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Asian Jr. of Microbiol. Biotech. Env. Sc. Vol. 17, No. (4) : 2015 : 831-838
© Global Science Publications
ISSN-0972-3005
*Corresponding author’s email: ruguruwanjohi@gmail.com
ISOLATION AND IDENTIFICATION OF BACTERIA WITH
BIOREMEDIATION POTENTIAL OF OIL SPILLS IN LAKE
NAKURU, KENYA
LUCY WANJOHI*1a, LIZZY MWAMBURI1b, EMILY TOO1b, BECKY ALOO1b AND JANET KOSGEI1b
1aDepartment of Environmental Biology and Health, School of Environmental Studies University of
Eldoret, P.O. Box 1125-30100, Eldoret
1bDepartment of Biological Sciences, School of Science, University of Eldoret, P.O Box 1125-30100, Eldoret
(Received 22 March, 2015; accepted 17 May, 2015)
Key words : Bacteria, Bioremediation, Oil spills, Environment
Abstract–Although the role of bacteria in all spheres of humanity has been documented, new challenges are
emerging with researchers querying the importance that bacteria could play. Moreover, some ecosystems
especially the alkaline lakes are known to be rich in microbes that are a potential solution to the emerging
challenges. One such new challenge is the management of non-bio-degradable fossil fuel products within
the agri-ecosystems and natural environment. Microbes with enzymes capable of degrading such products
have been isolated in some of the saline lakes in Kenya. Studies on bacterial ecology of Lake Nakuru have
not been well covered. The high demand of energy from fossil fuels results in accidental and associated
petroleum oil spills which results in ecological risks and long term environmental disturbance. The waters
of Lake Nakuru may be rich in halophilic bacteria capable of degrading such pollutants. The aim of the
study was to determine bacteria biodiversity in Lake Nakuru and their potential utilization in oil spill
bioremediation on agricultural land and marine environment. Water samples were collected in five selected
points that reflected different catchment areas. Sampling was done from a boat on a monthly basis for six
months consecutively from December 2010 to May 2011. Three water samples of 500 ml were collected at
each site at ten centimeters depth. Serial dilution, culturing and isolation of bacteria were done.
Identification was through physical characterization and biochemical tests. Twenty-one bacteria isolates
were identified. Oil degradation was determined by greasy spot test. Bacteria identified as having high
potential in oil bioremediation management includes Providencia rettgeri, Morganella morganii, Tatumella
ptyseas, Bacillus anthracoides, Chryseobacterium indologenes, Streptococcus pyogenes, Chryseobacterium
meningosepticum, Pseudomonas cepacia, Proteus penneri, Alcaligene sp., Moraxella sp., Providencia stuarti and
Sphingomonas paucimobilis. Novel species in this study were Tatumella ptyseas, Streptococcus pyogenes, Proteus
penneri, Providencia stuarti and Yersinia pseudotuberculosis.
INTRODUCTION
Aromatic hydrocarbons are environmental
pollutants with toxic, carcinogenic and mutagenic
properties (Mastrangela et al., 1997). This
environmental pollution occurs when trying to meet
the high demand of petroleum oil as a source of
energy. During the process of exploration,
production, transportation and storage, accidental
and associated petroleum oil spills occur. Petroleum
oil spills from transportation accidents, pipeline
breakages and tank leakages can be considered as
the most frequent causes of aromatic hydrocarbon
release (Bossert et al., 1984). Marine environments
are usually vulnerable to oil contamination since oil
spills of coastal regions and open sea are poorly
containable and mitigation is difficult. Oil spills
result in ecological risks and long-term
environmental disturbance. The impact associated
with oil seepages on land include loss of soil
fertility, water holding capacity, permeability and
binding capacity (Bossert et al., 1984).
The search for effective and efficient methods of
oil removal from contaminated sites has intensified
(Hoff, 1993). One promising method is the biological
degradation of oil by bacteria that metabolize oil
that is rich in carbon to satisfy their cell growth and
energy needs (Coral et al., 2005). Oil biodegradation
832 WANJOHI ET AL.
is a large component of oil weathering, a natural
process whereby bacteria alter and break down
organic molecules into other substances, producing
fatty acids and carbon dioxide (Atlas, 1991).
Many microorganisms possess the enzymatic
capability to degrade petroleum hydrocarbons.
Some microorganisms degrade alkanes, others
aromatics and others both paraffinic and aromatic
hydrocarbons. Often the normal alkanes in the range
of C10 to
C26 are viewed as the most readily
degraded, but low- molecular- weight aromatics,
such as benzene, toluene and xylene, which are
among the toxic compounds found in petroleum
are also very readily biodegraded by marine
microorganisms. More complex structures are more
resistant to biodegradation. This means that fewer
microorganisms can degrade them and the rate of
biodegradation are lower than biodegradation rates
of the simpler hydrocarbon structure, the higher the
number of methyl branched constituents or
condensed aromatic rings, the slower the rate of
biodegradation (Atlas, 1991).
There is a lag time after oil spill before
indigenous microorganisms begin to break down
the oil molecules (Hoff, 1993). This lag time is
related to the initial toxicity of the volatile fractions
of the oil, which evaporates in a few days after a
spill. Microbes must begin to use the oil and expand
their population before measurable degradation
takes place, a period usually lasting several days
(Pothuluri et al., 1994).
Microbes play a major role in removal of poly
aromatic hydrocarbons (PAHs) from contaminated
environments due to their advantages such as cost
effectiveness and more complete clean up (Pothuluri
et al., 1994). Biodegradation of petroleum in the
marine environment is carried out largely by diverse
bacterial population, including various Pseudomonas
species (Vila et al., 2010). Marine oil spills are
eliminated by hydrocarbon-degrading activities of
microbial communities, particularly by specialist
bacteria called hydrocarbonoclastic bacteria (HCB)
(Santos et al., 2006). The hydrocarbon-degrading
microorganisms are ubiquitously distributed in the
marine environment. Generally in the pristine
environments, the hydrocarbons degrading bacteria
comprise of less than 1% of the total bacterial
population (Atlas, 1991). These bacteria presumably
utilize hydrocarbons that are naturally produced by
plants, algae and other living organisms. They also
utilize other substrates such as carbohydrates and
proteins. The proportion of hydrocarbon degrading
(HCD) microorganisms increases rapidly when an
environment is contaminated with petroleum. In
particular, in marine environments contaminated
with hydrocarbons, there is an increase in the
proportion of bacterial populations with plasmids
containing genes for hydrocarbon utilization.
Biodegradation of oil in soil is carried out by
bacteria that are found in oil contaminated sites,
such as vehicle service stations. HCD bacteria from
these sites have been reported to utilize aromatic
hydrocarbons at wide range of temperature and pH
(Swamakaran and Panchanathan, 2011). However,
oil is composed only of hydrogen and carbon, and
the bacteria need additional nutrients to grow. The
inorganic nutrients and oxygen should be provided
in order to provide the bacteria with nitrogen and
several essential minerals (Hoff, 1993). The complete
biodegradation (mineralization) of hydrocarbons
produces the non-toxic end products, carbon
dioxide and water, as well as cell biomass which are
largely proteins which can be safely assimilated into
the food chain (Hoff, 1993).
Oil spills occur in Nakuru town garages and are
washed by the run off during the rainy season to
Lake Nakuru. Oil spills results in ecological risks
and long term environmental disturbance. Saline
lakes in Kenya have microbes which are able to
degrade organic and inorganic materials. Such
microbes have been isolated in some of the saline
lakes in Kenya especially lake Bogoria and lake
Elementaita. Studies on bacterial ecology of lake
Nakuru have not been well covered although a lot of
studies on zooplankton ecology in lake Nakuru
have been undertaken by Yasindi et al., (2002) and
phytoplankton ecology by Oduor and Schagerl
(2007). The waters of Lake Nakuru may be rich in
halophilic bacteria which may be important in
bioremediation. There is need to therefore identify
novel bacteria from Lake Nakuru which are able to
remediate our environment from plastic and
petroleum oil pollutants.
MATERIALS AND METHODS
Study area
Lake Nakuru is in the Nakuru County, Kenya, at an
altitude of 1,759 m. It is within Lake Nakuru
national park. It consists of a shallow pan of water
lying on salt impregnated clay which retains coarser
polar sediments. The surface area is 40-60 km2 but is
subject to marked fluctuations as lake level is
constantly rising and falling. The average depth is 1
m (Kairu, 1991). The length of shoreline is 27 km.
Isolation and Identification of Bacteria with Bioremediation Potential of Oil Spills in Lake 833
The water level is unregulated. It has a catchment
area of 1,800 km2 (Vareshi, 1982).
Sampling Locations
Five sites were selected and geo-referenced using
Geographical Positioning System (GPS). The sites
selected were; the middle of the lake (Jetty mid)
(latitude -0.354781 and longitude 36.093118), Hippo
point (latitude -0.319546 and longitude 36.105102)
and near the mouth of three rivers feeding the lake.
The site near river Enderit lies at latitude -0.386313
and longitude 36.110497, river Makalia (latitude -
0.391499 and longitude 36.083254) and river Njoro
(latitude -0.331833 and longitude 36.092667). These
sampling points were selected to reflect different
catchment areas.
Methods
Sampling was done from a boat in the five sites on a
monthly basis for six months consecutively from
December 2010 to May 2011. The samples were
collected randomly. At each sampling, three water
samples of 500 mL each were collected using sterile
plastic bottles at each site by hand dipping the
bottles at ten centimeters (10 cm) beneath the water
surface. The samples were kept in a cool box under
ice at 4oC during transportation. These samples were
used for serial dilution and culturing of the bacteria.
Culturing and counting the colony forming units
(CFU)
Nutrient agar media was sterilized and serial
dilution carried out up to 10 6. About 1 mL of the
diluted sample water was inoculated on sterile
Nutrient agar media. The inoculated plates were
incubated upside down at 35°C for 24 hours. The
petri dishes were sealed using adhesive tape to
prevent contamination. The number of colonies
formed (CFU) was counted per site to determine
bacterial load. Single colonies were picked using a
sterile wire loop and streaked on sterile media to
obtain pure cultures.
Identification of bacteria
The isolated bacteria were identified based on
physical characterization and biochemical tests
using Bergey’s manual of determinative bacteriology
(Holt, 1994). Morphological characteristics such as
shape and size were determined under a light
microscope and using Gram staining.
Motility test was determined by microscopically
observing the bacteria in a wet mount. An inoculum
from a freshly prepared culture was used to prepare
the wet mount. The inoculum was transferred to a
drop of water on a microscope slide, mixed and
covered using a cover slip. The slide was observed
under light microscope. The bacteria that were
observed to swim randomly against the current of
water streaming across the slide surface were
positive for this test.
Lactose test was conducted in order to determine
whether the bacteria fermented carbohydrate as
carbon source. An inoculum from a pure culture
was transferred aseptically to a sterile tube of phenol
red lactose broth. The inoculated tube was incubated
at 35°C for 24 hours. A positive test was indicated by
colour change from red to yellow.
Hydrogen sulfide gas production test was carried
out to determine whether the bacteria were able to
reduce sulfur containing compounds to sulfides
during metabolism process. An inoculum from a
pure culture was transferred aseptically to a sterile
triple sugar ion agar slant. The inoculated tube was
incubated at 35°C for 24 hours. A black colour in the
agar slant media indicated a positive test.
Citrate test was carried out to determine the
ability of the bacteria to utilize sodium citrate as the
only source of carbon. A sterile wire loop was used
to inoculate 3 mL of sterile Koser citrate medium
with a broth culture of bacteria. The inoculated
broth was incubated at 35° C for 72 hours. A change
of colour from green to blue indicated a positive
result.
Serotyping was carried out using Analytical
Profile Index (API) kit. API kits used were enteric
(API 20E) non-enteric (API 20NE) and Streptococcus
kits (API 20strep.) (Biomerieux Inc. USA).
Degradation of oil
Oil degradation experiment was done as described
by Spring (1994). The experiment was set up in four
replicates. Each set up of the main experiment
contained 2 mL of paraffin, 150 mL of distilled
water, the bacterium being tested and a mixture of
inorganic nutrients comprising of 0.011 M
ammonium phosphate, 0.002 M magnesium sulfate,
0.012 M potassium phosphate and 0.144 M non-
iodinated sodium chloride. These were put in a
conical flask that was sealed using parafilm to
provide aeration, avoid contamination and
evaporation.
The negative control comprised of 2 mL of
paraffin, 150 mL distilled water and the inorganic
nutrients. The positive control comprised of 2 mL of
834 WANJOHI ET AL.
paraffin, 150 mL of distilled water, inorganic
nutrients and 5 grams of soil sample from Huruma
service station in Eldoret, Kenya. This soil was
collected at a depth of 5 cm, placed in sterile
polythene bags and taken to the laboratory for the
purpose of the culturing, isolation and identification
of the bacteria present. The soil was autoclaved after
bacteria isolation and later inoculated with these
bacteria. These were put in conical flasks that were
covered using parafilm.
Results obtained from each conical flask were
recorded on a weekly basis for seven weeks. A
“greasy spot” test was performed by dividing a
brown paper bag into 5.08×5.08 cm squares and
labeling as per the conical flasks. A small quantity of
liquid just below the surface of oil from each jar was
drawn using a dropper. Two drops of this liquid
were put onto the center of the square on the brown
paper. This procedure was repeated twice for each
jar. A few minutes after the water evaporated a
greasy spot in each small square was observed. The
circumference of each greasy spot was marked using
a pencil and its diameter recorded. The initial
diameter measurement of the greasy spot formed by
the drop of oil was carried out immediately after
setting up the degradation experiment. The initial
mean diameter of all the set up was 5 cm. Subse-
quent diameter measurements were carried out on
weekly basis for seven weeks.
Data Analysis
The diameters obtained from the oil degradation
experiments were averaged for each bacterium
species for each week. A One-way Analysis of
Variance (ANOVA), was carried out to determine
whether the degradation of oil by each bacteria
species was significant. Time was used as the
independent variable while bacteria degradation
was the dependent variable. A Post hoc test was
carried out to separate the means of the greasy spot
formed by each bacterium. Means were separated
using Duncan’s multiple range test (DMRT) at 95%
level of significance
RESULTS
Biodiversity and Identification of Bacteria
The waters of Lake Nakuru were found to have
diverse bacteria (Figs. 1 to 6). Twenty one species
were isolated and identified from the waters of Lake
Nakuru (Table 1). The bacteria were classified
according to their different morphological
characteristics. There were 19 Gram negative and 2
Gram positive bacteria consisting of 17 bacilli, 1
coccus, 1 coccobacillus, 1 vibrio and 1 filamentous.
They were further identified using other
biochemical tests and API kits (Table 1). There were
12 enteric and 9 non enteric bacteria (Table 1).
Fifteen bacteria were motile while 6 were non motile
(Table 1). The bacteria isolated and identified from
Eldoret service station were Bacillus sp., Pseudomonas
sp., and Micrococcus sp.
Fig. 1. Bacteria streaks showing a variation of bacteria
growing on a petri dish
Fig. 2. Filamentous, Gram negative bacteria isolated from
Lake Nakuru
Fig. 3. Gram Negative rods isolated from Lake Nakuru
Isolation and Identification of Bacteria with Bioremediation Potential of Oil Spills in Lake 835
Biodegradation of oil
All bacteria species showed a progressive reduction
of the amount of oil in each set up (Table 2). After 1
week, Streptococcus pyogenes and Chryseobacterium
indologenes caused a higher reduction of the greasy
spot diameter than the rest with a diameter of 3.800
cm and 3.225 cm respectively (Table 6). The negative
control also showed a slight change in the size of the
greasy spot formed with a diameter of 4.950cm
(Table 2). Chryseobacterium indologenes caused the
highest change in the size of the greasy spot formed.
Providencia rettgeri, C. indologenes, and S. pyogenes
caused a significant reduction in the size of greasy
Fig. 4. Gram positive rods isolated from Lake Nakuru.
Fig. 5. Gram positive coccus isolated from Lake Nakuru.
Table 1. Bacteria species isolated from Lake Nakuru, classified according to shape, gram staining and biochemical tests
Bacteria species Gram Citrate H2S Fermentation Motility Enteric/non Shape
stain test production test enteric
Bacillus anthracoides + - - + + Non enteric Rods
Streptococcus pyogenes + - - - - Non enteric spherical
Erwinia mallotivora - - - - + Enteric Rods
Erwinia amylovora - - + + + Enteric Rods
Sphingomonas paucimobilis - - - - + Non enteric Rods
Morganella morganii - - + + - Enteric Rods
Enterobacter or Pantonea agglomerans - - - + - Enteric Rods
Yersinia pseudotuberculosis - - - + - Enteric Rods
Chryseobacterium meningosepticum - - - - - Non enteric Rods
Providencia stuarti - + - - + Enteric Rods
Vibrio vulnificus - - - + + Non enteric Comma
Pseudomonas cepacia - - - - + Enteric Rods
Proteus penneri - - - - + Enteric Rods
Erwinia nigrifluence - - - + - Enteric Rods
Agrobacterium radiobacter - - - + + Non enteric Rods
Providencia rettgeri - + - + + Non enteric Rods
Alcaligen sp. - - - + + Enteric Rods
Tatumella ptyseas - - - + + Enteric Rods
Moraxella sp. - - - + + Non enteric Spherical
and rods
Chryseobacterium indologenes - - - - - Non enteric Filamentous
Acinetobacter sp. - - - - - Enteric Rods
Fig. 6. Gram negative coccobacillus isolated from Lake
Nakuru.
836 WANJOHI ET AL.
spot diameter after two weeks. Providencia rettgeri
had the smallest diameter overall of 2.250cm (Table
2). The bacteria from the Eldoret service station were
good in oil degradation as they had an average
diameter of 2.775 cm. These bacteria were Bacillus
sp., Pseudomonas sp., and Micrococcus sp.
After 3 weeks the greasy spot diameter
measurement P. rettgeri was still the most active in
oil degradation followed by C. indologenes (Table 2).
The greasy spot test taken after 4 weeks indicated
that P. rettgeri, C. indologenes and S. pyogenes had
degraded a large amount of oil. Providencia rettgeri
indicated the greatest change in the greasy spot size
formed, with a diameter of 0.875 (Table 2).
The fifth diameter measurement indicated that P.
rettgeri had the smallest diameter measurement over
all. Bacteria from vehicle service station and C.
indologenes showed less than 1cm of the greasy spot
formed. The diameter measurements taken after 6
weeks indicated that P. rettgeri, C. indologenes and the
bacteria from vehicle service station had degraded
all the oil as there was no greasy spot formed.
Streptococcus pyogenes had a diameter of 1.425 cm
while Morganella Morganii had a greasy spot
diameter of 0.675 (Table 2).
At week 7 the greasy spot test indicated that
Streptococcus pyogenes and Morganella Morganii had a
greasy spot diameter of less than 1cm. Bacillus
anthracoides had a greasy spot diameter of 1.5 cm. All
the bacteria had a greasy spot diameter of less than
3.5 cm (Table 2).
Analysis of variance indicated that Bacillus
anthracoides, Yersinia pseudotuberculosis, Providencia
rettgeri, Bacillus sp., Pseudomonas sp., Micrococcus sp,
Providencia stuarti, Morganella morganii,
Sphingomonas paucimobilis, Moraxella sp., Alcaligen
sp., Proteus penneri, Chryseobacterium
meningosepticum, Pseudomonas cepacia, Tatumella
ptyseas, Chryseobacterium indologenes and
Streptococcus pyogenes were significant in oil
degradation. Acinetobacter sp., E. amylovora and E.
nigrifluence were unable to degrade oil.
DISCUSSION
Biodiversity of Bacteria in Lake Nakuru
The results from this study indicate that Lake
Nakuru has a rich diversity of bacteria. This may be
due to diverse ecological niche of the lake. Twenty
one species of bacteria were identified. They had
different morphological and biochemical
characteristics. There were bacillus, coccus,
coccobacillus, vibrio and filamentous bacteria. Some
of the bacteria were inhabitants of the lake as they
are known to be found in various habitats including
salty waters for example Vibrio vulnicus. Other
bacteria were likely to be in the Lake as a result of
pollution for example Streptococcus pyogenes.
Vareshi, (1982) had reported that the ecosystem in
the lake may have changed due to pollution. Most of
the bacteria identified are human, animal or plant
pathogens but despite this, they are important in
degradation and bioremediation.
Oil Degradation
There was a lag time before the process of
degradation of oil commenced. The lag time may
have allowed for the evaporation of the initial
toxicity of the volatile fraction of oils and also
increase in bacteria population. This was in
agreement with the findings of Pothuluri, (1994)
who reported that microbial populations must begin
to use the oil and expand their population before
measurable degradation takes place, a period
usually lasting several days. Application of nitrogen
can stimulate the bioremediation of oil
contaminated sites (Drozdowicz et al., 2002). Atlas,
(1981) reported that in aerobic oil degradation
process, the hydrocarbon in oil is converted to
carbon dioxide and water by bacteria. The initial
step in aerobic biodegradation involves the
oxidation of the substrate by oxygenases for which
molecular oxygen is required. Hoff, (1993) reported
that nitrogen and other essential minerals need to be
provided in order to provide the bacteria with the
necessary minerals.
Alcaligenes sp. was able to degrade the oil
significantly. This bacterium can be used in
bioremediation of oil contaminated sites.
Drozdowicz et al. (2002) reported that two bacterial
strains which belong to the genera Agrobacterium
and Alcaligens exhibited the ability to degrade
gasoline aromatic compounds and fix nitrogen at
the same time.
Other bacteria that were able to degrade oil were;
Morganella morganii, Tatumella ptyseas, Bacillus
anthracoides, Chryseobacterium indologenes,
Streptococcus pyogenes, Chryseobacterium
meningosepticum, Pseudomonas cepacia, Proteus
penneri, Alcaligen sp., Yersinia pseudotuberculosis,
Moraxella sp., Providencia stuarti, and Sphingomonas
paucimobilis. These bacteria can be used to remediate
the environment from oil contaminants. Results for
the positive control showed that the bacteria in this
Isolation and Identification of Bacteria with Bioremediation Potential of Oil Spills in Lake 837
Table 2. Weekly diameter measurements (cm) of the greasy spot formed by the oil drop for each bacterium species. (Two decimal places is suffice)
Bacteria species Week 1 Week 2 Week 3 Week 4 Week 5 Week 6 Week 7
1st diameter 2nd diameter 3rd diameter 4th diameter 5th diameter 6th diameter 7th diameter
measurement measurement measurement measurement measurement measurement measurement
B. anthracoides 4.325±0.217b4.200±0.178b4.025±0.278b3.125±0.301b2.800±0.329c2.525±0.335c1.500±0.381c
Y. pseudotuberculosis 4.925±0.377a4.375±0.450b4.225±0.217c3.725±0.170b3.000±0.342b2.975±0.371b2.350±0.733c
P. rettgeri 4.275±0.214c2.250±0.104b1.950±0.220d0.875±0.515d0.325±0.320d0.000±0.000d0.000±0.000d
Positive control 4.300±0.173b2.775±0.330b2.625±0.450d2.265±0.450c0.525±0.197d0.000±0.000d0.000±0.000d
P. stuati 4.100±0.070c4.075±0.750b3.975±0.225b3.750±0.166b3.600±0.910b3.525±0.193b3.300±0.212b
M. morganni 4.500±0.180c4.150±0.086 b 3.325±0.250c2.300±0.122c1.850±0.275c0.675±0.312d0.325±0.236d
S. paucimobilis, 4.375±0.085b4.175±0.138b3.925±0.286b3.880±0.568b3.775±0.144b3.700±0.070b3.400±0.168b
Acinetobacter sp. 4.475±0.206b4.250±0.176b4.000±0.187b3.975±0.085b3.850±0.119b3.800±0.334b3.425±0.189b
Moraxella sp. 4.400±0.168b4.200±0.040b3.950±0.064b3.550±0.150b3.475±0.125b2.825±0.350c2.600±0.668c
E. amylovora 4.725±0.229b4.475±0.189b4.225±0.125b4.125±0.335b3.825±0.338b3.700±0.248b3.350±0.185b
Negative control 4.950±0.189a4.900±0.168a4.850±0.210a4.725±0.210a4.575±0.155a4.275±0.165a4.225±0.165a
Alcaligen sp. 4.475±0.301b4.175±0.315b4.025±0.278b3.650±0.132b3.450±0.210b3.350±0.222b2.075±0.687c
P. penneli 4.900±0.158a4.650±0.064b4.225±0.217b4.150±0.328b3.925±0.075b3.675±0.222b3.370±0.111b
C. meningosepticum 4.800±0.248ab 4.325±0.131b4.275±0.214b4.100±0.231b4.050±0.266b3.600±0.286b2.675±0.373c
P. cepacia 4.875±0.125a4.300±0.108b4.200±0.265b3.925±0.269b3.625±0.304b3.400±0.300b2.750±0.366c
E. nigrifluence 4.300±0.187b4.000±0.212b3.975±0.225b3.775±0.029b3.475±0.502b3.300±0.168b3.200±0.456b
T. ptyseas 4.725±0.111b4.600±0.108b4.500±0.108b4.050±0.104b3.800±0.268b3.450±0.104b2.900±0.212c
C. indologenes 3.225±0.175d2.575±0.440d2.125±0.568d1.575±0.421c0.950±0.210d0.000±0.000d0.000±0.000d
S. pyogenes 3.800±0.141c3.500±0.178c3.425±0.189c2.125±0.111c1.950±0.333c1.425±0.878c0.450±0.287d
Note: Means with similar letters within a column are not significantly different at P < 0.05. Means were separated using DMRT test
838 WANJOHI ET AL.
soil were able to degrade oil. Erwinia nigrifluence,
Erwinia amylovora and Acinetobactor sp. were unable
to degrade oil. The results for Acinetobacter sp. were
in contrast with what was reported by Head et al.,
(2003). According to the author’s findings,
Acinetobacter sp. was unable to degrade oil.
The following families are known to degrade oil
Chryseobactericeae, Bacillaceae, Staphylococcaceae,
Sphingomonadaceae, Alcaligenaceae,
Burkholderiaceae, Pseudomonadaceae and
Moraxellaceae. Apart from these families, other
genera that have not been reported in oil
degradation were Providencia, Tatumella,
Streptococcus and Yersinia.
Burkhoderia cepacia was able to degrade oil. This
bacterium is known to metabolize chlorinated
hydrocarbon. These hydrocarbons are commonly
found in commercial pesticides and herbicides. This
bacterium can be added to sites that are
contaminated by these toxins to clean up the
environment. Burkhoderia cepacia is one of the most
effective bacteria in degrading the chemicals found
in house hold herbicide (weed-B-Gone) (Springer,
1992).
In conclusion, the bacteria that were able to
degrade oil included Providencia rettgeri, Morganella
morganii, Tatumella ptyseas, Bacillus anthracoides,
Chryseobacterium indologenes, Streptococcus pyogenes,
Chryseobacterium meningosepticum, Pseudomonas
cepacia, Proteus penneri, Alcaligene sp., Moraxella sp.,
Providencia stuarti and Sphingomonas paucimobilis.
Providencia rettgeri was the best in oil degradation.
Among these Providencia, Tatumella, Streptococcus
and Yersinia were novel species.
ACKNOWLEDGEMENT
This research was funded in part by the National
Council for Science and Technology (NCST), Kenya.
The authors express their gratitude to NCST,
University of Eldoret, Kenya Medical Research
Institute, Kenya Wildlife Services, Nakuru Water
and Sanitation Company, and to all others who
participated in this research.
REFERENCES
Atlas, R. M. 1991. Microbial hydrocarbon degradation,
bioremediation of oil spills. J. Chem. Tech. Biotech. 52:
149–156.
Atlas, R. 1981. Microbial-degradation of petroleum-
hydrocarbons-an environmental perspective.
Microbiol. Rev. 45 : 180-209.
Bossert, M., Kachel, W. and Bartha, R. 1984. Fate of
hydrocarbons during oily sludge disposal in soil.
Applied env. Microbiology. v 47.
Coral, G. and Karagoz, S. 2005. Isolation of phenanthrene
degrading bacteria from petroleum refinary soil.
Annals of MB. 55 (4) : 255-259.
Drozdowickz, A., Leite, S., Rosado, A. and Prantera, M.
2002. Degradation of gasoline aromatic
hydrocarbons by two N-2-fixing, soil bacteria.
Biotechnol Lett. 24: 85
Head, I. M., Jones, D. M. and Larter, S. R. 2003. Biological
activity in the deep subsurface and the origin of
heavy oil. Nature. v. 426: p. 344-352.
Hoff, Rebecca, Z. 1993. Bioremediation: an overview of
its development and use for oil spill cleanup. Marine
Pollution Bulletin. 29: 476-481.
Holt, J.G. 1994. Bergey’s Manual of Determinative
Bacteriology.
Kairu, J. 1991. Studies of the Concentration of Organic
Chlorine Pesticides and Metal Residues In Fish and
Birds of Lake Nakuru, Kenya. MSc. Thesis
Agricultural University, Norway.
Mastrangela, G., Fadda, E. and Marzia, V. 1997. Polycyclic
aromatic hydrocarbons and cancer in man.
environmental health aspect. v. 104 no. 11.
Oduor, S. O. and Schagerl, M. 2007. Temporal trends of
ion contents and nutrients in three Kenyan Rift Valley
saline-alkaline Lakes and their influence on
phytoplankton biomass. Hydrobiologia. 585 : 59.
Pothuluri, J. V. and Cerniglia, C.E. 1994. Microbial
metabolism of polycyclic aromatic hydrocarbons.”
In G.R. Chaudry, ed. Biological Degradation and
Bioremediation of Toxic Chemicals, Chapman and Hall
London.
Santos, V.D., Bartels, D., Bekel, T., Brecht, M., Buhrmester,
J. and Schneiker, S. 2006. Genome sequence of the
ubiquitous hydrocarbon-degrading marine
bacterium Alcanivorax borkumensis. Nat Biotechnol.
24: 9.
Spring, http://www.accessexcellence.org/AB/BA/A
Students Experiment.php (1994) Date of access: 8
october 2010 2:30pm
Springer, V. 1992. Prokaryotes. Vol 3, 2nd edition
Swamakaran, Hemalatha and Panchanathan
Veeramanikandan. 2011. Characterization of
aromatic hydrocarbon degrading bacteria from
petroleum contaminated sites. J. of Environmental
Protection. 2: 243-254. Chennai,India.
Yasindi, A. W., Lynn, D. H. and Taylor, W. D. 2002. Ciliated
protozoa in Lake Nakuru, a shallow alkaline-saline
Lake in Kenya: Seasonal variation, potential
production and role in the foodweb. Archive of fur
Hydrobiologia. 154 (2): 311-325.
Vareschi, E. 1982. Ecology of the lake Nakuru (Kenya) iii.
Synopsis of Abiotic factors and primary production.
Oecologia. 55: 45-5.