Agro-Industrial Wastewaters as Feedstocks for Microbial Lipase Production: Integrating Enzyme Bioprocessing, Effluent Valorization and Wastewater Management

Abstract

Agro-industrial wastewaters contain substantial organic and lipid loads that pose environmental challenges but may also provide potentially low-cost substrates for microbial bioprocessing. This scoping review evaluated the use of agro-industrial wastewaters as fermentation feedstocks for microbial lipase production and their potential integration with wastewater treatment and resource recovery. Previous reviews have largely treated microbial lipase production, waste-derived fermentation substrates, and lipase-based wastewater treatment as separate topics. This review bridges these areas by focusing on liquid agro-industrial wastewaters as fermentation feedstocks and jointly evaluating lipase production, changes in wastewater quality, process conditions, analytical comparability, and technological readiness. Following PRISMA-ScR guidance, Scopus and the Web of Science were searched for studies published from 2016 to 2026, yielding 20 eligible articles. The evidence was heavily concentrated on olive mill wastewater and palm oil mill effluent, whereas other agro-industrial effluents were scarcely investigated. Yeasts, bacteria, filamentous fungi, mixed cultures, and recombinant whole-cell systems were employed, although cultivation strategies, wastewater conditioning, supplementation, and lipase assays varied substantially. Several studies combined lipase production with reductions in chemical oxygen demand, oil and grease, phenolic compounds, or other wastewater constituents. However, most processes remained at the shake-flask scale, while downstream recovery, detailed enzyme characterization, storage stability, catalyst reuse, and controlled scale-up were infrequently investigated. Wastewater-based lipase production therefore represents a promising circular-bioeconomy strategy, but further development requires standardized performance reporting, minimal-input cultivation, integrated final-effluent assessment, scale-up, and formal techno-economic and life-cycle assessment.

1. Introduction

The growing volume of wastewater poses a major environmental, public health, and resource management challenge. Population growth, accelerated urbanization, economic development, and the expansion of food-processing activities contribute to rising wastewater volumes and pollution loads worldwide [1]. Food-processing effluents commonly contain organic matter, nutrients, suspended solids, lipids, and other process-specific contaminants [2]. However, the global status of industrial wastewater treatment remains difficult to assess because of incomplete reporting. According to the most recent assessment of Sustainable Development Goal indicator 6.3.1, data were available for only 22 countries, representing approximately 8% of the global population, and only 27% of the reported industrial wastewater was classified as safely treated [3]. At the same time, wastewater management is increasingly shifting from pollutant removal toward resource recovery, with wastewater recognized as a potential source of reusable water, energy, nutrients, and value-added bioproducts [4].

Agro-industrial activities generate wastewater with a high and often variable organic load. Olive and palm oil processing, dairy manufacturing, slaughtering, fish processing, and edible-oil refining produce effluents containing lipids, free fatty acids, proteins, carbohydrates, phenolic compounds, and suspended organic matter. Consequently, these wastewaters are often characterized by elevated chemical oxygen demand (COD), biochemical oxygen demand (BOD), and oil and grease concentrations. Their composition varies with the raw material, production technology, water consumption, cleaning procedures, season, and geographical location, complicating the design and operation of conventional treatment systems [5].

Lipid-rich effluents are particularly difficult to manage because fats, oils, and grease can accumulate in wastewater infrastructure, clog pipes, form excessive scum and foam, reduce air–liquid oxygen transfer, and interfere with biological treatment [6]. During anaerobic treatment, the accumulation of long-chain fatty acids may inhibit microbial activity, while phenolic compounds, particularly those in olive mill wastewater, may further impair biodegradation and process stability [7]. Such effluents may therefore require pretreatment or carefully controlled operational strategies, including dilution, nutrient supplementation, intermittent feeding, or physicochemical conditioning, thereby increasing overall process complexity. Nevertheless, the organic compounds responsible for the high pollution load may also serve as carbon sources, nutrients, or metabolic inducers for selected microorganisms. Agro-industrial wastewater may thus function not only as a waste stream but also as a feedstock for integrated bioprocesses that combine pollutant removal with the production of valuable microbial metabolites. For example, fed-batch cultivation of lipolytic yeasts in olive mill wastewater enabled simultaneous lipase production and reductions in COD and lipid content exceeding 50% [8].

Lipases, or triacylglycerol acylhydrolases (EC 3.1.1.3), catalyze the hydrolysis of triacylglycerols and, under suitable conditions, esterification, transesterification, and interesterification [9]. Their catalytic versatility has enabled their use in food processing, detergents, pharmaceuticals, cosmetics, biodiesel production, oleochemical transformations, wastewater treatment, and the synthesis of structured lipids [10]. Microbial lipases are particularly attractive for industrial production because bacteria, yeasts, and filamentous fungi can be cultivated rapidly and yield enzymes with diverse substrate specificities, regioselectivities, and tolerance to temperature, pH, solvents, and surfactants [11].

The fermentation medium is a key determinant of the cost and overall sustainability of microbial enzyme production. Partial or complete replacement of refined carbon sources and lipid inducers with low-cost residual materials may improve process economics while supporting waste valorization. Accordingly, numerous solid agro-industrial residues have been investigated as alternative substrates for microbial lipase production [10]. Liquid agro-industrial effluents are a distinct and potentially advantageous feedstock category, as they can be used directly or after appropriate conditioning in submerged fermentation systems. Their residual oils, fatty acids, and glycerides may support microbial growth and stimulate lipase production, as demonstrated with olive mill wastewater used as a culture medium for lipolytic microorganisms [12]. In selected systems, wastewater-based fermentation may also reduce COD, lipid concentrations, oil and grease, or phenolic compounds, thereby combining enzyme production with partial effluent treatment [8,13].

However, implementing wastewater-based lipase production remains challenging. The variable composition of agro-industrial effluents and the presence of potentially inhibitory compounds may necessitate dilution, nutrient supplementation, pH adjustment, or process sterilization, all of which can substantially affect microbial growth and product yields [13,14]. Product recovery and purification may pose additional technical and economic bottlenecks, potentially diminishing the advantages of using low-cost waste-derived media [15]. Moreover, reductions in bulk pollution indicators, such as COD or lipid concentration, do not necessarily demonstrate that the resulting effluent meets discharge or reuse requirements. Its suitability must be evaluated for the intended application and may require consideration of residual toxicity, microbial contamination, persistent pollutants, and other chemical hazards [16]. Process scalability, microbial safety, and the possible need for additional treatment should therefore be included in assessments of the overall feasibility and sustainability of wastewater-based bioprocesses.

Previous reviews have covered microbial lipase production and its biotechnological and industrial applications [9,11], the use of agri-food wastes and renewable residues as alternative substrates for lipase production [10], and the application of microbial lipases in wastewater treatment [17]. However, these reviews do not synthesize studies in which liquid agro-industrial wastewater is intentionally used as a fermentation feedstock for microbial lipase production, nor do they jointly evaluate enzyme production, wastewater-quality changes, medium conditioning and supplementation, heterogeneity in lipase-activity determination, downstream processing, and process scale-up. The present scoping review addresses this gap by integrating these dimensions within a single evidence framework.

Specifically, this review addresses the following questions: (1) Which agro-industrial wastewaters and microbial systems have been investigated for lipase production? (2) How are these wastewaters conditioned, supplemented, and incorporated into cultivation processes? (3) How are lipase production and activity determined and reported, and to what extent are the resulting data comparable across studies? (4) To what extent is lipase production coupled with reductions in wastewater pollutants and with assessment of final-effluent quality? (5) What evidence exists for downstream recovery, catalyst stability and reuse, scale-up, and economic and environmental feasibility?

2. Materials and Methods

2.1. Review Design

This study was conducted as a structured scoping review because the objective was to map and critically characterize a heterogeneous body of evidence rather than to answer a narrowly defined effectiveness question or quantitatively synthesize directly comparable outcomes. The available literature varies substantially in wastewater type and composition, microbial platform, cultivation strategy, lipase localization and assay methodology, and the reporting of wastewater-treatment outcomes. A scoping review was therefore considered more appropriate than a conventional systematic review for identifying the breadth of available evidence, comparing methodological approaches, and defining current research gaps.

The review methodology was based on guidance from the Joanna Briggs Institute, and the reporting followed the Preferred Reporting Items for Systematic Reviews and Meta-Analyses extension for Scoping Reviews (PRISMA-ScR) [18,19]. The review question followed the Population–Concept–Context (PCC) framework. The Population comprised microbial lipase-producing systems, including bacteria, yeasts, filamentous fungi, mixed cultures, and recombinant whole-cell systems. The Concept was the use of agro-industrial wastewater or effluent as a fermentation feedstock for microbial lipase production, including cultivation conditions, enzyme-production outcomes, and, where reported, concurrent changes in wastewater quality. The Context comprised wastewater-based microbial cultivation conducted at the laboratory, pilot, or industrial scale. Accordingly, the primary review question was: What evidence has been reported on the use of agro-industrial wastewaters as fermentation feedstocks for microbial lipase production, and what microorganisms, process configurations, lipase-production outcomes, wastewater-treatment effects, and technological limitations have been described?

The review focused on studies published from January 2016 and 1 August 2026. The year 2016 was selected to provide a defined, approximately ten-year evidence window focused on contemporary wastewater valorization and microbial bioprocessing approaches. Earlier studies were excluded from the main evidence synthesis but were considered, where necessary, to provide historical context or to identify seminal developments.

2.2. Eligibility Criteria

Original peer-reviewed research articles were eligible if they investigated the use of agro-industrial wastewater or effluent as an intentional component of the microbial cultivation medium for lipase production. Wastewater was considered a fermentation feedstock when it was incorporated into the production medium as a source of carbon, nutrients, lipid-associated compounds, or other fermentation-relevant constituents, and lipase production or activity was evaluated in the wastewater-containing cultivation system. No fixed minimum wastewater percentage was imposed because the included literature used highly heterogeneous medium formulations and wastewater concentrations, and the functional role of the wastewater could not be reliably represented by a single numerical threshold.

Studies using untreated or pretreated wastewater were eligible, including those involving centrifugation, filtration, dilution, sterilization, pH adjustment, or nutrient supplementation. Studies in which wastewater constituted only part of the final cultivation medium were also eligible, provided that its use as a fermentation feedstock was an explicit component of the experimental design. In contrast, studies were excluded when wastewater was used only as a trace or incidental additive to an otherwise conventional synthetic or rich medium, and its role as a fermentation feedstock was not meaningfully evaluated.

Eligible studies included bacterial, yeast, fungal, recombinant, and mixed-culture systems conducted at laboratory, pilot, or industrial scales. Studies based primarily on solid-state fermentation of solid agro-industrial residues were excluded if wastewater served only as an auxiliary liquid rather than as the fermentation feedstock.

Studies were excluded if they (i) investigated only solid agro-industrial residues; (ii) used waste oils, oil cakes, food residues, or other by-products without a wastewater or effluent component; (iii) applied purified or commercial lipases solely for wastewater treatment; (iv) evaluated microbial wastewater treatment without measuring lipase production or activity; (v) focused exclusively on esterases or other hydrolytic enzymes; or (vi) were reviews, book chapters, conference abstracts, patents, or publications without accessible full text. Only articles published in English were included in the main analysis, which may have introduced language bias and is acknowledged as a limitation of the review. Review articles were used for background information and backward citation searching but were not included in the evidence tables. The search was designed to capture both generic terminology for agro-industrial effluents and specific wastewater streams known to contain lipid-rich or fermentation-relevant components.

2.3. Information Sources and Search Strategy

The literature search was conducted in Scopus and the Web of Science Core Collection, with the final search performed on 1 August 2026. These two multidisciplinary databases were selected for their broad and complementary coverage of peer-reviewed literature in biotechnology, food science, environmental science, and process engineering, the principal disciplines represented in the review topic. Review articles identified during the search were also used for backward citation searching to identify potentially relevant primary studies. The following search strategy was used in Scopus:

TITLE-ABS-KEY (((lipase* OR (lipolytic W/1 enzyme*)) AND ((agroindustr* W/3 wastewater*) OR (agroindustr* W/3 effluent*) OR (“agro-industrial” W/3 wastewater*) OR (“agro-industrial” W/3 effluent*) OR (“agro-food” W/3 wastewater*) OR (“agro-food” W/3 effluent*) OR (“food processing” W/3 wastewater*) OR (“food processing” W/3 effluent*) OR “olive mill wastewater” OR “olive oil mill wastewater” OR “palm oil mill effluent” OR “dairy wastewater” OR “dairy effluent” OR “cheese wastewater” OR “whey wastewater” OR “slaughterhouse wastewater” OR “abattoir wastewater” OR “poultry processing wastewater” OR “fish processing wastewater” OR “seafood processing wastewater” OR “edible oil refinery wastewater” OR “vegetable oil processing wastewater” OR “oil mill wastewater” OR “oily wastewater” OR “lipid-rich wastewater”) AND (produc* OR ferment* OR cultivat* OR biosynthes* OR bioprocess* OR feedstock* OR “culture medium” OR valorization OR valorisation))) AND PUBYEAR > 2015 AND PUBYEAR < 2027.

The search syntax was adapted to the requirements of Web of Science while retaining the same conceptual blocks, i.e., TS = ((lipase* OR (lipolytic NEAR/1 enzyme*)) AND (“olive mill wastewater” OR “olive oil mill wastewater” OR “palm oil mill effluent” OR “dairy wastewater” OR “dairy effluent” OR “cheese wastewater” OR “whey wastewater” OR “slaughterhouse wastewater” OR “abattoir wastewater” OR “poultry processing wastewater” OR “fish processing wastewater” OR “seafood processing wastewater” OR “edible oil refinery wastewater” OR “vegetable oil processing wastewater” OR “oil mill wastewater” OR “food processing wastewater” OR “food processing effluent” OR “agro-industrial wastewater” OR “agroindustrial wastewater” OR “agro-industrial effluent” OR “agroindustrial effluent” OR “agro-food wastewater” OR “agro-food effluent” OR “oily wastewater” OR “lipid-rich wastewater”) AND (produc* OR ferment* OR cultivat* OR biosynthes* OR bioprocess* OR feedstock* OR “culture medium” OR valorization OR valorisation)) AND PY = (2016–2026). The results were limited to publications from 2016–2026 and to English-language articles.

2.4. Study Selection

All retrieved records were exported, combined, and checked for duplicates. The selection process was conducted in two stages. First, titles and abstracts were screened against the eligibility criteria. Then, the full texts of potentially relevant publications were assessed for final inclusion. Screening was performed independently by two reviewers at both the title/abstract and full-text stages. Disagreements regarding study eligibility were resolved through discussion until consensus was reached. No formal inter-reviewer agreement statistic was calculated. Reasons for exclusion at the full-text stage were recorded. The study-selection process was summarized in a PRISMA-ScR flow diagram showing the numbers of identified, screened, excluded, and included records.

2.5. Data Extraction

Data were extracted using a standardized form developed for this review. The form was pilot-tested on a subset of eligible publications and adjusted as needed. One reviewer extracted the data, and a second reviewer checked it against the original publications. The reviewers resolved any discrepancies identified during verification through discussion. The following information was collected:

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Publication year and country;

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Type, origin, and initial characteristics of the wastewater;

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Pretreatment, dilution, sterilization, and supplementation procedures;

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Microorganism, strain, and inoculum characteristics;

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Fermentation mode, scale, duration, temperature, pH, aeration, and agitation;

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Carbon and nitrogen sources, lipid inducers, and other medium supplements;

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Lipase assay substrate, analytical conditions, and activity units;

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Reported maximum lipase activity, productivity, and specific activity;

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Changes in wastewater parameters, including COD, BOD, lipid content, oil and grease, and phenolic compounds;

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Enzyme recovery, concentration, purification, formulation, or direct use of crude enzymes or whole-cell biocatalysts;

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Reported limitations and proposed applications.

The primary outcomes of interest were microbial lipase production and activity in wastewater-containing cultivation systems and, where evaluated, concurrent changes in wastewater-quality parameters. Secondary outcomes included cultivation conditions, wastewater conditioning and medium supplementation, enzyme localization, downstream processing, biochemical characterization, catalyst stability and reuse, and reported applications. For studies examining multiple microorganisms or fermentation strategies, each system was documented separately when sufficient data were available.

2.6. Data Synthesis

The extracted data were synthesized descriptively and organized into predefined comparison domains reflecting the main sources of heterogeneity across the evidence base: (i) wastewater type and degree of wastewater use in the cultivation medium; (ii) microbial group and enzyme localization; (iii) cultivation mode, scale, and process conditions; (iv) lipase assay methodology and activity-expression basis; (v) wastewater-conditioning and supplementation strategies; (vi) concurrent wastewater-quality changes; and (vii) downstream processing, characterization, and catalytic application.

Lipase activities were retained in the units reported by the original authors because substantial differences in assay substrates, reaction conditions, enzyme fractions, and unit definitions frequently precluded reliable conversion. Values were standardized only when the original data and unit definitions allowed an unambiguous conversion. Accordingly, activity values were treated as study-specific outcomes and were not used to rank microorganisms or wastewater substrates. When outcomes were reported using directly comparable measurement bases, numerical ranges were summarized descriptively. Otherwise, results were presented separately without quantitative ranking. No categorical thresholds for low, moderate, or high lipase activity or productivity were applied because such categories would not be valid across the heterogeneous assay systems.

The findings were presented in comparative tables and figures summarizing wastewater streams, microbial systems, cultivation conditions, lipase-production outcomes, wastewater-treatment indicators, and downstream applications. Where relevant, results were distinguished based on whether pollutant reduction occurred concurrently with lipase-producing cultivation or during a separate post-production application of harvested cells or enzymes. Different wastewater-treatment endpoints, such as COD reduction, O&G (oil and grease) removal, triglyceride degradation, and phenolic-compound reduction, were treated as distinct outcomes and were not considered directly interchangeable.

No meta-analysis was performed because the included studies were highly heterogeneous in wastewater composition, cultivation conditions, enzyme localization, lipase assays, outcome definitions, and reporting formats. A formal risk-of-bias assessment was not conducted because the purpose of this scoping review was to map and critically characterize the breadth and methodological features of the available evidence rather than to estimate a pooled intervention effect or comparative treatment efficacy. Instead, methodological characteristics affecting the interpretability and comparability of individual studies, i.e., including wastewater characterization, medium composition, cultivation scale, assay methodology, and completeness of wastewater-quality assessment, were extracted and incorporated into the descriptive synthesis and critical discussion. These methodological differences were also considered when interpreting apparent differences in process performance, and no microorganism or technological approach was designated as universally superior on the basis of the reported activity or treatment values alone.

3. Results

3.1. Literature Search Results and Study Selection

The database search identified 222 records, including 118 from Scopus and 104 from the Web of Science Core Collection. After removing 89 duplicate records, 133 unique publications remained for title and abstract screening. Of these, 112 records were excluded because they did not meet the predefined eligibility criteria. At the title/abstract screening stage, records were excluded when they clearly fell outside the predefined scope, for example, because they did not involve agro-industrial wastewater as a fermentation feedstock, did not evaluate microbial lipase production, focused primarily on solid residues or waste oils without a wastewater component, or represented non-eligible publication types. Individual exclusion reasons were not coded at this preliminary screening stage. The full texts of 21 potentially relevant articles were retrieved and assessed for eligibility. One article was excluded because the wastewater was used only as a minor additive rather than as an eligible fermentation feedstock. Consequently, 20 original research articles were included in the final qualitative synthesis. The study identification and selection process is presented in Figure 1.

3.2. General Characteristics of the Included Studies

The 20 studies included in the review, published between 2016 and 2025, investigated microbial lipase production using agro-industrial wastewaters either as the primary cultivation medium or as a major component of the fermentation system [20,21,22,23,24,25,26,27,28,29,30,31,32,33,34,35,36,37,38,39]. Their general characteristics, including wastewater type, microbial platform, culture system, and primary research objective, are summarized in

Table 1. General characteristics of studies investigating microbial lipase production using agro-industrial wastewaters.

Country	Wastewater Type	Microorganism	Microbial Group	Culture System	Main Study Objective	Reference
Tunisia	OMW	Fusarium solani	Filamentous fungus	Monoculture; batch shake-flask cultivation; biotreatment assay	Lipase production in an OMW-based medium combined with evaluation of COD, oil, sugar, and phenolic-compound removal	[20]
Thailand	POME	Yarrowia lipolytica TISTR 5151	Yeast	Initial strain screening followed by monoculture optimization in POME	Valorization of POME into microbial lipid and cell-bound lipase for direct biodiesel production	[21]
Portugal	OMW, winery vinasse, and an OMW–vinasse mixture	Different strains of Aspergillus ibericus; A. uvarum; A. niger	Filamentous fungi	Separate fungal monocultures; batch cultivation in individual and mixed effluents	Combined bioremediation of olive-mill and winery wastewaters with production of lipase, protease, and tannase	[22]
Thailand	POME	Candida rugosa CU1; Y. lipolytica rM-4A; their co-culture	Yeasts	Single cultures and co-culture in undiluted POME; parallel treatment with crude extracellular enzymes	Comparison of live-cell and crude-enzyme approaches for POME biodegradation, including lipase production and pollutant removal	[23]
India	POME	F. solani NFCCL 4084	Filamentous fungus	Monoculture; shake-flask cultivation; Plackett–Burman, OFAT, and face-centred CCD	Statistical optimization of low-cost halophilic lipase production in a POME-based medium	[24]
Indonesia	POME	Pseudomonas aeruginosa B2290	Bacterium	Monoculture; submerged shake-flask fermentation; OFAT optimization; spray drying	Optimization of lipase production in POME and preparation of a dry crude-lipase extract	[25]
Malaysia	POME	Meyerozyma guilliermondii	Yeast	Monoculture; aerobic shake-flask cultivation in unsupplemented and supplemented POME	Evaluation of an indigenous yeast for POME bioremediation and extracellular lipase production	[26]
Nigeria	POME	Mutant Bacillus niacini EMB-5	Bacterium	Isolate screening and mutagenesis; POME cultivation; downstream enzyme purification	Enhancement of POME-based lipase production and characterization of the purified alkaline thermostable enzyme	[27]
Iran	VORW	Y. lipolytica CBS 6303	Yeast	Monoculture; shake-flask optimization followed by bioreactor cultivation	Simultaneous production of lipase, microbial oil, and lipid-rich biomass with reductions in wastewater COD	[28]
Portugal	OMW	M. capitatus JT5	Yeast	Isolation and screening; shake-flask optimization; stirred-tank bioreactor	Isolation of an OMW-adapted lipase-producing yeast, optimization of extracellular lipase production, and bioreactor scale-up	[29]
Portugal	OMW	C. tropicalis ATCC 750	Yeast	Monoculture; shake-flask optimization followed by stirred-bioreactor cultivation	Combination of OMW detoxification with production of lipase, protease, microbial lipids, and biomass	[30]
Indonesia	POME	Recombinant A. oryzae expressing C. antarctica lipase B	Recombinant filamentous fungus/whole-cell biocatalyst	Whole-cell cultivation and immobilization in biomass-support particles using POME as carbon source	Use of POME to support immobilized whole-cell lipase preparation and its application in biodiesel synthesis	[31]
Thailand	POME blended with waste frying oil	M. spicifer AW2; Staphylococcus hominis AUP19	Yeast–bacterium co-culture	Synthetic co-culture; statistically optimized blended-waste medium; subsequent application assays	Optimization of cell-bound lipase production and demonstration of applications in biodiesel synthesis and POME bioremediation	[32]
Thailand	POME	M. spicifer AW2; S. hominis AUP19	Yeast–bacterium system	Pure cultures and synthetic co-culture; batch cultivation in diluted POME	Production of cell-bound lipases integrated with POME bioremediation and biodiesel synthesis	[33]
Greece	OMW	B. aryabhattai BA03	Bacterium	Monoculture; shake-flask cultivation; sterile and non-sterile media; factorial optimization	Evaluation of OMW as a low-cost lipase-production medium and assessment of wastewater concentration, pH, temperature, and sterilization	[34]
Algeria	OMW	A. oryzae 1042.72	Filamentous fungus	Monoculture; submerged fermentation; Plackett–Burman design and CCD/RSM	Optimization of OMW-based medium components for enhanced extracellular lipase production	[35]
Algeria	OMW	Streptomyces sp. SC1	Bacterium (actinobacterium)	Monoculture; submerged shake-flask fermentation; Box–Behnken design/RSM	Isolation of lipase-producing actinobacteria and optimization of OMW-based bacterial lipase production	[36]
Thailand	POME	M. spicifer AW2; Staphylococcus hominis AUP19	Yeast–bacterium co-culture	Synthetic 1:1 co-culture; submerged batch cultivation in POME50; PBD followed by RSM-CCD	Integrated optimization of cell-bound lipase and biomass production with COD/O&G removal, followed by storage-stability evaluation and solvent-free biodiesel synthesis	[37]
Indonesia	POME	Pichia sp. Y1440; T. coremiiforme Y771	Yeasts	Separate monocultures; shake-flask cultivation; OFAT and central composite design/RSM	Optimization of POME concentration and pH for lipase production by two lipolytic yeasts	[38]
Algeria	OMW; waste frying oil as comparator	Y. lipolytica L2	Yeast	Monoculture; shake-flask cultivation under increasing ethanol stress	Assessment of ethanol-stress effects on lipase production, microbial growth, lipid accumulation, and fatty-acid remodeling in OMW-based cultivation	[39]

Note: Country refers primarily to the origin of the wastewater used in the study. Where the wastewater source was not explicitly stated, the country of the experimental study was used. Abbreviations: OMW, olive mill wastewater; POME, palm oil mill effluent; VORW, vegetable oil refinery wastewater; O&G, oil and grease; OFAT, one-factor-at-a-time; CCD, central composite design; RSM, response surface methodology.

. Unless otherwise stated, quantitative counts in this section refer to publications rather than individual experimental systems. A study evaluating multiple microorganisms, strains, culture configurations, or operating conditions was therefore counted once at the study level, while the individual experimental systems are described separately in the corresponding tables.

The available evidence was heavily concentrated on wastewater generated by the vegetable oil industry. Eleven of the 20 included studies (55%) investigated POME-containing systems, including ten studies using POME as the principal wastewater feedstock [21,23,24,25,26,27,31,33,37,38] and one using POME blended with waste frying oil [32]. Eight studies (40%) investigated OMW-containing systems [20,22,29,30,34,35,36,39], and among these, one additionally examined winery vinasse and an OMW–winery vinasse mixture [22], while another compared OMW with a waste-oil substrate [39]. One study reported VORW (5%) [28]. No eligible studies involving dairy, slaughterhouse, poultry-processing, or fish-processing wastewaters were identified within the present search strategy and eligibility criteria.

The predominance of POME and OMW likely reflects their high organic and lipid loads, residual lipids that can support or induce lipolytic metabolism, and their availability in major palm- and olive-oil-producing regions. However, this concentration of evidence also limits the generalizability of conclusions to other agro-industrial effluents. Accordingly, the absence of eligible studies for other wastewater categories in this review should not be interpreted as evidence that no such research exists, but rather as indicating limited evidence identified within the defined review scope.

At the study level, yeast-only systems were investigated in eight studies (40%) [21,23,26,28,29,30,38,39], including species of Yarrowia, Candida, Meyerozyma, Magnusiomyces, Pichia, and Trichosporon. Bacterial monocultures accounted for four studies (20%) [25,27,34,36], involving Pseudomonas, Bacillus, and Streptomyces. Filamentous-fungal systems were investigated in five studies (25%) [20,22,24,31,35], including the recombinant A. oryzae whole-cell system reported in [31]. Three studies (15%) examined synthetic yeast–bacterium systems based on M. spicifer AW2 and AUP19 [32,33,37]. Several publications evaluated more than one strain, monoculture, or co-culture configuration, and these were treated as separate experimental systems in the detailed synthesis but not as separate studies in the percentages reported above.

Most of the evidence remained at the laboratory flask scale. Only four of the 20 studies (20%) included a controlled stirred-tank bioreactor stage [25,28,29,30], whereas the remaining studies were conducted exclusively in flask-based, static, or other laboratory-scale systems. More advanced downstream or application-oriented steps were even less common and included enzyme purification or spray drying [25,27], whole-cell immobilization [31], and direct catalytic use of cell-bound or immobilized lipases [21,31,32,33,37].

The evidence map in Figure 2 highlights the uneven distribution of research across wastewater types and microbial groups. Available evidence is concentrated primarily on OMW- and POME-based processes [20,21,22,23,24,25,26,27,28,29,30,31,32,33,34,35,36,37,38,39], whereas VORW was represented by only one study [28] and combined waste streams were investigated only sporadically [22,32]. Yeasts show the broadest coverage across wastewater categories [21,23,26,28,29,30,32,33,37,38,39], whereas bacterial and filamentous fungal systems are largely confined to OMW- and POME-based media [20,22,24,25,27,31,34,35,36]. Mixed yeast–bacterium systems have been evaluated only in POME-containing processes, including POME alone and a POME–waste frying oil blend [32,33,37]. This distribution indicates that the current knowledge base is strongly focused on oil-processing effluents and that the potential of other agro-industrial wastewaters for microbial lipase production remains largely unexplored.

3.3. Wastewater Characteristics and Conditioning Prior to Fermentation

The physicochemical characteristics and conditioning procedures of the agro-industrial wastewaters used in the included studies are summarized in

Table 2. Physicochemical characteristics and conditioning of agro-industrial wastewaters used for microbial lipase production.

Wastewater Type and Source	Reported Initial Characteristics	Wastewater Proportion and Use In Culture Medium	Pretreatment and Storage	Sterilization	pH Management	Supplementation	References
OMW. Local olive-oil manufacturer, Sfax, Tunisia	pH: 5.46 ± 0.04; COD: 80.59 ± 0.25 g/L; Lipids: 2.40 ± 0.36 g/L; phenolics: 12.24 ± 0.15 g/L (gallic acid equivalents); other: Total sugars: 40.11 ± 0.29 g/L; total nitrogen: not detected	Undiluted OMW; 50 mL in 250-mL flasks	Centrifuged at 4000× g for 15 min to remove solids; stored at −20 °C	Sterilized before cultivation	Adjusted to pH 6 before cultivation; no further pH correction	Nitrogen sources tested at 5 g/L: yeast extract, soy peptone, and (NH4)2SO4; yeast extract selected	[20]
POME. Industrial palm-oil plants, Songkhla Province, Thailand	pH: 4.3–4.5; COD: 37–48 g/L; Oil and grease: 0.200–0.282 g/L; phenolics: NR; other: TKN: 0.103–0.262 g/L	Twofold-diluted and undiluted POME were compared; undiluted POME selected for optimization	Centrifuged before use	Autoclaved before cultivation	Raw pH 4.3 and adjusted pH 5.0, 5.5, and 6.0 tested; pH 5.5 used in the optimized process	(NH4)2SO4 added at 1 g N/L to undiluted POME	[21]
OMW; VS. OMW: olive mill in Trás-os-Montes, Portugal; VS: winery in Minho, Portugal	pH: NR; COD: OMW: 122.9 ± 0.42 g/L; VS: 48.07 ± 1.43 g/L; OMW lipids: 4.10 ± 0.42 g/L; VS: not detected; phenolics: OMW: 5.91 ± 0.09 g/L; VS: 0.54 ± 0.03 g/L; other: OMW: TOC 21.4 ± 1.2, sugars 12.7 ± 1.3, solids 15.48 ± 0.54 g/L, N 5.2 ± 0.2 mg/L; VS: TOC 3.53 ± 0.02, sugars 0.68 ± 0.01, solids 22.24 ± 0.09 g/L, N 218.67 ± 35.27 mg/L	Undiluted OMW; OMW:nutrient medium 1:1 (v/v); OMW:VS 1:1 (v/v)	Effluents homogenized and stored separately at −20 °C	121 °C for 15 min	No explicit pH adjustment reported	Nutrient medium: NaNO3 3 g/L, K2HPO4 1 g/L, KCl 0.5 g/L, MgSO4·7H2O 0.5 g/L, CaCl2·2H2O 0.5 g/L, and trace-metal solution	[22]
POME. Palm-oil mill, Chonburi Province, Thailand	pH: 4.3; COD: 34,584 mg/L; Oil and grease: 2762 mg/L; phenolics: 9782.5 mg/L; other: BOD 33,090; TS 56,100; TSS 12,430; VS 46,930; TKN 1693; total P 44.3; reducing sugars 3449.8 mg/L	Live-cell cultures: undiluted POME; separate crude-enzyme treatment also used twofold-diluted POME	Stored at −20 °C; diluted-enzyme-treatment arm was centrifuged at 4000 rpm for 15 min	Live-cell undiluted POME was autoclaved; sterile and non-sterile enzyme-treatment arms were also examined	Raw pH used for live-cell cultivation; diluted enzyme-treatment medium adjusted to pH 5.6	Live-cell cultivation supplemented with 0.5% (w/v) palm oil; no additional nutrient supplementation	[23]
POME. Palm-oil plant near Kuvempu University, Shankaraghatta, India	pH: NR; COD: NR; NR; phenolics: NR; other: Initial physicochemical composition not reported	POME varied in statistical screening and OFAT experiments (approximately 0.25–1.25%, w/v)	Collected in a clean container and stored at 4 °C	121 °C, 15 psi, 20 min	Adjusted with 1 M HCl or NaOH; pH ranges defined by experimental designs	Malt extract, (NH4)2SO4, CaCl2, MgSO4, olive oil, peptone, K2HPO4, NaNO3, and Tween 80; K2HPO4, NaNO3, and Tween 80 subsequently optimized	[24]
POME. Palm-oil company, Bengkulu, Indonesia	pH: NR; COD: NR; NR; phenolics: NR; other: Initial physicochemical composition not reported	POME used as the basal medium (100 mL per flask)	Stored at 4 °C	Sterilized POME used	NR	Initial medium: olive oil 0.2% (v/v), peptone 0.5% (w/v), CaCl2·2H2O 10 mM, Tween 80 0.7% (v/v); reported optimum: olive oil 0.4%, peptone 0.9%, Ca2+ 4 mM, Tween 80 0.9%	[25]
POME. Local palm-oil mill, Johor, Malaysia	pH: 4.6 ± 0.1; COD: 35,983.5 ± 110 mg/L; Oil and grease: 3750 ± 11 mg/L; phenolics: Phenolic compounds analyzed qualitatively by FTIR/GC–MS; initial total concentration NR; other: Total N 833.4 ± 12; ammoniacal N 91.7 ± 4; TOC 3119 ± 22; phosphate 581.7 ± 8 mg/L	Undiluted POME used as the sole carbon source	Transported on ice and stored at 4 °C; autoclaved and then centrifuged at 4000 rpm, 4 °C, 15 min to remove particulates	121 °C for 15 min	Main treatment used raw POME without pH adjustment; acclimatization medium was adjusted to pH 4.5	Carbon sources screened at 1.0% (w/v) and nitrogen sources at 0.5% (w/v); glycerol 1% and yeast extract 0.5% were most effective	[26]
POME. Oil-palm processing industries, Ipinsa, Akure, southwest Nigeria	pH: NR; COD: NR; NR; phenolics: NR; other: Initial physicochemical composition not reported	Quantitative screening: 10 mL POME in 50 mL basal medium; production-scale description reports 10 mL POME in approximately 1 L medium	NR	Not explicitly reported	Adjusted to pH 5.0	(NH4)2SO4, Na2HPO4, KH2PO4, MgSO4, and CaCl2; POME was emulsified with Tween 80	[27]
VORW. Pars Vegetable Oil Company, Tehran, Iran	pH: NR; COD: NR; Oil and grease: 3–4 g/L; palmitic 36.68%, stearic 28.18%, oleic 32.00%, linoleic 3.14%; phenolics: NR; other: Initial COD and other wastewater-composition data not reported	Primary medium: 30 mL/L; RSM range: 25–75 mL/L; optimized at 75 mL/L	NR	NR	Optimized at pH 6; bioreactor pH maintained at 6 using KOH or H3PO4	Primary medium included yeast extract, peptone, KH2PO4, NaH2PO4, and MgSO4·7H2O; optimized medium reported as yeast extract 0.5 g/L and peptone 4 g/L	[28]
OMW. Three-phase olive mill, Algarve, Portugal; sampled during the final month of the campaign	pH: 4.99; COD: 55.2 g/L; Lipids: 3.20 g/L; phenolics: 3.97 g caffeic acid equivalents/L; other: Reducing sugars 35 g/L; TSS 21.9 g/L; NO3− 1.42 g/L; NH4+ 1.70 g/L	Growth screening: 10–75% OMW in YEP; lipase-production and bioreactor experiments: undiluted OMW	Filtered through gauze and centrifuged at 8600× g for 15 min	121 °C for 20 min	Adjusted to pH 6.8 in shake-flask production and pH 6.1 before bioreactor sterilization	Yeast extract 2 g/L; olive oil 1–3 g/L; NH4Cl 0.2–2.8 g/L	[29]
OMW (three batches). Olive mills in northern Portugal	pH: OMW-1: 5.03 ± 0.01; OMW-2: 4.83 ± 0.02; OMW-3: 4.94 ± 0.09; COD: OMW-1: 108.7 ± 0.4; OMW-2: 51.3 ± 1.3; OMW-3: 36.7 ± 3.0 g/L; NR; phenolics: OMW-1: 4.83 ± 0.05; OMW-2: 2.60 ± 0.10; OMW-3: 1.80 ± 0.10 g/L; other: OMW-1/2/3, respectively: TOC 46.6/20.1/15.0 g/L; reducing sugars 36.3/13.2/10.1 g/L; TN 285.7/152.4/626.8 mg/L	Screening: 5–50% (v/v); main shake-flask and bioreactor experiments: 50% OMW	NR	Sterile OMW used in screening; sterilization details for the main flask/bioreactor media were not explicitly stated	Initial pH values of 5 and 7 tested; maintained at pH 5.5 or 7 in the bioreactor using HCl/NaOH	YNB without amino acids; NH4Cl, (NH4)2SO4, or urea supplied at an equivalent 1.06 g N/L; bioreactor medium contained 4.05 g/L NH4Cl	[30]
POME. PT Agricinal Palm Oil Mills, Bengkulu, Indonesia	pH: NR; COD: NR; Acid value 169.72 ± 0.46 mg KOH/g oil; saponification value 211.70 ± 8.51 mg KOH/g oil; iodine value 53.54 ± 1.50 g I2/100 g lipid; phenolics: NR; other: Total carbon 15.82 ± 0.01 g/L; TOC 15.07 ± 0.01 g/L	Whole POME without separation, 2 g per 100 mL medium (2%, w/v), used instead of glucose	Whole POME used without separation	Not explicitly reported for the POME-based culture medium	NR	Per 100 mL: polypeptone 2 g, KH2PO4 0.5 g, NaNO3 0.1 g, MgSO4·7H2O 0.05 g; polyurethane biomass-support particles	[31]
POME blended with WFO. POME: Larp Tavee Palm Oil Co., Ltd., Satun, Thailand; WFO: local fried-chicken stalls, Songkhla, Thailand	pH: POME50: approximately 3.8 before adjustment; COD: POME50: 29,747 mg/L; POME50 oil and grease: 5562 mg/L; phenolics: NR; other: POME50 TKN: 277 mg/L	Base medium: centrifuged POME diluted 1:1 to 50% (POME50)	Raw POME centrifuged at 4000× g, 4 °C, 15 min; centrifuged sample stored at −20 °C	121 °C, 15 psi, 15 min	Production media adjusted to pH 7.0	WFO, crude glycerol, or molasses initially tested at 2% (v/v); nitrogen sources at 0.5% (w/v); surfactants at 0.1%. Optimized medium contained WFO, (NH4)2SO4, and gum Arabic	[32]
POME. Palm-oil industrial plant, Satun Province, Thailand	pH: Raw POME 3.84 ± 0.02; POME100 3.80 ± 0.03; POME50 3.80 ± 0.00; POME25 3.92 ± 0.02; COD: Raw 67,145 ± 563; POME100 50,581 ± 350; POME50 29,747 ± 557; POME25 18,889 ± 278 mg/L; O&G: raw 11,382 ± 300; POME100 7733 ± 197; POME50 5562 ± 106; POME25 2783 ± 399 mg/L; phenolics: NR; other: TKN: raw 1462 ± 167; POME100 526 ± 25; POME50 277 ± 38; POME25 114 ± 6 mg/L; TS and TSS also reported	POME100, POME50 (1:1 dilution), and POME25 (1:3 dilution) compared	Raw POME centrifuged at 4000× g, 4 °C, 15 min; stored at 4 °C before processing and at −20 °C after centrifugation	121 °C, 15 psi, 15 min	Natural pH approximately 3.8 and pH-adjusted media at 7.0 were compared	No external nutrients in POME media; standard synthetic BSM served as comparator	[33]
OMW. Three-phase olive mill, Kalamata, Peloponnese, Greece	pH: Initial wastewater pH NR; COD: NR; NR; phenolics: Initial concentration NR; sterilization increased measured phenols by 17.25 ± 0.05%; other: Sterilization reduced total sugars by 14.99 ± 0.10%	Undiluted, 50%, and 25% OMWW; best activity obtained with 50% non-sterile OMWW	Frozen at −20 °C until use	Sterile and non-sterile media compared; sterilization at 121 °C for 20 min	Adjusted to pH 6, 7, or 8	Yeast extract 10 g/L	[34]
OMW. Disposal ponds of a three-phase olive-oil process, Jijel, Algeria	pH: NR; COD: NR; Lipids: 1.50 ± 0.30% (as reported); phenolics: NR; other: Sugars 0.12 ± 0.04%; total nitrogen 3.06 ± 0.50%; minerals 6.15 ± 0.06%	OMW-based medium; exact OMW fraction not reported	Centrifuged at 3800× g for 20 min	Not explicitly reported	pH 5–6 evaluated in Plackett–Burman design	Peptone, glucose, CaCl2, Tween 80, NaNO3, and yeast extract; glucose, yeast extract, and CaCl2 selected for CCD/RSM	[35]
OMW. OMW source and collection procedure not reported; actinobacteria were isolated from olive-mill-associated sites in Béjaïa, Algeria	pH: NR; COD: NR; NR; phenolics: NR; other: Initial physicochemical composition not reported	OMW 5–45% (v/v) in Box–Behnken design; optimum predicted/validated at approximately 6% (v/v)	NR	NR	Initial pH 4–10 studied; optimum at pH 4	OMW described as the sole carbon source; other production-medium constituents were not clearly specified for the optimization stage	[36]
POME. Larp Tavee Palm Oil Co., Ltd., Satun, Thailand	POME100 after centrifugation: pH 3.80; COD 50,581 ± 350 mg/L; O&G 7733 ± 198 mg/L; TKN 526 ± 25 mg/L. POME50: pH 3.80; COD 29,747 ± 557 mg/L; O&G 5562 ± 106 mg/L; TKN 277 ± 38 mg/L	POME100 was diluted 1:1 with deionized water to obtain POME50; POME50 was used as the cultivation medium	POME centrifuged at 4000× g, 4 °C, 15 min; supernatant designated POME100 and diluted 1:1 with deionized water	Autoclaved at 121 °C for 15 min	Initial pH was included in PBD and RSM-CCD; optimized process at pH 7.0	(NH4)2SO4 evaluated as nitrogen source; optimized concentration 1.23% (w/v)	[37]
POME. POME source not reported	pH: NR; COD: NR; NR; phenolics: NR; other: Initial physicochemical composition not reported	Initial comparison used 50% POME; OFAT tested 10, 25, 50, and 75%; RSM extended the tested range; confirmation optima were 80% for Pichia sp. and 53.1% for T. coremiiforme	NR	Media prepared fresh and autoclaved at 121 °C for 15 min	OFAT tested pH 5–8; RSM optima were pH 5 for Pichia sp. and pH 8 for T. coremiiforme	Enriched 50% POME medium contained peptone 10 g, yeast extract 0.5 g, Tween 80 1.5 mL, and olive oil 2.5 mL per 100 mL POME–water base	[38]
OMW; WCO as a separate comparator. OMW obtained in May 2024 from an olive-oil extraction facility in Béjaïa, Algeria	pH: NR; COD: NR; NR; phenolics: Polyphenols: 2 g/L; other: Other initial OMW characteristics not reported	OMW added at 200 g/L as the hydrophobic carbon source in YP medium	No OMW pretreatment or storage conditions reported	Not explicitly reported for OMW	NR	YP medium: peptone 20 g/L and yeast extract 10 g/L; ethanol added after 24 h at 0, 3, 5, or 7% (v/v)	[39]

Note: Values were retained in the units and terminology used by the original authors. NR indicates not reported. Where several wastewater batches, dilutions, or experimental variants were examined, the relevant range or principal production condition is shown. Where reported, the proportion of wastewater in the final cultivation medium is provided. Where the exact proportion was not stated in the original study, the reported qualitative description of wastewater use was retained. Abbreviations: OMW, olive mill wastewater; POME, palm oil mill effluent; VORW, vegetable oil refinery wastewater; VS, winery vinasse; WFO, waste frying oil; WCO, waste cooking oil; COD, chemical oxygen demand; TOC, total organic carbon; TKN, total Kjeldahl nitrogen; O&G, oil and grease; YNB, yeast nitrogen base; YP, yeast extract–peptone medium.

. Considerable heterogeneity was observed both across wastewater types and within batches of the same effluent. Reported COD values for OMW-containing streams ranged from approximately 36.7 to 122.9 g/L [22,30], whereas POME generally exhibited COD values of approximately 29.7–67.1 g/L, depending on its origin, dilution level, and pretreatment [21,23,26,32,33,37]. The effluents also differed markedly in lipid or oil-and-grease content, nitrogen availability, reducing sugars, suspended solids, and phenolic-compound concentrations [20,21,22,23,26,29,30,32,33,37]. In some studies, separate batches or concentrations of the same wastewater showed substantially different compositions, confirming that wastewater origin, sampling period, processing history, and dilution can influence its suitability as a fermentation substrate [30,33]. Importantly, high concentrations of organic matter, lipids, or phenolic compounds should not be interpreted as inherently advantageous for fermentation. Although these constituents may provide carbon sources or induce lipase production, high organic loads and inhibitory constituents can also limit microbial growth and increase process-control requirements.

The completeness of wastewater characterization varied widely across the included studies. Several publications reported relatively detailed initial values for COD, BOD, oil and grease, nitrogen, solids, sugars, or phenolic compounds [20,21,22,23,26,29,30,32,33,37]. For example, the POME used by Theerachat et al. contained 34,584 mg/L COD, 2762 mg/L oil and grease, and 9782.5 mg/L total phenolic compounds [23]. In contrast, other studies reported only selected parameters or omitted the initial physicochemical composition of the wastewater [24,25,27,28,34,35,36,38,39]. Such incomplete characterization is a major limitation of the existing evidence base because the reproducibility and comparability of wastewater-based fermentation processes cannot be adequately assessed when the initial feedstock composition is poorly described. Future studies should, at minimum, report wastewater pH, COD, BOD (where relevant), TOC, oil and grease or total lipids, total phenolic compounds (where applicable), total nitrogen or TKN, and TSS. When evaluating wastewater treatment, determine the same core parameters after cultivation to enable a meaningful assessment of treatment performance. Differences in analytical terminology and reporting units were also evident, particularly for lipids, oil and grease, and phenolic compounds, limiting direct quantitative comparisons across studies [20,21,23,29,31,35].

Wastewater conditioning ranged from minimal preparation to extensive modification of the fermentation matrix. Common procedures included centrifugation or filtration to remove suspended solids, refrigerated or frozen storage, dilution with water, pH adjustment, and thermal sterilization [20,21,22,23,26,29,32,33,34,37,38]. Several microorganisms were cultivated in undiluted OMW or POME, demonstrating tolerance to high organic loads and, in some cases, elevated concentrations of phenolic compounds [20,21,23,26,29]. In the study by Theerachat et al., the yeast cultures were grown in undiluted POME supplemented with 0.5% (w/v) palm oil, while no other nutrient supplementation was reported [23].

Other studies used diluted wastewater, with dilution ranging from low concentrations to approximately 50–80% of the final medium [24,27,28,30,31,33,34,36,37,38]. Fibriana et al. demonstrated in two related studies that twofold-diluted POME (POME50) adjusted to pH 7.0 supported cell-bound lipase production by the M. spicifer AW2–S. hominis AUP19 co-culture [33,37]. In the later study, POME50 was supplemented only with ammonium sulfate as the additional nutrient, and its concentration was subsequently optimized alongside process variables [37]. These results indicate that dilution may help reduce the organic load and other potentially inhibitory characteristics of concentrated POME, although it also increases water consumption and reduces the concentration of wastewater-derived nutrients.

Sterilization was used in most cultivation systems for which this information was reported, typically by autoclaving [20,21,22,23,24,25,26,29,32,33,34,37,38]. However, the feasibility of non-sterile processing was examined only occasionally. Paz et al. directly compared sterilized and non-sterilized OMW-based media and achieved the highest lipase activity with 50% non-sterile OMW at pH 8 and 27 °C [34]. This result suggests that sterilization may not always be required, although the reproducibility, contamination risk, and long-term stability of non-sterile processes remain insufficiently investigated.

Although wastewater generally served as the principal carbon source or fermentation matrix, supplementation was common. Nitrogen sources such as yeast extract, peptone, ammonium salts, and urea were frequently added, along with mineral salts, additional oils, surfactants, or emulsifying agents [20,21,22,24,25,26,28,29,30,31,32,34,35,37,38,39]. In the POME–waste frying oil system developed by Fibriana et al., supplementation with waste frying oil, ammonium sulfate, and gum Arabic yielded a maximum cell-bound lipase activity of 4708.9 U/L [32]. In a subsequent POME50-based process using the same M. spicifer AW2–S. hominis AUP19 co-culture, the medium was simplified to ammonium sulfate supplementation at an optimized concentration of 1.23% (w/v) [37]. This later study therefore exemplifies reducing the number of externally added medium components while retaining POME as the principal fermentation matrix. Similarly, OMW-based cultivation of C. tropicalis involved nitrogen supplementation and pH control in both shake-flask and bioreactor experiments [30]. In the study by Laribi et al., ethanol at 3–7% (v/v) was also used as a controlled stress factor to redirect the metabolism of Y. lipolytica, thereby modulating lipase production, lipid accumulation, and fatty-acid remodeling [39].

A distinction should therefore be made between wastewater-based media, in which wastewater constitutes a major proportion of the cultivation matrix or fermentation-relevant substrates, and wastewater-supplemented media, in which the effluent is incorporated into an otherwise nutrient-rich or synthetic formulation. These approaches are not equivalent in terms of waste valorization. Supplementation with yeast extract, peptone, mineral salts, additional oils, surfactants, or emulsifiers may improve microbial growth and lipase production, but it also increases material costs and may reduce the extent to which the process depends on wastewater-derived nutrients. Consequently, high lipase productivity in a heavily supplemented medium should not, by itself, be interpreted as evidence of a low-cost or resource-efficient wastewater-valorization process.

At the study level, direct use of untreated wastewater was less common than conditioned or supplemented cultivation. Undiluted wastewater was used or evaluated as a cultivation condition in at least six of the 20 studies (30%) [20,21,22,23,26,29]. Thermal sterilization was explicitly reported in 13 studies (65%) [20,21,22,23,24,25,26,29,32,33,34,37,38], while external supplementation with nutrients, carbon sources, lipid inducers, minerals, surfactants, or emulsifiers was used or evaluated in 18 studies (90%) [20,21,22,23,24,25,26,27,28,29,30,31,32,34,35,37,38,39]. One POME-based study explicitly avoided external nutrient supplementation [33], whereas the complete production medium composition was insufficiently reported in another study [36]. These frequencies demonstrate that wastewater conditioning and medium modification were common features of the published processes rather than exceptional interventions.

The principal wastewater-conditioning steps therefore entail clear trade-offs. Dilution can reduce inhibitory loads, viscosity, and solids-related limitations, but it increases water consumption and lowers the concentration of wastewater-derived nutrients. Centrifugation or filtration can improve medium homogeneity and facilitate microbial cultivation, but it introduces additional separation operations and associated energy requirements. Thermal sterilization reduces the risk of microbial contamination and improves experimental reproducibility, but it increases energy demand and may alter wastewater composition. Similarly, pH adjustment can improve microbial growth and enzyme production, but it requires additional chemicals, whereas nutrient or lipid supplementation may enhance productivity at the expense of higher material inputs and reduced reliance on the wastewater itself. These trade-offs should be considered when assessing whether improved biological performance translates into a genuinely resource-efficient wastewater-valorization process.

Overall, the reviewed studies demonstrate that agro-industrial wastewaters can provide carbon, lipids, minerals, and other nutrients that support microbial growth and lipase production. However, wastewater valorization should not automatically be equated with a sustainable process. The environmental and economic performance of such systems depends not only on replacing conventional fermentation substrates with wastewater but also on the material and energy inputs required for dilution, solids removal, sterilization, pH adjustment, nutrient or lipid supplementation, and downstream processing, as well as on the generation and management of residual biomass and other secondary waste streams. Consequently, the sustainability of wastewater-based lipase production should be evaluated based on the overall material, water, energy, and waste balance rather than lipase productivity or wastewater utilization alone.

3.4. Microbial Cultivation Strategies and Lipase Production

The microbial cultivation conditions and lipase-production outcomes reported in the included studies are summarized in

Table 3. Microbial cultivation conditions and lipase-production outcomes in agro-industrial wastewater-based media.

Microorganism	Culture Mode and Scale	Principal Cultivation Conditions	Optimization Strategy	Lipase Localization	Activity-Assay Method	Maximum Lipase Activity	Time to Maximum Activity	Reference
F. solani	Submerged batch culture in 250-mL shake flasks; 50 mL working volume	Undiluted OMW supplemented with a nitrogen source (5 g/L); yeast extract was selected; 1% (v/v) inoculum. Temperature: 30 °C; pH: Initial pH 6; mixing/aeration: 160 rpm; cultivation duration: 5 d.	Comparison of yeast extract, soy peptone, and (NH4)2SO4; no formal statistical design	Extracellular; crude culture supernatant	Potentiometric pH-stat titration of fatty acids released from an olive-oil emulsion at 37 °C. 1 U = 1 µmol fatty acid released per min.	Approximately 14 U/mL	5 d	[20]
Y. lipolytica TISTR 5151	Submerged batch culture in 250-mL shake flasks; 50 mL working volume	Undiluted POME supplemented with (NH4)2SO4 at 1 g N/L; 10% (v/v) seed culture. Temperature: 30 ± 2 °C; pH: Raw pH and pH 5.0–6.0 tested; optimum pH 5.0; mixing/aeration: 140 rpm; cultivation duration: 72 h.	Strain screening followed by comparisons of POME dilution, nitrogen supplementation, and initial pH	Cell-bound lipase	Hydrolysis of palm oil by wet yeast cells; released free fatty acids quantified as palmitic acid. 1 U = 1 µmol free fatty acid released per min.	4081 ± 48 U/L	48 h	[21]
A. ibericus, A. uvarum, and A. niger	Submerged batch culture in 250-mL flasks; 100 mL working volume; static incubation	Undiluted OMW, OMW:nutrient medium 1:1 (v/v), or OMW:winery vinasse 1:1 (v/v); inoculum 2 mL at 107 spores/mL. Temperature: 25 °C; pH: No explicit pH adjustment reported; mixing/aeration: No agitation reported; cultivation duration: 10 d.	Comparative screening of three Aspergillus species and three wastewater-based media; no formal statistical design	Extracellular	Spectrophotometric hydrolysis of p-nitrophenyl butyrate in acetate buffer; absorbance measured at 405 nm. Activity defined by the authors from p-nitrophenol release per min.	1253.7 ± 161.2 U/L (A. ibericus in undiluted OMW)	10 d	[22]
C. rugosa CU1 and Y. lipolytica rM-4A, individually and as a co-culture	Submerged batch culture in shake flasks; single cultures and 1:1 co-culture	Sterile undiluted POME supplemented with 0.5% (w/v) palm oil; no additional nutrient supplementation. Temperature: 30 °C; pH: Raw POME pH used for live-cell cultivation; mixing/aeration: 200 rpm; cultivation duration: 120 h.	Direct comparison of single cultures, co-culture, and separately produced crude enzyme preparations	Extracellular lipase	Spectrophotometric p-nitrophenyl butyrate assay at 37 °C and pH 7.2. Defined by the authors from butyric-acid release per min.	NR for POME-grown cultures; activity was presented mainly in relative terms	Monitored up to 120 h	[23]
F. solani NFCCL 4084	Submerged batch culture in 100-mL shake flasks; 30 mL working volume	POME-based medium containing statistically screened carbon, nitrogen, mineral, oil, and surfactant components; inoculated with a 6-mm mycelial disc. Temperature: 28 °C; pH: Included as a design factor; mixing/aeration: 130 rpm; cultivation duration: 5 d.	Plackett–Burman screening → OFAT range selection → face-centred central composite design	Extracellular; culture filtrate	Spectrophotometric hydrolysis of p-nitrophenyl palmitate; absorbance measured at 410 nm. 1 U = 1 µmol p-nitrophenol released per min.	7.80 U/mL predicted; 6.93 U/mL in the confirmation experiment	5 d	[24]
P. aeruginosa B2290	Shake-flask submerged fermentation; subsequently scaled to a 30-L bioreactor with 20 L medium	POME basal medium; optimized additions: 3% (v/v) inoculum, 4 mM Ca2+, 0.4% (v/v) olive oil, 0.9% (w/v) peptone, and 0.9% Tween 80. Temperature: 30 °C; pH: NR; mixing/aeration: 170 rpm in shake flasks; bioreactor agitation NR; cultivation duration: 96 h.	One-factor-at-a-time optimization of inoculum and medium supplements	Extracellular; cell-free supernatant; spray-dried downstream	Alkali titration of fatty acids released from an olive-oil substrate. Volumetric activity reported in U/mL; dry-extract activity in U/g.	1.327 U/mL in crude supernatant; 28.5 U/g in the spray-dried extract	96 h	[25]
M. guilliermondii	Aerobic submerged batch treatment in shake flasks	Undiluted POME used as the sole carbon source; carbon and nitrogen supplements were also screened; glycerol 1% and yeast extract 0.5% gave the best treatment response. Temperature: 30 °C; pH: Raw POME pH; no adjustment in the principal treatment; mixing/aeration: 150 rpm; cultivation duration: 7 d.	Comparative screening of carbon and nitrogen supplementation; no formal statistical design	Extracellular lipolytic activity inferred from the culture	Qualitative Tween 20–CaCl2 agar assay based on formation of an insoluble calcium–fatty acid precipitate. No quantitative enzyme-activity unit reported.	Qualitative activity only; no U value reported	Treatment evaluated after 7 d	[26]
Mutant B. niacini EMB-5	Shake-flask submerged culture; screening at 50 mL and larger-scale production at approximately 1 L	POME in a mineral-salt basal medium and emulsified with Tween 80. Temperature: 37 °C; pH: Initial pH 5.0; mixing/aeration: 150 rpm; cultivation duration: Up to 72 h; production harvest at 36 h.	Isolation and screening followed by ethidium-bromide mutagenesis; no statistical medium design	Extracellular; cell-free supernatant; subsequently purified	Spectrophotometric hydrolysis of p-nitrophenyl palmitate. Activity reported in U/mL.	38.01 U/mL for mutant EMB-5; 29.23 U/mL for the wild-type strain	36 h for mutant; 42 h for wild type	[27]
Y. lipolytica CBS 6303	Shake-flask batch culture; optimized medium subsequently evaluated in a 5-L bioreactor with 2.5 L working volume	Primary medium contained 30 mL/L VORW plus yeast extract, peptone, phosphate salts, and MgSO4·7H2O; RSM later optimized the medium primarily for microbial oil production. Temperature: 29 °C; pH: Bioreactor maintained at pH 6; mixing/aeration: 200 rpm in flasks; 500 rpm and 1 vvm in bioreactor; cultivation duration: 48 h in flasks; 20 h in bioreactor.	Central composite design/response surface methodology focused mainly on microbial oil production	Culture-derived lipase; enzyme fraction not described in sufficient detail	Titration-based lipase-activity assay. Activity reported in U/mL.	8 U/mL in the primary shake-flask medium	48 h	[28]
M. capitatus JT5	Shake-flask batch culture followed by cultivation in a 2-L stirred-tank bioreactor with 1 L working volume	Undiluted OMW supplemented with yeast extract 2 g/L, NH4Cl 2.8 g/L, and olive oil; bioreactor experiments compared 1 and 3 g/L olive oil. Temperature: 30 °C; pH: Initial pH 6.8 in flasks and 6.1 in the bioreactor; mixing/aeration: 180 rpm in flasks; 100 rpm and 1 vvm in bioreactor; cultivation duration: 120 h in initial flasks; ≥96 h in bioreactor.	Doehlert response-surface design for oxygen availability and NH4Cl concentration; bioreactor scale-up and olive-oil comparison	Extracellular; cell-free culture supernatant	Spectrophotometric hydrolysis of p-nitrophenyl butyrate. 1 U = 1 µmol p-nitrophenol released per min.	3.96 U/mL in the stirred-tank bioreactor with 3 g/L olive oil	96 h; activity was still increasing at the final reported time	[29]
C. tropicalis ATCC 750	Shake-flask batch cultures followed by a 2-L stirred-tank bioreactor with 0.5 L working volume	50% (v/v) OMW with YNB and nitrogen supplementation; bioreactor medium contained OMW-3, YNB, and NH4Cl. Temperature: 30 °C; pH: Initial pH 5 or 7; controlled at pH 5.5 or 7 in the bioreactor; mixing/aeration: 200 rpm in flasks; 200 or 500 rpm and 1 vvm in bioreactor; cultivation duration: 6 d.	Comparison of OMW batches, nitrogen sources, pH, and agitation; PCA used for descriptive interpretation	Extracellular	Spectrophotometric p-nitrophenyl butyrate assay at 37 °C; absorbance measured at 410 nm. 1 U = 1 µmol p-nitrophenol released per min.	203 ± 18 U/L in the bioreactor at pH 5.5 and 200 rpm	Exact time NR; maximum reported for the 6-d cultivation	[30]
Recombinant A. oryzae expressing C. antarctica lipase B	Whole-cell cultivation and simultaneous immobilization in polyurethane biomass-support particles; 500-mL Sakaguchi flasks with 100 mL medium	POME at 2% (w/v), used instead of glucose, with polypeptone, KH2PO4, NaNO3, and MgSO4·7H2O. Temperature: 30 °C; pH: NR; mixing/aeration: 150 oscillations/min; cultivation duration: 96 h.	Direct comparison of POME and additional carbon-source concentrations; no formal statistical design	Immobilized whole-cell, membrane-bound recombinant CALB	Titrimetric hydrolysis of olive oil at 40 °C using immobilized whole-cell particles. Activity reported as U/mg of whole-cell biocatalyst.	2.23 ± 0.02 U/mg	96 h	[31]
M. spicifer AW2 and S. hominis AUP19 co-culture	Submerged batch co-culture in shake flasks; yeast:bacterium inoculum ratio 1:1	Twofold-diluted POME supplemented with 2.08% (v/v) waste frying oil, approximately 1.72% (w/v) (NH4)2SO4, and 0.1% (w/v) gum Arabic. Temperature: 30 ± 2 °C; pH: Initial pH 7.0; mixing/aeration: 150 rpm; cultivation duration: 5 d.	Nutrient screening followed by central composite design/response surface methodology	Predominantly cell-bound; extracellular activity also monitored	Modified cupric-acetate method using palm oil in isooctane; released fatty acids measured at 715 nm. 1 U = 1 µmol palmitic acid released per min; cell-bound activity converted to U/L.	4708.9 U/L	24 h	[32]
M. spicifer AW2 and S. hominis AUP19, individually and as a co-culture	Submerged batch cultures in shake flasks; synthetic 1:1 yeast–bacterium co-culture	POME concentrations of 25%, 50%, and 100% and natural versus pH-adjusted media were evaluated; POME50 at pH 7 was selected. Temperature: 30 ± 2 °C; pH: Natural pH approximately 3.8 or adjusted to pH 7.0; mixing/aeration: 150 rpm; cultivation duration: 120 h.	Strain screening followed by comparisons of POME concentration, initial pH, and mono- versus co-culture	Predominantly cell-bound; extracellular activity also measured	Modified cupric-acetate assay with a palmitic-acid calibration standard. 1 U = 1 µmol palmitic acid released per min.	3860 U/L for the co-culture	24 h	[33]
B. aryabhattai BA03	Submerged batch culture in shake flasks under sterile and non-sterile conditions	Undiluted, 25%, or 50% OMW supplemented with yeast extract 10 g/L. Temperature: 20, 27, and 37 °C tested; pH: Initial pH 6, 7, and 8 tested; optimum pH 8; mixing/aeration: 150 rpm; cultivation duration: Up to 7 d; recommended process 4 d.	Factorial comparisons of wastewater concentration, pH, temperature, and sterilization status	Extracellular; cell-free supernatant	Spectrophotometric hydrolysis of p-nitrophenyl ester substrate at 37 °C; absorbance measured at 410 nm. 1 U = 1 µmol p-nitrophenol released per min.	28.34 ± 1.15 U/mL in 50% non-sterile OMW	4 d	[34]
A. oryzae 1042.72	Submerged batch culture in 250-mL shake flasks; 50 mL working volume	OMW-based medium; optimized additions were glucose 3.49 g/L, yeast extract 5.58 g/L, and CaCl2 0.72 g/L; inoculum 106 spores/mL. Temperature: 30 °C; pH: Initial pH 6; mixing/aeration: 200 rpm; cultivation duration: 72 h.	Plackett–Burman design followed by central composite design/response surface methodology	Extracellular; crude enzyme extract	Titrimetric hydrolysis of an olive-oil emulsion at 37 °C; liberated fatty acids titrated with NaOH. 1 U/mL = 1 µmol fatty acid released per min.	212.53 U/mL	48 h	[35]
Streptomyces sp. SC1	Submerged shake-flask culture	OMW used as the sole carbon source; optimum approximately 6% (v/v) OMW with an inoculum of 1.7 × 107 spores/mL. Temperature: 30 °C; pH: Initial pH 4; mixing/aeration: Agitation NR; cultivation duration: 9 d.	Box–Behnken design/response surface methodology evaluating incubation time, pH, temperature, inoculum size, and OMW concentration	Extracellular; culture supernatant	Titrimetric hydrolysis of an olive-oil/gum-arabic emulsion at 37 °C. Activity reported in U/mL from NaOH consumption.	8.4 U/mL measured; 8.82 U/mL predicted by the model	9 d	[36]
M. spicifer AW2 + S. hominis AUP19	Submerged batch co-culture in 250-mL shake flasks; yeast:bacterium inoculum ratio 1:1	POME50 supplemented with 1.23% (w/v) (NH4)2SO4; 10% (v/v) inoculum; 30 ± 2 °C; initial pH 7.0; 150 rpm; time-course cultivation up to 144 h	Plackett–Burman screening followed by RSM-CCD; ammonium sulfate concentration, incubation time, and initial pH optimized	Predominantly cell-bound; extracellular activity also monitored	Modified cupric-acetate assay; 1 U = activity releasing 1 µmol palmitic acid per min	4103 U/L	24 h	[37]
Pichia sp. Y1440 and T. coremiiforme Y771	Submerged batch culture on a rotary shaker	Pure and nutrient-enriched POME were initially compared; subsequent OFAT/RSM experiments varied POME concentration and pH. Temperature: Room temperature; pH: pH 5–8 examined; RSM optima approximately pH 5 for Pichia and pH 8 for T. coremiiforme; mixing/aeration: 150 rpm; cultivation duration: 72 h.	OFAT followed by response surface methodology	Extracellular; centrifuged culture extract	Spectrophotometric hydrolysis of p-nitrophenyl palmitate at 410 nm. Activity reported in U/mL.	Observed OFAT maxima: 11.6 U/mL (Pichia) and 30 U/mL (T. coremiiforme)	72 h	[38]
Y. lipolytica L2	Submerged batch culture in 250-mL shake flasks; 100 mL working volume	YP medium containing OMW at 200 g/L; ethanol added progressively 24 h after inoculation at 0, 3, 5, or 7% (v/v). Temperature: 29 ± 1 °C; pH: NR; mixing/aeration: 180 rpm; cultivation duration: 96 h.	One-factor comparison of ethanol-stress levels and incubation times; OMW and waste cooking oil studied as separate carbon-source systems	Extracellular; cell-free supernatant	Spectrophotometric hydrolysis of p-nitrophenyl laurate at 37 °C; absorbance measured at 410 nm. 1 U = 1 µmol p-nitrophenol released per min.	0.55 ± 0.11 U/mL in OMW medium with 5% ethanol	48 h	[39]

Note: Activity values should not be interpreted as a ranking of microorganisms or wastewater substrates because the studies used different enzyme fractions, assay substrates, reaction conditions, unit definitions, and expression bases. Reported values and units were retained as presented by the original authors. Abbreviations: NR, not reported; OMW, olive mill wastewater; POME, palm oil mill effluent; VORW, vegetable oil refinery wastewater; OFAT, one factor at a time; RSM, response surface methodology; YNB, yeast nitrogen base; YP, yeast extract–peptone medium.

. Most experiments were conducted as submerged batch cultures in shake flasks, typically at temperatures of 25–30 °C and agitation rates of approximately 130–200 rpm [20,21,23,24,25,26,28,29,30,32,33,34,35,36,37,39]. Static cultivation was used in one study involving Aspergillus spp. [22], whereas B. niacini was cultivated at 37 °C [27]. The tested initial pH values spanned a broad range, from acidic conditions around pH 4 to alkaline conditions around pH 8, reflecting differences in wastewater composition and microbial physiology [24,27,30,33,34,35,36,37,38]. Cultivation times varied substantially, with maximum lipase activities reached within 24–48 h in several yeast and yeast–bacterium systems [21,28,32,33,37,39], whereas longer cultivation periods of up to 9–10 days were reported for Streptomyces sp. and filamentous fungi [22,36].

Although shake-flask cultivation predominated, only a few studies advanced to controlled bioreactor systems. Larger-scale cultivation was reported for P. aeruginosa in a 30-L bioreactor [25], Y. lipolytica in a 5-L reactor [28], and M. capitatus and C. tropicalis in 2-L stirred-tank bioreactors [29,30]. These studies examined the scale-up of wastewater-based microbial processes beyond the shake-flask scale. However, the bioreactor experiments often addressed multiple outputs, including biomass formation, microbial lipid accumulation, wastewater remediation, and enzyme production, and lipase activity was not always the primary process response [25,28,29,30].

The approaches used to improve enzyme production also varied considerably. Several studies relied on direct comparisons or one-factor-at-a-time experiments to examine wastewater concentration, pH, temperature, inoculum level, nitrogen supplementation, or the addition of lipid inducers [20,21,23,25,26,31,33,34,39]. More structured optimization strategies included Plackett–Burman screening, central composite design, Box–Behnken design, Doehlert design, and response surface methodology [24,28,29,32,35,36,37,38]. In the later M. spicifer–S. hominis study, Plackett–Burman screening was followed by RSM-CCD to optimize cell-bound lipase production, biomass formation, and COD/O&G removal [37]. Such multivariable designs allowed the influence of, and in several studies the interactions among, medium and process variables to be evaluated simultaneously.

Most of the microorganisms investigated produced extracellular lipases, which were evaluated in culture supernatants or filtrates [20,22,23,24,25,27,29,30,34,35,36,38,39]. In contrast, Y. lipolytica TISTR 5151 and the yeast–bacterium systems based on M. spicifer AW2 and S. hominis AUP19 were primarily evaluated as sources of cell-bound lipases [21,32,33,37]. Rachmadona et al. used a distinct whole-cell approach in which recombinant A. oryzae expressing C. antarctica lipase B was cultivated and simultaneously immobilized within polyurethane biomass-support particles [31]. These differences are technologically relevant because cell-bound and immobilized lipases can be used directly as whole-cell biocatalysts, whereas extracellular enzymes generally require separation from microbial biomass and, depending on the intended application, may require further concentration or purification.

The reported maximum lipase activities varied widely across the included studies. Volumetric extracellular activities, expressed in U/mL, ranged from approximately 14 U/mL for F. solani [20] to 212.53 U/mL for A. oryzae [35], with values of 38.01 U/mL for mutant B. niacini EMB-5 [27], 28.34 U/mL for B. aryabhattai BA03 [34], 8.4 U/mL for Streptomyces sp. SC1 [36], and 0.55 ± 0.11 U/mL for ethanol-stressed Y. lipolytica cultivated in OMW [39]. Cell-bound activities were commonly reported in U/L, including 4081 ± 48 U/L for Y. lipolytica TISTR 5151 [21], 3860 U/L for the M. spicifer–S. hominis co-culture in POME [33], 4708.9 U/L for the related POME–waste frying oil system [32], and 4103 U/L for the subsequently optimized POME50-based co-culture [37]. The recombinant immobilized whole-cell system was reported on a mass basis, reaching 2.23 ± 0.02 U/mg [31]. One study reported only qualitative lipolytic activity [26], whereas another did not provide an absolute activity value for the POME-grown cultures [23].

Unfortunately, a direct comparison of reported lipase activities is inappropriate because the studies differed substantially in enzyme localization, assay substrate, reaction pH and temperature, unit definition, and the basis for expressing activity. Natural oils, including olive and palm oil, were used in titrimetric or colorimetric assays in several studies [20,21,25,31,32,33,35,36,37], whereas others employed p-nitrophenyl esters such as butyrate, palmitate, or laurate [22,23,24,27,29,30,34,38,39]. Moreover, activities were expressed as U/mL, U/L, U/g, or U/mg, depending on the enzyme preparation and assay system. Consequently, the activity values presented in Table 3 should be interpreted as study-specific indicators of process performance rather than as a direct ranking of microorganisms or wastewater substrates.

3.5. Wastewater-Treatment Performance

Wastewater-treatment outcomes associated with microbial lipase production are summarized in

Table 4. Wastewater-treatment outcomes associated with microbial lipase production in agro-industrial wastewater-based media.

Wastewater	Microorganism	Assessment Configuration	Treatment Conditions and Duration	COD/Organic-Load Reduction	Oil, Grease, or Lipid Reduction	Phenolic-Compound Reduction	Other Reported Changes	Integration with Lipase Production	Reference
OMW	F. solani	Simultaneous cultivation, lipase production, and OMW treatment	Undiluted OMW supplemented with yeast extract; 30 °C, 160 rpm, 5 d	24.1% reduction after 5 d	100% oil removal after 5 d	Maximum 46% removal	Total sugars decreased by 68%; maximum sugar reduction was reported on day 2	Yes—treatment occurred during lipase-producing fungal growth	[20]
POME	Y. lipolytica TISTR 5151	Simultaneous cultivation, cell-bound lipase/lipid production, and COD removal	Undiluted POME with ammonium sulfate; 30 ± 2 °C, 140 rpm, 72 h; initial pH varied	Maximum 93.4 ± 5.1% at initial pH 5.0; 92.4 ± 6.7% at raw pH 4.3; 81.2 ± 7.6% with ammonium sulfate at pH 5.5	NR	NR	Final pH increased in most cultures	Yes—COD removal was measured in the same cultures used for lipase and lipid production	[21]
OMW; OMW–winery vinasse mixture	A. ibericus, A. uvarum, and A. niger	Simultaneous submerged fungal treatment and enzyme production	Undiluted OMW, twofold-diluted OMW with nutrients, or OMW:vinasse 1:1; 25 °C, static, 10 d	Best reduction: 66.9 ± 2.0% with A. niger in OMW:vinasse; 65.1 ± 3.1% with A. uvarum in OMW:vinasse	NR	Best reduction: 43.3 ± 0.8% with A. uvarum in OMW:vinasse	Maximum colour reduction with A. uvarum in OMW:vinasse: 56.1 ± 0.6% at 395 nm and 71.6 ± 1.5% at 525 nm	Yes—lipolytic, proteolytic, and tannase activities were detected during bioremediation	[22]
POME	C. rugosa CU1 and Y. lipolytica rM-4A, alone and in co-culture	Both simultaneous live-cell treatment and separate application of pre-produced crude lipase/laccase	Live cells: sterile undiluted POME, 30 °C, 200 rpm, up to 120 h. Enzyme arm: separate crude enzyme preparations applied to diluted non-sterile or undiluted sterile POME	Live-cell co-culture: 60.30 ± 4.89% after 120 h. Separate enzyme arm: 64.1% in non-sterile diluted POME and 39.5% in sterile undiluted POME using lipase + laccase	Live-cell co-culture: 98.53 ± 1.70% triglyceride degradation after 120 h. Separate C. rugosa lipase: 93% after 48 h	Live-cell Y. lipolytica: 36.1% removal after 96 h; C. rugosa 28.4%; co-culture 31.7%. Separate enzyme arm: maximum 17.5% with laccase	Reducing sugars decreased during live-cell cultivation; no decolorization was observed	Partly—live-cell results were simultaneous, but crude-enzyme treatment used enzymes produced previously in synthetic media	[23]
POME	F. solani NFCCL 4084	Lipase-production optimization only	POME-based shake-flask fermentation; 28 °C, 130 rpm, 5 d	NR	NR	NR	No wastewater-quality parameters were monitored after cultivation	No treatment outcome assessed	[24]
POME	P. aeruginosa B2290	Lipase-production and downstream spray-drying study	Shake-flask and 30-L bioreactor cultivation; 30 °C, up to 96 h	NR	NR	NR	No post-cultivation wastewater-treatment parameters were reported	No treatment outcome assessed	[25]
POME	M. guilliermondii	Simultaneous aerobic bioremediation and extracellular lipolytic activity	Undiluted POME; 30 °C, 150 rpm, 7 d; nutrient supplementation was also evaluated	72% reduction	92.4% oil-and-grease removal	Disappearance of several detected phenolic/hydrocarbon compounds by GC–MS; total phenolic removal not quantified	Total N 49.2%; ammoniacal N 45.1%; TOC 46.6%; phosphate 60.6% removal. pH increased from 4.6 to 8.2; seed-germination index increased from 59.1 to 74.3%	Yes—pollutant removal and extracellular lipolytic activity occurred in the same aerobic treatment	[26]
POME	Mutant B. niacini EMB-5	Lipase production, purification, and biochemical characterization	POME-containing submerged fermentation; 37 °C, 150 rpm	NR	NR	NR	No wastewater-treatment indicators were measured	No treatment outcome assessed	[27]
VORW	Y. lipolytica CBS 6303	Simultaneous wastewater valorization, microbial-oil production, and COD reduction	Optimized shake-flask and 5-L bioreactor cultures; 29 °C; COD evaluated after fermentation	Approximately 80% reduction after 20 h	NR as a treatment endpoint	NR	Microbial oil and lipid-rich biomass were produced; no additional effluent-quality indicators were reported	Yes—COD reduction occurred as part of the wastewater-based fermentation	[28]
OMW	M. capitatus JT5	Lipase-production optimization and bioreactor scale-up	Undiluted OMW; shake flasks and 2-L stirred-tank reactor	NR	NR	NR	Initial OMW composition was characterized, but changes after cultivation were not reported	No quantitative treatment outcome assessed	[29]
OMW	C. tropicalis ATCC 750	Simultaneous OMW bioconversion, pollutant removal, and production of lipase, protease, biomass, and lipids	50% OMW; shake flasks and 2-L bioreactor; 30 °C, 200 or 500 rpm, 1 vvm, 6 d	Maximum 68.2 ± 6.0% in the bioreactor at pH 7 and 500 rpm	NR	Maximum 39.1 ± 0.2% in the bioreactor at pH 7 and 500 rpm	Reducing sugars 86.4 ± 0.7%; TOC 58.7 ± 4.1%; total N maximum 52.7 ± 3.7% reduction	Yes—wastewater-component consumption and enzyme production were monitored in the same cultures	[30]
POME	Recombinant A. oryzae expressing C. antarctica lipase B	Whole-cell lipase production/immobilization followed by a separate biodiesel application	POME-based culture for 96 h; immobilized cells subsequently used to esterify POME-derived lipids	NR	No removal percentage reported as a wastewater-treatment endpoint	NR	No conventional wastewater-remediation outcome was assessed	No conventional remediation assessment; downstream valorization was evaluated separately	[31]
POME blended with WFO for production; POME for subsequent treatment	M. spicifer AW2 + S. hominis AUP19	Combined production-medium valorization plus separate post-production application of harvested cell-bound lipases	Production: optimized POME50–WFO medium, 24 h. Separate treatment: harvested wet cells added at 10% (v/v) to heat-treated POME for 72 h	Separate treatment: 73.6% removal in twofold-diluted POME after 72 h	Production fermentation: 58.3% O&G removal after 24 h. Separate treatment: 73.3% O&G removal after 72 h	NR	During separate treatment, pH increased from approximately 3.8 to 6.7–7.9 depending on POME concentration	Both—O&G removal occurred during production, whereas the highest COD/O&G treatment values came from a separate application	[32]
POME	M. spicifer AW2 + S. hominis AUP19	Simultaneous co-culture growth, cell-bound lipase production, and POME treatment	POME50 adjusted to pH 7.0; 30 ± 2 °C, 150 rpm, 120 h	75.9 ± 2.8% with the co-culture; concentration decreased from 29,747 to 7169 mg/L	80.1 ± 1.3% O&G removal with the co-culture; concentration decreased from 5562 to 1107 mg/L	NR	Final pH 8.17 ± 0.07 under the optimal POME50 co-culture condition	Yes—treatment and cell-bound lipase production occurred in the same POME culture	[33]
OMW	B. aryabhattai BA03	Simultaneous non-sterile/sterile cultivation, lipase production, and reductions in OMW constituents	Treatment outcomes quantified in 25% OMW at initial pH 8 and 27 °C; 150 rpm, 4 d	NR	NR	21.19 ± 0.70% in sterile medium and 27.81 ± 0.33% in non-sterile medium	Sugars metabolized: 66.41 ± 1.45% in sterile medium and 82.37 ± 0.13% in non-sterile medium; pH increased to >8	Yes—phenol and sugar reductions accompanied lipase production	[34]
OMW	A. oryzae 1042.72	Statistical optimization of lipase production	OMW-based shake-flask cultivation; 30 °C, 200 rpm, 72 h	NR	NR	NR	No before/after wastewater-quality measurements were reported	No treatment outcome assessed	[35]
OMW	Streptomyces sp. SC1	Lipase-production optimization with OMW described as undergoing biodegradation	Approximately 6% OMW; 30 °C, initial pH 4, 9 d	NR	NR	NR	The authors stated that OMW biodegradation was followed, but no quantitative pollutant-removal result was presented	Potentially simultaneous, but treatment performance was not reportable	[36]
POME	M. spicifer AW2 + S. hominis AUP19	Simultaneous co-culture growth, cell-bound lipase production, biomass formation, and POME bioremediation	POME50 + 1.23% (w/v) (NH4)2SO4; initial pH 7.0; 30 ± 2 °C; 150 rpm; optimized confirmation at 140 h	84.5 ± 1.4%	87.9 ± 2.3% O&G removal	NR	Cell biomass 13.8 ± 1.2 g/L; CBL activity 3391 ± 83 U/L under the combined optimized condition	Yes—CBL production, biomass formation, COD removal, and O&G removal were evaluated in the same cultivation process	[37]
POME	Pichia sp. Y1440 and T. coremiiforme Y771	Lipase-production optimization with carbohydrate consumption monitored	Pure or nutrient-enriched POME; room temperature, 150 rpm, 72 h	NR	NR	NR	Pure POME carbohydrate concentration decreased from 119.1 to 36.3 mg/L for Pichia sp. and from 122.7 to 51.9 mg/L for T. coremiiforme between 24 and 72 h; smaller declines occurred in enriched POME	Limited—substrate consumption was monitored during lipase production, but standard wastewater-treatment endpoints were not assessed	[38]
OMW	Y. lipolytica L2	Lipase/lipid production under ethanol stress	OMW-containing YP medium; 29 ± 1 °C, 180 rpm, 96 h; ethanol 0–7% (v/v)	NR	NR as wastewater-treatment performance	NR	The study evaluated microbial lipid accumulation, fatty-acid remodeling, lipase activity, and oxidative stability rather than final effluent quality	No treatment outcome assessed	[39]

Note: NR indicates not reported. A missing value means that the parameter was not quantified and does not imply absence of treatment. Where a publication included both microbial cultivation and a separate application of previously produced cells or enzymes, these outcomes are explicitly distinguished. The highest values reported for different pollutants may originate from different strains or operating conditions and should not be treated as a single combined process result. Changes in color, pH, germination index, or other auxiliary indicators are reported as complementary process outcomes and should not be directly compared with conventional wastewater-quality parameters such as COD, BOD, or O&G. Abbreviations: OMW, olive mill wastewater; POME, palm oil mill effluent; VORW, vegetable oil refinery wastewater; WFO, waste frying oil; CBL, cell-bound lipase; COD, chemical oxygen demand; TOC, total organic carbon; O&G, oil and grease.

. The scope of environmental assessment varied widely across the included studies. Some authors quantified several indicators, including COD, oil and grease, phenolic compounds, sugars, nitrogen, phosphorus, TOC, color, and phytotoxicity [20,21,22,23,26,30,33,34,37], whereas others focused primarily on lipase production and did not assess wastewater quality after cultivation [24,25,27,29,31,35,36,39]. One additional study monitored carbohydrate consumption during POME-based cultivation but did not evaluate conventional wastewater-treatment indicators such as COD or O&G [38]. Consequently, the absence of a reported removal value should be interpreted as a lack of measurement rather than as evidence of no treatment effect. Throughout this section, the terms reduction, removal, degradation, and consumption are used according to the analytical endpoint and terminology reported in the original study and are not treated as interchangeable.

COD reduction was the most commonly reported measure of organic-load removal. The highest reported value was observed during cultivation of Y. lipolytica TISTR 5151 in POME, reaching 93.4 ± 5.1% at an initial pH of 5.0 [21]. The optimized M. spicifer AW2–S. hominis AUP19 co-culture subsequently achieved 84.5 ± 1.4% COD removal in POME50 [37], whereas Y. lipolytica cultivated in vegetable oil refinery wastewater showed approximately an 80% decrease after 20 h [28]. In an earlier POME-based study, the M. spicifer–S. hominis co-culture reduced COD by 75.9 ± 2.8% [33]. Other reported reductions included 72% for M. guilliermondii [26], 68.2 ± 6.0% for C. tropicalis in a controlled bioreactor [30], and up to 66.9 ± 2.0% during fungal treatment of an OMW–winery vinasse mixture [22]. The co-culture of C. rugosa and Y. lipolytica rM-4A reduced COD by 60.30 ± 4.89% in undiluted POME [23], whereas F. solani achieved a 24.1% reduction in undiluted OMW [20].

Removal of the lipid fraction was particularly pronounced in several oil-rich wastewater systems. Complete oil removal was reported during cultivation of F. solani in OMW [20], whereas the C. rugosa–Y. lipolytica co-culture degraded 98.53 ± 1.70% of triglycerides in POME after 120 h [23]. M. guilliermondii removed 92.4% of the oil-and-grease fraction [26]. In the M. spicifer–S. hominis systems, O&G removal reached 80.1 ± 1.3% in POME50 without external nutrient supplementation [33] and 87.9 ± 2.3% in the subsequently optimized POME50 medium supplemented with ammonium sulfate [37]. These findings are consistent with concurrent microbial lipolytic activity and removal of hydrophobic wastewater constituents. However, the reported endpoints were not equivalent, because some studies measured total oil and grease, whereas others quantified triglycerides or selected lipid fractions. Moreover, the observed reductions cannot be attributed exclusively to lipase activity because microbial uptake, biomass formation, and other metabolic processes occurred simultaneously.

Phenolic-compound removal was primarily investigated in OMW- and POME-based processes. The highest reported reduction was approximately 46% during F. solani cultivation in OMW [20]. Treatment of the OMW–winery vinasse mixture with A. uvarum decreased phenolic compounds by 43.3 ± 0.8% [22], whereas C. tropicalis achieved up to 39.1 ± 0.2% removal in the bioreactor [30]. In POME, Y. lipolytica rM-4A removed 36.1% of the total phenolic content, compared with 31.7% for the yeast co-culture and 28.4% for C. rugosa alone [23]. B. aryabhattai BA03 reduced phenolic compounds by 27.81 ± 0.33% under non-sterile conditions and by 21.19 ± 0.70% in sterile OMW-based medium [34]. In the study involving M. guilliermondii, GC–MS revealed the disappearance of several phenolic and hydrocarbon compounds, but the quantitative percentage of total-phenol removal was not reported [26]. A reduction in total phenolic-compound concentration should not, however, be interpreted as evidence of detoxification, because it does not establish the toxicity of residual or newly formed metabolites. Similarly, the disappearance of individual compounds detected by GC–MS should be distinguished from quantitative removal of total phenolic compounds.

Several studies also demonstrated broader changes in wastewater composition. F. solani reduced total sugars in OMW by 68% [20], while fungal treatment of combined OMW and winery vinasse decreased wastewater color by up to 56.1 ± 0.6% at 395 nm and 71.6 ± 1.5% at 525 nm [22]. Treatment of POME with M. guilliermondii reduced total nitrogen by 49.2%, ammoniacal nitrogen by 45.1%, TOC by 46.6%, and phosphate by 60.6%; it also increased the seed-germination index from 59.1 to 74.3%, suggesting a partial reduction in phytotoxicity [26]. In the bioreactor study with C. tropicalis, reductions reached 86.4 ± 0.7% for reducing sugars, 58.7 ± 4.1% for TOC, and 52.7 ± 3.7% for total nitrogen [30]. Carbohydrate consumption was also observed during POME-based cultivation of Pichia sp. and T. coremiiforme, although conventional treatment indicators such as COD or O&G were not measured [38]. Changes in color, seed germination, pH, or other auxiliary indicators provide complementary information on process effects but are not directly comparable with conventional wastewater-quality parameters such as COD, BOD, or O&G.

An important distinction should be made between pollutant removal during microbial cultivation and the separate application of previously produced cells or enzymes. In the study by Theerachat et al. [23], the live-cell co-culture simultaneously produced enzymes and treated POME, whereas additional experiments used crude lipase and laccase preparations produced in synthetic media. Similarly, in the study by Fibriana et al. [32], O&G removal occurred during production of cell-bound lipases (CBLs) in the blended POME–waste frying oil medium, but the highest COD and O&G removal values were obtained in a separate treatment step in which harvested wet cells were added to a new batch of heat-treated POME. In contrast, the subsequent POME50-based co-culture study integrated CBL production, biomass formation, and COD and O&G removal within the same cultivation process [37]. Under the combined optimized condition, COD and O&G removal reached 84.5 ± 1.4% and 87.9 ± 2.3%, respectively [37]. The time-course data also revealed a temporal difference among these outputs: CBL activity peaked at 24 h, whereas COD and O&G removal continued to increase for approximately six days [37]. These distinctions are important because treatment achieved during enzyme-producing fermentation represents direct wastewater valorization, whereas post-production application of harvested cells or enzymes constitutes a separate bioremediation step.

The available evidence indicates that microbial cultivation in agro-industrial wastewaters can support lipase production while contributing to reductions in organic load and selected wastewater constituents. However, treatment performance was not consistently evaluated, and only a limited number of studies assessed multiple wastewater-quality parameters. Moreover, treated effluents were rarely compared with applicable discharge or reuse standards, and residual toxicity, microbial safety, and suitability for reuse were seldom investigated. Therefore, reductions in COD, O&G, or other bulk parameters should be interpreted as evidence of partial bioremediation and wastewater valorization rather than as validation of complete wastewater treatment.

3.6. Lipase Recovery, Characterization and Applications

The downstream processing, biochemical characterization, and applications of the produced lipases are summarized in

Table 5. Downstream processing, biochemical characterization, and applications of lipases produced in agro-industrial wastewater-based media.

Microorganism	Lipase Form	Recovery or Downstream Processing	Purification or Immobilization	Biochemical Characterization	Application Tested	Main Application or Downstream Outcome	Reference
F. solani	Extracellular lipase in crude culture supernatant	Culture liquid used as a crude enzyme source; no concentration or formulation reported	None reported	No biochemical characterization of the enzyme was performed	No separate post-production application; lipase production occurred during OMW biotreatment	The cultivation combined lipase production with 100% oil removal, 24.1% COD reduction, and partial removal of phenols and sugars	[20]
Y. lipolytica TISTR 5151	Cell-bound lipase associated with wet yeast cells	Wet cells harvested by centrifugation and used directly; drying and lipid extraction were avoided	Natural cell-surface immobilization; no enzyme purification	Optimum pH 7.5 and temperature 30 °C; greatest pH stability around pH 6–7; high thermal stability at 25–35 °C, with rapid activity loss at higher temperatures	Direct extraction and transesterification of intracellular yeast lipids using methanol in a solvent-free whole-cell process	FAME yield reached 32.1% after 24 h and 40.9% after 72 h	[21]
A. ibericus, A. uvarum, and A. niger	Extracellular lipolytic activity in crude culture liquid	Culture liquid analyzed directly; no concentration or formulation reported	None reported	No biochemical characterization of the lipase was performed	No separate enzyme application; lipolytic activity was generated during fungal bioremediation	The process simultaneously produced hydrolytic enzymes and reduced COD, phenols, and colour, particularly in the OMW–vinasse mixture	[22]
C. rugosa CU1 and Y. lipolytica rM-4A	Extracellular lipase in live POME cultures; separate crude enzyme preparations were also examined	Culture broths were centrifuged and filtered through a 0.2-µm membrane to obtain crude extracellular enzyme preparations	None reported	No biochemical characterization beyond activity determination with p-nitrophenyl butyrate	Live mono- and co-cultures were used for POME treatment; a separate treatment arm applied crude lipase and laccase produced previously in synthetic media	The live co-culture degraded 98.53 ± 1.70% of triglycerides and reduced COD by 60.30 ± 4.89%; the separate C. rugosa lipase preparation degraded 93% of triglycerides after 48 h	[23]
F. solani NFCCL 4084	Extracellular lipase in crude culture filtrate	Culture filtrate used directly for the activity assay	None reported	No systematic biochemical characterization; the study focused on production optimization and described the enzyme as alkaline/halophilic	No application evaluated in the reported experiments	The optimized process increased crude lipase activity to 7.80 U/mL, with 6.93 U/mL obtained in the confirmation experiment	[24]
P. aeruginosa B2290	Extracellular lipase in cell-free supernatant and spray-dried extract	Fermentation broth centrifuged to obtain cell-free supernatant; supernatant subsequently spray-dried	No chromatographic purification or immobilization	No biochemical characterization of pH, temperature, stability, kinetics, or substrate specificity	No catalytic application tested	Spray drying yielded 15.643 g of dry extract from 500 mL supernatant; activity was 28.5 U/g and specific activity 2.417 U/g protein	[25]
M. guilliermondii	Extracellular lipolytic activity detected qualitatively	No enzyme recovery beyond culture-based detection	None reported	No quantitative enzyme characterization; activity was demonstrated using a Tween 20–CaCl2 plate assay	No separate lipase application; lipolytic activity accompanied aerobic POME bioremediation	The live treatment removed 92.4% O&G and 72% COD and reduced nitrogen, TOC, phosphate, hydrocarbons, and phytotoxicity	[26]
Mutant B. niacini EMB-5	Purified extracellular alkaline lipase	Cells removed by centrifugation; enzyme recovered from cell-free supernatant	60% ammonium-sulfate precipitation, dialysis, DEAE-Sephadex A-50 ion exchange, and Sephadex G-100 gel filtration; 19.05-fold purification, 7.18% yield, 641.42 U/mg; apparent molecular mass 59 kDa	Optimum pH 9.0 and temperature 40 °C; alkaline stability at pH 8–11; substantial thermal stability up to 90 °C; Ca2+, Na+, K+, Mn2+, and Al3+ stimulated activity, whereas Cu2+, Hg2+, EDTA, and β-mercaptoethanol inhibited it; Km 6.1 mg/mL and Vmax 59 µmol/min/mg	Hydrolysis of different natural oils	Relative hydrolytic activity was highest for olive oil (96%), followed by vegetable oil (92%), palm oil (89%), corn oil (78%), soybean oil (70%), and canola oil (61%)	[27]
Y. lipolytica CBS 6303	Culture-derived lipase reported as volumetric activity	No downstream recovery procedure described for the lipase	None reported	No biochemical characterization of the lipase	No application of the produced lipase was evaluated	The study focused on simultaneous production of lipase, microbial oil, and lipid-rich biomass and on COD reduction	[28]
M. capitatus JT5	Extracellular lipase in cell-free culture supernatant	Culture samples centrifuged and supernatant used as the crude enzyme source	None reported	No biochemical characterization; the study focused on production optimization and bioreactor scale-up	No catalytic application evaluated	Bioreactor cultivation increased extracellular activity to 3.96 U/mL in undiluted OMW supplemented with olive oil	[29]
C. tropicalis ATCC 750	Extracellular lipase in crude culture broth	Activity measured in culture samples; no concentration or formulation reported	None reported	No enzyme-focused biochemical characterization	No separate lipase application; production was integrated with OMW bioconversion	The process produced lipase and protease while reducing COD, phenols, sugars, TOC, and nitrogen and generating biomass and microbial lipids	[30]
Recombinant A. oryzae expressing C. antarctica lipase B (r-CALB)	Immobilized recombinant whole-cell lipase	Whole cells cultivated and retained within polyurethane biomass-support particles; particles recovered as the catalyst	Simultaneous cell growth and immobilization in polyurethane foam; no enzyme purification	SEM confirmed cell immobilization; hydrolytic activity reached 2.23 ± 0.02 U/mg in POME medium; activity decreased to 1.59 U/mg after ten cycles, corresponding to 71.47% residual activity	Ethanolysis/esterification of POME to fatty acid ethyl esters; comparison with commercial Novozym 435; repeated-batch reuse	FAEE content increased to 97.52 ± 0.21%; more than 95% FAEE was maintained after the tenth batch, and the catalyst showed a better conversion rate than Novozym 435 under the tested conditions	[31]
M. spicifer AW2 + S. hominis AUP19	Mixed cell-bound lipases in wet co-culture biomass	Cells harvested from the optimized broth by centrifugation and used directly as wet whole-cell catalysts	Natural cell-bound immobilization; no enzyme purification	Hydrolytic activity quantified, but no systematic pH, temperature, kinetic, or stability characterization	Solvent-free esterification of oleic acid, transesterification of palm oil, and separate POME bioremediation	FAME reached 76.2 ± 0.1% in esterification and 87.7 ± 0.5% in transesterification; separate whole-cell treatment removed 73.3% O&G and 73.6% COD	[32]
M. spicifer AW2 + S. hominis AUP19	Mixed cell-bound lipases in wet yeast–bacterium biomass	Wet cells harvested by centrifugation and used directly	Natural cell-bound immobilization; no enzyme purification	Hydrolytic, esterification, and transesterification activities were evaluated; no broader biochemical characterization	Solvent-free esterification of oleic acid and transesterification of palm oil; simultaneous POME treatment was also assessed	After 72 h, mixed cells produced 73.5 ± 3.3% FAME from oleic acid and 82.5 ± 0.3% from palm oil, outperforming both monocultures	[33]
B. aryabhattai BA03	Extracellular lipase in cell-free supernatant	Cells removed and crude supernatant used for the activity assay; no further processing reported	None reported	No biochemical characterization of the produced enzyme	No separate lipase application; production accompanied the reduction of sugars and phenols in OMW	The study established the feasibility of producing lipase in non-sterile OMW-based medium but did not evaluate a recovered enzyme preparation	[34]
A. oryzae 1042.72	Extracellular lipase in crude enzyme extract	Crude enzyme preparation used directly for activity determination	None reported	No biochemical characterization; work was limited to statistical optimization of production	No application evaluated	Optimized medium increased crude activity from 110.21 to 212.53 U/mL	[35]
Streptomyces sp. SC1	Extracellular lipase in culture supernatant	Culture supernatant used as the crude enzyme source	None reported	No biochemical characterization; production variables were optimized by response surface methodology	No application evaluated	The measured and model-predicted activities were approximately 8.4 and 8.82 U/mL, respectively	[36]
M. spicifer AW2 + S. hominis AUP19	Mixed CBLs associated with yeast–bacterium biomass	Cells harvested by centrifugation and used directly as wet whole-cell catalysts; for storage-stability testing, the CBL preparation was air-dried for 1 h and stored at 4 °C or room temperature for 5 weeks	No enzyme purification or external immobilization; lipase remained cell-bound	Storage stability evaluated for 5 weeks; approximately 80% residual activity at 4 °C and 60% at room temperature after 5 weeks	Solvent-free esterification of oleic acid and transesterification of palm oil using fresh and stored CBLs	Fresh CBLs yielded 78.1% FAME from oleic acid and 86.8% from palm oil after 72 h; after 5 weeks at 4 °C, yields remained 77.3% and 86.4%, respectively	[37]
Pichia sp. Y1440 and T. coremiiforme Y771	Extracellular lipase in centrifuged crude culture extract	Culture broth centrifuged and supernatant used directly	None reported	No biochemical characterization of the enzymes	No catalytic application evaluated	The work focused on optimizing POME concentration and pH for crude lipase production	[38]
Y. lipolytica L2	Extracellular lipase in cell-free supernatant	Culture samples centrifuged and supernatant used for activity measurement; no enzyme formulation reported	None reported	No biochemical characterization of the lipase; ethanol-stress effects on activity were assessed	No application of the produced lipase was tested	The study focused on stress-modulated lipase secretion and the production and characterization of intracellular microbial lipids	[39]

A statement that no purification, characterization, or application was reported reflects the scope of the original study and does not imply that the enzyme lacks such properties. Whole-cell systems contain naturally cell-bound or engineered immobilized lipases and therefore avoid conventional enzyme purification. Abbreviations: NR, not reported; OMW, olive mill wastewater; POME, palm oil mill effluent; VORW, vegetable oil refinery wastewater; WFO, waste frying oil; FAME, fatty acid methyl esters; FAEE, fatty acid ethyl esters; CBL, cell-bound lipase; COD, chemical oxygen demand; O&G, oil and grease.

. Most studies assessed lipase activity directly in crude culture supernatants, culture filtrates, or microbial biomass without further concentration, purification, or formulation [20,22,23,24,26,28,29,30,34,35,36,38,39]. In these investigations, the primary objective was typically to confirm lipase production, optimize cultivation conditions, or assess wastewater bioconversion. Only a limited number of studies extended the process to enzyme drying, multistep purification, whole-cell immobilization, storage-stability evaluation, or practical catalytic applications [21,25,27,31,32,33,37].

For extracellular lipase, downstream processing was typically limited to biomass removal by centrifugation or filtration, followed by direct analysis or use of crude supernatants or filtrates [20,22,23,24,27,29,34,35,36,38,39]. Most studies did not report enzyme recovery yield, concentration losses, formulation, storage stability, or long-term catalytic performance. More advanced downstream processing was reported only occasionally. The P. aeruginosa B2290 lipase preparation was spray-dried [25], whereas the most extensive conventional purification was reported for the extracellular lipase from mutant B. niacini EMB-5 [27]. The latter achieved 19.05-fold purification but only 7.18% overall recovery, illustrating the potential trade-off between enzyme purity and product recovery during multistep downstream processing.

Detailed biochemical characterization was also uncommon and was reported primarily for the cell-bound lipase of Y. lipolytica TISTR 5151 and the purified B. niacini EMB-5 lipase [21,27]. For most other systems, substrate specificity, kinetic parameters, pH and thermal stability, solvent tolerance, and related enzyme properties were not systematically investigated [20,22,23,24,25,26,28,29,30,31,32,33,34,35,36,38,39]. A different type of stability assessment was reported for the mixed cell-bound lipases (CBLs) of M. spicifer AW2 and S. hominis AUP19, which retained approximately 80% of their initial hydrolytic activity after five weeks of storage at 4 °C [37].

Whole-cell and cell-bound systems were the most application-oriented strategies in the included studies because they can reduce or eliminate conventional enzyme purification and, in some cases, external immobilization. Wet cells of Y. lipolytica TISTR 5151 were used directly for lipid conversion [21], while recombinant A. oryzae expressing C. antarctica lipase B provided the most extensively evaluated immobilized whole-cell system [31]. The latter maintained more than 95% FAEE production after ten batches and retained 71.47% of its initial hydrolytic activity after ten cycles, providing the strongest evidence of repeated catalyst use among the included studies [31].

Mixed CBLs from M. spicifer AW2 and S. hominis AUP19 were also evaluated directly as wet-cell catalysts in several related studies [32,33,37]. These systems supported biodiesel synthesis and, in one study, separate POME bioremediation [32]. The later POME50-based study further demonstrated storage stability, with approximately 80% of the initial hydrolytic activity retained after five weeks at 4 °C, while biodiesel yields remained essentially unchanged [37]. Importantly, storage stability and repeated catalytic reuse are distinct performance characteristics and should not be interpreted as equivalent evidence of operational durability.

Other application studies were more limited. The purified B. niacini lipase was evaluated for hydrolysis of several natural oils [27], whereas in several studies the lipolytic system was used primarily for wastewater treatment rather than as a separately recovered catalyst [20,22,26,30,34]. The crude lipase and laccase preparations applied separately to POME in [23] had been produced in synthetic media and therefore do not represent downstream applications of enzymes produced by wastewater-based fermentation. Likewise, the microbial-lipid characterization reported for ethanol-stressed Y. lipolytica concerned an intracellular co-product rather than an application of the extracellular lipase [39].

Overall, the included studies demonstrate three principal routes for wastewater-derived lipolytic systems: crude extracellular preparations, purified soluble lipases, and cell-bound or immobilized whole-cell catalysts. Nevertheless, most investigations ended at the stage of production optimization and activity measurement. Data on recovery efficiency, formulation, long-term stability, repeated use, mass balance, product quality, continuous operation, and scale-up validation remain scarce. Whole-cell systems are particularly promising from a downstream perspective, but their practical value still requires confirmation under repeated, continuous, and larger-scale processing conditions.

3.7. Technological Readiness and Research Gaps

The technological limitations and research priorities identified across the included studies are summarized in

Table 6. Current technological limitations and research priorities for wastewater-based microbial lipase production.

Process Area	Current Evidence	Readiness Assessment	Principal Limitation	Recommended Research Priority	Evidence Basis
Wastewater feedstock diversity	The evidence base is concentrated on OMW and POME. VORW and combined effluent streams were investigated only sporadically.	Narrow proof-of-concept evidence	The technological potential of many other agro-industrial wastewaters remains unknown, and conclusions are dominated by oil-processing effluents.	Expand screening to other lipid-containing and nutrient-rich agro-industrial effluents and compare their performance under a common experimental framework.	[20,21,22,23,24,25,26,27,28,29,30,31,32,33,34,35,36,37,38,39]
Wastewater characterization and batch variability	Some studies reported COD, lipids or O&G, phenols, sugars, nitrogen, solids, and pH, but several publications provided only partial or no initial characterization.	Laboratory characterization is inconsistent	Incomplete feedstock data and limited information on batch, seasonal, and storage-related variability restrict reproducibility and cross-study interpretation.	Adopt a minimum characterization panel covering pH, COD, BOD where relevant, TOC, lipids/O&G, phenols, sugars, nitrogen, phosphorus, solids, conductivity, and batch-to-batch variability.	[20,21,22,23,26,29,30,31,32,33,34,35,37,38,39]
Wastewater conditioning and medium formulation	Centrifugation, filtration, dilution, autoclaving, pH correction, nitrogen supplementation, mineral salts, oils, surfactants, and emulsifiers were frequently used.	Optimized laboratory media	Additional water, chemicals, and energy may offset the environmental and economic benefit of replacing conventional fermentation substrates.	Develop minimally conditioned, low-supplement, and non-sterile processes; quantify water, chemical, and energy inputs through complete process mass and energy balances.	[20,21,22,23,24,25,26,27,28,29,30,31,32,33,34,35,36,37,38,39]
Microbial robustness and process reproducibility	Yeasts, bacteria, filamentous fungi, mixed cultures, and one recombinant whole-cell system were evaluated, usually in single-batch laboratory experiments.	Strain-level feasibility demonstrated	Tolerance to fluctuating wastewater composition, contamination, repeated cultivation, and long-term operation was rarely tested.	Validate selected strains and consortia across multiple independently collected wastewater batches and extended cultivation campaigns, including contamination resilience and genetic or phenotypic stability.	[20,21,22,23,24,25,26,27,28,29,30,31,32,33,34,35,36,37,38,39]
Fermentation engineering and scale-up	Most studies used shake flasks. Only a few progressed to stirred-tank bioreactors, generally at bench scale.	Limited bench-scale validation	Information on oxygen transfer, aeration efficiency, mixing, foaming, rheology, heat transfer, online monitoring, and scale-dependent productivity is scarce.	Perform controlled scale-up studies reporting kLa or equivalent oxygen-transfer indicators, OUR/CER where feasible, power input, foam control, mixing time, and volumetric productivity; evaluate fed-batch and continuous operation.	[25,28,29,30]
Lipase activity measurement and reporting	Different natural oils and p-nitrophenyl esters, assay conditions, enzyme fractions, unit definitions, and expression bases were used.	Analytical methods are not standardized	Activities reported in U/mL, U/L, U/g, and U/mg cannot be used for reliable ranking of strains, wastewaters, or process configurations.	Report complete assay conditions and unit definitions together with volumetric activity, specific activity, productivity, protein or biomass concentration, recovery yield, and enzyme output per unit volume or organic load of wastewater processed.	[20,21,22,23,24,25,26,27,28,29,30,31,32,33,34,35,36,37,38,39]
Integrated wastewater-treatment validation	Several studies reported reductions in COD, O&G or triglycerides, phenols, sugars, TOC, nitrogen, phosphorus, color, or phytotoxicity.	Partial bioremediation demonstrated	Pollutant removal was monitored inconsistently, and treated effluents were rarely compared with discharge or reuse criteria.	Evaluate a standardized final-effluent panel, including residual COD/BOD, O&G, phenols, nutrients, solids, color, toxicity, microbial safety, and compliance with locally relevant discharge or reuse standards.	[20,21,22,23,26,28,30,31,32,33,34,37,38]
Downstream recovery, formulation, and storage	Most extracellular lipases remained as crude supernatants or filtrates. Spray drying, multistep purification, whole-cell immobilization, and explicit storage-stability testing were reported only occasionally.	Downstream proof-of-concept	Recovery yield, concentration losses, formulation performance, storage life, and downstream costs remain poorly characterized, although one mixed CBL system retained substantial activity during five weeks of storage.	Determine enzyme mass balances, recovery and activity yields, concentration factors, storage stability, formulation performance, and the cost and environmental burden of each downstream operation.	[21,25,27,31,32,33,37]
Catalytic application and catalyst reuse	Selected cell-bound, immobilized, or purified lipases were tested in biodiesel synthesis, natural-oil hydrolysis, and POME bioremediation. Storage stability was evaluated for one additional whole-cell system, whereas repeated catalytic reuse was investigated comprehensively in only one immobilized recombinant whole-cell system.	Application-level proof-of-concept	Most studies did not evaluate repeated use, long-term operational stability, continuous processing, product quality, or performance against commercial catalysts.	Conduct repeated-batch and continuous trials, quantify activity decay and catalyst lifetime, verify product specifications, and benchmark wastewater-derived systems against commercial lipases under identical conditions.	[21,27,31,32,33,37]
Biosafety and regulatory suitability	The reviewed studies focused primarily on enzyme production and pollutant removal; systematic assessment of production-strain safety, residual viable cells, microbial contaminants, or intended-use regulatory requirements was uncommon.	Not sufficiently evaluated	Use of opportunistic or non-food-grade microorganisms may constrain enzyme handling, effluent discharge, and applications in food, feed, pharmaceutical, or environmental settings.	Select appropriate production hosts, define containment and inactivation procedures, assess residual viable cells and relevant microbial contaminants, and align product specifications with the intended application.	[20,21,22,23,24,25,26,27,28,29,30,31,32,33,34,35,36,37,38,39]
Techno-economic and environmental performance	Formal techno-economic assessments and life-cycle assessments were absent. One study reported a simplified laboratory-scale profitability analysis for CBL and biodiesel production [37].	Preliminary economic indication; industrial feasibility not demonstrated	The simplified analysis considered mainly raw-material and utility costs and excluded labor and facility-maintenance costs. Capital expenditure, scale-dependent costs, sensitivity analysis, environmental impacts, and comparisons with conventional production routes remain unknown.	Perform formal techno-economic and life-cycle assessments, including sensitivity analysis and complete carbon, water, nutrient, and energy balances, using conventional lipase production and wastewater treatment as reference scenarios.	[20,21,22,23,24,25,26,27,28,29,30,31,32,33,34,35,36,37,38,39]

Note: Readiness assessments are descriptive syntheses of the evidence reported in the included studies and are not formal Technology Readiness Level (TRL) assignments. The evidence basis column refers to the reference numbering used in this review. Abbreviations: OMW, olive mill wastewater; POME, palm oil mill effluent; VORW, vegetable oil refinery wastewater; O&G, oil and grease; COD, chemical oxygen demand; BOD, biochemical oxygen demand; TOC, total organic carbon; OUR, oxygen uptake rate; CBL, cell-bound lipase; CER, carbon dioxide evolution rate; kLa, volumetric oxygen-transfer coefficient.

. Although the available evidence demonstrates the technical feasibility of using agro-industrial wastewater as substrates for microbial lipase production, most of the investigated processes remain at an early laboratory stage. The current literature predominantly provides strain- or process-level proof of concept rather than complete, scalable production platforms. Consequently, the readiness categories presented in Table 6 should be interpreted as a descriptive synthesis of the available evidence rather than as formal Technology Readiness Level assignments.

The available evidence remains dominated by laboratory-scale proof-of-concept studies focused primarily on OMW and POME. Only a few studies progressed to controlled stirred-tank bioreactors, and variability in wastewater characterization, medium conditioning, cultivation conditions, lipase assay methodology, and activity expression continues to limit cross-study comparisons and process generalization [20,21,22,23,24,25,26,27,28,29,30,31,32,33,34,35,36,37,38,39]. Validation across independently collected wastewater batches, larger-scale systems, and prolonged or continuous operation remains scarce.

Major translational gaps also persist in downstream recovery, catalyst formulation, storage stability, repeated use, final-effluent characterization, biosafety, and economic and environmental validation. Although selected studies demonstrated whole-cell reuse, storage stability, or preliminary profitability, formal techno-economic assessment and life-cycle assessment are still absent. Accordingly, the readiness categories summarized in Table 6 should be interpreted as a descriptive synthesis of the current evidence rather than as formal Technology Readiness Level assignments.

Overall, wastewater-based microbial lipase production is supported by substantial laboratory-scale proof-of-concept evidence, but progress toward industrial implementation will require integrated validation of feedstock variability, microbial robustness, scale-up, enzyme productivity, wastewater treatment performance, downstream processing, catalyst stability and reuse, biosafety, and economic and environmental performance.

4. Discussion

4.1. Agro-Industrial Wastewaters as Fermentation Feedstocks for Lipase Production

The present review demonstrates that lipid-rich agro-industrial wastewaters can serve not only as waste streams requiring treatment but also as fermentation feedstocks for microbial lipase production. This concept aligns with the broader transition from end-of-pipe wastewater treatment toward resource recovery and wastewater-based biorefineries. In particular, POME has been recognized as a potential fermentation substrate for recovering enzymes, microbial biomass, lipids, biofuels, and other value-added products, while OMW has similarly been considered a source of carbon, minerals, lipids, and bioactive compounds suitable for biological valorization [41,42]. In the studies included in the present review, this potential was reflected in the predominance of OMW- and POME-based processes and in the ability of diverse yeasts, bacteria, filamentous fungi, and mixed microbial systems to produce lipases in these matrices [20,21,22,23,24,25,26,27,28,29,30,31,32,33,34,35,36,37,38,39].

The suitability of these effluents for lipase production is closely tied to their composition. Unlike conventional carbohydrate-rich fermentation residues, OMW and POME contain substantial amounts of lipid-associated compounds that can serve as both carbon sources and signals that promote lipolytic metabolism. This is particularly relevant because microbial lipase production is often enhanced by oils, fatty acids, triglycerides, or other hydrophobic substrates. More generally, reviews of microbial lipase production from renewable residues have identified lipid-rich agro-industrial wastes and industrial effluents as promising alternatives to refined fermentation substrates, although supplementation with nitrogen sources or additional lipid inducers is frequently required [9,10,13].

However, the same characteristics that make these wastewaters attractive as fermentation feedstocks can also restrict microbial growth. OMW frequently combines a high organic load with substantial concentrations of phenolic compounds and relatively limited nitrogen availability, whereas POME may contain high concentrations of residual oil, suspended matter, organic acids, and other components that can affect mass transfer or microbial physiology [13]. Polyphenols present in food-processing wastewaters are particularly relevant because, although they represent potentially valuable compounds, they may also exert antimicrobial effects and interfere with biological wastewater treatment [42]. Similarly, Cheng et al. [43] identified several potential limitations of microbial POME conversion, including oil-associated inhibition, chemical inhibition, acidic pH, solids, and process-dependent oxygen limitations, and proposed dilution, pretreatment, supplementation, acclimation, mixed cultures, and pH control as possible mitigation strategies. These observations provide a broader explanation for the considerable variation in wastewater conditioning observed in the present review.

An important distinction emerging from the included studies is the difference between using wastewater as a genuine fermentation medium and incorporating wastewater as one component of a substantially modified culture medium. Several microorganisms were successfully cultivated in undiluted OMW or POME [20,21,23,26,29], demonstrating that intensive dilution is not universally required. In contrast, other studies used wastewater at considerably lower proportions or combined it with conventional nutrient mixtures, additional carbon or nitrogen sources, oils, surfactants, and mineral salts [24,27,28,30,31,32,33,34,35,36,37,38,39]. At the extreme, POME constituted only a relatively small fraction of the culture medium in some systems [24,31]. Consequently, the term wastewater-based medium is more appropriate for the overall evidence base than implying that all investigated processes used wastewater as the sole fermentation substrate.

This distinction is particularly relevant when assessing the proposed economic advantage of waste-derived media. Conventional fermentation media are a major component of enzyme production costs, so replacing refined substrates with inexpensive residues is often proposed to reduce lipase production costs [9,10]. Nevertheless, supplementation can substantially alter the extent of waste valorization. Nitrogen sources such as yeast extract, peptone, ammonium salts, or urea were frequently required in the reviewed studies [20,22,28,29,30,34,35,37,38,39]. Additional oils and lipid-derived compounds were also commonly added, including olive oil, palm oil, waste frying oil, Tween 80, and other emulsifying agents [23,24,25,29,32,35]. Although these components may increase lipase production, their presence complicates attribution of enzyme induction specifically to the wastewater itself.

This issue is particularly important for lipid inducers. The intrinsic lipid fraction of OMW, POME, and VORW may provide both substrate and induction signals, but when an additional oil is supplied, the relative contribution of the wastewater becomes difficult to assess. Future studies would benefit from factorial designs that independently evaluate wastewater concentrations, intrinsic lipid content, added lipid inducers, and nitrogen supplementation. Such experiments could determine whether the wastewater merely replaces part of the basal medium or whether its native composition provides a genuine functional advantage for lipase biosynthesis. This distinction would also enable more meaningful comparisons with conventional synthetic media.

Wastewater dilution presents a similar trade-off. Dilution can reduce concentrations of inhibitory compounds, mitigate viscosity and suspended-solids effects, and improve oxygen transfer, but it also increases water demand and lowers the concentration of potentially useful nutrients. The broader POME literature likewise identifies dilution and debris removal as common strategies to overcome microbial inhibition, while emphasizing that additional pretreatment itself introduces process requirements [43]. The results of the present review indicate that high dilution is not always necessary: several strains tolerated undiluted effluents, and B. aryabhattai BA03 produced substantial lipase activity even in non-sterile OMW-based medium [34]. This suggests that selecting or adapting robust microorganisms may be preferable to extensive wastewater conditioning in future process development.

The predominance of OMW and POME in the available literature should also be interpreted cautiously. Their frequent use likely reflects both their high lipid content and the regional importance of olive- and palm-oil industries, rather than evidence that these are inherently the most suitable wastewaters for lipase production. POME has already been extensively discussed as a multiproduct biorefinery feedstock, with possible conversion to enzymes, microbial lipids, biofuels, biomass, and other products [41]. In contrast, the potential of other agro-industrial wastewaters remains largely unexplored for microbial lipase production. The identification of only one VORW-based study [28] and very few combined-wastewater systems in the present review therefore points to a substantial opportunity to expand the substrate spectrum beyond olive- and palm-oil processing effluents.

Microbial Diversity and Physiological Adaptation to Wastewater Matrices

The microbial groups identified in this review differ not only taxonomically but also in the physiological strategies that may support growth in lipid-rich, chemically complex wastewater matrices. Yeasts and bacteria typically grow as dispersed cells and are readily cultivated under submerged conditions, which facilitates mixing, aeration, and process control. Among yeasts, Y. lipolytica is particularly well adapted to hydrophobic substrates because extracellular or cell-associated hydrolysis of triacylglycerols can be followed by fatty-acid uptake, peroxisomal β-oxidation, and, depending on nutritional conditions, intracellular storage in lipid bodies [44,45]. These characteristics provide a physiological basis for its repeated use in lipid-rich POME, OMW, and refinery-wastewater systems.

Filamentous fungi such as Aspergillus and Fusarium spp. employ a distinct growth strategy. Hyphal growth and extensive secretion of extracellular hydrolytic and oxidative enzymes can facilitate colonization and conversion of heterogeneous or particulate substrates, and may confer an advantage in complex matrices. These characteristics also explain the frequent use of filamentous fungi in static or solid-associated fermentation systems. However, filamentous morphology may be disadvantageous in submerged bioreactors because dispersed mycelia can increase broth viscosity and impair mixing and oxygen transfer, whereas large pellets may develop internal mass-transfer limitations. Thus, the relative advantage of yeasts, bacteria, or filamentous fungi depends strongly on wastewater composition and reactor configuration rather than on microbial taxonomy alone [46].

Resistance to inhibitory wastewater constituents is likewise strain- and taxon-dependent. Phenolic compounds and organic acids may impair membrane integrity, intracellular pH homeostasis, and oxidative-stress balance. Filamentous fungi may have an additional advantage through extracellular oxidative enzyme systems, including laccases and peroxidases, which can participate in the transformation of aromatic and phenolic compounds [47]. However, these mechanisms were not directly characterized in the Aspergillus and Fusarium studies included in the present review [20,22,24,35]. Therefore, growth in phenol-containing wastewater should be regarded as evidence of phenotypic tolerance rather than as direct proof that a specific oxidative detoxification pathway was responsible.

Overall, agro-industrial wastewater should be viewed as a heterogeneous fermentation feedstock rather than a standardized, inherently low-cost medium. Its value lies in the simultaneous availability of organic carbon, lipids, minerals, and other nutrients, but successful utilization depends on balancing these resources against inhibitory compounds and nutritional deficiencies. The most promising future processes are likely those in which the intrinsic wastewater composition supplies most of the microbial carbon and induction requirements while minimizing dilution, sterilization, and external supplementation. Demonstrating this balance is essential before wastewater-based lipase production can be considered a genuinely resource-efficient alternative to conventional fermentation.

4.2. Coupling Lipase Production with Wastewater Treatment

A major advantage of wastewater-based lipase production is the potential to generate a value-added enzyme while simultaneously reducing the pollutant load. Several studies reviewed here showed that microbial growth and lipase production were accompanied by substantial decreases in COD and lipid fractions, and, in some cases, reductions in phenolic compounds, sugars, TOC, nitrogen, phosphorus, color, or phytotoxicity [20,21,22,23,26,28,30,33,34,37]. This distinguishes wastewater-based fermentation from conventional enzyme production, in which the culture medium itself generally requires management after fermentation.

This concept aligns with the broader use of microbial lipolytic systems to treat fat- and oil-rich wastewaters. Kumar et al. [48] highlighted the potential of microbial lipases as biocatalysts to accelerate the degradation of lipid pollutants, while the broader POME biorefinery literature emphasizes a transition from pollutant removal alone to simultaneous resource recovery [41]. In this context, lipase production can be regarded as a form of product-oriented wastewater treatment, in which part of the organic load is converted into microbial biomass, enzymes, lipids, or other potentially recoverable products rather than being removed solely as waste.

The later M. spicifer–S. hominis study offers a particularly informative example of this integration because CBL production, biomass formation, and COD and O&G removal were optimized within the same POME-based cultivation process [37]. However, the time-course results also showed that these objectives were not temporally aligned: maximum CBL activity occurred after 24 h, whereas COD and O&G removal continued to increase during prolonged cultivation. This illustrates an important process-design trade-off, as the optimal harvest time for enzyme production may differ from the cultivation period required to maximize wastewater treatment. Future process optimization should therefore treat enzyme productivity and remediation performance as separate responses rather than assuming that both reach their optima simultaneously. A plausible biological explanation for this temporal separation is that lipase production and bulk organic-matter removal reflect distinct stages and processes of microbial metabolism. Cell-bound lipase activity may reach an early maximum once the lipolytic system is induced, whereas hydrolyzed lipids and other wastewater-derived organic compounds can continue to be taken up and metabolized during subsequent biomass growth. Fatty acids released from triacylglycerols may undergo intracellular catabolism, including β-oxidation in organisms capable of this pathway, while COD reduction also reflects the assimilation or transformation of non-lipid organic constituents. Therefore, the delayed maximum in COD and O&G removal cannot be attributed solely to lipase activity. Because the underlying metabolic fluxes were not directly measured in [37], this interpretation should be regarded as a plausible mechanistic explanation rather than an experimentally demonstrated mechanism.

Nevertheless, high pollutant-removal percentages should not be taken as evidence of complete wastewater treatment. Only a minority of the included studies comprehensively characterized the final effluent, and compliance with discharge or reuse standards was rarely demonstrated. Moreover, reductions in COD or O&G during microbial cultivation cannot necessarily be attributed directly to lipase activity because substrate assimilation, biomass formation, biosorption, and other extracellular or intracellular metabolic processes occur simultaneously. Therefore, lipase production and wastewater-treatment performance should be evaluated as complementary but distinct process outputs. Future studies should therefore adopt standardized final-effluent monitoring and compare residual pollutant levels against locally relevant discharge or reuse criteria, while experimental designs should distinguish lipase-mediated hydrolysis from broader microbial assimilation and metabolic processes.

4.3. From Lipase Production to Functional Biocatalysts

Although all included studies demonstrated lipolytic activity, relatively few advanced from enzyme production to the evaluation of a functional biocatalyst. Most lipases were studied in crude culture supernatants or filtrates, whereas enzyme purification, formulation, immobilization, storage-stability assessment, catalytic reuse, and product-oriented applications were considerably less common [21,25,27,31,32,33,37]. This represents an important gap because high activity in a fermentation broth does not necessarily translate into a technically useful enzyme preparation.

Among application-oriented strategies, cell-bound and immobilized whole-cell lipases are particularly attractive. Whole-cell systems can eliminate several conventional downstream operations, including enzyme purification and, in some cases, external immobilization, while facilitating catalyst recovery from the reaction mixture. These advantages align with advances in enzymatic biodiesel technology, where immobilization and whole-cell catalysis have been proposed to improve operational stability and enable catalyst reuse [49,50]. Hama et al. [49], for example, emphasized the role of lipase technology in biodiesel production from diverse feedstocks, whereas Quayson et al. [50] identified catalyst stability, immobilization, and reuse as central factors influencing the feasibility of enzymatic biodiesel processes.

The results obtained with Y. lipolytica, recombinant A. oryzae, and the M. spicifer–S. hominis co-culture illustrate this direction particularly well [21,31,32,33,37]. These systems enabled microbial biomass to be used directly in transesterification or esterification reactions, thereby avoiding conventional enzyme purification. The recombinant A. oryzae catalyst also demonstrated repeated use, retaining substantial activity over ten reaction cycles [31]. The M. spicifer–S. hominis system provided complementary evidence of stability: after five weeks of storage at 4 °C, approximately 80% of the initial hydrolytic activity remained, and biodiesel yields were essentially maintained [37]. One plausible explanation for this stability is the cell-associated localization of the lipases. In whole-cell systems, the enzyme remains associated with the cellular envelope and surrounding macromolecular structures, which can serve as a natural immobilization environment, restrict conformational mobility, and protect the enzyme from external destabilizing conditions. This protective effect may contribute to greater storage stability than would be expected for an unprotected soluble enzyme. However, the structural basis of the observed stability was not directly investigated in [37], so the contribution of the cellular envelope should be regarded as a plausible explanation rather than a demonstrated mechanism. Together, these studies indicate that operational reuse and storage stability should be treated as distinct but complementary characteristics when assessing the practical value of whole-cell catalysts.

By contrast, conventional multistep purification was reported only for the B. niacini lipase [27], where a 19.05-fold increase in specific activity was accompanied by a final recovery of only 7.18%. This illustrates the potential trade-off between enzyme purity and overall product recovery. In wastewater-based processes, where low production cost is a key proposed advantage, extensive purification may therefore diminish some of the economic benefit associated with using inexpensive feedstocks.

Future development should focus not only on maximizing lipase activity during fermentation but also on recoverable catalytic activity, formulation, storage stability, operational lifetime, and reusability. For low-cost wastewater-based systems, a moderately active crude or whole-cell catalyst that can be efficiently recovered, stored, and reused may ultimately be more attractive than a highly purified enzyme that requires multiple downstream operations. Comparative studies should therefore assess not only enzyme activity but also catalyst yield, stability, number of usable cycles, regeneration potential, catalyst cost, product quality, and performance relative to commercial preparations.

4.4. Sustainability and Prospects for Integrated Wastewater Biorefineries

Wastewater is often presented as an inherently low-cost and sustainable fermentation feedstock. However, replacing a refined carbon source with wastewater is only one component of the overall process. The studies reviewed here frequently included wastewater transport and storage, centrifugation or filtration, dilution, sterilization, pH adjustment, nutrient supplementation, aeration, and, in some cases, additional oils or surfactants. Consequently, the environmental and economic benefits of wastewater-based lipase production cannot be established solely by substrate replacement.

This distinction aligns with the broader principles of sustainable biocatalysis, which emphasize assessing environmental performance at the level of the complete process rather than solely on the renewable or waste-derived origin of a substrate [51]. For wastewater-derived lipases, such an assessment should include the energy required for sterilization and aeration, the water used for dilution, the production and consumption of nutrient supplements, downstream enzyme recovery, treatment of the remaining effluent, and the value and lifetime of the resulting catalyst. Whole-cell approaches may be advantageous in this respect by reducing downstream processing, but this potential benefit also requires quantitative verification.

Formal techno-economic assessments and life-cycle assessments remain absent from the current evidence base. However, one study provided a simplified laboratory-scale profitability analysis for CBL and biodiesel production [37]. Profitability indices above unity were reported for the optimized processes, but the calculation considered primarily raw-material and utility costs and excluded labor and facility-maintenance costs. Consequently, these results should be regarded as a preliminary economic indication rather than evidence of full process feasibility. Future studies should integrate lipase productivity with wastewater-treatment efficiency, material and energy balances, catalyst recovery and reuse, capital and operating costs, sensitivity analysis, downstream processing, residual-effluent treatment, and co-product generation. Comparisons should ideally include conventional lipase fermentation and conventional wastewater treatment as reference scenarios. Accordingly, the technological value of wastewater-derived lipase systems should be assessed based on the combined performance across enzyme production, wastewater-quality improvement, catalyst recovery and lifetime, downstream requirements, and overall economic and environmental burdens, rather than on any single process outcome.

Ultimately, the greatest potential may lie not in producing lipase as a single product but in developing an integrated wastewater biorefinery that converts organic pollutants into multiple recoverable outputs. Depending on the microbial platform, these outputs may include lipases, microbial biomass, intracellular lipids, biodiesel precursors, and other metabolites, alongside partial wastewater remediation. Future research should therefore move beyond demonstrating microbial growth in wastewater toward processes in which enzyme production, co-product recovery, wastewater treatment, and water reuse are designed and evaluated as components of an integrated system.

5. Conclusions

This scoping review shows that agro-industrial wastewaters can support microbial lipase production and, in selected systems, simultaneous reductions in wastewater pollutant loads. However, the current evidence remains dominated by laboratory-scale studies using OMW and POME, with limited representation of other agro-industrial effluents. Differences in wastewater conditioning, lipase localization, assay methods, unit definitions, and activity-expression bases further limit direct comparisons among studies. Moreover, reductions in COD, O&G, phenolic compounds, or other wastewater constituents should not be interpreted as evidence of complete wastewater treatment or compliance with discharge or reuse requirements.

Most investigated processes remain at the proof-of-concept stage, with limited scale-up validation and insufficient information on enzyme recovery, formulation, long-term catalyst stability, repeated use, biosafety, and process economics. Future research should therefore prioritize standardized analytical and activity-reporting methods, minimally supplemented cultivation, pilot- and industrial-scale validation, long-term catalyst performance, comprehensive final-effluent assessment, and formal techno-economic and life-cycle assessments. Although wastewater-based microbial lipase production is a promising circular-bioeconomy strategy, the available evidence does not yet demonstrate that it is economically or environmentally superior to conventional enzyme-production and wastewater-treatment processes.