Bioethanol: A New Synergy between Marine Chitinases from Bacillus haynesii and Ethanol Production by Mucor circinelloides

Abstract

The fourth generation of bioethanol production is on a lookout for non-lignocellulosic biomass waste. One such candidate is chitin, the second most abundant biopolymer on earth. However, the crystalline nature of chitin hinders its application potential for bioethanol production. This limitation can be circumvented by hydrolysing this polymer into oligomers using chitinases. We used this hypothesis and isolated a Bacillus haynesii, a marine bacterium that utilizes colloidal chitin as a substrate and produces chitin oligosaccharides. Further, we utilized Mucor circinelloides to produce bioethanol using the chitin oligosaccharides in the shake flask. We investigated the effect of inoculum age, filling volume, different substrates, and substrate concentration on bioethanol production using Mucor circinelloides from Bacillus haynesii-produced chitin oligosaccharides. Bacillus haynesii demonstrated a maximum chitinase activity of 3.08 U/mL with specific activity of 96 U/mg at the 90th h. Chitin oligosaccharides produced by Bacillus haynesii were confirmed using mass spectrometry. Bioethanol concentration was determined using dichromate oxidation assay as well as gas chromatography. The research resulted in 7.4 g/L of ethanol from 30 g/L of chitin oligosaccharides, with a maximum ethanol yield of 0.25 g of ethanol/g substrate at the 55th h with 48 h inoculum in 80 mL of fermentation medium. Results suggest that chitin oligosaccharides from Bacillus haynesii are an effective and renewable substrate for bioethanol production.

1. Introduction

Renewable energy resources aid in alleviating climatic change and securing the access to energy. The primary energy consumption was estimated as 11,295 Mt annually in 2008. Usage of fossil fuels remains a global threat for climatic change and has a major impact on nature and human system. Global warming due to greenhouse gas emissions could be fortunately altered if fossil fuels are replaced by renewable resources. Bioethanol, an alternative fuel, serves as a superior substitute for fossil fuels and can meet energy demands if produced in sufficient quantities. In 2016, the global bioethanol production was 100.2 BL. It is estimated that the global bioethanol production and its consumption will reach 134.5 BL by 2024. For bioethanol production, raw materials such as sugarcane and starch containing feed stocks were explored. It is known that raw material sums up to 40–75% of the total production cost [1,2,3]. As an initiative, the first-generation bioethanol from food crops sparked the food-versus-fuel debate. In addition, large land and water resources were required for these crops. To address the limitation, the second-generation biofuels were based on non-agricultural crops, agricultural residues, and wood-processing waste. As a result, in 2017, United States increased cellulosic bioethanol production to 38 ML [4]. Bioethanol production from feedstock such as corn stover, barley straw, and corn meal was 5.85 L/kg [5], 0.054 L/kg [6], and 9.67±0.11 L/kg [7] of dried biomass, respectively. Mithra et al. reported a bioethanol yield of 34–44 g/L from lignocellulos-starch biomass by hydrolysis followed by fermentation in fed-batch mode [8]. Annually, 5–8 Mt of lignocellulosic biomass is generated from agriculture and deforestation, but the generated biomass is lower than the consumption of crude oil per annum [9]. In addition, second-generation biomass requires a pre-treatment process in order to convert the rigid recalcitrance structure of lignocellulosic biomass to fermentable sugar [10]. In Brazil, for a conventional ethanol production plant, the electricity consumption is about 28 kWh/t of canes, and steam consumption (2.5 bar) is about 372 kg/t of canes [11]. Hence, second-generation biofuels involve high capital cost and complicated processing equipment.

The quest for fresh avenues for bioethanol production has led to exploration of alternatives, leading to the third- and fourth-generation bioethanol producers.

The insoluble chitin polymer is gaining importance in bio-refinery, as it can serve as an inherent substrate for production of ethanol. Kamal et al. reported that insect-based chitin for bioethanol production could be an excellent alternative for energy crop-based raw materials. Chitin can be a potential cheap and renewable source for bioethanol production [12]. Chitin, the second most abundant renewable polysaccharide, is composed of N acetyl-D-glucosamine residues linked with β-(1,4) glycosidic bonds. It is a major component in shrimps, gut lining of insects and arthropods, crab shells, marine invertebrates, and cell wall of fungi and in nematode eggs [13,14,15]. While 70% of the total body weight of shellfish is considered waste, 20–58% of the said waste contains chitin [16]. As an insoluble polymer, the disposal of marine waste from food-processing industries remains challenging. It is estimated that around 10¹¹ t of chitin is produced annually in biosphere [17]. Degradation of chitin into soluble chitin oligomers by chitinase enzyme from marine bacteria serves as a promising solution for the disposal of chitin wastes as well as the production of valuable products. Interestingly, in marine sediment, the chitin content is less, and the reason behind this is the utilization of chitin by marine bacteria possessing chitinolytic enzymes.

Chitinolytic enzymes belong to the glycosyl hydrolase family, which hydrolyse the β-1,4-glycosidic bonds between N- acetyl- D- glucosamine residues in the chitin chain, resulting in chitin oligosaccharides as products. Hence, chitinase-producing marine bacteria help in the bioconversion of chitin to organic compounds, which can serve as a natural carbon and nitrogen source for other organisms [18]. In addition to this, marine bacterial chitinases possess activity in wide range of pH and temperature. Studies conducted on chitinase from a marine bacterium, Microbulbifer sp. BN3, have reported stable activity at temperatures up to 50 °C and pH range of 4.0–9.0 [19]. The chitinase from Pseudoalteromonas sp. DL-6 [20] and Alteromonas sp. 0–7 [21] have exhibited activity at pH 8 with specific activity of 42.17 U/mg and 4.5 U/mg, respectively. Other examples of marine bacterial chitinase reported include Moritella marina [22], Bacillus sp. R2 [23] and Micrococcus sp. AG84 [24]. An earlier study by our group reported chitinase from Bacillus aryabhattai with a pH optimum at 7. The enzyme exhibited activity of 146.2 U/mL, 114.9 U/mL, and 175.4 U/mL with chitin powder, chitin flakes, and colloidal chitin, respectively [25].

To incorporate chitin-based biorefinery, the selection of a micro-organism capable of utilizing the chitin and its oligosaccharides is important. Production of bioethanol from chitin requires enzymatic degradation of chitin to N-acetyl glucosamine (NAG), followed by fermentation of NAG. Endo chitinases from micro-organisms randomly cleave the chitin and result in chitin oligosaccharides. Inokuma et al. reported that 18.6 g/L and 16.9 g/L of ethanol was produced from 50 g/L of NAG using Mucor circinelloides NBRC6746 and Mucor ambiguous NBRC8092. M. circinelloides is known to produce bioethanol from NAG and chitin waste [12,26]. Until now, ethanol production from chitin oligosaccharides has not been reported. Generation of bioethanol from chitin oligosaccharides will pave the way for the utilization of waste to create value-added product. In a commercial production of bioethanol, incorporating micro-organisms will result in reducing fermentation steps, which in turn impacts the capital cost of fuel production. In this study, an enzymatic process is employed for chitin oligosaccharide production from chitin using marine bacterium chitinase. The produced chitin oligosaccharides were utilized for the production of ethanol using Mucor circinelloides, and the impact of process parameters such as inoculum age, filling volume, effect of different substrates, and substrate concentration on ethanol production was investigated.

2. Materials and Methods

2.1. Chemicals

The following chemicals were purchased from Hi media (Thane, India); agar (GRM026); chitin flakes from shrimp shells (GRM1356); chitin powder (GRM10909); phenol crystal (AS022); sulphuric acid (AS015); NaCl (MB023); KCl (MB043); CaCl₂.2H₂O (MB034); MgCl₂.6H₂O (MB040); MgSO₄.7H₂O (GRM683); M9 minimal medium salts (5X) (G013); Tris Cl pH 8 (ML013); 3,5-dinitro salicylic acid, A.G (GRM-1582); potassium dichromate, A.R (GRM699); and glucosamine hydrochloride (TC129). Chitin oligosaccharides (C2762) were purchased from TCI chemicals (Tokyo, Japan), and we also used NaOH (28151190, Loba, Mumbai, India); N acetyl glucosamine AR, 99% (59012, SRL chemicals, Mumbai, India); and ethanol, absolute, A.G.

2.2. Colloidal Chitin Preparation

Colloidal chitin was prepared according to the method described by Murthy et al., 2012, with 5 g of chitin flakes in 60 mL of concentrated HCl (37% v/v). The mixture was stirred for 1.5 h–2 h at 3 °C to obtain a homogenous solution. The solution was subjected to filtration using a muslin cloth to remove the undissolved particles, and the obtained filtrate was added to 800 mL of pre-chilled water and stored at 4 °C overnight. After filtration, the precipitate was collected and centrifuged at 10,000 rpm for 10 min. The collected pellet was washed with distilled water, and pH was adjusted to 7 using 4 N NaOH, and this made up to 100 mL and was stored at 4 °C for further use [27].

2.3. Quantification of Chitin Content in Colloidal Chitin

Chitin concentration was determined according to Henriques et al., 2020. Different concentrations of chitin powder ranging from 1 mg to 20 mg were taken in 2 mL microfuge tubes. Then, 400 µL of distilled water and phenol (5% w/v) were added; 1 mL of concentrated sulphuric acid was added, and the resultant mixture was incubated at 30 °C for 10 min. The tubes were mixed well and incubated further for 10 min. The absorbance of the solution was measured at 490 nm using the UV spectrophotometer. A standard graph was plotted between chitin weight and absorbance at 490 nm. The chitin content in the colloidal chitin was determined using Equation (1):

$$\mathit{Chitin}\left(\mathrm{mg}\right)=\frac{\mathit{Absorbance} \mathit{at} 490 \mathrm{nm}}{0.1037}$$

(1)

Similarly, 400 µL of colloidal chitin was taken, and its chitin content was calculated from the standard graph using the same procedure [28].

2.4. Screening and Isolation of Marine Bacteria

Deep-sea sediment sample was collected in a sterile container from the Arabian Sea at a depth of 40 m, with 12°48’ N and 74°40’ E as co-ordinates using an Ekman grab sampler. The microbes were isolated using the serial dilution plating technique and purified with four rounds of re-streaking [29]. These isolates were screened for chitinase activity by plating on M9 minimal media with artificial seawater (MgSO₄.7H₂O, 0.99 g/L; CaCl₂, 0.25 g/L; MgCl₂. 6H₂O, 1.5 g/L; NaCl, 26.29 g/L; KCl, 0.74 g/L) supplemented with 1% colloidal chitin. Artificial sea water was prepared according to Cold Spring Harbor protocol [30]. Agar at a concentration of 1.5% was used as a solidifying agent. The Petri plates were incubated at 37 °C for 48 h and flooded with Lugol’s iodine solution to visualize the chitinase-producing microbes. The positive isolates were subjected to biochemical characterization according to Bergey’s manual of determinative bacteriology [31]. Further, 16S rRNA molecular characterization was performed.

2.5. Production and Quantification of Chitin Oligomers

Colloidal chitin served as a sole carbon source as well as an inducer for chitinase production. Pre-inoculum media for chitinase production of minimal salt (M9) (Na₂HPO₄, 6.7 g/L; KH₂PO_4, 3 g/L; NaCl, 0.5 g/L; NH₄Cl, 1 g/L), supplemented with 1% colloidal chitin, was prepared using artificial sea water. Artificial sea water and M9 media with 1% colloidal chitin were separately sterilized and mixed after sterilisation. Chitin oligosaccharides were produced by enzymatic reaction with 1% colloidal chitin as substrate in the presence of chitinase from the screened bacterium. After enzymatic reaction, the reaction mixture was centrifuged at 10,000 rpm for 10 min. The supernatant was filtered through 3 KDa filters (Pall Filters, Grand Island, NE, USA) and subjected to centrifugation at 7000 rpm for 15 min. The resulting chitin oligosaccharides were quantified by DNSA and analysed using mass spectrometry (Xevo QToF, Waters, Milford, MA, USA).

2.6. Ethanol Production from Chitin Oligosaccharides Using Mucor Circinelloides

Chitin oligosaccharides produced from enzymatic reaction were used as substrate for ethanol production. The reaction mixture was inoculated with the ethanol-producing microorganism, Mucor circinelloides, a dimorphic microorganism that uses chitinous products as a carbon source. The spore suspension (1 ± 0.16 × 10⁶ spores/mL) was prepared by inoculating freeze-dried fungus Mucor circinelloides on potato dextrose agar slant at pH 5.6 for 96 h at 25 °C. Then, 1 mL of sterile distilled water was added to the slant and shaken vigorously for collection of fresh spore suspension. After spore suspension inoculation, preculture medium was cultured aerobically under shaking conditions (120 rpm) at 25 °C. After 2 days, mycelial biomass was harvested from culture medium and washed with isotonic water under aseptic condition, which later was used as an inoculum for ethanol fermentation. The harvested fungal biomass was directly transferred to the enzymatic reaction mixture in the 100 mL screw-cap bottle and cultured micro-aerobically under shaking conditions (120 rpm) at 25 °C. Further, the samples were centrifuged at 7000 rpm for 5 min at 4 °C, and the supernatant was concentrated using rotary vacuum evaporator (Hei-VAP-core, Heidolph, Schwabach, Germany) and used for the ethanol determination. Influence of different substrates, volume of fermentation medium, substrate concentration, and inoculum age on ethanol production were investigated.

2.7. Chitinase Assay

Chitinase activity was assayed using 1% colloidal chitin as substrate based on the estimation of reducing sugar. Enzyme and substrate were taken in the ratio of 1:1 and incubated with 1 mL of Tris- Cl buffer (10 mM, pH 7) at 37 °C for 60 min. The same procedure was followed for the enzyme blank and substrate blank. The reaction mixture was centrifuged at 10,000 rpm for 10 min. The obtained supernatant was subjected to dinitro salicylic acid (DNSA) method.

2.8. DNSA

DNSA reagent was added to the supernatant and incubated at 95 °C for 10–15 min. After cooling, absorbance at 595 nm was measured using UV–visible spectrometer (Thermofisher Scientific, Waltham, MA, USA). Concentration of N acetyl glucosamine (NAG) released was calculated using Equation (2) obtained from the standard graph plotted using the N acetyl glucosamine of different concentration (2–10 mM). One unit of chitinase enzyme is defined as the amount of enzyme that released one micromole of N acetyl glucosamine per minute under standard assay conditions [32].

$$\mathit{Concentration} \mathit{of} NAG \left(\mathrm{mM}\right)=\frac{\mathit{Absorbance} \mathit{at} 540 \mathrm{nm}}{0.65}$$

(2)

2.9. Ethanol Quantification

2.9.1. Analytical Method

Mucor circinelloides was separated and washed twice with double-distilled water and kept at 60 °C until reaching constant weight. The biomass was determined by measuring its dry weight.

The sugar concentration was estimated by DNSA method at the absorbance of 540 nm. The ethanol concentration was determined by colorimetric method of dichromate oxidation assay, where 1 mL of sample supernatant was added with 170 µL of potassium dichromate solution (0.1 g/mL), followed by 200 µL of sulphuric acid, incubated at room temperature in dark for 5 min; then, the absorbance was measured at 580 nm. Since dichromate assay can also measure the concentration of reducing sugar in a sample, the following steps were followed to determine ethanol concentration [33].

The ethanol concentration in the fermentation medium was quantified in five steps. Step 1: the concentration of reducing sugar in the sample was calculated by DNSA assay. Step 2: the absorbance value contributed by the respective concentration of reducing sugar in the sample was measured by the dichromate oxidation assay with NAG as standard. This was noted as A. Step 3: the absorbance obtained from the dichromate oxidation assay for the sample was noted as B. Step 4: the absorbance value contributed by ethanol in the sample was calculated by subtracting A from B. Step 5: the ethanol concentration in the sample was calculated by the dichromate oxidation assay standard curve for ethanol, as represented in Equation (3) [34].

$$\mathit{Ethanol} \mathit{concentration} \left(\frac{\mu g}{\mathrm{ml}}\right)=\frac{\mathit{Absorbance} \mathit{at} 580 \mathrm{nm}}{0.2897}$$

(3)

2.9.2. Gas Chromatography

Gas chromatography (Finnican Trace GC ultra, Thermo Scientific, Waltham, MA, USA) equipped with HP-INNOWAX (Agilent Technologies, Santa Clara, CA, USA) capillary column was employed for ethanol quantification. The capillary column length, diameter, and film thickness were 30 m, 0.320 mm, and 0.50 µm, respectively. The GC oven (column temperature) was held at 70 °C for 5 min and increased to 180 °C at the rate of 5 °C/min. A flame ionization detector (FID) was employed. Samples were introduced into the GC column in split mode with the split ratio of 37:1, and the spit flow was 130 mL/min and with the total flow of 151 mL/min. The injector temperature was held at 200 °C while the detector was operated at 260 °C. Hydrogen flow rate was maintained at 40 mL/min. The sample injection volume was 0.1 µL. Nitrogen was used as carrier gas. The total analysis time was 33 min. The ethanol in sample was analysed using Equation (4) by comparing the retention time with the ethanol standards from 0.1% (v/v) to 0.5% (v/v) [35].

$$\mathit{Ethanol} \mathit{concentration}\% \left(\frac{v}{v}\right)=\frac{\mathit{Area} \left(\mathrm{mV}\ast S\right)}{428.95}$$

(4)

2.9.3. Mass Spectrometry (MS) Analysis

Next, 1 µL of sample volume was injected for MS analysis, and its mass spectrum was obtained using mass spectrometer (Xevo QToF, Waters, USA). The source of ionisation was electron spray ionization, and the detector was Xevo G2-XS QTof (Waters, USA). The hydrolysed product of enzymatic reaction was injected to mass spectrometry. Isocratic mobile-phase water:acetonitrile was used in a ratio of 30:70 with the flow rate of 0.1 mL/min. Both the mobile phases contained 0.1% (v/v) formic acid to improve ionization. The detector was regulated with capillary voltage of +5 KV and capillary temperature of 275 °C. Nebuliser pressure and nitrogen flow rate were 100 psi and 10 L/min, respectively. Analysis was performed in both positive ion mode and negative ion mode. Mass spectra was analysed in the range of m/z 50–2000. A full scan was performed for the duration of 5 min.

3. Results and Discussion

3.1. Screening, Isolation, and Identification of Bacillus haynesii

Seven different strains were isolated for chitinolytic enzymes production from the Arabian Sea sediment sample on minimal salts medium supplemented with colloidal chitin as the sole carbon source. The initial qualitative screening with Lugol’s iodine solution hinted towards isolate SEDS2 as the highest producer of the chitinase enzyme (Figure S1); hence, further characterization based on biochemical (Table S1) and molecular identification was undertaken. The biochemical tests revealed that the SEDS2 strain was a motile, Gram-positive bacterium. The isolate showed positive results for catalase, MR-VP, and starch hydrolysis. The strain SEDS2 was fermentative without H₂S gas production and negative for the indole test. 16S rRNA sequence homology was analysed using BLAST algorithm. The BLAST results with the highest score were taken for phylogenetic analysis. The strain was identified as Bacillus haynesii as represented in Figure 1 based on the phylogenetic analysis using MEGA 7.0 software [36]. The strain was submitted to the National Centre for Microbial Resource (NCMR) with the catalogue number MCC 4009.

3.2. Production of Chitin Oligomers

Bacillus haynesii culture was maintained at 37 °C in 120 rpm for the production of chitinase. The enzyme showed highest activity of 3.08 U/mL at the 90th h of pre-inoculum medium at pH 7. Further, the broth containing chitinase enzyme was used for enzymatic reaction, which resulted in a chitin oligosaccharides yield of 2.4 g/L (0.95 g/g), which is more or less in the same range as given in the literature. Wang et al. reported the concentration of N-acetyl glucosamine (NAG) as 2.65 g/L from chitin powder using Chitinolyticbacter meiyuanensis SYBC-H1 [14]. Chitinibacter tainanensis produced NAG from α- chitin and β-chitin at the concentration of 0.75 g/g and 0.98 g/g, respectively [37]. The maximum NAG yield of 12.785 ± 0.77 g/L was reported at pH 7 using Trichoderma harzianum AUMC 5408 using colloidal chitin as substrate [12].

3.3. Mass Spectrometry Analysis

The chitin oligosaccharides produced from the enzymatic reaction were analysed using Xevo G2-XS QTof mass spectrometry. Enzymatic reaction was performed with colloidal chitin as substrate for 6 h, and the supernatant was passed through 3 KDa centrifugal filters and analysed using mass spectrometry. Mass spectra confirmed N-acetyl glucosamine and glucosamine (GlcN) residues were generated from enzymatic reaction of 6 h. Mass spectra of 6 h of enzymatic reaction revealed the presence of N- acetyl glucosamine and glucosamine residues (Figure 2). The glucosamine could have formed by the enzymatic action of chitin deacetylase. The activity of chitin deacetylase was about 4.92 U/ mL.

In positive ion mode, the highest intensity m/z of 164.92 corresponds to NAG monomer with the loss of NHCOCH₃. Kim et al. reported that MS spectrum of chito-oligosaccharides can result in a neutral loss of NH₃ group, a H₂O group, or combination of two groups [38]. The peak at 447.14 m/z corresponded to a dimer of NAG/GlcN with ACN as an adduct. The m/z of 606.53 corresponds to NAG/NAG/GlcN trimer with a potassium adduct. The peak at 1042.29 corresponds to hexamer of NAG/NAG/GlcN/GlcN/GlcN/GlcN with the removal of C₂H₄ group. Kamst et al. reported that an m/z ratio of 1034 represented the presence of five N acetyl glucosamine residues [39].

In negative ion mode, the m/z of 792.40 corresponds to the presence of tetramer of glucosamine and n-acetyl glucosamine with a formic acid adduct. The highest-intensity m/z of 268.78 corresponds to NAG monomer with formic acid as an adduct. The m/z of 620.5300 corresponds to a trimer of NAG/NAG/GlcN residues. The mass spectra of 6 h of enzymatic reaction in positive mode and negative mode are represented in Figure 2. Figure 3 represents the MS spectra of substrate blank or only broth supernatant in positive ion mode and negative ion mode. There are no common peaks between the substrate blank and enzymatic reaction mixture. The mass spectra of substrate blank proved that chitin oligosaccharides produced were from enzymatic reaction and not from broth supernatant. The maximum-intensity peaks of monomers in mass spectrometry results revealed that the chitinase from B. hynesii could be exochitinase. Further, the produced chitin oligosaccharides were used for ethanol production.

3.4. Ethanol Production from Chitin Oligomers: Effect of Various Process Parameters on Ethanol Production

3.4.1. Effect of Inoculum Age on Ethanol Production

Impact of inoculum age on ethanol production is represented in Figure 4. The maximum production was observed using 48 h inoculum, yielding 7.4 g/L ethanol. A further increase in inoculum age to 60 h resulted in decreased ethanol production (7 g/L). The effect of inoculum age on ethanol production using chitin/chitin oligomers as substrate is not reported in the research, to the best of our knowledge. However, a similar observation using grape juice as ethanol-producing substrate was reported by Kaur et al. in Saccharomyces cerevisiae MTCC170 and MTCC11815 strains, where the inoculum age of 48 h resulted in maximum ethanol production [40]. Manikandan et al. investigated the effect of inoculum age on ethanol production using corn flour. They observed a maximum ethanol concentration of 63 g/L after 48 h of inoculation in fermentation medium with 24-hour-old slant using Saccharomyces cerevisiae [41]. This decline in ethanol production after 48 h may be due to the depletion of nutrients. Another reason for the decrease could be hindered cell interactions and space limitations due to high cell densities, growth, and metabolism, as shown in [40,41,42,43].

Thus, for further experiments, the inoculum age of 48 h was maintained.

3.4.2. Effect of Volume of Fermentation Medium on Ethanol Production

Decreasing the oxygen availability in the head space of the system is important to avoid the high degree of aerobic metabolism that utilizes sugar for maintenance and growth, which will eventually block the pathway to produce ethanol. The effect of the volume of fermentation medium on the ethanol production was investigated. The highest ethanol production of 7.6 g/L was achieved after 40th hour in both the 60% medium volume and 80% medium volume when 30 g/L of chitin oligosaccharides was used as a substrate, as shown in the Figure 5. The maximum yield from substrate was 0.15 g of ethanol/g of colloidal chitin. Hence, 80% media volume was taken for further experiments, as overall ethanol quantity is greater with more volume (Figure 6).

3.4.3. Effect of Substrate Concentration on Ethanol Production

In a fermentation process, substrate is a key material for nutrients, and its concentration is important for making the process economical. With lesser substrate concentration, few substrates can bind to the enzyme’s active site, and an increase in substrate concentration can result in increase in product formation. Primarily, the concentration of substrate has an impact on uptake rates and product rate kinetics [44]. Production of a high concentration of bioethanol decreases the energy cost of distillation and hence influences economic efficiency [45]. However, high substrate concentration may result in a low titre of ethanol production due to the production of inhibitory compounds or repression of glycolytic enzymes [45,46,47].

Hence, the experiments were performed with various concentrations of chitin oligosaccharides to enhance the ethanol production. Initial chitin oligosaccharides concentration taken were 10, 30, 50, and 70 g/L in the 80 mL fermentation media. The results are summarized in Figure 5. It shows the change in the ethanol concentration with varying substrate concentration. The ethanol production was affected by substrate concentration between 10–70 g/L. It was observed that the maximum ethanol concentration of 7.52 and 7.6 g/L was produced in the 30 and 50 g/L of substrate. Ethanol fermentation from 30 g/L of substrate at the 40th hour produced the maximum amount of ethanol, with a yield coefficient of 0.25 g of ethanol/g; on the other hand, 50 g/L of substrate produced ethanol with a yield coefficient of 0.152 g of ethanol/g.

The obtained results were confirmed by gas chromatography. Under the optimized conditions, the major peak at the retention time of 15.16 min corresponds to ethanol. The ethanol peak was observed at all concentrations of ethanol from 0.1% (v/v) to 0.5% (v/v). The standard graph was plotted based on the peak area for each standard. Supplementary Figure S2 represents the gas chromatogram of 1% (v/v) ethanol. For each sample, the concentration of ethanol was calculated from the standard graph using the peak area. An ethanol peak with the peak area of 324.8209 was observed from fermented medium with 30 g/L chitin oligomer as substrate, which corresponds to 7.4 g/L of ethanol. Figure 7 represents the gas chromatogram of ethanol production from 30 g/L of chitin oligosaccharides using M. circinelloides (Figure 7A). Lubbehusen et al. reported ethanol production of 0.3 g/L h from glucose by M. circinelloides [48]. As the substrate concentration increased, the ethanol production was affected. The gas chromatogram of ethanol production from 50 g/L of chitin oligosaccharides using M. circinelloides is represented in Figure 7B. Since there was no further improvement in ethanol production, we used 30 g/L of substrate as the initial substrate concentration for the further experiments. A further increase in concentration decreased the ethanol conversion efficiency, which might be due to the feedback inhibition imposed by ethanol. A similar study was conducted by Mardawati et al., who found 0.25 g/L of ethanol production with 9% oil palm empty fruit bunches concentration, but a further increase in substrate concentration resulted in lowering this ethanol yield [46]. There are no reports on the effect of chitin/chitin oligomer substrate concentration on the ethanol production, to the best of our knowledge. However, the effect of substrate concentration on ethanol production is reported using other different substrates. Triwahyuni et al. investigated the effect of substrate concentration (15%, 20%, and 25%) on bioethanol production from oil palm empty fruit bunches by solid-state fermentation. Substrate concentration of 25% resulted in decreased ethanol production (45.50 g/L) as compared with ethanol production of 78.25 g/L for 20% substrate concentration [45]. Increase in glucose concentration had negative impact on ethanol yield, as it caused catabolite repression [49,50,51,52].

3.4.4. Effect of Different Substrates on Ethanol Production

In the co-culture micro fermentation set-up, the chitin hydrolysates produced by B. haynesii were used for the ethanol production by M. circinelloides. Thus, we wanted to probe whether the fungus in an anaerobic condition can utilize chitin or its hydrolysates for ethanol production. Patil et al. reported that incorporating chitin supplements in media led to shortened fermentation time and indirectly reduced the cost of ethanol production [53]. To partially simulate the conditions of the co-culture, the M. circinelloides were fed with different substrates including colloidal chitin, chitin powder, chitin oligosaccharides, and NAG. In microaerobic fermentation, M. circinelloides produced 7 g/L of ethanol from chitin oligosaccharides. The theoretical maximum ethanol yield from NAG by the catabolic and glycolysis pathway via fructose-6-phosphate to produce 2 mol of ethanol per mol of NAG consumed is 0.417 g of ethanol/g of NAG consumed [26]. Among all four substrates, NAG and chitin oligosaccharides gave the higher ethanol production within the 55th hour. This was in tune with the products formed after the first enzymatic hydrolysis by B. haynesii chitinases. Inokuma et al. reported that 14.5 ± 0.2 g/L and 16.9± 0.2 g/L of ethanol was produced from M. circinelloides NBRC 4572 and M. ambiguous NBRC 8092 from NAG at 72 h and 96 h, respectively [26]. Wendland et al. reported that ethanol production from NAG using genetically engineered Saccharomyces cerevisiae resulted in 3 g/L of ethanol after 11 days of fermentation [54]. Kamal et al. reported the production of about 11.9 g/L bioethanol on the sixth day using M. circinelloides (AUMC 6017) from a low-cost chitin source, namely insect waste [12]. Cody et al. reported that an ethanol production of 598 µg/mL was achieved using 10 mg/mL of glucosamine using Z. mobilis [55].

In the present study, the maximum ethanol production from colloidal chitin (CC) and chitin by M. circinelloides was approximately 0.57 g/L of ethanol after the 60th hour, and ethanol yield from substrate was 0.01 g of ethanol/g of colloidal chitin and 0.47 g/L of ethanol from chitin after the 55th hour. The ethanol yield from the polymeric substrate was 0.009 g of ethanol/g of chitin, as can be observed from Figure 8. It is evident from the results obtained that M. circinelloides can also produce ethanol from chitin as well as colloidal chitin by microaerobic fermentation but with reduced productivity as compared to chitin oligosaccharides and NAG. This reduction in the ethanol yield could be due to negligible chitinase activity in the anaerobic condition (less than 0.1 unit per mL). Hence, M. circinelloides could be used in conjunction with B. haynesii to improve the ethanol yield.

4. Conclusions

Chitinase enzymes from marine bacteria Bacillus haynesii serve as an excellent candidate for production of chitin oligomers from chitin. MS results show the conversion of chitin into chitin oligomers. Chito oligosaccharides produced by Bacillus haynesii were utilised for ethanol production, which resulted in an increase in ethanol production fermentation efficiency by 2-fold from 30 g/L of substrate. Its maximum ethanol yield production was 0.25 g of ethanol/g of colloidal chitin at the 55th hour. However, the observed ethanol production from chitin substrate was very low due to its reduced availability of chitinolytic enzyme activity. The impact of various process parameters such as inoculum age, filling volume, different substrates, and substrate concentration were explored. Therefore, in order to increase ethanol production, optimization of all aspects of fermentation, characterization of chitinolytic enzyme, and screening of chitinase for the addition and construction of a genetically engineered Mucor strain has to be a research focus.