As inferred by 16S rRNA gene analysis, strain S5 showed maximum sequence identity (100%) with corresponding gene sequences of the reference strain A. aegrifaciens LMG 26852 (NCBI Accession No.NR_117707). Thus, strain S5 was identified as A. aegrifaciens.
Unrooted phylogenetic tree (Fig. 1) based on comparison of 16S rRNA gene sequences demonstrating the evolutionary relationships between strain S5 and relating species. The GenBank accession number for each microbe used in the analysis is shown in parentheses after the species name. Bootstrap values (%) are indicated at the nodes. The evolutionary history was inferred using the Neighbor-Joining method (Saitou and Nei1987).The optimal tree with the sum of branch length = 0.26080621 is shown. The percentage of replicate trees in which the associated taxa clustered together in the bootstrap test (10000 replicates) is shown next to the branches (Felsenstein1985). The evolutionary distances were computed using the Maximum Composite Likelihood method (Tamura et al. 2004) and are in the units of the number of base substitutions per site. This analysis involved 12 nucleotide sequences. Codon positions included were 1st + 2nd + 3rd + Noncoding. All positions containing gaps and missing data were eliminated (complete deletion option). There were a total of 441 positions in the final dataset. Evolutionary analyses were conducted in MEGA X (Kumar et al. 2018).
3.1.2. Biofilm forming capacity and motility
In 2004, Johnsen and Karlson reported that biofilms increased the solubility of PAHs and subsequently their mass transfer from recalcitrant crystals to cells for biotransformation. In the present study, we are interested in evaluating the biofilm forming capacity of A. aegrifaciens.
The ability of A. aegrifaciens to produce biofilm on polystyrene microplate was tested under two temperatures (20 and 30°C) and two incubation times (24 and 48 hours). We found that the examined strain was able to produce biofilm at both tested conditions.The results of the optical density at 550 nm presented in (Fig. 2a), showed a significant(p < 0.001) increase in biofilm biomass by 59% at 20°C compared to 30°C. This proves that biofilm production among A. aegrifaciens is temperature dependent. This result agrees with previous studies, Ben Miloud Yahia et al. (2018) showed that the ability of Salmonella Hadar to produce biofilm could be reduced by lowering the temperature. In this context, Pinel et al. (2021) suggested that heat induces a change in biofilm bacterial community and EPS composition caused by a relatively lower extracellular sugar production at elevated temperatures. Furthermore, results showed that the amount of biofilm was strongly reduced by 47%, after incubation for 48 h (Fig. 2b), due to the dispersion of biofilm normally occurring in the stationary phase and enhanced with nutrient starvation (Gjermansen et al. 2010).
As shown on (Fig. 2c), A. aegrifaciens can also form unattached floating biofilm at air-liquid (A-L) interface, which is generally referred to as “pellicle”. The earliest experimental observations of these were probably made for Bacterium aceti and Bacterium xylinum in 1886 (Brown 1886). Most bacteria when propagated in static liquid culture grow within the broth phase or quickly sediment to the bottom. There are few bacteria reported to be capable of colonizing the air-liquid (A-L) interface (Spiers et al. 2003).
To determine the exopolysaccharides (EPS) production in isolated strain, the Congo red assay was carried out. As shown in (Fig. 2d), the culture of A. aegrifaciens, produced an amount of CR-binding material that can bind approximately 68% of the initial amount of CR. Ghafoor et al. (2011) have shown that the EPS overproduction led to an increased Congo red binding.
It was shown that Achromobacter spp. can form biofilm on abiotic surfaces such as contact lens, central venous catheters, and urinary catheters (Kim et al. 2008; Choe et al. 2012; Konstantinovic et al. 2017). As well, Achromobacter species, recognized as emerging pathogens isolated from patients with cystic fibrosis, can form biofilm in the respiratory tract in patients (Nielsen et al. 2016;Damar-Celik et al. 2021).
As bacterial motility could contribute to survival and initiate the first phase of biofilm colonization, we investigated the swarming, swimming, and twitching motility of A. aegrifaciens after 24- and 48hours incubation at 30°C.For the swarming (a flagella-driven movement), the zone of motilitywas6 ± 0 mm at 24 h and 9.7 ± 1.5mm at 48 h. The swimming zone was 12.67 ± 3.1 mm at 24 h and 30.7 ± 5,1mm at 48 h. The twitching (a type IV pili-dependent motility) was 6.7 ± 2.9 mm and 9.3 ± 2.3 mm after 24 and 48 h, respectively.
Bacterial motility is often crucial for diverse processes such as biofilm formation. Ditmarsch et al., (2013) have found that evolution in swarming colonies reliably produces evolution of poor biofilm formers which supports the existence of an evolutionary trade-off between motility and biofilm formation. It was also reported that swimming motility is an important factor in the formation of biofilm as it contributes to the early development of the biofilm architecture (Srey et al. 2013). However, Nielsen et al. (2019) have found that inactivation of the flagellar M-ring protein, FliF, in A. xylosoxidans resulted in a 39% reduction of in vitro biofilm formation on peg-lids. Moreover, Khademi et al. (2021) showed that the swimming motility of A. xylosoxidans is gradually lost over time following the initial colonization.
3.1.3. Screening of Biosurfactant production
Production of the biosurfactant serves as an essential aid to swarming motility by acting as a wetting agent to overcome the surface tension of water and facilitate movement across the moist surface (Van Alst et al. 2007). In addition, it enhances the bioavailability, solubilization, and biodegradation of hydrophobic pollutants during the degradation of insoluble petroleum hydrocarbons (Tian et al. 2016).
Figure 3 shows the results of oil displacement test of the strain A. aegrifaciens grown on basal medium containing motor oil (1%, v/v)as the sole carbon and energy source, at 120 rpm and 30°C for4 days. Halo zones were observed with diameters over10 mm, suggesting a biosurfactant synthesis (p < 0.05).
Emulsifying ability of the cell-free culture broth, determined as emulsification index (E24), was tested with variety of hydrophobic compounds including mineral oil, kerosen and olive oil. Emulsion formation degrees by biosurfactants against olive oil (53%) were higher than those of kerosene and mineral oil0.35% and 11%, respectively (p < 0.0001) (Fig. 4).
Recently, more attention has been focused on the important environmental value of biosurfactants produced by microorganisms (Subashchandrabose et al. 2019; Li et al. 2020). Achromobacter sp. LH-1 grown in MSM with PHE as the sole carbon source produce biodemulsifier achieved 95.6% demulsification efficiency for W/O model emulsions within 24 hours (Hou et al. 2018). A biosurfactant-producing strain Achromobacter AC15, isolated from mangrove soil, utilized pyrene as the sole carbon source, decreased the surface tension of the culture medium to 33.2 mN m− 1 after 6 days of cultivation (Li et al. 2020).
Biosurfactant-producing microorganisms, including bacteria and fungi, are important in solubilizing hydrophobic contaminants affording their ultimate degradation, which can be used to speed up the remediation of organic and metal-contaminated sites.4
3.2 Chrysene degradation assay
A. aegrifaciens was isolated from PAHs polluted seawater and incubated for 7 days with chrysene as the sole carbon source. Biodegradation of chrysene by the bacterial isolate was indicated based on an increase in the total viable count and confirmed by GC-MS analyses.
From (Fig. 5) a growth linked chrysene degradation pattern can be observed. Both growth and degradation rate increased gradually, with a maximum growth on the6th day over 25.1032 CFU/mL and a maximum degradation (86%) on day7. An authentic standard of chrysene was used, and the retention time was determined to be 17.40 minutes.
Since chrysene is a PAH of four fused benzene rings, it has a very poor solubility in water, which results reduce in bioavailability, and inhibits microbial utilization. A. aegrifaciens was compared with previous chrysene-degrading bacteria, such as Sphingomonas sp. which was able to degrade 97.5% ofchrysene from an initial concentration of 500 mg/L in 35 days (Willison 2004) ,while Achromobacter xylosoxidans utilized 56% of 50 mg/L chrysene within 15 days (Ghevariya et al. 2011). In addition, a consortium of Rhodococcus, Bacillus, and Burkholderia utilized 96% of 10 mg/L chrysene in 8 days(Vaidya et al.2018).
Furthermore, Al Farraj et al. (2019) reported that under optimum conditions, Hortaea sp. B15exhibited 77%chrysene degradation from an initial concentration of 100 mg/Lin 20 days. Recently, Bacillus halotolerans exhibited 90%of 100 mg/L chrysene degradation in 6 days, under optimum conditions (Thomas et al.2021).Therefore, this study demonstrated that A. aegrifaciens is an efficient chrysene-degrading bacterium, that utilized 86% of 10mg/L chrysene in 7 days.
Identification of metabolites
To identify the metabolites of chrysene degradation and determine the degradation pathway, GC-MS was used. The conditions for GC-MS consisted of the use of a scan-interval of 0.05 s, and a mass range of 40–450.Results showed that A. aegrifaciens, grown in BH broth amended with chrysene(10 mg/L)for 7 days, was able to degrade chrysene to Benzoic acid, 3,5-dicyclohexyl-4-hydroxy-, methyl ester by a major peak at 25.89 min and Phenol, 2,4-bis(1,1-dimethlethy) by a peak at 10.28 min.
Other intermediates detected in the spectrum include eicosane (RT 24.901), heptadecane, 9-octyl- (RT 23.88), tetracosane, 11-decyl- (RT 24.133) (Fig. 6).
A previous study has found out that Polyporus sp. S133 degraded chrysene via 1-hydroxy-2-naphthoic acid to phthalic acid and salicylic acid (Hadibarata et al. 2009). As well, Al Farraaj et al.(2019) revealed the metabolic products produced during the degradation of chrysene by Hortaea sp. B15 such as phthalic acid (RT13.9) and1-hydroxy-2-naphthoic acid (1H2NA) (RT 8.6). Recently, Thomas et al. (2021) established that during the chrysene biodegradation by Bacillus halotolerans, the main metabolite detected by GCMS was esters of phthalic acid, which had an Rf value of 25.107,other intermediates detected in the spectrum include eicosane (Rf-12.066), 2,4 dibutylphenol (Rf-12.46), isobutyl phthalate (Rf-17.403), dibutyl phthalate (Rf-18.63), heneicosane (Rf-19.121), eicosane,7-hexyl (Rf-21.444), heptadecane (Rf-22.531), eicosane,10-methyl (Rf-23.577), 2-methyloctacosane (Rf-24.658) and Tritetracontane (Rf-25.921).
Actually, according to the presence or absence of oxygen, there are two principal strategies to degrade PAHs. In the aerobic catabolism, the oxygen is the final electron acceptor and a co-substrate for the hydroxylation and oxygenolytic ring cleavage of the aromatic. Ring hydroxylation is the initial and rate-limiting step in PAH biodegradation and is carried out by di- or mono-oxygenases that produce metabolites with one or two –OH radicals (Wang et al. 2018). On the other hand, the anaerobic catabolism of aromatic compounds uses a quite different procedure based on reductive reactions (Ghosal et al. 2016).
However, little is known regarding the degradation mechanisms for PAHs by A. aegrifaciens. We have found that A. aegrifaciens is capable to transform chrysene into Benzoic acid, 3,5-dicyclohexyl-4-hydroxy-, methyl ester (Fig. 7). Therefore, the effective degradation of chrysene, and its metabolites, by strain A. aegrifaciens, suggests that it is a potential bacterium for the bioremediation of PAHs and other sites contaminated with hydrocarbon.