bioremediation of crude oil-contaminated soil in … 2/bioremediation...absence and presence of...

Journal of Environmental Treatment Techniques 2019, Volume 7, Issue 1, Pages: 179-195

179

Bioremediation of Crude Oil-Contaminated Soil

in the Presence of Nickel, Zinc and Cadmium

Heavy Metals Using Bacterial and Fungal

Consortia-Bioaugmentation Strategy

Samuel Enahoro Agarry1*, Ganiyu Kayode Latinwo1, Ebenezer Olujimi Dada1, Chiedu

Ngozi Owabor2

1Biochemical and Bioenvironmental Engineering Laboratory, Department of Chemical Engineering, Ladoke Akintola University of Technology, Ogbomoso, Nigeria

2Department of Chemical Engineering, Federal University of Petroleum Resources, Effurun, Delta State, Nigeria

Received: 10/01/2019 Accepted: 27/02/2019 Published: 01/06/2019

Abstract The study evaluated the effectiveness of indigenous bacterial consortia (Pseudomonas aeruginosa, Bacillus subtilis and

Micrococcus letus) and fungal consortia (Aspergillus niger, Aspergillus carmari and Penicillium notatum) as well as their combination (bacterial-fungal consortia) as bioaugmentation agents in the soil bioremediation of petroleum hydrocarbons in the absence and presence of nickel, zinc and cadmium heavy metals. Bioremediation was carried out in 10% w/w crude oil-contaminated soil microcosms for 35 days in the absence and presence of nickel, zinc, and cadmium bioaugmented with or

without bacterial, fungal and bacterial-fungal consortia, respectively. In the heavy metal-free soil microcosms, 72.5%, 64% and 90.7% total petroleum hydrocarbon (TPH) biodegradation were attained with bacterial, fungal and bacterial-fungal consortia, respectively, while 45% TPH biodegradation was achieved in the non-bioaugmented soil microcosm. In the heavy metal-soil microcosms: nickel, zinc, cadmium and mixed form (nickel + zinc + cadmium), 79.2%, 81.4%, 75.3% and 68.2% TPH biodegradation was correspondingly obtained with bacterial consortia; 69.4%, 66.4%, 68.2%, and 60.6% with fungal consortia; while 99%, 98.5%, 95.7%, and 100% was respectively attained with bacterial-fungal consortia. The kinetics of TPH biodegradation were adequately described by the first-order kinetics and half-life times were estimated. Soil microcosm bioaugmented with bacterial-fungal consortia displayed the highest biodegradation rate constant with the lowest half-life times in

the absence and presence of heavy metals. Therefore, the results suggest that microbial consortia (bacterial and fungal) could be very effective for soil bioremediation of crude oil in the presence of heavy metals.

Keywords: Bacteria; Bioaugmentation; Bioremediation; Crude oil; Fungi; Heavy metals

1 Introduction1

Nigeria is the largest oil producer in Africa with a maximum oil production capacity of 2.5 million barrel per day and the sixth largest oil producing country in the world [1]. All over the world, oil is transported through pipelines, vessels/ships, road, and rail, and as a result poses serious danger to the environment in case of spills. In 2005, almost every day, about nine incidents of oil pollution were

reported around the world [2]. Since 1960 till date, over

Corresponding author: Samuel Enahoro Agarry, Biochemical and Bioenvironmental Engineering Laboratory, Department of Chemical Engineering, Ladoke Akintola University of Technology, Ogbomoso, Nigeria. E-mail: [email protected].

four thousand incidents of crude oil spills have been estimated to have occurred in the Niger Delta region of Nigeria, amounting to several million barrels of crude oil [3]. About 90% of the contaminated sites in the USA are

soils that are contaminated with petroleum hydrocarbons [4]. Crude oil consists of complex mixture of aliphatic and aromatic hydrocarbons, asphaltene and resins which are considered to be environmental pollutants [5]. In addition, trace amount of heavy metals have been recognized to exist in crude oils [3, 6]. The metals commonly considered as pollutants includes aluminium, arsenic, cadmium,

chromium, copper, mercury, nickel, lead, iron, zinc and some radionuclides [7]. These hydrocarbons and heavy metals have been found to be toxic, mutagenic and carcinogenic [8]. However, some of these metals such as

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copper, chromium, nickel, zinc and iron play an integral role as micronutrients in the life metabolic processes of microbes and can as well at higher concentrations become toxic and inhibit various cellular or biochemical processes by forming unspecific complex compounds within the

microbial cell [9]. While several other metals like cadmium, lead, silver and mercury are not essential, do not have any biological role and are potentially toxic to microorganisms [9]. The toxicity of these hydrocarbons and metals necessitated the need to develop methods or technologies that are environmentally-friendly to remediate oil spills contamination of the environment (soil or water). One of

such technology is bioremediation due to its efficiency and cost-effectiveness as compared to physicochemical technologies that are expensive [10]. Microbial species that utilizes hydrocarbons as a source of carbon and energy are referred to as hydrocarbonoclastic microorganisms. They are the major agents of bioremediation and thus petroleum hydrocarbons degradation due to their associated metabolic abilities [2]. Reviews of literature on the petroleum

hydrocarbons degradation have confirmed that several microorganisms majorly bacteria and fungi are capable of utilizing petroleum hydrocarbons as the sole source of carbon and energy [2, 11, 12]. The desire for environmentally-friendly and sustainable technology towards contaminated environment has made the evolving bioremediation technology a standard practice for the treatment and restoration of contaminated

environment. Over three decades, considerable studies have been carried out in both the laboratory and field in the area of bioremediation with different modifications such as nutrient supplementation (biostimulation) [13] and microbial inoculation (bioaugmentation) for the remediation of diverse range of contaminants [14 – 16]. In all of these studies which did not involve heavy metals interference, bioaugmentation strategy has been found to be successful.

Bioaugmentation is normally recommended for contaminated sites where the autochthonous microbial populations is insufficient for contaminants degradation and/or those in which the indigenous microorganisms do not possess the necessary catabolic pathways for contaminants metabolism [17, 18]. However, for environment such as soil and water co- contaminated with both metals and organic compounds, they are considered to

be crucially difficult to treat due to the mixed nature of the pollutants [19]. Thus a co-contaminated environment represent a serious problem in the bioremediation processes [20]. This is because metals cannot be biologically degraded or modified like toxic organic compounds, but their speciation and bioavailability may change with varying environmental factors [21, 22]. Also, metals may inhibit the hydrocarbon biodegradation through its

interaction with microbial enzymes that are directly involved in the biodegradation or through its interaction with microbial enzymes involved in general metabolism [23]. Nevertheless, to overcome this difficulty is to either employ the use of bioaugmentation with microorganisms that has the ability to resist and detoxify metal as well as the use of organic compound-degrading microorganisms

[24] or the use of phytoremediation [25, 26]. There are certain number of metal-resistant microorganisms that can detoxify metals, such as cadmium [27, 28]. These metal-resistant and detoxifying microorganisms possesses the capability to exhibit some

metal resistance mechanisms that allows them to function in a metal co-contaminated environments and these include: intracellular and extracellular sequestration; extracellular precipitation; redox transformation; membrane efflux system; exclusion by permeability barrier; and enzymatic detoxification [9]. Despite the bioremediation successes that have so far been recorded using bioaugmentation strategies, very few studies involving bioaugmentation

strategy with the use of either single strain/pure culture or mixed culture of two microbial strains for the bioremediation of petroleum hydrocarbons such as polycyclic aromatic hydrocarbons (PAHs) and heavy metals co-contamination have been carried out [20, 29 – 32]. From these studies, varying forms and degrees of organic compound degradation have been reported. For instance, Alisi et al. [20] reported that complete

degradation of hydrocarbon n-C12–20 and total disappearance of phenanthrene in diesel oil as well as 75% diesel oil reduction was achieved by bacterial bioaugmentation in the presence of heavy metals. Owabor et al. [29] also reported that the degradation of naphthalene in soil by inoculated microbial consortium made up of Bacillus and Aspergillus niger species was gradually inhibited as the concentration of each of the heavy metals

(Pb, Hg, Ni and Cr) increased from 40 to 200 mg/L. Bioaugumentation strategies involving the use of a consortium consisting of Acremonium sp. and Bacillus subtilis demonstrated high degradation efficiency in soil that is heavily contaminated with crude oil [33], while the presence of heavy metals have been revealed to impact on the fungal-bacterial synergism in polycyclic aromatic hydrocarbons (PAHs) degradation [31]. In spite of these information that revealed that soil

bioremediation in the presence of heavy metals can be achieved with bioaugmentation strategy, the strategy is still faced with a number of challenges with regard to the presence of heavy metals, which often result in the toxicity and inhibition of microbial growth and limit degradation ability. There is still paucity of information on the influence of heavy metals as single or mixture form on bioaugmentation strategy using mixed bacterial consortia,

fungal consortia and bacterial-fungal consortia in the bioremediation of petroleum hydrocarbons in crude oil contaminated soil. Hence, this present study aimed to explore the effectiveness of mixed indigenous bacterial consortia, mixed indigenous fungal consortia and mixed bacterial-fungal consortia as bioaugmentation agents in the soil bioremediation of petroleum hydrocarbons in crude oil in

the absence and presence of heavy metals (nickel, zinc and cadmium) in single or mixture form. The rates of petroleum hydrocarbons biodegradation were determined from the application of first-order kinetics and the biodegradation half-life times were estimated.


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2. Materials and Methods 2.1 Chemicals and Reagents

All the chemicals and reagents are of analytical grade

such as: n-hexane, calcium chloride solution, zinc chloride,

cadmium sulphate, nickel nitrate, nutrient agar (NA), and

potato dextrose agar (PDA).

2.2 Sample Collection

Soil samples were obtained from a farmland in Oleh

town (Latitude 5.4589° N and Longitude 6.2031° E) of

Delta State in the Niger-Delta region of Nigeria. It has no

pollution history and devoid of hydrocarbon contamination.

It was the top soil layer of the farmland not exceeding a

depth of 25 cm from the surface that was excavated with a

shovel, collected into a black polyethylene bag and brought

to the laboratory. Crude oil used for artificial pollution was

obtained from Warri Refinery and Petrochemical Company

located in Warri (5.5544° N, 5.7932° E) in the Niger-Delta

region of Nigeria. The microbial strains used in this study

were cultured from the soil sample obtained from a

previously oil contaminated soil in the Niger-Delta region

of Nigeria.

2.3 Characterization of Soil

Physical, chemical and microbiological properties of the

top soil samples at the onset of the remediation were

characterized. The soil samples were sieved, pulverized and

air-dried. Then, the samples were analyzed for pH,

moisture content, total organic carbon, total nitrogen, and

available phosphorus using standard methods [34]. The soil

was also analyzed for total hydrocarbon degrading bacteria

(THDB) and total hydrocarbon degrading fungi (THDF)

count using pour plate method [28]. The metals (calcium,

magnesium, nickel, zinc and cadmium) concentrations in

the soil samples were determined after wet digestion of 0.5

g of the soil n 10 mL of concentrated HNO3 and HClO4

(7:1 v/v) and the concentrations measured using atomic

absorption spectrometry (AAS) with a flame furnace

nebulizer (Perkin-Elmer). The physical and chemical

characteristics of the soil were as follows: pH (6.8 ± 0.3),

moisture content (10.2 ± 0.2%), total organic carbon (0.76

± 0.03%), total nitrogen (0.12 ± 0.01%), available

phosphorus (0.46 ± 0.24%), nickel (< 0.01 mg/kg), zinc (<

0.01 mg/kg), cadmium (< 0.01 mg/kg), calcium

(14.24meq/100g) and magnesium (9.85meq/100g). The

microbiological properties of the soil were as follows:

THDB count (0.8 ± 0.01 ×102 cfu/g) and THDF count (0.2

± 0.03 ×102 cfu/g). This revealed that the population of the

hydrocarbon degrading microorganisms in the test soil

sample is very small and thus there is the need to bio-

augment the soil with more hydrocarbon degrading

microorganisms.

2.4 Source of Bacterial and Fungal Consortia

The bacteria and fungi species used for this study were

isolated from the collected oil polluted soil samples using

the serial dilution and pour or spread plate method [28].

The soil samples were sieved and 10 g of it were added into

90 mL de-ionized water in a conical flask, vigorously

shaken and left overnight. For bacteria isolation, 0.1 mL

aliquots of clear soil suspension and its dilutions (from 10-1

– 10-3) were poured and spread on a NA plate containing

crude oil droplets, Nystatin (to prevent fungal growth) and

the plate was then incubated at 37°C for 24 hours; and the

grown bacterial colonies were isolated [35]. For fungi

isolation, 0.1 mL aliquots of the soil suspension and its

dilution (10-1 – 10-3) were introduced into sterile Petri

dishes containing PDA and droplets of crude oil and then

incubated at ambient temperature (28 ± 2oC) for 72 – 120

hours. Streptomycin antibiotic (10 mg/L) was added into

the PDA medium to inhibit any form of bacterial growth.

After 3-5 days of fungal growth, the spore bearing mycelia

were then carefully sectioned, teased out and stained on a

slide using lactophenol cotton blue stain and later observed

with a light microscope. The bacterial and fungal colonies

with different morphologies obtained after incubation were

sub-cultured and purified repeatedly by streaking on sterile

NA and PDA plates, respectively. Pure cultures of isolated

bacterial species were identified and characterized on the

basis of Bergey’s manual [36]. Standard conventional

methods that involved various cultural, morphological,

physiological and biochemical tests were performed

namely gram staining, catalase test, indole test, citrate

utilisation test, urease test, motility test, oxidase test,

coagulase test, glucose utilization, fructose utilization,

lactose utilization, lipase test [37]. The isolated bacterial

species were identified as Pseudomonas aeruginosa,

Bacillus subtilis, Staphylococcus aureus, Staphylococcus

epidermidis, Escherichia coli and Micrococcus letus.

Pseudomonas, Bacillus, Staphylococcus, E.coli and

Micrococcus species have been reported to have the

potential to biodegrade petroleum hydrocarbons [38, 39].

The fungal isolates were identified and characterized on the

basis of cultural, microscopic (septation of mycelium,

shape, form, diameter and texture of spore/conidia), and

macroscopic (pigmentation, shape, diameter, colony

appearance and texture) features [28, 40]. The cultural and

morphological features of the fungal isolates were then

compared with those described by Samson et al. [41] as

well as using the keys of Mackie and McCartney [42]. The

fungal isolates were identified as Aspergillus niger,

Aspergillus carmari, and Penicillium notatum. Aspergillus

and Penicillium species have been confirmed to have the

potential to biodegrade the petroleum hydrocarbons in

crude oil [43, 44].

2.5 Heavy Metal Tolerance Index of Bacterial and Fungal

Isolates

Isolated bacterial and fungal species were assessed

either as pure isolates or as mixed isolates (consortium) for


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their heavy metal tolerance or resistance to Cd2+, Zn2+ and

Ni2+. Eight mm diameter circular extract or disks from 10

day old pure cultures of each bacterial isolates and fungal

isolates were inoculated into separate corresponding NA

and PDA Petri-dishes or plates supplemented with each of

the sterilized 31.13 mg/L Ni (NO3)2, 20.84 mg/L ZnCl2 and

18.55 mg/L CdSO4 salts solution corresponding to 10 mg/L

each of Ni2+, Zn2+ and Cd2+ heavy metals, respectively. The

inoculated plates were incubated at ambient temperature

(28±2°C) for at least 7 to 10 days to establish their growth.

The experiments were performed in triplicates with

bacterial isolates in NA plate and fungal isolates in PDA

plate without supplementation with heavy metals that

served as control. The radial growth was determined by

measurement of the culture spread from the center of the

colony or inoculated portion. The inoculated portion

diameter was subtracted from the diameter of growth [45].

The tolerance index (TI), which is an indication of the

microbes response to the stress of heavy metal was

calculated from the growth of microbial isolates exposed to

the heavy metals divided by the growth of the microbial

isolates in the absence of heavy metals [27]. The microbial

isolates heavy metal tolerance can be rated as follows [28]:

0.00–0.39 (very low tolerance), 0.40–0.59(low tolerance),

0.60–0.79 (moderate tolerance), 0.80–0.99 (high tolerance)

and 1.00- >1.00 (very high tolerance). The higher the TI

values, the higher the microbial species tolerance or

resistance to the heavy metal.

2.6 Preparation of Bacterial and Fungal Inoculum

The bacterial and fungal inoculums were prepared

according to the method of Ma et al. [15]. To prepare and

generate the bacterial inoculum required for

bioaugmentation, the bacterial isolates were separately

grown in a NA broth at 30 oC and shaken in an orbital

shaker at 120 rpm for 24 h. Thereafter, the cultures were

centrifuged at 10,000 rpm for 10 min. Then, the cell pellets

obtained were washed twice with phosphate buffer (0.1 M,

pH 7.0), and again re-suspended in a fresh phosphate buffer

and the resulted suspensions were mixed together in equal

proportions and used as mixed bacterial culture (bacterial

consortia) for the study. Similarly, to prepare fungal

inoculums for bioaugmentation, the fungal isolates were

grown separately in a PDA liquid medium at 30 oC and

shaken in an orbital shaker at 100 rpm for 72 h. Then, the

cultures were centrifuged and the fungal mycelia obtained

were washed twice with phosphate buffer, and thereafter re-

suspended in a fresh phosphate buffer and the resultant

suspensions were mixed in equal proportions and used as

mixed fungal culture (fungal consortia). The volume of the

liquid medium was 30 mL with 106 cfu/ml of either

bacteria or fungi to be added to each of the contaminated

soil microcosms.

2.7 Experimental Design and Bioremediation Protocol

The soil sample from the farmland was sieved with a 5

mm mesh sieve to remove large stones and debris.

Thereafter, the sieved soil samples were sun-dried by

spreading on a flat clean surface for one week and then

pulverized. A known weight of the soil sample (150 g) was

each introduced into sixteen separate clean dry plastic

containers (i.e. soil microcosm) and moistened to about

20% water holding capacity with sterilized de-ionized

water. Thereafter, 18 mL of crude oil corresponding to 15 g

(i.e. 10% w/w) was measured and introduced into each of

the soil in the sixteen plastic containers and the contents

were thoroughly stirred. The sixteen soil microcosms were

labelled B-T1 to B-T16. The 10% w/w artificial

contamination was adopted so as to achieve severe

contamination because beyond 3% w/w concentration, oil

has been reported to be increasingly deleterious to soil biota

and crop growth [13, 46]. Soil microcosm B-T1 served as

the control (i.e. natural bioattenuation). After the oil

contamination, 31.13 mg/L of nickel nitrate (Ni(NO3)2),

18.55 mg/L cadmium tetraoxosulphate (VI) (CdSO4), and

20.84 mg/L zinc chloride (ZnCl2) solutions each of which

corresponded to 10 mg/L of Ni2+, Cd2+ and Zn2+

concentrations, respectively, were added singly into the

contaminated soil microcosms while 70.52 mg/L of mixed

heavy metal salts solution corresponding to 30 mg/L (Ni2+

+ Zn2+ + Cd2+) concentration was added as shown in Table

1. The soil microcosms were left for 14 days to allow for

aging or equilibration. After day 14, mixed bacterial

(Pseudomonas aeruginosa, Bacillus subtilis and

Micrococcus letus) and mixed fungal (Aspergillus niger,

Aspergillus carmari, and Penicillium notatum) inoculums

were inoculated into the contaminated soil microcosms as

also shown in Table 1 and thoroughly mixed together with

a stirring rod. The inoculated contaminated soil

microcosms were covered with aluminum foil and then

incubated for 35 days. At intervals of 3 days, small volume

(30 mL) of sterilized de-ionized water was added to the

contaminated soil microcosms and stirred together so as to

maintain the moisture content. At intervals of 7 days, soil

sample was taken to carry out analyses for residual TPH,

microbial (bacterial and fungal) count and pH, respectively.

2.8 pH Measurement

The pH of the soil samples was measured in the course

of the bioremediation studies according to the following

procedure: 10 g of the soil sample was accurately weighed

and added into 10 mL of de-ionized water in a conical

flask. The mixture was allowed to stand for 15 min after

which the flask was placed in an orbital shaker (SSL1-

model) and agitated at 150 rpm for 30 min. At the end of

the agitation, the mixture was allowed to stand for 10 min

and the pH value was read on an already calibrated pH

meter (JENWAY 3020-model).


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Table 1: Soil microcosm artificially contaminated with crude oil and heavy metal and bioaugmented with microbial species

Soil Microcosm Code Description

B-T1 (Control) Soil + Oil (Natural Bioattenuation)

B-T2 Soil + Oil + Bacterial Consortium B-T3 Soil + Oil + Bacterial Consortium + Nickel B-T4 Soil + Oil + Bacterial Consortium + Zinc B-T5 Soil + Oil + Bacterial Consortium + Cadmium B-T6 Soil + Oil + Bacterial Consortium + Nickel + Zinc + Cadmium B-T7 Soil + Oil + Fungal Consortium B-T8 Soil + Oil + Fungal Consortium + Nickel B-T9 Soil + Oil + Fungal Consortium + Zinc

B-T10 Soil + Oil + Fungal Consortium + Cadmium B-T11 Soil + Oil + Fungal Consortium + Nickel + Zinc + Cadmium B-T12 Soil + Oil + Bacterial- Fungal Consortium

B-T13 Soil + Oil + Bacterial-Fungal Consortium + Nickel B-T14 Soil + Oil + Bacterial-Fungal Consortium + Zinc B-T15 Soil + Oil + Bacterial-Fungal Consortium + Cadmium B-T16 Soil + Oil + Bacterial-Fungal Consortium + Ni + Zn + Cd

2.9 Determination of Total Petroleum Hydrocarbon

The total petroleum hydrocarbon (TPH) content of the

artificially contaminated soil samples was determined

according to the following procedure: 5 g of the soil sample

was weighed into a plastic bottle and 25 mL of n-hexane

was added to the soil sample and the mixture shaken on an

orbital shaker (SSL1-model) at 250 rpm for 20 min and

then allowed to stand. Filtration of the mixture was done

and the hexane solvent in the filtrate was allowed to

evaporate at room temperature in a fume hood. The

concentration of extracted residual TPH in the filtrate was

then determined or measured at an absorbance of

wavelength 400 nm using the UV-VIS Spectrophotometer

(JENWAY 6715-model).

2.10 Determination of Total Hydrocarbon-Degrading

Bacterial and Fungal Count

Quantification of the total hydrocarbon-degrading

bacteria (THDB) and total hydrocarbon-degrading fungi

(THDF) present in the soil samples was determined by the

pour plate count method [28]. Soil samples (10 g) was

transferred into sterilized Erlenmeyer conical flasks

containing 90 mL of sterile 0.9% (m/v) NaCl solution and

then shaken in an orbital shaker at 150 rpm for 15 min.

Samples (1mL) were subjected to a serial 10-fold dilution

procedure and cultivated in a NA medium for THDB at 30 oC for 48 h and in a PDA medium for THDF at 30 oC for 72

h, respectively. The number of colony forming units (cfu)

was counted in each sample and expressed as colony-

forming units per gram of dry soil (cfu/g dry soil). All

microbiological counts and experiments were carried out in

triplicate. The TPH concentration was expressed as mg of

petroleum hydrocarbons per kg of dry soil [13]. Eq. (1) was

employed to calculate the percent TPH biodegradation [13]:

100%

o

fo

TPH

TPHTPHTPH (1)

where oTPH and fTPH are the initial and residual TPH

concentrations, respectively.

2.11 Determination of TPH Biodegradation Rate and

Half-life Time

The rate of TPH biodegradation was determined from

the application of first-order kinetic model equation as

given in Eq. (2) [13, 47]:

kt

ot eTPHTPH (2)

where TPHO and TPHt are the initial TPH and residual TPH

concentration (mg/kg) at time t in soil (mg/kg), k is the

biodegradation rate constant (day−1), and t is time (day).

The biodegradation half-life time (t1/2) was calculated from

the biodegradation rate constant (k) using Eq. (3) [13, 47]:

k

2ln (3)

3. Results and Discussion 3.1 Heavy Metal Tolerance Index of Bacterial and Fungal

Isolates

In ascertaining the tolerance of the bacterial and fungal isolates to heavy metals (Ni2+, Zn2+ and Cd2+) of 10 mg/L concentration, the tolerance index (TI) was estimated and the values are presented in Table 2. The tolerance rating of Pseudomonas aeruginosa and Bacillus subtilis (being

bacterial isolates) as well as Aspergillus niger, Aspergillus carmari and Penicillium notatum (fungal isolates) to 10 mg/L each of Ni2+, Zn2+ and Cd2+ were observed to be high, with TI ranging between 0.80 and 0.99. Similar observations have been reported for Aspergillus sp and


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Penicillium sp as well as Pseudomonas aeruginosa and Bacillus subtilis using higher concentration of Cd [48, 49], Ni [49] and Zn [49], respectively. In addition, high tolerance of Pseudomonas sp., Bacillus sp, Aspergillus sp.,and Penicillium sp. for heavy metals such as Zn, Ni and

Cd has been reported [50, 51]. Meanwhile, the tolerance rating of Pseudomonas aeruginosa and Bacillus subtilis (being bacterial isolates) as well as Aspergillus niger, Aspergillus carmari and Penicillium notatum (fungal isolates) to 30 mg/L of the combined or mixed heavy metals (Ni2+ + Zn2+ + Cd2+) were observed to be moderately low, with TI ranging between 0.10 and 0.62.

Pseudomonas aeruginosa showed a relatively higher TI

for Ni2+, Zn2+ and Cd2+ and this was followed by Bacillus subtilis, Micrococcus letus, Escherichia coli, Staphylococcus aureus and Staphylococcus epidermidis, respectively. Aspergillus niger showed a relatively higher TI for Zn2+ and Cd2+ and then followed by Aspergillus carmari and Penicillium notatum, respectively. Furthermore, the heavy metal tolerance by the six bacteria species (Pseudomonas aeruginosa, Bacillus subtilis,

Micrococcus luteus, Escherichia coli, Staphylococcus aureus and Staphylococcus epidermidis) is in this decreasing order of Zn > Ni > Cd while the metal tolerance by the three fungal species (Aspergillus niger, Aspergillus carmari and Penicillium notatum) is in this decreasing order of Zn > Cd > Ni. A similar observation has been reported by Oaikhena et al. [52] for metal tolerance by five bacteria species (Pseudomonas aeruginosa, Staphylococcus

aureus, Escherichia coli, Proteus vulgaris and Klebsiella pneumoniae). The higher tolerance levels for zinc and nickel could be attributed to their classification as micronutrients that are needed by the bacteria in trace

amount [53]. On the other hand, Smrithi and Usha [54] have reported the metal tolerance by bacteria isolates from tannery effluent in the decreasing order of Ni > Zn > Cd while Amalesh et al. [55] stated that the decreasing order for metal tolerance is Cd > Ni.

On the other hand, the tolerance ratings of the bacterial consortia (Pseudomonas aeruginosa, Bacillus subtilis and Micrococcus letus), fungal consortia (Aspergillus niger, Aspergillus carmari and Penicillium notatum) and bacterial-fungal consortia (Pseudomonas aeruginosa, Bacillus subtilis, Micrococcus letus, Aspergillus niger, Aspergillus carmari and Penicillium notatum) to 10 mg/L each of Ni2+, Zn2+ and Cd2+ were observed to be very high,

with TI ranging between 1.81 and 2.38. These very high TI values showed that microbial consortia can tolerate the presence of heavy metals more than the individual component in the consortium. In addition, the TI for the combined mixture of heavy metals (Ni2+ + Zn2+ + Cd2+) was found to be 0.83, 0.80 and 1.66 for bacterial, fungal and bacterial-fungal consortia, respectively. These relatively high TI values indicate that microbial consortia

can tolerate the presence of combined or mixed heavy metals more than their separate components in the consortium.

3.2 Bacterial and Fungal Consortia Degradation of

Petroleum Hydrocarbons Figure 1 shows the level of TPH biodegradation in non-bioaugmented soil microcosm (B-T1) and in soil microcosm

bio-augmented alone with bacterial consortium (microcosm B-T2), fungal consortium (microcosm B-T7) and bacterial-fungal consortium (B-T12), respectively.

Table 2: Tolerance index of bacterial and fungal isolates in growth media supplemented with 10 ppm of heavy metal

Microbial Isolates Tolerance Index

Cd2+ Ni2+ Zn2+ Ni2+ + Zn2+ + Cd2+

Pseudomonas aeruginosa 0.88 0.90 0.92 0.62 Bacillus subtilis 0.80 0.88 0.85 0.58 Micrococcus luteus 0.60 0.77 0.80 0.44 Escherichia coli 0.55 0.75 0.70 0.30 Staphylococcus aureus 0.25 0.45 0.56 0.15 Staphylococcus epidermidis 0.20 0.45 0.54 0.10

Aspergillus niger 0.85 0.80 0.99 0.61 Aspergillus carmari 0.82 0.80 0.94 0.60 Penicillium notatum 0.80 0.80 0.90 0.56 Bacterial consortium 1.86 1.88 1.90 0.83 Fungal consortium 1.84 1.81 1.87 0.80 Bacterial-fungal consortium 2.25 2.34 2.38 1.66


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Figure 1: TPH biodegradation profile by bacterial consortium, fungal consortium and bacterial-fungal consortium in the absence of heavy metals

(Nickel, Zinc and Cadmium). Bars indicate the average of triplicate samples while the error bars show the standard deviation.

As shown in Figure 1, within the first 21 days of remediation in the soil microcosms B-T2, B-T7 and B-T12, a rapid decrease in TPH amounting to over 50% degradation was attained. The degradation efficiency of these consortia increased with time and at the end of day 35 remediation period, 100, 000 mg/kg of TPH concentration was reduced to 27,500 ± 1750, 36,000 ± 1900 and 9,300 ± 1100 mg/kg

TPH corresponding to 72.5%, 64% and 90.7% biodegradation or reduction in soil bio-augmented with bacterial consortium (B-T2), fungal consortium (B-T7) and bacterial-fungal consortium (B-T12), respectively. Meanwhile, in the non-bioaugmented soil microcosm (B-T1), the TPH concentration of 100,000 mg/kg was reduced to 55,000 mg/kg amounting to 45% TPH biodegradation at the end of day 35 remediation period. The bacterial

consortium, fungal consortium and bacterial-fungal consortium correspondingly resulted in an increase of 61%, 42.2% and 101.5% TPH biodegradation in relation to the percent TPH biodegradation obtained in the non-bioaugmented soil microcosm (natural bioattenuation) or control. The increased degradation by the consortium in the bio-augmented soil microcosms as compared to the non-bio-augmented microcosm may be attributed to greater

population of TPH degraders. Similarly, the bacterial-fungal consortium resulted in an increase of 25% and 41.7% TPH biodegradation with respect to the percent degradation attained by the bacterial consortium and fungal consortium, respectively. These increases in TPH biodegradation represent a significant difference (p < 0.05)

in the different consortium biodegradation efficiency. The higher percent TPH biodegradation observed in bacterial-fungal consortium might be ascribed to the synergistic metabolic activities by the consortium of three bacteria and three fungi isolates with larger number of different types of enzymes which might have enhanced the degradation process than separate bacterial and fungal consortiums with

fewer types of enzymes. These results thus suggest that the consortium which consists of both bacterial and fungal TPH degraders may lead to very rapid and more complete degradation as compared to separate consortium of bacteria or consortium of fungi and it emphasized the role of fungal species with degradation capability in the synergistic TPH degradation. Also, the bacterial consortium yielded an increase of 11% compared to the fungal consortium.

3.3 Influence of Heavy Metals on Bacterial and Fungal

Consortia Degradation of TPH The influence of the presence of heavy metals (Ni2+, Zn2+ and Cd2+) as individual metals and as combined or mixture form on bacterial consortium, fungal consortium and bacterial-fungal consortium degradation of TPH was evaluated. Figure 2(A) shows the level of TPH

biodegradation in soil microcosm bio-augmented alone with bacterial consortium (B-T2), bacterial consortium + Ni2+ (B-T3), bacterial consortium + Zn2+ (B-T4), bacterial consortium + Cd2+ (B-T5), and bacterial consortium + Ni2+ + Zn2+ + Cd2+ (B-T6), respectively.

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Figure 2: (A) TPH biodegradation profile by bacterial consortia in the absence and presence of heavy metals (Nickel, Zinc and Cadmium). (B)

Amount of heavy metal (Nickel, Zinc and Cadmium) bioavailable to bacterial consortia. Bars indicate the average of triplicate samples while the

error bars show the standard deviation.

A rapid decrease in TPH amounting to over 50% reduction was attained within the first 21 days of the remediation in all the soil microcosms augmented with bacteria and heavy metals. At the end of day 35 remediation period, 100,000 mg/kg of TPH concentration was reduced to 27,500 ± 1750, 20,800 ± 1500, 18,600 ± 1450, 24,700 ± 1300 and 31,800 ± 2000 mg/kg TPH corresponding to 72.5%, 79.2%, 81.4%, 75.3% and 68.2%

degradation or reduction in soil bio-augmented with bacterial consortium-metal free, bacterial consortium + Ni2+, bacterial consortium + Zn2+, bacterial consortium + Cd2+, and bacterial consortium + Ni2+ + Zn2+ + Cd2+, respectively. The results revealed that the individual presence of Ni2+, Zn2+ and Cd2+ exerted a significant positive influence on the TPH bacterial consortium degradation by increasing the TPH percent degradation by 9.2%, 12.3% and 3.9%, respectively, relative to the TPH

degradation by the bacterial consortium that is heavy metal-free. The relatively enhanced TPH biodegradation or reduction may be due to the relative bioavailability of the individual metals to the bacterial consortium (Figure 3(A)) which tend to stimulate growth as well as to the high metal TI of the bacterial consortium (Table 2). While the combined equal concentration mixture of Ni2+, Zn2+ and Cd2+ (30 mg/L) exerted a considerable negative impact on

the TPH bacterial consortium degradation by significantly (p < 0.05) suppressing or decreasing the TPH bacterial degradation by 5.9%. This indicated that the presence of mixed heavy metals (or multi-heavy metals) acts synergistically to impose a greater inhibitory response to the growth of the bacterial consortium thereby imposing a relatively partial inhibition to the degradation than that imposed by single or individual heavy metals. A similar

observation has been reported [56]. The different bioaugmentation treatments ranking with respect to increasing TPH biodegradation level was B-T6 < B-T2 < B-T5 < B-T3 < B-T4. That is, in all the soil microcosms bio-

augmented with bacterial consortium, the highest percentage of TPH biodegradation or reduction was achieved in soil microcosm that contained the presence of Zn2+ (81.4%). This is relatively and closely followed by that of bacterial consortium + Ni2+ (79.2%), bacterial consortium + Cd2+ (75.3%), bacterial consortium alone (72.5%), and bacterial consortium + Ni2+ + Zn2+ + Cd2+ (68.2%), respectively. Thus there are significant differences

(p < 0.05) in the percent TPH degradation influenced by the individual and combined mixture of Zn2+, Ni2+ and Cd2+ heavy metals. This observation means that the degree of bioavailability of the heavy metals to the bacterial consortium differs as shown in Figure 2(B). Figure 2(B) reveals that Zn2+ was more bioavailable to the consortium and followed by Ni2+ and Cd2+, respectively. Therefore, the potential bioavailability of Ni2+, Zn2+ and Cd2+ in soil strongly decreased after bioaugmentation with bacterial

consortium after 35 days’ incubation. However, bioavailable mixed heavy metals (Ni2+ + Zn2+ + Cd2+) in soil microcosm only decreased slightly after 35 days’ incubation. Thavamani et al. [30] reported that there was complete phenanthrene biodegradation in soil solution by bacterial consortium (Alcaligenes sp., Pseudomonas sp., Pandoraea sp. and Paenibacillus sp) in the presence of 5 mg/L of cadmium. Nonetheless, according to Sandrin and

Hoffman [57] making comparisons between heavy metals concentrations that inhibit organic compounds biodegradation as reported by different researchers is difficult. This is because more often than not, significant variation is usually observed in the inhibitory range for heavy metals due to different conditions of experiments used such as time of exposure, pH etc. [58]. Figure 3(A) shows the TPH biodegradation profile in soil microcosms

bio-augmented with fungal consortium-metal free, fungal consortium + Ni2+, fungal consortium + Zn2+, fungal consortium + Cd2+, and fungal consortium + Ni2+ + Zn2+ + Cd2+, respectively.

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Figure 3: (A) TPH biodegradation profile by fungal consortia in the absence and presence of heavy metals (Nickel, Zinc and Cadmium). (B)

Amount of heavy metal (Nickel, Zinc and Cadmium) bioavailable to fungal consortia. Bars indicate the average of triplicate samples while the

error bars show the standard deviation.

Similarly, within the first 21 days of the remediation period the decrease observed in TPH resulted to about 50% reduction in all the soil microcosms augmented with fungal consortium and heavy metals. At day-35 remediation period, the TPH concentration of 100,000 mg/kg was reduced to 36,000 ± 1900, 30,600 ± 1700, 34,200 ± 1600, 31,800 ± 1500 and 39,400 ± 1850 mg/kg TPH which corresponded to 64%, 69.4%, 66.4%, 68.2%, and 60.6%

TPH biodegradation in soil augmented with fungal consortium, fungal consortium + Ni2+, fungal consortium + Zn2+, fungal consortium + Cd2+, and fungal consortium + Ni2+ + Zn2+ + Cd2+, respectively. Similarly, in all the soil microcosms bio-augmented with fungal consortium, the microcosm containing Ni2+ had the highest percentage of TPH reduction (69.4%) and was relatively followed by that of fungal consortium + Cd2+ (68.2%), fungal consortium +

Zn2+ (66.4%), fungal consortium-metal free (64%) and fungal consortium + Ni2+ + Zn2+ + Cd2+ (60.6%), respectively. Hence, the bioaugmentation treatments ranking with respect to increasing TPH biodegradation level is B-T11 < B-T7 < B-T9 < B-T10 < B-T8. Thus there are significant differences (p < 0.05) in the percent TPH biodegradation influenced by the individual and combined mixture of Ni2+, Zn2+ and Cd2+ heavy metals. This

observation indicated that Ni2+, Zn2+ and Cd2+ exerted a relatively positive influence and thus enhanced the TPH biodegradation by the fungal consortium by 8.4%, 6.6%

and 3.8%, respectively, relative to the percent TPH biodegradation attained by the fungal consortium-metal free. This enhancement could also be due to the relative bioavailability of the individual metals to the fungal consortium as shown in Figure 3(B) as well as the concentration tolerance of the individual heavy metals by the consortium (Table 2). Figure 3(B) reveals that Ni2+ was more bioavailable to the consortium and followed by Cd2+

and Ni2+, respectively. Meanwhile, the combined mixture of Ni2+ + Zn2+ + Cd2+ of 30 mg/L concentration exerted a considerable negative impact on the TPH fungal consortium degradation by significantly (p < 0.05) decreasing the TPH fungal consortium degradation by 5.3%. This observation also revealed that the presence of mixed heavy metals with relatively higher concentration acts synergistically to impose a greater inhibitory response

to the growth of the fungal consortium hence imposing a partial inhibition to the TPH degradation than that imposed by single heavy metals. A similar observation has been reported (Anahid et al., 2011).

The TPH biodegradation profile in soil microcosms bio-augmented with bacterial-fungal consortium (metal-free), bacterial-fungal consortium + Zn2+, bacterial-fungal consortium + Ni2+, bacterial-fungal consortium + Cd2+, and

bacterial-fungal consortium + Ni2+ + Zn2+ + Cd2+, respectively, is shown in Figure 4(A).

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Figure 4: (A) TPH biodegradation profile by bacterial-fungal consortium in the absence and presence of heavy metals (Nickel, Zinc and

Cadmium). (B) Amount of heavy metal (Nickel, Zinc and Cadmium) bioavailable to bacterial-fungal consortium. Bars indicate the average of

triplicate samples while the error bars show the standard deviation.

Similarly, a fast decrease in TPH concentration resulting to over 70% reduction was achieved within day 21 of the remediation period in all the soil microcosms bio-augmented with bacterial-fungal consortium and heavy

metals. At the end of day 35, the TPH concentration (100,000 mg/kg) significantly reduced to 9,300 ± 1100, 1,000 ± 850, 1,500 ± 900, 4,300 ± 1000 and 0 mg/kg TPH corresponding to 90.7%, 99%, 98.5%, 95.7% and 100% degradation in soil bio-augmented with bacterial-fungal consortium (metal free), bacterial-fungal consortium + Ni2+, bacteria-fungal consortium + Zn2+, bacterial-fungal consortium + Cd2+, and bacterial-fungal consortium + Ni2+

+ Zn2+ + Cd2+, respectively. The mixture of Ni2+, Zn2+ and Cd2+ of 30 mg/L concentration elicited a complete TPH biodegradation by the bacterial-fungal consortium. This indicated that the bacterial-fungal consortium acted synergistically to pose strong resistance to the synergistic toxicity and inhibitory effect imposed by the presence of the mixed heavy metals. In addition, the results also revealed that the single presence of Ni2+, Zn2+ and Cd2+ and

their mixture exerted a significant positive effect on the TPH biodegradation by bacterial-fungal consortium by increasing the percent TPH degradation by 9.2%, 8.6%, 5.5% and 10.3%, respectively, relative to the percent TPH degradation attained by the bacterial-fungal consortium (metal free). The observed relative enhancement of TPH biodegradation may be due to high relative bioavailability of the individual metals and their mixture as presented in

Figure 4(B) as well as the relative high combined or mixed metal concentration tolerance of the bacterial-fungal consortium (Table 2). It has been reported that Ni2+ supplementation at a concentration of 5 mM (294 mg/L) suppressed significantly (p≤ 0.01) fluorene, phenanthrene and anthracene removal from broth media, while it increased fluoranthene degradation by 4.8 % relative to the control using fungus-bacterial consortium (Bacillus subtilis

and Acremonium sp.) [59]. In addition, Shen et al. [60] have reported that 10 mg/L concentration of cadmium inhibited the biodegradation of phenanthrene by indigenous microorganisms in soil microcosms where the soil pH is

8.18 while on the other hand, 1 mg/L of cadmium inhibited phenanthrene biodegradation by indigenous microorganisms in soil of pH 7.6 [61].

The ranking of the different bacterial-fungal consortium bioaugmentation treatments with respect to TPH reduction or biodegradation was B-T12 < B-T15 < B-T14 < B-T13 < B-T16. The ranking revealed that in all the soil microcosms bio-augmented with bacterial-fungal consortium, the

highest percent TPH biodegradation was attained in soil microcosm containing the presence of the mixed heavy metals (Ni2+ + Zn2+ + Cd2+) (i.e. 100%). This is closely followed by bacterial-fungal consortium + Ni2+ (99%), bacterial-fungal consortium + Zn2+ (98.5%), bacterial-fungal consortium + Cd2+ (95.7%) and bacterial-fungal consortium-metal free (90.7%), respectively. Furthermore, in comparison with bacterial and fungal consortia, the

bacterial-fungal consortium in the absence and presence of heavy metals achieved a higher percent TPH biodegradation than that due to bacterial and fungal consortia, respectively. This is respectively followed by that due to bacterial consortium and fungal consortium in the absence and presence of heavy metals. This observation suggest that both bacteria and fungi species play very significant role in TPH degradation or removal, and that the

presence of both types of microorganisms may result in synergy of degradation activity during bioaugumentation [15]. Nevertheless, the bacterial consortium, fungal consortium and bacterial-fungal consortium have demonstrated their ability to degrade TPH with high degradation efficiency even in the presence of heavy metals in relation to the non-bioaugmented soil microcosm (control or natural bioattenuation).

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Figure 5: (A) Growth profile of bacterial consortium, fungal consortium and bacterial-fungal consortium in the absence of heavy metals (Nickel,

Zinc and Cadmium). (B) Growth profile of bacterial consortium in the absence and presence of heavy metals (Nickel, Zinc and Cadmium). (C)

Growth profile of fungal consortium in the absence and presence of heavy metals (Nickel, Zinc and Cadmium). (D) Growth profil e of bacterial-

fungal consortium in the absence and presence of heavy metals (Nickel, Zinc and Cadmium).

3.4 Microbial Growth Profile Figure 5 shows the profiles of microbial growth in the non-augmented soil microcosm and soil microcosms augmented with bacterial and fungal consortia in the presence and absence of Ni2+, Zn2+ and Cd2+ heavy metals. The profiles of bacterial growth in the non-augmented soil microcosm and soil microcosms augmented with bacterial consortium-metal free, fungal consortium-metal free and bacterial-fungal consortium-metal free as shown in Fig.

5(A) indicates that the bacterial counts generally increased from day 0 to day 21 in each of the augmented and non-augmented soil microcosms. For soil microcosm bio-augmented with bacterial consortium-metal free, fungal consortium-metal free and bacterial-fungal consortium-metal free, the bacterial counts increased from 4.8 to 12.8 × 106cfu-g−1, 4.3 to 7.8 × 106cfu-g−1 and 3.2 to 16.9 × 106cfu-

g−1 which corresponded to a percentage increase of 167%, 81.4% and 423%, respectively. On the other hand, the bacterial count and the fungal count in the non-augmented soil microcosm correspondingly increased from 8.1 to 8.9 × 104cfu-g−1 and from 2.8 to 3.5 × 104cfu-g−1corresponding to a percentage increase of 10 and 25%, respectively. This low percentage increase in both bacterial and fungal growth showed that the non-augmented soil microcosm had more of non-hydrocarbon utilizing microbes than the

hydrocarbon utilizing bacteria and fungi when compared to the soil microcosm bio-augmented with bacterial and fungal consortia that are hydrocarbon utilizers. This accounted for the low TPH biodegradation achieved in the non-augmented soil microcosm (natural bioattenuation).

Figure 5(B) shows the profiles of bacterial growth in the soil microcosms augmented with consortia of bacteria,

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Bacteria Count in Control

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190

Ni2+, Zn2+, Cd2+ and the mixture of Ni2+, Zn2+ and Cd2+, respectively. As shown in Figure 5, it is seen that the bacterial counts generally increased from day 0 to day 21 in each of the augmented soil microcosms. For soil microcosm bio-augmented with bacterial consortium-metal

free, bacterial consortium + Ni2+, bacterial consortium + Zn2+, bacterial consortium + Cd2+, and bacterial consortium+ Ni2+ + Zn2+ + Cd2+, the bacterial counts increased from 4.8 to 12.8 × 106cfu-g−1, 3.2 to 16.9 × 106cfu-g−1, 2.2 to 11.5 × 106cfu-g−1, 2.1 to 9.8 × 106cfu-g−1, and 4.2 to 10.6 × 106cfu-g−1 which corresponded to a percentage increase of 167%, 428%, 423%, 367% and 152%, respectively. This results revealed that soil

microcosm bio-augmented with bacterial consortia in the presence of Zn2+ heavy metal exhibited the highest bacterial growth. This is closely followed by soil microcosms’ bio-augmented with bacterial consortium + Ni2+, bacterial consortium + Cd2+, bacterial consortium-metal free and bacterial consortium + Ni2+ + Zn2+ + Cd2+, respectively. This observation suggests that the 10 mg/L concentration of the single heavy metals (Ni2+, Zn2+ and Cd2+) used in this

study had enhancement effect on the growth of the bacteria population with Zn2+ displaying the most growth enhancement while the combined mixture effect of the three heavy metals suppressed the growth of the bacteria population. The results also suggest that the combined mixture of heavy metals acts in synergy to impose more toxic effect to bacterial growth than the single heavy metal. A similar observation of growth suppression for Bacillus

sp. CPB4 in the presence of combined mixture of lead (Pb), copper (Cu), zinc (Zn) and cadmium (Cd) has been reported [62]. The suppression or inhibition of the bacterial growth exhibited by the combined mixture of Zn2+, Ni2+ and Cd2+ may be due to the fact that the bioavailable mixed heavy metals of 30 mg/L concentration might probably be high for the bacterial consortium to exert a synergistic toxic effect. It has been documented in the literature that all heavy metals exhibit toxicity to living microbial cells at

certain concentration [35, 63]. This could have been responsible for the lower TPH biodegradation observed in the soil microcosm containing combined mixture of the heavy metals (Ni + Zn + Cd) bio-augmented with bacterial consortium.

Figure 5(C) shows the growth profiles of fungi in the soil microcosms augmented with fungal consortium-metal free, fungal consortium + Ni2+, fungal consortium + Zn2+,

fungal consortium + Cd2+ and fungal consortium + Ni2+ + Zn2+ + Cd2+, respectively. Generally, it is seen that the fungal counts generally increased from day 0 to day 21 in each of the augmented soil microcosms. The fungal counts increased from 4.3 to 7.8 × 106cfu-g−1, 1.9 to 6.4 × 106cfu-g−1, 4.4 to 9.8 × 106cfu-g−1, 3.9 to 10.7 × 106cfu-g−1, and 3.8 to 6.5 × 106cfu-g−1, and which corresponded to a percentage increase of 81.4%, 237%, 123%, 174% and

71.1%, for soil microcosm bio-augmented with fungal consortium-metal free, fungal consortium + Ni2+, fungal consortium + Zn2+, fungal consortium + Cd2+ and fungal consortium + Ni2+ + Zn2+ + Cd2+, respectively. This results indicated that bio-augmented soil microcosm (B-T8) containing fungal consortium + Ni2+ displayed the highest fungal growth. This is relatively followed by soil

microcosms’ bio-augmented with fungal consortium + Cd2+ (B-T10), fungal consortium + Zn2+ (B-T9), fungal consortium-metal free (B-T7) and fungal consortium + Ni2+

+ Zn2+ + Cd2+ (B-T11), respectively. This observation suggests that the 10 mg/L concentration of each Ni2+, Zn2+

and Cd2+ enhanced the growth of the fungal population while the mixed heavy metals (Ni2+ + Zn2+ + Cd2+) of 30 mg/L total concentration suppressed the growth. The enhanced growth of the fungal consortium may be due to the synergistic interaction of the fungal species in the consortium to positively tolerate 10 mg/L concentration each of Ni2+, Zn2+ and Cd2+ respectively. While the fungal consortium growth suppression due to the presence of

combined mixture of heavy metals (Ni2+ + Zn2+ + Cd2+) may probably be due to the concentration (30 mg/L) being too high to act synergistically to impose a greater inhibitory response to the growth of the fungal consortium.

Figure 5(D) shows the growth profiles of bacteria and fungi in the soil microcosms augmented with bacterial-fungal consortium-metal free, bacterial-fungal consortium + Ni2+, bacterial-fungal consortium + Zn2+, bacterial-fungal

consortium + Cd2+ and bacterial-fungal consortium + Ni2+ + Zn2+ + Cd2+, respectively. Generally, it is seen that the bacterial-fungal counts generally increased from day 0 to day 21 in each of the augmented soil microcosms. The bacterial-fungal counts increased from 7.6 to 17.2 × 106cfu-g−1, 4.3 to 19.4 × 106cfu-g−1, 5.5 to 17.8 × 106cfu-g−1, 5.0 to 15.8 × 106cfu-g−1, and 6.7 to 30.7.3 × 106cfu-g−1, and which corresponded to a percentage increase of 126%,

351%, 224%, 216% and 358%, for soil microcosm bio-augmented with bacterial-fungal consortium-metal free, bacterial-fungal consortium + Ni2+, bacterial-fungal consortium + Zn2+, bacterial-fungal consortium + Cd2+, and bacterial-fungal consortium + Ni2+ + Zn2+ + Cd2+, respectively. This results indicated that bio-augmented soil microcosm (B-T16) containing bacterial-fungal consortium + Ni2+ + Zn2+ + Cd2+ displayed the highest bacterial-fungal growth. This is closely followed by soil microcosms’ bio-

augmented with bacterial-fungal consortium + Ni2+ (B-T13), bacterial-fungal consortium + Zn2+ (B-T14), bacterial-fungal consortium + Cd2+ (B-T15) and bacterial-fungal consortium-metal free (B-T12), respectively. This observation suggests that the 10 mg/L concentration of each Ni2+, Zn2+ and Cd2+ as well as the 30 mg/L of the mixed heavy metals enhanced the growth of the bacterial-fungal population. The enhanced growth of the bacterial-fungal consortium may be

due to the synergistic interaction of the bacterial-fungal species in the consortium to positively tolerate 10 mg/L concentration each of Ni2+, Zn2+ and Cd2+ and the 30 mg/L of the mixed heavy metals, respectively.

3.5 pH profile during TPH biodegradation The un-impacted soil pH before crude oil

contamination was 5.1. However, after crude oil

contamination, the soil pH reduced to 4.5. This observed reduction in soil pH as a result of crude oil contamination is in agreement with the findings of Osuji and Nwoye [64]. The reduction in soil pH implies increased soil acidity which is a problem for agricultural soils.


191

Figure 6. (A) Soil pH profile in the course of TPH biodegradation by bacterial consortium, fungal consortium and bacterial-fungal consortium in

the absence of heavy metals (Nickel, Zinc and Cadmium). (B) Soil pH profile in the course of TPH biodegradation by bacterial consortium in the

absence and presence of heavy metals (Nickel, Zinc and Cadmium). (C) Soil pH profile in the course of TPH biodegradation by fungal

consortium in the absence and presence of heavy metals (Nickel, Zinc and Cadmium). (D) Soil pH profile in the course of TPH biodegradation

by bacterial-fungal consortium in the absence and presence of heavy metals (Nickel, Zinc and Cadmium).

The resulting increased acidity could be due to the fact that hydrocarbons contain many free cations causing them

to have properties of a weak acid. Figure 6 shows the soil pH profile in the course of TPH biodegradation in the absence and presence of heavy metals and microbial consortium. The results in Figure 6 as depicted for microbial consortium in the absence of heavy metals (Figure 6(A)), bacterial consortium in the presence of heavy metals (Figure 6(B)), fungal consortium in the presence of heavy metals (Figure 6(C)) and bacterial-fungal

consortium in the presence of heavy metals (Figure 6(D)) revealed that the pH of the crude oil contaminated soil microcosms increased in the course of the TPH biodegradation. The observed changes in the soil pH values of the various soil microcosms were due to the removal or degradation of the crude oil contaminant. The range of soil

pH values (6.6 – 7.8) obtained for the various bioaugmented and non-bioaugmented soil microcosms

were within the range for optimum microbial activities.

3.6 Kinetics and half-life of TPH biodegradation First-order kinetics model equation (Eq. 2) fitted to the

biodegradation data was used to determine the rate of TPH biodegradation (i.e. biodegradation rate constants (k)) in the various bioaugmentation treatments. The results are presented in Table 3. The results in Table 3 as revealed by

high correlation coefficient (R2) indicated that the TPH biodegradation data fitted well to the first-order kinetic model. The half-life time (t1/2) of TPH biodegradation in the absence and presence of heavy metals was calculated using Eq. (3), and the results are given in Table 3.

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Control (Natural Bioattenuation)


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Table 3: Biodegradation rate constants and correlation coefficients obtained from the fitting of first-order kinetic model to TPH biodegradation data and the calculated half-life time

Soil Treatment k (day-1) 2/1t (days)

2R

B-T1 (Control) 0.010 69.3 0.9553

Bacterial Consortium (B-T2) 0.043 16.1 0.9952 Bacterial Consortium + Nickel (B-T3) 0.049 14.1 0.9981 Bacterial Consortium + Zinc (B-T4) 0.051 13.6 0.9994 Bacterial Consortium + Cadmium (B-T5) 0.046 15.1 0.9974 Bacterial Consortium + Nickel + Zinc + Cadmium (B-T6) 0.040 17.3 0.9942 Fungal Consortium (B-T7) 0.035 19.8 0.9930 Fungal Consortium + Nickel (B-T8) 0.040 17.3 0.9933 Fungal Consortium + Zinc (B-T9) 0.037 18.7 0.9956

Fungal Consortium + Cadmium (B-T10) 0.039 17.8 0.9946 Fungal Consortium + Nickel + Zinc + Cadmium (B-T11) 0.032 21.7 0.9894 Bacterial- Fungal Consortium (B-T12) 0.060 11.6 0.9966 Bacterial-Fungal Consortium + Nickel (B-T13) 0.129 5.4 0.9977 Bacterial-Fungal Consortium + Zinc (B-T14) 0.118 5.9 0.9949 Bacterial-Fungal Consortium + Cadmium (B-T15) 0.084 8.3 0.9972 Bacterial-Fungal Consortium + Ni + Zn + Cd (B-T16) 0.158 4.4 0.9992

It is important to note that a higher biodegradation rate constant indicates a faster or higher rate of biodegradation and subsequently a lower half-life time. It could be seen from Table 3 that for soil microcosms augmented with microorganisms in the absence of heavy metals (B-T2, B-T7 and B-T12) and non-augmented soil microcosm, the soil microcosm augmented with bacterial-fungal consortium had a higher k (0.060 day-1) and lower t1/2 (11.6 days) than

that augmented with bacterial consortium (k= 0.043 day-1 and t1/2= 16.1 days) and fungal consortium (k= 0.035 day-1 and t1/2= 19.8 days), respectively. While the non-augmented soil microcosm had the least k value of 0.010 day-1 and the highest t1/2 value of 69.3 days. The value obtained for the non-bioaugmented soil microcosm is close to the k value of 0.015 day-1 obtained for the natural bioattenuation of soil microcosm contaminated with lubricating motor oil [13].

For soil microcosms augmented with bacterial

consortium in the presence of heavy metals, the soil augmented with bacterial consortium + Zn2+ had a relatively higher k (0.051 day-1) and lower t1/2

(13.6 days)

than the one augmented with bacterial consortium + Ni2+ (k = 0.049 day-1 and t1/2 = 14.1 days), bacterial consortium + Cd2+ (k = 0.046 day-1 and t1/2 = 15.1 days), bacterial consortium (metal free) (k = 0.043 day-1 and t1/2= 16.1 days) and bacterial consortium + Ni2+ + Zn2+ + Cd2+ (k=

0.040 day-1 and t1/2= 17.3 days), respectively. Furthermore, for soil microcosms augmented with fungal consortium in the presence of heavy metals, the soil augmented with fungal consortium + Ni2+ had a relatively higher k (0.040 day-1) and lower t1/2

(17.3 days) than the one augmented

with fungal consortium + Cd2+ ( k = 0.039 day-1 and t1/2=

17.8 days), fungal consortium + Zn2+ (k = 0.037 day-1 and t1/2 = 18.7 days), fungal consortium (metal free) (k = 0.035 day-1 and t1/2 = 19.8 days) and fungal consortium + Ni2+ + Zn2+ + Cd2+ (k = 0.032 day-1 and t1/2 = 21.7 days), respectively.

Finally, for soil microcosms augmented with bacterial-fungal consortium in the presence of heavy metals, the soil augmented with bacterial-fungal consortium + Ni2+ + Zn2+

+ Cd2+ had a higher k (0.158 day-1) and lower t1/2 (4.4 days)

than that augmented with bacterial-fungal consortium + Ni2+ (k = 0.129 day-1 and t1/2 = 5.4 days), bacterial-fungal consortium + Zn2+ (k = 0.118 day-1 and t1/2 = 5.9 days), bacterial-fungal consortium + Cd2+ (k = 0.084 day-1 and t1/2

= 8.3 days) and bacterial-fungal consortium (metal free) (k = 0.060 day-1 and t1/2 = 11.6 days), respectively.

4 Conclusions From this study, it can be concluded that the bacteria species (Pseudomonas aeruginosa, Bacillus subtilis and Micrococcus letus) and fungi species (Aspergillus niger,

Aspergillus carmari and Penicillium notatum) used as consortia in this study showed high tolerance and resistance to nickel, zinc and cadmium heavy metals in soil. The addition of hydrocarbon utilizing and metal-resistant microorganisms in the form of consortia either as bacterial consortia (Pseudomonas aeruginosa, Bacillus subtilis and Micrococcus letus), fungal consortia (Aspergillus niger, Aspergillus carmari and Penicillium notatum) or bacterial-

fungal consortia (Pseudomonas aeruginosa, Bacillus subtilis, Micrococcus letus, Aspergillus niger, Aspergillus carmari and Penicillium notatum) provided an enhanced TPH biodegradation (>60%) in the presence and absence of single or individual heavy metals (nickel, zinc and cadmium) with the bacterial-fungal consortia providing the highest enhancement. Complete TPH biodegradation (100%) and enhanced microbial growth in soil can be

achieved in the presence of mixed heavy metals(Nickel + Zinc + Cadmium) using bacterial-fungal consortia while a reduction in TPH biodegradation and decreased microbial growth is attained with the use of bacterial or fungal consortia. These features of a bacterial-fungal consortium such as mixed-heavy metal resistance and high TPH removal ability make it a very attractive candidate for TPH biodegradation in co-contaminated soil environment. The

rate of TPH biodegradation in the presence and absence of heavy metals and inoculated microbial consortia can be described by biodegradation rate constant obtained from the


193

application of first order kinetics. The rate constant ( k )

ranges between 0.035 day-1 and 0.060 day-1 for soil

microcosm bioaugmented with bacterial, fungal and bacterial-fungal consortia, respectively in the absence of heavy metals and 0.010 day-1 for non-bioaugmented soil microcosm (natural bioattenuation). A half-life time (t1/2) of 69.3 days was attained for TPH biodegradation in non-bioaugmented soil microcosm. This t1/2 was reduced to between 16.1, 19.8 and 11.6 days in soil microcosm bioaugmented with bacterial consortia, fungal consortia and bacterial-fungal consortia, respectively. Similarly, the rate

constant (k) ranges between 0.032 day-1 and 0.158 day-1 for soil microcosm bioaugmented with bacterial, fungal and bacterial-fungal consortia, respectively in the presence of single and mixed heavy metals Therefore, the inoculation of bacterial consortia, fungal consortia and bacterial-fungal consortia that possesses heavy metal resistance and petroleum hydrocarbons degradation ability is a promising bioaugmentation strategy to enhance in-situ and ex-situ soil

bioremediation of soil contaminated with both petroleum hydrocarbons and heavy metals.

Aknowledgment The authors wish to thank the staff of Thermosteel

Laboratories, Warri, Delta State of Nigeria for providing the facilities used for the soil physical, chemical and microbial analyses.

Ethical issue Authors are aware of, and have complied with the best

practice in publication ethics specifically with regard to authorship, dual submission, and manipulation of figures, competing interests and compliance with policies on research ethics. Authors have adhered to publication requirements that this submitted work is original and has not been published elsewhere in any form of language.

Competing interests The authors wish to declare that there is no conflict of

interest in this research work.

Authors’ contribution All the authors of this study have completely contributed to the data collection, data analyses and manuscript writing.

References 1. Nigerian National Petroleum Corporation (NNPC)

(2016). Oil Production. http://www.nnpcgroup.com/nnpcbusiness/upstreamventures/oilproduction.aspx.

2. Koshlaf, E., and Ball, A. S. (2017). Soil bioremediation approaches for petroleum hydrocarbon polluted environments. AIMS Microbiology 3(1), 25-49.

3. Nduka, J. K. C., Constance, E., and Obiakor, E. (2006). Selective bioaccumulation of metals by different parts

of some fish species from crude oil polluted water, Bull. Environ. Contam. Toxicol. 77, 846-853.

4. Das, N., and Chandran, P. (2011). Microbial degradation of petroleum hydrocarbon contaminants: An overview. Research Int. 2011, Article ID 941810, 13 pages. Doi:10.4061/2011/941810.

5. Yasin, G., Bhanger, M. I., Ansari, T. M. et al. (2013).

Quality and chemistry of crude oils. J. Pet. Technol. Altern. Fuels 4, 53–63.

6. Ogbo, E. M., and Okhuoya, J. A. (2011). Bioavailability of some heavy metals in crude oil contaminated soils remediated with Pleurotus tuber-regium Fr. Singer. Asian J. Biological Sci. 4, 53-61

7. Shtangeeva, I. (2006). Phytoremediation of trace element contaminated soil with cereal crops: role of

fertilizers and bacteria on bioavailability. In: Trace Elements in the Environment. Biogeochemistry, Biotechnology and Bioremediation, Prasad, M.N.V., K.S. Sajwan and R. Naidu (Eds.). CRC Press, Taylor and Francis Group, Boca Raton, pp: 549-581.

8. Samanta, S. K., Singh, O. V., and Jain, R. K. (2002). Polycyclic aromatic hydrocarbons: environmental pollution and bioremediation. Trends Biotechnol. 20,

243–248. 9. Said, W. A., and D. A. Lewis. (1991). Quantitative

assessment of the effects of metals on microbial degradation of organic chemicals. Appl. Environ. Microbiol. 57, 1498–150

10. Ojo, O. A. (2006). Petroleum hydrocarbon utilization by native bacterial population from a wastewater canal in Southwest Nigeria. Afr. J. Biotechnol. 5(4), 333–337.

11. Ameen, F., Moslem, M., Hadi, S., et al. (2016). Biodegradation of diesel fuel hydrocarbons by mangrove fungi from Red Sea coast of Saudi Arabia. Saudi. J. Biol. Sci. 23(2), 211–218.

12. Zhang, J. H., Quan-Hong, X., Hui, G., et al. (2016). Degradation of crude oil by fungal enzyme preparations from Aspergillus spp. for potential use in enhanced oil recovery. J. Chem. Technol. Biotechnol. 91, 865–875.

13. Agarry, S. E., Aremu, M. O., and Aworanti, O. A.

(2013). Kinetic modelling and half-life study on bioremediation of soil co-contaminated with lubricating motor oil and lead using different bioremediation strategies. Soil and Sediment Contam.- An Int. J. 22(7), 800 – 816.

14. Iordache, M., and Borza, I. (2012). The bioremediation potential of earthworms (Oligochaeta: Lumbricidae) in a soil polluted with heavy metals. J. Food Agric.

Environ. 10(2), 1183–1186. 15. Ma, X-K., Ding. N., and Peterson, E. C. (2015).

Bioaugmentation of soil contaminated with high-level crude oil through inoculation with mixed cultures including Acremonium sp. Biodegradation 26 (3), 259-269.

16. Patowary, K., Patowary, R., Kalita, M. C., and Deka, S. (2016). Development of an efficient bacterial

consortium for the potential remediation of hydrocarbons from contaminated sites. Front. Microbiol. 7, 1092. doi: 10.3389/fmicb.2016.01092

17. Mrozik, A., and Piotrowska-Seget, Z. (2010). Bioaugmentation as a strategy for cleaning up of soils contaminated with aromatic compounds. Microbiological Res. 165, 363-375.


194

18. Cycoń, M., Mrozik, A., and Piotrowska-Seget, Z. (2017). Bioaugmentation as a strategy for the remediation of pesticide-polluted soil: A review. Chemosphere 172, 52-71.

19. Roane, T. M., Josephson, K. L., and Pepper, I. L.

(2001). Dual-bio augmentation strategy to enhance remediation of co-contaminated soil. Appl. Environ. Microbiol. 67, 3208–3215.

20. Alisi, C., Musella, R., Tasso, F., Ubaldi, C., Manzo, S., Cremisini, C. and Sprocati, A.R. (2009). Bioremediation of diesel oil in a co-contaminated soil by bioaugumentation with a microbial formula tailored with native strains selected for heavy metals resistance.

Science of the Total Environment 407, 3024-3032. 21. Shukla, K. P., Singh, N. K., and Sharma, S. (2010).

Bioremediation: developments, current practices and perspectives. Genet. Eng. Biotechnol. J. 2010, 1–19.

22. Das, M., and Adholeya, A. (2011). Role of microorganisms in remediation of contaminated soil T. In: Satyanarayana et al. (eds.), Microorganisms in Environmental Management: Microbes and

Environment, pp81-111 23. Thavamani, 1. P., Megharaj, M., and Naidu, R. (2012a).

Bioremediation of high molecular weight polyaromatic hydrocarbons co-contaminated with metals in liquid and soil slurries by metal tolerant PAHs degrading bacterial consortium. Biodegradation 23, 823-35.

24. Thavamani, P., Malik, S., Beer, M., Megharaj, M., and Naidu, R. (2012b). Microbial activity and diversity in

long-term mixed contaminated soils with respect to polyaromatic hydrocarbons and heavy metals. J. Environ. Manage. 99, 10–17.

25. Wu, F., Yang, W., Zhang, J. And Zhou, L. (2010). Cadmium accumulation and growth responses of a poplar (Populus deltoids×Populus nigra) in cadmium contaminated purple soil and alluvial soil. Journal of Hazardous Materials, 177 (1-3), 268-273.

26. Agarry, S. E., Aremu, M. O., and Aworanti, O. A.

(2014). Biostimulation and phytoremediation treatment strategies of gasoline-nickel co-contaminated soil. Soil and Sediment Contam.: An Int. J. 23 (3), 227-244.

27. Tesfalem, B W. (2017). Isolation, Screening and Identification of Cadmium Tolerant Fungi and Their Removal Potential. J. Forensic Sci. & Criminal Inves. 5(1), 555-656.

28. Oladipo, O. G., Awotoye, O. O., Olayinka, A.,

Bezuidenhout, C. C., and Maboeta, M. S. (2018). Heavy metal tolerance traits of filamentous fungi isolated from gold and gemstone mining sites. Brazilian Journal of Microbiology 49, 29–37.

29. Owabor, C. N., Onwuemene, O. C., and Enaburekhan, I. (2011). Bioremediation of polycyclic aromatic hydrocarbon contaminated aqueous-soil matrix: effect of co-contamination. J. Appl. Sci. Environ. Manage. 15

(4), 583 – 588. 30. Thavamani, P., Megharaj, M., Venkateswarlu, K., and

Naidu, R. (2013). Mixed contamination of polyaromatic hydrocarbons and metals at manufactured gas plant sites: toxicity and implications to bioremediation. In: Environmental Contamination:

Health Risks, Bioavailability and Bioremedation, ed: Wong MH, Taylor and Francis, pp: 347–368

31. Ma, X. K., Ding, N., Peterson, E. C., and Daugulis, A. J. (2016). Heavy metals species affect fungal-bacterial synergism during the bioremediation of fluoranthene.

Appl. Microbiol. Biotechnol. 100, 7741-50. 32. Wang, C., Gu, L., Ge, S., Liu, X., Zhang, X., and Chen,

X. (2018). Remediation potential of immobilized bacterial consortium with biochar as carrier in pyrene-Cr (VI) co-contaminated soil. Environmental Technology, DOI: 10.1080/09593330.2018.1441328

33. Kim, Y. M., Ahn, C. K., Woo, S. H., Jung, G. Y., and Park, J. M. (2009). Synergic degradation of

phenanthrene by consortia of newly isolated bacterial strains. J. Biotechnol. 144, 293-298.

34. Jackson, M. H. (1967). Soil chemical analysis. Prentice-Hall of India (Pvt.Ltd.), New Delhi, India.

35. Mustafa, S., Al-Douseri, A., Majki, K., and Al-Saleh, E. (2013). Potential of crude oil-degrading bacteria to co-resist heavy metals in soil. WIT Transactions on Ecology and the Environment, 173 (VI), 697-705.

36. Krieg, N. R., Holt, J. G., Sneath, P. H.A., Stanley, J. T., and Williams, S. T. (1994). Bergey’s Manual of Determinative Bacteriology, 9th ed., Williams and Wilkins, Baltimore.

37. Saroj, A., and Keerti, D. (2013). Isolation and characterization of hydrocarbon degrading microorganisms from petroleum oil contaminated soil sites. Bull. Environ. Sci. Res. 2 (4), 5-10.

38. Ahmed, R. Z., Ahmed, N. and Gadd, G. M. (2010). Isolation of two Kocuria species capable of growing on various polycyclic aromatic hydrocarbons. Afr. J. Biotechnol. 9, 3611-3617.

39. Giwa, O. E., and Ibitoye, F. O. (2017). Bioremediation of heavy metals in crude oil contaminated soil using isolated indigenous microorganism cultured with E coli DE3 BL21. Int. J. Eng. Appl. Sci. 4 (6), 67-70.

40. Richa, S., Subhash, C., and Amrita, S. (2013). Isolation

of microorganism from soil contaminated with degraded paper in Jharna village. J. Soil Sci. Environ. Manage. 4 (2), 23-27.

41. Samson, R. A., Hoekstra, E. S., van Oorschot C. A. N. (1984). Introduction of Food-Borne Fungi. Amsterdam, The Netherlands: Institute ofthe Royal Netherlands Academy of Arts Science; pp 248.

42. Mackie, L. and McCartney, M. (1999). Practical

medical Microbiology, 14th Ed.Harcourt brace and Company limited.

43. Obahiagbon, K.O. and Owabor, C.N. (2008). Biotreatment of crude oil polluted water using mixed microbial populations of P.aureginosa, Pennicillium notatum, E.coli and Aspergillus niger. Adv. Mater. Res. 4, 802-807.

44. EL-Hanafy, A-M., Anwar, Y., Sabir, J. S. M.,

Mohamed, S. A., Al-Garni, S. S. M., Abu Zinadah, O. A. H. and Ahmed, M. M. (2017). Characterization of native fungi responsible for degrading crude oil from the coastal area of Yanbu, Saudi Arabia. Biotechnol. Biotechnological Equipment 31 (1), 105–111.

45. Carrillo, G. R, and Gonzalez, C. M. C. (2012). Tolerance to and accumulation of cadmium by the


195

mycelium of the fungi Scleroderma citrinum and Pisolithus tinctorius. Biol. Trace Elem. Res. 146, 388-395.

46. Osuji, L. C., Egbuson, E. J. G., and Ojinnaka, C. M. (2005). Chemical reclamation of crude-oil-inundated

soils from Niger Delta, Nigeria. Chem. Ecol. 21(1), 1-10.

47. Zahed, M. A., Abdul Aziz, H., Isa, M. H., Mohajeri, L., Mohajeri, S., and Kutty, S. R. M. (2011). Kinetic modeling and half-life study on bioremediation of crude oil dispersed by Corexit 9500. J. Hazard Mater. 185, 1027–1031.

48. Rasool, A., and Irum, S. 2014. Toxic Metal Effect on

Filamentous Fungi Isolated from the Contaminated Soil of Multan and Gujranwala. J. Bioresource Manage. 1 (2), 38-51.

49. Godheja, J., Shekhar, S. K., Satyanarayan, G. N. V., Singh, S. P., and Modi, D. R. (2017). Antibiotic and heavy metal tolerance of some indigenous bacteria isolated from petroleum contaminated soil sediments with a study of their aromatic hydrocarbon degradation

potential. Int. J. Curr. Microbiol. App. Sci. 6(3), 194-211.

50. Ezzouhri, L., Castro, E., Moya, M., Espinola, F., and Lairini, K. (2009). Heavy metal tolerance of filamentous fungi isolated from polluted sites in Tangier, Morocco. African Journal of Microbiology Research 3 (2), 035-048

51. Chanda, D., Sharma, G. D., Jha, D. K., and Hijri, M.

2017. Tolerance of Microorganisms in Soil Contaminated with Trace Metals: An Overview. In: Shukla P. (eds) Recent advances in Applied Microbiology, pp 165-193.

52. Oaikhena, E. E., Makaije, D. B., Denwe, S. D., Namadi, M. M., and Haroun, A. A. (2016). Bioremediation potentials of heavy metal tolerant bacteria isolated from petroleum refinery effluent. American J. Environ. Protect. 5 (2), 29-34.

53. Nies, D., H. (1999). Microbial heavy-metal resistance. Appl. Microbial Biotechnol. 51, 730–750.

54. Smrithi, A., and Usha, K. (2012). Isolation and characterization of chromium removing bacteria from tannery effluent disposal site. Int. J. Adv. Biotechnol. Res. 3(3), 644-652.

55. Amalesh S., ParamitaBera, MahamudaKhatun, Chandrima, S., Pinaki, P., Asif, L., Anurup, M. (2012).

An investigation on heavy metal tolerance and antibiotic resistance properties of bacterial strain Bacillus sp. Isolated from municipal waste. J. Microbial. Biotech. Res. 2(1), 178 - 189.

56. Anahid, S., Yaghmaei, S., and Ghobadinejad, Z. (2011). Heavy metal tolerance of fungi. Scientia Iranica 18, 5028.

57. Sandrin, T.R., and Hoffman, D. R. (2007).

Bioremediation of organic and metal co-contaminated environments: effects of metal toxicity, speciation, and bioavailability on biodegradation. In: Environmental Bioremediation Technologies, (ed) Singh SN, Tripathi RD, Springer, Berlin. pp. 1–34.

58. Semerci N., and Çeçen, F. (2007). Importance of Cd speciation in nitrification inhibition. J. Hazard Mater. 147, 503–512.

59. Ma, X-K., Li, T-t., Farm, H., Peterson, E, C., Zhao, W-w., Guo, W., and Zhou, B. (2017). The influence of

heavy metals on the bioremediation of polycyclic aromatic hydrocarbons in aquatic system by a bacterial-fungal consortium. Environmental Technology, DOI: 10.1080/09593330.2017.1351492. http://dx.doi.org/10.1080/09593330.2017.1351492

60. Shen, G., Cao, L., Lu, L.Y., and Hong, J. (2006). Combined effect of heavy metals and polycyclic aromatic hydrocarbons on urease activity in soil.

Ecotoxicol. Environ. Saf. 63, 474–480. 61. Maslin, P., and Maier, R. M. (2000). Rhamnolipid-

enhanced mineralization of phenanthrene in organic-metal co-contaminated soils. Bioremediation J. 4, 295–308.

62. Kim, S. U., Cheong, Y. H., Seo, D. C., Hur, J. S., Heo, J. S., and Cho, J. S. (2007). Characterisation of heavy metal tolerance and biosorption capacity of bacterium

strain CPB4 (Bacillus spp.). Water Sci. Technol. 55, 105-11.

63. Sandrin, T. R., and Maier, R. M. (2003). Impact of metals on the biodegradation of organic pollutants. Environ. Health Persp. 111, 1093–1101.

64. Osuji, L., and Nwoye, I. 2007. An appraisal of the impact of petroleum hydrocarbons on soil fertility: the Owaza experience. Afr. J. Agric. Res. 2, 318-324.

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