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Bacteriological and polycyclic aromatic hydrocarbon evaluation of mangrove phyllosphere and rhizosphere from Qua Iboe river estuary, Nigeria.


Mangrove ecosystems are heterogeneous habitats with an unusual variety of animals and plants adapted to the environmental conditions of highly saline, frequently inundated, soft bottomed anaerobic mud (Clough, 1992). Rhizophora racemosa belongs to the family Rhizophoraceae. They are the most dominant tropical mangroves worldwide and are generally believed to play a vital role for mangrove ecosystems including shoreline protection, enhancement of water quality in near shore environments and support of estuarine and marine food chains (Duke and Bunt, 1999). As keystone specie, one that influences the community by its activity or role, not its numerical dominance (Newton, 2010), Rhizophora racemosa is a unique member of the metacommunity of biological species.

The Qua Iboe River mangrove estuary is mesotidal and like other estuaries in the Niger Delta, Nigeria, it is exceptionally productive and provides critical habitat for fishes, shellfishes, migratory water birds and a diversity of wildlife. The impact of the estuary on the lifestyle of the people in the area cannot be overemphasized. Oil and gas are among the natural resources in the Qua Iboe River Estuary (QIRE) and contributes substantially to the economic growth of the area, particularly Nigeria. However, oil and gas exploration and production is mainly to blame for oil spills and associated damages to plants, aquatic life, and air pollution from gas flaring, produced water and contaminants from drilling fluids discharged into the ecosystem. In addition, pollutant in storm water from urban and agricultural runoff and sewage are discharged directly into the estuary with its attendant detrimental effect. Environmental pollution in the Qua Iboe mangrove estuary is primarily a result of increasing concentration of inorganic and organic compound including polycyclic aromatic hydrocarbons.

PAHs are formed mainly as a result of pyrolytic processes, especially the incomplete combustion of organic materials during industrial and other human activities, such as processing of coal and crude oil, combustion of natural gas, combustion of refuse, vehicle traffic, cooking and tobacco smoking, as well as in natural processes such as carbonization (Allain, 1994; ATSDR, 1994).

PAHs constitute a group of priority pollutants which are of increasing environmental concern because of their adverse effect on humans, animals and plants. Soils, sediments, and broad leaf generally serve as a sink of PAHs which leads to the bioaccumulation of PAHs at contaminated sites. However, despite the common use and cost effectiveness of bioremediation, it is generally observed that a residual fraction remains undegraded even when optimal biodegradation condition has been provided. In many cases the recalcitrance of this residual fraction is caused by a limited bioavailability (Cuypers, 2001).

The nature and degree of oil spills, which constitute PAHs impacts to mangrove are related to three variables: The type of oil, the life stage, and the topographic elevation on which the mangroves are colonizing. In general, the bioaccumulation varies with the degree of contamination. Mangrove can suffer impacts when their roots are coated or smothered with oil, preventing gas exchange. The mangrove may be directly exposed or poisoned via adsorption of the toxic soluble fractions of PAHs through the pneumatophores and prop roots. Mangroves may also be affected by chronic exposures to PAHs--contaminated sediments. Effects of bioaccumulation of PAHs in mangroves may exhibit one or more of the following symptoms: Partial to complete defoliation, low survival of propagules, leaf deformities, reduced leaf size and flower production, increased insect infestation, death of the individual plant (Bender et al., 1980).

The effect of human activities in the QIRE increasingly threaten environmental health. Of special concern is biodiversity and habitat loss by native biota with invasive species rapidly colonizing the area putting many unique species at risk. This study was carried out to determine the densities and distribution of PAH-utilizing bacteria and PAH levels in the rhizosphere and phyllosphere of Rhizophora racemosa with a view to ascertain the degradative and bioaccumulative potentials of the bacteria and mangrove plants respectively. The interaction of the biota and pollutants in the mangrove ecosystem and possible ecotoxicological effect due to bioaccumulation are highlighted.

Study Area

Qua Iboe River Estuary is located on latitude [4.sup.0] [30.sup.I] to [4.sup.0] [45.sup.I] N and longitude [7.sup.0] [31.sup.I] to [8.sup.0] [45.sup.I] E in the south-eastern part of Nigeria at Ibeno Local Government Area in Akwa Ibom State (Fig.1). It has a permanent human settlement of mainly fishermen around the river bank. Qua Iboe River Estuary is a mesotidal estuary having tidal amplitude of 1m and 3m at neap and spring phases, respectively. Although sandy beaches are known to develop in some portions of the estuary, most of them are fringed with tidal mud flats and mangrove swamps in which Rhizophora racemosa, R. harrisoni, R. mangle, Avicennia and the brackish water palm, Nypa fructicans are the dominant flora species (Chapman, 1975). Like other estuaries in the Niger Delta, the Qua Iboe Estuary experiences great fluctuations in salinity between the wet and the dry seasons, as well as the normal gradient extending up-stream from the mouth of the river (Essien et al., 2007). Ecological habitats therefore vary from purely marine to those of brackish and freshwater. Tidal currents which are strong at the mouth of the estuary but weak along its upper ridges and creeks; play an important role in the biota distribution including the distribution of algae in the coarse sandy recreational beaches (Ukpong, 1995).


Collection of Sample

The rhizosphere, leaf and root samples were taken from mature R. racemosa (21-25cm stem diameter at base just above the highest prop root, 6-8m tall) which are indigenous in the QIRE. The species of Rhizophora have dense distribution in the seaward outer fringes of the shoreline where the population of invasive N. fructicans is low. Rhizosphere and root samples were collected by excavating the root system of the plant during low tide. Samples for chemical analysis were collected into Amber glass containers with Teflon-lined cap, stored in the dark at 4[degrees]C with a maximum holding time of 6 h before extraction. Prop roots and leaves of the mangrove (R. racemosa) were obtained with the aid of machete into polythene bags. Sampling was done during the rainy season (months of June - July). Equal amount of sediment samples and mangrove rhizosphere soil were collected from each location (at approximate distance of 10m, 100m and 200m from the flare point) and formed into a composite sample to reduce the total number of samples and associated cost of the analysis. The composite sample was thoroughly homogenized. Precisely 10g subsample of the homogenized sample was serially diluted for microbiological analysis (Harrigan and McCance, 1990)

Enumeration of Heterotrophic, Hydrocarbon- and PAH- utilizing bacteria.

The bacteria isolates on the leaf surface were enumerated by counting the viable colonies using the "leaf imprint technique". Sterile molten nutrient agar cooled to about 40[degrees]C was poured in 15ml amount to sterile Petri-dishes. The leaf sample was moistened with sterile water and used to make an imprint of the phyllosphere and phylloplane on the agar. The counts of total heterotrophic bacteria (THB) in the sediment and mangrove rhizosphere was enumerated by pour plate technique (Harrigan and McCance, 1990) using diluents prepared with 25% Ringer's solution and cultured on nutrient agar (Difco) and starch nitrate agar. The hydrocarbon utilizing bacteria (HUB) were enumerated by the spread plate technique using oil-mineral salt medium (MSM). The media were supplemented with cycloheximide (100[micro]g/ml and benomyl (50[micro]g/ml) to prevent fungal growth (Kinkle et al, 1995) The crude oil used was sterilized by filtering through Millipore filter (0.45[micro] pore size) and stored in sterile glass bottles. Bushnell-Haas (BH) minimal medium (Sigma-Aldrich) supplemented with 1.5% NaCl was used for the enrichment and isolation of PAH-utilizing Vibrio species.

PAH-utilizing bacteria (PAHUB) were isolated using enrichment cultures in 100ml Erlenmeyer flasks containing 20ml of M9 medium (Maniatis et al., 1982) supplemented with 1g/l of pure crystal naphthalene (Sigma) to avoid solvent as potential substrates and incubated at 28 [+ or -] 2 [degrees]C for 28 day on orbital shaker at 90rpm. Uninoculated medium served as control for each culture. Inoculated THB, HUB and PAHUB plates were incubated aerobically at room temperature (28 [+ or -] [2.sup.0]C) for 24 hours, 5 to 14 days and 21-28 days respectively and thereafter enumerated (Harrigan and McCance, 1990; Amadi and Braide, 2003).Growth to turbidity after incubation confirmed the presence of organisms capable of utilizing PAH as sole carbon source. Representative bacterial colonies were purified by repeated subculturing and maintained as stock on nutrient agar slants. The identification of the isolates was done by comparing the cultural, morphological and biochemical characteristics of the cultures with the characteristics of known taxa using the Bergey's manual of determinative bacteriology (Holt et al., 1994) and Cowan and Steel's manual for the identification of medical bacteria (Barrow and Feltham, 1992).

Chemical Analysis

The analysis of PAH concentration was carried out using standard procedure (APHA 1998; Radojevic and Bashkin, 1999). Briefly, each of dried and ground sample spiked with squalene and 32-alkane were serially extracted with 100 mL methyl isobutyl ketone (Analar grade). Each extract was allowed to settle, centrifuged for 5 min and decanted. The extracts were concentrated on a rotatory evaporator maintained at 20[degrees]C to a volume of about 5ml. A sample volume of 1[micro]l each of the extract was subjected to a GC-MS (Hewlett Packard model 5890). Concentrations of polycyclic aromatic hydrocarbons were quantified relative to the total peaks as these were converted to weights using hydrocarbon standard calibration (FEPA, 2001). Duplicates and method blanks were similarly treated using the same reagents to test for the precision, accuracy and reagent purity used in the analytical procedure.

Determination of Bioaccumulation Factor

The bioaccumulation factor (BAF), a dimensionless inde x indicating that the factor relates accumulated pollutant to that in sediment (Spacie et al., 1995) expressed as:

BAF = [C.sub.organism]/[C.sub.source]

Where [C.sub.organism] is concentration in the organism resulting from uptake from food and water sources and [C.sub.source] is concentration in the reference source of pollutant. The concentration of PAHs in the ambient air was not determined in this work and thus bioaccumulation of the pollutant was based on plant-sediment interaction. If the BAF is greater than 1, biomagnification may be occurring. If it is less than 1, trophic dilution is suggested although other factors such as allometric processes or growth dilution may be contributing to the changes (Newman, 2010).

Statistical Analysis

Correlation analysis of data were performed using Analyse-It General 1.73 statistical software[R] on Log--transformed estimates of bacterial densities (Log cfu/g) with levels of significance maintained at 95% for each test. Interpretation was done based on Hinkle et al., (1994) rule of thumb for interpreting the size of a correlation coefficient.

Results and Discussion

Culture-dependent bacteriological analysis of the samples revealed that there was spatial variability in the bacterial density in the rhizosphere and phyllosphere of the plant as well as the sediment. The bacterial density was in the order phyllosphere < sediment < rhizosphere (Table 1). The phyllosphere HET, HUB, PAHUB and TVC in the three locations ranged from 92/0.54 to 142/0.62 cfu/[m.sup.2], 74/0.58 to 82/0.62 cfu/[m.sup.2], 47/0.56 to 52/0.62 cfu/[m.sup.2] and 00/0.56 to 21/0.(50 cfu/[m.sup.2]. The rhizosphere HET, HUB, PAHUB and TVC ranged from 12.5 x [10.sup.5] to 13.2 x [10.sup.5] cfu/g, 9.4 x [10.sup.5] to 10.9 x [10.sup.5] cfu/g, 7.9 x [10.sup.5] to 11.9 x [10.sup.5] cfu/g and 5.8 x [10.sup.5] to 7.9 x [10.sup.5] cfu/g whereas the counts in the sediment ranged from 10.6 x [10.sup.5] to 12.1 x [10.sup.5] cfu/g, 7.6 x [10.sup.5] to 9.2 x [10.sup.5] cfu/g, 7.1 x [10.sup.5] to 9.7 x [10.sup.5] cfu/g and 4.7 x [10.sup.5] to 7.4 x [10.sup.5] cfu/g respectively. This is indicative of the influence of the mangrove plant root in altering the physicochemical properties of its microhabitat allowing diverse microbes to proliferate. The microbial load as expected was in the order: location 1(10m) < location 2 (60m) < location 3 (110m) from the flare point though with no cultural difference in the microbial diversity indicating ubiquity of the isolates. The low microbial load at location 1 could have been as a result of heat from the infernal flame though an environmentally sensitive organism like E. coli was isolated at location 1 indicating recent fecal contamination of the site. There was a moderate to very high positive correlation (r = 0.69, 0.97 and 0.97) in the microbial load of the rhizosphere--sediment samples in locations 1, 2 and 3 respectively.

The high count is a strong evidence of microbial diversity and abundance in the ecosystem most of which were found to have the ability to utilize PAH, indicated by the level of turbidity produced in the PAH-mineral salt medium compared with the uninoculated control (Table 3). Among the isolates, Pseudomonas aeruginosa, P. putida, Micrococcus varians, Bacillus subtilis, Acinetobacter iwoffii, Nocardia sp, Anabaena and Nodularia exhibited profuse growth on the fourteenth day of incubation indicating strong potential to utilize the low molecular weight PAH. Moderate growth and utilization was exhibited by Vibrio alginolyticus, Chromobacterium violaceum, Vibrio. estuarianus, Alcaligenes denitrificans, Sarcina sp and Flavobacterium breve. Despite the abundance of the isolates in the samples, Serratia marcescens, Escherichia coli, Enterobacter aerogenes, Chromatium sp, and Erwinia amylovora exhibited minimal growth towards the end of incubation period indicating weak ability to utilize the PAH. V. alginolyticus, Anabaena, Nocardia and E. coli were not isolated from the phyllosphere of the plant (Table 2).The ability of these organisms to survive the toxic nature of PAH could probably be due to pre-exposure and/or acquisition of metabolic plasmids in the environment and could be involved in cometabolic utilization of PAH given their abundance in the samples. This result is in agreement with the findings of Udotong et al., (2008) that pre-exposure to petrogenic waste in the environment could enhance the degradation potential of the organisms.

Chemical analysis also revealed that there was variability in the concentration of PAH in the rhizosphere, phyllosphere and sediment of the study area. Spatial differences in PAH levels in the plant leaf, roots and rhizosphere sediment revealed that the plant does not excrete PAH as in the case of salt, instead accumulates them. This was evident in the elevated PAH concentration in the leaf that ranged from 18.65mg/kg to 25.21 mg/kg compared to the root (9.92 to 11.13mg/kg) against a background concentration of 8.92mg/kg to 9.55mg/kg (sediment) and 7.26mg/kg to 7.51mg/kg (rhizosphere). The low sediment and rhizosphere PAH level (Table 4) could be as a result of photo-oxidation, volatilization and rhizodegradation before uptake of the bioavailable pollutants.

Three possible reasons could be deduced for the elevated concentration of PAH in the mangrove phyllosphere. First, prolonged uptake and accumulation of the contaminant and poor phytodegradative potential of the mangrove plant. Second, the low rate of evapo-transpiration and unavailability of the contaminant for interaction with phyllosphere bacterial species due to cuticular separation. Third, the leaves of the plant receive daily atmospheric deposition of soot and diffusion of volatile PAHs primarily through open stomatal pores exacerbated by elevated CO2 concentration which encourages photosynthesis. Also, the phyllosphere is a harsh environment that does not favor enhanced cooperative interactions between microbial species that could degrade the pollutant through cometabolism. Though plant exposure to both gaseous and solid phase PAHs in ambient air was not determined in this study, the process of accumulation is reported to be affected by several abiotic and biotic factors including vapor-particulate partitioning in the atmosphere, ambient temperature, octanol-air partitioning coefficients, leaf surface area and lipid concentration in the plant tissues (Slaski et al., 2000).

Nestled among the fast spreading Nypa palms are patches and isolated stands of the mangrove plants at the fringes of the shoreline close to the water atrophied and overshadowed by the invasive Nypa fructicans. The species richness of the Qua Iboe River estuarine ecosystem is therefore affected by the N. fructican which contributes to the steady asphyxiation and differential mortality of the mangrove trees. This is in addition to the major long term local threat to the mangrove plant from clear-cutting and clearing for use as fire wood and subsistent agricultural practices in an effort to survive the persistent wave of lack, poverty and poor living conditions prevalent in the area changes the estuarine community structure. Defoliation by anthropogenic pressure and leaf fall returns the accumulated PAH to the sediment and biota. Foliar herbivores such as swimming crab (Callinectes latimanus) on the mangrove plant and other lower level biota in the food chain are therefore exposed to high PAH concentrations (Eduok et al., 2010) that may rise to toxic levels if metabolism is not sufficient to prevent rapid accumulation that may result in biomagnification along the food chain. The BAF for the three locations (Table 5) indicated that values for phenanthrene, pyrene, benzo(a) anthracene, chrysene, benzo(b) fluoranthene, benzo(k) fluoranthene, benzo(a)pyrene, dibenzo(a,h)anthracene and benzo (g,h,l) perylene were higher than 1. This is suggestive that biomagnification of PAHs with values higher than 1 is occurring along the food chain in the mangrove ecosystem. This finding is very significant in view of the fact that humans depend on the aqua-terrestrial resources for dietary protein supplement may have elevated tissue burden of PAH with adverse toxicological implications as most of the PAH are known carcinogens. The leaves of Rhizophora racemosa has ethno-medicinal value to the people of south eastern Nigeria (Obute and Adubor, 2007), however, the use of the plant parts could have negative reaction in body tissues with grave consequences considering the PAH accumulation in the mangrove plant.

Apart from diffuse sources from surface runoff, sewage and bush burning, significant PAH loading of the estuarine ecosystem is from the pyrogenic and petrogenic activities of ExxonMobil and allied companies with the emission of soot, smoke and particulate matter in the area. It has been demonstrated that certain lower molecular weight (LMW), non carcinogenic PAHs, at environmentally realistic levels were acutely toxic to aquatic organisms, or produced deleterious sublethal responses (Neff, 1985). The most obvious could be the low population densities of the mangrove oyster compared with other oyster beds in the estuary where the populations of R. racemosa in the ecosystem are not asphyxiated (Eduok et al., 2010). Most biologist would agree that a 50% reduction in species richness is a clear indication of diminished health of an ecological community. Ecological overshoot, using resources faster than they can be regenerated (Newman, 2010) by man is rapidly decimating the mangrove species in the QIRE. The average ambient wet season air temperature of the estuary was 43[degrees]C at about 10m (Location 1) from a flare point and decreased inversely with distance from the source. This in addition to global warming, rise in sea level, coastal erosion and Nypa fruticans encroachment makes the mangrove plant more vulnerable to PAH-induced defoliation, falling, anoxia and rapid decline in population.

Although efficient and rapid PAH degradation generally depends on molecular oxygen availability (Cerniglia, 1992, Chung and King, 1999, Leahy and Olsen, 1997), the low availability of molecular oxygen in the anoxic estuarine sediment reduces the rate of PAH degradation limiting it to those environmental matrices with rapid oxygen exchange as indicated by the low PAH levels in the rhizosphere. The rhizosphere, the soil directly under the influence of the plant root harbours a wide diversity in species and number of organisms with modified physicochemical and nutritional properties that encourage interaction of the biotic and abiotic environmental components. Several reasons could be deduced for the low PAH level in the rhizosphere: i) Plants may enhance rates of PAH degradation by stimulating microbial growth and activity, and/or by increasing PAH availability in the rhizosphere (Binet et al., 2000). ii) The presence of plants may influence carbon dioxide concentration, pH, osmotic potential, redo x potential and oxygen concentration (Anderson et al., 1993). iii) Plant characteristics such as species, age, root type, soil type, and the weathered age of the contaminant will determine the structure of microbial communities in the rhizosphere (Sylvia et al., 1998). iv) The size of root impact zone in the rhizosphere is determined by the concentration and type of carbon exudates (Taiz and Zeiger, 1998), and expands with increasing root hairs, root overlap and fibrous roots (Sylvia et al., 1998). The degradation of comple x compounds such as PAHs may be enhanced by the presence of root exudates because of increased interaction between microbes, nutrients and contaminants (Reilley et al., 1996; Cunningham et al., 1996). The aerating prop roots of the mangrove plant (Rhizophora racemosa) above the ground or especially above water with its rich production of exudates, death and decay, are extensively interwoven in close physical proximity to the negatively orthogravitropic pneumatophores of Avicennia sp supplying air through the soft, spongy tissue to the roots and microorganisms in the anoxic region of the mud. The combined effect of the prop root and pneumatophores of Rhizophora and Avicennia respectively, could be responsible for the supply of the essential requirements needed for aerobic PAH degradation in the rhizosphere of the plant. This result is indicative of the in situ functional role of R. racemosa and its microbial rhizosphere flora in the sequestration and degradation of organic pollutants such as PAHs. The findings of the present study is consistent with previous reports (Aprill and Sims, 1990, Lee and Banks, 1993; Chaineau et al., 1997, Leyval and Binet, 1998; Daane et al., 2001; Musella, 2006) suggesting that plant cropping could be used as an alternative technique to reduce PAH levels in soils due to enhanced microbial activity in the rhizosphere. R. racemosa as keystone specie should be adequately protected in the general interest and health of the estuary and humans.


Qua Iboe River Estuary and the Niger Delta in general have experience prolonged pyrogenic and petrogenic contamination that pose serious health risk to the environment and human. Though awareness and concern about environmental issues is changing, little has been done to reduce pollutant emission/discharge into the environment or engage the society in managing the natural environment. This study has shown the rhizosphere-microbe effect on PAH-utilization indicated by the low PAH level in the rhizosphere compared to the non-rhizosphere sample. This is suggestive that the plant and its associated microbes could be effective in phytoremediative clean-up of the PAH contaminated QIRE and other estuaries in the Niger Delta. The PAH level in the sediment may represent a baseline concentration of the available PAH stressor in the estuarine ecosystem. The global climate change in addition to anthropogenic pressures, heat and salt stress, and Nypa palm asphyxiation have resulted in loss of mangrove-dependent productivity and biodiversity with negative socioeconomic and ecological impacts. The effect is long term and steady PAH accumulation, biomagnification and destruction of spawning sites of many estuarine-dependent biota such as crabs, oysters and fishes. Prohibition of gas flaring and cutting of existing mangrove for charcoal production and re-vegetation of intertidal areas by suitable species of mangrove especially the fast growing Rhizophora species appears to be the most plausible approach to curb the extinction of the mangrove and biodiversity that are already endangered. Also, biomagnification of PAH along the food chain with potential adverse toxicological effects on humans in the estuarine environment would be minimized. Rhizophora racemosa could serve as indicator of exposure for retroactive risk assessment of ambient air and sediment contamination with PAH in estuarine ecosystem.


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Samuel I. Eduok * (1), Basil N. Ita (2), Uwa O. Uye (3) and Nkoyo M. Utuk (4)

(1), (4) Department of Microbiology, University of Uyo, P.M.B. 1017,Uyo, Nigeria

(2) Department of Chemistry, University of Uyo, P.M.B. 1017, Uyo, Nigeria.

(3) Ministry of Health, Damaturu, Yobe State, Nigeria.

* Author for Correspondence E-mail:
Table 1: Microbial counts.

Sample Location 1a


P(cfu/[m.sup.2]) 92/ 74/ 47/ 0.56 00/
 0.54 0.58 0.56

R(x [10.sup.5]cfu/g) 12.9 9.4 7.9 5.8
 (6.11) (5.97) (5.89) (5.76)

S(x [10.sup.5]cfu/g) 10.6 7.6 7.1 4.7
 (6.03) (5.88) 5.85) (5.67)

Sample Location 2b


P(cfu/[m.sup.2]) 112/ 82/ 52/0.62 13/
 0.60 0.62 0.64

R(x [10.sup.5]cfu/g) 13.2 10.9 8.3 7.3
 (6.12) (6.04) (5.92) (5.86)

S(x [10.sup.5]cfu/g) 12.1 8.7 8.9 6.9
 (6.08) (5.94 (5.95) 5.84)

mple Location 3c


P(cfu/[m.sup.2]) 142/ 76/ 45/0.62 21/
 0.64 0.60 0.60

R(x [10.sup.5]cfu/g) 12.5 9.8 11. 9 7.9
 (6.90) (5.99) (6.08) (5.89)

S(x [10.sup.5]cfu/g) 11.3 9.2 9.7 7.4
 (6.05) (5.96) (5.99) (5.87)

Counts are mean of three determinations, Value in parenthesis are
log10, a = 10m ([43.sup.0]C), b = 60m ([38.sup.0]C), c = 110m
([25.sup.0]C). P = phyllosphere, R = rhizosphere, S = Sediment,
HET = heterotrophic bacterial count, HUB = Hydrocarbon utilizing
bacterial count, PAHUB = polycyclic aromatic hydrocarbon utilizing
bacteria, TVC = Total Vibrio count.

Table 2: Percentage frequency of Isolate occurrence.

Organism Phyllosphere Sediment Rhizosphere
 (N=9) (N=9) (N=9)

Vibrio alginolyticus 00 88.9 100
Pseudomonas aeruginosa 100 88.9 100
Vibrio estuarianus 11.1 100 100
P. putida 88.9 88.9 100
Alcaligenes denitrificans 66.7 88.9 100
Chromobacterium violaceum 88.9 100 88.9
Nocardia sp 00 88.9 100
Micrococcus varians 44.4 100 100
Acinetobacter iwoffii 77.8 88.9 100
Serratia marcescens 88.9 100 100
Bacillus subtilis 8.9 100 100
Chromatium sp 88.9 100 100
Escherichia coli 00 88.9 88.9
Anabaena 00 100 100
Enterobacter aerogenes 22.2 88.9 100
Sarcina sp 66.7 88.9 77.8
Flavobacterium breve 33.3 66.7 88.9
Nodularia 11.1 100 100
Erwina amylovora 44.4 66.7 88.9

N = Number of sample

Table 3: Screening test for PAH utilization by bacterial isolates.

Organism Incubation (Days)

 7 14 21 28

Vibrio alginolyticus + + ++ ++
Pseudomonas aeruginosa + ++ ++ +++
Vibrio estuarianus + ++ ++ ++
P. putida + +++ +++ +++
Alcaligenes denitrificans + +++ ++ +++
Chromobacterium violaceum + + ++ ++
Nocardia sp + ++ +++ +++
Micrococcus varians + +++ +++ +++
Acinetobacter iwoffii + +++ +++ +++
Serratia marcescens - - + +
Bacillus subtilis + +++ +++ +++
Chromatium sp - - + +
Escherichia coli - - - +
Anabaena + +++ +++ +++
Enterobacter aerogenes - - - +
Sarcina sp + + ++ ++
Flavobacterium breve + + ++ ++
Nodularia + +++ +++ +++
Erwina amylovora - - + +
Control - - - -

+++ = Profuse growth, ++ = Moderate growth, + = Scanty growth,
- = No growth

Table 4: Polycyclic Aromatic Hydrocarbon levels in the samples.

Parameters Concentration (mg/kg)

 Benthic Sediment

 L1 L2 L3

Naphthalene 1.00 1.00 1.00
2-Methylnaphthalene 0.20 0.20 0.20
Acenaphthylene 1.00 1.00 1.00
Acenaphthene 0.70 0.70 0.70
Fluorene 0.70 0.70 0.70
Phenanthrene 0.20 0.27 0.20
Anthracene 0.40 0.40 0.40
Fluoranthene 0.20 0.48 0.20
Pyrene 0.20 0.46 0.20
Benzo(a)anthracene 0.33 0.53 0.56
Chrysene 0.40 0.40 0.75
Benzo(b)fluoranthene 0.74 0.58 0.91
Benzo(k)fluoranthene 0.25 0.37 0.46
Benzo(a)pyrene 0.20 0.39 0.31
Dibenzo(a,h)anthracene 0.47 0.20 0.59
Benzo(g,h,l)perylene 0.48 0.20 0.20
Indeno(1,2,3-d)pyrene 1.53 1.04 1.17
Total PAH 9.00 8.92 9.55

Parameters Concentration (mg/kg)


 L1 L2 L3

Naphthalene 1.00 1.00 1.00
2-Methylnaphthalene 0.39 0.23 0.23
Acenaphthylene 1.00 1.00 1.00
Acenaphthene 0.70 0.70 0.70
Fluorene 0.83 0.81 0.78
Phenanthrene 2.19 1.76 1.04
Anthracene 0.46 0.42 0.47
Fluoranthene 0.65 0.53 0.58
Pyrene 0.20 0.26 0.32
Benzo(a)anthracene 0.69 0.53 0.63
Chrysene 1.89 1.42 1.54
Benzo(b)fluoranthene 0.33 0.30 0.35
Benzo(k)fluoranthene 1.39 1.16 1.21
Benzo(a)pyrene 5.52 4.89 4.27
Dibenzo(a,h)anthracene 2.87 2.07 2.42
Benzo(g,h,l)perylene 3.57 1.09 1.02
Indeno(1,2,3-d)pyrene 1.53 1.32 1.09
Total PAH 25.21 19.49 18.65

Parameters Concentration (mg/kg)


 L1 L2 L3

Naphthalene 1.00 10.. 1.00
2-Methylnaphthalene 0.20 0.20 0.20
Acenaphthylene 1.00 1.00 1.00
Acenaphthene 0.70 0.70 0.70
Fluorene 0.70 0.70 0.70
Phenanthrene 0.40 0.25 0.20
Anthracene 0.20 0.36 0.39
Fluoranthene 0.20 0.20 0.20
Pyrene 0.64 0.45 0.29
Benzo(a)anthracene 0.47 0.43 0.41
Chrysene 0.37 0.57 0.39
Benzo(b)fluoranthene 0.86 0.73 0.76
Benzo(k)fluoranthene 0.53 0.59 0.54
Benzo(a)pyrene 1.17 1.09 1.03
Dibenzo(a,h)anthracene 0.58 0.54 0.52
Benzo(g,h,l)perylene 0.61 0.66 0.61
Indeno(1,2,3-d)pyrene 1.50 1.03 0.98
Total PAH 11.13 10.50 9.92

Parameters Concentration (mg/kg)


 L1 L2 L3

Naphthalene 1.00 1.00 1.00
2-Methylnaphthalene 0.20 0.20 0.20
Acenaphthylene 1.00 1.00 1.00
Acenaphthene 0.70 0.70 0.70
Fluorene 0.7 0.70 0.70
Phenanthrene 0.20 0.23 0.20
Anthracene 0.40 0.25 0.22
Fluoranthene 0.20 0.20 0.20
Pyrene 0.20 0.27 0.23
Benzo(a)anthracene 0.43 0.34 0.30
Chrysene 0.22 0.22 0.22
Benzo(b)fluoranthene 0.49 0.45 0.47
Benzo(k)fluoranthene 0.74 0.73 0.74
Benzo(a)pyrene 0.43 0.46 0.44
Dibenzo(a,h)anthracene 0.20 0.24 0.21
Benzo(g,h,l)perylene 0.20 0.20 0.20
Indeno(1,2,3-d)pyrene 0.20 0.26 0.23
Total PAH 7.51 7.45 7.26

L = Location

Table 5: Bioaccumulation Factor (BAF) of PAHs in Rhizophora racemosa.

Parameter BAF (Sediment--Root)

 L1 L2 L3

Naphthalene 1.00 1.00 1.00
2-Methylnaphthalene 1.00 1.00 1.00
Acenaphthylene 1.00 1.00 1.00
Acenaphthene 1.00 1.00 1.00
Fluorene 1.00 1.00 1.00
Phenanthrene 2.00 * 0.93 1.00
Anthracene 0.50 0.90 0.98
Fluoranthene 1.00 0.42 1.00
Pyrene 3.20 * 0.98 1.45 *
Benzo(a)anthracene 1.42 * 0.81 0.73
Chrysene 0.93 1.43 * 0.47
Benzo(b)fluoranthene 1.16 * 1.26 * 0.84
Benzo(k)fluoranthene 2.12 * 1.59 * 1.17 *
Benzo(a)pyrene 5.85 * 2.79 * 3.32 *
Dibenzo(a,h)anthracene 1.23 * 2.70 * 0.88
Benzo(g,h,l)perylene 1.27 * 3.30 * 3.05 *
Indeno(1,2,3-d)pyrene 0.98 0.99 0.83
Total PAH 1.24 1.18 1.04

* = Biomagnification of individual PAH may be occurring.
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Author:Eduok, Samuel I.; Ita, Basil N.; Uye, Uwa O.; Utuk, Nkoyo M.
Publication:International Journal of Applied Environmental Sciences
Date:Dec 1, 2010
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