Role of crude enzymes and microbial consortia in the degradation of organic matter

*Corresponding author: Himanshu Pundir Copyright © 2025 Author(s) retain the copyright of this article. This article is published under the terms of the Creative Commons Attribution License 4.0. Role of crude enzymes and microbial consortia in the degradation of organic matter Himanshu Pundir *, Rahil Balu, Pratibha Shah, and Rajitha Satish Department of Microbiology, Kishinchand Chellaram College, Churchgate, Mumbai-400020. World Journal of Biology Pharmacy and Health Sciences, 2025, 24(01), 287-298 Publication history: Received on 11 September 2025; revised on 17 October 2025; accepted on 20 October 2025 Article DOI: https://doi.org/10.30574/wjbphs.2025.24.1.0914 Abstract The rapid expansion of agricultural and industrial sectors has resulted in the accumulation of lipidand polysacchariderich wastes, posing substantial environmental challenges. Microbial biodegradation employing crude enzymes presents a sustainable and eco-friendly approach to mitigating these pollutants. In this study, soil and sludge samples collected from sugarcane and mustard cultivation fields, along with associated processing units, were enriched in selective media to isolate potent enzyme-producing microorganisms. The isolates were screened for lipase, amylase, and cellulase activity using tributyrin agar, carboxymethyl cellulose (CMC) agar, and starch agar, respectively. Enzymatic activity was quantified through titrimetric and DNS assays, while MALDI-TOF analysis confirmed the taxonomic identities. Lipase purification was carried out using ammonium sulfate precipitation, followed by immobilization in calcium alginate beads to assess stability and application potential. A microbial consortium was subsequently developed to evaluate synergistic degradation of organic waste. Seven bacterial isolates belonging to Pseudomonas, Bacillus, Enterococcus, Micrococcus, and Providencia genera exhibited significant enzymatic activity. Lipase activity ranged between 476–588 U/ml, with purification enhancing it up to 1064 U/ml. Although immobilization slightly reduced activity, microbial consortia demonstrated enhanced biodegradation efficiency. Notably, consortia 1 and 4 achieved the highest waste reduction (~11 g over 45 days). These findings highlight the biotechnological potential of enzyme-producing microbes and their consortia in sustainable waste degradation, offering promising applications in industrial effluent treatment and agricultural residue management. Keywords: Lipase; Amylase; Cellulase; Enzyme immobilization; Biodegradation 1. Introduction The exponential increase in global population and industrialization has resulted in escalating environmental challenges, particularly due to the accumulation of organic waste from agricultural and industrial sources. Wastes rich in lipids and polysaccharides represent a major contributor to soil and water pollution, necessitating the development of innovative, sustainable, and biologically compatible waste management strategies. Agricultural systems, though essential for global food security, generate vast quantities of biodegradable residues, including damaged grains and plant matter, which, upon decomposition, release organic pollutants into the environment. Similarly, industrial operations such as food processing and petroleum refining produce large volumes of lipid-laden effluents that are difficult to manage using conventional treatment methods. Traditional approaches to treating lipid-containing wastewater, such as grease traps, incineration, and landfill disposal, are often ineffective, costly, and environmentally detrimental. In contrast, microbial biodegradation offers a sustainable World Journal of Biology Pharmacy and Health Sciences, 2025, 24(01), 287-298 288 and efficient alternative. Several microorganisms, including Pseudomonas aeruginosa, Bacillus spp., and various yeast strains, have demonstrated strong lipid-degrading capabilities. Notably, Bacillus subtilis BN1001 has been successfully utilized for treating lipid-rich industrial effluents. Polysaccharides such as cellulose and starch also constitute a major fraction of agricultural waste. Cellulases and amylases, enzymes secreted by bacteria and fungi, play pivotal roles in the hydrolysis of these polymers into simple sugars, enabling their conversion into renewable energy substrates and industrial feedstocks. Enzymatic degradation of damaged grains, for instance, can yield fermentable sugars, representing an eco-friendly approach to waste valorization. This study investigates the enzymatic potential of microorganisms isolated from sugarcane fields, mustard oil processing units, and related organic waste environments. These naturally adapted microbes were screened and characterized through cultural, biochemical, and molecular analyses for their ability to degrade lipidic and polysaccharide substrates. Furthermore, enzyme immobilization using calcium alginate beads was explored to enhance catalytic stability and facilitate reuse in continuous degradation systems. By elucidating the enzymatic profiles and synergistic activity of these isolates, this research contributes to the advancement of microbial biocatalysis for sustainable environmental remediation and organic waste management. 2. Materials and methods 2.1. Sample Collection Soil and sludge samples were selected as sources for isolating cellulase, amylase, and lipase-producing bacteria. Sampling was strategically performed at agricultural and industrial sites rich in organic matter: the sugarcane field at PJF9+P97, Sisoni, and the sugarcane factory at Isharpur, Noorpur Dehat, Uttar Pradesh, was targeted for the isolation of polysaccharide-degrading enzymes (amylase, cellulase). For lipid-degrading isolates, soil samples were obtained from the mustard field at PJF9+P97, Sisoni, and an oil production factory at PJC7+8HP, Sisoni. To enhance environmental diversity in the study, sludge was collected from a water source in Churchgate, Mumbai. All samples were aseptically gathered using sterile plastic bags and bottles. Immediate sealing and refrigerated transport preserved sample integrity for downstream microbiological analysis. 2.2. Enrichment of the Sample To facilitate the proliferation of target enzyme-producing microbial populations from diverse environmental matrices, enrichment was conducted using selective liquid media optimized for distinct enzyme activities. Specifically, for the isolation of cellulase producers, carboxymethyl cellulose (CMC) broth was employed. Amylase producers were targeted using starch broth, while lipase-producing organisms were enriched in both Tween 80 and Tributyrin broths. For each collected sample, precisely 1 gram of soil or 1 ml of sludge was introduced into 150 ml of the designated enrichment medium. The enrichment cultures were incubated at ambient temperature under continuous agitation at 150 rpm for a period extending up to 30 days, thereby maximizing the selective amplification of the desired bacterial communities. During this incubation, periodic aliquots were aseptically transferred onto corresponding selective agar plates, Tween 80 agar and Tributyrin agar for lipase producers, CMC agar for cellulase, and starch agar for amylase isolates. The manifestation of hydrolysis zones, visualized using appropriate reagents (such as Gram iodine for starch and CMC hydrolysis), served as an initial qualitative indicator of functional enzyme production and allowed for the preliminary selection of potent isolates. 2.3. Screening of Enzyme-Producing Organisms 2.3.1. Primary Screening A primary screen was performed on morphologically distinct colonies obtained from enriched cultures. Each prospective isolate was evaluated based on its colonial morphology, Gram staining characteristics, and functional enzyme activity using substrate hydrolysis tests (starch, cellulose, lipid). The generation of clear zones surrounding colonies on indicator medium signified hydrolytic activity, thus presumptively identifying enzyme producers. 2.3.2. Production and Extraction of the Crude Enzymes For quantitative assessment, selected isolates underwent further incubation via the submerged fermentation process: World Journal of Biology Pharmacy and Health Sciences, 2025, 24(01), 287-298 289 • Inoculum Preparation: Isolates were cultured in their respective inoculum preparation media (olive oil broth for lipase, CMC inoculum broth for cellulase, starch inoculum broth for amylase) under shaking conditions for three days to achieve substantial microbial mass. • Fermentation: Following the inoculum phase, 5 ml were transferred into 150 ml of sterile production media in 250 ml flasks and incubated for five additional days under shaking conditions. • Extraction (By cold centrifugation): The resulting broth was subjected to cold centrifugation (2000 rpm, 3 hours, 0–4 °C) in 15 ml conical tubes. The resulting clear, cell-free supernatant represented the crude enzyme extract, which was retained for subsequent activity assays and secondary screening of enzyme-producing organisms. 2.3.3. Secondary Screening: The organisms screened through primary screening underwent secondary screening with the help of respective assay methods for crude enzymes. Cellulase and Amylase enzyme Assay (By DNS method) [10]: For both cellulase and amylase, enzyme activity was measured spectrophotometrically via the dinitrosalicylic acid (DNS) reduction method. In each assay, 0.5 ml of crude enzyme extract was incubated with 0.5 ml of 1000 μg/ml substrate in 0.1 M sodium phosphate buffer (pH 7.0) at 37°C for 60 minutes. The reaction was terminated by the addition of 2 ml of 3,5-dinitrosalicylic acid reagent, and absorbance was measured at 550 nm. One unit (U) of enzyme was defined as the amount catalyzing the release of 1 μ/mol of reducing sugar per minute under assay conditions. Lipase assay (By Titrimetric method) [10]: For the lipase assay, 1 ml of crude enzyme extract was mixed with 1 ml of 0.1 M Tris-HCl buffer (pH 8.0), 2.5 ml of distilled water, and 1 ml of olive oil. The reaction mixture was homogenized and incubated at 37 °C for 30 minutes under controlled conditions, with parallel test and blank reactions prepared. After incubation, 3 ml of 95% ethanol was added to terminate the reaction. The liberated free fatty acids were quantified by titration against 0.1 M NaOH using phenolphthalein as an indicator, where the appearance of a stable pink colour denoted the endpoint, indicating neutralization of the fatty acids. 2.4. Identification: Final screening of enzyme-producing bacterial isolates was accomplished via Matrix-Assisted Laser Desorption Ionization-Time of Flight (MALDI-TOF) mass spectrometry. This powerful analytical technique, outsourced to an accredited laboratory, enables rapid and reliable taxonomic identification by generating unique mass spectral fingerprints of microbial proteins. Compared to conventional biochemical and molecular methods, MALDI-TOF delivers accurate genus and species-level classification, thereby confirming the identity of the isolates. Notably, strains belonging to Enterococcus, Pseudomonas, Micrococcus, Providencia, and Bacillus were successfully characterized using this method. 2.5. Lipase purification (By ammonium Sulphate precipitation) [14]: Cell-free supernatants exhibiting significant lipase activity were subjected to protein purification via ammonium sulfate precipitation, a classical "salting-out" technique for enzyme isolation. The process was based on the Ammonium Sulfate Fractionation Table to determine precise saturation concentrations. Typically, the procedure involved placing 100 ml of the cell-free enzyme solution over a magnetic stirrer in an ice bath for temperature control. Incremental addition of solid ammonium sulfate (to reach 60–70% saturation) facilitated gradual precipitation of proteins, including lipase, while maintaining low speeds and consistent mixing. The suspension was allowed to equilibrate overnight at 4°C. Centrifugation (2400 rpm, 3 hours, 0–4°C) separated the precipitated protein pellet, which was subsequently solubilized in minimal Tris-HCl buffer (pH 8.0) and dialyzed to remove residual salts, monitored using barium chloride and dilute HCl for complete desalting. The purified enzyme was then assayed for enhanced catalytic activity. 2.6. Enzyme Immobilization [3]: The lipase enzyme purified by ammonium sulfate precipitation was employed for immobilization using the calcium alginate method, leveraging gel matrix entrapment for enhanced stability and reusability. For each assay, 10 ml of 4% sodium alginate and 150 ml of 6% calcium chloride were autoclaved. The cooled solutions were mixed with 1 ml purified enzyme in sodium alginate, and beads were formed by dispensing the mixture into calcium chloride with continuous swirling. After overnight hardening at refrigeration temperatures, the beads were washed thoroughly with distilled water. Immobilized enzyme beads from three selected isolates were then packed into a sterile bioreactor column and World Journal of Biology Pharmacy and Health Sciences, 2025, 24(01), 287-298 290 tested for continuous lipase activity. Olive oil, emulsified with Tween 80, served as the substrate in Tris-HCl buffer (pH 8.0). Titrimetric assays were performed by collecting reaction aliquots at intervals to quantify fatty acid production, confirming the efficacy of immobilization for potential biotechnological applications. 2.7. Consortium Preparation [8][15]: To evaluate the efficiency of microbial consortia in degrading organic household waste, selected bacterial isolates were grouped based on known enzymatic profiles, comprising three lipase producers, two cellulase producers, and two amylase producers. Four unique consortia were formulated by strategic combinations to harness synergistic enzymatic activity. A mixture of commonly encountered household biowastes, potato peels, coriander leaves and roots, spinach leaves, spring onion, red spinach, and pea pods, was used as the substrate. Approximately 22 grams of autoclaved waste material were dispensed into pre-weighed sterile Petri plates. As a lipid source, 3 ml of olive oil was added to each experimental plate. Plates assigned as controls received sterile distilled water in place of the consortium inoculum. 2.7.1. Preparation and Activation of Consortium Inoculum Sterile saline suspensions of each isolate were prepared and adjusted to an optical density of 0.1 at 540 nm. One milliliter of this suspension was introduced into 10 ml of nutrient media and incubated at room temperature for 24 hours to activate growth. Subsequently, 1 ml of the incubated media was transferred to specific activation media and incubated again for 24 hours under shaker conditions (150 rpm). Finally, 3 ml of the prepared consortium mixture was dispensed onto each sterile waste-containing Petri plate. Control plates received an equivalent volume of sterile distilled water. 2.7.2. Experimental Setup and Incubation Procedure All experimental and control plates were meticulously weighed after the addition of inoculum and securely wrapped to prevent external contamination. Incubation was maintained at ambient temperature (25–28°C). The degradation process was monitored by recording the weight of each plate at 5-day intervals during the initial 15 days and finally on the 45th day post-inoculation. The difference in plate weight before and after incubation provided a quantitative measurement of organic matter degradation. Consortium efficacy was determined by calculating the percentage weight reduction, accounting for initial and final masses 3. Results 3.1. Screening Twenty-one distinct bacterial strains were isolated from various environmental sources based on colony morphology and Gram reaction. Of these, ten exhibited lipase activity, indicated by hydrolysis zones on tributyrin agar; eight with the largest clear zones were selected for further study. Screening for cellulase and amylase producers yielded five and six isolates, respectively, all showing clearance zones on indicator agar after Gram’s iodine flooding. In total, six cellulase and amylase producers with prominent activity were selected. Fourteen promising isolates from the primary screen underwent secondary screening and were confirmed as pure enzyme producers after further purification on Nutrient and MacConkey agar. For tracking, isolates were labeled using source-based initials with numerical subscripts for multiple isolates from the same origin, as detailed in the table below. Table 1 List of labelled isolates Lipase-producers Cellulase and amylase producers S₁, S₂, S₆, SF₁, SF₂, SF₃, LF₁, LM₃ S, SM₅, SF₂, SF₃, LM₁, SF₁B SFSugarcane Field, SMSugar refinery (Manufacturer), LFMustard (Lipid) Field, LMMustard oil (Lipid) refinery (Manufacturer) World Journal of Biology Pharmacy and Health Sciences, 2025, 24(01), 287-298 291 Figure 1 Labelled isolates from screening 3.2. Measurement of Enzyme Activity (Secondary Screening) In the secondary screening phase, fourteen isolates from the initial screening underwent submerged fermentation for enzyme production. Enzyme-specific assays evaluated crude enzyme extracts. 3.2.1. Lipase Assay: Enzyme activity was determined using a titrimetric method, in which sodium hydroxide (NaOH) was titrated against the liberated fatty acids. This allowed identification of isolates with the highest lipase potential. From the readings, 5 isolates were selected for further testing and applications. The lipase activity was assayed for all the selected isolates, and it was as follows: Enzyme Activity (U/ml) = (𝑁𝑎𝑂𝐻)(𝑀𝑜𝑙𝑎𝑟𝑖𝑡𝑦 𝑜𝑓 𝑁𝑎𝑂𝐻)(1000)(2)(𝑑𝑓) (1) Table 2 Lipase Enzyme Activity Sample Calculation Result S₁ 1.7 * 0.1 * 1000 * 2 * 1.4 / 1 476 U/ml S₂ 2.1 * 0.1 * 1000 * 2 * 1.4 / 1 588 U/ml S₆ 2 * 0.1 * 1000 * 2 * 1.4 / 1 560 U/ml World Journal of Biology Pharmacy and Health Sciences, 2025, 24(01), 287-298 292 LF₁ 1.7 * 0.1 * 1000 * 2 * 1.4 / 1 476 U/ml LF₃ 1.7 * 0.1 * 1000 * 2 * 1.4 / 1 476 U/ml The mean and the standard deviation obtained for the enzyme activity are 515.2 ± 54.58. 3.2.2. Cellulase Assay: Cellulase production was quantified using a spectrophotometric assay, with concurrent substrate blanks to correct for any turbidity in crude enzyme samples. This approach minimized error in absorbance readings. Based on these measurements, three isolates were selected for their high cellulase activity. Enzyme activity (U/ml) = (μmoles of glucose released)(1) (0.5) (30) μmoles of glucose released =Glucose concentration (μg/ml) Molecular weight Table 3 Cellulase Enzyme Activity Sample Calculation Result SF₂ 2.61 * 1 / 0.5 * 30 0.174 U/ml SM₅ 2.79 * 1 / 0.5 * 30 0.186 U/ml SF₃ 2.70 * 1 / 0.5 * 30 0.180 U/ml The mean and the standard deviation for the spectrophotometric readings are 0.27 ± 0.052 3.2.3. Amylase Assay: Amylase activity was determined by enzyme-specific assay, with substrate blanks included for all crude enzyme measurements to enhance the reliability. Of the six isolates tested, 3 showed significant amylase activity and were selected for further study. Amylase activity for all the selected isolates was calculated: Enzyme activity (U/ml) = (μmoles of Maltose released)(1) (0.5) (30) μmoles of Maltose released =Glucose concentration (μg/ml) Molecular weight Table 4 Amylase Enzyme Activity Sample Calculation Result SF₁B 1.08 * 1 / 0.5 * 30 0.072 U/ml SM₅ 0.93 * 1 / 0.5 * 30 0.062 U/ml SF₃ 1.52 * 1 / 0.5 * 30 0.101 U/ml The mean and the standard deviation for the spectrophotometric readings are 0.208 ± 0.044. The analysis of enzyme activity among the six tested isolates revealed notable overlap in both cellulase and amylase production. Isolates SM5 and SF3 exhibited high levels of activity for both enzymes. Additionally, SF2 was distinguished by significant cellulase production, while SF1B showed marked amylase activity. As a result, four isolates were selected for further study: two (SM5, SF3) for concurrent cellulase and amylase production, and one each (SF2, SF1B) for exclusive cellulase or amylase activity, respectively. The results demonstrated overlap in cellulase and amylase activity among the six isolates: World Journal of Biology Pharmacy and Health Sciences, 2025, 24(01), 287-298 293 • SM5 and SF3 showed strong activity for both enzymes. • SF2 was notable for cellulase production. • SF1B displayed high amylase activity. Consequently, two isolates were chosen for concurrent production, and two for singular enzyme production, totaling four for further study. 3.3. Identification: Isolates confirmed for the production of the desired enzymes were identified using morphological and cultural tests and the MALDI-TOF technique, up to genus level. The results were as follows: Table 5 List of organisms identified through MALDI-TOF Sample Taxonomic Identification S₂ Pseudomonas aeruginosa LF₁ Bacillus cereus group LM₃ Enterococcus gattinarum SM₅ Micrococcus luteus SF₃ Micrococcus luteus SF₂ Providencia rettgeri SF₁B Not identified 3.4. Lipase Purification: Purification aimed to isolate lipase from crude enzyme extracts and remove residual media and byproducts. Crude enzyme was produced via submerged fermentation (5 days), extracted by cold centrifugation, and initially assayed using a titrimetric method. Three isolates with notable lipase activity were selected, cultured again for 5 days, and subjected to ammonium sulfate precipitation to selectively "salt out" lipase protein. The resulting protein precipitate was resuspended and desalted to eliminate trace salts, enhancing enzymatic activity and purity for downstream analysis. Enzyme Activity (U/ml) = (𝑁𝑎𝑂𝐻)(𝑀𝑜𝑙𝑎𝑟𝑖𝑡𝑦 𝑜𝑓 𝑁𝑎𝑂𝐻)(1000)(2)(𝑑𝑓) (1) Table 6 Purified Lipase Enzyme Activity Sample Calculation Result LF₁ 3.8 * 0.1 * 1000 * 2 * 1.4 / 1 1,064 U/ml LM₃ 2.1 * 0.1 * 1000 * 2 * 1.4 / 1 588 U/ml S₂ 2.5 * 0.1 * 1000 * 2 * 1.4 / 1 700 U/ml The mean and standard deviation for the purified lipase enzyme activity are 784 ± 248.87. 3.5. Lipase Immobilization: The activity of the entrapped lipase enzyme was assayed using a bioreactor containing the beads in a solution of TrisHCl buffer containing olive oil as a lipid source and Tween80 as the emulsifier. The 1 ml reaction solution was eluted every 10 mins and assayed with the titrimetric method used before. The mean and the standard deviation for the titrimetric readings at 60 mins were observed to be 2.17 ± 0.15. The observations made were as follows: Table 7 Bioreactor Trimetric Assay World Journal of Biology Pharmacy and Health Sciences, 2025, 24(01), 287-298 294 Time (mins) LF₁ LF₃ S₆ 0 0 0 0 10 1.2 1.2 1.3 20 2.0 3.0 2.5 30 1.7 2.1 1.7 40 1.9 1.9 1.8 50 1.7 2.1 2 60 2.0 2.2 2.3 Figure 2 Bioreactor 3.6. Consortium Treatment Efficiency: To assess the consortium's effectiveness in treating biodegradable waste, the weight of the plates and the waste were initially measured separately. Subsequently, the waste was added to the plates, sterilized, and inoculated with the consortium. Weight measurements were taken at five-day intervals until the 15th day, after which readings were directly recorded on the 45th day. The following data were obtained from these measurements. Table 8 List of data consisting of the weight of the Petri plate taken for 45 days Consortium Organism Mixture (1 ml each) Day 1 (gm) Day 5 (gm) Day 10 (gm) Day 15 (gm) Day 45 (gm) Difference Day 1–45 (gm) 1 LF₁ + SF₁B + SF₂ 107.547 111.739 112.208 109.092 97.392 10.155 2 S₂ + SF₁B + SF₂ 105.143 109.688 109.065 108.345 101.286 3.857 3 LF₁ + SM₅ + SF₃ 118.191 122.466 121.808 121.176 111.387 6.804 4 S₂ + SF₃ + SM₅ 108.796 112.945 111.714 110.564 98.07 10.726 Control (5) Distilled water 109.06 113.259 110.435 111.086 99.834 9.226 World Journal of Biology Pharmacy and Health Sciences, 2025, 24(01), 287-298 295 Following the analysis of the data presented in the table, a graph was constructed, and several observations were derived. These observations aided in determining which consortium of organisms holds the potential for the degradation of organic matter, thus aiding future research endeavours and potential applications. Note: It was observed that the control plate exhibited fungal growth on the 45th day, potentially contributing to discrepancies in total plate weight and the rate of organic matter degradation. These deviations may have led to variations from the anticipated plate weight. Figure 3 Graph representing the variation in weight over 45 days Analysis of the graph revealed Consortium 1 as the most efficient in waste degradation, closely followed by Consortium 4. The initial 15-day weight increase across all plates suggested microbial culture proliferation. Consortia 1 and 4 notably reduced weight by nearly 11 grams, while Consortia 2 and 3 decreased by approximately 2 and 7 grams, respectively. The control plate unexpectedly reduced by almost 10 grams. Consortium 1 Consortium 1 Control (5) Control (5) Figure 4 Petri plates - Day 1 Vs Day 45 4. Discussion In The findings of this study corroborate a growing consensus in the scientific community regarding the central role of microbial consortia and specialized bacterial isolates in the bioremediation of organic wastes. The results demonstrate that purposefully selected and optimized microorganisms can substantially enhance the degradation of complex substrates such as fats, oils, grease, and lignocellulosic agricultural residues.