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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/IJLTEMAS.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 log
10
3.90±1.00 cfu/g (Small and Medium
Scale- Sapele Road) to log
10
4.66±3.0 cfu/g (Large Scale Enterprise- Okomu). The total fungal counts ranged
from log
10
3.78±1.00 cfu/g (Small and Medium Scale- Aduwawa) to log
10
4.34±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). A general 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: A significant 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 are the most researched polymeric biosurfactants. The
effective extracellular polyanionic amphipathics heteropolysaccharide bioemulsifier is produced by
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 50
0
C, 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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18
0
C 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
A colony 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 140
0
C 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 - 90
0
C. 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 % CH
4
, 35 % CO
2
and traces of H
2
S, according to Yacob et al. (2005). It is anticipated that one tonne
of palm oil mill effluent can yield about 28 m
3
of 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 – 37
0
C) or thermophilic (50 – 60
0
C)
temperatures. According to Najafpour et al. (2006), the effluent from palm oil milling is released at a high
temperature of roughly 90
0
C, 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 37
0
C, Yu et al. (2002) found that operating at 55
0
C 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
55
0
C produced more gas than one running at 35ºC.
Furthermore, distillery waste digested at anaerobic digestion temperatures of 35 – 55
0
C produced the highest
amount of methane and total biogas at a digester temperature of 50
0
C, 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
5cm Apart) 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 121
0
C 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 37
0
C for 24 hours, while the Potato Dextrose Agar
plates were incubated at 28
0
C 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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𝑐𝑓𝑢/𝑚𝑙 =
𝑛𝑢𝑚𝑏𝑒𝑟 𝑜𝑓 𝑐𝑜𝑙𝑜𝑛𝑖𝑒𝑠 𝑥 𝑑𝑖𝑙𝑢𝑡𝑖𝑜𝑛 𝑓𝑎𝑐𝑡𝑜𝑟
𝑣𝑜𝑙𝑢𝑚𝑒 𝑜𝑓 𝑖𝑛𝑜𝑐𝑢𝑙𝑢𝑚
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 37
0
C 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 H
2
O
2
2 H
2
O + O
2
Oxidase Test
A diluted 1% solution of oxidase reagent, prepared according to standard protocols, was utilized. A small 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
37
0
C 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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NH
2
CO.NH
2
+ H
2
O 2NH
3
+ CO
2
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 37
0
C 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 37
0
C 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 121
0
C. After preparing and sterilizing a 1 % sugar solution at 121
0
C 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 37
0
C. 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. A clean 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 28
0
C. 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 37
0
C, 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 65
0
C 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 –20
0
C 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 System Thermalcycler (Applied Biosystems Inc., USA) with the following profile: initial denaturation
at 94
0
C for 5 mins; followed by 30 cycles consisting of 94
0
C for 30 sec, 50°C for 60 sec and 72
0
C for 1 min 30
sec; and a final termination at 72
0
C for 10 mins, then held at 4
0
C.
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 60
0
C 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 –20
0
C for at least 30 mins.
Centrifugation for 10 mins at 13000g and 4
0
C 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 4
0
C.
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 –20
0
C 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 37
0
C.
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 37
0
C:
- 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 OD
600
(= 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 35
0
C 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 Log
10
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 Log
10
sfu/ml, while Sapele road samples ranged from 3.78 ±
0.58 to 4.08 ± 0.58 Log
10
sfu/ml, Ovbiogie samples ranged from 3.74 ± 0.00 to 3.98 ± 0.26 Log
10
sfu/ml. POME
samples from Aduwawa had both the highest and least heterotrophic fungal count of 4.34 ± 0.76 and 3.93 ± 1.70
Log
10
sfu/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
(Log
10
Cfu/ml)
Okomu Oil
(Log
10
Cfu/ml)
Aduwawa
(Log
10
Cfu/ml)
Ovbiogie
(Log
10
Cfu/ml)
Sapele Road
(Log
10
Cfu/ml)
Top
4.34±0.32
a
4.62±0.79
a
4.51±0.71
a
4.33 ± 1.02
a
4.18 ± 0.88
a
Middle
4.28±0.51
a
4.66±0.83
b
4.52±1.34
b
4.31 ± 1.10
b
3.95 ± 1.16
b
Bottom
4.10 ±1.39
b
4.06±0.88
b
4.02±1.00
b
3.93 ± 1.09
b
3.90 ± 1.49
b
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 (Log
10
sfu/ml)
Okomu Oil
(Log
10
sfu/ml)
Aduwawa
(Log
10
sfu/ml)
Ovbiogie
(Log
10
sfu/ml)
Sapele Road
(Log
10
sfu/ml)
Top
4.29±0.24
a
4.26±0.62
a
4.34±0.76
a
3.98 ± 0.26
b
4.08 ± 0.58
a
Middle
4.23±0.55
a
3.98±0.90
b
4.23±1.43
a
3.93 ± 0.33
b
4.02 ± 0.33
a
Bottom
4.11±1.57
a
3.85±0.23
b
3.93±1.70
a
3.74 ± 0.00
b
3.78 ± 0.58
a
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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
Bacteria (%)
Fungal (%)
Sapele Road
72.29
61.36
Aduwawa
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
Bacteria (%)
Fungal (%)
NIFOR
43.18
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
Bacterial identity
Query cover (%)
Percent identity (%)
Accession No.
LSE01
Bacillus cereus
99.0
99.80
CP053954.1
SME02
Pseudomonas
aeruginosa
100.00
100.00
MK875779.1
SME03
Bacillus
amyloliqefaciens
99.00
99.93
CP054415.1
LSE04
Escherichia coli
100.00
100.00
MK371829.1
LSE06
Klebsiella aerogenes
100.00
100.00
CP048598.1
Key:
LSE01: Bacillus cereus
SME02: Pseudomonas aeruginosa
SME03: Bacillus amyloliqefaciens
LSE04: Escherichia coli
LSE06: 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)
Pseudomonas sp.
Bacillus amyloliqefaciens
Bacillus cereus
Klebsiella sp.
Escherichia sp.
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)
Pseudomonas sp.
Bacillus
amyloliqefaciens
Bacillus cereus
Klebsiella sp.
Escherichia sp.
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
-
-
-
-
-
Sample depths
Sample depths
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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
Aspergillus niger
Fusarium solani
Penicillium chrysogenum
Microsporum sp
Penicillium citrinum
Aspergillus flavus
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)
Aspergillus niger
Fusarium solani
Penicillium
chrysogenum
Microsporum sp
Penicillium citrinum
Aspergillus flavus
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)
Biosurfactant Production
Hemolytic activity
Pseudomonas
aeruginosa
12.00
+
+
Bacillus
amyloliqefaciens
15.00
+
+
Klebsiella aerogenes
7.00
+
-
Escherichia coli
0.00
-
+
Bacillus cereus
16.00
+
+
Aspergillus niger
14.00
+
+
Fusarium solani
10.00
+
+
Penicillium
chrysogenum
7.00
+
+
Microsporum sp.
0.00
-
+
Keys: + = positive, - = negative
Sample depths
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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
Pseudomonas aeruginosa
0.644±1.22
Bacillus amyloliqefaciens
1.01±2.30
Bacillus cereus
0.91±0.33
Klebsiella aerogenes
0.41±0.50
Escherichia coli
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 (1.01 ± 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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