Next Article in Journal
Exploring Volatile Profiles in Cactus-Based Fermented Beverages: Effects of Fermentation Method
Previous Article in Journal
Evaluation of Carboxymethyl Cellulose as an Additive for Selective Protein Removal from Wine
Previous Article in Special Issue
Advancing Thermophilic Anaerobic Digestion of Corn Whole Stillage: Lignocellulose Decomposition and Microbial Community Characterization
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Fermentation
Volume 11
Issue 5
10.3390/fermentation11050274
fermentation-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Improved Cellulolytic Activity of Alternaria citri: Optimization and EMS Treatment for Enhanced Cellulase Production

by
Sibtain Ahmed
*,
Hina Andaleeb
*,
Aqsa Aslam
,
Junaid Ahmad Raza
,
Sheikh Muhammad Yahya Waseem
,
Atayyaba Javaid
and
Chand Talib
Department of Biochemistry, Bahauddin Zakariya University, Multan 60800, Pakistan
*
Authors to whom correspondence should be addressed.
Fermentation 2025, 11(5), 274; https://doi.org/10.3390/fermentation11050274
Submission received: 27 March 2025 / Revised: 7 May 2025 / Accepted: 8 May 2025 / Published: 11 May 2025
(This article belongs to the Collection Bioconversion of Lignocellulosic Materials to Value-Added Products)
Browse Figures
Versions Notes

Abstract

:
Fungal species secrete various enzymes and are considered the primary sources of industrially important cellulases. Cellulases are essential natural factors for cellulose degradation and have attracted significant interest for multiple applications. However, reducing the cost and enhancing cellulase production remains a significant challenge. Mutagenesis has opened a new window for enhancing enzyme secretion by modifying the organism’s genome. In this study, cellulases from Alternaria citri were produced and characterized, and the optimization for ideal fermentation conditions was performed for three types of cellulases (endoglucanase, exoglucanase, and β-glucosidase) by a wild-type (A. citri) and a mutant strain (A. citri 305). Ethyl methanesulfonate, a chemical mutagen, was used to enhance cellulase production by A. citri. The results demonstrate the improved cellulolytic ability of the mutant strain A. citri 305 utilizing lignocellulosic waste substances, particularly, orange-peel powder, wheat straw, sugarcane bagasse, and sawdust, making this study economically valuable. This evokes the potential for multi-dimensional applications in enzyme production, waste degradation, and biofuel generation. This study highlights that the activity of cellulases to hydrolyze various lignocellulosic substrates is enhanced after mutagenesis.

1. Introduction

Due to their natural abundance, microorganisms are considered natural producers of enzymes and are, therefore, important for biomass conversion [1]. Other than some bacterial species and yeast genera, filamentous fungi of Aspergillus, Chaetomium, Paecilomyces, Penicillium, and Trichoderma genera have also been used for enzymes and biomass production [2,3]. Many fungi are well-known producers of cellulases and xylanases [4]. Among all other sources of lignocellulolytic enzymes, microbial sources are considered primary sources [5]. Microbial cellulases have become the most popular biocatalysts because of their complexity [6] and broad-spectrum industrial applications [7,8]. Different fungal species have been reported to secrete cellulases and are being used as commercial sources of cellulases at the industrial level [1]. Filamentous fungi are well known for the production of enzymes [9]. Alternaria citri, belonging to the phylum Ascomycota, class Dothideomycetes, and order Pleosporales, causes Brown rot of citrus fruits, which has proven to be among the most serious and widely studied issues [10]. A. citri is found in all regions where citrus fruits are cultivated and harvested around the world. Due to its easy and rapid growth cycle, it has gained significant interest among researchers. Every year, a considerable amount of citrus fruit yield is lost due to this pathogenic fungus. It secretes various enzymes, including endogenous polygalacturonase and cellulases, which can degrade the plant cell wall. In this study, we explored the potential of wild and mutant strains of Alternaria citri for cellulase production.
Cellulose, an unbranched biopolymer [11] and a fundamental polysaccharide comprising β-1,4 glycosidic linkages, is a significant component of bacterial and plant cell walls [12,13]. Around 45% of all complex organic matter accounts for cellulose [14], which is known to be a prestigious and sustainable biopolymer on Earth [15,16]. The environmentally friendly hydrolysis of cellulose to gain reducible fermentable sugars is a significant process that finds applications in bioethanol production [17,18]. The most preferred method for the hydrolysis of cellulose to fermentable sugars is the enzymatic conversion of lignocellulosic biomass to valuable products over chemical conversions [19], because it promotes the green industry and is more environmentally friendly [5] and economically feasible [20] and requires less energy [21,22].
Many microbial genera from bacteria and fungi have been found to produce industrially essential enzymes [23]. Cellulases, including exoglucanase (EC 3.2.1.74), endoglucanase (EC 3.2.1.4), and β-glucosidases (EC 3.2.1.21), work synergistically [24] and find a wide range of applications in the biorefinery [25,26], beverage, and fruit juice industries [6]; bioethanol production [27,28]; textile industries [29]; paper and pulp manufacturing industries [30]; and pharmaceutical industries [31]. Because of their vital role in the natural carbon cycle [32], the industrial demands of cellulases are increasing daily; hence, the enhancement of cellulase production has grabbed the attention of researchers [33]. Due to their wide range of applications, cellulases are recorded as the third most industrially used enzyme group [34]. Cellulases produced by fermentation technologies are widely used in various industries, such as textiles, pulp and paper, and bioenergy production [35]. Because of the low availability [36] and high cost of cellulases [34,37], the cost reduction of cellulases has become a promising challenge [38].
Due to increasing industrialization and various household activities, much agricultural, industrial, and municipal waste is produced on Earth [39]. Bioconversion of agricultural waste through microbial fermentation is the natural way to recover resources [40]. The agro-industrial waste left on farmlands comprises cellulose and hemicellulose, which can be utilized as a substrate for enzyme production [41]. Although fungal sources produce enzymes in bulk [42], many filamentous fungi yield less cellulase because of their high viscosity during their growth period [43]. Great efforts have been made to enhance enzyme production through various strategies to minimize the gap between demand and the production of industrially important enzymes. One well-known strategy is the alteration of the microbial genome through mutagenesis [44]. A significant enhancement in enzyme secretion occurs through physical and chemical mutagenesis [45]. Mutagens are physical or chemical agents that alter the genome of living organisms, naturally causing the organism to undergo mutation. Compared to physical mutagens, chemical mutagens cause more gene mutations but fewer chromosomal mutations [46]. Ethyl methanesulfonate (EMS) is a chemical mutagen that owes its biological reactivity to its ethyl group. EMS alkylates guanine bases and due to this ethyl group, alkylated G pairs with T instead of C, primarily resulting in G/C-to-A/T transitions [47,48].
Several fungal strains of the genus Alternaria have been reported for cellulase production, but the data on cellulase production by Alternaria citri is very scarce. This study provides information on the industrial potential of A. citri (accession no. FCBP-PTF-1092) for cellulase production due to its high production yield and ease of cultivation. EMS treatment has been extensively used in strain improvement and enzyme production enhancement in various genera and species; however, exposing Alternaria citri to EMS for enhancing cellulase (carbohydrase) production contributes to the novelty of this study and distinguishes it from existing commercial sources of cellulases. Furthermore, this strain utilizes agro-industrial waste, including wheat straw, sugarcane bagasse, orange peel, and sawdust as substrates for cellulase production, making this study economically valuable.
This study contributes to overcoming the gap between increasing industrial demands and limited availability of efficient microbial sources for a commercially and industrially important enzyme system: cellulases. Cellulases were optimally produced from Alternaria citri utilizing lignocellulosic biomass, and their cellulolytic potential was enhanced through chemical mutagenesis.

2. Materials and Methods

2.1. Microorganism and Culture Conditions

Alternaria citri (accession no. FCBP-PTF-1092), isolated from citrus fruit, from Lahore (24–25 ℃, 29.12.09), was obtained from the First Fungal Culture Bank of Pakistan (FCBP), Department of Plant Pathology, Faculty of Agricultural Science (IAGS), University of the Punjab, Lahore, Pakistan. The strain was streaked onto potato dextrose agar (PDA) plates and incubated for 5 days at 25 ± 0.5 ℃. Isolated spores were maintained on a Malt extract agar slant and stored at 4 ℃ [49].

2.2. Screening of A. citri Cellulases

The cellulase production by A. citri was determined by growing it on a Carboxymethyl cellulose–potato dextrose agar (CMC-PDA) medium [8,50], and the plates were incubated for 120 h at 25 ℃. After incubation, the petri plates were flooded with a Congo red solution and left for 15 min [51]. The plates were washed with 1M of a saline solution to observe the zone of hydrolysis around the fungal spot [52].

2.3. Chemical Mutagenesis Using EMS

The strain improvement of A. citri was performed by chemical mutagenesis through EMS [37]. The EMS concentration range selected for mutagenesis was 500 µg/mL to 10 mg/mL (500 µg/mL, 1 mg/mL, 3 mg/mL, 5 mg/mL, 7 mg/mL, and 10 mg/mL). All of these EMS solutions were added to the spore suspension and left for around 60 min at room temperature. The tubes were centrifuged at 12,000 rpm for 15 min. The supernatant was discarded, sterile saline water was added, and the tubes were centrifuged again at 12,000 rpm for 15 min. After washing the pellets twice with saline water [53], the pellets were streaked on a PDA medium and incubated for 6 days at 25 ± 0.5 ℃.

2.4. Screening and Selection of Mutant Strains

All the mutant spores were streaked on the CMC-PDA medium and were incubated for 6 days at 28 ± 0.5 ℃. The plate assays for cellulase production from mutant strains were also performed by the Congo red method [52]. After the confirmation of cellulase production by A. citri through plate assays, the enzyme was produced in liquid media through submerged fermentation [54] using the Mandels and Weber method [55] for secondary screening. The mutant with the highest cellulolytic activity was selected for further studies and given the name A. citri 305. The spore suspension was inoculated into a production medium and incubated at 25 ± 0.5 ℃ in an orbital shaker incubator at 110–120 rpm [56]. The culture was harvested at 24 h intervals and centrifuged at 6,000 rpm for 15 min. The supernatant was used as a crude extracellular enzyme for the enzyme assays. The enzyme was stored at −20 ℃ in a freezer after adding 5% glycerol [56].

2.4.1. Endoglucanase Assay

Endoglucanase (CMCase activity) was determined using 1% w/v CMC [57,58] prepared in 0.05 M of a sodium citrate buffer (pH 5) [41]. The CMC solution was pre-incubated, followed by adding an equal volume of enzyme extract, and the reaction mixture was incubated at 50 ℃ for 20 min [59].
After the enzyme substrate reaction, a 3,5-dinitrosalicylic acid reagent was added to the reaction mixture, and the solution was boiled for 10 min [59]. The tubes were placed in ice cold water, and the absorbance was recorded at a 540 nm wavelength in a UV–visible spectrophotometer (JENWAY, Model 6300, Manufactured in the UK by Cole-Parmer Ltd., Stone, Staffs, UK, ST15 0SA) [60]. The amount of reducing sugars released in the test samples was determined based on their absorbance values using the glucose standard curve [61]. The unit of endoglucanase activity taken was IU/mL (International unit per milliliter). One International unit (IU) of endoglucanase activity was defined as the amount of enzyme that released 1 µmol of reducing sugars per minute under the assay conditions [62,63].

2.4.2. Exoglucanase Assay

Exoglucanase (FPase activity) was quantified using the fine powder of filter paper as a substrate [64], according to the standard method [65,66]. The filter paper solution was pre-heated, followed by the addition of enzyme extract, and incubated at 50 ℃ for 30 min [33,34,62]. After the enzyme substrate reaction, the exoglucanase activity was determined by a DNS reagent following the similar protocol used for endoglucanase activity. One unit (U) of exoglucanase activity was defined as the amount of enzyme that released 1 µmol of reducing sugars per minute under the assay conditions [66,67].

2.4.3. β-Glucosidase

β-glucosidase (βGLase/cellobiase) activity was determined by using cellobiose as a substrate [68,69,70]. The substrate was pre-incubated, and the enzyme was added to it. The reaction mixture was incubated at 50 ℃ for 20 min. After the enzyme substrate reaction, a GOD-POD reagent [70] was added, and the absorbance was checked at 515 nm. One unit of β-glucosidase activity was defined as the amount of enzyme necessary to liberate 2 mol of glucose per minute during cellobiose breakdown [37].

2.5. Optimization of Physical Factors for Cellulase Production

The one-factor-at-a-time method was utilized to check the effect of various physical parameters on the production of all three types of cellulases from the wild A. citri and mutant A. citri 305 strains. The major factors to be investigated were temperature, pH, and the incubation period [6].
To select the optimum temperature for cellulase production, both the wild and the mutant strains were incubated at varying temperatures (20 ℃, 25 ℃, 30 ℃, 35 ℃, and 40 ℃).
The optimum pH for cellulase production in liquid culture media was selected by growing A. citri and A. citri 305 in a range of pH 4 to 6 (4, 4.5, 5, 5.5, and 6). The enzyme activity was determined by harvesting the enzyme and then performing its quantitative analyses [71].
The incubation period ranged from 3 to 8 days to check the optimum incubation period. Enzyme activity assays were carried out at regular 24 h intervals [72].
The physical parameters were optimized for all three cellulases, where CMC was used for endoglucanase activity, filter paper for exoglucanase activity, and cellobiose for β-glucosidase activity.

2.6. Optimization of Nutritional Requirements for Cellulase Production

To obtain the optimum substrate concentration, a specific concentration range of substrate was used, and the enzymatic activity was observed. To check the best carbon source for cellulase production by the wild and mutant strains of A. citri, the flasks were supplied with 0.1% glucose, maltose, sucrose, galactose, and lactose [50]. The culture was grown at optimum conditions. The enzyme activity was determined to obtain the most effective carbon source for cellulase production [51]. To optimize the nitrogen source requirements, four nitrogen sources, including peptone, ammonium sulfate, sodium nitrate, and urea, were added to the cellulose-containing culture medium in varying concentration ranges [73].

2.7. Cellulase Purification from Wild A. citri and Mutant A. citri 305

The enzyme was purified by ammonium sulfate precipitation [74]. The crude enzyme supernatant was subjected to 0–40% and 40–70% saturation. Ammonium sulfate was added gradually with continuous shaking. The saturated enzyme solution was centrifuged at 10,000 rpm for around 10–15 min. The pellets were re-suspended in a 50 mM sodium citrate buffer, and desalting was carried out by dialyzing the samples [51] against the same buffer for 24 h, with the buffer changing every 6 h [8].

2.8. Determination of Protein Concentration

The protein concentration of the enzyme solutions was determined by the Lowry method, using bovine serum albumin (BSA) as a standard [51].

2.9. Enzyme Characterization from Wild A. citri and Mutant A. citri 305

2.9.1. Effects of Temperature, pH, Metal Ions, and Additives on Purified Cellulase Activity

The effect of varying temperatures on the activity of cellulases was analyzed at a temperature range of 20–70 ℃, with a constant gap of 10 ℃ [75]. After the optimization of the temperature parameter, the thermo-stability of the enzyme was analyzed by incubating the enzyme at the temperatures of 40 ℃, 50 ℃, and 60 ℃ for 0–4 h by measuring its activity after every 30 min.
The effect of varying pHs on the activity of cellulases was analyzed at the pH range of 3–8 [76]. The pH was maintained by using different buffers (a citrate buffer for pH 3–5, a phosphate buffer for pH 6–7, and a tris HCl buffer for pH 8) [18]. To observe the pH stability of the enzyme, it was pre-incubated at 50 ℃ for 0–4 h at pH 5, 6, and 7, and then, its activity assays were performed every 30 min. The activities were analyzed against the following substrates: CMC, filter paper powder, and cellobiose.
To observe the effects of metal ions on cellulase activity, 10% of metal ions, namely, NaNO3, KH2PO4, CaCl2, MgSO4, FeSO4, MnSO4, CoCl2, and ZnSO4, were incubated with the reaction mixture, and the enzyme activity was recorded. A total of 5 mM of various additives, such as Tween 80, SDS, urea, and EDTA, was used to evaluate their effects on cellulolytic activity [8,77].

2.9.2. Enzyme Kinetics of Purified Cellulases from Wild A. citri and Mutant A. citri 305

The kinetic parameters Km and Vmax of cellulases were calculated by plotting initial enzyme reaction velocities against the substrate concentration in the Lineweaver–Burk transformations of the Michaelis–Menten equation [78,79]. The substrates used for endoglucanase, exoglucanase, and β-glucosidase were CMC, filter paper powder, and cellobiose, respectively [37]. The reactions were carried out under recently optimized conditions.

2.10. Statistical Analysis

All of the assays of this research were conducted in triplicate. The mean and standard deviation were calculated using Microsoft Excel, and the results are presented as the mean ± standard deviation (SD). The standard deviation represents experimental variations (SD < 5%). The data were analyzed statistically using the GraphPad Prism 9 software (Version 9.5.1 (733). The statistical significance of the data (p <0.05) was determined by applying One-way ANOVA with Tukey’s post hoc analysis. All of the graphs were created using OriginPro (Origin 2024).

3. Results and Discussion

3.1. Effects of EMS Mutagenesis and Cellulolytic Screening of Mutants

A. citri spores were treated with different EMS concentrations for 30, 60, and 90 min of incubation. As the exposure time increased (0, 30, 60, and 90 min), the survival rate started to decrease, indicating that after a specific time, the EMS started to kill the fungal spores. The best outcomes of EMS treatment were observed for the 60 min exposure, and cellulase secretion was also observed to be elevated. The mutants of 60 min of exposure time, treated with 0.05%, 0.1%, 0.3%, 0.5%, and 0.7% EMS, were further selected for cellulase production. These mutants were named M1, M2, M3, M4, and M5, respectively; re-streaked on PDA plates; and stored in the refrigerator for future use. The highest fungal growth among the mutants was observed in the M1 (EMS0.05%-60min) mutant. As the EMS concentration increased, the number of fungal colonies started to decrease (Figure 1B). Minimum growth was observed for M5 (EMS1%-60min), which was considered to be a lethal dose for A. citri.
In the cellulase plate assays, the cellulase production by the wild and all mutant strains was confirmed by forming a hydrolysis zone (Figure 1A). In a quantitative assay, the maximum cellulase production was observed in the M4 mutant treated with 0.5% EMS (Figure 1B), which was selected as the best mutant strain and named A. citri 305.
Each fungal strain has its own lethal dose of EMS. As the EMS concentration increases, the germination rate of the fungal spores and seedlings decreases [79]. Higher EMS concentrations can cause more severe damage and even lethality in some cases. On the other hand, optimum EMS concentrations can induce significant changes and mutations in fungi without compromising viability [80].
The viability of mutant strains for cellulase production is an important factor to be considered, which is typically evaluated through successive sub-culturing of the mutants for multiple generations [81]. Random mutagenesis enhances enzyme production in microorganisms that can be observed through the formation of hydrolysis zones in plate assays [82,83]. Mutagenesis by different concentrations of EMS enhances enzyme production in several microorganisms [84,85], as in Aspergillus terreus and Trichoderma viride, whose production of cellulases (CMCase, FPase, and β-glucosidase) enhanced significantly after EMS treatment [65,84].

3.2. Optimization of Physical Parameters for Cellulase Production by Wild A. citri and Mutant A. citri 305

The maintenance of physical conditions is essential for the physiology of microorganisms, specifically, their growth and enzyme production. The production of cellulases, including endoglucanase, exoglucanase, and β-glucosidase, is reported to be influentially increased when the conditions and growth parameters are optimized [50,70].

3.2.1. Temperature Optimization

The optimum temperature for the growth and cellulase production by wild A. citri and mutant A. citri 305 was checked by growing both strains at different temperatures, and the enzyme activity was quantified at all temperatures. Among several temperature values, ranging between 20 and 45 ℃, the temperature of 25 ℃ was observed to be the optimum temperature for cellulase production, and enzyme production decreased with a further temperature rise (Figure 2).
Temperature is regarded as a key parameter that affects and controls microbial growth, cell development, various metabolic reactions, and enzyme production by altering the physical and physiological characteristics of microbes. Therefore, the release of extracellular enzymes is found to be greatly dependent on temperature [86]. At higher temperatures, the loss of enzyme activity is observed as the enzymes lose their functional characteristics [87]. Fungi produces maximum cellulases at an optimum pH of 4–4.5 and at a temperature of 28 ℃ [88].

3.2.2. Optimization of pH

The optimum pH required for growth and cellulase production by wild A. citri and mutant A. citri 305 was determined by growing the fungus in a range of pH values at the optimum temperature. The optimum pH for cellulase production was found to be 4.5, and the cellulolytic activity decreased with an increasing pH (Figure 3).
pH influences microbial cellulase-producing capabilities by affecting the surface charge and nature of cell membranes. This is a fact that lies behind the varying capacity of microbes to absorb required nutrients and, hence, to grow well and secrete enzymes [16]. The growth of fungus has been found to be in direct relation to enzyme production. Therefore, the higher the growth rate of the fungus, the greater the cellulase secretion [8]. Most fungi grow and secrete metabolites at a pH range of 4.5–5.5, and enzymes are mostly stable at this pH [89]. Aspergillus niger, and several other strains show the highest cellulase production at pH 5 [90]. The production of cellulase by Trichosporon insectorum was reported to reach its maximum at pH 4 [35].

3.2.3. Incubation Period Optimization

The optimization of the incubation period was carried out by incubating both A. citri and A. citri 305 for 24–168 h at optimum conditions. The enzyme was harvested at every 24 h interval, and the activity was analyzed. Cellulase production increased with the incubation time, and the maximum enzyme was secreted on the 5th day (120 h) by the wild-type strain and on the 6th day (144 h) of incubation by the mutant strain. After the optimum incubation period, cellulase production was observed to be decreased (Figure 4).
Different fungi produced maximum cellulases at incubation periods of 96–144 h. Aspergillus sp. also gave maximum cellulase production at 5 days of incubation under solid state fermentation [6]. A. niger and A. flavus also produced maximum cellulases at 5 days of incubation, and after that, it continued to decrease [90]. Several fungal species secretes exoglucanase at pH 5 and at 6 days of incubation [91]. A mutant strain of T. harzianum and Penicillium funiculosum was reported to exhibit maximum cellulase secretion on the 6th day of culturing [37,83].

3.3. Optimization of Nutritional Requirements for Cellulase Production by A. citri and A. citri 305

3.3.1. Substrate Optimization

The substrate concentration has a great impact on enzyme production by any microorganism. Different substrates, including Carboxymethyl cellulose, wheat straw, orange-peel powder, sugarcane bagasse, and sawdust of Pine trees (Pinus roxburghii) and Gum Arabic trees (Vachellia nilotica), were used as a source of cellulose in submerged fermentation to optimize the best substrate for maximum cellulolytic activity. Maximum cellulase production was obtained by using CMC as a substrate, which was named as a standard substrate. Both A. citri and A. citri 305 exhibited significant cellulases activity using orange-peel powder and the sawdust of Pine trees and Gum Arabic trees, and lesser cellulase activity was obtained using wheat straw and sugarcane as a source of cellulose (Figure 5A). Therefore, orange peel was utilized as a cost-effective alternative to CMC. Since orange-peel powder was the best substrate for cellulase production, it was used in different concentrations to obtain the optimum substrate concentration. The optimum substrate concentration for maximum cellulase production was 1% for wild A. citri and 1.5% for the mutant A. citri 305 (Figure 5B).
The highest cellulase activity was obtained with 1% substrate [35]. maximum cellulase production from Aspergillus terreus was observed when cultivated on a 1% substrate [59]. Aspergillus niger was reported to exhibit higher cellulase production with corn stubble and sorghum bagasse as substrates [41]. Aspergillus uvarum expresses maximum cellulase using sorghum straw as a substrate [18]. CMC has been identified as the best optimized substrate for cellulase production by Cohnella xylanilytica [16].

3.3.2. Carbon Source Optimization

Carbon sources play a crucial role in the growth and development of microbial cultures. Microorganisms require carbon, nitrogen, and oxygen for nourishment and to produce their necessary life products. Their metabolic reactions utilize several organic compounds and produce others required for their survival. Carbon is a major element of life, as it is involved in the energy balance cycle.
The optimization of the best carbon source for cellulase production by A. citri and the mutant A. citri 305 was carried out by testing five carbohydrates. Maximum cellulase production was observed by utilizing glucose as a carbon source (Figure 6A). Therefore, the optimum glucose concentration for cellulase production was checked. It was observed that 0.2g/L of glucose is the optimum glucose concentration for cellulase production (Figure 6B).
Different organisms prefer different compounds as carbon sources. Glucose was used as the carbon source for the secretion of cellulases from Trichoderma afroharzianum [44] and various other microbial strains [92]. Alternaria brassicicola [51], Aspergillus chevalieri, and Trichoderma sp. show maximum cell growth and enzyme production when utilizing sucrose as a carbon source [93,94].

3.3.3. Nitrogen Source Optimization

A sufficient amount of nitrogen sources is necessary for the growth and development of microbes. Enzyme production is also dependent on several nitrogen sources. Ammonium sulfate (1.5 g/L) and sodium nitrate (1.5 g/L) significantly enhanced cellulase production. The optimum concentration of peptone for the production of cellulases was 0.1 g/L and 0.3 g/L, and urea was also observed to have an enhancing effect on cellulase production (Figure 7).
The cellulase production by different microbial strains was positively influenced using ammonium sulfate and sodium nitrate [70,95]. Along with some other factors, 0.125% peptone was sufficient for enhancing cellulase production by Aspergillus niger [87].
Biosynthetic capabilities of A. niger to produce cellulases were increased when urea was used a nitrogen source [96].

3.4. Cellulase Purification from Wild A. citri and Mutant Strain A. citri 305

A summary of the purification process of cellulases produced by A. citri and A. citri 305 is described in Table 1. The recovery yield of purified cellulases was maximum for endoglucanase at a value of 62.79% with a purification fold of 2.12 for A. citri and 43.07% with a purification fold of 3.95 for A. citri 305. However, the maximum specific activity was obtained from β-glucosidase (Table 1).
The enzyme purification experiments of B. tequilensis led to purified cellulase with a yield of 10% and a 9.89 purification fold [97]. The cellulase of B. licheniformis gave a purification fold of about 1.96 with a specific activity of 333.4 U/mg and a 51.1% yield after ammonium sulfate precipitation [98]. The purification fold of the CMCase B. subtilis subsp. subtilis was 5.7 times with a recovery yield of 0.73% [99]. The ammonium sulfate precipitation of thermophillic cellulase produced by a novel cellulolytic strain, Paenibacillus barcinonensis, led to a recovery yield of 25.59% with 2.97 purification folds [100].

3.5. Characterization of Purified Cellulases from Wild A. citri and Mutant Strain A. citri 305

3.5.1. Effect of Temperature on the Activity of Purified Cellulases

A broad temperature range (20–70 ℃) was used to check the influence of temperature on cellulase activity. The enzyme activity was found to increase with a rise in temperature of up to 50 ℃, the optimum temperature, and then decreased subsequently (Figure 8).
Acinetobacter junii also produced cellulases that had the highest activity at 50 ℃ [101]. The cellulases produced by Aspergillus uvarum showed maximum activity at 50 ℃ [18]. The optimum temperature for the cellulolytic activity of Bacillus tequilensis was reported at 50 ℃ [8].

3.5.2. Effect of pH on Purified Cellulase Activity

Enzyme activity is majorly affected by the pH of the substrate. The cellulolytic activity of purified cellulases extracted from wild A. citri and mutant A. citri 305 was analyzed at range of pH 3–8 to determine the optimum pH for cellulase activity. The maximum activity of cellulases was obtained at pH 6, and this was regarded as the optimum pH for cellulase activity (Figure 9).
Bacillus mycoides secretes cellulase that is active over a range of pH of 4–6 [102]. The cellulolytic activity of bacterium Bacillus tequilensis has also been observed to be maximum at pH 6.5 [8].

3.5.3. Effects of Metal Ions on Purified Cellulase Activity

This study involves an investigation of the impact of Sodium dodecyl sulfate (SDS), Tween 80, Ethylenediaminetetra acetic acid (EDTA), and urea. The influence of several metal ions and additives on cellulase activity was evaluated in the current study, and the results are depicted in Table 2. Some metal ions enhanced the activity, including Fe2+, Mg2+, and Zn2+. These metals increased the activity of cellulase -produced by both wild A. citri and the mutant A. citri 305. However, CoCl2 decreased cellulolytic activity significantly. NaNO3 decreased the activity of endoglucanase while stimulating the activity of exoglucanase and β-glucosidase. Only Tween 80 enhanced cellulolytic activity, while it was significantly decreased when pre-treated with urea. SDS and EDTA also decreased the activity of cellulases (Table 2).
Metal ions stabilize the enzyme substrate complex and work as cofactors of some enzymes and highly influence their activity, because most metal ions possess more than one positive charge [103]. The impact of surfactants has been observed to be an enhancing one, and these are categorized as inducers of enzyme activity [104]. The activity of cellulases produced by bacteria also increase after being treated with MgSO4, ZnSO4, and MnSO4 [8]. Fe2+ was also reported to enhance the cellulolytic activity of Aspergillus niger, while Co2+ decreases enzyme activity [105]. Na+ has been reported to have a negligible effect on the cellulase activity of Aspergillus niger [75].
Non-ionic detergents, like Tween 20, Tween 80, Span 80, and PEG 400, have been reported to be stimulants of cellulases [18], because these substances maximize the conversion of cellulose to reducing sugars, like glucose, hence enhancing the enzyme output [106]. SDS causes some unintended bonding in the proteins’ tertiary structures, hence disrupting the active sites of the enzymes. Ultimately, the proteins become unfolded, and their structure is distorted [107], and the enzymes lose their activity to a significant level. EDTA inactivates the cellulase enzyme by chelating the metal ions as enzyme cofactors, thereby inhibiting the cellulolytic potential [108]. Urea negatively influences the cellulolytic potential by decreasing cellulase activity, as it is known to disrupt the tertiary structure of proteins. The activity of cellulase produced by Bacillus tequilensis decreases in the presence of urea and EDTA [8].

3.5.4. Thermo-Stability Studies on Cellulases from Wild A. citri and Mutant Strain A. citri 305

Thermo-stability is actually the capacity of an enzyme to be active at a specific temperature for a specific time period. Thermo-stability assays were carried out at 40 ℃, 50 ℃, and 60 ℃ for the time period of 4 h. The activity was measured at every 30 min.
The β-glucosidase enzyme of the wild type A. citri and its mutant strain A. citri 305 is more heat stable at 40 ℃ and 50 ℃, as it retained more than 80% activity even after 4 h of incubation. But at 60 ℃, its activity dropped down to a significant level. The results of the thermo-stability assays of endoglucanase and exoglucanase were observed to be virtually similar, as both of these showed a significant decrease in the activity of the enzymes produced by the wild-type strain, and the mutant strain’s enzymes were notably more heat stable, as nearly 80% activity was recorded after 4 h of incubation (Figure 10). Therefore, it was concluded that the mutant strain secreted more heat-stable cellulases compared to the wild-type strain.

3.5.5. pH Stability Studies on Purified Cellulases from Wild A. citri and Mutant Strain of A. citri 305

The cellulases showed optimum activity at pH 6. Therefore, the pH stability assays were carried out at three pH values, including the optimum pH 6, its immediately lower value 5, and its immediately higher value 7.
All three enzymes were more stable at pH 6. Above and below this pH level, the wild-type enzymes were less stable and lost their activity over time, while the cellulases of the mutant strain were comparatively more stable and resistant to changes in pH (Figure 11). This is in accordance with studies where it has been reported that the cellulase of Aspergillus uvarum retains nearly 90% activity after 4 h of incubation and after 20 h of incubation, wherein the activity decreases to around 50% [18].

3.6. Kinetics Analysis of Purified Cellulase

The kinetic parameters Vmax and Km of all three cellulases were calculated by the Lineweaver–Burk plot at a range of substrate concentrations (0.5–10 mg/mL). The rate of reaction started to increase linearly with an increasing substrate concentration. The enzymes attained a maximum value at a specific substrate concentration, and afterward, it became almost constant, turning the curve of the graph into straight line. The maximum rate of reaction was attained by the enzyme of wild A. citri at a lower substrate concentration, while the enzyme of the mutant A. citri 305 attained the maximum reaction rate at a relatively higher substrate concentration.
The cellulases of the mutant strain show lower Michealis–Menten constant Km values (0.031± 0.001 for β-glucosidase, 0.295 ± 0.006 for endoglucanase, and 0.241 ± 0.007 for exoglucanase) than that of the wild A. citri (0.048 ± 0.001 for β-glucosidase, 0.492 ± 0.004 for endoglucanase, and 0.278 ± 0.011 for exoglucanase), indicating that the cellulases of the mutant strain have a higher substrate-binding capacity. As it requires a comparably lower substrate concentration to attain half maximum velocity, it has a higher efficiency value than that of the wild-type cellulases. The mutant strain’s cellulases have a relatively higher Vmax value (1.1281 ± 0.0041 for β-glucosidase, 0.060 ± 0.000 for endoglucanase, and 0.023 ± 0.000 for exoglucanase) than that of the wild-type strain (0.754 ± 0.011 for β-glucosidase, 0.039 ± 0.000 for endoglucanase, and 0.023 ± 0.000 for exoglucanase), supporting the high enzyme efficiency by explaining the fact that they possess the capability to convert substrates into products at a comparatively faster rate than that of the cellulases of wild A. citri, which have lower Vmax values (Table 3).

4. Conclusions

This study highlights the cellulolytic potential of A. citri utilizing lignocellulosic biomass, particularly, orange peel, wheat straw, sugarcane bagasse, and sawdust, and its enhancement by EMS treatment. As a low-cost substrate, lignocellulosic biomass has demonstrated its feasibility in enzyme production, waste management programs, and biofuel generation industries. In this study, the optimization of cellulase production was performed by a one-factor-at-a-time approach. The lower Km and the higher Vmax values of the cellulases secreted by the mutant A. citri 305 suggested that these cellulases are much more efficient than those of the wild A. citri. The expressed enzymes, endoglucanase, exoglucanase, and β-glucosidase, emphasized substrate specificities and are favorable for the degradation of various biomasses. The optimum cellulase activity was observed at a temperature of 50 ℃ and pH 6. Some metal ions, such as Fe2+, Mg2+, and Zn2+, enhanced the cellulases’ activity, while others, like Co2+, inhibited enzyme activity. Tween 80 was observed to increase enzyme activity, while SDS, EDTA, and urea were observed to be enzyme deactivators. The characterization studies of the cellulases indicated significant thermo-stability and pH stability, which are important parameters for enzyme storage and transport. Overall, this research contributes to meeting the gap between ever-rising commercial demands and limited effective microbial sources available for the production of cellulases and helps to reduce the cost of cellulases by utilizing waste substances to provide a multi-purpose approach where the waste degradation process is also considered along with industrially important enzyme production.

Author Contributions

S.A. and H.A.: writing—review and editing, writing—original draft, visualization, methodology, investigation, formal analysis, data curation, and conceptualization. S.A. and H.A.: investigation and formal analysis. A.A.: investigation and formal analysis. J.A.R., S.M.Y.W., A.J. and C.T.: data curation and conceptualization. S.A and H.A.: data curation, writing—review and editing. A.A. and J.A.R.: writing—original draft. A.A., S.A. and H.A.: writing—review and editing, writing—original draft, funding acquisition, conceptualization, supervision, and resources. All authors have read and agreed to the published version of the manuscript.

Funding

This research was supported by the Office of Research, Innovation and Commercialization (ORIC)-Bahauddin Zakariya University, Multan (Pakistan).

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

The original contributions presented in this study are included in the article. Further inquiries can be directed at the corresponding author.

Acknowledgments

The authors acknowledge the Office of Research, Innovation and Commercialization (ORIC)-Bahauddin Zakariya University, Multan, and the Department of Biochemistry, Bahauddin Zakariya University, Multan, Pakistan, for supporting this research.

Conflicts of Interest

The authors declare no conflicts of interest.

Abbreviations

The following abbreviations are used in this manuscript:
EMSEthyl methanesulfonate
CMCCarboxymethyl cellulose
OPOrange peel
SBSugarcane bagasse
WSWheat straw

References

  1. Ahmed, S.; Riaz, S.; Jamil, A. Molecular Cloning of Fungal Xylanases: An Overview. Appl. Microbiol. Biotechnol. 2009, 84, 19–35. [Google Scholar] [CrossRef] [PubMed]
  2. Ahmed, S.; Mustafa, G.; Arshad, M.; Rajoka, M.I. Fungal Biomass Protein Production from Trichoderma harzianum Using Rice Polishing. BioMed Res. Int. 2017, 2017, 6232793. [Google Scholar] [CrossRef] [PubMed]
  3. Ahmed, S.; Jabeen, A.; Jamil, A. Xylanase from Trichoderma harzianum: Enzyme Characterization and gene isolation. J. Chem. Soc. Pak. 2007, 29, 176–182. [Google Scholar]
  4. Ahmed, S.; Imdad, S.S.; Jamil, A. Comparative Study for the Kinetics of Extracellular Xylanases from Trichoderma harzianum and Chaetomium thermophilum. Electron. J. Biotechnol. 2012, 15, 3. [Google Scholar] [CrossRef]
  5. Benatti, A.L.T.; Polizeli, M.L.T. M Lignocellulolytic Biocatalysts: The Main Players Involved in Multiple Biotechnological Processes for Biomass Valorization. Microorganisms 2023, 11, 162. [Google Scholar] [CrossRef]
  6. Boondaeng, A.; Keabpimai, J.; Trakunjae, C.; Vaithanomsat, P.; Srichola, P.; Niyomvong, N. Cellulase Production under Solid-State Fermentation by Aspergillus Sp. IN5: Parameter Optimization and Application. Heliyon 2024, 10, e26601. [Google Scholar] [CrossRef]
  7. Korsa, G.; Konwarh, R.; Masi, C.; Ayele, A.; Haile, S. Microbial Cellulase Production and Its Potential Application for Textile Industries. Ann. Microbiol. 2023, 73, 13. [Google Scholar] [CrossRef]
  8. Malik, W.A.; Javed, S. Enhancement of Cellulase Production by Cellulolytic Bacteria SB125 in Submerged Fermentation Medium and Biochemical Characterization of the Enzyme. Int. J. Biol. Macromol. 2024, 263, 130415. [Google Scholar] [CrossRef]
  9. Liu, S.; Ahmed, S.; Zhang, C.; Liu, T.; Shao, C.; Fang, Y. Diversity and antimicrobial activity of culturable fungi associated with sea anemone Anthopleura xanthogrammica. Electron. J. Biotechnol. 2020, 44, 41–46. [Google Scholar] [CrossRef]
  10. Iqbal, S.; Atiq, M.; Sahi, S.T.; Akbar, N.; Rajput, N.A. Progressive Alterations in Mineral Contents in Citrus Genotypes toward Alternaria citri Causing Brown Spot of Citrus. PLoS ONE 2024, 19, e0306031. [Google Scholar] [CrossRef]
  11. Li, H.; Zhang, M.; Zhang, Y.; Xu, X.; Zhao, Y.; Jiang, X.; Zhang, R.; Gui, Z. Characterization of Cellulose-Degrading Bacteria Isolated from Silkworm Excrement and Optimization of Its Cellulase Production. Polymers 2023, 15, 4142. [Google Scholar] [CrossRef] [PubMed]
  12. Jakob, M.; Mahendran, A.R.; Gindl-Altmutter, W.; Bliem, P.; Konnerth, J.; Mueller, U.; Veigel, S. The Strength and Stiffness of Oriented Wood and Cellulose-Fibre Materials: A Review. Prog. Mater. Sci. 2022, 125, 100916. [Google Scholar] [CrossRef]
  13. Doan, C.T.; Tran, T.N.; Pham, T.P.; Tran, T.T.T.; Truong, B.P.; Nguyen, T.T.; Nguyen, T.M.; Bui, T.Q.H.; Nguyen, A.D.; Wang, S.-L. Production, Purification, and Characterization of a Cellulase from Paenibacillus Elgii. Polymers 2024, 16, 2037. [Google Scholar] [CrossRef] [PubMed]
  14. Pata, U.K. Linking Renewable Energy, Globalization, Agriculture, CO2 Emissions and Ecological Footprint in BRIC Countries: A Sustainability Perspective. Renew. Energy 2021, 173, 197–208. [Google Scholar] [CrossRef]
  15. Sharma, A.; Tewari, R.; Rana, S.S.; Soni, R.; Soni, S.K. Cellulases: Classification, Methods of Determination and Industrial Applications. Appl. Biochem. Biotechnol. 2016, 179, 1346–1380. [Google Scholar] [CrossRef]
  16. Boro, M.; Verma, A.K. Optimization of Cellulase Production by Cohnella Xylanilytica RU-14 Using Statistical Methods. Appl. Biochem. Biotechnol. 2024, 196, 2757–2770. [Google Scholar] [CrossRef]
  17. Abdelhamed, E.A.; Madian, H.R.; Salem, A.A.; Sidkey, N. Bioethanol Production from Hydrolysis and Fermentation of Rice Straw Using a Combination of Microbial Isolates. Egypt. J. Chem. 2024, 67, 167–180. [Google Scholar] [CrossRef]
  18. Kaur, G.; Taggar, M.S.; Kalia, A.; Krishania, M.; Singh, A. Fungal Secretomes of Aspergillus terreus Repertoires Cultivated on Native and Acid/Alkali Treated Paddy Straw for Cellulase and Xylanase Production. BioEnergy Res. 2024, 17, 145–159. [Google Scholar] [CrossRef]
  19. Ascencio, J.J.; Magalhães, L.S.; Ferreira, F.B.; Heinz, O.; Ferraz, A.; Chandel, A.K. Surfactant-Enhanced Enzymatic Hydrolysis of Eucalyptus Kraft Pulp: The Interrelationship Between Lignin Reduction and Sugar Recovery. Catalysts 2025, 15, 47. [Google Scholar] [CrossRef]
  20. Feng, J.; Techapun, C.; Phimolsiripol, Y.; Phongthai, S.; Khemacheewakul, J.; Taesuwan, S.; Mahakuntha, C.; Porninta, K.; Htike, S.L.; Kumar, A. Utilization of Agricultural Wastes for Co-Production of Xylitol, Ethanol, and Phenylacetylcarbinol: A Review. Bioresour. Technol. 2024, 392, 129926. [Google Scholar] [CrossRef]
  21. Rosales-Calderon, O.; Trajano, H.L.; Duff, S.J.B. Stability of Commercial Glucanase and β -Glucosidase Preparations under Hydrolysis Conditions. PeerJ 2014, 2, e402. [Google Scholar] [CrossRef] [PubMed]
  22. Hong, M.-R.; Kim, Y.-S.; Park, C.-S.; Lee, J.-K.; Kim, Y.-S.; Oh, D.-K. Characterization of a Recombinant β-Glucosidase from the Thermophilic Bacterium Caldicellulosiruptor saccharolyticus. J. Biosci. Bioeng. 2009, 108, 36–40. [Google Scholar] [CrossRef] [PubMed]
  23. Mushtaq, A.; Ahmed, S.; Mehmood, T.; Cruz-Reyes, J.; Jamil, A.; Nawaz, S. Cloning, Expression, and Characterization of a Metalloprotease from Thermophilic Bacterium Streptomyces thermovulgaris. Biology 2024, 13, 619. [Google Scholar] [CrossRef] [PubMed]
  24. Jalak, J.; Kurašin, M.; Teugjas, H.; Väljamäe, P. Endo-Exo Synergism in Cellulose Hydrolysis Revisited. J. Biol. Chem. 2012, 287, 28802–28815. [Google Scholar] [CrossRef]
  25. Gan, J.; Iqbal, H.M.N.; Show, P.L.; Rahdar, A.; Bilal, M. Upgrading Recalcitrant Lignocellulosic Biomass Hydrolysis by Immobilized Cellulolytic Enzyme–Based Nanobiocatalytic Systems: A Review. Biomass Convers. Biorefinery 2024, 14, 4485–4509. [Google Scholar] [CrossRef]
  26. Ahmed, S.; Bashir, A.; Saleem, H.; Saadia, M.; Jamil, A. Production and purification of cellulose-degrading enzymes from a filamentous fungus Trichoderma harzianum. Pak. J. Bot. 2009, 41, 1411–1419. [Google Scholar]
  27. Boro, M.; Verma, A.K.; Chettri, D.; Yata, V.K.; Verma, A.K. Strategies Involved in Biofuel Production from Agro-Based Lignocellulose Biomass. Environ. Technol. Innov. 2022, 28, 102679. [Google Scholar] [CrossRef]
  28. Da Silva, M.F.; Flaibam, B.; De Mélo, A.H.F.; Sampaio, U.; Clerici, M.T.P.S.; Goldbeck, R. Optimization of Enzymatic Hydrolysis of Cellulose Extracted from Bamboo Culm for Bioethanol Production by Saccharomyces cerevisiae Modified via CRISPR/Cas9. Food Res. Int. 2024, 192, 114768. [Google Scholar] [CrossRef]
  29. Singh, A.; Bajar, S.; Devi, A.; Pant, D. An Overview on the Recent Developments in Fungal Cellulase Production and Their Industrial Applications. Bioresour. Technol. Rep. 2021, 14, 100652. [Google Scholar] [CrossRef]
  30. Reis, C.E.R.; Libardi Junior, N.; Bento, H.B.S.; Carvalho, A.K.F.D.; Vandenberghe, L.P.D.S.; Soccol, C.R.; Aminabhavi, T.M.; Chandel, A.K. Process Strategies to Reduce Cellulase Enzyme Loading for Renewable Sugar Production in Biorefineries. Chem. Eng. J. 2023, 451, 138690. [Google Scholar] [CrossRef]
  31. Chukwuma, O.B.; Rafatullah, M.; Tajarudin, H.A.; Ismail, N. Lignocellulolytic Enzymes in Biotechnological and Industrial Processes: A Review. Sustainability 2020, 12, 7282. [Google Scholar] [CrossRef]
  32. Faheina Junior, G.S.; Sousa, K.A.; Zilli, J.E.; Vergara, C.; Pinto, G.A.S.; Santiago-Aguiar, R.S. Enhanced Cellulase Production by Talaromyces Amestolkiae CMIAT055 Using Banana Pseudostem. Waste Biomass Valorization 2022, 13, 3535–3546. [Google Scholar] [CrossRef]
  33. Fatima, N.; Ahmad, I.; Shakir, H.A.; Khan, M.; Franco, M.; Irfan, M. Production and characterization of cellulases from Aspergillus niger under static fermentation. Cellul. Chem. Technol. 2024, 58, 705–712. [Google Scholar] [CrossRef]
  34. Santos, G.B.; de Sousa Francisco Filho, Á.; da Silva Rodrigues, J.R.; de Souza, R.R. Cellulase Production by Aspergillus niger Using Urban Lignocellulosic Waste as Substrate: Evaluation of Different Cultivation Strategies. J. Environ. Manag. 2022, 305, 114431. [Google Scholar] [CrossRef] [PubMed]
  35. Touijer, H.; Benchemsi, N.; Irfan, M.; Tramice, A.; Slighoua, M.; Mothana, R.A.; Alanzi, A.R.; Dalila, B.; Bekkari, H. Statistical Optimization and Purification of Cellulase Enzyme Production from Trichosporon insectorum. Fermentation 2024, 10, 453. [Google Scholar] [CrossRef]
  36. Infanzón-Rodríguez, M.I.; del Moral, S.; Gómez-Rodríguez, J.; Faife-Pérez, E.; Aguilar-Uscanga, M.G. Second-Generation Bioethanol Production and Cellulases of Aspergillus niger ITV02 Using Sugarcane Bagasse as Substrate. BioEnergy Res. 2024, 17, 160–172. [Google Scholar] [CrossRef]
  37. Chavan, S.; Shete, A.; Dharne, M.S. Strain Improvement for Cellulolytic Enzymes for Effective Saccharification of Lignocellulosic Biomass by Mutant of Penicillium funiculosum NCIM 1228. Syst. Microbiol. Biomanufacturing 2024, 4, 716–730. [Google Scholar] [CrossRef]
  38. Liu, F.; Wang, Z.; Manglekar, R.R.; Geng, A. Enhanced Cellulase Production through Random Mutagenesis of Talaromyces pinophilus OPC4-1 and Fermentation Optimization. Process Biochem. 2020, 90, 12–22. [Google Scholar] [CrossRef]
  39. Asghar, A.; Afzaal, M.; Saeed, F.; Ahmed, A.; Ateeq, H.; Shah, Y.A.; Islam, F.; Hussain, M.; Akram, N.; Shah, M.A. Valorization and Food Applications of Okara (Soybean Residue): A Concurrent Review. Food Sci. Nutr. 2023, 11, 3631–3640. [Google Scholar] [CrossRef]
  40. Ahmed, S.; Ahmad, F.; Hashmi, A.S. Production of Microbial biomass protein by sequential culture fermentation of Arachniotus sp. and Candida utilis. Pak. J. Bot. 2010, 42, 1225–1234. [Google Scholar]
  41. Miranda-Sosa, A.; Del Moral, S.; Infanzón-Rodriguez, M.I.; Aguilar-Uscanga, M.G. Reappraisal of Different Agro-Industrial Waste for the Optimization of Cellulase Production from Aspergillus niger ITV02 in a Liquid Medium Using a Box–Benkhen Design. 3 Biotech 2024, 14, 278. [Google Scholar] [CrossRef] [PubMed]
  42. Dixit, M.; Shukla, P. Multi-Efficient Endoglucanase from Aspergillus niger MPS25 and Its Potential Applications in Saccharification of Wheat Straw and Waste Paper Deinking. Chemosphere 2023, 313, 137298. [Google Scholar] [CrossRef] [PubMed]
  43. Adsul, M.G.; Dixit, P.; Saini, J.K.; Gupta, R.P.; Ramakumar, S.S.V.; Mathur, A.S. Morphologically Favorable Mutant of Trichoderma reesei for Low Viscosity Cellulase Production. Biotechnol. Bioeng. 2022, 119, 2167–2181. [Google Scholar] [CrossRef] [PubMed]
  44. Peng, Z.-Q.; Li, C.; Lin, Y.; Wu, S.-S.; Gan, L.-H.; Liu, J.; Yang, S.-L.; Zeng, X.-H.; Lin, L. Cellulase Production and Efficient Saccharification of Biomass by a New Mutant Trichoderma afroharzianum MEA-12. Biotechnol. Biofuels 2021, 14, 219. [Google Scholar] [CrossRef]
  45. Hidayati, F.L.N.; Cahyanto, M.N. Enhancement of Indigenous Fungal Cellulase Production by Gamma Rays. In IOP Conference Series: Materials Science and Engineering; IOP Publishing: Bristol, UK, 2021; Volume 1192, p. 012004. [Google Scholar]
  46. Türkoğlu, A.; Haliloğlu, K.; Tosun, M.; Bujak, H.; Eren, B.; Demirel, F.; Szulc, P.; Karagöz, H.; Selwet, M.; Özkan, G.; et al. Ethyl Methanesulfonate (EMS) Mutagen Toxicity-Induced DNA Damage, Cytosine Methylation Alteration, and iPBS-Retrotransposon Polymorphisms in Wheat (Triticum aestivum L.). Agronomy 2023, 13, 1767. [Google Scholar] [CrossRef]
  47. Sega, G.A. A Review of the Genetic Effects of Ethyl Methanesulfonate. Mutat. Res. Genet. Toxicol. 1984, 134, 113–142. [Google Scholar] [CrossRef]
  48. Xu, L.; Wang, W.; Wu, J.; Shin, J.H.; Wang, P.; Unarta, I.C.; Chong, J.; Wang, Y.; Wang, D. Mechanism of DNA Alkylation-Induced Transcriptional Stalling, Lesion Bypass, and Mutagenesis. Proc. Natl. Acad. Sci. USA 2017, 114, E7082–E7091. [Google Scholar] [CrossRef]
  49. Sangkharak, K.; Vangsirikul, P.; Janthachat, S. Strain Improvement and Optimization for Enhanced Production of Cellulase in Cellulomonas Sp. TSU-03. Afr. J. Microbiol. Res. 2012, 6, 1079–1084. [Google Scholar] [CrossRef]
  50. El-Khamisi, E.F.; Soliman, E.A.M.; El-Sayed, G.M.; Nour, S.A.; Abdel-Monem, M.O.; Hassan, M.G. Optimization, Gene Cloning, Expression, and Molecular Docking Insights for Enhanced Cellulase Enzyme Production by Bacillus amyloliquefaciens Strain Elh1. Microb. Cell Factories 2024, 23, 191. [Google Scholar] [CrossRef]
  51. Deep, S.; Sharma, P.; Behera, N. Optimization of Extracellular Cellulase Enzyme Production from Alternaria brassicicola. Int. J. Curr. Microbiol. App. Sci. 2014, 3, 127. [Google Scholar]
  52. Cherkupally, R.; Amballa, H.; Bhoomi, N.R. In Vitro Screening for Enzymatic Activity of Trichoderma Species for Biocontrol Potential. Ann. Plant Sci. 2017, 6, 1784–1789. [Google Scholar] [CrossRef]
  53. Haq, I.; Ali, S.; Saleem, A.; Javed, M. Mutagenesis of Bacillus licheniformis through Ethyl Methanesulfonate for Alpha Amylase Production. Pak. J. Bot. 2009, 41, 1489–1498. [Google Scholar]
  54. Infanzón-Rodríguez, M.I.; Ragazzo-Sánchez, J.A.; Del Moral, S.; Calderón-Santoyo, M.; Aguilar-Uscanga, M.G. Enzymatic Hydrolysis of Lignocellulosic Biomass Using Native Cellulase Produced by Aspergillus niger ITV02 under Liquid State Fermentation. Biotechnol. Appl. Biochem. 2022, 69, 198–208. [Google Scholar] [CrossRef] [PubMed]
  55. Mandels, M.; Weber, J. The Production of Cellulases. In Cellulases and Their Applications; Advances in Chemistry; American Chemical Society: Washington, DC, USA, 1969; Volume 95, pp. 391–414. [Google Scholar] [CrossRef]
  56. Reczey, K.; Szengyel, Z.; Eklund, R.; Zacchi, G. Cellulase Production by T. reesei. Bioresour. Technol. 1996, 57, 25–30. [Google Scholar] [CrossRef]
  57. Liu, J.; Wang, Y.; Lu, S.; Liu, X.; Zhang, H.; Wang, W. Neg1 Overexpression in Trichoderma harzianum T4 Enhanced Its Ability to Produce Spores and Antagonistic Activity against Phytopathogenic Fungi. Arch. Microbiol. 2024, 206, 365. [Google Scholar] [CrossRef]
  58. Kamande, S.M.; Omwenga, G.I.; Ngugi, M.P. Production of Cellulases by Xylaria Sp. and Nemania Sp. Using Lignocellulose Substrates for Bioethanol Production from Maize Cobs. Heliyon 2024, 10, e36802. [Google Scholar] [CrossRef]
  59. Kadoguchi, E.A.; Velasco, J.; Silva, S.S.D.; Ingle, A.P.; Segato, F.; Chandel, A.K. Optimization of Cellulase Production from Agri-Industrial Residues by Aspergillus terreus NIH2624. Processes 2024, 12, 2169. [Google Scholar] [CrossRef]
  60. Saroj, P.; P, M.; Narasimhulu, K. Biochemical Characterization of Thermostable Carboxymethyl Cellulase and β-Glucosidase from Aspergillus fumigatus JCM 10253. Appl. Biochem. Biotechnol. 2022, 194, 2503–2527. [Google Scholar] [CrossRef]
  61. Gad, S.; Ayakar, S.; Adivarekar, R. Formulation and Characterization of Bacterial Consortium for Efficient Lignocellulosic Waste Degradation. J. Environ. Chem. Eng. 2024, 12, 112619. [Google Scholar] [CrossRef]
  62. Kumar, A.K.; Parikh, B.S.; Singh, S.P.; Shah, D. Use of Combined UV and Chemical Mutagenesis Treatment of Aspergillus terreus D34 for Hyper-Production of Cellulose-Degrading Enzymes and Enzymatic Hydrolysis of Mild-Alkali Pretreated Rice Straw. Bioresour. Bioprocess. 2015, 2, 35. [Google Scholar] [CrossRef]
  63. Bhardwaj, S.; Mishra, A.; Ghosh, D.; Sinha, A.K. Magnetically Separable Spent Coffee Grounds as a Potential Novel Support for the Covalent Immobilization of β-glucosidase for Cellobiose Hydrolysis. J. Chem. Technol. Biotechnol. 2024, 99, 1984–1995. [Google Scholar] [CrossRef]
  64. Msango, K.; Gouda, M.N.R.; Ramakrishnan, B.; Kumar, A.; Subramanian, S. Variation and Functional Profile of Gut Bacteria in the Scarab Beetle, Anomala dimidiata, under a Cellulose-Enriched Microenvironment. Sci. Rep. 2024, 14, 22400. [Google Scholar] [CrossRef] [PubMed]
  65. Ghose, T.K. Measurement of Cellulase Activities. Pure Appl. Chem. 1987, 59, 257–268. [Google Scholar] [CrossRef]
  66. Sun, J.; Wang, W.; Hao, J. A Novel GH1 β-Glucosidase from an Arctic Bacterium: Characterization and Secretory Expression in Bacillus subtilis. Process Biochem. 2024, 140, 108–116. [Google Scholar] [CrossRef]
  67. Prakash, S.; Kumari, H.; Pasrija, R.; Kumar, A. Unraveling the Intricacies of Carbon Catabolic Repression and Glucose in Fungal Pathogenesis. Hum. Fungal Dis. 2025, 1, 169–182. [Google Scholar] [CrossRef]
  68. Ten Kate, G.A.; Sanders, P.; Dijkhuizen, L.; Van Leeuwen, S.S. Kinetics and Products of Thermotoga maritima β-Glucosidase with Lactose and Cellobiose. Appl. Microbiol. Biotechnol. 2024, 108, 349. [Google Scholar] [CrossRef]
  69. Gawade, D.B.; Perane, R.R.; Deokar, C.D.; Raghuwanshi, K.S.; Suryawanshi, A.P. Effect of Physiological Factors on Production of Cellulolytic Enzymes by Rhizoctonia bataticola. Indian Phytopathol. 2018, 71, 555–561. [Google Scholar] [CrossRef]
  70. Sulyman, A.O.; Igunnu, A.; Malomo, S.O. Isolation, Purification and Characterization of Cellulase Produced by Aspergillus niger Cultured on Arachis Hypogaea Shells. Heliyon 2020, 6, e05668. [Google Scholar] [CrossRef]
  71. Sarsan, S.; Merugu, R. Role of Bioprocess Parameters to Improve Cellulase Production: Part II. In New and Future Developments in Microbial Biotechnology and Bioengineering; Elsevier: Amsterdam, The Netherlands, 2019; pp. 77–97. [Google Scholar] [CrossRef]
  72. Gaur, R.; Tiwari, S. Isolation, Production, Purification and Characterization of an Organic-Solvent-Thermostable Alkalophilic Cellulase from Bacillus vallismortis RG-07. BMC Biotechnol. 2015, 15, 19. [Google Scholar] [CrossRef]
  73. Ahmed, K.; Munawar, S.; Naz, S. Purification and Characterization of cellulase from Aspergillus. Chem. Bio. Environ. Sci. 2015, 27, 4341–4344. Available online: https://api.semanticscholar.org/CorpusID:212459652 (accessed on 12 October 2024).
  74. Jin, X.; Song, J.; Ma, J.; Liu, G.-Q. Thermostable β-Xylosidase from Aspergillus fumigatus: Purification, Characterization and Potential Application in Lignocellulose Bioethanol Production. Renew. Energy 2020, 155, 1425–1431. [Google Scholar] [CrossRef]
  75. Taha, A.S.J.; Taha, A.J.; Faisal, Z.G. Purification and Kinetic Study on Cellulase Produced by Local Trichoderma viride. Nat. Sci. 2015, 13, 87–90. [Google Scholar]
  76. Regmi, S.; Choi, Y.S.; Kim, Y.K.; Khan, M.M.; Lee, S.H.; Cho, S.S.; Jin, Y.-Y.; Lee, D.Y.; Yoo, J.C.; Suh, J.-W. Endoglucanase Produced by Bacillus subtilis Strain CBS31: Biochemical Characterization, Thermodynamic Study, Enzymatic Hydrolysis, and Bio-Industrial Applications. Biotechnol. Bioprocess Eng. 2020, 25, 104–116. [Google Scholar] [CrossRef]
  77. Ribeiro, O.; Magalhães, F.; Aguiar, T.Q.; Wiebe, M.G.; Penttilä, M.; Domingues, L. Random and Direct Mutagenesis to Enhance Protein Secretion in Ashbya gossypii. Bioengineered 2013, 4, 322–331. [Google Scholar] [CrossRef]
  78. Chen, L.; Duan, L.; Sun, M.; Yang, Z.; Li, H.; Hu, K.; Yang, H.; Liu, L. Current Trends and Insights on EMS Mutagenesis Application to Studies on Plant Abiotic Stress Tolerance and Development. Front. Plant Sci. 2023, 13, 1052569. [Google Scholar] [CrossRef]
  79. Fonseca, R.; Capel, C.; Nieto-Canseco, R.; Ortiz-Atienza, A.; Bretones, S.; López-Fábregas, J.D.; Quevedo-Colmena, A.S.; Lebrón, R.; Barragán-Lozano, T.; Villalobos-Ramírez, V.; et al. A Tomato EMS-Mutagenized Population Provides New Valuable Resources for Gene Discovery and Breeding of Developmental Traits. Plants 2022, 11, 2453. [Google Scholar] [CrossRef]
  80. Wang, H.; Zhai, L.; Geng, A. Enhanced Cellulase and Reducing Sugar Production by a New Mutant Strain Trichoderma harzianum EUA20. J. Biosci. Bioeng. 2020, 129, 242–249. [Google Scholar] [CrossRef]
  81. Sharma, A.K.; Bhati, N. Enhancement of Lipase Production by Ethyl Methane Sulfonate Mutagenesis of Soil Fungal Isolate. Plant Sci. Today 2019, 6, 600–606. [Google Scholar]
  82. Aljammas, H.A.; Yazji, S.; Azizieh, A. Enhancement of Protease Production from Rhizomucor miehei by Mutagenesis with Ethyl Methanesulfonate, Ultraviolet, and Microwaves—A Preliminary Study. Bioresour. Technol. Rep. 2022, 20, 101287. [Google Scholar] [CrossRef]
  83. Elakkiya, P.; Muralikrishnan, V. Cellulase Production and Purification of Mutant Strain Trichoderma viride. Int. J. Curr. Microbiol. App. Sci. 2014, 3, 720–727. [Google Scholar]
  84. Pramanik, S.K.; Mahmud, S.; Paul, G.K.; Jabin, T.; Naher, K.; Uddin, M.S.; Zaman, S.; Saleh, M.A. Fermentation Optimization of Cellulase Production from Sugarcane Bagasse by Bacillus pseudomycoides and Molecular Modeling Study of Cellulase. Curr. Res. Microb. Sci. 2021, 2, 100013. [Google Scholar] [CrossRef] [PubMed]
  85. Nehad, E.A.; Yoness, M.F.; Reem, A.A. Optimization and Purification of Cellulase Produced by Penicillium decumbens and Its Application. Egypt. Pharm. J. 2019, 18, 391–402. [Google Scholar]
  86. Acharya, P.B.; Acharya, D.K.; Modi, H.A. Optimization for Cellulase Production by Aspergillus niger Using Saw Dust as Substrate. Afr. J. Biotechnol. 2008, 7, 4147–4152. [Google Scholar]
  87. Latifian, M.; Hamidi-Esfahani, Z.; Barzegar, M. Evaluation of Culture Conditions for Cellulase Production by Two Trichoderma reesei Mutants under Solid-State Fermentation Conditions. Bioresour. Technol. 2007, 98, 3634–3637. [Google Scholar] [CrossRef]
  88. Abd Elrsoul, R.; Bakhiet, S.E.A. Optimization of Factors Influencing Cellulase Production by Some Indigenous Isolated Fungal Species. Jordan J. Biol. Sci. 2018, 11, 31–36. [Google Scholar]
  89. Dutt, D.; Kumar, A. Optimization of Cellulase Production under Solid-State Fermentation by Aspergillus flavus (AT-2) and Aspergillus niger (AT-3) and Its Impact on Stickies and Ink Particle Size of Sorted Office Paper. Cellul. Chem. Technol. 2014, 48, 285–298. [Google Scholar]
  90. Ng, I.-S.; Chen, P.T.; Ju, Y.-M.; Tsai, S.-W. Novel Cellulase Screening and Optimal Production from the Wood Decaying Xylariaceae: Daldinia Species. Appl. Biochem. Biotechnol. 2024, 196, 1–11. [Google Scholar] [CrossRef]
  91. Sethi, S.; Datta, A.; Gupta, B.L.; Gupta, S. Optimization of Cellulase Production from Bacteria Isolated from Soil. ISRN Biotechnol. 2013, 2013, 985685. [Google Scholar] [CrossRef]
  92. Kaur, H.P.; Joshi, D. Optimization of Cellulase Produced by Fungus Isolated from Water. World J. Pharm. Sci. 2015, 4, 521–534. [Google Scholar]
  93. Gautam, S.P.; Bundela, P.S.; Pandey, A.K.; Awasthi, M.K.; Sarsaiya, S. Optimization of the Medium for the Production of Cellulase by the Trichoderma viride Using Submerged Fermentation. Int. J. Environ. Sci. 2010, 1, 656–665. [Google Scholar]
  94. Vijayaraghavan, P.; Vincent, S.P.; Dhillon, G.S. Solid-Substrate Bioprocessing of Cow Dung for the Production of Carboxymethyl Cellulase by Bacillus halodurans IND18. Waste Manag. 2016, 48, 513–520. [Google Scholar] [CrossRef] [PubMed]
  95. Saleem, A.; El-Said, A.H.M.; Maghraby, T.A.; Hussein, M.A. Pathogenicity and Pectinase Activity of Some Facultative Mycoparasites Isolated from Vicia Faba Diseased Leaves in Relation to Photosynthetic Pigments of Plant. J. Plant Pathol. Microbiol. 2012, 3, 141. [Google Scholar]
  96. Ghori, M.I.; Ahmed, S.; Malana, M.A.; Jamil, A. Corn stover-enhanced cellulase production by Aspergillus niger NRRL 567. Afr. J. Biotechnol 2011, 10, 5878–5886. [Google Scholar]
  97. Dar, M.A.; Pawar, K.D.; Rajput, B.P.; Rahi, P.; Pandit, R.S. Purification of a Cellulase from Cellulolytic Gut Bacterium, Bacillus tequilensis G9 and Its Evaluation for Valorization of Agro-Wastes into Added Value Byproducts. Biocatal. Agric. Biotechnol. 2019, 20, 101219. [Google Scholar] [CrossRef]
  98. Elsababty, Z.E.; Abdel-Aziz, S.H.; Ibrahim, A.M.; Guirgis, A.A.; Dawwam, G.E. Purification, Biochemical Characterization, and Molecular Cloning of Cellulase from Bacillus licheniformis Strain Z9 Isolated from Soil. J. Genet. Eng. Biotechnol. 2022, 20, 34. [Google Scholar] [CrossRef]
  99. Kim, B.-K.; Lee, B.-H.; Lee, Y.-J.; Jin, I.-H.; Chung, C.-H.; Lee, J.-W. Purification and Characterization of Carboxymethylcellulase Isolated from a Marine Bacterium, Bacillus subtilis Subsp. Subtilis A-53. Enzym. Microb. Technol. 2009, 44, 411–416. [Google Scholar] [CrossRef]
  100. Asha, B.M. Purification and Characterization of a Thermophilic Cellulase from a Novel Cellulolytic Strain, Paenibacillus barcinonensis. J. Microbiol. Biotechnol. 2012, 22, 1501–1509. [Google Scholar] [CrossRef]
  101. Banerjee, S.; Maiti, T.K.; Roy, R.N. Production, Purification, and Characterization of Cellulase from Acinetobacter junii GAC 16.2, a Novel Cellulolytic Gut Isolate of Gryllotalpa Africana, and Its Effects on Cotton Fiber and Sawdust. Ann. Microbiol. 2020, 70, 28. [Google Scholar] [CrossRef]
  102. Steiner, E.; Margesin, R. Production and Partial Characterization of a Crude Cold-Active Cellulase (CMCase) from Bacillus mycoides AR20-61 Isolated from an Alpine Forest Site. Ann. Microbiol. 2020, 70, 67. [Google Scholar] [CrossRef]
  103. Mohapatra, S.; Padhy, S.; Mohapatra, P.K.D.; Thatoi, H.N. Enhanced Reducing Sugar Production by Saccharification of Lignocellulosic Biomass, Pennisetum Species through Cellulase from a Newly Isolated Aspergillus fumigatus. Bioresour. Technol. 2018, 253, 262–272. [Google Scholar] [CrossRef]
  104. Das, S.; Bhattacharya, A.; Haldar, S.; Ganguly, A.; Gu, S.; Ting, Y.P.; Chatterjee, P.K. Optimization of Enzymatic Saccharification of Water Hyacinth Biomass for Bio-Ethanol: Comparison between Artificial Neural Network and Response Surface Methodology. Sustain. Mater. Technol. 2015, 3, 17–28. [Google Scholar] [CrossRef]
  105. Okonkwo, I.F. Effect of Metal Ions and Enzyme Inhibitor on the Activity of Cellulase Enzyme of Aspergillus flavus. Int. J. Environ. Agric. Biotechnol. 2019, 4, 727–734. [Google Scholar] [CrossRef]
  106. Pino, M.S.; Michelin, M.; Rodríguez-Jasso, R.M.; Oliva-Taravilla, A.; Teixeira, J.A.; Ruiz, H.A. Hot Compressed Water Pretreatment and Surfactant Effect on Enzymatic Hydrolysis Using Agave Bagasse. Energies 2021, 14, 4746. [Google Scholar] [CrossRef]
  107. Singh, B. Purification and Characterization of a Protease-Resistant Phytase of Aspergillus oryzae SBS50 Whose Properties Make It Exceptionally Useful as a Feed Supplement. Int. J. Biol. Macromol. 2017, 103, 458–466. [Google Scholar] [CrossRef]
  108. Abhishek Walia, A.W.; Preeti Mehta, P.M.; Anjali Chauhan, A.C.; Saurabh Kulshrestha, S.K.; Shirkot, C.K. Purification and Characterization of Cellulase-Free Low Molecular Weight Endo β-1, 4 Xylanase from an Alkalophilic Cellulosimicrobium cellulans CKMX1 Isolated from Mushroom Compost. World J. Microbiol. Biotechnol. 2014, 30, 2597–2608. [Google Scholar] [CrossRef]
Fermentation 11 00274 g001
Figure 1. (A). Primary cellulolytic screening and (B) effect of increasing EMS concentrations on fungal survival rate and quantitative cellulase screening for wild-type and mutant strains (M1: 0.05%, M2: 0.1%, M3: 0.3%, M4: 0.5% and M5: 0.7%) of A. citri.
Figure 1. (A). Primary cellulolytic screening and (B) effect of increasing EMS concentrations on fungal survival rate and quantitative cellulase screening for wild-type and mutant strains (M1: 0.05%, M2: 0.1%, M3: 0.3%, M4: 0.5% and M5: 0.7%) of A. citri.
Fermentation 11 00274 g001
Fermentation 11 00274 g002
Figure 2. Optimization of temperature for cellulase production by A. citri and A. citri 305.
Figure 2. Optimization of temperature for cellulase production by A. citri and A. citri 305.
Fermentation 11 00274 g002
Fermentation 11 00274 g003
Figure 3. Optimization of pH for cellulase production by A. citri and A. citri 305.
Figure 3. Optimization of pH for cellulase production by A. citri and A. citri 305.
Fermentation 11 00274 g003
Fermentation 11 00274 g004
Figure 4. Optimization of incubation period for cellulase production by A. citri and A. citri 305.
Figure 4. Optimization of incubation period for cellulase production by A. citri and A. citri 305.
Fermentation 11 00274 g004
Fermentation 11 00274 g005
Figure 5. (A) Effect of substrates (SB: sugarcane bagasse; WS: wheat straw; OP: orange peel; PT: Pine trees; GA: Gum Arabic trees; CMC: Carboxymethyl cellulose) on cellulase production and (B) effect of substrate (orange-peel powder) concentration on cellulase production.
Figure 5. (A) Effect of substrates (SB: sugarcane bagasse; WS: wheat straw; OP: orange peel; PT: Pine trees; GA: Gum Arabic trees; CMC: Carboxymethyl cellulose) on cellulase production and (B) effect of substrate (orange-peel powder) concentration on cellulase production.
Fermentation 11 00274 g005
Fermentation 11 00274 g006
Figure 6. (A) Effect of carbon sources on cellulase production and (B) glucose concentration optimization for cellulase production.
Figure 6. (A) Effect of carbon sources on cellulase production and (B) glucose concentration optimization for cellulase production.
Fermentation 11 00274 g006
Fermentation 11 00274 g007
Figure 7. Optimization of nitrogen sources for cellulase production.
Figure 7. Optimization of nitrogen sources for cellulase production.
Fermentation 11 00274 g007
Fermentation 11 00274 g008
Figure 8. Effect of temperature on enzymatic activity profile of cellulases from wild A. citri and mutant A. citri 305 at pH 6 using 1% Carboxymethyl cellulose (endoglucanase), filter paper (exoglucanase), and cellobiose (β-glucosidase).
Figure 8. Effect of temperature on enzymatic activity profile of cellulases from wild A. citri and mutant A. citri 305 at pH 6 using 1% Carboxymethyl cellulose (endoglucanase), filter paper (exoglucanase), and cellobiose (β-glucosidase).
Fermentation 11 00274 g008
Fermentation 11 00274 g009
Figure 9. Effect of pH on enzymatic activity profile of cellulases from wild A. citri and mutant A. citri 305 at 50 ℃ using 1% Carboxymethyl cellulose (endoglucanase), filter paper (exoglucanase), and cellobiose (β-glucosidase).
Figure 9. Effect of pH on enzymatic activity profile of cellulases from wild A. citri and mutant A. citri 305 at 50 ℃ using 1% Carboxymethyl cellulose (endoglucanase), filter paper (exoglucanase), and cellobiose (β-glucosidase).
Fermentation 11 00274 g009
Fermentation 11 00274 g010
Figure 10. Thermo-stability assays of cellulases at temperatures of (A) 40 ℃, (B) 50 ℃, and (C) 60 ℃ of cellulases using 1% Carboxymethyl cellulose (endoglucanase), filter paper (exoglucanase), and cellobiose (β-glucosidase) at a temperature of 50 ℃ and pH 6.
Figure 10. Thermo-stability assays of cellulases at temperatures of (A) 40 ℃, (B) 50 ℃, and (C) 60 ℃ of cellulases using 1% Carboxymethyl cellulose (endoglucanase), filter paper (exoglucanase), and cellobiose (β-glucosidase) at a temperature of 50 ℃ and pH 6.
Fermentation 11 00274 g010
Fermentation 11 00274 g011
Figure 11. pH stability assays of cellulases produced by wild A. citri and mutant A. citri 305 at different pH, (A) pH 5, (B) pH 6, and (C) pH 7, using 1% Carboxymethyl cellulose (endoglucanase), filter paper (exoglucanase), and cellobiose (β-glucosidase) at a temperature of 50 ℃ and pH 6.
Figure 11. pH stability assays of cellulases produced by wild A. citri and mutant A. citri 305 at different pH, (A) pH 5, (B) pH 6, and (C) pH 7, using 1% Carboxymethyl cellulose (endoglucanase), filter paper (exoglucanase), and cellobiose (β-glucosidase) at a temperature of 50 ℃ and pH 6.
Fermentation 11 00274 g011
Table 1. A summary of the purification of cellulases from A. citri and A. citri 305.
Table 1. A summary of the purification of cellulases from A. citri and A. citri 305.
CellulasesParametersCulture SupernatantPartially Purified Enzyme
A. citriA. citri 305A. citriA. citri 305
TP (mg/mL)1.07 ± 0.021.55 ± 0.010.35 ± 0.010.17 ± 0.02
EndoglucanaseTA (U)0.43 ± 0.030.65 ± 0.040.27 ± 0.010.28 ± 0.01
SA (U/mg)0.40 ± 0.030.42 ± 0.0260.85 ± 0.031.65 ± 0.18
PF112.123.95
R (%)10010062.7943.07
ExoglucanaseTA (U)0.26 ± 0.020.27 ± 0.050.09 ± 0.0030.10 ± 0.01
SA (U/mg)0.24 ± 0.020.18 ± 0.0340.28 ± 0.020.58 ± 0.06
PF111.173.34
R (%)10010034.6137.03
β-glucosidase TA (U)8.59 ± 1.4412.85 ± 0.913.88 ± 0.311.49 ± 0.26
SA (U/mg)8.02 ± 1.358.29 ± 0.58812.26 ± 1.058.76 ± 1.79
PF111.521.05
R (%)10010045.1611.59
(TP: total protein, TA: total activity, SA: specific activity, PF: purification fold, and R: recovery).
Table 2. Effects of metal ions and additives on cellulase activity of wild A. citri and mutant A. citri 305.
Table 2. Effects of metal ions and additives on cellulase activity of wild A. citri and mutant A. citri 305.
Metal Ions
(10%)		Relative ActivityRelative ActivityRelative Activity
(%)(%)(%)
β-glucosidaseEndoglucanaseExoglucanase
A. citriA. citri 305A. citriA. citri 305A. citriA. citri 305
Control100 ± 0100 ± 0100 ± 0100 ± 0100 ± 0100 ± 0
NaNO3104 ± 1103 ± 196 ± 196 ± 1126 ± 1104 ± 3
FeSO4127 ± 1139 ± 3203 ± 1137 ± 1275 ± 1285 ± 2
MgSO4107 ± 1135 ± 2119 ± 2113 ± 1135 ± 2205 ± 2
ZnSO4120 ± 1108 ± 2131 ± 2112 ± 1156 ± 0195 ± 2
KH2PO479 ± 390 ± 496 ± 194 ± 0115 ± 1118 ± 0
CoCl253 ± 173 ± 370 ± 173 ± 195 ± 178 ± 3
Additives
(5 mM)		β-glucosidaseEndoglucanaseExoglucanase
(%)(%)(%)
A. citriA. citri 305A. citriA. citri 305A. citriA. citri 305
Control100 ± 0100 ± 0100 ± 0100 ± 0100 ± 0100 ± 0
SDS56 ± 296 ± 163 ± 160 ± 141 ± 254 ± 1
Tween 80107 ± 2108 ± 0118 ± 2112 ± 1126 ± 2165 ± 1
EDTA67 ± 397 ± 177 ± 183 ± 178 ± 290 ± 1
Urea68 ± 359 ± 296 ± 375 ± 086 ± 264 ± 4
Table 3. Kinetics studies of cellulases produced by wild A. citri and mutant A. citri 305.
Table 3. Kinetics studies of cellulases produced by wild A. citri and mutant A. citri 305.
CellulasesStrainsKmVmaxEfficiency
(mg/mL)(mg/mL/min)(/min)
EndoglucanaseA. citri0.492 ± 0.0040.039 ± 0.0000.078 ± 0.001
A. citri 3050.295 ± 0.0060.060 ± 0.0000.202 ± 0.004
ExoglucanaseA. citri0.278 ± 0.0110.023 ± 0.0000.084 ± 0.003
A. citri 3050.241 ± 0.0070.023 ± 0.0000.094 ± 0.004
β-glucosidaseA. citri0.048 ± 0.0010.754 ± 0.01115.683 ± 0.099
A. citri 3050.031 ± 0.0011.128 ± 0.00436.765 ± 1.598
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Ahmed, S.; Andaleeb, H.; Aslam, A.; Raza, J.A.; Waseem, S.M.Y.; Javaid, A.; Talib, C. Improved Cellulolytic Activity of Alternaria citri: Optimization and EMS Treatment for Enhanced Cellulase Production. Fermentation 2025, 11, 274. https://doi.org/10.3390/fermentation11050274

AMA Style

Ahmed S, Andaleeb H, Aslam A, Raza JA, Waseem SMY, Javaid A, Talib C. Improved Cellulolytic Activity of Alternaria citri: Optimization and EMS Treatment for Enhanced Cellulase Production. Fermentation. 2025; 11(5):274. https://doi.org/10.3390/fermentation11050274

Chicago/Turabian Style

Ahmed, Sibtain, Hina Andaleeb, Aqsa Aslam, Junaid Ahmad Raza, Sheikh Muhammad Yahya Waseem, Atayyaba Javaid, and Chand Talib. 2025. "Improved Cellulolytic Activity of Alternaria citri: Optimization and EMS Treatment for Enhanced Cellulase Production" Fermentation 11, no. 5: 274. https://doi.org/10.3390/fermentation11050274

APA Style

Ahmed, S., Andaleeb, H., Aslam, A., Raza, J. A., Waseem, S. M. Y., Javaid, A., & Talib, C. (2025). Improved Cellulolytic Activity of Alternaria citri: Optimization and EMS Treatment for Enhanced Cellulase Production. Fermentation, 11(5), 274. https://doi.org/10.3390/fermentation11050274

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop