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Highlights in BioScience
ISSN: 2682-4043
DOI:10.36462/H.BioSci.20218

Highlights in BioScience August 2020| Volume 3
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Research Article

Open Access

Department of Science Technology, Akwa Ibom
State Polytechnic, Ikot Osurua, P.M.B 1200,
Ikot Ekpene, Akwa Ibom State, Nigeria.

Contacts of Authors

* To whom correspondence should be
addressed: Aniefon Alphonsus Ibuot

Citation: James II, Ben MG, Jones AM, Akpan
PS, Eka II, Oruk AE, Ibuot AA (2020).
Characterization of hydrocarbon utilizing
bacteria in waste engine oil-impacted sites.
Highlights in BioScience Volume 3. Article ID
20218. dio:10.36462/H.BioSci.20218

Received: June 6, 2020

Accepted: August 18, 2020

Published: August 31, 2020

Copyright: © 2020 James et al. This is an open
access article distributed under the terms of the
Creative Commons Attribution License, which
permits unrestricted use, distribution, and
reproduction in any medium, provided the
original author and source are credited.

Data Availability Statement: All relevant data
are within the paper and supplementary
materials.

Funding: The authors have no support or
funding to report.

Competing interests: The authors declare that
they have no competing interests.

Characterization of hydrocarbon utilizing bacteria in
waste engine oil-impacted sites

Iniobong Ime James, Mayen Godwin Ben, Agnes Monday Jones,Patience
Saturday Akpan, Idorenyin Idorenyin Eka, Albert Ema Oruk, and Aniefon
Alphonsus Ibuot*

Abstract
Changes in soil physicochemical properties and bacterial species

present in soil contaminated with waste engine oil were evaluated at three
auto-mechanical workshops in Uyo, Nigeria. This work was aimed at
isolating and identifying hydrocarbon degrading bacteria from waste
engine oil polluted soil, and assessing their hydrocarbon-utilizing ability.
Waste engine oil pollution affected soils significantly with increases in soil
physicochemical properties, and heterotrophic bacterial population counts.
Eight bacterial species Corynebacterium kutscheri, Pseudomonas
aeruginosa, Flavobacterium aquatile, Serratia odorifera, Micrococcus
agilis, Staphylococcus aureus, Micrococcus luteus and Bacillus substilis
were isolated by the selective enrichment technique and screened for
hydrocarbon utilization capability in mineral salt media with 1% (v/v)
waste engine oil as a sole carbon and energy source. The extent of bacterial
growth observed was related to the ability of organisms to biodegrade
hydrocarbons present in the medium bacterium species, which showed
varying hydrocarbon utilization during the 15 days of incubation. Growth
in hydrocarbon medium was the most efficient in cultures of
Corynebacterium kutscheri. All isolates also showed variable
emulsification ability, with Corynebacterium kutscheri, showing the
highest ability. These results demonstrate the presence of indigenous
bacteria in hydrocarbon-polluted soils and the potential toward the
remediation of hydrocarbons.

Keywords: Hydrocarbon-utilizing bacteria, selective enrichment technique,
Corynebacterium kutscheri

Introduction
Petroleum utilization as fuel and petroleum products leads to severe

environmental pollution [1]. Large-scale accidental spills pose a great threat to
the ecosystem [2]. Soil pollution by petroleum hydrocarbons has been shown
to produce pronounced changes in the physicochemical and microstructure of
the oil-contaminated soil [3]. This affects parameters such as soil porosity,
bulk density, and adsorption [4-5]. Fresh spills and/or high levels of pollutants
may often result in the reduction of large sectors of soil microbial population,
although soils with lower levels or old pollution may show an increase in
numbers and diversity of microorganisms [6-7]. The diversity and the number
of microorganisms at polluted soil sites may assist in the characterization of
such a site, such as the toxicity of petroleum hydrocarbons to the microbiome,
age of the spill and concentration of the pollutant [8].

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Additionally, microorganisms in soils exposed to
hydrocarbon pollution usually exhibit a higher potential for
biodegradation of such pollutant compounds than others with
no history of such exposure. Percentages of hydrocarb-
onoclastic microbes are quite low in soil when there is no oil
spill, but may increase 1,000 fold after oil spill [9].

Conventional remediation methods do not seem to be
able to address this problem, or tends to aggravate the
problem [10]. Mechanical methods such as incineration,
excavation and/or burial in secure land fill, as well as a host
of other chemical decomprelocates osition methods are
expensive, time consuming and only the pollution [11]. An
efficient way of remediating the oil-contaminated sites could
be employment of microorganisms, such as bacteria,
microscopic algae, and fungi, isolated from polluted
environments or enhanced from the organisms already
present in the same environment [12-13]. Waste engine
oil-polluted soils also serve as a source of indigenous
bacteria capable of hydrocarbon degradation.

The employment of microorganisms in the
biodegradation of hydrocarbons over chemical or
conventional treatment is preferred for many reasons; end
products are comparatively safer and cost-effectiveness [11].
Ogunbayo et al., [18] evaluated the effectiveness of bacteria
indigenous to soil in remediating engine oil-polluted, soil and
isolated Bacillus, Pseudomonas, Flavobacterium, Microc-
occus and Rhodococcus species, with Pseudomonas and
Rhodococcus species giving most favorable degradation
effectiveness and efficiencies.

This study therefore considered the isolation of
indigenous bacterial communities in waste engine
oil-polluted soil using selective enrichment technique, and
the assessment of hydrocarbon-utilization capability in waste
engine oil-augmented mineral salt medium.

Materials and Methods
Sample collection

Waste engine oil –contaminated soil samples used in
this study were collected from three auto-mechanic
workshops within the mechanic village, Uyo, Akwa Ibom
State, Nigeria. Composite soil samples were obtained at each
sampling point using a soil auger from 0-10 cm below the
soil surface. The soils were labeled “Unpolluted” for the
unpolluted sample, “MA” for the mechanic workshop 1
sample, “MB” for the mechanic workshop 2 samples, and
“MC” for the mechanic workshop 3 samples. This was
followed by bulking and transportation to the laboratory in
sterile polythene bags within six hours for isolation of
organisms.

Physico-chemical analysis of soil samples
The soil pH was measured using HANNA

Instruments Model 209 pH meter [14]. Moisture content was
calculated on the basis of the air dry weight as described by
AOAC [15]. Total organic carbon was calculated by
weighing exactly 0.5 g of the soil sample into a flask, and 10
ml of 1.0 M K2Cr2O7 was added and swirled to mix. 20 ml

conc. H2SO4 was added, gently swirled for a minute and
allowed to stand for 20 minutes. The suspension was diluted
to about 100 ml of distilled water. Five drops of
o-phenanthroline indicator were added to each sample and
was titrated with 0.5 M ferrous ammonium sulfate to a light
blue end point. The reagent blank was also run and the titre
values recorded, and used to calculate the organic carbon
content [15].

The total hydrocarbon content (THC) was determined
by first extracting hydrocarbons by acidifying 2 g of
representative soil samples using H2SO4, and extracting upon
addition of 20 ml of toluene in a separatory funnel. The
contents of the funnel were shaken, and allowed to settle into
two layers. The absorbance of the supernatant (extract) was
read at 420 nm with UNICAM UV/VIS spectrophotometer
(Spectronic 20D). Readings were recorded from the
spectrophotometer and using the determined curve to obtain
the figure [16].

Phosphorus was determined using the ascorbic acid
method as described by AOAC, [15]. 50 ml of the soil
dilution was pipetted into 250 ml Erlenmeyer flask, and 1
drop of phenolphthalein indicator was added. Exactly 5 N
H2SO4 (148 ml conc. H2SO4 in 100ml H20) is added
drop-wise to develop a red colour. Exactly 8 ml of combined
reagents made up of 50 ml of 5 N H2SO4, 5 ml potassium
antimonyl tartrate solution (1.372 g potassium antimonyl
tartrate in 500 ml distilled water); 15 ml ammonium
molybdate solution (20 g ammonium molybdate crystal in
500 ml distilled water); were added and thoroughly mixed,
and allowed to stand for 20 min. The phosphorus content was
determined by measuring the absorbance of the sample at
880 nm.

The nitrogen concentration was determined according
to the methods of Bremmer and Mulvaney, [17]. One
milliliter of the soil sample was introduced into the standard
kjeldahl flask containing 1.5 g CuSO4, and 1.5 g Na2SO4 as
catalyst, alongside concentrated H2SO4. The flask was gently
heated on a heating mantle, taking care to prevent frothing.
The solution was transferred after heating to a 100 ml
standard flask and made up to the mark with distilled water.
A portion of this digest was pipetted into a semi
micro-kjeldahl distillation apparatus and treated with 30 ml
of 40% NaOH solution. The ammonia evolved was
steam-distilled into a 100 ml conical flask containing 10 ml
solution of saturated boric acid to which 4 drops of Tashirus
indicator had been previously added. The tip of the condenser
was immersed in the boric acid solution and the distillation
continued until about two-thirds of the original volume was
obtained. The tip of the condenser was finally rinsed with a
few milliliters of distilled water. The distillate was then
titrated with 0.1N HCl until a purple-pink end point was
observed. A blank determination was also carried out in a
similar manner without the sample, and the calculation done
as follows:

Nitrogen (%) = (Real titre – Blank titre) x 0.1 x 0.014 x 100
Weight of the sample

Enumeration of total heterotrophic bacteria (THB)
The THB population in the soil samples was

enumerated by adopting the standard plate counts technique
using the spread plate method as described by Ogunbayo et
al., [18]. These involved spreading aliquots of a serially

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diluted 0.1 ml of 10-5 dilutions of the soil sample suspension
on nutrient agar plates and the plates were incubated at 30 oC
for 24 h. Similar aliquots were also incubated in minimal salt
agar plates containing used engine oil as the sole source of
carbon and energy. The plates were all incubated aerobically
at 30 oC. The percentages of hydrocarbon-utilizing bacteria
(HUB) relative to the total heterotrophic counts were noted.

Enumeration of hydrocarbon utilizing bacteria (HUB)
Oil-utilizing bacteria were isolated from polluted soil

samples by enrichment in mineral salt medium (MSM)
modified from Okpokwasili and Nwosu, [19] using waste
engine oil as a carbon and energy source. The soil samples
were sieved using a 2 mm mesh sieve. 10 g of the sieved soil
samples was inoculated into 100 ml sterile MSM. 1 ml of the
waste engine oil was added to the medium as a sole source of
carbon and energy, and the culture was incubated on a rotary
shaker at 170 rpm for 1 week.

The enrichment procedure was repeated for three cycles.
At the end of each enrichment cycle, 1 ml of the culture was
diluted serially 10-fold down the gradient to 10-5 and plated.
Pure cultures of the isolates were obtained by plating 1 ml of
the 10-5 dilution of the third enrichment cycle onto MSM
agar plates, and incubating at 30 oC (± 2) for 48 h. Pure
cultures obtained by this procedure were stored in slants at 4
oC until further identification.

Characterization and identification of bacteria
Isolates were identified on the basis of colonial

characteristics, Gram’s reaction and cell biochemical
reactions as described by Cheesbrough [20]. Identification
used the taxonomic schemes of Holt et al. [21].

Hydrocarbon utilization screening of bacteria
To determine the ability of the isolates to utilize engine

oil as the sole carbon and energy source, the growth patterns
of isolates in mineral salt medium in the presence of 1% (v/v)
of the waste engine oil (5.0 mL in 100 mL MSM) were
determined according to Onuoha et al. [22]. Waste engine
oil-augmented MSM was dispensed into 250 ml Erlenmeyer,
and inoculated with 0.1 ml of 24 h cultures of the bacterial
isolates. Incubation was done at 30 oC for 15 days. Growth
patterns were determined monitoring changes in pH, optical
density and total viable count at 5-day intervals during the
incubation. The pH of the medium was measured using the
pH meter (HANNA Instruments). Growth was also
monitored by measuring the optical density (OD) at 600 nm
using the spectrophotometer (Spectrumlab). Total viable
counts of the cultures were obtained by incubation of 0.1 ml
of the cultures using the spread plate technique on nutrient
agar plates at 30 oC for 24 h.

Emulsification activity of bacteria
The emulsification index (E24) of the isolates was

determined according to the methods of Ganesh and Lin, [23],
by adding 1ml of waste engine oil to the same amount of
culture media as used for degradation assay, mixing the
vortex for 2 min and leaving to stand for 24 h. The percentage
of emulsification index was obtained as follows:

E24 = Height of the emulsified layer x 100
Total height of the liquid column

Results
Physicochemical properties of soil samples

The results of the physicochemical analysis of the
different soil sample are shown in Table 1. The high
amounts of organic carbon (5.32 ± 2.65% in MA, 9.79 ±
0.51% in MB and 7.29 ± 3.09% in MC), and THC (2933.76 ±
404.27 mg/kg in MA, 3122.72 ± 131.00 mg/kg in MB and
3202.61± 675.07 mg/kg in MC), compared to the unpolluted
soil sample (3.7 ± 2.43% organic carbon content and 39.97±
13.49 mg/kg THC) is indicative of heavy pollution of the
mechanical workshop samples with petroleum hydrocarbons.
Soil samples from mechanical workshop 1 contained higher
amounts of nitrates (0.25 ± 0.03 mg/g), while samples from
mechanical workshop 2 contained the highest amounts of
phosphates (10.74 ± 0.88 mg/g) and THC (3202.61± 675.07
mg/kg). The pH values of the soil samples indicate all soil
samples as moderately acidic to acidic (from pH 5.78 to pH
6.79).

Bacterial count of soil samples
The total heterotrophic bacterial count and hydrocarbon

utilizing bacterial count of the original soil samples is shown
in Table 2. A higher THB count was recorded in polluted soil
samples (4.4 ± 1.90 x 107CFU/g from MA sample, 6.0 ± 0.23
x 107 CFU/g from MB sample and 4.5 ± 0.03 x 107 CFU/g
from MC sample) than in the unpolluted soil sample (1.9 x
107 CFU/g). Higher THB (6.0 ± 0.23 x 107CFU/g) and HUB
(5.2 ± 0.25 x 107 CFU/g) counts were observed in the MB
sample than in other similar polluted samples indicative of its
extent of pollution. Hydrocarbon utilising bacterial counts
were slightly lower in all samples than the corresponding
heterotrophic bacterial counts.

Characterization and identification of bacteria
The identified bacterial isolates were Corynebacterium

kutscheri, Pseudomonas aeruginosa, Micrococcus agilis,
Flavobacterium aquatile, Staphylococcus aureus,
Micrococcus luteus, Serratia odorifera and Bacillus substilis,
as shown in Table 3 and 4.

Hydrocarbon utilization potential of bacteria
Table 5 shows the changes in pH of MSM during growth

of bacteria isolates in hydrocarbon. Decreases in pH (to <pH
7.00) were observed in medium containing isolates,
Corynebacterium kutscheri, Micrococcus agilis, Serratia
odorifera and Bacillus substilis, (which, however, increased
to above pH 7.00). The greatest decrease in pH occurred in
cultures of Corynebacterium kutscheri (from 7.00 a.m. on
Day 0 to 5.83 ± 0.34 on Day 15). Medium containing other
isolates showed slight increases in pH over time. The
changes in the total viable counts of bacterial isolates during
15 days of growth in waste engine oil-augmented MSM are
shown in Table 6. Total viable counts were higher in cultures
containing Corynebacterium kutscheri (1.94 ± 0.12 x 108

CFU/ml on Day 5, 6.73 ± 0.45 x 108 CFU/ml on Day 10 and
3.13 ± 0.02 x 108 on Day 15). Growth of Staphylococcus
aureus showed the lowest decreases (0.31± 0.11 x 108on Day

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5, 0.44 ± 0.01 x 108 on Day 10 and 0.42 ± 0.02 x 108 on Day
15).

Table 7 shows the results of turbidity measurement
(OD600) of the medium during incubation. The highest
increases in turbidity were recorded in cultures with
Corynebacterium kutscheri, (0.189 ± 0.04 on Day 0 to
0.301± 0.11 on Day 15), and Bacillus substilis, (0.165 ± 0.04
on Day 0 to 0.341± 0.02 on Day 15). The lowest turbidity

was observed with Staphylococcus aureus (0.20 ± 0.04 on
Day 0 to 0.14 ± 0.02 on Day 15).

Emulsification activity of bacteria

Figure 1 shows the emulsification index (E24) of the
bacterial isolates in waste engine oil. Corynebacterium
kutscheri had the highest emulsification index (52%). The
lowest emulsification index was observed for
Staphylococcus aureus (8%).

Table 1: Physicochemical properties of different soil samples contaminated with waste engine oil.

Values represent means of triplicate determinations ± SD. MA = Mechanic Workshop 1, MB = Mechanic Workshop 2, MC = Mechanic Workshop 3. THC =
Total hydrocarbon content.

Table 2: Bacterial count of soil samples contaminated with waste engine oil.

Values represent means of triplicate determinations ± SD. MA = Mechanic Workshop 1, MB= Mechanic Workshop 2, MC= Mechanic Workshop 3.

Table 3 : Biochemical characteristics of spore (Sp), catalase (Ca), motility (Mo), oxidase (Ox), litmus reaction (Lr), urease (Ur),
gelatin (Ge), citrate (Ci), glucose (Gl), maltose (Mal), mannitol (Man), lactose (La), sucrose (Su), arabinose (Ar) and xylose (Xy)
for studied bacterial isolates from soil contaminated with waste engine oil.

Keys: AG = acid and gas production; A = acid production only; + = positive reaction; - = negative.

Table 4: Phenotypic characteristics of bacterial isolates from soil contaminated with waste engine oil.

Parameters Unpolluted (Control) Polluted
MA MB MC

pH 6.79 ± 0.24 6.18 ± 0.13 5.78 ± 0.03 6.44 ± 0.07
Nitrate (mg/g) 0.14 ± 0.02 0.25 ± 0.03 0.23 ± 0.02 0.20 ± 0.02
Phosphate (mg/g) 6.65 ± 1.72 6.92 ± 2.92 10.74 ± 0.88 8.49 ±1.9
Moisture content (%) 32.25 ± 9.30 43.09 ± 0.65 45.13 ± 7.02 27.72 ± 9.09
Organic carbon content (%) 3.7 ± 2.43 5.32 ± 2.65 9.79 ± 0.51 7.29 ± 3.09
THC (mg/kg) 39.97 ±13.49 2933.76 ± 404..27 3202.61± 675.07 3122.72±131.00

Parameter Unpolluted (Control) Polluted
MA MB MC

THB (CFU/g) 1.9 ± 0.02 x 107 4.4 ±1.90 x 107 6.0 ± 0.23 x 107 4.5 ± 0.03 x 107
HUB (CFU/g) 6.5 ± 0.04x 106 3.2 ± 0.05 x 107 5.2 ± 0.25 x 10 7 1.9 ± 0.10 x 106

Isolates Isolate code Sp Ca Mo Ox Lr Ur Ge Ci Gl Mal Man La Su Ar Xy

1 MA3, MB1, MC4 - + - - - - - + A A A - - - AG

2 MA2, MB2 - + + + + - + - - A A A - A A

3 MA1 - + - - - - - - A - A - - AG AG

4 MA4 - + - + + - - - A A - A A - -

5 MB3, MC1, MC2, MA5 - + - - + - - - - - - - A - -

6 MA6 - + - + - - - - - - A - A - -

7 MC3 - + + - - + + + A A A A A A -

8 MB4, MC5 + + + - + - + + A A A - A A AG

Isolates Isolate code Gram reaction/Cell shape Probable Bacteria

1 MA3, MB1, MC4 Gram-positive Rods Corynebacterium kurtscheri

2 MA2, MB2 Gram-negative Rods Pseudomonas aeruginosa

3 MA1 Gram-positive Cocci Micrococcus agilis

4 MA4 Gram-negative Rods Flavobacterium aquatile

5 MB3, MC1, MC2, MA5 Gram-positive Cocci Staphylococcus aureus

6 MA6 Gram-positive Cocci Micrococcus luteus

7 MC3 Gram-negative Rods Serratia odorifera

8 MB4, MC5 Gram-positive Rods Bacillus substilis

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Table 5 : pH of the medium during the growth of bacterial isolates in hydrocarbon medium.

Isolates Days
0 5 10 15

Micrococcus agilis 7.00 6.51 ± 0.01 6.38 ± 0.03 6.04 ± 0.05
Pseudomonas aeruginosa 7.00 7.06 ± 0.07 7.78 ± 0.01 7.71 ± 0.06
Corynebacterium kutscheri 7.00 6.51± 0.01 6.38 ± 0.00 5.83 ± 0.34
Flavobacterium aquatile 7.00 7.31± 0.23 7.50 ± 0.16 7.17 ± 0.28
Micrococcus luteus 7.00 6.13± 0.07 7.23 ± 0.03 7.37 ± 0.06
Staphylococcus aureus 7.00 7.23± 0.03 7.31± 0.03 7.23 ± 0.07
Serratia odorifera 7.00 7.09 ± 0.01 6.73 ± 0.04 6.42 ± 0.07
Bacillus substilis 7.00 6.85 ± 0.13 6.78 ± 0.18 7.24 ± 0.02
Data represent means of triplicate determinations ± SD.

Table 6: Total viable count (TVC) of bacterial isolates during growth in hydrocarbon medium (x 108 CFU/ml).

Isolates
Days

0 5 10 15
Micrococcus agilis 0 0.63 ± 0.03 2.46 ± 0.02 0.87 ± 0.06
Pseudomonas aeruginosa 0 0.54 ± 0.02 2.13 ± 0.16 0.45 ± 0.02
Corynebacterium kutscheri 0 1.94 ± 0.08 5.27 ± 0.07 3.18 ± 0.07
Micrococcus luteus 0 0.95 ± 0.01 2.49 ± 0.03 1.43 ± 0.03
Flavobacterium aquatile 0 1.94 ± 0.12 6.73 ± 0.45 3.13 ± 0.02
Staphylococcus aureus 0 0.31± 0.11 0.44 ± 0.01 0.42 ± 0.02
Serratia odorifera 0 0.54 ± 0.03 2.23 ± 0.25 0.48 ± 0.04
Bacillus substilis 0 1.39 ± 0.41 4.08 ± 0.65 2.47 ± 0.07
Data represent means of triplicate determinations ± SD.

Table 7: Turbidity measurement of bacterial growth in hydrocarbon medium (OD600).

Isolates
Hours

0 5 10 15
Microccoccus agilis 0.97 ± 0.01 0.154 ± 0.01 0.209 ± 0.12 0.201± 0.03
Pseudomonas aeruginosa 0.033 ± 0.001 0.035 ± 0.10 0.182 ± 0.10 0.097± 0.05
Corynebacterium kutscheri 0.189 ± 0.04 0.252 ± 0.07 0.386 ± 0.04 0.301± 0.11
Flavobacterium aquatile 0.085 ± 0.01 0.153 ± 0.08 0.196 ± 0.07 0.227± 0.05
Micrococcus luteus 0.040 ± 0.02 0.083 ± 0.06 0.162 ± 0.03 0.183± 0.04
Staphylococcus aureus 0.20 ± 0.04 0.053 ± 0.10 0.084 ± 0.05 0.014± 0.02
Serratia odorifera 0.40 ± 0.02 0.172 ± 0.11 0.121 ± 0.001 0.096± 0.01
Bacillus substilis 0.165 ± 0.05 0.204 ± 0.01 0.413 ± 0.05 0.341± 0.04
Data represent means of triplicate determinations ± SD.

Figure 1 : Emulsification activity (E24) of bacterial isolates.

Discussion
This study on the assessment of hydrocarbon utilizing

the potential of bacteria isolated from waste engine
oil-polluted soil on hydrocarbons reveals the presence of
hydrocarbon-utilizing bacteria in waste engine oil-polluted
soil environment as well as the potential of one of these
isolates Corynebacterium kutscheri to degrade hydrocarbons
under a variety of experimental conditions.

The results of the physicochemical analysis of the soil
samples, as presented in Table 1 showed higher levels of
properties in the polluted soil samples when compared with
the unpolluted soil sample. The results support the results of
Chikere [24], and Chikere and Ekwuabu [25], which showed
high physicochemical parameters in polluted soil samples
compared to unpolluted samples determined and indicated
previous exposure of polluted samples to hydrocarbon
contamination with traces of other organic and inorganic
contaminants.

The high bacterial counts recorded in polluted soil
samples, as presented in Table 2, compared with the
unpolluted control samples could be attributed to the myriad
of nutrients, high organic matter concentration and other
ecological factors that influence the survival of soil
microorganisms that play important roles in the
decomposition and recycling of nutrients [26]. Luepromchai
et al. [27] has reported an increase in the numbers of
hydrocarbon degraders in soil in the presence of PAHs,
without any impact on the overall bacterial numbers. The
difference between THB and HUB counts was observed to be
minimally insignificant, which suggests that most of the
micro-organisms present in various polluted sample sites are

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hydrocarbon degraders that can withstand the concentrations
of hydrocarbons and use them as a source of carbon [25].

The investigation of the morphology and type of
bacterial colonies obtained from the oil-polluted soil showed
seven bacterial species, which are Corynebacterium
kutscheri, Pseudomonas aeruginosa, Micrococcus agilis,
Flavobacterium aquatile, Staphylococcus aureus,
Micrococcus luteus, Serratia odorifera, and Bacillus
substilis. This shows the majority of bacteria isolated as
being Gram-positive species. Some researchers have,
however, reported oil-polluted soils as being dominated by
Gram-negative bacteria [25]

The results of the emulsification test as shown in Figure
1 demonstrates that the isolates produced emulsifying
compounds. A large variety and number of biosurfactant
producers have been isolated from hydrocarbon-impacted
sites [28], although they have also been identified from soils,
which are unconnected to hydrocarbon contamination [29].
Corynebacterium kutscheri showed the highest
emulsification (52%) at 1% waste engine oil, while the least
was Staphylococcus aureus (8%). Corynebacterium sp has
also been reported by Onuoha et al. [22] with high
emulsification ability among three hydrocarbon-degrading
bacterial species isolated from soil.

Decrease in the pH of the medium (as shown on Table
5), increase in total viable counts (Table 6), as well as in
turbidity of the medium (Table 7) were regarded as
indicators of degradation. The correlation between the pH of
the medium and cell growth with hydrocarbon utilization has
been previously reported by Patila et al., [30]. The
preliminary estimation of the isolate’s degradative ability
during growth in hydrocarbon supplemented medium
suggested the effectiveness of growth in hydrocarbon
medium by Corynebacterium kutscheri, and thus its potential
as a candidate for further biodegradation studies.

Conclusion
The result of this study indicates that the contaminated

environments possess indigenous populations within them,
as seen in bacterial count results of microbiological growth
of mineral salt cultures with waste engine oil as the sole
carbon source. These indigenous bacteria isolated in this
study show promising, ability to degrade petroleum
hydrocarbons with Corynebacterium kutscheri as a potential
candidate for further biodegradation efficiency studies.

Acknowledgments
The authors are grateful to the Laboratory Technologist

of Microbiology Laboratory, Michael Okpara University,
Umudike, Nigeria for their support.

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