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Journal of Environmental Management 162 (2015) 30e36

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Journal of Environmental Management

journal homepage: www.elsevier .com/locate/ jenvman

Research article

Biosurfactant from red ash trees enhances the bioremediation of PAHcontaminated soil at a former gasworks site

Warren Blyth, Esmaeil Shahsavari*, Paul D. Morrison, Andrew S. BallCentre for Environmental Sustainability and Remediation, School of Applied Sciences, RMIT University, Bundoora, Victoria 3083, Australia

a r t i c l e i n f o

Article history:Received 9 May 2015Received in revised form15 July 2015Accepted 17 July 2015Available online xxx

Keywords:Polycyclic aromatic hydrocarbonsBioremediationBiosurfactantRed ash treeDGGEqPCR

* Corresponding author.E-mail address: esmaeil.shahsavari@rmit.edu.au (E

http://dx.doi.org/10.1016/j.jenvman.2015.07.0410301-4797/© 2015 Elsevier Ltd. All rights reserved.

a b s t r a c t

Polycyclic aromatic hydrocarbons (PAHs) are persistent contaminants that accumulate in soil, sludge andon vegetation and are produced through activities such as coal burning, wood combustion and in the useof transport vehicles. Naturally occurring surfactants have been known to enhance PAH-removal fromsoil by improving PAH solubilization thereby increasing PAHemicrobe interactions. The aim of thisresearch was to determine if a biosurfactant derived from the leaves of the Australian red ash (Alphitoniaexcelsa) would enhance bioremediation of a heavily PAH-contaminated soil and to determine how themicrobial community was affected. Results of GC-MS analysis show that the extracted biosurfactant wassignificantly more efficient than the control in regards to the degradation of total 16 US EPA priority PAHs(78.7% degradation compared to 62.0%) and total petroleum hydrocarbons (TPH) (92.9% degradationcompared to 44.3%). Furthermore the quantification of bacterial genes by qPCR analysis showed thatthere was an increase in the number of gene copies associated with Gram positive PAH-degradingbacteria. The results suggest a commercial potential for the use of the Australian red ash tree as asource of biosurfactant for use in the accelerated degradation of hydrocarbons.

© 2015 Elsevier Ltd. All rights reserved.

1. Introduction

Polycyclic aromatic hydrocarbons (PAHs) are significant organiccontaminants that typically have a structure that is composed of2e6 aromatic rings (Sun et al., 1998). PAHs are produced by oilrefineries and arise as a result of the incomplete combustion oforganic compounds from activities such as coal burning, wasteincineration, wood combustion, aluminum production and trans-portation (Essumang et al., 2012). While there are potentiallyhundreds of PAHs, the US EPA has a priority list of the 16 chemicalsthat pose the most concern to the environment (Ball andTruskewycz, 2013).

There is mounting evidence that suggests that the exposure toPAHs is detrimental to human health. Research has shown that PAHexposure has been linked to obesity, cancer, lower IQ, inhibitedgrowth, endocrine disruption and diabetes (Anyakora et al., 2004).The two most common points of entry into the body are inhalationof airborne particles and the ingestion of contaminated food. PAHsare highly hydrophobic and this determines the most significant

. Shahsavari).

physical and chemical aspects of their interactions with the envi-ronment. Irigaray et al. (2006) showed that benzo[a]pyreneimpaired adipose (fat) tissue lipolysis which led to weight gain andincreased appetite in mice. Interestingly, after the withdrawal ofbenzo[a]pyrene, the mice retained their elevated weight (Irigarayet al., 2006).

The historical methods for the removal of PAHs and other suchorganic compounds include chemical oxidation, thermal treatmentand solvent washing. PAHs have been successfully been ‘washed’from contaminated soils with solvents including synthetic surfac-tants (Lau et al., 2014), peanut oil, vegetable oils (Gong et al., 2005)and cyclodextrins (Viglianti et al., 2006). These techniques havebeen shown to be satisfactory; however, they tend to require sig-nificant post-removal clean-up and in the case of thermal treat-ment the soil is potentially left with limited nutrients and devoid ofmicrobial life (Gan et al., 2009). The removal of PAHs from soils inparticular poses multiple challenges. When designing a remedia-tion strategy the following variables must be considered: soil pH,salinity, temperature, nutrient level, soil type, the nature of indig-enous microorganisms, the effect on the surrounding environment,in-situ or ex-situ processing and the cost (Ferradji et al., 2014).

In recent times, bioremediation has been used to ‘naturally’clean up contaminated soil. These processes include

W. Blyth et al. / Journal of Environmental Management 162 (2015) 30e36 31

biostimulation, bioaugmentation, phytoremediation or a combi-nation of two methods (e.g. phytoremediation in conjunction withbiostimulation). In bioremediation, plants (Cofield et al., 2008;Huang et al., 2004; Liste and Alexander, 2000; Rezek et al., 2008),bacteria (Li et al., 2008; Mao et al., 2012) and fungi (Cajthaml et al.,2002; Leonardi et al., 2007; Sasek et al., 2003) have all been widelyused to enhance the removal of contaminants from soil, water orsludge. The advantages of these methods are that they are lesslikely to be environmentally damaging, they are more successful insitu and cost-wise they compete well with traditional approaches(Ortega-Calvo et al., 2013).

Numerous studies have confirmed that certain microorganismsare capable of using PAHs as ‘food’ by oxidizing the aromatic ringsvia ring-hydroxylating dioxygenase (RHD) enzymes. However,because of their strongly hydrophobic nature and low water solu-bility, PAHs are known to adhere to the small pockets in betweensoil particles which can result in strong absorption to clays, organicmatter, coal tar and soot (Niqui-Arroyo et al., 2006). This reducesthe bioavailability of the contaminants to breakdown by PAH-degrading organisms. A recent method for improving bioavail-ability that researchers have used is washing a surfactant throughthe contaminated soil. Surfactants are compounds that areamphiphilic in nature. This means that their structure has bothhydrophobic and hydrophilic (polar) aspects (Delgado-Balbuenaet al., 2013). The effectiveness of a surfactant can be determinedby its ability to lower surface tension and form micelles with thehydrophobic part concentrated at the surface and the hydrophilicportion oriented towards the solution (Mulligan, 2004). Surfactantsmay be synthetic or naturally occurring (biosurfactants) and havebeen used to promote solubilization, emulsification and wetting ofsoils with the desired outcome being the removal of recalcitrantcontaminants (Delgado-Balbuena et al., 2013).

The problem with synthetic biosurfactants is that unwanted(toxic) by-products may be incompatible with naturally biologicalsystems (Urum et al., 2006). Also, studies have shown that the useof some synthetic surfactants may actually result in the surfactantbeing adsorbed onto the soil matrix with the PAH attached whichthen increases sorption of PAHs into the soil rather than removal(Zhou and Zhu, 2007).

The use of biosurfactants has increased because they tend tohave advantages such as retention of high activity at extremes ofpH, temperature and salinity, low operating costs and the fact theyare readily biodegradable (Urum et al., 2006). In many cases bio-surfactants derived from microorganisms have been used toremediate soil contaminated with PAHs (Cui et al., 2008; Husain,2008; Pei et al., 2010). These include microorganisms such asPseudomonas aeruginosa (rhamnolipids), Bacillus subtilis (surfac-tins) and Candida bombicola (sophorolipids) that secrete bio-surfactants which enable these microbes to metabolize certainhydrocarbons by reducing the surface tension at the oil/waterinterface (Mulligan, 2004).

However using such compounds derived from plants is lesscommon. Maity et al. (2013) reported that the saponin isolatedfrom Soapberry (Sapindus mukorissi) enhanced the removal ofnickel and chromium from contaminated soil (Maity et al., 2013). Inanother study, Zhou et al. (2011) found that saponin had a highermolar solubilization ratio (MSR) than synthetic (Brij 58, Triton X-100, Tween 80) and naturally occurring (rhamnolipids and dir-hamnolipids) surfactants when their phenanthrene-solubilizingproperties were measured (Zhou et al., 2011). Furthermore,studies by Kobayashi et al. (2012) indicated that aqueous saponinprepared from Quillaja bark can significantly enhance the degra-dation of pyrene spiked soils. Regardless of concentration(0.025e0.5 g/l) there was a 40e60% range of pyrene degradation inonly 60 h, significantly greater reduction relative to the control soil

(27%) (Kobayashi et al., 2012).In tropical regions of Australia, Polynesia and Southeast Asia,

Alphitonia excelsa, commonly known as the red ash, soap tree or thesilver leaf is common. The indigenous peoples of Australia haveused its berries to stun fish and as a headache remedy. They havealso produced a soap/foam by removing leaves from the tree andrubbing them with water between the hands. The biosurfactantreleased from red ash leaves has been identified as a triterpenoidsaponin and its PAH-solubilization properties are of significant in-terest (Branch et al., 1972).

The aims of this research are to (i) investigate whether thebiosurfactant derived from red ash leaves will enhance or inhibitPAH-degradation of heavily contaminated soil and (ii) to under-stand how the extracted biosurfactant affects the indigenous mi-crobial community.

2. Methods

2.1. Experimental set up

Heavily contaminated soil was obtained from the Environ-mental and Earth Sciences group (ESSI) in Cootamundra, NewSouth Wales, Australia. The contaminated soil originated from aformer gas production plant. The initial contaminated soil (~7 kg)was passed through a 6 mm sieve to remove stones, large soilparticles and other debris. A sample (300 g) of the contaminatedsoil was analyzed by SESL Australia Pty Ltd for analysis of totalcarbon, nitrogen and phosphorus. Other soil parameters weredetermined using standard methods (Rayment and Higginson,1992). Physio-chemical characteristics of the soil are presented inSupplementary Table 1.

2.2. Preparation of surfactant

Leaves of red ash treewere donated from a specimen that growsin the Royal Botanic Gardens in Melbourne, Victoria. Upon removalfrom the tree the leaves were prepared by macerating in a foodmixer for 60 s on high and then further breaking down with amortar and pestle (5 min) to release the biosurfactant. The sur-factant solution for the treatment was made using the maceratedred ash leaves. A concentration of 25 g of leaves per kg of soil waschosen based on initial foaming experiments, the practical amountof leaves available and values obtained in the literature (Kobayashiet al., 2012). The solution of biosurfactant was prepared by mixingmacerated leaves (25 g) with milliQ water (500 mL) (50 g/L). Thissolutionwas then agitated at 150 rpm for 15 min and the liquid wasthen separated from the leaves via filtration. For each treatment, analiquot of the 50 g/L solution was added to each pot, giving a finalconcentration of 25 g/kg of soil.

2.3. Analysis of PAHs and total petroleum hydrocarbon (TPH) in soilsamples

Hydrocarbons in soil were extracted with hexane as describedby Mansur et al. (2014). The soil extracts taken from weeks 0, 4, 8and 12 for each treatment were subjected to GC-MS analysis. Usingan Agilent 6890 GC, an aliquot (1 mL) was injected into a capillarycolumn with a length of 30 m, a diameter of 250 mm and filmthickness of 0.25 mm. Heliumwas used as the carrier gas (3mL/min)and the injection temperature was 325 �C. Similarly, TPH (C10eC40)was determined as previously described (Makadia et al., 2011;Mansur et al., 2014).

Fig. 1. PAH (a) and TPH (b) levels (%) remaining in the amended with surfactant andcontrol soils over 12 weeks of experiment (mean ± SE, n ¼ 3). * indicates a significantlydifferent value compared to the control soil at each time point (P < 0.05).

W. Blyth et al. / Journal of Environmental Management 162 (2015) 30e3632

2.4. Culturable microbial abundance

At each time point and at the beginning of the experiment theabundance of culturable bacteria and fungi were quantified innutrient agar and Czapek Dox agar, respectively, using a plate counttechnique (Foght and Aislabie, 2005).

PAH-degrading bacterial abundance was quantified by platingappropriate soil dilutions onto Bushnell Haas agar using a pyreneand phenanthrene spraying method as described by Foght andAislabie (2005).

2.5. Quantification of total bacteria and PAH-ring hydroxylatingdioxygenase genes

DNA from soil samples was extracted using a MoBio Powersoil®

DNA Isolation Kit according to the manufacturer's guideline atselected time points. Once the extraction steps were complete, theindividual tubes continuing DNA were stored at �20 �C for lateruse.

To evaluate the effects of biosurfactant on total bacterial andalso PAH-ring hydroxylating dioxygenase genes (PAH-RHDa, i.e.genes involved in the degradation of PAHs), the DNA for soil sam-ples from treated and control soil at each point time was subjectedto Real-timeeqPCR quantification using a QIAGEN Rotor-Genemachine. For quantification of total bacteria, universal primers341F and 518R (Sch€afer and Muyzer, 2001) were used to amplify16S rRNA genes. PAH-RHDa genes from Gram positive (PAH-RHDaGP) and negative bacteria (PAH-RHDa GN) were quantified usingprimer sets developed by C�ebron et al. (2008). In brief, for eachqPCR reaction a total volume of 20 mL was used in each tube con-sisting of molecular-biology-grade water (8.2 mL), Kapa SYBR FastqPCR Master Mix (10 mL, �2, Kapa Biosystems), forward primer(0.4 mL, 10 pmol/mL), reverse primer (0.4 mL, 10 pmol/mL) and DNAsample (1 mL). DNA (1 mL) was obtained from 1/10 dilutions of theoriginal DNA extractions performed earlier. Real-time-qPCRamplification conditions for bacteria were an initial denaturationstep at 95 (5min) followed by 40 cycles of 95 �C denaturation (10 s),annealing at 55 �C (30 s), 72 �C extension (30 s), 80 �C primer dimerremoval and signal acquisition (10 s) (Shahsavari et al., 2013). ForPAH-RHDa genes, the cycling conditions consisted of 40 cycles of95 �C denaturation (30 s), annealing at 54 �C for PAH-RHDa GP or57 �C for PAH-RHDa GN (30 s), 72 �C extension (30 s), 80 �C primerdimer removal and signal acquisition (10 s) as described by C�ebronet al. (2013).

2.6. Microbial community analysis

To assess total bacterial community, PCR amplification of 16SrDNA gene was performed with universal primer 341F-GC and907R according to the method of Sch€afer and Muyzer (2001). Thethermocycling program consisted of one cycle of 5 min at 95 �C, 33cycles of 30 s at 94 �C, 30 s at 52 �C, 1 min at 72 �C and a finalextension at 72 �C for 10 min.

For fungi, Internal Transcribed Spacer regions (ITS) amplificationof soil DNA extracts was carried out via a nested reaction withITS1F- ITS4 and ITS1F-GC primer sets as previously described(Anderson and Parkin, 2007). The first reactionwas carried out withITS1F and ITS4 (without a GC clamp). The second reaction wascarried out with GC primer ITS1F GC and ITS2. Product from thefirst PCR (2 mL) was used as template DNA in the second PCR. Theprogram for fungal amplifications was 1 cycle of 5 min at 95 �C, 35cycles of 45 s at 94 �C, 45 s at 58 �C and 45 s at 72 �C and a finalextension at 72 �C for 10 min. Agarose gel electrophoresis wasperformed to confirm DNA amplification after performing PCR.

DGGE was carried out on selected PCR amplicons on a Universal

Mutation Detection System D-code apparatus (Biorad, CA, USA)with a 6% polyacrylamide gel using a denaturing gradient of40e60% for bacteria and 42e52% for fungi, with gels runs at 60 �Cfor 18 h. Upon completion of electrophoresis, the gels were silverstained as described elsewhere (Girvan et al., 2003) and saved asTiff files with an Epson V700 scanner. The images were thenanalyzedwith Phoretix 1D software and dendrogramswere createdto represent the change in the microbial community over thecourse of the experiment.

2.7. Data analysis

Statistical analyses were carried out using XLStat 2014 software(Addinsoft). The data were subjected to using one way analysis ofvariance (ANOVA) followed by Tukey test or T test (P < 0.05). Datafrom plate counting and qPCR were log transformed prior toanalysis.

3. Results and discussion

3.1. Effects of extracted biosurfactant on degradation of PAHs andTPH

Examination of the amended soil in this experiment revealedthat the initial total concentration of the 16 US EPA PAHs was2,525 mg/kg. This compares to other studies where the initialconcentration of PAHs was 364 mg/kg (Sun et al., 2012) and1279e1369 mg/kg (Usman et al., 2012). This suggests that the testsoil was heavily contaminated.

Fig. 1 illustrates the changes in the concentration of total PAHsand TPH during the 12 week incubation. The results showed that

Fig. 2. Percentage remaining of 3, 4 and 5-ringed PAHs of soil under the influence ofthe surfactant treatment compared to the control after 12 weeks treatment(mean ± SE, n ¼ 3). * indicates significantly different values to control soil in eachdifferent ring PAH (P < 0.05).

W. Blyth et al. / Journal of Environmental Management 162 (2015) 30e36 33

after only four weeks there was a marked decrease in the con-centration of total PAHs in the biosurfactant treatment compared tothe control (Fig. 1a). The PAH concentration in biosurfactantamended soil was reduced to 832.5 mg/kg (67% reduction) afteronly 4 weeks compared with a 52% reduction in the control soil(after 8 weeks). After 12weeks further biodegradation of total PAHsfor both the control (62%) and the surfactant treatment (79%) wasobserved. This is typical of remediation experiments such as theone conducted by Contreras-Ramos et al. (2008) which focused on

Fig. 3. Microbial population of soil amended with surfactant and control soils; totalbacteria (a), PAH-degrading bacteria (b) and fungi (c) (mean ± SE, n ¼ 3). * indicates asignificantly different value compared to the control soil at each time point (P < 0.05).

soil, sludge and vermicompost spiked with anthracene, phenan-threne and benzo[a]pyrene. The researchers found that the con-centration of the first two contaminants decreased substantially inthe first 7 days of the experiment (Contreras-Ramos et al., 2008). Interms of TPH (Fig. 1b), there was a significant level of degradationmeasured after four weeks for the biosurfactant treatment, (16.2%,4385 mg/kg) while an increase in TPH concentration was observedin the control. This is likely due to the fact that TPH by-productsmay be present following the partial degradation of high molecu-lar weight molecules (above C40). The final concentrations of TPHwere reduced by 44% for the control (16,626mg/kg) and 93% for theextracted biosurfactant (2119 mg/kg) after 12 weeks. In both casesthere was a significant difference between the control and surfac-tant for total PAHs and TPH.

3.2. Effect of extracted biosurfactant on 3, 4 and 5-ringed PAHs

The percentage reduction in 3, 4 and 5-ringed PAHs for bothcontrol and biosurfactant wash treatment is compared in Fig. 2.There were reductions of 75 (370.7 mg/kg), 76 (1073.0 mg/kg) and85 (304.5 mg/kg) percent for 3, 4 and 5-ringed PAHs respectively inthe biosurfactant treatment compared with 42 (208.8 mg/kg), 60(874.1 mg/kg) and 82 (293.7 mg/kg) percent for the control. Theresults showed that the addition of biosurfactant led to enhanceddegradation of 3 and 4-ringed PAHs. However the difference be-tween the control and biosurfactant treatment was not significantin terms of 5-ringed PAHs. These results correlate to similar out-comes when coal tar-spiked soil was phyto-remediated with a

Fig. 4. The number of gene copies of total bacteria (a), PAH-RHDa GN (b) and PAH-RHDa GP (c) bacteria per g dry soil in soil amended with surfactant and control soil(mean ± SE, n ¼ 3). * indicates a significantly different value compared to the controlsoil at each time point (P < 0.05).

W. Blyth et al. / Journal of Environmental Management 162 (2015) 30e3634

variety of plants. A study by Smith et al. (2011) showed that thedegradation of 2e3 ringed PAHs (78e100%) was significantlyhigher than 4-ringed (36e85%) and 5e6 ringed (8e33%) PAHs(Smith et al., 2011).

One of the main issues with the biodegradation of PAHs and alsoTPH is bioavailability of these compounds. Petroleum hydrocarboncompounds, especially PAHs tend to absorb to soil particles and inthis case, even if the microbial communities have the potential todegrade PAHs, hydrocarbon degraders cannot use the PAHs as en-ergy and food source. Surfactants are promising agents forincreasing the bioavailability of petroleum hydrocarbon-by prod-ucts. It has been shown that addition of different synthetic sur-factants had a positive impact on the degradation of petroleumhydrocarbons including PAHs (Adetutu et al., 2012; Hultgren et al.,2009; Zhang et al., 2010). Research by Franzetti et al. (2008)compared the diesel-degradation abilities of the ‘Brij’ family ofsynthetic surfactants to the ‘Tween’ family and found that in

Fig. 5. Cluster analysis using UPGMA method of bacterial (a) and fungal (b) communities insimilarity).

conjunctionwith a liquid bacterial culture the residual hydrocarbonpercentages were 36 (Brij), 9 (Tween) and 37 (control) after 7 daysincubation.

In regard to biosurfactant-assisted remediation there arenumerous research papers addressing the treatment of soilcontaminated with a range of organic pollutants. Research by Cuiet al. (2008) found that a rhamnolipid biosurfactant isolated fromSphingomonas sp. 12A and Pseudomonas sp. 12B increased the sol-ubility of anthracene significantly compared to the control. After 18days of treatment the percentage solubilization was determined as39 (Sphingomonas sp. 12A), 52 (Pseudomonas sp. 12B) and 34(control). Pei et al. (2010) compared the level of phenanthrenedegradation by Sphingomonas sp. under the influence of addedbiosurfactant or synthetic surfactant (Tween-80). The researchersconcluded that after 10 days treatment there was 83.6% minerali-zation of phenanthrene with the biosurfactant and only 33.5% withthe Tween-80.

soil amended with surfactant and control soils through weeks 0, 4, 8 and 12 (scale is

W. Blyth et al. / Journal of Environmental Management 162 (2015) 30e36 35

3.3. Effect of extracted biosurfactant on microbial abundance

To determine whether the differences in TPH and PAH degra-dation over the 12 weeks between the two treatments reflectedincreased microbial biomass, viable microbial counts in the soilduring treatment were undertaken (Fig. 3). The results showed thatthe addition of surfactant to the soil increased the number of mi-croorganisms present after 12 weeks incubation. The heterotrophicbacteria (Fig. 3a) and PAH-degrading bacteria (Fig. 3b) showed asignificant increase in the number of colony forming units (CFU) insoils amended with the biosurfactant treatment compared to thecontrol while fungal abundance (Fig. 3c) did not show any signifi-cant differences.

Molecular analysis of the soil microbial community began withqPCR analysis for total bacteria (16S rDNA), Gram negative PAH-degrading bacteria and Gram positive PAH-degrading bacteria(Fig. 4aec). Real time-qPCR revealed that gene copies of 16S rDNAand PAH-RHDa of Gram positive bacteria were higher than control.For example, by week 12, gene copies of 16S rDNA and PAH-RHDaGram positive bacteria in the biosurfactant treatment showed 42and 6-fold increases relative to control soil. In contrast there was a5-fold decrease in gene copies associated with Gram negative PAH-degrading bacteria during the biosurfactant treatment incubation.C�ebron et al. (2008) showed that Gram-positive and negativebacteria initiate PAH metabolism with a similar RHD-enzyme butwith different substrate specificity. The genes sequences coding thealpha subunit of this enzyme are broadly grouped into those Grampositive and negative bacterial groups. This may explain the dif-ference in numbers of gene copies that were found to be present inthis study (C�ebron et al., 2008).

3.4. Microbial community structure

Having established that significant changes in the total numberof bacteria were occurring in the presence of the biosurfactant,community fingerprinting was undertaken using PCR-DGGE toassess the impact of the treatment of community diversity. Thediversity of the bacterial community (Fig. 5a) showed that the di-versity of bacteria in the control soil had close similarity at differenttime points suggesting that the predominant bacteria early in theincubation remained dominant throughout the experiment. Thesamples associated with the biosurfactant wash were closelygrouped together and separated from the control, confirming thatthe addition of surfactant extract to PAH-contaminated soil causedsome changes in bacterial community. However, cluster analysis offungi (Fig. 5b) showed that the changes in the fungal communitywere less than bacteria in soil amended with biosurfactant extract.

4. Conclusion

This research project sought to investigate whether the bio-surfactant contained within the leaves of the red ash wouldenhance or inhibit the biodegradation of soil contaminated withPAHs. Overall, the results indicate there was an enhanced reductionin the concentration of PAHs in heavily contaminated soil amendedwith extracted biosurfactant from red ash leaves compared to thecontrol as well as TPH. There may be a number of reasons for thisobservation including increased solubilization of PAHs and theirsubsequent increased bioavailability to PAH-degradingmicrobes. Inthe presence of biosurfactant, the microbial community waschanged in some instances and the abundance of bacteria increasedfollowing the addition of the biosurfactant extract. Also, it wasshown that the biosurfactant initiated increased growth of Grampositive PAH-degrading bacteria which can be linked with signifi-cant reduction in total PAHs and TPH concentrations. The findings

from this study suggest a possible commercial application of redash leaves as a source of biosurfactant for use in the bioremediationof PAH contaminated soils. This application provides an eco-friendly, sustainable, biodegradable and low cost alternative tothe use of chemical surfactants.

Acknowledgments

This work was supported by the award of a grant from theAustralian Research Council to AS Ball (Grant No. LP110201130).

Appendix A. Supplementary data

Supplementary data related to this article can be found at http://dx.doi.org/10.1016/j.jenvman.2015.07.041.

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