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Introduction

The existence of petroleum predates man, but the present day petroleum industry was established in 1859 by Colonel E.A. Drake in Romania [1]. The early usage of crude oil was primarily for lighting as it conveniently replaced whale oil which was expensive; nonetheless, today, it is used as fuel and it is at present Nigeria’s and indeed, the world’s leading energy source [2]. Petroleum is the name given to a mix of condensate, natural gas and crude oil. Crude oil is a complex combination of varying molecular weight hydrocarbons comprising hydrogen and carbon in the ratio of 2:1. It also consists of about 3% (v/v) oxygen, nitrogen and sulphur; trace amounts of phosphorus and heavy metals namely nickel and vanadium [1,3-6]. There are 4 groups of hydrocarbons in crude oil viz. the saturated hydrocarbons and the aromatics; the more polar, non hydrocarbon components; the resins and the asphaltenes. Moreover, crude oil can be categorized according to individual distillation residues as naphthenes, paraffins and aromatics; and on the basis of heavy molecular weight components as heavy, medium or light; and also based on the age, location and depth of the oil well as paraffin-based, asphalt-based or mixed based [7,8]. In Nigeria, oil fields litter the Niger-Delta region, having a large network of pipelines which convey crude oil to various refineries including the Kaduna refinery up north. These pipes get vandalized occasionally by hoodlums thus resulting in oil spillage and environmental pollution. Oil spillage is therefore, the discharge of oil into the natural environment with its consequent perils. Bioremediation is the application of micro organisms, plants, or microbial or plant enzymes to decontaminate and reclaim an environment [9]. The concept embodies a host of other processes including biodegradation. Biodegradation is the biologically catalyzed break down in the structural and molecular complexity of compounds into smaller components such as carbondioxide and minerals by enzymatic or metabolic processes in the environment [10]. Nonetheless, in marine, saline and hypersaline environments, biodegradation is an uphill task owing to the harmful impact of salt on micro organisms and as such in order to successfully bioremediate such environments, dilution is done which is costly; therefore, to clean up this type of environment, a distinct bioremediation approach employing halophiles and halotolerant micro organisms capable of biodegrading crude oil in saline or hypersaline conditions becomes necessary [11]. Much of the earth’s surface is occupied by water and a greater percentage of the earth’s hydrosphere is salt water (Oceans and seas), in spite of this, literature on crude oil degradation by halophilic and halotolerant micro organisms especially molds remains meager [12]. Of this paltry amount of literature on crude oil degradation by halophilic micro organisms, those associated with molds are relatively insufficient, regardless of the fact that they are decomposers.

Materials and Methods

Isolation, Characterization and Identification of Halophilic Fungal Species

Fungal Isolation and Characterization

The water samples obtained from the lakes quarterly from January to December were stored in ice chests and transported to the laboratory before being transferred to refrigerators. Exactly 0.1ml of the water samples were transferred into the centre of already prepared agar plates using sterile pipettes. With the aid of a sterile glass spreader, the aliquot was spread evenly on the surface of the agar plate. All fungal media were amended with 0.5mg/ml of Chloramphenicol to inhibit bacterial growth. Plates were incubated at room temperature for 10 days each. Developing fungal isolates were purified by repeated subculture technique and transferred to Bijou bottles with agar slopes for identification and storage. Czapex-Dox Agar (CzA) and SDA prepared with the lake water were used for the isolation of halophilic fungal species.

Identification of Fungal Isolates

Preliminary fungal characterization were done by studying the cultural characteristics and employing the slide culture wet mount technique for evaluating the fungal microscopic features with reference to the Manual of Fungal Atlases according to [13-16]. The identities of 7 most halophilic isolates were confirmed at Macrogen Inc., 10F, 254 Beotkkot-ro, Geumcheon-gu, Seoul, Republic of Korea, using the ITS rDNA Sequence Analyses. Molecular assays were carried out on each sample using nucleic acid as a standard. A proprietary formulation [microLYSIS®-PLUS (MLP), Microzone, UK] was subjected to the rapid heating and cooling of a thermal cycler, to lyse cells and release deoxyribonucleic acid (DNA). Following DNA extraction, Polymerase Chain Reaction (PCR) was employed to amplify copies of the rDNA in vitro. The quality of the PCR product was assessed by undertaking gel electrophoresis. PCR purification step was carried out to remove unutilized dNTPs, primers, polymerase and other PCR mixture compounds and obtain a highly purified DNA template for sequencing. This procedure also allowed concentration of low yield amplicons. Sequencing reactions were undertaken using BigDye® Terminator v3.1 kit from Applied Biosystems (Life Technologies, UK) which utilises fluorescent labelling of the chain terminator ddNTPs, to permit sequencing. Removal of excess unincorporated dye terminators was carried out to ensure a problem-free electrophoresis of fluorescently labelled sequencing reaction products on the capillary array AB 3130 Genetic Analyzer (DS1) DyeEx™ 2.0 (Qiagen, UK). Modules containing pre-hydrated gel-filtration resin were optimized for clean-up of sequencing reactions containing Big Dye® terminators. Dye removal was followed by suspension of the purified products in highly deionized formamide Hi-Di™ (Life Technologies, UK) to prevent rapid sample evaporation and secondary structure formation. Sample was loaded onto the AB 3130 Genetic Analyzer and sequencing undertaken to determine the order of the nucleotide bases, adenine, guanine, cytosine, and thymine in the DNA oligonucleotide. Following sequencing, identifications were undertaken by comparing the sequence obtained with those available from the European Molecular Biology Laboratory (EMBL) database via the European Bioinformatics Institute (EBI). The strains were identified using Inter specific region sequencing analyses [17].

Halotolerance Test of the Isolates

Salt tolerance of the isolates was checked by inoculating the developing cultures in triplicates on CzA amended with salt up to concentrations of 0, 5, 10, 15, 20, 25, 30, 35, 40, 45, and 50 % w/v. Growth was recorded after 7 days incubation in terms of colony diameter. Plates that did not show growth up to 7 days were further incubated till the fifteenth day to check for delayed growth. Thereafter, the colony diameter of the molds in millimetres plotted against percentage salt concentration in the medium was used to determine their salt tolerance [18]. The salt tolerance study was used to classify the isolates as slight halophiles (0.2-0.5 or even 0.85 M salt equivalent to 1-5%), moderate halophiles (0.5-2.5 M or 0.85-3.4 M salt equivalent to 5-20%), borderline extreme halophiles (1.5-4.0 M salt equivalent to 9-23%) and extreme halophile (2.5-5.2 M salt equivalent to 15-30%) according to various classification schemes proposed by [19-23]. The most halophilic isolate was then used for the biodegradation studies.

Growth Tolerance of Aspergillus flavipes to different Percentage of Crude Oil.

The Bonny light crude oil used for this study was obtained from from Warri Refinery and Petrochemicals. Three set of 100ml conical flasks were prepared containing 49 ml of 2%, 4% and 6% crude oil in Bushnell-Haas broth. Exactly 1 ml of the 10-2 dilution tube of the 24 hours Sabourand dextrose broth culture of Aspergillus flavipes was used to seed each flask, incubated at room temperature and the developing colonies counted after 48 hours. Upon completion of the 48 hours of incubation period, 0.1 ml aliquots of the 2%, 4%, 6% of crude oil amended Bushnell-Haas broth culture were plated out on Bushnell-Haas agar and incubated at room temperature for 48 hours and the total fungal count obtained.

Preliminary Hydrocarbon Degradation Studies

Modified method of [12] was employed for this study. The concentration of crude oil that gave the best growth rate on agar plate (Bushnell-Haas amended with 2% crude oil) was used for this study. Four Erlen-meyer flasks (100ml) containing 50 ml Bushnell-Haas broth amended with 2% crude oil, were inoculated with 1ml of 24 hours broth culture of Aspergillus flavipes and incubated in a rotary shaker for two months. Biweekly, one flask was withdrawn and the crude oil degradation rates evaluated gravimetrically using a separating funnel and also chromatograpically by GC-FID technique.

Evaluation of Hydrocarbon Degradation Rate by Gravimetry

The hydrocarbon degradation potential of the mold was evaluated using the modified gravimetric analysis method according to [25]. Exactly 5 ml of n-hexane was added to the fermentation flask containing the crude oil degradation set up and the contents transferred to a separating funnel. Extraction was done thrice to ensure thorough oil recovery. The extract was however mixed with 0.4g of anhydrous sodium sulphate to remove moisture and carefully transferred into a beaker leaving the sodium sulphate behind. This was evaporated to dryness by heating in a rotary evaporator. The amount of residual oil was measured after extraction of oil from the medium and evaporating it to dryness.

The crude oil degradation equation was derived thus:

W₄ = W₃ ― W₁

W₅ = W₂ ― W₁

Where:

W1: Weight of empty beaker

W2: Weight of beaker + crude oil before degradation

W3: weight of beaker + recovered crude oil after degradation

W4: Residual crude oil

W5: Original weight of crude oil before degradation

Preparation of Samples for GC Analysis TPH

Florisil Clean Up

Standard methods of [27] was used for this study. Florisil was heated in an oven at 130 0C overnight (ca.15h) and transferred to a 250ml size beaker and placed in a desicator. Then, 0.5g anhydrous NaSO4 was added to 1.0g of activated flosiril (florisil charged with magnesium silicate) with mesh size of 60–100nm on an 8ml column plugged with glass wool. Packed column was filled with 5ml n–hexane for conditioning and the stopcock opened to allow n–hexane run out until it just reaches the top of the sodium sulphate into a receiving vessel whilst tapping gently the top of the column till the florisil settled well in the column. Extract was transferred into the column with disposable Pasteur pipette from an evaporating flask. Each evaporating flask was rinsed twice with 1ml portions of n–hexane to dislodge any residual sample and added to column. Eluent was collected into an evaporating flask and evaporated to dryness in a rotary evaporator. Dry eluent was dissolved in 1ml acetone for Chromatographic analysis using Buck Gas Chromatograph Model No: 910 equipped with an HP 88 capillary on-column (100ml x 0.25μm film thickness,) CA, USA, automatic injector, Flame Ionisation Detector (Detector Temperature A: 250 ⁰C, Injector temperature: 22 ⁰C), Integrator chart speed: 2cm/min, Oven temperature: 180 ⁰C.

Results

Halotolerance Test of the Isolates

The isolates recovered in descending order of salt tolerance were Aspergillus flavipes (13mm at 40%), Penicillium citrinum (10mm at 40%), Aspergillus ochraceus (9mm at 40%), Aspergillus nomius (15mm at 35%), Microsphaeropsis arundinis (12mm at 35%), Aspergillus sydowi (28mm at 30%), Penicillium janthinellum(26mm at 30%), Mucor sp (13mm at 30%), Aureobasidium sp (12mm at 30%), Trichoderma sp (9mm at 30%), Alternaria sp. (22mm at 25%), Aspergillus sp (18mm at 25%), Penicillium sp (20mm at 20%), Cladosporium sp. (7mm at 15%) as seen in figures 1 to 14. From the above conclusions, it can be inferred that Aspergillus flavipes, Penicillium citrinum, Aspergillus ochraceus, Aspergillus nomius, Microsphaeropsis arundinis, Aspergillus sydowi, Penicillium janthinellum, Mucor sp, Aureobasidium sp, Trichoderma sp, Alternaria sp, and Aspergillus sp belonged to the extreme halophiles class, whereas, Penicillium sp and Cladosporium sp belonged to the borderline extreme halophiles and moderate halophiles respectively. No slight halophile was isolated in this work as seen in Table 1.

Growth Tolerance of Aspergillus flavipes to different Percentages of Crude Oil

The growth tolerance of Aspergillus flavipes on different percentage concentrations of crude oil (2%, 4% and 6%) in Bushnell-Haas broth was studied. There was insignificant growth on 4% and 6% crude oil broth as shown by total fungal count. Only 2% crude oil broth had significant confluent growth with a fungal count of 2.56x105 CFU/ml. thus the biodegradation study was performed using 2% crude oil in broth, as shown in Table 3.

Bi-weekly Degradation Rate of of crude oil by Aspergillus flavipes as shown by Gravimetry

The crude oil degradation rate of Aspergillus flavipes was evaluated by gravimetric method bi-weekly. After the first two weeks, it was observed that the crude oil had been degraded by 36%, by 4 weeks, it was 67%; by 6 weeks, it was 89%; and upon completion of the 8 weeks, the percentage degradation had reached 91% as shown in Figure 1.

Bi-weekly Total Petroleum Hydrocarbon Degradation of crude oil by Aspergillus flavipes as shown by GC-FID Analysis

The bi-weekly total petroleum hydrocarbon (TPH) degradation was investigated for a period of 8 weeks and the following inferences were drawn. Fractions C10 and C11 were significantly degraded. Whereas, fractions C12-C20 were moderately degraded; however, fractions C26-C34 were insignificantly degraded as shown in Figure 2.

Discussion

A little above two decades ago, scientists believed that hypersaline environments were occupied by archaea,bacteria and a eukaryote, the alga named Dunaliella salina. But today, several researchers such as [28-33] have isolated a plethora of eukaryotic fungal species predominantly Cladosporium, different species within the anamorphic Aspergillus and Penicillium, the teleomorphic Emericella and Eurotium, certain species of non-melanized yeasts represented by black, yeast-like hyphomycetes: Hortaea werneckii, Phaeotheca triangularis, Trimmatostroma salinum, and Aureobasidium pullulans, together with phylogenetically closely related Cladosporium species, all belonging to the order Dothideales, and Wallemia sp from various hypersaline waters of the world and this work has corroborated that fact by isolating numerous halophilic fungi from uburu and Okposi salt lakes (Tables 1 and 2). A total of 14 different fungal isolates namely Aspergillus flavipes (13mm at 40%), Penicillium citrinum (10mm at 40%), Aspergillus ochraceus (9mm at 40%), Aspergillus nomius (15mm at 35%), Microsphaeropsis arundinis (12mm at 35%), Aspergillus sydowi (28mm at 30%), Penicillium janthinellum (26mm at 30%), Mucor sp (13mm at 30%), Aureobasidium sp (12mm at 30%), Trichoderma sp (9mm at 30%), Alternaria sp. (22mm at 25%), Aspergillus sp (18mm at 25%), Penicillium sp (20mm at 20%) and Cladosporium sp. (7mm at 15%) were isolated from Uburu and Okposi salt lakes throughout the 4 quarters and two seasons of the the year (Table 2). The seasons were divided in to rainy (April-September) and dry (October-March). Aspergillus and Penicillium species occurred throughout the study period in both lakes. However, the top 7 extreme halophilic fungi viz. Aspergillus flavipes, Penicillium citrinum, Aspergillus ochraceus, Aspergillus nomius, Microsphaeropsis arundinis, Aspergillus sydowi and Penicillium janthinellum were characterized and identified phenotypically and genotypically, while, the rest were characterized only phenotypically (Tables 1 and 2). Of the 14 isolates, 12 were extremely halophilic, one was a borderline extreme halophile and another was a moderate halophile; moreover, Gunde-Cimerman et al. (2009) stated that fungi isolated from hypersaline habitats of 1.7 M ( ̴10%) salt concentration and can grow invitro at or above 3 M ( ̴18%) salt concentration should be considered as a halophilic fungus (Table 2). All the organisms isolated in this study tolerated this salinity range, and some of them exceeded the salinity range as well. No slight halophile was isolated in this work as seen in table 2. These findings agrees to some extent to the work of Mansour (2017) who collected sandstone samples from the Medamoud, Egypt, and halophilic fungi (Aspergillus nidulans, Aureobasidium pullulans and Cladosporium sphaerospermum) from a local culture bank. Mansour estimated salinity tolerance by growing these fungi at 0% to 25% of NaCl concentration supplemented on potato dextrose agar and broth and posited that 5% of NaCl was the best suited growth supplement for halophilic fungi and that halophiles showed better tolerance to salt as compared to the solid media; and that, 25% of NaCl concentration was found to inhibit any fungal growth. The fact that no slight halophilic fungus was isolate in this work is also in concordance with the reports of [12,19,20,23], who stated that micro organisms belonging to the slight halophiles are mostly marine bacteria such as Vibrio. The most halophilic fumgi that was isolated in this study was Aspergillus flavipes which had a colony diameter of 13mm at 40% salt concentration; hence it was used for biodegradation studies.

The growth tolerance of Aspergillus flavipes on different percentage concentrations of crude oil (2%, 4% and 6%) in Bushnell-Haas broth was studied. There was insignificant growth on 4% and 6% crude oil broth as shown by total fungal count. Only 2% crude oil broth had significant confluent growth with a fungal count of 2.56x105 CFU/ml. thus the biodegradation study was performed using 2% crude oil in broth, as shown in Table 3. Obuewe et al. (2005) have demonstrated that two halophilic fungi namely Fusarium lateritium and Drechslera sp. were able metabolize crude oil, with degradation efficiency improved more than thrice if petroleumwas an additional source of carbon and energy instead of being the exclusive one [24-35]. However, in this work crude oil was an exclusive source of carbon and energy and hence reason why Aspergillus flavipes tolerated 2% crude oil concentration out of the 3 concentrations assayed (Table 3).

The crude oil degradation rate of Aspergillus flavipes was evaluated gravimetrically and chromatographically bi-weekly. The gravimetric analysis showed that after the first two weeks, the crude oil had been degraded by 36%, by 4 weeks, it was 67%; by 6 weeks, it was 89%; and upon completion of the 8 weeks, the percentage degradation had reached 91% as shown in Figure 1. However, the bi-weekly total petroleum hydrocarbon (TPH) degradation was investigated for a period of 8 weeks also by way of Gas chromatograph coupled with flame ionization detection, and the following inferences were drawn. Fractions C10 and C11 were significantly degraded. Whereas, fractions C12-C20 were moderately degraded; and fractions C26-C34 were insignificantly degraded as shown in Figure 2. This agrees with the findings of [37, 38] who stated that hydrocarbons differ in their susceptibility to microbial attack and generally degrade in the following order of decreasing susceptibility: n-alkanes >branched alkanes>low molecular weight aromatics >cyclic alkanes, >polyaromatic hydrocarbons.

Conclusion

Crude oil contamination affects the environment and in return, the environmental conditions impact its bioremediation processes. As a complex mixture, different hydrocarbon groups have varying bioavailability and biodegradability leading to different intrinsic bioremediation applicability. Aspergillus flavipes tested in the present study can be used in oil bioremediation programmes as it has the activity to grow in crude oil amended Bushnell-Haas broth as evidenced by the decline in the total petroleum hydrocarbon content of bonny light crude oil over an eight week period with 89 % TPH degradation rate, as shown by gravimetry and gas chromatography.