Characterization of culturable microbial community in oil contaminated soils in Greater Port Harcourt Area, Nigeria

1 Department of Microbiology, Faculty of Science, University of Port Harcourt, Nigeria. 2 Department of Animal Science, Faculty of Agriculture, University of Port Harcourt, Nigeria. 3 Center of Excellence in Phytochemicals, Textiles and Renewable Energy, Faculty of Science, Moi University, Eldoret, Kenya. 4 Department of Plant Science and Biochemistry, Faculty of Science, University of Port Harcourt, Nigeria. 5 Department of Physical and Biological Sciences, Faculty of Science, Bomet University College, Kenya.


INTRODUCTION
Rapid population growth and an immense industrial revolution, even though beneficial in the civilization of human living standards, have jeopardized the state of the environment (Zhao et al., 2017;Jacob et al., 2018;Liu et al., 2019) by introducing a variety of toxic substances such as total petroleum hydrocarbons (TPH), polycyclic aromatic hydrocarbons (PAHs), pesticides, heavy metals, synthetic pigments, and polychlorinated biphenyls (PCBs) (Bilal et al., 2017;Barrios-Estrada et al., 2018). In Nigeria, the Niger Delta region is a major centre of activities of oil mining and its' associated industrial sectors. This oil-rich region accounts for more than three quarters of Nigeria's total annual revenue. This industrial growth has led to environmental degradation and left vast footprints of hydrocarbons in the environment (Lindén and Pålsson, 2013). Exploration of oil in Nigeria started in the 1950s and large processing facilities were built to harvest this mineral resource. The eruption of oil spills is on the rise due to increased exploration and insufficient environmental management strategies and this has led to the accumulation of total petroleum hydrocarbons (TPH) over time in sensitive natural habitats. Such chemical spills have intensified, contaminating soils not just in industrial areas but also in the agricultural areas. The consequences of pollution have left undesirable environmental and socio-economic issues leading to loss of ecological resources, poverty and public health concerns (UNEP, 2011;Nkonya et al., 2016;Wali et al., 2019). Further, projections show that the global population may exceed 9 billion by 2050 and that agricultural production would have to rise by 70 to 100% to support the growing population (National Geographic Society, 2020). Yield improvements cannot be accomplished unless the ecosystem is controlled to protect the integrity of the soil ecosystem.
Microorganisms are ubiquitous and are an integral part of the environment since they play a vital role in maintaining processes including biogeochemical cycles in the ecosystem. Soil microorganisms' biogeography is essentially distinct from their counterparts found in animals and plants, and is thus still poorly understood (Whitman et al., 1998). It is vital to assess the impact of anthropogenic activities on the structure of the soil microorganism community in order to provide a basis for reference to the positive and negative impacts that may occur in soils. Microbial diversity of soil must be maintained at its optimum level in order to achieve longterm agricultural productivity. Also, knowledge of soil quality is important for the effective management of farms as it provides baseline data on strategies to maintain and improve soil fertility (Zhou et al., 2014). Soil microorganisms' metabolic activities are mainly driven by temperature and physicochemical parameters (Yang et al., 2020). Bacteria containing nirK, nirS, and nosZ-I genes often have a unique composition in farmland soils as compared to wetland soils, with nirK and nirS being particularly distinct from those containing nosZ-I (Bowen et al., 2020). A reduction in soil pH decreases the abundance of genes and changes the composition of nirK and nirS in agricultural and wetland soil, and raises the ratio of N 2 O: (N 2 +N 2 O) in agricultural soils (Bowen et al., 2020). Agricultural practices have a significant influence on chemical and microbiological soil parameters affecting Wanjala et al. 33 soil fertility (Bowen et al., 2020). The patterns of microorganisms in soil polluted with petroleum products vary depending on the chemical composition of the soil and the type of petroleum products. Escherichia coli, Pseudomonas species, Bacillus species, Proteus species, and Penicillin species, have been identified to exist in soil contaminated with cyanide (Eze and Onyilide, 2015). The analysis shows the presence of microorganisms in soils contaminated with cyanide at a concentration of 3.0 mg/kg, showing that microorganisms can survive in cyanide contaminated habitats. Soil pollution affects the population and diversity of soil microorganisms. Microbial diversity is declining with an increase in contamination of the environment (Xie et al., 2016). The presence of low levels of microorganisms is related to increased intoxication of cadmium in soils (Xie et al., 2016). Exposure of microorganisms to the concentration of pollutants in soil is therefore causative to the development of adaptive characteristics among the various species found in contaminated soils. Acquisition of new genes that are responsible for resistance for toxicants is an option for microorganisms in the environment. Heavy metal contaminated soil in the marketplaces (Uyo, Umuabia, Sokoto and Oka) in Nigeria has been shown to influence the diversity and distribution of soil microorganisms (Akpoveta et al., 2010;Ogbemudia and Mbong, 2013;Eze et al., 2013;Imarhiagbe et al., 2017). Furthermore, the growth of microalgae (Microcystis flos-aquae) in crude oil contaminated media show an exponential growth and reduction of crude oil in the media, an indication of the potential of microorganisms for oil degradation in polluted environments (Ifeanyi and Ogbulie, 2016), and adaptation by shifts in microbial populations, species richness and diversity, thus the role played by microorganisms is diverse. The use of oil spills in Calabar Cross River State in Nigeria has been shown to influence the distribution of microorganisms in soil (Unimke et al., 2017). Some heterotrophic bacteria isolated from these soils included: Pseudomonas spp., Bacillus spp., Klebsiella species, Proteus spp., Enterococcus faecalis and Flavobacterium species (Unimke et al., 2017). The total hydrocarbon utilizing bacteria (THB) include Bacillus spp., Pseudomonas spp., and Micrococcus species (Unimke et al., 2017). Highly prevalent genera were Pseudomonas spp., and Bacillus spp., indicating that oil degradation microbes are more abundant in oil contamination areas (Unimke et al., 2017). Arthrobacter species, strain YC-RL1, could use bisphenol A (BPA) as a carbon source to grow in contaminated soil (Ren et al., 2016). Sourced from soils that were contaminated with crude oil, Planococcus maritimus Isolate Y42 was able to use crude oil as its sole source of energy carbon (Yang et al., 2018).
Pseudomonas, Rhizobium, Rhodococcus, Sphingomonas, Enterobacter, Acinetobacter, Bacillus, Paenibacillus, and Variovorax species were found in various petroleum contaminated soils and had high biodegradability on alkane mixtures with diverse lengths of the carbon chain ranging between C9 to C30 (Zheng et al., 2018). The microbial diversity in petroleum contaminated soils may be different in soils with similar or different types of contaminants, as most contaminated soils are also polluted by other industrial wastes and chemicals. These studies show that anthropological activities are a threat to soil ecosystem integrity and it is important to periodically monitor the concentration of pollutants in the soil and their effects on soil microorganisms.
The objective of this study was to determine microbial diversity in oil-contaminated soils in three broad sites; urban, industrial and agricultural. Morphological and biochemical experiments have been used to classify a variety of cultivable microbes. The findings of this study could be useful in the production of highly efficient isolates for bioremediation of soils contaminated with petroleum oil.

Description of the study site
This study was conducted in nine selected test and three control sites in Port Harcourt, the Capital of Rivers State, Nigeria ( Figure  1). The study sites were grouped into three areas, including urban (GRA phase 2, Diobu-Mile 1 and Mguoba), industrial (Eleme hosting NNPC Refinery, Agbada-SPDC-flow station) and agricultural (Aluu, Oquwi-Eleme, Emuoha-Eu). The study sites were characterized with different economic activities shown in Table 1.

Sampling
In the wet season (April to October 2018), composite samples were collected by random sampling from each of the three areas; urban, industrial, and agricultural. Five (5) samples were collected at random around each test field. The five individual samples were thoroughly mixed in a sterile jar by coning and quartering to achieve a homogeneous composite blend. A total of 12 composite samples; A1, A2, A3, I1, I2, I3 U1, U2 and U3 as test samples, and CA, CI and CU as control samples (Table 1), were collected simultaneously. The samples were obtained at a depth of 0 to 15 cm from the top of the soil using a regular auger three times during the rainy season. Homogenized composite samples (400 g) were then wrapped using a sterile wooden shovel into polyethylene bags. Samples were collected for microbial analysis using pre-sterilized materials to prevent sample contamination. The locations of the sampling sites were determined using the GPS and the measurements were recorded. Samples were taken to the laboratory in an ice box for examination.

Determination of total petroleum hydrocarbon (TPH) content of soil
The Hewlett Packard 5890 Series II Gas Chromatograph FID method was used. In this method, 1 g of well-mixed sample was weighed into Acetone rinsed beaker. Then, 1 g of anhydrous sodium sulphate was added to the soil sample and 5 ml of solvent (1:1 of dichloromethane and acetone) was added and stirred for 15 min using a magnetic stirrer and the ensuing mixture was poured into a round bottom flask. This was repeated once more by adding 5 ml of mixed solvent. It was stirred and permitted to stand/settle and then decanted into another round bottom flask. The solvent was concentrated with 1 ml hexane to exchange it and it was reconcentrated to 2 ml. The columns were eluted (washed off) with 10 ml n-hexane. 1 ml of the extract was pipetted into the column and 10 ml of n-hexane was used to collect the aliphatic components. The extract was concentrated to 1 ml and poured into a glass vial for Gas Chromatography.

Enumeration of total heterotrophic bacteria (THB)
Heterotrophic bacteria were enumerated by pour plate method (APHA, 1998). One gram of soil sample was weighed into 9 ml sterile diluent (0.85% NaCl) under aseptic condition (laminar bench floor). It was then homogenized using a laboratory vortex mixer (Model: 10101001, IP42) and serially diluted. Then 0.1 ml aliquot of the inoculum was collected using a sterile pipette, inoculated on Nutrient Agar (NA) medium. The inoculum was spread evenly using a sterile glass spreader stick. Plates were then incubated at 37°C for 24 h. Thereafter, colonies were counted and expressed as colony forming units (CFUs/mg of soil) value per gram of soil sample. Distinct colonies with different morphological patterns (color, size, shape, edge, elevation, surface and opacity) were picked and streaked or subculture on freshly prepared nutrient agar medium in order to obtain pure culture after 24 h of incubation at 37°C. The pure cultures were Gram stained for microscopic examination and were further subjected to biochemical tests.

Enumeration of hydrocarbon utilizing bacteria
Hydrocarbon utilizing bacteria (HUB) were enumerated by the pour plate method (APHA, 1998) method. 1 g of soil sample was weighed into a 9 ml sterile diluent (0.85% NaCl) under aseptic conditions. The sample was then homogenized using a laboratory vortex mixer (Model: 10101001, IP42) and serially diluted. Then 0.1 ml aliquot of the inoculum was inoculated on Mineral Salt Agar (MSA) medium containing g/l of MgSO 4 .7H 2 O 0.42 g, KCl 0.29 g, K 2 HPO 4 1.25 g, KH 2 PO 4 0.83 g, NaNO 3 0.42 g, NaCl 10 g and Agar Powder 18 g, using the spread technique. Sterile filter paper (Whatman 540) was soaked with crude oil and placed in the lid of petri dish. Plates were incubated in inverted position at room temperature for 5 days until there was observable growth. Thereafter, distinct colonies were purified by sub-culturing on a freshly prepared medium and incubated for 24 h, from which microscopic examination and biochemical tests.

Enumeration of total fungi
Total fungi were performed using a pour plate method (APHA, 1998). Under aseptic conditions, one gram of soil sample was weighed in a 9 ml sterile diluent (0.85 per cent NaCl). The sample was then homogenized using a vortex mixer (Model 10101001, IP42) and diluted in series using sterile pipettes. Thereafter, 0.1 ml of the inoculum aliquot was inoculated on Potato Dextrose Agar (PDA) mixed with an antibacterial reagent (Normocure TM ) to inhibit bacterial growth and allow only fungal growth. Then, the inoculated plates were incubated for 5 to 7 days at ambient temperature. To obtain colony forming unit per gram (CFU/g) of the soil, colonies were enumerated using a colony counter.

Enumeration of hydrocarbon utilizing fungi
Hydrocarbons utilizing fungi (HUF) were cultured using the pour plate method (APHA, 1998). Under aseptic conditions (laminar flow bench), 1 g of soil sample was weighed into a 9 ml sterile diluent (0.85% NaCl). The sample was then homogenized using a laboratory vortex mixer (Model: 10101001, IP42) and serially diluted using sterile pipettes. 0.1 ml aliquot of inoculum was then inoculated on Mineral Salt Agar (MSA) mixed with an antibacterial reagent (Normocure™) in order to inhibit the growth of bacteria and allow for only growth of fungi. Sterile filter paper (Whatman 540) was subsequently soaked with crude oil and put in the petri dish cover. At room temperature, the plates were then incubated in an inverted position for 5 to 7 days. Colonies were counted using a colony counter to get colony forming units per gram of soil. Cultural characteristics (colour and microscopic observations) of the isolates were then observed and purified by sub-culturing on freshly prepared medium and incubated again for 3 to 5 days. From the pure cultures, microscopic examination was done using lactophenol cotton blue stain and observed under ×400 magnification.

Determination of % hydrocarbon utilizing fungi and bacteria
Percent hydrocarbon utilizing fungi and bacteria were expressed as a fraction of the total heterotrophic viable count using the formula:

Characterization and identification of THB and TF fungi
The fungal and bacterial isolates were identified morphologically (color, size, shape, edge, elevation, surface and opacity). Further, bacterial isolates were identified biochemically and characterized according to the scheme of Bergey's manual of Determinative Bacteriology (Holt et al., 1994) Table 2 shows the ranks of prevalence, diversity of THB, Table 2. Variation in prevalence of THB in soil from agricultural, industrial and urban areas in Greater Port Harcourt Area, Nigeria.

Prevalence and diversity of HUB in sampling sites
(3) between all the industrial and the control site; however, there was a difference in diversity between the controls and contaminated sites in urban and agricultural areas. Table 4 shows the ranks of prevalence and diversity of TF in the sampling sites. The most prevalent fungal isolate includes Aspergillus niger, Aspergillus flavus and Candida torulopsis which were prevalent in 12, 6 and 5 sites, respectively (Table 3). The highest diversity was observed in A2 (Eleme) with 5 isolates A. niger, Mucor mucedo, Saccharomyces cerevisiae, Paecilomyces species, and Geotrichium species (Table 3). The list diversity was observed in A3 (Emuoha), I1 (Onne) and U3 (Mgbuoba) with 3 isolates each. Table 5 shows the different hydrocarbon utilizing fungi (HUF) that were isolated from the soil samples. A. niger