concentrations is depicted in Figure 3b.

At all fig concentrations in Figure 3b, butanol concentration increased with the increase in butyric acid concentration up to about 2 g/L due to butyric acid limitation and then decreased at higher butyric acid levels, probably because of butyric acid inhibition. In the experiment where no fig was added, butanol was produced because butyric acid, added externally, was reduced and converted into butanol. This showed that the most convenient butanol production could be achieved by gradually feeding butyric acid at around 2 g/L. Where both fig and butyric acid are present, higher butanol formation is observed, except for 30 g/L fixed fig concentration, indicating that a harmonic composition of fig and butyric acid contributed well to butanol formation. For all fig levels examined, at a fixed butyric acid concentration of 2 g/L, the fig/butyric acid ratio (w/w) was in the range of 0–15, and the highest butanol formation was obtained at 15 g/L fig, where this ratio was 7.5 (w/w).

When Figure 3a,b are evaluated together, it can be said that butanol production increases when fig and butyric acid concentrations are close to the midpoints of the inspected ranges. As the values for fig and butyric acid approach the extremes, especially at high concentrations, butanol production is the lowest.

Triplicate confirmation experiments were conducted at 16 g fig/L and 1.6 g butyric acid/L, which produced the highest butanol formation, in order to assess the model’s dependability. Figure 4 shows the variation in butanol, total volatile fatty acids (TVFA), and total sugar (TS) concentrations with time for the confirmation experiment and variation in pH and ORP. Figure 4a shows the TS concentration was 9.12 g/L at the beginning and decreased to 0.34 g/L at the end, indicating effective consumption of TS by T. thermosachharolyticum. TS was first converted to fatty acids and then to solvents. During the experimental period, 1.6 g/L butyric acid was added 10 times in equal amounts. When the figure is examined, it is seen that the TVFA concentration increases regularly and rises to 1.94 g/L at the end of the experiment. Since the volatile fatty acid (VFA) type could not be monitored during the course of the experiment, it is not shown in the figure. If there was no external butyric acid addition, we would be able to see that butyric acid would first be formed and then converted into butanol. We are unable to see this rise or fall in butyric acid here because TVFA stands for total volatile fatty acids. Butanol concentration increased quite rapidly in the first 2 days and reached 0.32 g/L at the end of the experiment. The predicted and experimental results for butanol concentration were 0.36 and 0.32 g/L, respectively, indicating 89% reliability of the model. The butanol yield was 0.034 g butanol/g TS.

In addition to the studies, experiments were carried out with dried figs from the market without aflatoxin under optimum experimental conditions. In the experiment where 16 g/L dried figs were used and 1.6 g/L butyric acid was added in equal volume a total of 10 times, a butanol concentration of 0.45 g/L was obtained. This result showed that aflatoxin content had an inhibitory effect on butanol production and reduced butanol production by about 30% compared to the experiment with figs without aflatoxin.

Figure 4b shows the variation in pH and ORP over the 5-day test period. The initial pH value was set to 7. The pH was measured and recorded every 24 h during the experimental period and kept at around 7. The pH value decreased in the first days due to VFA formation, and at the end of the second day, the pH value was less than 5. After day 2, the decrease in pH was less, and the experiment was complete at pH 6 at the end of day 5. The ORP value varied between −100 mV and −200 mV, indicating anaerobic conditions.

Table 4. Comparison of reported butanol production studies by strains of Thermoanaerobacterium thermosaccharolyticum.

Bacterial Strain	Genotype	Substrate	Substrate Concentration, g/L	Temperature, °C	Butanol Concentration, g/L	Ref.
T. thermosaccharolyticum DSM571	non-GM strain	Starch	20	58	1.24	[37]
T. thermosaccharolyticum strain 021	non-GM strain	Starch	20	58	1.98	[37]
T. thermosaccharolyticum TG57	non-GM strain	Cellulose	30	53	1.93	[62]
T. thermosaccharolyticum TG57	non-GM strain	Xylose	30	53	3.63	[62]
T. thermosaccharolyticum GSU5	GM strain	Glucose	10	60	0.33	[63]
T. thermosaccharolyticum GSU5	GM strain	Xylose	10	60	0.26	[63]
T. thermosaccharolyticum DSM571	non-GM strain	Waste Fig	16	55	0.32	This study

summarizes the reported studies on butanol production by Thermoanaerobacterium thermosaccharolyticum, including results obtained with both genetically modified (GM) and non-genetically modified (non-GM) strains. While metabolic engineering provides an effective approach to enhance the yield of target products using GM organisms, non-GM organisms are generally considered more reliable due to their ability to provide more stable and consistent production compared to their GM counterparts [61]. Freier-Schröder et al. [37] obtained the highest butanol concentration of 1.26 g/L with the natural strain Thermoanaerobacterium thermosaccharolyticum DSM 571 from 20 g/L starch. In the same study and under the same conditions, the obtained butanol was 1.92 g/L with the genetically modified strain. Using cellulose, a non-food substance, as a substrate, Li et al. [62] reported that Thermoanaerobacterium thermosaccharolyticum TG57, isolated from waste generated after mushroom collection, produced 1.93 g/L butanol as the single final product. Peña et al. [63], discovered that Thermoanaerobacterium thermosaccharolyticum GSU5, isolated from animal manure, is capable of producing butanol. The highest butanol production was 0.33 g/L when 10 g/L glucose was used as a substrate, followed by 0.26 g/L butanol when 10 g/L xylose was used as a substrate. Thermoanaerobacterium thermosaccharolyticum DSM 571 was also used in the study, where a butanol concentration of 0.034 g/L was obtained by using 10 g/L glucose as a substrate. In this study, 0.32 g/L butanol production was achieved by Thermoanaerobacterium thermosaccharolyticum DSM 571 from 16 g/L waste figs and gradually fed butyric acid of 1.6 g/L. Compared to similar studies in the literature, it can be said that butanol production at this level is reasonable under thermophilic conditions. However, making a one-to-one performance comparison is not correct due to the difference in experimental conditions and cultures used. The reason for low butanol formation might be the result of compounds undefined in this study in the fig or because of a shift in the microbial pathway.

4. Conclusions

Butanol production by thermophilic fermentation was studied using Thermoanaerobacterium thermosaccharolyticum DSM 571. The effect of fig and butyric acid concentration on butanol formation was assessed using a Box–Wilson statistical experiment design, and the most convenient conditions were determined accordingly. The high sugar, vitamin, and mineral content of dried fig waste makes it a potential substrate for fermentation. However, it was found that fig alone was not enough for obtaining effective butanol formation. When it is mixed with the proper amount of butyric acid, then butanol formation increases at the same time. However, we observed that gradual feeding of butyric acid could be used to prevent a shocking load of butyric acid and a fast drop in pH. By gradually supplying the right amount of butyric acid, the microbial culture was able to oxidize the media’s accumulated reducing power. The important thing here is to ensure the balance of supply and demand. The presence of butyric acid, appropriate to the amount of electrons and protons released during the decomposition of figs, allows the reducing power to reduce butyric acid to butanol. Of course, microbial conditions must be suitable for this trail to occur. Deficiency or excess of fig or butyric acid concentration may cause limitations or inhibitions. When the literature is examined, it is seen that thermophilic butanol production studies generally report butanol production from cultures subjected to genetic modification in rich nutrient media. This study investigated butanol production from waste figs using a non-genetically modified culture. When the results are examined, it is seen that the butanol concentration did not reach the inhibition levels reported for mesophilic ABE fermentation. In fact, simultaneous removal of butanol from the medium during ABE fermentation is recommended to eliminate this inhibition. Therefore, in situ removal may be a useful way to obtain higher butanol titers. To our knowledge, no study on butanol production from fig waste by thermophilic fermentation has been reported in the literature. At the same time, the number of studies on butanol production using Thermoanaerobacterium thermosaccharolyticum DSM 571 bacteria is also very limited. Therefore, the results of this study provide sound information for thermophilic butanol production. The experimental results of this study show that high concentrations of fig and butyric acid lead to inhibition, while low concentrations of fig and butyric acid cause limitation. Butyric acid did have a more profound effect on butanol formation than fig alone, but the presence of both compounds produced better results. The most convenient butanol production of 0.32 g/L was obtained when 16 g fig/L was gradually fed with 1.6 g/L butyric acid. The model generated by the Box–Wilson design was proven with 89% reliability.