INTERNATIONAL JOURNAL OF LATEST TECHNOLOGY IN ENGINEERING,

MANAGEMENT & APPLIED SCIENCE (IJLTEMAS)

ISSN 2278-2540 | DOI: 10.51583/IJLTEMAS | Volume XV, Issue II, February 2026

Assessment of Bio-Surfactant Producing Microorganisms from Palm Oil

Mill Effluents in Edo State, Nigeria

Ekhator Stanley Osarobo.

School of General Studies Edo State College of Health Sciences and Technology, Edo State, Nigeria

DOI: https://doi.org/10.51583/IJ L T EMAS.2026.15020000060

Received: 25 February 2026; Accepted: 02 March 2026; Published: 16 March 2026

ABSTRACT

Palm oil mill effluent (POME) is a wastewater generated from palm oil milling activities which requires effective

treatment before being discharged into the watercourses due to its highly polluting properties. Hence this study

was aimed at evaluating the biosurfactant-producing microorganisms from POME at different depths from large

and small/medium scale enterprises in Edo State, Nigeria.

POME was aseptically collected using sterile bottles from various depths: top, middle and bottom in selected

palm oil companies across Edo State, Nigeria. The companies were categorized into large-scale enterprises

(L.S.E.), which included Okomu Oil Palm and NIFOR and small and medium-scale enterprises (S.M.E.),

comprising Ovbiogie, Sapele Road and Aduwawa oil palm companies.

The bacteria isolated were Bacillus cereus, Pseudomonas aeruginosa, Bacillus amyloliqefaciens, Klebsiella

aerogenes and Escherichia coli. Fungi like Aspergillus niger, Fusarium solani, Penicillium chrysogenum,

Microsporum sp., Penicillium citrinum and Aspergillus flavus were also isolated from these samples using the

Pour plate techniques. The bacterial species obtained in the pure culture of substrate were identified using

standard bacterial and fungal techniques. Isolated organisms were screened for their ability to produce

biosurfactants using oil spreading assay, hemolytic, and emulsification activity. The test of how susceptible the

isolates were to antibiotics was conducted with the aid of the Kirby-Bauer disk diffusion assay. The data obtained

were analyzed using Microsoft Excel 2019 and PhyloT software to establish the relationship between isolated

microorganisms from POME.

The results of the total heterotrophic bacterial counts ranged from log₁₀3.90±1.00 cfu/g (Small and Medium

Scale- Sapele Road) to log₁₀4.66±3.0 cfu/g (Large Scale Enterprise- Okomu). The total fungal counts ranged

from log₁₀3.78±1.00 cfu/g (Small and Medium Scale- Aduwawa) to log₁₀4.34±2.00 cfu/g (Small and Medium

Scale- Aduwawa). The difference in percentage reduction in the density of microbes between the top and bottom

depth ranged from 43.18% (NIFOR) to 72.29 % (Okomu). Also, a significant difference (p<0.05) between the

microbial diversity of large-scale and small-scale oil-producing enterprises was observed. The isolated bacteria

included Bacillus cereus, Pseudomonas aeruginosa, Bacillus amyloliqefaciens, Klebsiella aerogenes and

Escherichia coli.

The isolated fungi were Aspergillus niger, Penicillium citrinum, Penicillium chrysogenum, Aspergillus flavus,

Microsporum sp. and Fusarium solani. Biosurfactant screening results revealed that most microbial isolates were

potential biosurfactant producers, with Bacillus sp. showing the highest clear zone of oil spread assay. However,

specific isolates like E. coli and Microsporum sp. did not produce any clear zone for oil spread assay. More so,

Bacillus sp. was found to be the best biosurfactant producer due to its hemolytic activity and the assay with the

highest zone (10mm) of displacement. POME is home to many microorganisms of importance to both industrial

and environmental processes. This research has demonstrated that POME serves as a reservoir for

microorganisms capable of producing biosurfactants.

Keywords: Palm Oil Mill Effluent (POME), Biosurfactant-producing microorganisms, Microbial diversity,

Bacillus species, Oil spreading assay, Antibiotic susceptibility testing.

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INTRODUCTION

Biosurfactants are group of surface-active molecules that are synthesized by different microorganisms, and thus

have uniquely diverse structure (Nitschke and Costa, 2007). They are amphiphilic in nature, possessing both

water-loving (hydrophilic) and water-repelling (hydrophobic) polar properties. This unique structural feature

makes them suitable surface tension reducing agent in different phases of fluid. They are very useful in many

commercial processes that have been reported in different fields and these include the bioremediation of

recalcitrant pollutants, microbial-enhanced oil recovery, food and cosmetic industries (Nitschke and Pastore,

2006). Biosurfactants have become very popular due to their inherent health, commercial and environmental

benefits such as their low toxic levels, environmental preservation quality, easy sources and low cost of

production including renewable materials or agricultural waste products (Saharan et al., 2011). However, the

application of biosurfactants in the wider industry is still limited due to economic and/or operational concerns

(Saharan et al., 2011).

Presently, biosurfactants have low potential benefits and competitive advantages when compared with chemical

surfactants. This is mainly because its production output is low, relative to costs (Makkar and Cameotra, 2002).

Efforts to address these challenges signaled the alternative use of agro-wastes or feed stocks obtained at little or

no cost, as sources of biosurfactant-producing organisms, in many studies (Joshi et al., 2008, Sobrinho et al.,

2008, Saimmai et al., 2012). These approaches are unconventional and still have not addressed the challenges

leading to the adoption of appropriate wastes as substrates (Nawawi et al., 2010). The unavailability of waste

substrates which have a uniform composition of important macromolecules such as carbohydrates and lipids,

and therefore have the benefits of ensuring optimal microbial growth and biosurfactant outputs, is still a serious

challenge for researchers (Makkar and Cameotra, 1999).

A wide range of waste products are usually generated in large quantities from oil extraction processes and these

include residual oil that can potentially cause water and soil pollution (Singh et al., 2011). However, oil residues

can be absorbed, dispersed or made soluble by microorganisms which live in soil and water and survives by

producing biosurfactants (Nerurkar et al., 2009). Organic waste products from palm oil production are generally

difficult to manage, raising serious environmental concerns in the areas where production takes place (Sulaiman

et al., 2011). Ameliorative actions would require approaches that are viable economically and also practical in

their implementation (Puetpaiboon and Chotwattanasak, 2004). Ageneral name for the waste product from palm

oil extraction is palm oil mill effluent (POME). POME is a mixture of over 90% water, less than 1% oil and

about 4.5% % total solids (Ma, 2000). Over 4,000 mg/l of oil and grease (which can occur as oil droplets in a

water–oil emulsion) can be an important constituent of the colloidal suspension (Alhaji et al., 2016). This study

presents POME as a new and promising source for producing biosurfactants. Using agricultural wastes as

substrates in the biotechnology industry has led to significant cost reductions in biosurfactant production and

has also driven the development of innovative and effective waste management techniques (Banat et al., 2010).

Biosurfactants are produced by a diverse group of microorganisms which occur in diversity and are secreted or

found on the cell surface of substrates that cannot mix with water, especially during the growth phase (Singh et

al., 2010). In this investigation, POME was utilized as a unique source for organisms capable of producing

biosurfactants.

LITERATURE REVIEW

Introduction to Surfactants

Surfactants are compounds that reduce the surface tension of a liquid, making it easier for the liquid to spread or

mix with other substances. Common examples include detergents, emulsifiers, wetting agents, foaming agents

and dispersants. (Rosen and Kunjappu, 2012).

Definition and Composition of Biosurfactants

Surfactants of biological nature are those that are produced by microorganisms, and are therefore referred to as

biosurfactants or microbial surfactants. These surfactants of biological origin can as well reduce the tension or

force acting on the surface of a liquid or between two phases of liquids that do not form a uniform mixture

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(Mulligan and Catherine 2005). Their occurrence is as complex lipid macromolecules including phospholipids,

fatty acids, glycolipids, and lipoproteins (Banat et al., 2000).

Advantages of Biosurfactants over Synthetic Surfactants

Biosurfactants are not toxic, they are more effective, and poses no threat to the environment, unlike the

alternative synthetic surfactants (Ron and Rosenberg 2001). Also, biosurfactants could be produced from cheap

agricultural materials unlike the chemical surfactants that are gotten from petroleum feedstock (Magalhaes et

al., 2018). Additionally, microbial surfactants are more stable at under extreme conditions of temperature,

salinity, and pH; and this makes them more commercially viable than synthetic surfactants (Gutnick and Bach,

2002; Mulligan and Catherine, 2005). These characteristics are very important in both food and non-food

manufacturing processes including drug formulations, environmental bioremediation, and enhanced oil recovery.

Factors Affecting Biosurfactant Production

Historically, the production of surface-active molecules has gained increasing traction because of the potential

benefits in food and non-food industrial processes. Some important factors that can affect the production of

microbial surfactants include aeration, temperature, nitrogen, carbon, and other trace elements (Roy, 2017).

Nevertheless, the type and quantity of biosurfactants generated typically rely on the specific biosurfactant-

producing organism (Marchant and Banat 2012). The solubility of oil can also be increased by biosurfactants,

and this can potentially increase their bioavailability as important sources of carbon and energy (Mulligan, 2009).

Biosurfactants are amphiphilic in nature, possessing both water-loving (hydrophilic) and water-repelling

(hydrophobic) polar properties. This unique structural feature makes them suitable surface tension reducing

agent in different phases of fluid (Nayak et al., 2009).

Health, Economic and Environmental Significance

Biosurfactants have become very popular due to their inherent health, commercial and environmental benefits

such as their low toxic levels, environmental preservation quality, easy sources and low cost of production

including renewable materials or agricultural waste products, and possibly under extreme conditions of

temperature, salinity and pH levels (Pansiripat et al., 2010). Surfactants from microorganisms have very unique

properties and are therefore a great fit for new applications. The evidence of this specificity has been adduced in

many previous studies on the relevance of biosurfactants in industrial sectors (Perfumo et al., 2010) and in

environment protection (Das and Mukherjee, 2007), over the last decade.

Environmental Applications

Water repellent contaminants in petroleum, soil and water environments hinder microbial degradation. However,

when these contaminants are solubilized, often through the action of biosurfactants, they become more

bioavailable, facilitating easier and more efficient microbial breakdown (Metcalfe et al., 2008). Naturally-

occurring surfactants are able to increase the surface area of water-repelling surfaces, and thereby increasing

their solubility in water.

This property is particularly useful in environments contaminated with substances like pesticides in soil and

water. For this reason, surfactants might contribute significantly to the degradation of polluting agents by

microbes (Murphy et al., 2005). The mechanisms or models for identifying and classifying microbial surfactants

which are produced from different microorganisms have recently undergone broad review (Ying, 2006).

Various organic compounds serve as crucial carbon and energy sources for microbial proliferation, readily

diffusing into cells with the assistance of microorganisms. This is essentially possible when the carbon and

energy sources exists in a form that cannot dissolve in water, such as a hydrocarbon (CxHy); and thus their

diffusion into the cell is aided by their production of biosurfactants. Usually, the CxHy active agents in the

growth medium are blended by the release of ionic surface molecules (surfactants) by some fungi and bacteria,

including yeast (Zheng et al., 2008).

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Examples of Microbial Surfactants and Producing Organisms

Examples of these microbial surfactants include rhamnolipids or sophorolipids which are produced by various

Pseudomonas sp. and Torulopsis sp., respectively (Walter et al., 2010). Moreover, certain microorganisms

possess an intrinsic capability to modify their cell wall structure through the production of macromolecules like

nonionic surfactants. Examples include the cell wall binding lipopolysaccharides produced by Candida lipolytica

and Candida tropicalis, followed by Rhodococcus erythropolis and several strains of Mycobacterium sp. and

Arthrobacter sp. that creates nonionic trehalose corynomycolates (Tabatabaee et al., 2005 and Makkar et al.,

2011). Similarly, Emulsan, derived from species of Acinetobacter and lipoproteins such as Surfactin and

Subtilisin, are examples of some lipopolysaccharides that are produced by Bacillus subtilis (Gorkovenko et al.,

1997). Additional examples of microbial surfactants that demonstrate significant efficacy include:

(i) Mycolates and Corynomycolates, synthesized by Rhodococcus sp., Corynebacteria sp., Mycobacteria sp.,

and Nocardia sp.

(ii) Ornithinlipids, derived from Pseudomonas rubescens, Gluconobacter cerinus and Thiobacillus ferroxidans

(Okoliegbe and Agarry, 2012).

Screening Methods for Biosurfactant-Producing Microorganisms

A variety of screening methods used in identifying biosurfactant-producing organisms exist, and examples

include methylene blue assay/Centriamide test (CTAB), β haemolysis test, oil displacement test, drop collapsing

and emulsification index test (Satpute et al., 2010). However, it is not easy to precisely know the type of

biosurfactant derived from the microorganisms by using only one method. This is basically as a result of their

unique chemical and functional characteristics; which altogether makes it necessary to consider the application

of a combination of screening methods to be able to sufficiently understand the biosurfactant-producing ability

and mechanisms of individual microorganism. The research conducted by Satpute et al. (2010) also adduced

evidence that the application of single screening method is not suitable for identifying all types of microbial

surfactants, and thus recommended the adoption of a combination of (more than one) screening methods for the

identification of potential biosurfactants-producing microbes.

Advantages of Biosurfactants

When compared to surfactants that are chemically synthesized, biosurfactants have the following advantages:

1. They are Biodegradable

This is because they are not very toxic, their chemical structure is simple, they do not accumulate in the

environment because they degrade easily.

2. They are compatible with the biological environment and are easily digested (biocompactability

and digestibility)

They are compatible with living systems and thus can be used in both food and non-food materials

including additives, drugs and cosmetics; this is essentially because they are of biological origin.

3. Raw materials abundance in supply

The raw materials for biosurfactants production are abundant in supply. The microbial production process

can involve the separate or combined use of the carbon sources which include carbohydrates, lipids or

hydrocarbons.

4. Production economics of scales

Biosurfactants can typically be derived from industrial wastes which are potentially resourceful means

of producing large quantities of microbial surfactants, when the intended use is permitted.

5. Environmental control

Biosurfactants promotes bioremediation, control of oil spills, and are useful in biodegrading, detoxifying

and stabilizing emulsions or effluents from industrial processes.

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6. Specificity

Biosurfactants possess distinct functional groups that enable them to perform specific tasks, such as de-

emulsifying industrial emulsions and detoxifying particular pollutants.

Other applications encompass the formulation of specialized cosmetics and their utilization in tailored

pharmaceutical and food processing. The notable resilience of biosurfactants under extreme conditions of

temperature, pH, and salinity underscores their significant characteristics (Murphy et al., 2005).

Classification of Biosurfactants

The microbial origin of biosurfactants and what they are chemically composed of, are the basis of their

classification. This is unlike chemical surfactants whose classification is based on the nature or type of polar

groups they possess.

Biosurfactant Classification based on molecular weight

Biosurfactants can be categorized into two groups: low molecular mass molecules, which effectively reduce

surface and interfacial tension, and large molecular-mass polymers, which serve as more efficient emulsion-

stabilizing agents (Rosenberg and Ron, 1999). Important macromolecules such as phospholipids, lipopeptides,

and glycolipids are the main classes of surfactants with low molecular mass (Mukherjee et al., 2006).

Biosurfactants majorly contain negative charges or they can be neutral. Similarly, the presence of compounds

derived from fatty acids makes majority of biosurfactants have water-repelling structures, while the water-loving

structures are usually composed of amino acid, phosphate, carbohydrate, or cyclic peptide (Banat et al., 2014).

Low molecular weight biosurfactants:

These compounds effectively reduce surface and interfacial tension at the air/water interface. Low molecular-

weight biosurfactants typically consist of glycolipids or lipopeptides. Among the extensively researched

glycolipids are rhamnolipids, trehalolipids, and sophorolipids, which are disaccharides acylated with long-chain

fatty acids or hydroxy fatty acids (Fracchia et al., 2015).

High- molecular weight biosurfactants:

These are also known as bioemulsans, and are more efficient oil in water emulsions stabilizers. Their effective

emulsifying property is based on their ability to show considerable specificity for substrate, as well as work at

low concentrations (Uzoigwe et al., 2015). Previous studies have reported that polymeric surfactants that are

made up of large macromolecules such as polysaccharides, proteins, and lipopolysaccharides, are usually

produced on the outer cell surface by a large number of bacterial species from different genera (Rosenberg and

Ron, 1999).

Furthermore, biosurfactants can be classified based on the nature of their polar groups, resulting in either anionic

or neutral characteristics. Their hydrophobic structure is determined by compounds derived from fatty acids or

the presence of long-chain fatty acids. The hydrophilic region can include carbohydrates, amino acids,

phosphates, or cyclic peptides. Generally, biosurfactants exhibit the following structural components: a

hydrophilic moiety composed of amino acids or peptide anions or cations; a hydrophobic moiety comprising

unsaturated, saturated, or derivative fatty acids and mono-, di-, or polysaccharides.

Classification Based On Chemical Composition

Glycolipids:

In biosurfactants, glycolipids make up the majority. Several carbohydrates, when paired with one or more

aliphatic carboxylic acids, hydroxy fatty acids, or fatty alcohols, can produce glycolipids. These biosurfactant

compounds hold significant potential for commercial applications due to their high production yields and

capability to be synthesized from renewable substrates (Marchant and Banat 2012). Microbial species such as

Pseudomonas sp.

synthesise rhamnolipids, Pseudozyma antarctica synthesise mannosylerythriol lipids,

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Nocardia sp. and Mycobacterium sp., Rhodococcus sp. synthesises trehalose and Candida sp. synthesizes

sophorolipids (Zheng et al., 2008).

Rhamnolipids: The most widely researched type of lipids are glycolipids, which are made up of at least one or

more than one rhamnose molecules bound to at least one or more than one β-hydroxydecanoic acid molecules

(Abdel-Mawgoud et al., 2010). One of the hydroxyl groups from the acid is involved in forming a glycosidic

bond with the reducing end of the rhamnose disaccharide, while the second hydroxyl group from the acid is used

to create ester bonds (Abdel-Mawgoud et al., 2010). It was initially investigated the synthesis of glycolipid-

containing rhamnose in Pseudomonas aeruginosa. Major glycolipids produced by P. aeruginosa are L-

Rhamnosyl-Lrhamnosyl-β-hydroxydecanoyl-β-hydroxydecanoate

and

Lrhamnosyl-β-hydroxydecanoyl-β-

hydrtocydecanoate, also known as rhamnolipids 1 and 2 (Nitschke and Pastore 2006). It is believed that the

phosphatidylethanolamine component in living membrane systems associate with the rhamnolipid molecular

entity and as a result, according to Sanchez et al. (2006), they possess antibacterial action against both Gram

positive and Gram negative microorganisms. Additionally, rhamnolipids have important use in beauty products,

drugs, and food industries (Sánchez et al., 2006).

Trehalolipids: There are reports of several molecular variants of microbiological trehalolipid biosurfactants.

Most taxonomic categories within Mycobacterium, Corynebacterium and Nocardia are associated with the

disaccharide trehalose, which is linked to mycolic acid at positions C-6 and C-6 (Daffé and Draper, 1998). The

long, α-branched chain, and β-hydroxy fatty acids molecular entity are called 2-alkyl, 3-hydroxy long-chain fatty

acids (Brennan and Nikaido, 1995). The degree of branching or unsaturation, the total amount of carbon

molecules, and molecular makeup of 2-alkyl, 3-hydroxy long-chain fatty acids in trehalolipids derived from

different living organisms are all distinct. The surface energy and interfacial surface tension within the culture

medium were reduced by trehalose lipids obtained from Rhodococcus erythropolis and Arthrobacter sp. (White

et al., 2013).

Mannosylerythritol lipids: The fungus Pseudozyma antarctica forms mannosylerythritol lipids (MEL) as a

combination of four component parts: MEL-A and MEL-B consist of the principal derivatives, while MEL-C

and MEL-D are the secondary derivatives (Konishi et al., 2007). These complexes' foundation is a

mannoseerythritol disaccharide, which happens to be acetylated to form short carbon molecules of two to eight

or long carbon molecules of ten to eighteen fatty acid chain length (Kitamota et al., 1990).

The variety of molecular properties exhibited by MEL, such as binding affinity of proteins to immunoglobulin

G and adhesin and promotion of cell division in relation to distinct mammalian cells (Im et al., 2001). They can

also reduce the surface energy of water approximately to 35mN/m (Fischer and Zettl, 2000).

Both pharmaceutical and medical industries are very interested in MELs due to their intriguing biological

behavior.

Sophorolipids: Yeast like Torulopsis bombicola, T. petrophilum, and T. apicola are the dominant species that

synthesizes glycolipids, and are made up of a hydroxyl fatty acid of long-chain length and a dimer

of carbohydrate sophorose linked together by a glycosidic bond (Bajaj and Annapure, 2015). Sophorolipids are

typically found in combination with macrocyclic lactones and free acid state (Hirata et al., 2009). Evidence has

shown that the sophorolipid's lactone state is crucial for a number of different purposes (Hu and Ju, 2001). A

minimum of six to nine different hydrophobic sophorolipids are combined to form these biosurfactants (Hu and

Ju, 2001).

Lipopeptides and lipoproteins: Typically, each of the molecules in this group of biosurfactants are made up of

peptides that are cyclically connected to a fatty acid. These bactericidal-like compounds are produced by a

number of microorganisms including Bacillus subtilus (Malfanova et al., 2012). At 0.005 % level of

concentration, this biosurfactant, which is among the strongest, reduces surface energy from 72.8 to 27.9 mN/m

(Nguyen and Sabatini 2023).

Surfactin: One of the most notable biosurfactants is surfactin, a cyclic lipopeptide manufactured by Bacillus

subtilis. It comprises a fatty acid chain linked to a cyclic structure of 7 amino acids through lactone bonding.

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Even in small concentrations as low as 0.005 %, it effectively reduces surface energy from 72 to 27.9 mN/m

(Singh and Cameotra, 2004).

Lichenysin: The bacterium Bacillus licheniformis generates a range of biosurfactants known for their

remarkable balance in pH, temperature, and salt tolerance, working synergistically with each other. Their

physical, chemical and structural characteristics are likewise comparable to those of surfactin. The surface

energy of water can be lowered to 27 mN/m and the tension between it and its interface of water and n-

hexadecane to 0.36 mN/m by the surfactants that are generated by B. licheniformis (Coronel-León et al., 2015).

Neutral lipids, phospholipids and fatty acids: Asignificant quantity of phospholipid surfactants and fatty acids

are produced by several bacteria and yeast during their growth phase on n-alkanes. The length of the hydrocarbon

chain in each structure determines the equilibrium between affinity for water and affinity for lipids (HLB), which

has directly proportionality. Phosphatidylethanolamine-abundant sacs that create optically transparent alkane

solubilized oil are produced by Acinetobacter sp. Evidence has shown that the tension between the interface

of hexadecane and water is lowered by phosphatidylethanolamine produced by R. erythropolis cultured on n-

alkane, to a level below 1 mN/m, and an essential associated colloidal system concentration (Jorge et al., 2018).

Polymeric biosurfactants: Alasan, liposan, lipomanan emulsan and a few additional protein–

polysaccharide interactions

effective extracellular polyanionic

are

the

most

researched

heteropolysaccharide

polymeric

biosurfactants.

is produced

The

by

amphipathics

bioemulsifier

Acinetobacter calcoaceticus RAG-1. The molecules of hydrocarbon in water can be effectively emisfied by

emulsan, regardless of minimal concentrations ranging from as 0.001 to 0.01% (Desai and Banat, 1997).

Candida lipolytica produces liposan, an extracellular dispersible emulsifier that is 83% carbohydrate and 17%

protein (Danyelle et al., 2016).

Particulate biosurfactants: Microbial cells' absorption of alkanes is greatly aided by the microemulsion that is

created when hydrocarbons are partitioned by extrinsic membrane compartments. Acinetobacter sp.

compartments have a density of 1.158 cubic g/cm buoyancy, 20–50nm diameter, and a composition of

lipopolysaccharide endotoxin, Phosphatidic acids, and protein (Makula et al., 1975).

Characteristics of Biosurfactants

Due to the widening range of compounds that are becoming obtainable, biosurfactants are becoming more

appealing for application commercially. When chemically juxtaposed with the inorganic, biosurfactants possess

a number of advantageous superiorities. The following list includes a synopsis of each of the primary

characteristics that set biosurfactants apart:

Surface and Frontier Activity

The tension in the interfacial space between water and hexadecane can be reduced from 40 to 1 mN/m while the

surface energy of water from 72 to 35 mN/m by using a suitable surfactant. The B. subtilis-derived surfactin may

lower the tension in the interfacial space of water and hexadecane to less than 1 mN/m and the surface energy

of water to 25 mN/m (Gudiña, 2012). P. aeruginosa rhamnolipids reduce water's surface energy to 26 mN/m and

the tension in the interfacial space of water and hexadecane to less than 1 mN/m (Mendes et al., 2015). The

tension across the interfacial space drops to 5 mN/m and the surface energy drops to 33 mN/m by T. bombicola's

sophorolipids (Pakshirajan and Daverey 2010). Surfactants chemically produced takes a greater volume to

achieve the highest reduction in surface energy; in contrast, biosurfactants generally has a greater degree of

effectiveness, with a CMC that is roughly 10–40 times less (Sajadi et al., 2024).

Temperature, Ph and Ionic Strength Tolerance

The temperature and pH levels of the surrounding environment have little effect on the surface activity of a

variety of surfactants of biological origin. Evidence from the study carried out by Coronel-León et al. (2015)

showed that the quantities of Ca (50g/l) and NaC (25 g/l), and temperature of up to 50⁰C, had no effect on the

lichenysin derived B. licheniformis. After subjecting to autoclave at 121°C and subsequently at six months at

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18⁰C for 20 min, a lipopeptide derived from B. subtilis remained stable; Its surface activity remained consistent

within a pH range of 5 to 11, and its effectiveness was unaffected by NaCl concentrations up to 20 %.

Biodegradability

Microbially derived surfactants, in contrast to chemical surfactants, are readily broken down and primarily used

in ecological activities including oil spill dispersal and bio-remediation processes (Banat et al., 2014).

Low Toxicity

Since they are typically regarded as safe by-product, biosurfactants can be used in food, beauty products and

drugs production. Evidence from research proposed that a chemically synthesized anionic surfactant (Corexit)

compared to Photobacterium phosphoreum derived rhamnolipids resulted in 50 % death of sample population

(LC50), which is 10 times higher than rhamnolipids, indicating the higher lethality of the chemically synthesized

surfactant (Nash et al., 2014). The toxicological properties of six microbially derived surfactants, four chemical-

based surfactants, and two industrial dispersing agents were evaluated. It became apparent that the majority

of the microbially derived surfactants broke down more quickly, with the exception of a chemical-

based emulsifier or sucrose-stearate, which broke down more quickly than biological derived glycolipids

with displayed structural identity to glycolipids (Bhardwaj and Sharma, 2013). With respect to lethality and

mutation causing capabilities, a chemically derived surfactant with frequent usage in industrial processes was

juxtaposed with a biological surfactant derived from P. aeruginosa (Cooper and Cavalero, 2003).

According to Vijayakumar and Saravanan (2015), surfactant derived from biological sources was found to

be marginally non-toxic and non-mutagenic, while the surfactant derived from chemical sources showed

mutagenicity and lethality of a greater degree in the two tests. It is possible to create persistent emulsifiers that

degrade and form composites emulsion that persist for long period of time running into years. Surfactants of

microbial origin can operate as emulsifiers or destabilizers, depending on the state of the emulsion.

Overall, emulsifiers with higher molecular weight have superior performance than those with lower molecular

weight surfactants of biological origin (Mnif and Ghribi, 2015). Although T. bombicola-produced sophorolipids

can lower interfacial and surface energy, they are not effective emulsifiers (Cooper and Cavalero, 2003). Liposan,

on the other hand, has been effectively employed in the emulsification of edible oils indicating inability to

decrease surface energy (Paximada et al., 2021). The fact that surfactants of polymer source cover tiny beads of

oil to create steady emulsions gives them extra advantageous superiority: a characteristic that finds applicability

in Making oil/water emulsions for food production and beauty products.

Chemical heterogeneity

Chemical heterogeneity refers to the diverse and complex molecular structures present in biosurfactants, which

are surface-active compounds synthesized by various microorganisms, including bacteria, fungi, and yeasts (Dini

et al., 2024). This structural diversity arises from variations in the hydrophilic and hydrophobic moieties of

biosurfactant molecules, such as differences in fatty acid chain lengths, sugar residues, amino acids or peptide

structures. These molecular variations significantly influence the physicochemical properties of biosurfactants,

including their surface tension reduction, emulsification capacity and critical micelle concentration. The

chemical heterogeneity of biosurfactants is a key factor that enables them to function effectively under a wide

range of environmental conditions and interact with a variety of substrates. Consequently, this feature enhances

their applicability in numerous fields such as bioremediation, enhanced oil recovery, pharmaceuticals, cosmetics

and food processing (Dini et al., 2024). The ability to tailor biosurfactant properties by leveraging their inherent

chemical diversity makes them a valuable alternative to synthetic surfactants in both sustainable and specialized

applications.

Economical and Highly Promising Substrates

Like most biotechnology processes, the primary hurdle in the manufacture of biological surfactants is operational

cost. The quantity of the input/raw resources and the type of resources can frequently have a significant impact

on operational costs; as with other biotechnological processes, input/raw resources are thought to be responsible

for between 10% and 30% of overall operational costs. The use of inexpensive input/raw resources to produce

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the appropriate biological surfactants is therefore preferable in order to lower the operational cost. A possible

approach that has been thoroughly investigated is the use of inexpensive input/raw resources of agricultural

source as substrates to produce biological surfactant. Production of biosurfactants can be aided by low-cost

input/raw resources from sources such as oils obtained from plants and oil wastes. Numerous studies involving

oils obtained from plants have demonstrated their potential as affordable and efficient input/raw resources for

the synthesis of biosurfactants (Nitschke et al., 2005).

factors affecting the production of biosurfactant

Composition of the growth

The composition of the growth, which include the type of carbon, nitrogen sources, and carbon itself, all affect

the generation of biological surfactant in addition to the genetic variant of the producer. The total volume of

biological surfactant derived as well as the type of bio-polymer formed are influenced by the nitrogen proportion,

nutritional supplements constraints, and physical-chemical parameters like temperature, pH, aeration, divalent

cations and salt (Ilori et al., 2005).

Carbon sources for biosurfactant production: Many research investigations have employed a wide range of

carbon sources to produce biological surfactant. Evidence suggesting a good supply of carbon substrate for the

synthesis of biological surfactants includes; crude oil, diesel, glucose, sucrose, and glycerol, exist in the literature

(Fagade et al., 2009). Though its significance varies depending on the microorganism, it is clear that carbon

substrate is essential to the derivation of biological surfactants. For example, in the case of Pseudomonas sp. the

chemical makeup of the biological surfactant production was influenced by the various carbon sources in the

medium, but the overall chain length of the fatty acid component parts in glycolipid was unaffected by the chain

length of the substrate (Sari, 2019).

Nitrogen sources for biosurfactant production: A supply of nitrogen is necessary for the synthesis of

biological surfactants. Nitrogen-containing medium is crucial for the growth of microorganisms since it is

necessary for the synthesis of proteins and enzymes. Biological surfactants have been produced using a variety

of nitrogen-rich sources, including meat extract, yeast extract, ammonium sulphate, ammonium nitrate, and

sodium nitrate (Sari, 2018). P. aeruginosa produces biological surfactants mostly from nitrate-rich sources

(Pacwa-Płociniczak et al., 2011); ammonium salts and urea are favoured as source of nutrient for Arthrobacter

paraffineus (Pacwa-Płociniczak et al., 2011). Monosodium glutamate (MSG) otherwise known as yeast extract

is a rather common source of nitrogen utilized for the synthesis of surfactants of biological origin. However, the

amount of this nitrogen depends on the microbe and the medium used for cultivation. According to a report,

during the stationary phase of cell growth, surface-active chemicals are frequently produced when the culture

medium's nitrogen supply is reduced (Wu et al., 2011).

Environmental factors: The total yield of the biological surfactant derived is highly dependent on

environmental conditions. The biological surfactant production process must be optimized to derive substantial

volume because variations in temperature, pH, air circulation speed, might have an impact on the final result

(Saharan et al., 2011). Although it is claimed that the vast majority of biological surfactant manufactures are

carried out in the 25–30 °C temperature range, this range of temperatures modified the chemical makeup of the

biological surfactant generated in A. paraffineus and Pseudomonas sp. variant DSM2874 (Pacwa-Płociniczak et

al., 2011). Zinjarde and Pant (2002) investigated the impact of pH on the amount of biological surfactant

generated and found that optimal yield happened at pH 8.0, corresponding to the normal pH of seawater, which

is the native ecosystem of Y. lipolytica. The generation of rhamnolipids by Pseudomonas sp. peaked at the pH

range of 6 to 6.5 and declined precipitously above pH 7. Stirring and air circulation play a significant role in the

synthesis of biological surfactants by facilitating oxygenation between the gaseous phase to the aqueous phase.

Evidence in the literature suggests that bio-emulsifier or stabilizers synthesis can improve the dissolution of

nutrients that are hydrophobic, hence facilitating the delivery of nutrients to microbial organisms, inferring that

the biological significance of microbiological emulsifiers or stabilizers could be attributable to their ability to

dissolve and deliver hydrophobic nutrients (Alizadeh-Sani et al., 2018). The synthesis of biological surfactants

is also significantly influenced by the salt concentration of a given medium or substrate. However, the results

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for certain biological surfactant derivatives showed that they were unaffected by salt levels as high as 10 %

(weight/volume), despite having indicated that the CMC was slightly reduced (Mahmoud et al., 2010).

Applications of Biosurfactants

Although, chemical synthesis typically used to create all surfactants before now. Interestingly, because of their

wide spectrum of advantageous characteristics, and the various ways that microbes can synthesize them,

biological surfactants have received a lot of research focus lately (Velioglu and Urek, 2016). The fact that they

are less hazardous than chemically manufactured surfactants and readily biodegrade makes them

environmentally acceptable, which is an especially important factor. Because of these special qualities, biological

surfactants can be used in many industrial processes as substitutes for surfactants that are chemically

manufactured (Zhao et al., 2017). In addition, they can be used in biological remediation and treatment of

wastewater, and are harmless on the environment. Removing heavy metals from polluted soil, hydrocarbon in

water bodies, hexa-chloro cyclohexane decomposition, microbial enhanced oil recovery, and hydrocarbon

decomposition in polluted soil are a few possible uses of biological surfactants in contamination control (Kumar

and Mandal, 2017).

Ingredients in Food Formulations

Evidence that exists in the literature has shown that surfactants have the capacity for lowering surface energy

and tension between surfaces, which makes emulsion derivation and stabilizing effect easier. Additionally,

surfactants can serve a number of purposes in the manufacture of food (Nitschke and Costa, 2007). In this regard,

to alter the viscoelasticity features of whole-wheat dough, stabilizing air-circulating systems, enhancing the

whole-wheat and preservation period of yields of starch content, preventing accumulation of tiny aggregates of

fat and enhancing the consistency and surface texture of calorie-dense food products (Wang et al., 2024).

Biological surfactants are stabilizers that help keep fats and oils from retrogradation while also allowing the

flavor oils to be dissolved and regulation in ice cream and pastry recipes (Zhao et al., 2017). Research

investigation showed that it is possible to enhance the size, surface texture, uniformity of dough, and preservation

of baked food products by including rhamnolipid surfactants (Wang et al., 2024). In addition, rhamnolipids may

be used to enhance the qualities of buttery cream and frosted food products (Wang et al., 2024). Significant

opportunity exists for L-rhamnose as a precursor to flavoring. Similar to furaneol, it is currently employed in

industry as an intermediate for premium flavoring ingredients (Roscher et al., 1997).

Adhesion-prevention agents

Acolony of bacteria on a surface is referred to as a biofilm. In addition to the bacteria, the biofilm also comprises

of any extrinsic material generated at the surface and any material substance confined in the structural matrix

that has developed (Flemming and Wingender 2010). According to Srey et al. (2013), bacterial biofilms found

on surfaces in the food manufacturing sector are probable causes of exposure that could cause food to deteriorate

and spread illness (Srey et al., (2013). Preventing bacteria from adhering to surfaces that come in touch with

food is therefore, crucial to giving customers high-quality and healthy food products. Research have shown how

biological surfactants affect microbial adherence to and disengagement from surfaces (Gudiña et al., 2013).

Streptococcus thermophilus produces a biosurfactant that has antimicrobial properties, including the ability to

inhibit the growth of various thermophilic strains of Streptococcus. These strains are commonly used as fat and

oil emulsifiers but may also contribute to the formation of foul odours. The biosurfactant from S. thermophilus

is currently being applied in industrial settings, such as in pasteurizers, to reduce offensive odors by preventing

bacterial spatter on heat-exchanger plates (Thando et al., 2017) One novel approach to lessen adhesion has been

proposed: the bio-treatment of surfaces via the application of surfactants of microbial origin (Thando et al.,

2017).

Therapeutic and biomedical applications and antimicrobial activity

Biological surfactants have shown microbicidal activity against various bacteria, algae, fungi, and viruses in

numerous studies. Significant antimycotic action was demonstrated by the lipopeptide iturin derived from

Bacillus subtilis (Rahman et al., 2007). It has been shown that 80 mM surfactin inactivates enveloped viruses,

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including herpes and retrovRhamnolipids, at concentrations ranging from 0.4 to 10.0 mg/l, were found to hinder

colony formation in Heterosigma akashivo and Protocentrum dentatum, both known toxic bloom algae (Wang

et al., 2005). Furthermore, a rhamnolipid mixture derived from Pseudomonas aeruginosa exhibited antifungal

properties against Chaetonium globosum, Penicillium crysogenum and Aspergillus niger (at 16 mg/ml),

Aureobasidium pullulants (at 32 mg/ml), as well as against the phytopathogens Botrytis cinerea and Rhizoctonia

solani, with significant effectiveness (Goswami et al., 2015). The rhamnolipid mixture exhibited antimicrobial

activity against various bacterial species, including Escherichia coli, Micrococcus luteus and Serratia

marcescens. It was also effective at concentrations of 32 mg/ml against Alcaligenes faecalis, 16 mg/ml against

Mycobacterium phlei and 8 mg/ml against Staphylococcus epidermidis (Banat et al., 2010). Potent antimycotic

medium targeting plant and seed fungal pathogens were discovered to be rhamnolipids and sophorolipids (Chen

et al., 2020). Glycolipid surfactant mannosylerythritolcations lipid (MEL) derived from Candida antartica

demonstrated significant microbicidal action against Gram positive bacteria such as Staphylococci, streptococci

and some listeria species (Kitamoto et al., 1993).

Anticancer activity

The impacts of 7 extracellular glycolipids synthesized by microorganisms, including mannosyl erythritol lipids-

A, mannosyl erythritol lipids-B, rhamnolipid, polyol lipid, sophorose lipid, and others, have been investigated.

With the exception of rhamnolipid, all these glycolipids were found to induce cell differentiation rather than cell

growth in the HL60 human promyelocytic leukemia cell line (Isoda et al., 1997). Specifically, sophorolipid and

mannosylerythritol lipid notably promoted characteristic differentiation features in monocytes and granulocytes,

respectively (Cameotra and Makkar, 2004). B16 cells exposed to MEL underwent chromatin condensation, DNA

cleavage and sub-G1 arrest (Kitamoto et al., 1993). This is the first demonstration that glycolipids can cause

growth inhibition, cell death, and differentiation in malignant mouse melanoma cells (Briem et al., 1999).

Furthermore, MEL enhanced the activity of acetylcholine esterase and arrested the cell cycle at the G1 phase in

PC12 cells, resulting in neurite outgrowth and partial cellular differentiation (Isoda et al., 2000). MEL, a

glycolipid surfactant sourced from Candida antarctica, additionally demonstrated antimicrobial effects against

Gram-positive bacteria (Kitamoto et al., 1993).

Anti-Human Immunodeficiency Virus and Sperm Immobilizing Activity

The significant occurrence of HIV/AIDS among women aged 15–49 years has underscored the necessity for a

safe, efficient, and female-controlled vaginal microbicide. In response to this concern, researchers have

investigated sophorolipid derived from C. bombicola and its derivatives for their potential to eliminate sperm,

HIV and vaginal cells (Shah et al., 2005). The sophorolipid diacetate ethyl ester derivative is the strongest

spermicide and virucide of the sophorolipids examined. It has comparable effects to nonoxynol in inactivating

HIV and human semen (Shah et al., 2005). Nevertheless, it induced considerable damage to vaginal cells, raising

doubts about its appropriateness for prolonged microbicidal contraceptive use (Shah et al., 2005).

Agents for respiratory failure

Breathing difficulties in premature infants result from the absence of pulmonary surfactant, a combination of

phospholipids and proteins (Nkadi et al., 2009). To address this, the genes responsible for producing surfactant

proteins can be extracted and inserted into bacteria, allowing for the proteins to be produced through fermentation

for medical use (Shah et al., 2005).

Agents that enhance the activity of skin fibroblast cells

Sophorolipids in lactone form contain a large amount of diacetyl lactones that can boost the metabolism of skin

dermal fibroblast cells and the formation of new collagen, at a concentration of 0.01 parts per million (ppm) at

5 %) (w/w) of dry matter in formulation (Borzeix and Concaix 2003). This finds utility in both the fields of

cosmetology and dermatology. The purified lactone sophorolipid compound holds significance in formulating

anti-aging dermal products due to its ability to stimulate dermal cell activity (Borzeix and Concaix 2003). By

encouraging the generation of fresh collagen fibers, purified lactone sophorolipids can help combat skin aging,

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and are utilized in various skincare products such as body creams, milks, lotions and gels (Borzeix and Concaix

2003).

Surgical antiadhesive agents: Surfactants derived from S. thermophilus, when applied to silicone rubber,

effectively prevented 85 % of C. albicans adhesion (Busscher et al., 1997). Similarly, surfactants obtained from

L. fermentum and L. acidophilus, when applied to glass surfaces, reduced the attachment of uropathogenic

Enterococcus faecalis cells by 77% (Velraeds et al., 1996). Additionally, the biosurfactant sourced from L.

fermentum inhibited S. aureus infection and adhered to surgical implants (Gan et al., 2009). Furthermore,

surfactin demonstrated a reduction in biofilm formation by Salmonella typhimurium, S. enterica, Escherichia

coli and Proteus mirabilis on PVC plates and vinyl urethral catheters (Rodrigues et al., 2006).

Palm Oil Mill Effluents

POME is recognized as one of the most environmentally harmful agro-industrial wastes, mainly because of its

high organic content. Among the most polluting agro-industrial wastes due to its high organic content. From

sterilization and clarification stages, palm oil mill effluent emerges as a dark brown, highly concentrated

colloidal mixture of water, oil, and fine cellulose materials (Madaki and Lau 2011). POME constitutes a colloidal

solution containing approximately 0.6 - 0.7 % oil, 95 – 96 % water and 4 – 5 % total solids (Ma, 2000). Oil palm

production in Nigeria witnessed an increase of 0.8 million tonnes from 1990 to 2001, reaching 9 million metric

tonnes (FAO, 2002). Of this production, approximately 43 – 45 % always remains as mill waste, comprising

Empty Fruit Bunches (EFB), Shell, Fibre and Palm Oil Mill Effluent (POME) (Madaki and Lau, 2011). As

production increases, these residues will continue to accumulate. Initiatives are underway to convert these waste

materials into valuable resources for energy generation, animal feed, and organic fertilizers. The oil extraction

procedure requires significant water usage for steam sterilization of palm fruit bunches and oil clarification. The

resulting wastewater sludge, termed palm oil mill effluent, is a brown sludge containing approximately 4 – 5 %

solids (predominantly organic matter), 0.5 – 1 % residual oil, and around 95 % water, with a high concentration

of organic nitrogen (Onyia et al., 2001). This effluent is a severe land and water pollutant when released directly

into the environment. In addition to lipids and volatile compounds, the adverse impacts of palm oil mill effluent

on living tissues may be attributed to water-soluble phenolic compounds (Radzia 2001, Perez et al., 1992). The

presence of ammonia in the effluent is undesirable as it contributes to high oxygen demand in water bodies.

Although palm oil mill effluent is a pollutant for the palm oil industry, it has great potential for improving animal

feed and soil quality (Binder et al., 2002). The quality of the raw material and the palm oil production processes

in the mills affect the characteristics of palm oil mill effluent. There are three main processing steps that result

in the POME according to Sethupathi (2004). The sterilization process of fresh fruit bunches (FFB), clarification

of crude palm oil (CPO), and the hydrocyclone separation of cracked kernel and shell mixture together account

for around 36%, 60 % and 4 % of palm oil mill effluent (POME) respectively within the oil mills. According to

Yacob et al. 2006), it is approximated that for every tonne of fresh fruit bunch processed, approximately 0.5 to

0.75 tonnes of palm oil mill effluent (POME) will be generated (Yacob et al., 2006).

Attributes of Palm Oil Mill Effluent (Pome)

There is a lot of waste produced by the palm oil mill industry. POME primarily originates from oil extraction,

washing, and purification processes within the mill. It comprises various components including cellulose

material, fats, oils, and greases (Agamuthu, 1995). Additionally, POME is laden with solids, encompassing both

suspended and dissolved particles, with concentrations ranging from 18,000 mg/L to 40,500 mg/L. These solids

are called palm oil mill sludges (POMS). Newly generated effluents, according to Ma, (2000), is a warm, acidic

liquid with a pH ranging from 4 to 5. It presents as a brownish colloidal suspension characterized by elevated

levels of organic matter, substantial concentrations inclusive of COD (50,000 mg/L), total solids (40,500 mg/L),

BOD (25,000 mg/L), and oil and grease (4,000 mg/L). Untreated or partially treated palm oil mill effluent

(POME) is characterized by a substantial concentration of readily degradable organic substances. Since no

chemicals are incorporated during the oil extraction process, POME is considered non-toxic. Nevertheless, in its

untreated state, POME plays a considerable role in aquatic pollution by diminishing dissolved oxygen levels in

water bodies (Khalid and Wan Mustafa, 1992). Conversely, it also contains significant amounts of essential

nutrients such as nitrogen (N), magnesium (Mg), phosphorus (P), potassium (K), and calcium (Ca) (Habib et al.,

1997; Muhrizal et al., 2006), which are crucial for plant growth and development. Because of its lack of toxicity

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and its potential as a fertilizer or alternative animal feed, POME can supply vital mineral nutrients. Additionally,

Agamuthu, (1995) observed that the elevated organic nitrogen content in POME enhances its efficacy as a

fertilizer.

According to Muhrizal, M., et al. (2006), POME has a higher aluminium (Al) concentration than composted

sawdust and chicken dung. Habib et al., (1997) stated that while lead (Pb) concentrations in palm oil mill effluent

(POME) typically remain above sub-lethal levels (more than 17.5 μg/g) (James et al., 1996), POME may contain

dangerous elements. James et al., (1996) explain that paints and glazing materials containing lead can

contaminate plastic or metal pipes, tanks, and containers used in palm oil processing, which may result in lead

leaching into the palm oil mill effluent (POME) (Okewole and Omin (2013).

Extraction of Crude Palm Oil

Harvested from oil palms, fresh fruit bunches (FFB) are processed in palm oil mills to produce crude palm oil

and palm kernel. These mills, which are typically found inside plantations, allow fresh fruit bunches (FFB) to

be moved about and processed. The primary process of palm oil milling primarily encompasses the physical

extraction of palm products (Hii et al., 2012).

The extraction of crude palm oil from FFB involves several processing stages. Sterilization is the initial step,

wherein freshly harvested fruit bunches are subjected to high-pressure steam (120 to 140⁰C at 40 psi or 275790

N/m²) promptly upon arrival at the mill. This procedure deactivates lipolytic enzymes accountable for oil

hydrolysis and fruit degradation, simultaneously averting the formation of free fatty acids, and priming the fruit

bunches for subsequent sub-processes (Igwe and Onyegbado, 2006). Bunch stripping follows, mechanically

separating fruits from bunch stalks. The sterilized and separated fruits then undergo digestion, achieved by

reheating them with steam to 80 - 90⁰C. This stage aids in oil extraction by rupturing oil-containing cells in the

mesocarp and separating the mesocarp from the nuts. The final stages involve oil extraction, clarification, and

purification, where crude oil is extracted from the digested fruit mash using a screw press without damaging the

kernel.

Initially, palm bunches are cut into quarters and left overnight to facilitate the separation of nuts from the spikelet.

The fruits are then boiled for 1 - 1.5 hours, crushed in a mortar or mashed with feet in a canoe-like container,

and water is added and thoroughly mixed. Subsequently, all nuts are meticulously removed by hand. The fibers

are vigorously shaken in the sludge until oily foams emerge on the surface. The foam is carefully collected in a

container until no more foam formation occurs. The collected foam is subsequently boiled for about 30 to 40

mins. The sludge sinks to the bottom, while the clean edible oil rises to the top. Occasionally, the oil extracted

from the sludge pit is reclaimed and blended with fiber to produce a combustible mixture known as a fire starter

cake, commonly referred to as flint. The sludge and the liquid waste, which is known as palm oil effluent, are

sometimes thrown on the plants and soil around them (Wu et al., 2007).

After extraction, the screw press separates the liquid from the nuts. However, the oil contains varying amounts

of water, solids, and impurities that must be eliminated. Fiber particles are removed from the crude oil using a

vibrating screen, while sand and dirt settle out. Water removal is accomplished through settling, centrifugation,

and vacuum drying processes. The clarified crude oil retains approximately 0.1 - 0.25 % moisture, which aids in

oxidative stability and prevents the formation of minute amounts of soluble solids known as gums (Shaaban et

al., 2004). The final product, crude palm oil, is either used locally or refined further (Igwe, 2006). Gunawan et

al. (2009) reported that approximately 22 kg of palm fruit oil and 1.6 kg of palm kernel oil can be extracted from

every 100 kg of fruit bunches. However, significant quantities of palm oil residues or pollutants are also

generated simultaneously, potentially leading to severe environmental pollution (Hii et al., 2012).

Effect of Palm Oil Mill Effluent on the Soil and River Quality

The cultivation and processing of oil palm, like many other agricultural and industrial activities, also contribute

to environmental problems. During oil processing, significant water volumes are utilized within mills for

extracting oil from palm fruits. Roughly 50 % of this water is transformed into palm oil mill effluent during

extraction. Estimates suggest that for every 1 tonne of crude palm oil produced, approximately 5 - 7.5 tonnes of

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water result in POME (Okwute and Isu (2007). In Nigeria, oil palm processing is a widespread practice among

many small-scale operators. In Nigeria’s palm oil industry, little or no treatment is done for most of the palm oil

mill effluent from small-scale traditional operators, and it is usually disposed off in the environment. Okwute

and Isu (2007) in their study reported the potential for the palm oil mill effluent to pollute nearby streams, rivers,

or surrounding land. As a result, river water often becomes brownish, emits unpleasant odors, and develops a

slimy texture. This phenomenon leads to the death of fishes and other aquatic life, depriving local communities

of access to clean water for household needs and fishing (Ezemonye et al., 2008). When discharged untreated

into local rivers or lakes, palm oil mill effluent (POME) emerges as a significant contributor to inland water

pollution. Comprising of lignocellulosic wastes containing a blend of carbohydrates and oil, POME exhibits

remarkably high levels of BOD and COD (Madaki and Lau, 2011). It is not uncommon for COD values to exceed

80,000 mg/L, a figure often associated with palm oil from the nut extracted insufficiently, which can significantly

elevate COD levels (Oswal et al.,, 2002). The aquatic life is being disrupted by the higher COD value

(Maygaonkar et al., 2012).

The application of palm oil mill effluent to soil can yield several advantageous soil chemical and physical

enhancements, including augmented levels of organic matter, organic carbon, major nutrients such as nitrogen

and phosphorus, as well as improvements in water-holding capacity and porosity (Okwute et al., 2023).

However, it also causes undesirable changes such as decreases in pH, and increase in salinity etc. (Onyia et al.,

2001). These effects occur very slowly and need many years to provide significant results. Soil microbiological

and biochemical properties have been recognized as precocious and responsive markers of soil modifications,

enabling the prediction of long-term trends in soil quality (Ros et al., 2003). POME is rich in organic content,

contains significant quantities of plant nutrients, and serves as a cost-effective source of these nutrients when

subjected to fermentation processes (Onyia et al., 2001). The detrimental impact of POME may be attributed to

phenols and other acids that are organic in nature and have phytotoxic and anti bacterial properties (Pascual et

al., 2007). Over time, the polyphenolic fraction breaks down and partly changes into humic substances. There is

limited understanding regarding the influence of POME on soil properties, particularly concerning biochemistry

and microbiology. Research indicates that the effects of waste application to soil are predominantly observed

during the initial weeks following the amendment (Binder et al., 2002).

The composition and quality of oil mill effluent vary depending on factors such as seasonal changes, raw material

standards, and current operational conditions. Typically, palm oil mill wastewater exhibits low pH levels, around

4-5, due to the presence of organic acids generated during fermentation. It additionally holds notably high total

solids (40,500 mg/L), as well as oil and grease (4000 mg/L) (Ma, 2000). Moreover, the wastewater comprises

dissolved components, such as elevated levels of proteins, carbohydrates, nitrogenous compounds, lipids, and

minerals. These substances have the potential to be transformed into valuable materials through microbial

activities (Alvionita et al., 2019).

The release of untreated effluents from palm oil mills can present significant environmental concerns (Singh et

al., 2010). Therefore, resolving the challenge of converting palm oil mill effluent (POME) into a sustainable

waste requires the implementation of efficient treatment and proper disposal methods.

Economic Importance of Palm Oil

Palm fruit oil ranks among the two most significant vegetable oils in the global oil and fats market, second only

to soybeans. The oil palm (Elaeis guineensis) stands as the most productive oil-producing plant globally, with

one hectare yielding between 10 and 35 tonnes of fresh fruit bunch (FFB) annually (Ma et al., 1996). Even

though palm trees can thrive for more than 200 years, their economic viability usually lasts between 20 to 25

years. The nursery phase typically spans 11 to 15 months, with the first harvest occurring 32 to 38 months after

planting, and peak yield reached 5 to 10 years post-planting (Igwe and Onyegbado, 2007). Harvested fruit

bunches yield oil extracted from the fleshy mesocarp, constituting at least 45 – 46 % of the total yield, while the

kernel comprises approximately 40-50 %. The nutrient needs of the palm tree fluctuate considerably, mainly

dictated by the genetic composition of the planting material and impacted by environmental factors like water

availability, sunlight exposure, and temperature (Igwe and Onyegbado, 2007).

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Processes and Microorganisms Involved in Pome Treatment

The biological ponding system has witnessed substantial adoption as a prevalent treatment approach for palm

oil mill effluent. Studies suggest that more than 85 % of palm oil mills exclusively utilize the biological ponding

system for effluent treatment (Najafpour et al., 2006). The typical components of this system include deoiling

ponds, anaerobic ponds, facultative ponds, and aerobic ponds (Mohammad et al., 2021). Successful operation of

the ponding system requires extended retention times of over 20 days, with biogas released into the atmosphere.

Yacob et al. (2006) reported that an average of 36 % methane gas is emitted into the atmosphere from open tank

digesters. Similarly, Shirai et al. (2003) discovered that methane gas production from open tank digesters and

lagoon systems is approximately 35 % and 45 %, respectively. A range of methods of treatment for the

wastewater from palm oil mills were examined in order to meet strict rules on discharge into watercourses.

According to Ahmad et al. (2003), treating the palm oil mill effluent with membrane technology in conjunction

with physical-chemical pretreatment resulted in significant reductions in turbidity, COD, and BOD, with

reductions of up to 100 %, 98.8 % and 99.4 %, respectively.

The effectiveness of various anaerobic treatment systems has been demonstrated through multiple studies. The

two-stage up-flow anaerobic sludge blanket system, as reported by Borja et al., (1996), can handle COD loading

rates of up to 30 g COD/L/day, resulting in over 90% methane yield and COD reduction. Similarly, single-stage

anaerobic tank digesters and anaerobic ponding systems, according to Ugoji (1997), achieve COD removal

efficiencies exceeding 94 % after a 10-day retention time. Research by Borja and Banks (1995) showcases COD

removal rates surpassing 90 % in both anaerobic filters and anaerobic fluidized bed reactors with an input rate

of 10 g COD/L/day. Najafpour et al. (2005) reported COD removal rates of up to 88 % with a hydraulic retention

time of 55 hours using attached growth on a rotating biological contactor. Additionally, Oswal et al. (2002)

achieved a 95 % reduction in COD through treatment with tropical marine yeast within a retention time of 2

days. Anaerobic digestion systems are increasingly utilized in wastewater treatment, particularly within the agro-

industry, due to their advantages over aerobic treatments. These benefits include the production of less waste

sludge, reduced energy requirements, and simpler restart procedures following extended shutdowns (Beccari et

al., 1996). The possibility of producing methane, a by-product of biogas, adds to the method's appeal. Laboratory

investigations show that the final result of anaerobic digestion of palm oil mill effluent is a biogas combination

with 65 % CH₄, 35 % CO₂and traces of H₂S, according to Yacob et al. (2005). It is anticipated that one tonne

of palm oil mill effluent can yield about 28 m³of biogas.

Prokaryotes Involved in Pome Degradation

This study reviews two domains of prokaryotic organisms: eubacteria and archaeabacteria, also known as

"ancient" bacteria. Both eubacteria and archaeabacteria are unicellular organisms, but archaeabacteria have

distinct cellular chemistry. Overall, these prokaryotes play crucial roles in biological wastewater treatment

processes. Archaeabacteria groups include halophiles, methanogens, and thermacidophiles (Gerardi, 2006).

Anaerobic Digestion

Suspended particles from dry plant matter and oil palm fruit debris make up the majority of organic components

found in raw palm oil mill effluent (POME). The first stage of POME breakdown is anaerobic digestion, which

entails removing bulk waste via a number of procedures. Anaerobic digestion is the process by which

biodegradable components in wastewater are biologically converted, in the absence of oxygen, into carbon

dioxide and methane (CH4) (Lam and Lee, 2011). The procedures encompass the breakdown of carbon

compounds through hydrolysis, fermentation, acetate formation, and methane production, facilitated by diverse

symbiotic microorganisms. The rich organic composition of POME, including cellulose, lignin, and residual oil,

creates favorable conditions for hydrolytic bacteria. These bacteria secrete extracellular enzymes like cellulase,

xylanase, and lipase to degrade carbon polymers into simpler compounds, thereby initiating the anaerobic

digestion of POME (Hassan et al., 2005). The outcomes of hydrolysis, such as monosaccharides, fatty acids, and

amino acids (from triglycerides), act as the materials for acidogenesis or fermentation in the subsequent phase.

During fermentation or anaerobic respiration, acidogenic bacteria break down carbohydrates and fatty acids into

simpler organic acids, such as lactic, propionic, and butyric acids, along with the production of hydrogen gas

(Chong et al., 2009). This is why they are called acidogenic bacteria. The organic acids are then transformed

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into acetate by acetogenic bacteria. Acetogenic bacteria frequently engage in syntrophic relationships with

certain methanogens, which utilize hydrogen gas to produce methane. Acetogenic bacteria and methanogens

engage in a mutualistic relationship. Methanogens reduce the hydrogen partial pressure, facilitating the oxidation

of organic acids into acetate, while relying on acetogenic bacteria to supply hydrogen gas for methanogenesis

(Ahmad et al., 2011). Additionally, certain acetogenic bacteria can reduce sulfate and utilize it as an electron

acceptor to produce sulfide gas (Wong et al., 2014). Finally, methanogens use the byproducts to produce

methane, which completes the transformation of organic matter into biogas (Slonczewski and Foster 2014).

Methanogens, classified as archaea, are typically categorized into two groups: acetotrophic and

hydrogenotrophic methanogens, distinguished by their respective substrates for methanogenesis (Demirel and

Scherer, 2008). Hydrogenotrophic methanogens utilize hydrogen gas as an electron acceptor during

methanogenesis. In contrast, acetotrophic methanogens convert acetate into methane (Demirel and Scherer

2008).

Nitrification, Denitrification, and Phosphorus Accumulation

Numerous studies have highlighted the nutrient richness of POME, including nitrogen and phosphorus content

(Chowdhury et al., 2007). Important steps in the breakdown of POME include nitrification, denitrification, and

phosphorus buildup, which remove phosphorus and inorganic nitrogen from wastewater.

Different nitrifiers carry out the oxidation of ammonium or ammonia to nitrate in two steps. Nitrosomonas sp.

are the bacteria that catalyze the first step, which is the conversion of ammonia to nitrite. They do this by

producing enzymes known as ammonia monooxygenase and hydroxylamine oxidoreductase (Hommes et al.,

2001). Nitrobacter bacteria are responsible for converting nitrite into nitrate using the enzyme nitrite

oxidoreductase (Bartosch et al., 1999). Denitrification occurs as denitrifying organisms reduce nitrate to nitrite,

then to nitrogen gas. This process involves the enzyme nitrate reductase and utilizes nitrate or nitrite as an

electron acceptor to generate energy, releasing nitrogen gas into the atmosphere (Daum et al., 1998). Meanwhile,

phosphorus removal from POME is facilitated by phosphorus-accumulating bacteria, which absorb excess

orthophosphate in the wastewater and store it within their cells. The removal of biomass from the wastewater

also eliminates the accumulated phosphorus (Bao et al., 2017).

Eukaryotes Involved in Pome Degradation

Fungi, algae, protozoa, and animals (rotifers, worms – nematodes and flatworms) are some of the eukaryotic

organisms that take part in POME treatment processes. Soil and water organisms infiltrate wastewater treatment

plants via inflow and infiltration pathways (Gerardi, 2006). Fungi or yeast and algae are the two eukaryotic

organisms that this review examines. They can be isolated from the POME, which is a liquid waste stream from

palm oil mills. By secreting extracellular enzymes, most fungi from the POME can break down lignocellulose

and lipids, which are complex polymers. Fungi play a vital role in breaking down lipids, not solely through the

action of the enzyme lipase, but also by secreting biosurfactants, as seen in certain species like Candida sp. (Kim

et al., 1999). These biosurfactants reduce surface tension and interfacial tension between water and lipid phases,

aiding in lipid degradation. Geotrichium candidum, for instance, can hydrolyze phenols and produce peroxidase

enzymes capable of breaking down various color dyes (Coulibaly et al., 2003). Similarly, Aspergillus fumigatus

demonstrates colour removal from POME, albeit through bioadsorption (Neoh et al., 2012). Additionally,

Chlorella pyrenoidosa and Chlorella vulgaris, two algae species isolated from POME, are involved in nitrogen

and phosphorus removal from the wastewater. Chlorella sp. rapidly takes up nitrogen and phosphorus from

POME for their growth and proliferation (Safi et al., 2014). These nutrients are used to build up phospholipids

and glycolipids which make up approximately 30% of their weight dry biomass (Lam and Lee, 2011).

Anaerobic Digestion Process

Anaerobic digestion emerges as a highly effective treatment approach for palm oil mill effluent (POME). In this

process, a diverse community of microorganisms orchestrates a series of complex biochemical reactions to

degrade organic matter, resulting in the production of methane and carbon dioxide (Borja et al., 1995).Achieving

stability and efficiency in this process relies on various factors, including reactor configurations, hydraulic

retention time, organic loading rates, pH levels, temperature, inhibitor concentrations, total volatile fatty acid

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(TVFA) levels, and substrate composition (Fikri et al., 2020). Thorough investigation and meticulous control of

these parameters are essential to prevent process failures or reduced efficiency, aiming to maintain them at or

close to optimal conditions.

These anaerobic digestions are usually carried out at mesophilic (30 – 37⁰C) or thermophilic (50 – 60⁰C)

temperatures. According to Najafpour et al. (2006), the effluent from palm oil milling is released at a high

temperature of roughly 90⁰C, which prepares the ground for the POME treatment at either mesophilic or

thermophilic temperatures. In a semicontinuous anaerobic reactor operating in a mesophilic environment, Cail

and Barford (1985) tested palm oil mill effluent, obtaining an approximate 75 % removal rate of chemical oxygen

demand (COD) with an organic loading rate (OLR) of 12.6 g[COD]/L/day and a 5.6-day hydraulic retention

period.

Similarly, employing a similar reactor design but operating under thermophilic conditions with a maximum OLR

of 15.1 g[COD]/L/day and a hydraulic retention period of 4.3 days, Padilla and Banks (1993) achieved an 85 %

removal of COD and a methane output of 295 ml/g[COD].

In comparison to running at 37⁰C, Yu et al. (2002) found that operating at 55⁰C resulted in a greater substrate

degradation rate, biogas generation rate, and specific rate of aqueous product creation. According to research by

De la Rubia et al. (2002), a reactor with OLRs of up to 2.19 kg m-3 d-3 COD and an operating temperature of

55⁰C produced more gas than one running at 35ºC.

Furthermore, distillery waste digested at anaerobic digestion temperatures of 35 – 55⁰C produced the highest

amount of methane and total biogas at a digester temperature of 50⁰C, according to Banerjee and Biswas, 2004).

These results show that, depending on the temperature, anaerobic bacteria can produce more or less methane

from organic waste. In actuality, if temperature rises are not controlled, biomass washout may occur, leading to

an accumulation of total volatile fatty acids (Lau and Fang, 1997).

Anaerobic microbes help this multi-stage process happen in the absence of oxygen. Having been in use for

almost a century, methanogenic anaerobic digestion of organic waste has a number of benefits over aerobic

treatment techniques, such as high rates of organic waste removal, low energy needs, less sludge creation, and

energy production (Choorit and Wisarnwan, 2007).

Each of the four main phases of anaerobic digestion hydrolysis, fermentation, acetogenesis, and methanogenesis

involves a different population of microbes. Hydrolytic bacteria convert polymeric organic molecules into

soluble monomers, such as glucose, fatty acids, and amino acids, during the hydrolysis stage (Menzel et al.,

2020). This procedure, which is essential for high levels of organic waste, could eventually become rate-limiting.

After hydrolyzed products are transformed by acid-forming bacteria into alcohols, aldehydes, ketones, ammonia,

carbon dioxide, water, and hydrogen, fermentation takes place. The result is the formation of organic acids such

as valeric acid, propionic acid, butyric acid and acetic acid (Zhang et al., 2020). However, methanogens are

unable to directly use volatile fatty acids with chains longer than four carbons (Wang et al., 1999).

In the acetogenesis stage that follows, obligatory hydrogen-producing acetogenic bacteria oxidize organic acids

to acetic acid and hydrogen. During acetogenesis, carbon dioxide and hydrogen are also used to produce acetate.

Acidogenesis and acetogenesis can occasionally coexist in a single stage (Aydin et al., 2017). Lastly, there are

two methods leading to the production of acetotrophic species convert acetate to carbon dioxide and methane,

and hydrogenotrophic organisms reduce carbon dioxide with hydrogen (Demirel et al., 2008).

Common methanogens found in biogas reactors comprise Methanobacterium, Methanothermobacter,

Methanobrevibacter, Methanosarcina, and Methanosaeta (formerly known as Methanothrix) (Sekiguchi et al.,

2001).

Single Phase and Two-Phase Arrangement

Acidification and methanogenesis in the typical anaerobic digestion process take place in a single reactor system,

or single-stage arrangement. However, because of their distinct physiologies, nutritional needs, development

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rates, and susceptibilities to environmental stimuli, acidogens and methanogens in such a system are difficult to

keep in balance (Demirel and Yenigün, 2002).

Anaerobic Sequencing Batch Reactor (Asbr)

This system was manufactured as a solution for effectively managing effluents with high suspended solids

content. Operating within a single reactor, the ASBR follows a four-step cycle (Angenent et al., 2004). Initially,

wastewater containing settled biomass is introduced into the reactor during the feeding stage. Subsequently, the

wastewater and biomass undergo intermittent mixing during the reaction process. Following this, the biomass

settles, and finally, the treated effluent is withdrawn from the reactor (Kannan and Singaram 2012).

According to Ratusznei et al. (2000), the ASBR system has a number of benefits, including better retention of

solids, effective operational management, high organic matter removal efficiency, ease of use, and the lack of a

settling tank.

Up-Flow Anaerobic Sludge Blanket (Uasb)

The Up-flow Anaerobic Sludge Blanket (UASB) reactor is a widely used system for anaerobic wastewater

treatment, applied in about 60 % of full-scale anaerobic treatment facilities globally (Angenent et al., 2004). In

this design, wastewater flows upward through a dense bed of anaerobic sludge granules, where microorganisms

break down organic matter and produce biogas.

The system’s efficiency largely depends on effective sludge retention. This is achieved through bacterial

entrapment within or between sludge particles, as well as bacterial immobilization via natural mechanisms like

biofilm formation and microbial aggregation within the sludge matrix (Lettinga, 1995).

MATERIALS AND METHODS

Study Area/Sample Collection

Samples Were Collected Aseptically Using a Calibrated Pipette from the Top, Middle and Bottom Layers (Each

5cmApart) From Palm Oil Effluents. Sampling Was Conducted at Both Large-Scale (Okomu Oil Palm Company

and The Nigerian Institute for Oil Palm Research) And Small-Scale (Ovbiogie, Sapele Road and aduwawa oil

Mills) Palm Oil Mill Effluents In Edo State. The Collected Samples Were Then Transported Under Sterile

Conditions to The Microbiology Laboratory at The University of Benin, Benin City, For Microbiological

Analysis.

Preparation and Sterilization af Culture Media

The preparation of all culture media (Nutrient Agar, Tryptone Soya Agar and Potato Dextrose Agar) adhered

strictly to the guidelines provided by the manufacturer. Sterilization procedures were conducted at 121⁰C at 15

psi pressure for a duration of 15 mins.

Isolation and Enumeration of Bacteria/Fungi from Samples

A volume of 30.0 ml of the palm oil mill effluent was transferred under sterile conditions into a conical flask

containing 270.0 milliliters of sterile distilled water. Subsequently, a tenfold serial dilution was conducted. An

aliquot of 0.1 ml from the 10^-3 dilution tube was plated onto Nutrient Agar, Tryptone Soya Agar, and Potato

Dextrose Agar (for fungal count). Each sample was inoculated in triplicate. The plates containing nutrient agar

and Tryptone Soya Agar were then placed in an incubator at 37⁰C for 24 hours, while the Potato Dextrose Agar

plates were incubated at 28⁰C for up to 5 days. Colony counts were determined for each plate, and the mean for

each sample was calculated using the formula described in equation (1) provided by Willey et al., (2008),

indicating the mean colony forming unit (cfu) and spore forming units (sfu) per milliliter:

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푛푢푚푏푒푟 표푓 푐표푙표푛푖푒푠 푥 푑푖푙푢푡푖표푛 푓푎푐푡표푟

푐푓푢/푚푙 =

푣표푙푢푚푒 표푓 푖푛표푐푢푙푢푚

Morphological and Biochemical Characteristics of Bacteria

Gram stain

Slides were prepared for each isolate by making smears and heat-fixing them on clean, grease-free slides. Crystal

violet, the primary stain, was applied to each smear for one minute, followed by rinsing with distilled water.

Subsequently, the smears were submerged in iodine solution for approximately one minute. After rinsing the

glass slide with distilled water, decolorization was carried out using a 95 % alcohol solution for 30 sec, followed

by another wash with distilled water. Counterstaining of the smears on the slides was performed using Safranin

solution for one minute. Finally, the slides were rinsed with distilled water, allowed to air dry, and examined

under a microscope at 1000x total magnification (Willey et al., 2008).

Motility /Test

Selected isolates were introduced into the medium (Motility Test Medium) using a sterile needle, which was

inserted approximately halfway into the medium. The tubes were then left uncovered and subjected to incubation

at 37⁰C for 18 to 24 hrs. Fuzzy growth extending from the point of inoculation signifies the organism's motility,

while growth confined strictly within the stab line signified non motility.

Biochemical Test

Catalase test

The purpose of this test is to determine the presence or absence of the catalase enzyme. Catalase facilitates the

decomposition of hydrogen peroxide into oxygen gas and water. A few drops of freshly prepared 3 % hydrogen

peroxide were added to the bacterial isolates smeared on a slide. The production of gas bubbles indicates a

positive result for the catalase enzyme (Olutiola et al., 1991).

2 H₂O₂

2 H₂O + O₂

Oxidase Test

Adiluted 1% solution of oxidase reagent, prepared according to standard protocols, was utilized. Asmall amount

of culture obtained from Nutrient Agar plate using a sterilized platinum wire loop was smeared onto a moistened

filter paper with an oxidase reagent. The appearance of a purple coloration indicates a positive result for the

oxidase test (Cheesbrough, 2006).

Coagulase Test

Numerous microorganisms, including Staphylococcus aureus, produce an enzyme known as coagulase. This

enzyme facilitates blood clotting by converting fibrinogen into fibrin. In the slide test method, a clean slide was

divided into two sections. On one section, a small amount of the test organism was emulsified in a drop of water

using a sterile wire loop; the other section contained only water and served as a negative control. A drop of

human plasma was added to both sections, and the slide was gently rocked for a few minutes. Agglutination

(clumping) observed only on the section with the test organism indicates a positive result for coagulase

production. The control section showed no agglutination, confirming the validity of the test.

Urease test

The isolates were introduced into slants of urea medium and placed in an upright position, then incubated at

37⁰C for 24-48 hours. Cultures testing positive for urease produced a red-pink coloration due to alterations in

the indicator's colour (Cheesbrough, 2005).

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NH₂CO.NH₂+ H₂O

2NH₃+ CO₂

Indole Test

The indole test was conducted to determine the isolates capable of converting tryptophan to indole. This test is

commonly employed to aid in distinguishing Gram-negative bacilli, particularly those belonging to the

Enterobacteriaceae family. Peptone water was prepared, and approximately 3 ml of it was dispensed into Bijou

tubes using a sterile pipette. Sterile loops were used to collect a well-isolated colony of bacteria, which was then

inoculated into the Bijou tubes. Subsequently, the tubes were incubated at 37⁰C for 48 hours. Following the

incubation period, 0.5 ml of Kovac’s Indole Reagent was added to each inoculated Bijou tube. The tubes were

gently shaken and observed for the appearance of a red coloration in the surface layer within 10 mins.

(Chessbrough, 2006). A red ring on the top of the tube indicated a positive indole reaction.

Citrate Utilization Test

The characteristic use of citrate as the only source of carbon, by some organisms, forms the basis of this test.

The process involved the inoculation of the test organism in a medium which contained Simon’s citrate, in a test

tube. The incubation temperature was set at 37⁰C for 24 to 48hours. The presence of a deep blue colour after

incubation indicated a positive result (Chessbrough, 2006).

Sugar Fermentation Test

The isolates in the test medium were tested whether they could, alongside the production of acid or gas or only

acid, ferment a sugar molecule. This test is based on the fact that most bacteria especially those of the Gram-

negative strain, use a variety of sugar as carbon sources and energy, and are also able to produce either acid and

gas or acid. Basically, this test serves as functionality test that helps to distinguish one bacteria strain from

another. Peptone water prepared in conical flask containing the indicator phenol red, was the growth medium

utilized in this study. Specialized tubes called Durhams tubes for the mixture were sterilized by an autoclave for

about 15 mins and at a temperature of 121⁰C. After preparing and sterilizing a 1 % sugar solution at 121⁰C for

approximately 10 mins, 5 ml of the solution was aseptically dispensed into tubes containing peptone solution

and indicators (phenol red). Subsequently, the tubes were inoculated with a young culture (fresh bacterial culture)

of the isolated organism and then incubated at approximately 37⁰C. After a 24-hour incubation period, the

production of acid and gas, or only acid, was observed. A change in the colour of the medium from light green

to yellow indicated acid production, while the presence of gas in the Durham tube indicated gas production

(Chessbrough, 2006).

Identification of Fungal Isolates

The identification of fungal isolates was carried out using the Lactophenol Cotton Blue (LPCB) staining

technique and microscopic examination. Aclean glass slide was prepared by placing a drop of lactophenol cotton

blue stain in the center using a sterilized needle or dropper. A small portion of mycelium was carefully taken

from the fungal culture using the same sterilized needle and transferred into the drop of stain on the slide.

Using the needle, the mycelium was gently spread out in the stain to ensure even distribution. A cover slip was

then placed over the preparation, applying light pressure to eliminate air bubbles and secure the sample for

viewing.

The slide was examined under a compound microscope, starting with the x10 objective lens for general

orientation and then under the x40 objective lens for detailed observation.

Observation: Under The Microscope, Fungal Structures Such as Hyphae, Conidiophores, Spores and Other

Morphological Features Were Observed. These Structures Were Compared with Standard Mycological

References to Identify the Fungal Species Based On Their Shape, Arrangement and Reproductive Structures.

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Molecular Identification

DNA Extraction

DNA was extracted using the protocol described by (Sambrook and Russell, 2011). Briefly, single colonies

grown on medium were transferred to 1.5ml of liquid medium, and cultures were grown on a shaker for 48 hours

at 28⁰C. After this period, cultures were centrifuged at 4600g for 5 mins. The resulting pellets were resuspended

in 520 µl of TE buffer (10 mM Tris-HCl, 1 mM EDTA, pH 8.0). Fifteen microliters of 20% SDS and 3µl of

Proteinase K (20 mg/ml) were then added. The mixture was incubated for 1 hour at 37⁰C, followed by the

addition of 100 µl of 5 M NaCl and 80µl of a 10 % CTAB solution in 0.7 M NaCl, and vortexed. The suspension

was incubated for 10 mins at 65⁰C and kept on ice for 15 mins. An equal volume of chloroform:isoamyl alcohol

(24:1) was added, followed by incubation on ice for 5 mins and centrifugation at 7200g for 20 min. The aqueous

phase was then transferred to a new tube, and isopropanol (1:0.6) was added to precipitate the DNA at –20⁰C for

16 hours. DNA was collected by centrifugation at 13000g for 10 mins, washed with 500µl of 70 % ethanol, air-

dried at room temperature for approximately three hours, and finally dissolved in 50 µl of TE buffer

Polymerase Chain Reaction (PCR)

The PCR sequencing preparation cocktail consisted of 10 µl of 5x GoTaq colourless reaction buffer, 3 µl of 25

mM MgCl₂, 1 µl of 10 mM dNTPs mix, 1 µl of 10 pmol each 27F 5’- AGA GTT TGA TCM TGG CTC AG-3’

and 1525R 5′-AAGGAGGTGATCCAGCC-3′ primers and 0.3 units of Taq DNA polymerase (Promega, USA)

made up to 42 µl with sterile distilled water, plus 8 µl of DNA template. PCR was carried out in a GeneAmp

9700 PCR SystemThermalcycler (Applied Biosystems Inc., USA) with the following profile: initial denaturation

at 94⁰C for 5 mins; followed by 30 cycles consisting of 94⁰C for 30 sec, 50°C for 60 sec and 72⁰C for 1 min 30

sec; and a final termination at 72⁰C for 10 mins, then held at 4⁰C.

Gel Electrophoresis

The integrity of the amplified approximately 1.5 Mb gene fragment was checked on a 1 % agarose gel to confirm

amplification. The buffer (1xTAE buffer) was prepared and subsequently used to prepare a 1.5 % agarose gel.

The suspension was boiled in a microwave for 5 mins. The molten agarose was allowed to cool to 60⁰C and

stained with 3 µl of 0.5 g/mL ethidium bromide (which absorbs invisible UV light and transmits the energy as

visible orange light). A comb was inserted into the slots of the casting tray and the molten agarose was poured

into the tray. The gel was allowed to solidify for 20 mins to form the wells.

The 1×TAE buffer was poured into the gel tank to barely submerge the gel. Two microliters of 10x blue gel

loading dye were added to 4 µl of each PCR product and loaded into the wells after the 100bp DNA ladder was

loaded into well 1. The gel was electrophoresed at 120V for 45 mins visualized by ultraviolet trans-illumination

and photographed. The sizes of the PCR products were estimated by comparison with the mobility of a 100bp

molecular weight ladder that was run alongside experimental samples in the gel.

Purification of Amplified Product

After gel integrity, the amplified fragments were ethanol-purified to remove the PCR reagents. Briefly, 7.6 µl of

3M Na acetate and 240 µl of 95 % ethanol were added to each approximately 40 µl PCR amplified product in a

new sterile 1.5ml Eppendorf tube, mixed thoroughly by vortexing, and kept at –20⁰C for at least 30 mins.

Centrifugation for 10 mins at 13000g and 4⁰C followed, with removal of the supernatant.

The pellet was washed by adding 150 µl of 70 % ethanol, mixed, then centrifuged for 15 mins at 7500g and 4⁰C.

Again, the supernatant was removed, and the tube was inverted on paper tissue to let it dry in the fume hood at

room temperature for 10–15 mins. The pellet was then resuspended with 20 µl of sterile distilled water and kept

at –20⁰C prior to sequencing. The purified fragment was checked on a 1.5 % agarose gel run at 110V for about

1 hour, as previously described, to confirm the presence of the purified product and quantified using a nanodrop

of 2000 spectrophotometer (Thermo Scientific).

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Sequencing Identification

All PCR products were purified with Exo sap and sent to Epoch Life science (USA) for Sanger sequencing.

Sequencing were identified using Gen Bank’s Basic Local Alignment Search Tool (BLAST) algorithm on

National Centre for Biotechnology and Information website. The corresponding sequences were identified using

the online blast search at (http://blast.ncbi.nlm.nih.gov/Blast.cgi).

Highly corresponding sequence were

recovered from NCBI and subjected to multiple sequence alignment using Bio edit software.

Blood Hemolysin Production

This qualitative screening method was employed to preliminarily assess the potential for biosurfactant

production by the organisms, utilizing blood agar as the culture medium. In this study, spot-inoculation of single

colonies of isolates was performed on agar plates containing blood, followed by incubation for approximately

48 hours at 37⁰C.

Following the incubation period, clear zones indicating the rupture of blood cells (hemolysis) were observed

around the colonies. Absence of hemolysis indicated gamma hemolysis, while partial hemolysis indicated alpha

hemolysis, and complete hemolysis indicated beta hemolysis of the blood culture medium (Satpute et al., 2010).

The forms of hemolysis were differentiated based on the appearance of zones surrounding the colonies on blood

agar plates after 48 hours of incubation at 37⁰C:

-

Beta (β) hemolysis: Characterized by a clear, transparent zone around the colony, indicating complete

lysis of red blood cells.

Alpha (α) hemolysis: Identified by a greenish or brownish discolouration around the colony, due to partial

lysis and oxidation of hemoglobin.

Gamma (γ) hemolysis: Indicated by no change in the medium around the colony, suggesting no hemolytic

activity.

This visual assessment was done under normal lighting conditions on the incubated blood agar plates.

Screening of Biosurfactant Producing Bacteria

The various source materials for inoculation were standardized to an optical density of OD₆₀₀(= 1.0) and derived

from selected bacterial cultures in nutrient broth. Approximately 5ml of the test bacterial cultures were

transferred into 100 ml of a nutritionally rich solution composed of 2 % (w/v) glucose in MSM in 500 ml

Erlenmeyer flasks. This solution served as the carbon source and was then incubated at 35⁰C with shaking at 200

rpm.

Assay for Biosurfactant Activity Via Oil Displacement/ Spread Assay

This step involved introducing the surfactant-containing medium (cell culture supernatant) to the interface

between oil and water. The test involved introducing a cell-free culture supernatant obtained by centrifuging

microbial cultures grown in production medium at 8,000 rpm for 10 mins onto the interface of oil and water. The

purpose was to observe the diameter of the clear zone formed as a result.

This was accomplished by adding 100 µl of kerosene to the surface of a Petri dish containing 15 ml of distilled

water. Subsequently, the oil surface was inoculated with 20 µl of cell culture supernatant. The presence of an

emulsified clear zone around the colonies served as a positive indication of biosurfactant production (Satpute et

al., 2010).

Data Analysis

Results were presented as mean of 3 replications. Analysis of variance were determined using the 1-way ANOVA.

Duncan’s multiple range tests were employed to assess mean differences (P<0.05) (Ogbeibu, 2014).

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RESULTS

Table 4.1 shows the total heterotrophic bacterial counts (THBC) observed in the POME. The THBC of POME

derived from NIFOR ranged from 4.10 ± 1.39 to 4.34 ± 0.32 (Log10 cfu/ml). Similarly, bacterial counts of

POME samples from Okomu Oil spanned from 4.06 ± 0.88 to 4.66 ± 0.83 (Log10 cfu/ml), whereas POME from

Aduwawa exhibited total bacterial counts varying between 4.02 ± 1.00 and 4.52 ± 1.34 (Log10 cfu/ml).

Moreover, Ovbiogie samples exhibited a range of 3.93 ± 1.09 to 4.33 ± 1.02 (Log10 cfu/ml), while Sapele Road

samples ranged from 3.90 ± 1.49 to 4.18 ± 0.88 (Log10 cfu/ml). Noteworthy is that POME samples from Okomo

Oil displayed the highest heterotrophic bacterial count (4.66 ± 0.83 Log10 cfu/ml), while those from Ovbiogie

exhibited the lowest heterotrophic bacterial count (3.93 ± 1.09 Log10 cfu/ml). Statistical analysis revealed

significant differences in bacterial density (P < 0.05) among the top, middle, and bottom POME samples

collected from different locations.

The heterotrophic fungal count (THFC) of Palm Oil Mill Effluents is shown in Table 4.2. The heterotrophic

fungal count of Palm Oil Mill Effluents (POME) from Aduwawa ranged from 4.34 ± 0.76 and 3.93 ± 1.70 Log₁₀

sfu/ml, fungal counts of POME samples from NIFOR ranged 4.11 ± 1.57 to 4.29 ± 0.24, POME samples from

Okomu oil ranged from 3.85 ± 0.23 to 4.26 ± 0.62 Log₁₀sfu/ml, while Sapele road samples ranged from 3.78 ±

0.58 to 4.08 ± 0.58 Log₁₀sfu/ml, Ovbiogie samples ranged from 3.74 ± 0.00 to 3.98 ± 0.26 Log₁₀sfu/ml. POME

samples from Aduwawa had both the highest and least heterotrophic fungal count of 4.34 ± 0.76 and 3.93 ± 1.70

Log₁₀sfu/ml respectively. Top POME samples from NIFOR, Okomu Oil, Aduwawa and Sapele Road were not

different significantly, while middle and bottom of POME samples from NIFOR, Aduwawa and Sapele Road

were different significantly (P<0.05) from Okomu Oil and Ovbiogie POME samples.

Table 4.1: The Heterotrophic bacterial counts (THBC) of Palm Oil Mill Effluents

Sample

depths

NIFOR

(Log₁₀

Cfu/ml)

Okomu Oil

Aduwawa

(Log₁₀

Cfu/ml)

Ovbiogie

Sapele

(Log₁₀Cfu/ml)

Road

(Log₁₀

Cfu/ml)

(Log₁₀

Cfu/ml)

Top

4.34±0.32^a

4.28±0.51^a

4.10 ±1.39^b

4.62±0.79^a

4.66±0.83^b

4.06±0.88^b

4.51±0.71^a

4.52±1.34^b

4.02±1.00^b

4.33 ± 1.02^a

4.31 ± 1.10^b

3.93 ± 1.09^b

4.18 ± 0.88^a

3.95 ± 1.16^b

3.90 ± 1.49^b

Middle

Bottom

Key: NIFOR: Nigeria Institute for Oil Palm Research. Values are presented as mean ± SEM; n=3. Mean values

with similar superscripts within a column across sampling depths are not significantly different, P>0.05.

Table 4.2: The Heterotrophic Fungal count (THFC) of Palm Oil Mill Effluents

Sample depths

NIFOR (Log₁₀Okomu Oil

Aduwawa

Ovbiogie

Sapele

Road

sfu/ml)

(Log₁₀sfu/ml)

3.98 ± 0.26^b

3.93 ± 0.33^b

3.74 ± 0.00^b

Top

4.29±0.24^a

4.23±0.55^a

4.11±1.57^a

4.26±0.62^a

3.98±0.90^b

3.85±0.23^b

4.34±0.76^a

4.23±1.43^a

3.93±1.70^a

4.08 ± 0.58^a

4.02 ± 0.33^a

3.78 ± 0.58^a

Middle

Bottom

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Key: NIFOR: Nigeria Institute for Oil Palm Research, Values are presented as mean ± SEM; n=3. Mean values

with similar superscripts within a column across sampling depths are not significantly different, P>0.05.

Table 4.3 shows the percentage bacterial and fungi reduction in small scale enterprise across the various depths

in each location. Percentage bacterial reduction of samples from Sapele Road was 72.29 %, Aduwawa was 67.69

% and Ovbiogie was 60.47 %. Percentage fungal reduction of samples from Sapele Road was 61.36 %, Aduwawa

was 42.11 % and Ovbiogie was 50.0 %. Samples from Sapele Road had highest bacterial and fungal reduction

(72.29 % and 61.36 %) respectively while samples from Ovbiogie and Aduwawa respectively had least bacterial

(60.47 %) and fungal (42.11 %) reduction respectively.

The percentage bacterial and fungi reduction in large scale enterprise is shown in Table 4.4. Percentage bacterial

reduction of samples from Sapele Road and Okomo oil were 43.18 % and 72.29 % respectively while percentage

fungal reduction of samples from Sapele Road and Okomo oil were 33.33 % and 61.11 % respectively with

Okomu oil samples having the highest bacterial (72.29 %) and fungal reduction (61.11 %) and NIFOR recorded

the least bacterial (43.18 %) and fungal reduction (33.33 %).

Table 4.5 shows the molecular identification of bacteriaL isolates obtained from palm oil mill effluents. Bacterial

identified were Bacillus cereus, Pseudomonas aeruginosa, Bacillus amyloliqefaciens, Escherichia coli and

Klebsiella aerogenes.

The distribution of bacterial isolates in large scale palm oil enterprise (LSE) is shown in Table 4.6. Pseudomonas

aeruginosa, Bacillus amyloliqefaciens, Bacillus cereus, Escherichia coli and Klebsiella aerogenes were present

in Okomu Oil (top) samples. Pseudomonas aeruginosa and Bacillus amyloliqefaciens were absent in NIFOR

bottom, Okomu middle and Okomu bottom samples.

Table 4.7 shows the distribution of bacterial isolates in small and medium scale palm oil enterprise (SME).

Bacillus amyloliqefaciens, Escherichia coli and Klebsiella aerogenes were absent in in Aduwawa and Ovbiogie

(middle and bottom), Sapele Road (bottom) samples while Bacillus amyloliqefaciens was present in Aduwawa

(bottom) and Ovbiogie (top and bottom) samples.

Table 4.3: Percentage bacterial and fungi reduction in small scale enterprise

Sample source

Sapele Road

Aduwawa

Bacteria (%)

72.29

Fungal (%)

61.36

67.69

42.11

Ovbiogie

60.47

50.0

The values above represent the average percentage reduction in bacterial and fungal counts from Palm Oil Mill

Effluent (POME), calculated across three sampling depths (top, middle and bottom) in small and medium scale

enterprises.

Table 4.4: Percentage bacterial and fungi reduction in large scale enterprise

Sample source

NIFOR

Bacteria (%)

43.18

Fungal (%)

33.33

Okomu

72.29

61.11

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The values above represent the average percentage reduction in bacterial and fungal counts from Palm Oil Mill

Effluent (POME), calculated across three sampling depths (top, middle and bottom) in large scale enterprises.

Table 4.5: Molecular identification of bacterial isolates obtained from Palm Oil Mill Effluents

Sample Code

LSE01

Bacterial identity

Bacillus cereus

Query cover (%)

99.0

Percent identity (%)

99.80

Accession No.

CP053954.1

Pseudomonas

aeruginosa

SME02

SME03

100.00

99.00

100.00

99.93

MK875779.1

CP054415.1

Bacillus

amyloliqefaciens

LSE04

LSE06

Escherichia coli

100.00

MK371829.1

CP048598.1

Klebsiella aerogenes

Key:

LSE01:

SME02:

SME03:

LSE04:

LSE06:

Bacillus cereus

Pseudomonas aeruginosa

Bacillus amyloliqefaciens

Escherichia coli

Klebsiella aerogenes

Plate 4.1 Agarose gel of 16s rRNA bacterial amplification of bacterial obtained from Palm Oil Effluents. Bands

A, B, C, D and E indicates the genomic DNAs of Bacillus cereus, Pseudomonas aeruginosa, Bacillus

amyloliqefaciens, Escherichia coli and Klebsiella aerogenes. "Con" indicates the control lane.

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Table 4.6: Distribution of bacterial isolates in large Scale Palm Oil Enterprise (LSE)

Sample depths

LSE 1 (NIFOR) Top

+

-

+

-

LSE 1(NIFOR) Middle

LSE 1 (NIFOR) Bottom

LSE 2 (OKOMU) Top

LSE 2 (OKOMU) Middle

LSE 2 (OKOMU) Bottom

+

-

+

-

+

-

+

-

+

-

+

-

+

Keys: + = present, - = absent

Top, middle and bottom indicates point of sample collection in the palm oil mill effluent

LSE 1 and LSE 2 represent the sampling depths.

Table 4.7: Distribution of bacterial isolates in Small and Medium Scale Palm Oil Enterprise (SME)

Sample depths

SME 1 (Aduwawa) Top

-

+

-

+

-

SME 1 (Aduwawa) Middle

-

+

SME 1 (Aduwawa) Bottom

-

+

-

SME 2 (Ovbiogie) Top

+

-

+

-

SME 2 (Ovbiogie) Middle

-

SME 2 (Ovbiogie) Bottom

-

+

-

SME 3 (Sapele Rd) Top

-

+

-

SME 3 (Sapele Rd) Middle

-

+

-

SME 3 (Sapele Rd) Bottom

-

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Key: + = present, - = absent

Top, middle and bottom indicates point of sample collection in the effluent

SME 1, SME 2 and SME 3 represent the sampling depths.

The distribution of fungal isolates in large scale palm oil enterprise (LSE) is shown in Table 4.8. Penicillium

chrysogenum and Penicillium citrinum were present in NIFOR (Bottom) samples and absent in NIFOR (Middle),

Okomu (Top and Middle) samples. Aspergillus niger, Fusarium solani, Penicillium chrysogenum, Microsporum

sp., Penicillium citrinum and Aspergillus flavus were absent in NIFOR (Middle) POME samples.

Table 4.9 shows the distribution of fungal isolates in small and medium scale palm oil enterprise (SME).

Aspergillus niger, Fusarium solani, Penicillium chrysogenum and Microsporum sp., were absent in Aduwawa

(Bottom), Ovbiogie (Middle and Bottom) POME samples.

Table 4.10 shows the zones of oil spreading assay of biosurfactant producing microorganisms from palm oil

effluents. Pseudomonas aeruginosa, Bacillus amyloliqefaciens, Bacillus cereus, Aspergillus niger, Fusarium

solani and Penicillium chrysogenum produced positive Biosurfactant and Hemolytic activity at different zone of

inhibitions (12.00mm, 15.00mm, 16.00mm, 14.00mm, 10.00mm and 7.00mm respectively) while Escherichia

coli and Microsporum sp. showed negative Biosurfactant Production at 0.00mm zone of inhibition.

Table 4.11 shows the emulsification activity of biosurfactant-producing microorganisms isolated from Palm Oil

Mill Effluents. The microorganisms were assessed for their emulsification ability at 610 nm, with results

expressed as average values with standard deviations. Bacillus amyloliquefaciens exhibited the highest

emulsification activity with a value of 1.01 ± 2.30, indicating strong biosurfactant production and emulsification

capacity. Bacillus cereus followed with an emulsification activity of 0.91 ± 0.33, reflecting its comparable

potential in biosurfactant production. Pseudomonas aeruginosa showed a slightly lower emulsification activity

(0.644 ± 1.22), suggesting moderate emulsification potential. Klebsiella aerogenes and Escherichia coli

exhibited the lowest emulsification activities, with values of 0.41 ± 0.50 and 0.39 ± 1.33, respectively.

Table 4.8: Distribution of fungal isolates in Large Scale Oil Enterprise (LSE)

Sample depths

LSE 1 (NIFOR) Top

+

-

+

-

LSE 1 (NIFOR) Middle

LSE 1 (NIFOR) Bottom

LSE 2 (Okomu) Top

-

+

-

+

-

+

-

+

-

+

-

LSE 2 (Okomu) Middle

LSE 2 (Okomu) Bottom

-

+

-

+

-

+

Key: + = present, - = absent

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Top, middle and bottom indicates point of sample collection in the effluent

LSE 1 and LSE 2 represents the sampling depths.

Table 4.9: Distribution of fungal isolates in Small and Medium Scale Enterprise (SME)

Sample depths

SME 1 (Aduwawa) Top

SME 1 (Aduwawa) Middle

SME 1 (Aduwawa) Bottom

SME 2 (Ovbiogie) Top

SME 2 (Ovbiogie) Middle

SME 2 (Ovbiogie) Bottom

SME 3 (Sapele Rd) Top

SME 3 (Sapele Rd) Middle

SME 3 (Sapele Rd) Bottom

-

+

-

+

-

+

-

+

-

+

-

+

-

+

-

+

-

+

-

Keys: + = present, - = absent

Top, middle and bottom indicates point of sample collection in the effluent

SME 1, SME 2 and SME 3 represents the sampling depths.

Table 4.10: Zones of Oil Spreading Assay of biosurfactant producing microorganisms from palm oil

effluents

Isolates

Zones (mm)

12.00

Biosurfactant Production

Hemolytic activity

Pseudomonas

aeruginosa

Bacillus

+

15.00

amyloliqefaciens

Klebsiella aerogenes

Escherichia coli

Bacillus cereus

Aspergillus niger

Fusarium solani

Penicillium

7.00

0.00

16.00

14.00

10.00

7.00

+

-

+

-

+

chrysogenum

Microsporum sp.

0.00

-

+

Keys: + = positive, - = negative

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Table 4.11: Emulsification activity of the microorganisms that produces biosurfactants obtained from

Palm Oil Mill Effluents

ISOLATES

Emulsification activity at 610nm

0.644±1.22

Pseudomonas aeruginosa

Bacillus amyloliqefaciens

Bacillus cereus

1.01±2.30

0.91±0.33

Klebsiella aerogenes

Escherichia coli

0.41±0.50

0.39±1.33

Values are presented as mean ± SEM; n=3.

DISCUSSION

The management and disposal of wastewater, particularly from palm oil mills, pose significant environmental

challenges for many villages reliant on small-scale palm oil production. While the practice of disposing untreated

palm oil mill waste has been ongoing in these communities for years, heightened scrutiny from environmental

regulatory bodies has brought attention to its adverse effects. The current study focused on identifying

biosurfactant-producing microorganisms in palm oil mill effluents which revealed levels of both bacterial and

fungal populations. Specifically, samples from Okomu oil exhibited the highest count of heterotrophic bacteria

(4.66±0.83 Log10 cfu/ml). This finding aligns with previous research by Eno et al. (2017), who documented

similarly high levels of total heterotrophic bacteria in palm oil mill effluents. Additionally, studies by Ibe et al.

(2014), Ohimain et al. (2013) and Ohimain et al. (2012) have reported comparable microbial counts in POME

samples, further supporting the findings of this investigation.

The elevated fungal count observed in the samples of POME utilized in this investigation suggests a stimulative

effect of POME on fungal proliferation. This is consistent with prior research indicating that POME harbors

metabolizable nutrients conducive to fungal growth (Nwago and Okolo, 2011). The variability in microbial

populations detected in our study may stem from various factors such as composition of nutrient, mineral content,

oxygen, temperature, acidity and the wastewater volume (Jeremiah et al., 2018). The abundance of bacteria in

POME could be attributed to potential contamination resulting from inadequate sanitation practices within the

mills (Okechalu et al., 2011), as well as the processing methods and environmental conditions prevailing in these

facilities. As noted by Ohimain et al. (2013), POME serves as a favorable habitation for lipolytic and cellulolytic

bacteria and fungi due to its nutrient-rich composition, including lipids and cellulose. The isolation of Aspergillus

niger and Aspergillus flavus from POME samples suggests their potential involvement in the biodegradation of

oily wastewaters, as previously reported in literature. However, further studies involving biodegradation assays

are necessary to confirm their functional role. Additionally, the identification of Penicillium chrysogenum and

Penicillium citrinum in POME aligns with previous research findings (Jeremiah et al., 2018). Similarly, the

isolation of Fusarium solani and Microsporum sp. in POME samples is consistent with the findings of Obire et

al. (2011), who also reported the presence of these fungal species in POME.

The distribution of biosurfactant-producing microorganisms within effluent systems presents significant

implications for both environmental management and microbial ecology. Observations indicate that these

microorganisms are more concentrated at the top of the effluent compared to the bottom, a phenomenon that can

be attributed to several interrelated factors (Wu et al., 2007). One primary reason for this stratification is the

differential availability of nutrients and oxygen in various layers of the effluent. The upper layers typically

receive more sunlight and oxygen diffusion, creating a more favourable environment for aerobic biosurfactant-

producing microorganisms, which thrive in such conditions (Wu et al., 2007). Additionally, hydrodynamic

factors play a crucial role in microbial distribution within effluents (Alvionita et al., 2019). The agitation caused

by inflow currents and turbulence tends to suspend lighter particles and microorganisms at higher levels, leading

to an increased concentration of active biosurfactant producers near the surface. This hydrodynamic behavior

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can foster competitive advantages for these microorganisms as they exploit available organic substrates more

efficiently than their counterparts residing deeper within the effluent column (Borja et al., 1995).

Furthermore, the metabolic activity of biosurfactant-producing microbes contributes to their own proliferation

at higher concentrations. As these organisms metabolize substrates and produce surfactants, they alter local

microenvironments, enhancing their growth potential relative to other microbial populations that may not share

such capabilities (Bajaj and Annapure, 2015). Consequently, this self-enhancing feedback loop further

consolidates their presence at elevated levels within the effluent system.

The molecular identification of bacterial isolates obtained from palm oil effluents revealed the presence of

various species, including Bacillus cereus, Pseudomonas aeruginosa, Bacillus amyloliqefaciens, Klebsiella

aerogenes and Escherichia coli. Fungi like Aspergillus niger, Fusarium solani, Penicillium chrysogenum,

Microsporum sp, Penicillium citrinum and Aspergillus flavus were also isolated from these samples. The

presence of Bacillus species, including Bacillus cereus, in POME samples may suggest their adaptability to

diverse environments, including those with high organic or lipid content (Imo and Ihejirika, 2021). While studies

by Bala et al. (2018) and Mukesh et al. (2012) have reported the lipolytic abilities of Bacillus species, further

assays would be necessary to confirm such activity under the conditions of this study.

Similarly, the isolation of bacteria like Bacillus species, Pseudomonas species, Klebsiella species, Escherichia

coli and fungi including Aspergillus niger, Fusarium solani, Penicillium chrysogenum, Microsporum sp.,

Penicillium citrinum and Aspergillus flavus in POME samples is consistent with studies conducted by Jeremiah

et al. (2014), Ohimain et al. (2012), and Okechalu et al. (2011). Bacillus spp were frequently isolated from

effluent samples across different scales of palm oil enterprises, while Penicillium chrysogenum exhibited the

highest frequency among fungal isolates. This aligns with previous findings by Ohimain et al. (2012) and

Ibegbulam and Achi (2014), which reported similar occurrences of Bacillus, Penicillium, and Aspergillus species

in palm oil effluent samples.

Furthermore, the study revealed that the isolated bacteria and fungi possess the capability to produce

biosurfactants, as evidenced by inhibition zone assays. This finding is in corroboration by Kanokrat et al. (2013),

who illustrated the biosurfactant-producing abilities of certain bacteria, including Pseudomonas sp. isolated from

palm oil-contaminated soils. Biosurfactants plays a very significant role in reducing surface tensions and

facilitating the desorption of POME pollutants from soil, as noted by Bustamante et al. (2012).

The presence of clear zones surrounding colonies on blood agar indicates the potential production of

biosurfactants by selected isolates, as noted by Popoola et al. (2023). Previous studies, such as those of Rajni et

al. (2016), have confirmed the ability to isolate fungal and bacterial strains capable of biosurfactant production

using similar methodologies.

Pseudomonas aeruginosa, as highlighted by Okwute and Ijah (2014), is naturally associated with the degradation

of palm oil and its derivatives, likely due to its capacity to metabolize oil as a carbon source. This species, along

with other Pseudomonas strains, possess the ability for utilizing hydrogen as sources of carbon and energy,

potentially leading to biosurfactant production Okwute et al. (2014).

The existence of Klebsiella aerogenes in POME samples is not unexpected, given its common occurrence in

various environmental sources such as soil, water, and animals. Contamination during oil extraction processes

could introduce Klebsiella aerogenes into POME samples.

This research sheds light on the pathogenic potential of bacterial isolates derived from palm oil mill effluents.

The virulence of pathogenic bacteria, such as Pseudomonas aeruginosa, Bacillus amyloliquefaciens, Escherichia

coli and Bacillus cereus, as well as fungal isolates including Aspergillus niger, Fusarium solani, Penicillium

chrysogenum, and Microsporum sp. is often attributed to their hemolytic activity (Oliveira et al., (2019). This

activity contributes to the pathogens' ability to overcome host defense mechanisms, facilitating colonization and

the establishment of infection (Oliveira et al., (2019).

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Escherichia coli, known as one of the most prevalent human pathogens, exhibited hemolytic activity in this

study. Its potential to cause various infections, from mild to severe, including food poisoning, underscores the

significance of understanding its pathogenic traits, such as its ability to produce hemolysin, as emphasized by

Sora et al. (2021).

The emulsification assay served as an indirect method for screening biosurfactant activity. In this study, Bacillus

amyloliquefaciens exhibited the highest emulsification potential (1.01ꢀ±ꢀ2.30%). Previous studies have

demonstrated the emulsifying capabilities of various microorganisms, especially bacteria, in the treatment of

palm oil mill effluent (POME), as reported by Sebiomo et al. (2011).

Similarly, hemolytic assay methods, as reported by Kawo et al. (2018) and Li et al. (2019), were utilized to

detect clear zones on blood agar plates, indicating biosurfactant production by Bacillus cereus and Aspergillus

niger, respectively. In contrast, Escherichia coli and Microsporum sp. did not exhibit biosurfactant-producing

capabilities in this study. However, Muneer et al., (2014) reported Microsporum sp. as a biosurfactant-producing

fungus, which contrasts with the findings observed in this study. It is essential that biosurfactants be able to

emulsify POME for the uptake and assimilation of hydrocarbons, suggesting that the isolates investigated in this

study hold promise for hydrocarbon degradation and potentially serve as sources for bioremediation of oil-

polluted environments.

Contributions to Knowledge

This study has contributed to knowledge in the following ways;

1. Palm Oil Mill Effluent is a repository for biosurfactant producing bacteria and fungi.

2. Biosurfactant-producing microbes are more concentrated at the top of the effluent compared to the

bottom.

3. Bacillus spp. amongst other isolates possessed higher potential for biosurfactant production.

CONCLUSION AND RECOMMENDATION

Palm oil enterprises in Edo state present a promising opportunity for biosurfactant production, given the

microorganisms found there and their capacity to produce biosurfactants. The presence of these microorganisms

underscores the potential of palm oil mill effluents as significant reservoirs for harnessing biosurfactant-

producing bacteria and fungi. Consequently, these isolates hold importance for synthesizing this valuable

compound, which finds diverse applications across various industries. It is advisable to conduct further research

to refine and optimize the production of biosurfactants by these organisms.

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