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Heliyon
Heliyon
Heliyon
2405-8440
Elsevier

S2405-8440(24)12833-2
10.1016/j.heliyon.2024.e36802
e36802
Research Paper
Production of cellulases by Xylaria sp. and Nemania sp. using lignocellulose substrates for bioethanol production from maize cobs
Kamande Stephen M. kamande.stephen@tukenya.ac.ke
ab⁎
Omwenga George I. b
Ngugi Mathew P. b
a Technical University of Kenya, Department of Biochemistry and Biotechnology, P.O. Box 52428-00200, Nairobi, Kenya
b Kenyatta University, Department of Biochemistry, Biotechnology and Microbiology, P.O. Box 43844-00100, Nairobi, Kenya
⁎ Corresponding author. Technical University of Kenya, Department of Biochemistry and Biotechnology, P.O. Box 52428-00200, Nairobi, Kenya. kamande.stephen@tukenya.ac.ke
30 8 2024
15 9 2024
30 8 2024
10 17 e3680211 5 2024
22 8 2024
22 8 2024
© 2024 The Authors
2024
https://creativecommons.org/licenses/by-nc-nd/4.0/ This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).
Two species of Xylaria (KM01, KM03) and Nemania sp.KM02 isolated from decaying plant biomass were evaluated for their ability to produce cellulases on maize cob, eucalyptus, and cypress substrates under solid-state fermentation. A total of 10 fungal samples from decaying plant biomass were collected from Karura forest based on morphological variations. The fungi isolated were screened for cellulase activity and positive isolates were selected for the study. ITS4 R and ITS86 F primers were used to identify the fungal isolates with accuracy ranging from 98 % to 100 %. The crude cellulases produced was assayed for FPase, exoglucanase, endoglucanase and β-glucosidase. Cellulases of Xylaria sp. KM01 produced higher FPase and exoglucanase (2.01 ± 0.13 IU/ml and 0.94 ± 0.08 IU/ml) on pretreated maize cobs with 0.1M HCl at 121oC, while that of Xylaria sp.KM03 produced higher β-glucosidase and endoglucanase (588.6 ± 64.2 IU/ml and 3.59 ± 0.02 IU/ml) on maize cobs pretreated with 0.1M NaOH at 121oC. However, cellulases of Xylaria sp. KM01 produced higher β-glucosidase and FPase (629.7 ± 20.2 IU/ml and 1.67 ± 0.03 IU/ml) on untreated maize cobs after the 9th day of incubation, whereas cellulases of Xylaria sp.KM03 and Nemania sp.KM02 produced higher endoglucanase and exoglucanase (2.80 ± 0.21 IU/ml and 0.83 ± 0.02 IU/ml) on untreated maize cobs after the 3rd and 6th day of incubation. Saccharification of maize cobs by cellulase of Xylaria sp.KM03 produced the highest reducing sugars at 8 % substrate loading (10.17 ± 0.37 mg/ml) after 72 h of incubation. Simultaneous hydrolysis and fermentation of maize cobs by cellulase of Nemania sp.KM02 and Saccharomyces cerevisiae yielded higher bioethanol (28.72 ± 3.82 mg/ml) after 96 h of fermentation. Maize cob is established as a suitable feedstock for cellulases and bioethanol production.

Keywords

Lignocellulose substrates
Cellulases
Saccharification
Bioethanol
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pmc1 Introduction

As the world's population grows, there is an increasing energy demand and limited fossil fuel (oil, coal, and gas) reserves to sustainably meet this demand [1]. The projected depletion of fossil fuels and the increase in global warming have led countries to explore the need for non-conventional and natural energy sources [2,3]. Serious environmental concerns posed by fossil fuels can be overcome by producing bioethanol from agricultural wastes. Pretreatment and enzyme optimization are key aspects of improving saccharification of biomass for bioethanol production. These two methods deal with the challenge by efficiently hydrolyzing cellulose (saccharification) into reducing sugars, [4]. The reducing sugars produced from lignocellulose hydrolysis can be fermented by; i) separate hydrolysis and fermentation, ii) simultaneous saccharification and fermentation (SSF), iii) solid-state fermentation (SSF), iv) simultaneous saccharification and co-fermentation (SSCF), v) consolidation bioprocessing (CBP) or v) batch fermentation [5]. In this study, separate hydrolysis and fermentation were used to produce ethanol from maize cobs.

Lignocellulose primarily consists of 35 %–50 % cellulose, 20 %–35 % hemicellulose, and 5 %–30 % lignin by dry weight and forms a major structural component of plant cell walls [[6], [7], [8]]. Cellulose consists of glucose molecules linked together by β-(1–4) glycosidic bonds with repeating cellobiose units [9,10]. Cellulose molecules (about 30 units) are packaged into elementary fibrils, which are reassembled into microfibrils, and finally assembled into cellulose fibers [11]. Cellulose degradation synergistically is done through the action of three enzymes (endoglucanase, exoglucanase and β-glucosidase) based on their catalytic action [12]. Endo-glucanase cleaves randomly in the internal D-glycosidic linkages preferentially in the amorphous units of the cellulose, easily rendering cellulose more prone for exo-glucanase cleavage to form free ends chain [13]. Exoglucanase (cellobiohydrolase, CBH) hydrolyzes cellulose to produce cellobiose units. CBH I cleave cellulose from the reducing end while CBH II cleaves from the non-reducing end. The presence of cellobiose, a by product of substrate breakdown will however prevent the activity of cellobiohydrolase [14]. β-glucosidase catalyzes the breakdown of cellobiose and oligosaccharides into glucose monosaccharides [15]. However, the production or presence of glucose competitively inhibits β-glucosidase activity [13].

Different microorganisms secret cellulolytic enzymes in nature, which can be purified and used to break down lignocellulose into simple fermentable sugars for biofuel production. Trichoderma sp. and Aspergillus sp. are commonly utilized species to source lignocellulases [16]. Despite numerous reports of isolation of lignocellulolytic enzymes from microorganisms in other parts of the world, the biotechnological benefit of novel lignocellulolytic enzymes from Kenyan microbial communities has not been fully exploited. Plant biomass like agricultural and forestry residues are renewable natural sources of organic matter that can serve as low-cost sources of feedstock that could be utilized to produce enzymes and alternative fuel among other commodity products [6]. It is, therefore, necessary to focus on isolating and characterizing new microorganisms from the local environment as sources of lignocellulolytic enzymes that hydrolyze lignocellulose biomass to reduce the cost and process of producing biofuels, among other products in today's market [[17], [18], [19]]. Cellulase enzyme production can be done using chemically defined media under submerged fermentation or by the use of lignocellulose materials under solid-state fermentation. Solid state fermentation (SSF) is a technique in which the microbial cell growth preferably fungi is performed in a fermenter with a moist substrate in no free-moving water environment [20,21]. This fermentation technology has been widely used for metabolite production from plant biomass such as lignocellulases, gibberellins, and biopesticides among others [22]. Recently, this technology has been adopted in cellulase production as compared to submerged fermentation systems due to high enzyme production [23].

Bioethanol is produced from the sugar component of agro-waste through fermentation under controlled environmental conditions releasing carbon dioxide (CO2) [24]. The fermentation process is anaerobically and follows the Embden-Meyerhoff pathway (EMP) which is catalyzed by bacterial and fungal enzymes [24]. In this process, the substrates are first hydrolyzed by chemical means or with cellulase enzymes to reduce sugars, followed by fermentation by ethanologenic microorganisms to produce bioethanol [25]. Bioethanol production from corn starch, though faces challenges due to the shortage of edible crops, but this can be substituted by using non-edible food crops and agro-waste. This would therefore facilitate proper farming for food security. Large amounts of agricultural waste are generated through human and animal activities and pose an environmental concern, a problem usually aggravated by biomass burning [26].

Several cellulosic materials widely used in SSF are sugarcane bagasse, wheat (bran and straw), rice (bran, straw, and husk) maize bran, soy hull, sawdust, and corncobs among others [16,27]. The choice of the substrate in a solid-state fermentation is dependent on many factors such as the cost of raw materials and substrate availability. In this study, we screened untreated and pretreated maize cob, cypress, and eucalyptus substrates for their suitability as carbon sources for cellulase enzyme production. The carbon source resulting in high cellulase production was used to produce enzymes for the saccharification of maize cobs for ethanol production using separate hydrolysis and fermentation technology.

2 Materials and methods

2.1 Sampling area and technique

The sampling area for the present study was Karura forest reserve, located north of central Nairobi (−1.250916S 36.845914 E) along Kiambu road. Ten (10) fungal samples from decaying plant biomass were collected from different sites in the forest. The distribution of the sample units was carried out randomly by locating the sites with fungal species found growing on decaying plant biomass. The specimens were carefully collected, placed into sample envelopes, and labeled before transporting them to the Microbial Biotechnology Laboratory, Department of Biochemistry, Microbiology and Biotechnology at Kenyatta University.

2.2 Experimental design

To examine the performance of cypress, eucalyptus, and maize cobs substrates for the production of cellulase enzymes utilizing the three isolated fungi, a multifactorial experimental design was used in the study. The tests were done in triplicates for reproducibility. The three independent variables for this study were the three isolated fungi, three substrates, and four pretreatment methods while the dependent variables were cellulase enzyme produced and ethanol.

2.3 Isolation and screening of fungi

The fruiting body and hyphae of the mushrooms were cut into 0.5 mm by 0.5 mm in size and sterilized with 1.5 % sodium hypochlorite for 30 s, rinsed 5–7 times with sterile distilled water before culturing on screening media [28]. Fungal strains were selected on their morphological variations and were cultured on 1 % CMC-Congo red medium according to Legodi et al. [29]. The composition of the medium was 1 % Carboxymethylcellulose powder (CMC) 10(g/l), glucose 1(g/l), agar 15(g/l), and Congo red 0.1(g/l) were placed in 2 L of Erlenmeyer flask and made to a liter with distilled H2O. Cultures were incubated at 25oC for 7 days and monitored daily to visualize the hydrolysis zones [28,30]. Three cultures with large clearing zones were selected for cellulase enzyme production. The cellulase-producing fungi were sub-cultured on potato dextrose agar (PDA). The medium was autoclaved at 121oC, 15 psi for 20 min. After cooling 2.5 ml of gentamycin antibiotic solution (100 mg/l) was added before dispensing into sterile petri dishes.

2.4 Culture maintenance

The selected fungal isolates were maintained on PDA by sub-culturing every 14 days and stored at 4 °C for future studies.

2.5 Molecular identification of fungi

Molecular characterization was based on the PCR amplification of the conserved nucleotide sequence of ribosomal internal transcribed spacer (ITS) using gene coding for the 18S, 5.8S, and 28S rRNA [31].

2.5.1 Fungal genomic DNA extraction

Mycelia of the three fungal strains were used for genomic DNA isolation by the Graham method.

[32]. Fungal mycelia were scraped with the help of a fine sterile scalpel followed by freezing in liquid nitrogen and suspended in 100 μl of extraction buffer solution [100 mM Tris-HCl, pH 8.0, 60 mM EDTA, pH8.0] in a sterile mortar and pestle and ground into fine powder. The powder was placed in 2000 μl microcentrifuge tubes and 400 μl of lysis buffer [400 mM Tris-HCl, pH 8.0, 60 mM EDTA, pH 8.0, 150 mM NaCl, 1 % sodium dodecyl sulfate] was added and the tube contents left at room temperature for 10 min. This was followed by adding 10 μl of Proteinase K [20 mg/ml], mixed gently, and incubated at 65 °C for 15 min in a water bath. An equal volume of chloroform: isoamyl alcohol (24:1 v/v) was added and centrifuged at 13,200 rpm for 5 min at 4 °C. The supernatant was transferred to a new tube and an equal volume of 3M Sodium acetate and isopropyl alcohol were added. The tubes were gently inverted 5 times. The tubes were centrifuged at 13,200 rpm for 10 min, and the supernatant was discarded. The resultant DNA pellet was washed with 300 μl of 70 % ethanol by spinning at 10,000 rpm for 1 min, the supernatant was discarded. The DNA pellet was air-dried and dissolved in 50 μl of 1 x Tris-EDTA, pH 8.0. The quality of DNA was checked on 1 % agarose gel.

2.5.2 PCR amplification

The internal transcribed spacer ITS target region coding gene for 5.8S rRNA was amplified using ITS4 (R) 5′TCCTCC GCT TAT TGA TAT GC 3′for the reverse strand and ITS86 (F)5′ GTG AAT CAT CGA ATC TTT GAA 3′ for the forward strand [31]. Amplification was done in 22 μl reaction mixture (3 μl of fungal DNA template, 0.4 μl Taq polymerase, and 4.0 μl buffer with dNTPs and MgCl2, 4 μl of both primers and 10.6 μl of PCR H2O). The thermocycler was initiated with a denaturation process at 95oC for 3 min, followed by 32 cycles of second denaturation at 95oC for 30 s, annealing at 55oC for 2 min, and elongation at 72oC for 1 min. Final elongation was done at 72oC for 10 min before maintaining at 4oC [33]. The quality of amplicons was checked on 1.2 % agarose gel.

2.5.3 Sequencing of PCR amplicons

The amplified DNA samples were packed in 1.0 ml microcentrifuge tubes and sent to Microgen Inc. in the Netherlands for sequencing using the Sanger deoxy chain termination method [33].

2.5.4 Phylogenetic analysis

Phylogenetic analysis of the ITS region of fungal isolates with sequences of related strains obtained from NCBI BLAST was done with MEGA11 software package, [34]. The phylogenetic tree was created using a matrix of pairwise distances estimated using the Neighbor-Joining method [35]. The robustness of the tree topologies was evaluated by bootstrap analysis based on 1000 re-samplings of the sequence alignment.

2.6 Solid-state fermentation (SSF)

Production of cellulases from three locally available substrates (maize cobs, eucalyptus, and cypress) as carbon sources was done in SSF. These substrates were dried, ground into fine powder by an electric mill, and sieved using a 2 mm sieve. Five (5) g of each substrate powder was weighed and dispensed into 350 ml bottles and 10 ml basal salt solution [1.25 g/l NaNO3, 1.0 g/l KH2PO4, 0.05 g/l CaCl2•6H2O, 0.1 g/l MgSO4•7H2O, 0.1 g/l Nacl] was added to each bottle. The bottles were plugged with cotton wool and autoclaved at 121o C, 15 psi for 15 min.

2.7 Inoculation of mycelium into SSF system

Inoculum was prepared from fungal cultures grown on a PDA medium after 7 days. Spores were suspended in 20 ml of 5 % sucrose solution. For fungi that did not sporulate, mycelium was scrapped from a PDA plate and blended in a sterilized blender for 30 s. The resulting mycelium extract was used as inoculum. Two (2) ml of the spore suspension or mycelia extract was transferred into each bottle in a biosafety cabinet, then thoroughly mixed with substrate and incubated at room temperature.

2.7.1 Effects of substrate pretreatment on cellulase enzyme production

Maize cob, cypress, and eucalyptus substrates were pretreated using 0.1M NaOH (w/v) and 0.1M HCl (v/v) at 60oC for 2 h and at 121oC (15 psi) for 15 min. After pretreatment, the substrates were washed with distilled water until neutral pH. The substrates were then dried at 70oC until constant weight is achieved. The pretreated substrates were inoculated with the three cellulolytic fungi as described in inoculation section above (2.7).

2.7.2 Effects of incubation time on cellulase enzyme production

The SSF process was carried out for 15 days to determine the effect of incubation time on cellulase enzyme production. Samples were taken at 3-day intervals for cellulase enzyme determination. All assays were carried out in triplicates.

2.8 Cellulase extraction and assays

The crude enzyme was extracted by adding 50 ml of 50 mM citrate buffer pH 4.8 to all bottles. Culture bottles were vigorously shaken and left to stand at room temperature for 2 h. The mixture was sieved through muslin cloth and the filtrate collected, was centrifuged for 10 min at 10,000 rpm. The crude enzyme extract was used for cellulolytic enzyme assays, using the DNS method [36].

2.8.1 Total cellulase activity (FPase) (filter paper unit (FPU))

FPase activity was determined by a strip of Whatman's No.1 filter paper (1 cm × 6 cm) approximately 50 mg according to Ghose [37]. The mixture contained 500 μl of 5 mM citrate buffer pH 4.8, filter paper strip, and 500 μl of crude enzyme extract in 50 mM citrate buffer, pH 4.8, accordingly. The mixture was incubated at 50 °C for 1 h in a water bath. The released reducing sugar was estimated by the addition of 3,5-dinitrosalicylic acid (DNS) reagent (10 g of 3,5-dinitrosalicylic acid, 0.5g of sodium sulfite, 10 g of sodium hydroxide, and 2 g of phenol were placed in a 1000 ml volumetric flask and dissolved with distilled water up to 1000 ml mark) with glucose as standard. The mixture was boiled for 5 min followed by adding 300 μl sodium potassium tartrate (400 g of potassium sodium tartrate powder was placed in a 1000 ml volumetric flask and dissolved with distilled water) after cooling. Reducing sugars produced during the reaction were assayed in triplicates using a Jenway 6300 UV/Vis spectrophotometer at 540 nm. One unit of FPU was defined as the amount of enzyme required to release 1 μmol of glucose (reducing sugar equivalents) per minute from filter paper under the assay conditions.

2.8.2 Exoglucanase assay

Exoglucanase activity was determined according to the method described by Ghose [37]. The reaction mixture contained 500 μl of 1.25 % Avicel (microcrystalline) 0.1M Na acetate buffer, pH 5.0, and 500 μl of crude enzyme. The mixture was incubated at 50∘C for 30min and the released reducing sugar was estimated as indicated in the assay for FPase above. One unit of exoglucanase activity was defined as the amount of enzyme releasing 1 μmol of glucose (reducing sugar equivalents) per minute per unit from avicel under the assay conditions.

2.8.3 Endoglucanase assay

Endoglucanase activity was determined according to the method described by Ghose [37]. The reaction mixture contained 500 μl of 1 % CMC in 0.1M Na acetate buffer, pH 5.0, and 500 μl of crude enzyme. The mixture was incubated at 50∘C for 30 min and the released reducing sugar was estimated as indicated in the assay for FPase above. One unit of endoglucanase activity was defined as the amount of enzyme releasing 1 μmol of glucose (reducing sugar equivalents) per minute per unit from CMC under the assay conditions.

2.8.4 β glucosidase assay

β-Glucosidase activity was determined according to the method described by Herr [38]. The reaction mixture contained 100 μl of 5 mM para-nitro phenol-β- D-glucopyranoside (pNPG) in 100 mM sodium acetate buffer, pH 4.8, 100 μl of crude enzyme solution, and 0.4 ml of 100 mM sodium acetate buffer, pH 4.8. The substrate control contained 0.2 ml of 5 mM pNPG prepared in 100 mM citrate buffer at pH 4.8 and 0.4 ml of 100 mM sodium acetate buffer, at pH 4.8. The mixture was incubated at 50 °C for 15 min in a water bath. The reaction was stopped by adding 800 μl ml of 50 mM NaOH-glycine buffer, pH 10.8. The enzyme activity indicated by the release of p-nitrophenol was determined at 420 nm using a Jenway 6300 UV/Vis spectrophotometer. One unit of β glucosidase activity was defined as the amount of enzyme required to release 1 μmol of p-nitrophenol per minute under the same assay conditions.

2.9 Concentration of crude enzymes for saccharification

The crude cellulases of isolated fungal strains from maize cob fermentation of higher enzyme activity were extracted through sieving with muslin cloth and separated using a high-speed centrifuge (10,000×g, 20 °C, 10 min). The crude extract was concentrated five-fold by freeze-drying using a BIOBASE™ tabletop freeze dryer as described by Jasińska et al. [39],.and used for the hydrolysis process.

2.10 Saccharomyces cerevisiae inoculum for ethanol production

Baker's yeast, Saccharomyces cerevisiae was used in the fermentation process for ethanol production. Dried yeast (1 g) was inoculated into a 5 % glucose solution in a sterile 50 ml Erlenmeyer flask and incubated in a shaker (150 rpm) at 25oC for 24 h. A loopful of the yeast colony was transferred on the agar plate and incubated at 25oC for 24 h to check for viability and contamination, [40]. The total viable yeast cells were then determined using a Neubauer chamber with 0.4 % trypan blue stain [41]. The yeast cells loading concentration used in the maize cobs hydrolysate (20 ml) was adjusted to 1.0 × 106 cells/ml.

2.11 Saccharification of maize cobs with cellulases of isolated fungi

Hydrolysis of maize cobs with crude cellulases of isolated fungi was done according to Mandels et al. [42]. Saccharification mixture comprised of untreated maize cobs, (4 %, 8 %, 18 %, & 20 % w/v) in 18 ml of citrate buffer at pH 4.8 and autoclaved at 121oC for 15 min. After cooling, 1 ml of crude enzyme extract (5 FPU), 0.5 ml of gentamycin (200 mg/l), and 0.5 ml of griseofulvin (200 mg/l) to inhibit microbial growth were added to make a total volume of 20 ml in 50 ml falcon tubes. The saccharification mixture was incubated at 25 °C for 72 h with agitation at 150 rpm. Five (5) FPU (Catalog #C1184 Sigma-Aldrich) cellulase and 100 IU/ml β-glucosidase (Catalog # 49290 Sigma-Aldrich) enzymes were used as positive control while citrate buffer at pH 4.8 was used as negative control. Samples (1 ml) were withdrawn aseptically from each tube at 12-h intervals and analyzed for reducing sugar by the DNS method described by Miller [36], to determine the best time for the maximum reducing sugar production.

2.12 Simultaneous saccharification and fermentation

Simultaneous saccharification and fermentation were used for ethanol production in 50 ml falcon tubes. The fermentation mixture comprised untreated maize cobs (4 %, 8 %, 18 %, & 20 % w/v) and 18 ml of citrate buffer at pH 4.8 and autoclaved at 121oC for 15 min. After cooling, 1 ml of crude enzyme extract (5 FPU, Catalog #C1184 Sigma-Aldrich), 0.5 ml of gentamycin (200 mg/l) to inhibit microbial growth, and 1 ml of Saccharomyces cerevisiae (1.0 × 106 cells/ml) were added and incubated at 37 °C for 96 h with agitation at 150 rpm. Five (5) FPU (Catalog #C1184 Sigma-Aldrich) cellulase and 100 IU/ml β-glucosidase (Catalog # 49290 Sigma-Aldrich) enzymes were used as positive control while citrate buffer at pH 4.8 was used as negative control. Samples (0.5 ml) were collected aseptically at 24-h intervals, and the amount of ethanol produced was determined by the acidified potassium dichromate method described by Ref. [43]. Ethanol sample (500 μl) was collected from fermentation mixture in a 2 ml micro centrifuge tube and 500 μl acidified potassium dichromate reagent (20 g of potassium dichromate (VI) dissolved in 1000 ml volumetric flask) was added. Concentrated sulfuric acid (100 ml) was slowly added to the solution at 60oC in a water bath. The mixture was diluted with 200 μl of sterile distilled water. Standard mixture was prepared from stock solution (20 mg/ml) at different concentrations (0.0, 0.2, 0.4, 0.8, 1.2, 1.6, and 2.0). The absorbance was measured at 600 nm using a Jenway 6300 UV/Vis spectrophotometer in triplicates and used in calculating the amount of ethanol (mg/ml) produced.

2.13 Data management and analysis

Enzyme activity absorbance data was converted into international units (IU/ml) using glucose and para-nitrophenol standard curves as described by Rahnama et al. [44]. Enzyme activity was calculated based on the following equations.i) FPase, Exoglucanase and Endoglucanase (IU/ml)

=finalabs−cm×dfsamplevolume×1time×1000μg1mg×1μmole180.16μg

ii) β-glucosidase (IU/ml)

=finalabs−cm×dfsamplevolume×1time×1000μg1mg×1μmole139.1μg

Where c = intercept, m = slope obtained from glucose and para-nitro phenol standard curves, ab is the absorbance, and df is the dilution factor. Time is the reaction time and is expressed in minutes.

Ethanol yield obtained in triplicates was entered into an Excel spreadsheet and converted into mg/ml of ethanol. The data was imported in R software version 3.5.1 [45] and checked for normality using Shapiro-Wilk and equality of variance using Levene's test. Data in triplicates were statistically analyzed using one-way ANOVA at P ≤ 0.05 significant level. Any significant differences in the factors affecting enzyme and ethanol production were determined by Tukey's HSD Post Hoc test. For molecular data analysis sequences were compared with the sequences in the nucleotide database (NCBI) using the BLAST technique. The MEGA software version X program was used for phylogenetic analysis and multiple alignments using CLUSTAL W [46]. Tables and figures were used to present the data.

3 Results

3.1 Isolation of cellulolytic fungi

The cellulolytic activities of the fungi were demonstrated on an agar medium containing 1 % carboxyl methyl cellulose and Congo red (Fig. 1). The ability to grow on a medium containing carboxyl methyl cellulose was taken as a measure of the cellulase activity of the fungi. This method has been used to measure the initial hydrolysis rate for cellulases using soluble cellulose derivatives [47,48], and is strongly recommended along with the determination of reducing sugar. The isolated fungi represent microorganisms obtained from decaying plant biomass from the Karura forest able to utilize and grow on lignocellulose [48]. These reports of their ability to utilize cellulose as a carbon source were confirmed in this project by their ability to exhibit growth on cellulolytic agrowastes such as sugarcane, cypress, eucalyptus, and maize cobs.Fig. 1 Cellulase activities and colony morphologies of Xylaria sp. KM01(A1, A2), Nemania sp. KM02 (B1, B2) and Xylaria sp. KM03 (C1, C2) cultured on 1 % CMC-Congo red agar and sub-cultured on PDA respectively. (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

Fig. 1

3.2 Molecular identification of the cellulolytic fungi

The fungal isolates were identified based on their molecular characteristics using ITS primers.

Fungal genomic DNA from the three fungi was isolated, amplified and amplicons sizes of 650 bp were obtained (Fig. 2). The fungal isolates were identified as; KM01, KM02 and KM03.Fig. 2 An image of an agarose gel indicating the amplification of ITS target region of Xylaria sp. KM01, Nemania sp. KM02 and Xylaria sp. KM03.

Fig. 2

The sequence data of the isolated fungal strains were submitted to the NCBI Gene Bank database for comparison with available sequences using BLAST (Fig. 3).Fig. 3 Phylogenetic tree for Xylaria sp. KM01, Nemania sp. KM02 and Xylaria sp. KM03. Cerrena unicolor isolate CFCC was used as an outgroup.

Fig. 3

The evolutionary history of Xylaria sp. KM01, Xylaria sp. KM03 and Nemania sp. KM02 were inferred using the Neighbor-Joining method [35]. The percentage of replicate trees in which the associated taxa clustered together in the bootstrap test (1000 replicates) are shown next to the branches Felsenstein [49]. The evolutionary distances in the units of the number of base substitutions per site were computed using the Kimura 2-parameter method (Kimura M [50]. There was inclusion of 1st+2nd+3rd + Noncoding codon positions. The bootstrap values are represented by figures at the nodes [34,35,49]. MEGA11 was used for evolutionary analyses (Tamura et al., 2021).

3.3 Effect of substrate pretreatment on cellulase production by Xylaria sp. KM01, Nemania sp. KM02 and Xylaria sp. KM03

The effect of pretreatment on FPase, exoglucanase, endoglucanase and β-glucosidase was investigated for the three fungal strains. The pretreatment was done at 60oC for 1 h and 121oC for 15 min with 0.1M HCl and 0.1M NaOH on cypress, eucalyptus and maize cob substrates under the SSF process. The crude enzyme was extracted on the 5th day of fermentation. A quantitative evaluation of cellulase production by the selected fungi was assayed using filter paper, avicel, CMC and pNPG as commercial substrates to assess the effect of pretreatment on FPase, exoglucanase, endoglucanase and β-glucosidase for the three fungal strains.

3.3.1 Effect of substrates pretreatment on total cellulase activity (FPase)

Maximum FPase was observed after 5 days for all the fungal strains (Fig. 4). On maize cob, Xylaria sp. KM01 and Nemania sp. KM02 produced 2.01 ± 0.131 IU/ml and 1.69 ± 0.085 IU/ml when pretreated with 0.1MHCl at 121oC. Xylaria sp. KM03 produced 1.98 ± 0.033 IU/ml when pretreated with 0.1M NaOH at 121oC (P < 0.05). Xylaria sp. KM01 produced 6.84 ± 0.040 IU/ml with untreated cypress while Xylaria sp. KM03 produced 0.6 ± 0.050 IU/ml followed by Nemania sp. KM02 with 0.53 ± 0.096 IU/ml when cypress pretreated with 0.1M NaOH at 121oC.on eucalyptus, Nemania sp. KM02 produced 0.91 ± 0.065 IU/ml when pretreated with 0.1M NaOH at 60oC. Xylaria sp. KM03 produced 0.83 ± 0.036 IU/ml followed by Xylaria sp. KM01 with 0.6 ± 0.050 IU/ml when pretreated with 0.1M NaOH at 121oC.Fig. 4 FPase activities of (A) Xylaria sp. KM01, (B) Nemania sp. KM02, and (C) Xylaria sp. KM03 cultured on pretreated cypress, eucalyptus and maize cob substrates. Values are means of three replicates ± SEM.

Fig. 4

3.3.2 Effect of substrate pretreatment on exoglucanase activity

The highest exoglucanase activity was recorded on maize cob for all the fungal strains (Fig. 5). Xylaria sp. KM01 and KM03 produced activities of 0.94 ± 0.077 IU/ml and 0.93 ± 0.044 IU/ml respectively, whereas Nemania sp. KM02 recorded 0.85 ± 0.096 IU/ml when pretreated with 0.1M HCl at 121oC. This was followed by eucalyptus with the highest recorded by Nemania sp. KM02 with 0.54 ± 0.6IU/ml when pretreated with 0.1M NaOH at 60oC, Xylaria sp. KM03 produced an amount of 0.38 ± 0.063 IU/ml when pretreated with 0.1M NaOH at 121oC and Xylaria sp. KM01 0.23 ± 0.015IU/ml when pretreated with 0.1M HCl at 60oC. The lowest activity noted on cypress of 0.28 ± 0.018 IU/ml produced by Nemania sp. KM02 when pretreated with 0.1M NaOH at 121oC, Xylaria sp. KM03 produced an amount of 0.23 ± 0.018 IU/ml when pretreated with 0.1M NaOH at 121oC, while Xylaria sp. KM01 produced 0.14 ± 0.042 IU/ml when pretreated with 0.1M NaOH at 121oC.Fig. 5 Exoglucanase activities of (A) Xylaria sp. KM01, (B) Nemania sp. KM02, and (C)Xylaria sp. KM03 cultured on pretreated cypress, eucalyptus and maize cob substrates. Values are means of three replicates ± SEM.

Fig. 5

3.3.3 Effect of substrate pretreatment on endoglucanase activity

The production of endoglucanase activity was higher on maize cob for all the fungal strains (Fig. 6). Xylaria sp. KM01 and Nemania sp. KM02 produced activities of 3.58 ± 0.12 IU/ml and 3.26 ± 0.34 IU/ml respectively, whereas Xylaria sp. KM03 recorded 2.80 ± 0.19 IU/ml when pretreated with 0.1M HCl at 121oC. On cypress, Xylaria sp. KM03 recorded 2.22 ± 0.12 IU/ml when pretreated with 0.1M HCl at 121oC, whereas Nemania sp. KM02 produced 1.85 ± 0.28 IU/ml with untreated cypress. The lowest activity of 1.31 ± 0.17 IU/ml was noted for Xylaria sp. KM03 when pretreated with 0.1M HCl at 121oC.Fig. 6 Endoglucanase activities of (A) Xylaria sp. KM01, (B) Nemania sp. KM02, and (C)Xylaria sp. KM03 cultured on pretreated cypress, eucalyptus and maize cob substrates. Values are means of three replicates ± SEM.

Fig. 6

3.3.4 Effect of substrate pretreatment on β-glucosidase activity

The levels of β-glucosidase produced vary among the fungal strains (Fig. 7). On maize cob, Xylaria sp. KM03, Nemania sp. KM02 and Xylaria sp. KM01 produced significantly higher activities of 588.6 ± 64.2 IU/ml when pretreated with 0.1M NaOH at 121oC, 418.7 ± 44.9 IU/ml when pretreated with 0.1M HCL at 60oC, 415.7 ± 29.4 IU/ml when pretreated with 0.1M HCL at 121oC respectively, compared to the other substrates tested (P ≤ 0.05). On eucalyptus, Xylaria sp. KM03, Nemania sp. KM02 and Xylaria sp. KM01 produced activities of 298.5 ± 57.9 IU/ml when pretreated with 0.1M NaOH at 121oC, 295.2 ± 30.0 IU/ml when untreated and 267.9 ± 15.9 IU/ml when pretreated with 0.1M NaOH at 60oC. On cypress, Nemania sp. KM02, Xylaria sp. KM01 and KM03 produced activities of 244.4 ± 32.3 IU/ml when pretreated with 0.1M HCL at 121oC, 236.0 ± 10.0 IU/ml when pretreated with 0.1M NaOH at 121oC, 190.9 ± 18.6 IU/ml when pretreated with 0.1M NaOH at 60oC.Fig. 7 β-Glucosidase activities of (A) Xylaria sp. KM01, (B) Nemania sp. KM02, and (C) Xylaria sp. KM03 cultured on pretreated cypress, eucalyptus and maize cob substrates. Values are means of three replicates ± SEM.

Fig. 7

3.4 Effect of incubation time on cellulase production by Xylaria sp.KM01, Nemania sp. KM02 and Xylaria sp. KM03

The production of cellulase with changes in incubation time was also investigated in this study (Fig. 8, Fig. 9, Fig. 10, Fig. 11). The fermentation was carried out at room temperature using cypress, eucalyptus and maize cob substrates under the SSF process. The crude enzyme was collected at 3-day intervals during fermentation. Quantitative estimation of cellulase produced by the selected fungi was assayed using filter paper, avicel, CMC and pNPG as commercial substrates to assess the effect of incubation time on FPase, exoglucanase, endoglucanase and β-glucosidase for the three fungal strains.Fig. 8 FPase activities of (A) Xylaria sp. KM01, (B) Nemania sp. KM02 and (C) Xylaria sp. KM03 cultured on untreated cypress, eucalyptus and maize cob substrates. Values are means of three replicates ± SEM.

Fig. 8

Fig. 9 Exoglucanase activities of (A) Xylaria sp. KM01, (B) Nemania sp. KM02 and (C) Xylaria sp. KM03 cultured on untreated cypress, eucalyptus and maize cob substrates. Values are means of three replicates ± SEM.

Fig. 9

Fig. 10 Endoglucanase activities of (A) Xylaria sp. KM01, (B) Nemania sp. KM02 and (C) Xylaria sp. KM03 cultured on untreated cypress, eucalyptus and maize cob substrates. Values are means of three replicates ± SEM.

Fig. 10

Fig. 11 β-Glucosidase Exoglucanase activities of (A) Xylaria sp. KM01, (B) Nemania sp. KM02 and (C) Xylaria sp. KM03 cultured on untreated cypress, eucalyptus and maize cob substrates. Values are means of three replicates ± SEM.

Fig. 11

3.4.1 Effect of incubation time on total cellulase activity (FPase)

Maximum FPase activities of 1.67 ± 0.03 IU/ml by Xylaria sp. KM01, 1.54 ± 0.1 IU/ml by Nemania sp. KM02 and 1.52 ± 0.11 IU/ml by Xylaria sp. KM03 were obtained when cultured on maize cob on the 9th day of incubation (P ≤ 0.05), compared to cypress with production amounts of 1.14 ± 0.12 IU/ml by Nemania sp. KM02, 1.18 ± 0.11 IU/ml by Xylaria sp. KM03 and 1.07 ± 0.09 IU/ml by Xylaria sp.KM01 and eucalyptus with production of 0.95 ± 0.04 IU/ml by Xylaria sp. KM01, 0.90 ± 0.06 IU/ml by Xylaria sp. KM03, 0.73 ± 0.14 IU/ml by Nemania sp. KM02 on the 3rd day of incubation respectively (Fig. 8).

3.4.2 Effect of incubation time on exoglucanase activity

The production of exoglucanase was higher on maize cob for all the fungal strains (Fig. 9). Xylaria sp. KM03 and KM01 produced higher activities of 0.83 ± 0.02 IU/ml and 0.62 ± 0.02 IU/ml on the 6th day of incubation respectively, whereas Nemania sp. KM02 produced activity of 0.72 ± 0.09 IU/ml on the 3rd day of incubation. On cypress, Xylaria sp. KM03 and Nemania sp. KM02 recorded 0.52 ± 0.073 IU/ml and 0.44 ± 0.05 IU/ml on the 6th day of incubation whereas Xylaria sp. KM01 recorded 0.34 ± 0.009 IU/ml on the 3rd day of incubation respectively. On eucalyptus, Xylaria sp. KM03 and KM03 recorded 0.41 ± 0.05 IU/ml and 0.18 ± 0.04 IU/ml on the 3rd day of incubation whereas Nemania sp. KM02 recorded 0.18 ± 0.00 IU/ml on the 9th day of incubation. Xylaria sp. KM03 recorded a steady decrease in activity after 3rd day of incubation on cypress and eucalyptus substrates.

3.4.3 Effect of incubation time on endoglucanase activity

The highest production of endoglucanase was also favored by the time of incubation for the three fungal strains (Fig. 10). On maize cob, Nemania sp. KM02 produced 2.80 ± 0.212 IU/ml on the 3rd day of incubation followed by Xylaria sp. KM01 with amounts of 2.53 ± 0.052 IU/ml on the 9th day of incubation and Xylaria sp. KM03 produced 2.51 ± 0.01 IU/ml on the 9th day of incubation respectively. On cypress, Xylaria sp. KM01 and KM03 produced 2.13 ± 0.12 IU/ml and 1.94 ± 0.28 IU/ml on the 3rd day of incubation respectively, whereas Nemania sp. KM02 produced 2.1 ± 0.16 IU/ml on the 9th day of incubation. On eucalyptus, Xylaria sp. KM01 and KM03 produced 2.4 ± 0.34 IU/ml and 1.54 ± 0.24 IU/ml, while Nemania sp. KM02 produced 0.7.28 ± 0.158 IU/ml on the 3rd day of incubation respectively. Both Xylaria sp. KM01 and KM03 recorded a steady decrease in activity after 3rd day of incubation on cypress and eucalyptus substrates.

3.4.4 Effect of incubation time on β-glucosidase activity

The levels of β-glucosidase produced vary among the fungal strains (Fig. 11). Generally, β-glucosidase activity was the highest on maize cob (P ≤ 0.05). Xylaria sp. KM01 produced higher activity of 629.7 ± 20.2 IU/ml on the 9th day of incubation followed by Xylaria sp. KM03 and Nemania sp. KM02 with amounts of 587.2 ± 29.7 IU/ml and 455.9 ± 28.9 IU/ml on the 12th day of incubation respectively. On cypress, Xylaria sp. KM01 and Nemania sp. KM02 produced 346.3 ± 53.2 IU/ml and 222.5 ± 19.6 IU/ml on the 9th day of incubation respectively. Xylaria sp. KM01 produced 243.1 ± 43.3 IU/ml on the 12th day of incubation. On eucalyptus, Xylaria sp. KM03 produced 396.0 ± 19.1 IU/ml on the 12th day of incubation, Nemania sp. KM02 produced 284.5 ± 17.4 IU/ml on the 6th day of incubation and Xylaria sp. KM01 produced 248.7 ± 17.3 IU/ml on the 9th day of incubation respectively.

3.5 Saccharification of maize cobs with cellulases of isolated fungi

The results show the amount of sugars during the fermentation period (Table 1). Maize cob hydrolysis by crude cellulases of isolated fungi produced higher reducing sugars. At 8 % concentration, cellulases of Xylaria sp. KM03 produced the highest amounts of sugar at 10.17 ± 0.37 mg/ml, Xylaria sp. KM01 produced yields of 9.48 ± 0.13 mg/ml, whereas Nemania sp. KM02 produced 9.41 ± 0.27 mg/ml after 72 h of fermentation period. It was observed that the sugar yield increased steadily reaching the peak after 72 h of fermentation at all the concentrations.Table 1 Sugar yield (mg/ml) from hydrolysis of untreated maize cobs of different concentration by cellulases of isolated fungi.

Table 1	Total reducing sugar (mg/ml)	
(Maize cob concentration %)	
4	8	18	20	Control (+)	Control (−)	
Fungi	Time (Hrs)	Mean ± SD	Mean ± SD	Mean ± SD	Mean ± SD	Mean ± SD	Mean ± SD	
Xylaria sp.
KM01	12	2.35 ± 0.11bcC	4.12 ± 1.21aB	3.58 ± 0.40abC	4.03 ± 0.48aD	4.53 ± 0.035aD	1.59 ± 0.05cD	
24	3.17 ± 0.73cC	8.21 ± 0.36aA	6.84 ± 0.13bB	6.42 ± 1.37bC	7.32 ± 0.45abC	1.85 ± 0.04dC	
36	7.54 ± 0.36bcB	8.78 ± 0.13aA	7.19 ± 0.2bcB	6.76 ± 0.03cBC	7.79 ± 0.55bBC	1.88 ± 0.00dBC	
48	8.37±0aAB	8.89 ± 0.00aA	7.55 ± 0.08abB	6.89 ± 0.06bBC	8.25 ± 0.86aBC	1.90 ± 0.00cBC	
60	8.51 ± 0.16abA	9.17 ± 0.04aA	8.37 ± 0.53bA	7.14 ± 0.23cAB	8.84 ± 0.38abB	1.95 ± 0.00dB	
72	8.83 ± 0.07cA	9.48 ± 0.13bA	8.84 ± 0.08cA	7.73 ± 0.28dA	11.21 ± 0.29aA	2.09 ± 0.00eA	
Nemania sp.
KM02	12	2.99 ± 0.11cD	5.51 ± 0.30aD	4.63 ± 0.45bC	4.40 ± 0.05bD	2.10 ± 0.07dD	1.63 ± 0.00dD	
24	3.50 ± 0.38bcD	6.40 ± 0.21aC	5.82 ± 0.03aBC	4.63 ± 0.01bCD	3.40 ± 0.91cD	1.90 ± 0.00dC	
36	5.12 ± 0.73bcC	7.10 ± 0.37aC	6.00 ± 0.14bBC	4.76 ± 0.06cC	5.84 ± 0.23bC	1.90 ± 0.00dC	
48	6.41 ± 0.43bB	8.52 ± 0.12aB	6.20 ± 0.00bcBC	5.19 ± 0.04cB	6.70 ± 0.37bC	1.90 ± 0.00dC	
60	7.73 ± 0.62abA	9.13 ± 0.44aAB	6.88 ± 0.70bcB	5.50 ± 0.20cB	9.18 ± 1.40aB	2.14 ± 0.00dB	
72	8.73 ± 0.12bA	9.41 ± 0.27bA	8.67 ± 0.29bA	6.07 ± 0.21cA	11.69 ± 0.56aA	2.23 ± 0.00dA	
Xylaria sp.
KM03	12	3.39 ± 0.35dE	7.15 ± 0.15aE	5.25 ± 0.27bD	4.31 ± 0.02cC	2.53 ± 0.62dE	1.63 ± 0.00eE	
24	4.27 ± 0.47cDE	7.51 ± 0.13aDE	6.13 ± 0.20bC	4.41 ± 0.08cC	4.76 ± 0.12cD	1.88 ± 0.00dD	
36	5.39 ± 0.57cCD	7.83 ± 0.15aD	6.55 ± 0.09bcBC	5.66 ± 0.91cB	7.38 ± 0.12abC	1.90 ± 0.00dC	
48	6.44 ± 0.32cBC	8.56 ± 0.28bC	6.67 ± 0.01cBC	6.45 ± 0.12cAB	9.75 ± 0.14aB	1.90 ± 0.00dC	
60	6.92 ± 0.28cAB	9.19 ± 0.12bB	6.94 ± 0.19cB	6.57 ± 0.00cAB	10.21 ± 0.40aB	1.93 ± 0.00dB	
72	8.13 ± 0.64cA	10.17 ± 0.37bA	9.07 ± 0.50cA	6.82 ± 0.17dA	12.81 ± 0.24aA	2.14 ± 0.00eA	
Table 1 shows amount of Sugar yield (mg/ml) from hydrolysis of untreated maize cobs of different concentration with partially purified cellulases of fungal isolates. Values are means of three replicates ± SEM and are expressed as mg/ml. Means expressed with different superscript capital letters in the same column are significantly different. Means expressed with different superscript small letters in the same row are significantly different at P < 0.05.

3.6 Ethanol production from maize cob using simultaneous hydrolysis and fermentation

Table 2 shows the amount of ethanol yield (mg/ml) produced from untreated maize cobs hydrolysate of different concentrations fermented with saccharomyces cerevisiae during the fermentation period (Table 2). Hydrolysis of maize cobs by cellulases of Xylaria sp. KM01, Nemania sp. KM02 and Xylaria sp. KM03 recorded higher amounts of ethanol after 96 h of fermentation. At 8 % concentration, cellulases of Xylaria sp. KM03 produced the highest amounts of ethanol at 28.59 ± 1.58 mg/ml, Nemania sp. KM01 produced yields of 28.72 ± 3.82 mg/ml, whereas Xylaria sp. KM02 produced 25.75 ± 0.21 mg/ml after 96 h of fermentation period. It was observed that there was a steady increase in ethanol yield with the maximum produced after 96 h of fermentation at all the concentrations. Table 3, adapted from Ref. [51] shows a comparison with a similar trend where at all concentrations of substrates, the ethanol yield increased steadily and recorded maximum yield after 72 h of fermentation. This shows that corn cobs can be one of the cheaper substrates for ethanol production.Table 2 Ethanol yield (mg/ml) produced from untreated maize cob hydrolysate fermented with saccharomyces cerevisiae.

Table 2	Ethanol yield (mg/ml)	
(Maize cob concentration %)	
4	8	18	20	Control (+)	Control (−)	
Fungi	Time(Hrs)	Mean ± SD	Mean ± SD	Mean ± SD	Mean ± SD	Mean ± SD	Mean ± SD	
Xylaria sp.
KM01	12	13.22 ± 1.66bF	13.35 ± 1.15bF	14.62 ± 0.46bD	9.03 ± 2.14cF	18.62 ± 0.46aE	8.25 ± 0.00cG	
24	17.20 ± 1.31bE	16.32 ± 1.74bE	15.93 ± 0.50bCD	14.33 ± 0.69bE	24.82 ± 3.66aD	8.59 ± 0.04cF	
36	18.71 ± 0.12bDE	18.75 ± 0.61bDE	16.76 ± 0.07bcC	15.89 ± 0.55cDE	29.65 ± 0.62aC	8.59 ± 0.00dF	
48	20.52 ± 0.50bCD	21.00 ± 0.42bCD	17.04 ± 0.09cC	17.07 ± 1.03cCD	30.90 ± 0.64aC	8.93 ± 0.00dE	
60	21.42 ± 0.17bBC	22.11 ± 0.17bC	19.17 ± 0.79cB	19.17 ± 0.69cBC	33.31 ± 1.36aC	9.61 ± 0.00dD	
72	22.05 ± 0.33bBC	23.15 ± 0.69bBC	20.23 ± 0.39cB	20.23 ± 0.17cAB	37.67 ± 0.55aB	9.73 ± 0.00dC	
84	23.20 ± 0.74bAB	24.29 ± 0.66bAB	21.06 ± 0.46cB	21.12 ± 0.48cAB	38.66 ± 0.54aB	9.90 ± 0.00dB	
96	24.61 ± 0.72bA	25.75 ± 0.21bA	24.40 ± 1.47bA	22.54 ± 0.41bA	43.09 ± 1.12aA	10.24 ± 0.00cA	
Nemania sp.
KM02	12	13.06 ± 1.43cG	16.32 ± 1.15bE	14.47 ± 0.23bcF	10.39 ± 0.07dF	23.81 ± 1.18aE	7.51 ± 0.00eE	
24	14.96 ± 0.51cF	17.88 ± 1.74bDE	16.82 ± 0.14bEF	11.13 ± 0.69dF	25.65 ± 0.59aE	7.51 ± 0.04eE	
36	16.40 ± 0.31cEF	18.62 ± 0.26bCDE	17.65 ± 0.23bDE	12.10 ± 0.58dEF	28.57 ± 0.72aD	7.79 ± 0.00eD	
48	17.39 ± 0.26dDE	20.38 ± 0.82bBCDE	18.62 ± 0.14cCDE	13.37 ± 0.21eDE	29.54 ± 0.53aD	8.25 ± 0.00fC	
60	18.47 ± 0.63bCD	21.75 ± 0.40bBCD	19.26 ± 0.39cCD	14.49 ± 0.32dCD	30.75 ± 0.17aCD	8.53 ± 0.00eB	
72	19.77 ± 0.16cBC	22.69 ± 0.46bBC	20.67 ± 0.76cBC	15.74 ± 0.89cBC	32.32 ± 0.96aC	8.70 ± 0.00dA	
84	20.40 ± 0.43cAB	24.10 ± 0.43bB	22.90 ± 1.31bcB	17.44 ± 0.71dB	35.27 ± 1.86aB	8.70 ± 0.00eA	
96	21.65 ± 1.79cA	28.72 ± 3.82bA	28.66 ± 1.79bA	20.02 ± 1.14cA	43.08 ± 0.41aA	8.70 ± 0.00dA	
Xylaria sp.
KM03	12	11.09 ± 0.55bE	15.24 ± 0.69aF	7.85 ± 1.08cF	9.86 ± 1.11bcE	17.04 ± 1.20aD	5.23 ± 0.00dG	
24	13.06 ± 0.16cD	18.37 ± 0.16bE	11.00 ± 2.09cdE	11.83 ± 0.64cdDE	23.23 ± 1.82aC	8.59 ± 0.00dF	
36	13.60 ± 0.16cD	20.06 ± 0.17bDE	13.80 ± 0.32cDE	13,73 ± 0.33cCDE	26.13 ± 0.60aBC	8.70 ± 0.00dE	
48	14.28 ± 0.12dCD	20.91 ± 0.48bCD	15.93 ± 1.10cCD	14.60 ± 0.43cdCD	27.07 ± 0.20aBC	8.99 ± 0.00eD	
60	15.45 ± 0.52dC	22.13 ± 0.41bBCD	18.34 ± 1.24cBC	15.24 ± 0.10dCD	28.21 ± 0.68aBC	9.73 ± 0.00eC	
72	16.72 ± 0.64dB	22.79 ± 0.17bBC	20.04 ± 0.09cAB	16.46 ± 0.42dBC	31.95 ± 1.06aAB	9.73 ± 0.00eC	
84	18.17 ± 0.27cA	24.04 ± 0.71bB	20.69 ± 0.39cAB	20.82 ± 1.53bcB	35.72 ± 2.38aA	10.18 ± 0.00dB	
96	19.19 ± 0.66dA	28.59 ± 1.58bA	22.49 ± 1.15cdA	27.02 ± 4.04bcA	38.54 ± 1.26aA	10.30 ± 0.00eA	
Table 2 shows the amount of ethanol yield (mg/ml) produced from untreated maize cob hydrolysate of different concentration fermented with saccharomyces cerevisiae. Values are means of three replicates ± SEM and are expressed as mg/ml. Means expressed with different superscript capital letters in the same column are significantly different at P < 0.05. Means expressed with different superscript small letters in the same row are significantly different at P < 0.05.

Table 3 Ethanol yield (%) produced from glucose (synthetic medium) and corn cob (complex medium) hydrolysate by cellulase of A niger fermented with saccharomyces cerevisiae.

Table 3Fermentation time		Ethanol yield (%)			
(Hrs)	(Glucose)	(Corn cobs)	
	1 %	10 %	1 %	10 %	
24	1.76	3.47	2.21	2.77	
48	2.38	5.21	3.44	4.80	
72	3.45	6.23	4.17	6.17	
96	2.85	6.01	3.89	5.11	
120	2.67	5.62	3.57	4.89	
(adapted from Ref. [51]).

4 Discussion

Fungal species are mostly used in cellulase production because of their capacity to provide large amounts of extracellular cellulases using cheap lignocellulose substrates. Cellulase enzyme production occurs through submerged fermentation or solid-state fermentation. The cost of cellulase enzymes prevents the economical application of ethanol production from lignocellulose. The cost could be reduced by culturing cellulase-producing fungi isolated from the local environment on cheap substrates. Our research was aimed at isolating and characterizing local fungi from the Karura forest for the production of cellulase enzymes using locally available substrates such as maize cobs, cypress and eucalyptus and subsequently used for saccharification of maize cobs for ethanol production using Saccharomyces cerevisiae. Considering the advantages of the solid-state fermentation process, we investigated several parameters that influence cellulase production, such as (i) time of incubation, (ii) pretreatment and (iii) substrate loading on maize cobs saccharification for ethanol production.

4.1 Isolation and screening of fungi

Ten (10) fungal samples from decaying plant biomass were collected from different sites in the forest. Growth on 1 % CMC Congo red agar was used for the isolation of cellulolytic fungal strains in this study. Three fungal strains were selected based on fast and abundant growth on CMC agar plates. This method provided a relatively easy tool for the specific detection of fungi that produce cellulase activity. A similar result was obtained by Ref. [29,48] where the isolated fungi were able to grow and secrete endoglucanase which hydrolyzed CMC bound to CR dye. This was revealed by appearance of pale yellow “halo zone” around the fungal growth, an indication of CMC hydrolysis.

4.2 Molecular identification of cellulase producing fungi

Molecular identification was based on the detection of conserved sequences in the 5.8S rRNA region and the amplification of the ITS2 region [31]. ITS4 R and ITS86 F primers were used to identify these fungal isolates by comparing sequences to those in NCBI databases using BLAST with accuracy ranging from 98 % to 100 %. The three fungal isolates were identified as Xylaria sp. KM01 (Genebank accession number: ON679521), Nemania sp. KM02 (Genebank accession number: ON678279), and Xylaria sp. KM03 (Genebank accession number: ON704639), (ncbi.nlm.nih.gov/blast)

4.3 Effect of pretreatment on cellulase production

Production of cellulase enzymes from complex lignocellulose is not significantly achieved by disrupting the recalcitrant structure through ordinary microbial attack. Therefore, the selection of a simple and effective method for treating lignocellulose materials should be considered. Several types of treatments were investigated for cellulose hydrolysis including acid and alkali pretreatments [[52], [53], [54], [55], [56]]. The proper mechanism is to attack hemicellulose for dissolution to initiate the loosening of the complex structure of lignocellulose.

A higher yield of FPase activity was produced by the three fungi on pretreated maize cob, Xylaria sp. KM01 produced the highest enzyme activity (2.11 ± 0.13 IU/ml) when pretreated with HCl at 121 °C. Xylaria sp. KM03 (1.98 ± 0.033 IU/ml) when pretreated with NaOH at 121oC Nemania sp. KM02 (16.9 ± 0.85 IU/ml) when pretreated with HCl at 121oC. Kakde et al. [57] reported similar findings when corn cob was pretreated with 5 % (w/v) HCl autoclaved at 121 °C for 60 min recorded maximum cellulase production with 3.46 FPU/ml. This was contrary to Xylaria sp. KM03 which recorded the maximum activity when pretreated with 0.1M NaOH at 121oC. A similar finding was reported by Ojumu et al. [58] where cellulase activity by Penicillium decumbens was recorded significantly higher on corn cob pre-treated with dilute alkali (NaOH).

Higher production of exoglucanase activities were produced by the three fungi on pretreated maize cob when pretreated with HCl at 121oC. Xylaria sp. KM01 produced the highest activity (0.94 ± 0.077 IU/ml), Xylaria sp. KM03 (9.28 ± 0.44 IU/ml) and Nemania sp. KM02 (0.85 ± 0.1 IU/ml), respectively [57]. reported similar findings when corn cob was pretreated with 5 % (w/v) HCl autoclaved at 121 °C for 60 min recorded maximum cellulase production with 3.46 FPU/ml.

Higher amounts of endoglucanase activity were produced by the three fungi on pretreated maize cob, Xylaria sp. KM03 produced the highest enzyme activity (3.59 ± 0.022 IU/ml) when pretreated with NaOH at 121oC. Xylaria sp. KM01 (3.58 ± 0.19 IU/ml) and Nemania sp. KM02 (3.26 ± 0.34 IU/ml) when pretreated with HCl at 121oC respectively. Kakde et al. [57] reported similar findings when corn cob was pretreated with 5 % (w/v) HCl autoclaved at 121 °C for 60 min recorded maximum cellulase production with 3.46 FPU/ml This was contrary to Xylaria sp. KM03 which recorded the maximum activity when pretreated with 0.1M NaOH at 121oC. A similar finding was reported by Ojumu et al. [58] where cellulase activity by Penicillium decumbens was recorded significantly higher on corn cob pre-treated with dilute alkali (NaOH).

Higher levels of β-glucosidase activity were produced by the three fungi on pretreated maize cobs, Xylaria sp. KM03 produced the highest enzyme activity (588.6 ± 64.2 IU/ml) when pretreated with NaOH at 121oC. Nemania sp. KM02 (418.7 ± 44.9 IU/ml) when pretreated with HCl at 60oC, and Xylaria sp. KM01 (415.7 ± 29.4 IU/ml) when pretreated with HCl at 121oC. Kakde et al. [57] reported similar findings when corn cob was pretreated with 5 % (w/v) HCl autoclaved at 121 °C for 60 min recorded maximum cellulase production with 3.46 FPU/ml. This was contrary to Xylaria sp. KM03 which recorded the maximum activity when pretreated with 0.1M NaOH at 121oC for 15 min. A similar finding was reported by Ojumu et al. [58] where cellulase activity by Penicillium decumbens was recorded significantly higher on corn cob pre-treated with dilute alkali (NaOH). In this study acid pretreatment was the most effective method for enzyme production as compared to alkali pretreatment. The possible mechanism involves the dissolution of hemicellulose causing loosening of the structure of raw material, which makes acid pretreatment an important method for the production of enzymes ([53,59]; Pandey et al. [60].

4.4 Effect of time of incubation on cellulase production

The optimal time of incubation for cellulase enzyme production depends on the type of substrate used and the fungal species involved [61]. The time of incubation significantly influenced cellulase enzyme activities from the 3rd day of incubation. The highest FPase activity for all three fungal strains were recorded on maize cob. Xylaria sp. KM01 produced the highest activity (1.67 ± 0.034 IU/ml), Nemania sp. KM02 (1.54 ± 0.01 IU/ml) and Xylaria sp KM03 (1.52 ± 0.11 IU/ml) on the 9th day of incubation respectively. These results were similar to Gautam et al., 2011 findings on Aspergillus niger which showed the highest cellulase activities were recorded after 5 days of incubation period cultured on a corn cob. However, contrary when they recorded a trend in which cellulase activities by Trichoderma viride cultured on corn cob peaked on the 3rd day of incubation and decreased afterward.

Exoglucanase activity produced by the three studied fungi recorded higher yields on maize cob. Xylaria sp. KM03 produced the highest activity (0.83 ± 0.023 IU/ml) on the 6th day of incubation. Nemania sp. KM02 (0.716 ± 0.089 IU/ml) on the 3rd day of incubation. Xylaria sp. KM01 (0.647 ± 0.028 IU/ml) on the 9th day of incubation. These results were similar to Ref. [62] findings on Aspergillus niger which showed the highest exoglucanase activity was recorded after 5 days of incubation and a similar trend in which cellulase activities by Trichoderma viride peaked on the 3rd day of incubation when cultured on a corn cob. Exoglucanase activity by Xylaria sp. KM03 recorded a steady decrease after 3rd day of incubation on cypress and eucalyptus substrates. This could be attributed to fungal species (genetic make-up) and substrates used for growth [63].

Endoglucanase produced by the three fungal strains recorded higher activity on maize cob. Nemania sp. KM02 (2.80 ± 0.212 IU/ml) on the 3rd day of incubation. Xylaria sp. KM01 (2.53 ± 0.052 IU/ml) on the 9th day of incubation. Xylaria sp. KM03 (2.51 ± 0.01 IU/ml) on the 6th day of incubation. These results were similar to Ref. [62] findings on Aspergillus niger which showed the highest cellulase activities were recorded after 5 days of incubation period cultured on a corn cob. However, endoglucanase activities by Xylaria sp. KM01 and KM03 recorded a steady decrease after 3rd day of incubation on cypress and eucalyptus substrates. This could be attributed to fungal species (genetic make-up) and substrates used for growth [63]. High viscosity of the substrate, decreases the oxygen supply to the organisms. It could also be inhibitory effects of accumulated cellobiose generated by the action of cellulases [14].

β-glucosidase activity produced by the three studied fungi was the highest on maize cobs. Xylaria sp. KM01 (629.7 ± 20.2 IU/ml) on the 9th day of incubation. Xylaria sp. KM03 (597.2 ± 29.7 IU/ml) on the 12th day of incubation and Nemania sp. KM02 (455.9 ± 28.7 IU/ml) on the 12th day of incubation [62]. recorded similar findings in which cellulase activities by Trichoderma viride cultured on corn cob peaked after the 5th day of incubation and decreased afterward. These differences could be due to the difference in the type and composition of substrates [64]. It could be also attributed to the maximum metabolic activity of the fungus attributed to the genetic make-up of the fungal strains as a result of adaptations to different habitats, and the concentration of soluble sugar in the substrate [65]. The low cellulase activities could be a result of an inhibitory effect of accumulated cellobiose [14].

4.5 Saccharification of maize cobs for sugar production

In the presence of a given amount of enzyme, the rate of an enzymatic reaction increases as the

substrate loading concentration increases until a limiting rate is reached, after which a further increase in the substrate concentration produces no significant change in the reaction rate. A higher yield of reducing sugars was optimized on substrate loading with 8 % at 72 h of saccharification by three fungi with the highest produced by Xylaria sp. KM03 (10.17 ± 0.37 mg/ml), Xylaria sp. KM01 (9.48 ± 0.13 mg/ml) and Nemania sp. KM02 (9.41 ± 0.27 mg/ml) respectively. A varying yield of sugars from corn cob has also been reported in other studies. Saliu and Sani [66]. recorded a yield of 7.63 mg/ml of reducing sugar produced on hydrolysis of corn cob with cellulases of A. niger within 48 h. Jaafaru and Fagade [67] recorded a similar trend when corn cob was pretreated with alkali (5.55 mg/ml) of reducing sugar on hydrolysis with culture filtrates of A. niger compared to 4.25 mg/ml from untreated corn cob of between 3 % and 5 % substrate concentration.

4.6 Ethanol production from maize cob in simultaneous hydrolysis and fermentation processes

Fermentation of maize cob hydrolysate by Saccharomyces cerevisiae yielded varying ethanol concentrations. A higher yield of ethanol was optimized on substrate loading with 8 % at 96 h of fermentation. The highest ethanol concentration was produced by cellulase of Nemania sp. KM02 (28.72 ± 3.82 mg/ml), followed by Xylaria sp. KM03 (28.59 ± 1.58 mg/ml) and Xylaria sp. KM01 (25.75 ± 0.21 mg/ml) respectively. Similar findings were recorded by Ref. [51] and Saliu and Sani [66] when corn cob was hydrolyzed by cellulases of A. niger, recording ethanol yield (6.17 % and 2.67 %) respectively after 72 h of fermentation by Saccharomyces cerevisiae. The presence of fermentable sugar in negative control also recorded low ethanol yield due to the presence of free soluble sugar in the substrate. Yeast, however, consumes sugar for the growth and production of other metabolic products. Moreover, since growth commences during the aerobic phase, some amount of sugar gets used up before the anaerobic stage which is characterized by ethanol production, [66].

Bioethanol yield generally increases during the fermentation period with maximum bioethanol production at 96 h of incubation. As ethanol level increases it reduces the pH that favors Saccharomyces cerevisiae which converts the sugar present in the medium to ethanol and also provides an acidic condition that prevents bacterial contamination during fermentation. Nester et al. [68] recorded similar results, however, the fermentation did not proceed after 96 h, and the projection was a decrease afterward as the yeast consumes these nutrients from the medium and depletes the amount of sugar as the fermentation progresses.

4.7 Conclusions

As a conventional plant, maize cob proved its feasibility as a viable feedstock and as a cheap substrate for cellulase and bioethanol production. In the study, the supremacy of the cellulase of Xylaria sp. KM03 in hydrolyzing maize cobs to fermentable sugars recorded 10.17 ± 0.37 mg/ml reducing sugars, while hydrolyzed by cellulase of Nemania sp. KM02 recorded 28.72 ± 3.82 mg/ml ethanol yield. From this study, it can be concluded that, Xylaria sp. KM01, Nemania sp.KM02 and Xylaria sp. KM03 isolated from decaying plant biomass are cellulase-producing fungi. Time of incubation significantly affected cellulase enzyme production by the three fungal isolates using cypress eucalyptus and maize cobs substrates. Pretreatment of substrates affected cellulase enzyme production by three isolated fungi. The crude enzymes produced by the three fungal isolates saccharified maize cobs to produce fermentable sugars. Hydrolysate from saccharified maize cobs was fermented to ethanol using Saccharomyces cerevisiae.

Data availability statement

The experimental data used to support the findings of this study are available from the corresponding author upon request and have not been deposited into a publicly available repository.

CRediT authorship contribution statement

Stephen M. Kamande: Writing – review & editing, Writing – original draft, Validation, Methodology, Investigation, Formal analysis, Data curation, Conceptualization. George I. Omwenga: Supervision, Project administration. Mathew P. Ngugi: Supervision.

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Appendix A Supplementary data

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Acknowledgments

This research was partially supported by an African Development Bank 10.13039/100013987 Government of Kenya grant (10.13039/100004425 ADB /GoK contact NO. MOHEST/HEST/31/2014–2015 ). Special thanks go to DAAD (German) and HELB (Kenya) secretariats for the scholarship grants to support this research work.

Appendix A Supplementary data to this article can be found online at https://doi.org/10.1016/j.heliyon.2024.e36802.
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References

1 World Bank World Development Report 2010: Development and Climate Change 2010 © Washington, DC http://hdl.handle.net/10986/4387License:CCBY3.0IGO
2 Tuo H. Energy and exergy-based working fluid selection for organic Rankine cycle recovering waste heat from high temperature solid oxide fuel cell and gas turbine hybrid systems International Journal of Energy Resources 37 14 2013 1831 1841
3 Zhao C. Ma Z. Shao Q. Li B. Ye J. Peng H. Enzymatic hydrolysis and physiochemical characterization of corn leaf after H-AFEX pretreatment Energy Fuels 28 7 2016 4288 4293
4 Kuhad R.C. Gupta R. Singh A. Microbial Cellulases and Their Industrial Application vol. 2011 2011 Enzyme Research 280696
5 Azhar S.H.M. Abdulla R. Jambo S.A. Marbawi H. Gansau J.A. Faik A.A.M. Rodrigues K.F. Yeasts in sustainable bioethanol production: a review Biochemistry and Biophysics Report 10 2017 52 61
6 Acharya B.K. Mohana S. Jog R. Divecha J. Madamwar D. Utilization of anaerobically treated distillery spent wash for production of cellulases under solid state fermentation J. Environ. Manag. 91 2010 2019 2027
7 Deswal D. Khasa Y.P. Kuhad R.C. Optimization of cellulose production by a brown rot fungus Fomitopsis sp. RCK under solid state fermentation Bioresour. Technol. 102 2011 6065 6072 21470856
8 Montoya S. Orrego C.E. Levin L. Growth, fruiting and lignocellulolytic enzyme production by the edible mushroom Grifola frondosa (maitake) World J. Microbiol. Biotechnol. 28 2011 1533 1541 22805935
9 Li H. Pu Y. Kumar R. Ragauskas A.J. Wyman C.E. Investigation of lignin deposition on cellulose during hydrothermal pretreatment, its effect of cellulose hydrolysis, and underlying mechanisms Biotechnol. Bioeng. 111 3 2014 485 492 24037461
10 Wang B.T. Hu S. Yu X.Y. Jin L. Zhu Y.J. Jin F.J. Studies of cellulose and starch utilization and the regulatory mechanisms of related enzymes in fungi Polymers 12 3 2020 530 32121667
11 Moon R.J. Martini A. Nairn J. Simonsen J. Cellulose nanomaterials review: structure, properties and nanocomposites Chem. Soc. Rev. 40 2011 3941 3994 21566801
12 Sawatdeenarunat C. Surendra K.C. Takara D. Oechsner H. Khanal S.K. s Anaerobic Digestion of Lignocellulosic Biomass: Challenges and Opportunitie 2015 Bioresource Technology 178 186
13 Guimarães L.H. Carbohydrates from biomass: sources and transformation by microbial enzymes Changed C. Carbohydrates - Comprehensive Studies on Glycobiology and Glycotechnology. Carbohydrates Technologies 2012 441 458 (Chapter 20)
14 Yue Z. Bin W. Baixu Y. Peiji G. Mechanism of cellobiose inhibition in cellulose hydrolysis by cellobiohydrolase Sci. China C Life Sci. 47 1 2004 18 24 15382672
15 Ouyang B. Wang G. Zhang N. Zuo J. Huang Y. Zhao X. Recent advances in β-glucosidase sequence and structure engineering: a brief review Molecules 25 13 2023 28
16 Pandey A. Nigam P. Selvakumar P. Soccol C.R. Solid state fermentation for the production of industrial enzymes Curr. Sci. 77 1999 149 162
17 Benatti A.L.T. Polizeli M. Lignocellulolytic biocatalysts: the main players involved in multiple biotechnological processes for biomass valorization Microorganisms 11 1 2023 162 36677454
18 Chukwuma O.B. Rafatullah M. Tajarudin H.A. Ismail N. Lignocellulolytic enzymes in biotechnological and industrial processes: a review Sustainability 12 2020 7282 2020
19 Ilić N. Milić M. Beluhan S. Branković S.D. Cellulases: From Lignocellulosic Biomass to Improved Production Energies 16 8 2023 3598
20 Archana A. Satyanarayana T. Xylanase production by thermophilic Bacillus licheniformis A 99 in solid state fermentation Enzym. Microb. Technol. 21 1997 12 17
21 Pandey A. Francis F. Soccol C.R. General aspects in solid state fermentation Pandey Ashok Concise Encyclopedia of Bioresource Technology 2004 Haworth Press New York 702 718
22 Raimbault M. General and microbiological aspects of solid substrate fermentation Electron. J. Biotechnol. 1 3 1998 174 189
23 Arora D.K. Bhatnagar D. Handbook of Fungal Biotechnology 2003 CRC press 471 493
24 Sun Y. Cheng J. Hydrolysis of lignocellulosic materials for ethanol production: a review Bioresour. Technol. 83 2 2002 1 11 12058826
25 Martín C. Galbe M. Wahlbom C.F. Hahn-Hägerdal B. Jönsson L.J. Ethanol production from enzymatic hydrolysates of sugarcane bagasse using recombinant xylose utilizing Saccharomyces cerevisiae Enzym. Microb. Technol. 31 2002 274 282
26 Iqbal H.M.N. Kyazze G. Keshavarz T. Advances in the valorization of lignocellulosic materials by biotechnology: an overview Bioresources 2 2013 3157 3176
27 Jacob N. Prema P. Influence of mode of fermentation on production of polygalacturonase by a novel strain of Streptomyces lydicus Food Biotechnol. 44 2 2006 263 267
28 Tilay A. Annapure U. Novel simplified and rapid method for screening and isolation of polyunsaturated fatty acids producing marine bacteria Biotechnology Research International 542721 2012 1 8
29 Legodi M.L. La Grange D. Jansen van Rensburg E.L. Ncube L. Isolation of Cellulose Degrading Fungi from Decaying Banana Pseudostem and Strelitzia Alba vol. 2019 2019 Enzyme Research 1390890
30 Sukesh K. An Introduction to Industrial Microbiology. S 2010 Chand Publishing New Delhi 22
31 White T.J. Bruns T. Lee S. Taylor J.W. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics Innis M.A. Gelfand D.H. Sninsky J.J. White T.J. PCR Protocols 1990 Academic Press New York, NY 315 322
32 Graham G.C. Meyers P. Henry R.J.A. Simplified method for the preparation of fungal genomic DNA for PCR and RAPD analysis Biotechniques 16 1994 48 50 8136138
33 Delgado-Serrano L. Restrepo S. Bustos J.R. Zambrano M.M. Anzola J.M. Mycofier: a new machine learning-based classifier for fungal ITS sequences BioMed Central Research Notes 9 2016 402 27516337
34 Tamura K. Stecher G. Kumar S. MEGA11: molecular evolutionary genetics analysis version 11 Mol. Biol. Evol. 38 7 2021 3022 3027 33892491
35 Saitou N. Nei M. The neighbor-joining method: a new method for reconstructing phylogenetic trees Mol. Biol. Evol. 4 1987 406 425 3447015
36 Miller G.L. Use of dinitrosalicylic acid reagent for determination of reducing sugar J. Anal. Chem. 31 1959 426 428
37 Ghose T.K. Measurement of cellulase activities Pure Appl. Chem. 59 1987 257 268 1987
38 Herr D. Secretion of cellulase and beta-glucosidase by Trichoderma viride ITCC-1433 in submerged culture on different substrates Biotechnol. Bioeng. 21 8 1979 1361 1371 1979 Aug 110377
39 Jasińska K. Zieniuk B. Nowak D. Fabiszewska A. Studies on the catalytic properties of crude freeze-dried preparations of yarrowia lipolytica extracellular lipases for geranyl ester derivative synthesis Biomolecules 11 6 2021 839 34200103
40 Suh S. Zhang N. Nguyen N. Gross S. Blackwell M. Lab Manual for Yeast Study. Mycology Lab 2007 Louisiana state university. Statistical assistance 200 208
41 Alfenore S. Molina-Jouve C. Guillouet S. Uribelarrea J.L. Goma G. Benbadis L. Improving ethanol production and viability of Saccharomyces cerevisiae by a vitamin feeding strategy during fed-batch process Appl. Microbiol. Biotechnol. 60 1 2002 67 72 12382043
42 Mandels M. Hontz L. Nystrom J. Enzymatic Hydrolysis of Waste Cellulose. Biotechnology and Bioengineering xvi 1974 U.S. Army Natick Laboratories 1471 1493 Natick Massachusetts 01760
43 Caputi A. Ueda M.T. Brown T. Spectrophotometric determination of ethanol in wine AJEV (Am. J. Enol. Vitic.) 19 1968 L160 L165
44 Rahnama N. Foo H.L. Abdul Rahman N.A. Saccharification of rice straw by cellulose from a local Trichoderma harzianum SNRS3 for biobutanol production BMC Biotechnol. 14 2014 103 25496491
45 R Core Team R: A Language and Environment for Statistical Computing_ 2018 R Foundation for Statistical Computing Vienna, Austria https://www.R-project.org/
46 Kumar S. Stecher G. Li M. Knyaz C. Tamura K. Mega X: molecular evolutionary genetics analysis across computing platforms Mol. Biol. Evol. 35 6 2018 1547 29722887
47 Zhang Y.H.P. Himmel M.E. Mielenz J.R. Outlook for cellulase improvement: screening and selection strategies Biotechnol. Adv. 24 2006 452 481 16690241
48 Zakpaa H.D. Mak-Mensah E.E. Johnson F.S. Production of bio-ethanol from corn cobs using Aspergillus Niger and saccharomyces cerevisae in simultaneous saccharification and fermentation Afr. J. Biotechnol. 8 2009 3018 3022
49 Felsenstein J. Confidence limits on phylogenies: an approach using the bootstrap Evolution 39 1985 783 791 28561359
50 Kimura M. A simple method for estimating evolutionary rate of base substitutions through comparative studies of nucleotide sequences J. Mol. Evol. 16 1980 111 120 7463489
51 Ado S.A. Kachalla G.U. Tijjani M.B. Aliyu M.S. Ethanol production from corn cobs by co-cultures of saccharomyces cerevisiae and aspergillus Niger Bayero J. Pure Appl. Sci. (Ankara) 2 2 2009 99 101
52 Forrest A.K. Hernandez J. Holtzapple M.T. Effects of temperature and pretreatment conditions on mixed-acid fermentation of water hyacinths using a mixed culture of thermophilic microorganisms Bioresour. Technol. 101 2010 7510 7515 20466539
53 Ma F. Yang N. Xu C. Yu H. Wu J. Zhang X. Combination of biological pretreatment with mild acid pretreatment for enzymatic hydrolysis and ethanol production from water hyacinth Bioresour. Technol. 101 2010 9600 9604 20705458
54 Gao J. Chen L. Yan Z. Wang L. Effect of ionic liquid pretreatment on the composition, structure and biogas production of water hyacinth (Eichhornia crassipes) Bioresour. Technol. 132 2013 361 364 23186677
55 Timung R. Mohan M. Chilukoti B. Sasmal S. Banerjee T. Goud V.V. Optimization of dilute acid and hot water pretreatment of different lignocellulosic biomass: a comparative study Biomass Bioenergy 81 2015 9 18
56 Yan J. Wei Z. Wang Q. He M. Li S. Irbis C. Bioethanol production from sodium hydroxide/hydrogen peroxide pretreated water hyacinth via simultaneous saccharification and fermentation with a newly isolated thermo tolerant Kluyveromyces marxianu strain Bioresour. Technol. 193 2015 103 109 26119051
57 Kakde P.R. Aithal S.C. Effect of pretreatment on agro-residues for cellulase production by solid state fermentation using fungi International Journal of Recent Scientific Research 11 1 2020 36903 36906 2020
58 Ojumu T.V. Solomon B.O. Betiku E. Layokun S.K. Amigun B. Cellulase Production by Aspergillus flavus Linn isolate NSPR 101 fermented in sawdust, bagasse and corn cob Afr. J. Biotechnol. 2 2003 150 152
59 Xia A. Cheng J. Song W. Yu C. Zhou J. Cen K. Enhancing enzymatic saccharification of water hyacinth through microwave heating with dilute acid pretreatment for biomass energy utilization Energy 61 2013 158 166
60 Pandey S. Shrivastava M. Shahid M. Kumar V. Singh A. Trivedi S. Srivastava Y.K. Trichoderma species cellulases produced by solid state fermentation J. Data Min. Genom. Proteonomics 6 2015 170
61 Ilyas U. Ahmed S. Majeed A. Nadeem M. Biohydrolysis of Saccharum spontaneum for cellulase production by Aspergillus terreus Afr. J. Biotechnol. 21 2012 4914 4920
62 Gautam S.P. Bundela P.S. Pandey A.K. Khan J. Awasthi M.K. Sarsaiya S. Optimization for the production of cellulase enzyme from municipal solid waste residue by two novel cellulolytic fungi Biotechnology Resources International 81 2011 4 25
63 Yadav P.R. Chauhan P.B. Gahlout M. Isolation, screening and optimization of process parameters for enhanced production of cellulase by solid state fermentation Int. J. Adv. Res. Biol. Sci. 3 2016 21 27
64 Bhat B.A. Khan G.R. Asokan K. Role of substrate effects on the morphological, structural, electrical and thermoelectrical properties of V2O5 thin films Resour. Adv. 5 2015 552 602
65 Yoon L.W. Ang T.N. Ngoh G.C. Chua A.S.M. Fungal solid-state fermentation and various methods of enhancement in cellulase production Biomass Bioenergy 67 2014 319 338
66 Saliu B.K. Sani A. Bioethanol potentials of corn cob hydrolysed using cellulases of Aspergillus Niger and Penicillium decumbens EXCLI J 11 2012 468 479 2012 Aug 15 27418920
67 Jaafaru M.I. Fagade O.E. Cellulase production and enzymatic hydrolysis of some selected local lignocellulosic substrates by a strain of Aspergillus Niger Res. J. Biol. Sci. 2 2007 13 16
68 Nester E.W. Anderson D.G. Roberts C.E. Pearsall N.N. Nester M.T. Dynamics of Prokaryotic Growth in Microbiology: A Human perspective.3 Rd 2001 McGraw-Hill New York 87 108
