Bioresource Technology 102 (2011) 772–778

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Effect of biosurfactant and fertilizer on biodegradation of crude oil by marine isolates of Bacillus megaterium, Corynebacterium kutscheri and Pseudomonas aeruginosa Rengathavasi Thavasi a,⇑, Singaram Jayalakshmi b, Ibrahim M. Banat c a b c

Department of Chemical and Biological Sciences, Polytechnic Institute of New York University, 6 Metrotech Center, Brooklyn, New York 11201, USA CAS in Marine Biology, Annamalai University, Parangipettai 608 502, Tamil Nadu, India School of Biomedical Sciences, University of Ulster, Coleraine BT52 1SA, Northern Ireland, UK

a r t i c l e

i n f o

Article history: Received 20 June 2010 Received in revised form 25 August 2010 Accepted 26 August 2010 Available online 31 August 2010 Keywords: Biosurfactants Bioremediation Corynebacterium kutscheri Bacillus megaterium Pseudomonas aeruginosa

a b s t r a c t This study was conducted to investigate the effects of fertilizers and biosurfactants on biodegradation of crude oil by three marine bacterial isolates; Bacillus megaterium, Corynebacterium kutscheri and Pseudomonas aeruginosa. Five sets of experiments were carried out in shake flask and microcosm conditions with crude oil as follows: Set 1–only bacterial cells added (no fertilizer and biosurfactant), Set 2–with additional fertilizer only, Set 3–with additional biosurfactant only, Set 4–with added biosurfactant + fertilizer, Set 5–with no bacterial cells added (control), all the above experimental sets were incubated for 168 h. The biosurfactant + fertilizer added Set 4, resulted in maximum crude oil degradation within shake flask and microcosm conditions. Among the three bacterial isolates, P. aeruginosa and biosurfactant produced by this strain resulted in maximum crude oil degradation compared to the other two bacterial strains investigated. Interestingly, when biosurfactant and bacterial cells were used (Set 3), significant oil biodegradation activity occurred and the difference between this treatment and that in Set 4 with added fertilizer + biosurfactant were only 4–5% higher degradation level in shake flask and 3.2–7% in microcosm experiments for all three bacterial strains used. It is concluded that, biosurfactants alone capable of promoting biodegradation to a large extent without added fertilizers, which will reduce the cost of bioremediation process and minimizes the dilution or wash away problems encountered when water soluble fertilizers used during bioremediation of aquatic environments. Ó 2010 Elsevier Ltd. All rights reserved.

1. Introduction Oil pollution in terrestrial and aquatic environments is a common phenomenon that causes significant ecological and social problems. The recent British Petroleum deep water horizon oil spill at the Gulf of Mexico during the summer of 2010 is a poignant example. The traditional available treatment processes used to decontaminate the polluted areas have been limited in their application (Perfumo et al., 2010b). Physical collection methods such as booms, skimmers and adsorbents typically recover no more than 10–15% of the spilled oil and the use of chemical surfactants as remediating agents is not favored due to their toxic effects on the existing biota in the polluted area. Therefore, despite decades of research, successful bioremediation of oil contaminated environment remains a challenge (Perfumo et al., 2010a).

⇑ Corresponding author. Tel.: +1 718 260 3408; fax: +1 718 260 3075. E-mail addresses: [email protected] (R. Thavasi), [email protected] (S. Jayalakshmi), [email protected] (I.M. Banat). 0960-8524/$ - see front matter Ó 2010 Elsevier Ltd. All rights reserved. doi:10.1016/j.biortech.2010.08.099

Bioremediation in aquatic environment is limited by the availability of nitrogen and phosphorous, which are necessary for initial microbial cell growth. The use of water-soluble salts containing nitrogen and phosphorus is effective under laboratory conditions, but are readily washed away by surface agitation and mixing in the aquatic environment. A possible alternate method could be through using biosurfactants along with nutrients in the form of fertilizers. Because biosurfactants rapidly emulsify the oil and therefore facilitate fast microbial growth and once the cell number reaches its peak, nutrient limitation would not be a problem to precede the biodegradation process. Biosurfactants are surface active compounds produced by microorganisms. There are many types of biosurfactants based on their chemical nature such as glycolipids, lipopolysaccharides, oligosaccharides, and lipopeptides have been reported to be produced by diverse bacterial genera (Banat et al., 2000, 2010; Franzetti et al., 2010). Biosurfactants received considerable attention in the field of environmental remediation processes such as bioremediation, soil washing, and soil flushing. Biosurfactants influence these processes because of their efficacy as dispersion and remediation

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agents and their environmentally friendly characteristics such as low toxicity and high biodegradability (Mulligan, 2005; Das et al., 2009; Sivapathasekaran et al., 2010; Kiran et al., 2010; Satpute et al., 2010). There are many reports describing the effect of exogenously added microbial biosurfactants in enhancing the bioremediation of crude oil-polluted soils by indigenous microbes (Abalos et al., 2004; Cubitto et al., 2004; Owsianiak et al., 2009) and how to purify and detect these biosurfactants (Satpute et al., 2009; Smyth et al., 2010a,b). As reported by Ron and Rosenberg (2002) biosurfactants are highly efficient emulsifiers that work at low concentrations i.e., 0.01–0.001%, representing emulsifier-tohydrocarbon ratios of 1:100–1:1000. This high active property at low concentration may reduce the addition of high volume of surfactants and fertilizers to remediate the oil polluted site. The purpose of this study was to evaluate the effect of biosurfactant and fertilizers on biodegradation of crude oil by three marine bacterial isolates namely, Corynebacterium kutscheri, Bacillus megaterium, and Pseudomonas aeruginosa in shake flask and microcosm experiments.

Table 1 Optimum culture conditions for biodegradation of crude oil (Thavasi, 2006).

*

Bacteria

Temperature (°C)

pH

Crude oil (%, v/v)

Salt (PPT or ‰)*

B. megaterium C. kutscheri P. aeruginosa

38 34 38

8.0 8.0 8.0

2.0 2.0 2.0

30 35 35

PPT or ‰, Parts per thousand.

aeruginosa. Urea and K2HPO4 (1:1, w/w) were used as fertilizers. The effect of culture conditions on biodegradation of crude oil and biosurfactant production was evaluated for each strain and reported elsewhere (Thavasi, 2006). Optimum culture conditions for each bacterial strain were outlined in Table 1. Degradation values from the uninoculated control flask (Set 5) was treated as natural weathering of crude oil and it was estimated as 5% and the value was subtracted from the results obtained in other experimental sets (Set 1–4). All the experiments were carried out in duplicate and the mean values were used as results.

2. Methods 2.3. Estimation of growth and crude oil degradation 2.1. Microorganisms and biosurfactants C. kutscheri, B. megaterium and P. aeruginosa used in this study were isolated and characterized as biosurfactant producing strains and reported earlier (Thavasi et al., 2007, 2008, 2010). The biosurfactants used in this study were produced through fermentation by the above bacterial strains using peanut oil cake as the carbon source. The chemical compositions of the biosurfactants are glycolipid, glycolipopetide and lipopeptide respectively for C. kutscheri, B. megaterium and P. aeruginosa. 2.2. Shake flask biodegradation experiments Shake flask biodegradation experiments were carried out in 500 mL Erlenmeyer flasks with 100 mL of mineral salt medium containing (g L1) 1.0 K2HPO4, 0.2 MgSO4 7H2O, 0.05 FeSO4 7H2O, 0.1 CaCl22H2O, 0.001 Na2MoO4 2H2O, 30 NaCl and crude oil (2.0%, w/v). Crude oil used in this study was obtained from Chennai Refineries Limited, Chennai, India with a specific gravity of 0.844 at 25 °C. Sterilized culture medium was inoculated with 1% (v/v) inoculum containing 105 bacterial cells mL1 and the culture flasks were maintained in a shaker for 168 h. Biodegradation experiments were conducted in five different sets using above culture medium: i. Bacterial cells + mineral salts medium + crude oil (normal)– Set 1 ii. Bacterial cells + mineral salts medium + fertilizer + crude oil – Set 2 iii. Bacterial cells + mineral salts medium + biosurfactant + crude oil – Set 3 iv. Bacterial cells + mineral salts medium + fertilizer + biosurfactant + crude oil – Set 4 v. Crude oil + mineral salts medium (No bacterial cells, control) – Set 5. The effect of fertilizer and biosurfactant concentration on biodegradation of crude oil was evaluated with different concentrations of fertilizer and biosurfactant (0.1, 0.5, 1.0, 1.5 and 2.0%, w/ v). The biosurfactants used in this study were produced by the same strain, i.e., glycolipid biosurfactant produced by C. kutscheri was used for biodegradation experiments with C. kutscheri, glycolipopetide and lipopeptide respectively for B. megaterium and P.

Crude oil degradation was estimated fluoremetrically as described in Intergovernmental Oceanographic Commission (IOC) Manuals and Guide No. 13 (1982). An amount of 5 mL culture medium was drawn from each above experimental sets and centrifuged at 6000 rpm to remove the bacterial cells. Crude oil residues from the cell free culture medium were extracted with n-hexane and total volume of the extract was made up to 10 mL. Crude oil content in the n-hexane extract was measured in a Varian, Cary Eclipse, fluorescence spectrophotometer at 310 nm excitation and at 374 nm emission wavelengths. Values were compared with a standard graph prepared with different concentration of crude oil using Cary Eclipse software and the degradation levels were expressed as percentage values. Bacterial cell growth was estimated using viable cell count method. Briefly, 1 mL of sample drawn from the culture broth was serially diluted and plated on nutrient agar plates and incubated for 24 h at 37 °C, colonies developed on the nutrient agar were counted and expressed as Log CFU mL1. 2.4. Gas chromatographic analysis of degraded residual crude oil Crude oil extract from culture broth was made up to 10 mL using n-hexane. A capillary column (30 m Fused Silica column, Restek Corporation, USA) and GC (Perkin–Elmer 8310) with flame ionisation detector were used for the analysis. The injection and detector temperature was 250 °C, and column temperature was programmed as 50 °C/4 min which was then increased at the rate of 10 °C/min to attain 330 °C and maintained at that temperature for 20 min. Florida Total Recoverable Petroleum Hydrocarbon (TRPH) standard (Restek-USA) was used to identify the compounds in the crude oil. 2.5. Laboratory scale microcosm experiment on biodegradation of crude oil with individual bacterial strains This experiment was carried out to investigate the influence of biosurfactant and fertilizers on biodegradation of crude oil in natural sea water. Plastic tanks with the capacity of 75 L were filled with 50 L of filtered and UV treated sea water having 30‰ (‰– parts per thousand) salt content, and pH 8.0. Experiments were conducted with five different sets as described in above methods Section 2.2. 2.0% (w/v) of crude oil was added to the filtered sea water and inoculated with 1% (v/v) of 24 h old bacterial culture

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containing 105 bacterial cells mL1. Continuous aeration was provided (1.5 L min1) using an oil free aerator and the experimental sets were maintained at room temperature for a period of 168 h (Thavasi et al., 2007). Biodegradation of crude oil was estimated fluoremeterically as described above in Section 2.3. An uninoculated control (Set 5) was maintained to assess the natural weathering of crude oil and it was estimated as 5% and the value was subtracted from the results obtained in other experimental sets (Set 1–4). 2.6. Bacterial adhesion to hydrocarbons (BATH assay) Cell hydrophobicity was measured by bacterial adherence to hydrocarbons (BATH) as described by Rosenberg et al. (1980). The cells were washed twice and suspended in a buffer salt solution (g L1 16.9 K2HPO4, 7.3 KH2PO4) to give an OD at 600 nm of 0.5. The cell suspension (2 mL) with 100 lL crude oil added was vortex-shaken for 3 min in 5 mL screw capped test tube. After shaking, crude oil and aqueous phase were allowed to separate for 1 h. The optical density (OD) of the aqueous phase was then measured at 600 nm in a spectrophotometer (Varian, Cary Eclipse, Spectrophotometer). For a given sample, three independent determinations were made and the mean value was calculated. Cells adhering to oil droplet were visualized by following a method described by Betts et al. (1989). Briefly, few drops of 2-(4-iodophenyl)-3-(4-nitrophenyl)-5-phenyltetrazolium chloride (INT) solution was added to the BATH assay culture broth and observed under the microscope. The INT turned red if it was reduced inside the cells, indicating the viability and adherence of cells with crude oil droplets. Hydrophobicity was expressed as the percentage of cell adherence to crude oil and calculated as follows:

% of bacterial adherence ¼ ð1  ðODshaken with oil =ODoriginal ÞÞ  100 where ODshaken with oil is OD of the mixture containing cells and crude oil. ODoriginal is OD of the cell suspension in the buffer solution (before mixing with crude oil). 2.7. Emulsification assay An amount of 500 lL of cell free culture broth was added to 5 mL of Tris buffer (pH 8.0) in a 30 mL screw capped test tube. Hydrocarbons such as waste motor lubricant oil, crude oil, diesel, kerosene, naphthalene, anthracene, xylene and peanut oil were tested for emulsification activity. 5 mg of hydrocarbon was added to the above solution and vortexed for 1 min and the mixture was allowed to stand for 20 min. The OD of the emulsified mixture was measured at 610 nm and the results were expressed as D610. 3. Results and discussion 3.1. Bacterial cell growth Results on bacterial cell growth in all the experiments showed that, all three bacterial strains utilized crude oil as their carbon and energy source, which was evident from the maximum cell growth observed after 128 h of incubation in all experiments, which is similar to earlier observation made by Thavasi et al. (2007, 2008, 2010) during biosurfactant production by these strains. Cell growth pattern for all three strains in shake flask and microcosm experiments were in the following order: biosurfactant + Fertilizer (Set 4) > biosurfactant (Set 3) > fertilizer (Set 2) > normal set (Set 1) (Fig. 1a, b, and c). The above information on higher growth observed in Set 4 with biosurfactant and fertilizer inferred that, biosurfactant facilitated the access of cells to crude oil

through emulsification and fertilizer provided additional essential nutrients to promote cell growth. Between the three bacterial strains used in this study, the growth pattern was in the order of P. aeruginosa > B. megaterium > C. kutscheri in all the treatments tested, which is similar to the observations reported by Sathishkumar et al. (2008) for Bacillus sp. IOS1–7, Corynebacterium sp. BPS2– 6, Pseudomonas sp. isolated from hydrocarbon contaminated area. Higher growth rate of P. aeruginosa compared to B. megaterium and C. kutscheri might be linked to more efficient breakdown and utilization of crude oil by P. aeruginosa (Das and Mukherjee, 2007). 3.2. Biodegradation with different concentrations of fertilizer Among the different fertilizer concentrations used 0.1% was the optimum level for biodegradation of crude oil for all three bacterial strains (Fig. 2a). Beyond 0.1% there was no significant change observed in bacterial cell growth and biodegradation rate. As observations made in this experiment, Kanaly et al. (2002) and Vyas and Dave (2010) reported that addition of excess nutrients beyond certain limit in bioremediation would have no impact on cell growth and biodegradation process and excess nutrient content can be toxic to cell growth. Thus, 0.1% fertilizer concentration was used as the optimum level for further shake flask and microcosm experiments carried out in this study. Maximum crude oil degradation was recorded with P. aeruginosa (89%), which indicates its suitability for use in environmental oil bioremediation and cleaning applications which may need limited nutrients addition. 3.3. Biodegradation with different concentrations of biosurfactant All three bacterial cultures used in this study showed maximum cell growth and biodegradation activity with 0.1% (w/v) of their relevant biosurfactant (Fig. 2b). There was no significant increase in degradation activity observed at concentrations above 0.1%; which indicates that at this concentration sufficient emulsification of the crude oil occurred making it bioavailable for degradation. Biosurfactants are known for their potential emulsification activity even at very low concentration (Raza et al., 2009). All the strains used in this study are biosurfactant producers and the additional initial biosurfactants added to the culture medium could be enough to facilitate the bacterial access to the crude oil followed with more biosurfactant production. Maximum crude oil degradation was recorded with P. aeruginosa (89%, with 0.1% biosurfactant), similar biosurfactant production and biodegradation activity of P. aeruginosa reported by many workers support the results obtained in this study (Das and Mukherjee, 2007; Cameotra and Singh, 2008; Obahiagbon and Akhabue 2009). Rhamnolipids are the main biosurfactant produced by P. aeruginosa reported in above studies, whereas the strain used in this study produces lipopeptide surface active molecules (Thavasi et al., 2010). In this study, we used the biosurfactants produced by the same strain in their biodegradation experiment; however detailed investigation are needed to determine the effect of biosurfactant produced by one bacterium on other bacteria to select the potential biosurfactant, most useful for all biodegrading microbes. 3.4. Biodegradation in shake flask and laboratory scale microcosm experiments Biodegradation of crude oil in shake flask (Fig. 3a) and laboratory scale experiment (Fig. 3b) showed maximum biodegradation when both biosurfactant + fertilizer (Set 4) were added for all strains used. In shake flask experiments, the difference in biodegradation levels when only bacterial cells were added (Set 1) and when additional biosurfactant + fertilizer (Set 4) was 19% (for all

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three bacterial strains). However there wasn’t much difference between the treatments with only biosurfactant (Set 3) and that with both biosurfactant + fertilizer (Set 4) was 4–5% for all three strains.

a

Biosurfactant

The difference was more significant when both biosurfactant and fertilizer were added (Set 4) and when fertilizer only was added (Set 2) where 11%, 12% and 9% for B. megaterium, C. kutscheri and

Fertilizer

Biosurfactant + Fertilizer

Normal

12

Log CFU/ml

10

8

6 4

2

0 0

24

48

72

96

120

144

168

Time (h)

b

Biosurfactant

Fertilizer

Biosurfactant + Fertilizer

Normal

10 9 8 Log CFU/ml

7 6 5 4 3 2 1 0 0

24

48

72

96

120

144

168

Time (h)

c

Biosurfactant

Fertilizer

Biosurfactant + Fertilizer

Normal

12

Log CFU/ml

10 8 6 4 2 0 0

24

48

72

96

120

144

168

Time (h) Fig. 1. (a) B. megaterium, (b) C. kutscheri, (c) P. aeruginosa cell growth in shake flask biodegradation experiments–normal (without biosurfactant and fertilizer) (Set 1), fertilizer (Set 2), biosurfactant (Set 3), and biosurfactant + fertilizer (Set 4).

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P. aeruginosa respectively. This indicated that the main factor improving oil degradation was the presence of biosurfactant rather than fertilizer. A similar trend was observed in the microcosm experiments were 19.3%, 14.7%, and 37.7% difference in degradation levels were observed between Set 1 and 4 for B. megaterium, C. kutscheri and P. aeruginosa respectively. Again there were much less different in biodegradation levels between Set 3 and 4 in microcosm with 7%, 3.5% and 3.2% difference for B. megaterium, C. kutscheri and P. aeruginosa respectively, indicating that biosurfactants alone can promote biodegradation to a greater extent without any fertilizer addition. The improved biodegradation levels obtained with biosurfactant indicated that they represent the most efficient accelerators for hydrocarbon biodegradation through increasing oil bioavailability (Perfumo et al., 2010a,b). The use of biosurfactant in combination with fertilizer could reduce the actual amount of fertilizer to be added to polluted sites. In some studies, water soluble fertilizers encountered problem such as washing away and rapid dilution in aquatic environment. Maki et al. (2003) reported that fertilizers only stimulate the early stage degradation rate of the oil and that the final degradation efficiencies with fertilizers were not significantly different from those where no fertilizers were used. It is important however to keep in mind that nutrients or fertilizers use may be essential in some environments with insufficient nutrient levels.

a

Bacillus megaterium

Gas chromatographic analysis of the degraded crude oil extracted from the culture medium was commensurate with the degradation values obtained by quantitative fluorometric analysis. All the bacterial strains used in this study were able to break the compounds present in crude oil. Preferential degradation of compounds by bacterial strains revealed that, B. megaterium utilized C18 compounds, C. kutscheri C12 compounds and P. aeruginosa C12 and C21 compounds completely (data not shown).

3.5. BATH assay BATH assay results showed that, P. aeruginosa had maximum cell adhesion with crude oil (95.3%), followed by C. kutscheri (49.7%), and B. megaterium (40.2%). Such high crude oil affinity observed with P. aeruginosa correlated with the maximum biodegradation potential observed for this strain. Cell surface properties are important factors that determine the rate of degradation of hydrophobic substrates (Franzetti et al., 2009). In an early investigation, cells exhibiting highest hydrophobicities were among the fastest hydrocarbon degraders (Zhang and Miller, 1994). Therefore, isolates with high hydrophobicity are likely to be more efficient degraders, as reported in this study for P. aeruginosa. Cell hydrophobicity is also an indication of biosurfactant production (Franzetti et al., 2009).

Corynebacterium kutscheri

Pseudomonas aeruginosa

% of Crude oil degradation

100 90 80 70 60 50 40 30 20 10 0 0.05%

0.10%

0.50%

1.00%

1.50%

2.00%

Fertilizer Concentration

b

Bacillus megaterium

Corynebacterium kutscheri

Pseudomonas aeruginosa

% of Crude oil degradation

100 90 80 70 60 50 40 30 20 10 0 0.05%

0.10%

0.50%

1.00%

1.50%

2.00%

Biosurfactant Concentration Fig. 2. Biodegradation of crude oil with different concentration of fertilizer (a) and biosurfactant (b).

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a

Bacillus megaterium

Corynebacterium kutscheri

Pseudomonas aeruginosa

% of Crude oil degradation

100 90 80 70 60 50 40 30 20 10 0 Normal

Fertilizer

Biosurfactant

Biosurfactant & Fertilizer

Experimental Setup

b

Bacillus megaterium

Corynebacterium kutscheri

Pseudomonas aeruginosa

100 % of Crude oil degradation

90 80 70 60 50 40 30 20 10 0 Normal

Fertilizer

Biosurfactant Experimental Setup

Biosurfactant & Fertilizer

Fig. 3. Biodegradation of crude oil in shake flask (a) and lab scale microcosm (b) experiments–normal (without biosurfactant and fertilizer) (Set 1), fertilizer (Set 2), biosurfactant (Set 3), and biosurfactant + fertilizer (Set 4).

3.6. Emulsification assay Emulsification activity of crude biosurfactants present in the cell free culture broth observed in this study was comparatively less than the emulsification activity recorded with the standard chemical surfactant Triton X-100 (Table 2). However, when considering the advantages of biosurfactants over chemically synthesized surfactants, such as lower toxicity, biodegradability and ecological acceptability, their use in bioremediation activity becomes more favorable. Emulsification of different hydrocarbons by the biosurfacTable 2 Emulsification of hydrocarbons by crude biosurfactant isolated from aB. megaterium, b C. kutscheri, cP. aeruginosa and standard surfactant Triton X-100.

4. Conclusion

Emulsification activity (D610)

Waste motor lubricant oil Crude oil Peanut oil Kerosene Diesel Xylene Naphthalene Anthracene

tants was in the order of waste motor lubricant oil > crude oil > peanut oil > kerosene > diesel > naphthalene > anthracene > xylene. Crude oil therefore was the second compound that has been highly emulsified by all three biosurfactants used in this study, which coincided with the enhanced biodegradation of crude oil obtained in Set 3 and 4 in this study. Fernandez-Linares et al. (1996) reported similar emulsification results by two marine strains, Pseudomonas nautical and Marinobacter hydrocarbonoclasticus, and concluded that emulsification is an essential process in alkane biodegradation. High emulsification activity at low concentration observed with biosurfactant produced by P. aeruginosa in this study could be used for environmental and crude oil tanker cleaning applications.

a

b

c

Triton X-100

1.86 1.72 1.42 1.01 0.85 0.53 0.46 0.42

1.72 1.69 1.31 0.95 0.81 0.51 0.41 0.41

2.01 1.97 1.95 1.12 0.90 0.61 0.51 0.45

1.93 1.85 1.56 1.12 0.94 0.78 0.63 0.58

The addition of biosurfactant along with fertilizer enhanced the oil degradation potential of all three bacterial isolates tested. Interestingly, biosurfactant addition alone to bacterial cells had significant oil biodegradation with little difference compared to the treatments where biosurfactant + fertilizer were added. Biosurfactants alone therefore are capable of promoting biodegradation process without using fertilizers. However, two important issues needs to be addressed before taking such technology to the field,

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(i) evaluation of the environment where such bioremediation process is needed for lack of essential nutrients, (ii) possible adverse effects of biosurfactants produced by some microorganisms to other bacteria employed in bioremediation. Acknowledgements We thank the authorities of Annamalai University for providing the facilities and Department of Ocean Development (DOD) and Council of Scientific and Industrial Research (CSIR), Government of India for providing financial support. References Abalos, A., Vinas, M., Sabate, J., Manresa, M.A., Solanas, A.M., 2004. Enhanced biodegradation of Casablanca crude oil by a microbial consortium in presence of a rhamnolipid produced by Pseudomonas aeruginosa AT 10. Biodegradation 15, 249–260. Banat, I.M., Makkar, R.S., Cameotra, S.S., 2000. Potential commercial applications of microbial surfactants. Appl. Microbiol. Biotechnol. 53, 495–508. Banat, I.M., Franzetti, A., Gandolfi, I., Bestetti, G., Martinotti, M.G., Fracchia, L., Smyth, T.J., Marchant, R., 2010. Microbial biosurfactants production, applications and future potential. Appl. Microbiol. Biotechnol. 87, 427–444. Betts, R.P., Bankers, P., Banks, J.G., 1989. Rapid enumeration of viable microorganisms by staining and direct microscopy. Lett. Appl. Microbiol. 9, 199–202. Cameotra, S.S., Singh, P., 2008. Bioremedation of oil sludge using crude biosurfactants. Int. Biodeterior. Biodegrad. 62, 274–280. Cubitto, M.A., Moran, A.C., Commendatore, M., Chiarello, M.N., Baldini, M.D., Sineriz, F., 2004. Effects of Bacillus subtilis O9 biosurfactant on the bioremediation of crude oil-polluted soil. Biodegradation 15, 281–287. Das, K., Mukherjee, A.K., 2007. Crude petroleum-oil biodegradation efficiency of Bacillus subtilis and Pseudomonas aeruginosa strains isolated from a petroleumoil contaminated soil from North-East India. Bioresour. Technol. 98, 1339–1345. Das, P., Mukherjee, S., Sen, R., 2009. Biosurfactant of marine origin exhibiting heavy metal remediation properties. Bioresour. Technol. 100, 4887–4890. Fernandez-Linares, L., Acquaviva, M., Bertrand, J.-C., Gauthier, M., 1996. Effect of sodium chloride concentration on growth and degradation of eicosane by marine halotolerent bacterium Marinobacter hydrocarbonoclastieus. Appl. Microbiol. 19, 113–121. Franzetti, A., Caredda, P., Colla, P.L., Pintus, M., Tamburini, E., Papacchini, M., Bestetti, G., 2009. Cultural factors affecting biosurfactant production by Gordonia sp. BS29. Int. Biodeterior. Biodegrad. 63, 943–947. Franzetti, A., Tamburini, E., Banat, I.M., 2010. Application of biological surface active compounds in remediation technologies. In: Sen, R. (Ed.), Biosurfactants: advances in Experimental Medicine and Biology, vol. 672. Springer-Verlag, Berlin Heidelberg, pp. 121–134. Intergovernmental Oceanographic Commission Manuals and Guide No.13, 1982. Manual for monitoring oil and dissolved/dispersed petroleum hydrocarbons in marine waters and beaches. UNESCO. Kanaly, R.A., Harayama, S., Watanabe, K., 2002. Rhodanobacter sp. strain BPC1 in a benzo(a)pyrene-mineralizing bacterial consortium. Appl. Environ. Microbiol. 68, 5826–5833. Kiran, S.G., Thomas, T.A., Selvin, J., Sabarathnam, B., Lipton, A.P., 2010. Optimization and characterization of a new lipopeptide biosurfactant produced by marine Brevibacterium aureum MSA13 in solid state culture. Bioresour. Technol. 101, 2389–2396. Maki, H., Hirayama, N., Hiwatari, T., Kohata, K., Uchiyama, H., Watanabe, M., Yamasaki, F., Furuki, M., 2003. Crude oil bioremediation field experiment in the Sea of Japan. Mar. Poll. Bull. 47, 74–77.

Mulligan, C.N., 2005. Environmental applications for biosurfactants. Environ. Pollut. 133, 183–198. Obahiagbon, K.O., Akhabue, C.E., 2009. Effect of microbial count of P. aeruginosa on biodegradation of crude oil contaminated water. Pet. Sci. Technol. 27, 1402– 1412. Owsianiak, M., Chrzanowski, Ł., Szulc, A., Staniewski, J., Olszanowski, A., OlejnikSchmidt, A.K., Heipieper, H.J., 2009. Biodegradation of diesel/biodiesel blends by a consortium of hydrocarbon degraders: Effect of the type of blend and the addition of biosurfactants. Bioresour. Technol. 100, 1497–1500. Perfumo, A., Smyth, T.J.P., Marchant, R., Banat, I.M., 2010a. Production and roles of biosurfactants and bioemulsifiers in accessing hydrophobic substrates. In: Timmis, K.N. (Ed.), Handbook of hydrocarbon and lipid microbiology. SpringerVerlag, Berlin Heidelberg, pp. 1501–1512. Perfumo, A., Rancich, I., Banat, I.M., 2010b. Possibilities and challenges for biosurfactants use in petroleum industry. In: Sen, R. (Ed.), Biosurfactants advances in Experimental Medicine and Biology, vol. 672. Springer-Verlag, Berlin Heidelberg, pp. 135–157. Raza, Z.A., Khalid, Z.M., Banat, I.M., 2009. Characterization of rhamnolipids produced by a Pseudomonas aeruginosa mutant strain grown on waste oils. J. Environ. Sci. Health Part A 44, 1367–1373. Ron, E.Z., Rosenberg, E., 2002. Biosurfactants and oil bioremediation. Curr. Opin. Biotechnol. 13, 24–252. Rosenberg, M., Gutnick, D., Rosenberg, E., 1980. Adherence to bacteria to hydrocarbons: a simple method for measuring cell-surface hydrophobicity. FEMS Microbiol. Lett. 9, 29–33. Sathishkumar, M., Binupriya, A.R., Baik, S.-H., Yun, S.-E., 2008. Biodegradation of crude oil by individual bacterial strains and a mixed bacterial consortium isolated from hydrocarbon contaminated areas. Clean 36, 92–96. Satpute, S.K., Banpurkar, A.G., Dhakephalkar, P.K., Banat, I.M., Chopade, B.A., 2009. Methods for investigating biosurfactants and bioemulsifiers: a review. Crit. Rev. Biotechnol. 30, 127–144. Satpute, S.K., Banat, I.M., Dhakephalkar, P.K., Banpurkar, A.G., Chopade, B.A., 2010. Biosurfactants, bioemulsifiers and exopolysaccharides from marine microorganisms. Biotechnol. Adv. 28, 436–450. Sivapathasekaran, C., Mukherjee, S., Ray, A., Gupta, A., Sen, R., 2010. Artificial neural network modeling and genetic algorithm based medium optimization for the improved production of marine biosurfactant. Bioresour. Technol. 101, 2884– 2887. Smyth, T.J.P., Perfumo, A., Marchant, R., Banat, I.M., 2010a. Isolation and analysis of low molecular weight microbial glycolipids. In: Timmis, K.N. (Ed.), Handbook of Hydrocarbon and Lipid Microbiology. Springer-Verlag, Berlin Heidelberg, pp. 3705–3723. Smyth, T.J.P., Perfumo, A., McClean, S., Marchant, R., Banat, I.M., 2010b. Isolation and analysis of lipopeptides and high molecular weight biosurfactants. In: Timmis, K.N. (Ed.), Handbook of Hydrocarbon and Lipid Microbiology. Springer-Verlag, Berlin Heidelberg, pp. 3689–3704. Thavasi, R., 2006. Biosurfactants from marine hydrocarbonoclastic bacteria and their application in marine oil pollution abatement. Ph.D thesis, Annamalai University, India, p. 162. Thavasi, R., Jayalakshmi, S., Balasubramanian, T., Banat, I.M., 2007. Biosurfactant production by Corynebacterium kutscheri from waste motor lubricant oil and peanut oil cake. Lett. Appl. Microbiol. 45, 686–691. Thavasi, R., Jayalakshmi, S., Balasubramanian, T., Banat, I.M., 2008. Production and characterization of a glycolipid biosurfactant from Bacillus megaterium. World J. Microbiol. Biotechnol. 24 (7), 917–925. Thavasi, R., V.R.M., Subramanyam Nambaru, Jayalakshmi, S., Balasubramanian, T., Banat, I.M, 2010. Biosurfactant production by Pseudomonas aeruginosa from renewable resources. Indian J. Microbiol. (Submitted for publication). Vyas, T.K., Dave, B.P., 2010. Effect of addition of nitrogen, phosphorus and potassium fertilizers on biodegradation of crude oil by marine bacteria. Indian J. Mar. Sci. 39, 143–150. Zhang, Y., Miller, R.M., 1994. Effect of Pseudomonas rhamnolipid biosurfactant on cell hydrophobicity and biodegradation of octadecane. Appl. Environ. Microbiol. 60, 2101–2106.

Bioresource Technology 102 (2011) 3366–3372

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Bioresource Technology journal homepage: www.elsevier.com/locate/biortech

Application of biosurfactant produced from peanut oil cake by Lactobacillus delbrueckii in biodegradation of crude oil Rengathavasi Thavasi a,⇑, Singaram Jayalakshmi b, Ibrahim M. Banat c a

Department of Chemical and Biological Sciences, Polytechnic Institute of New York University, 6 Metrotech Center, Brooklyn, NY 11201, USA CAS in Marine Biology, Annamalai University, Parangipettai 608 502, Tamil Nadu, India c School of Biomedical Sciences, University of Ulster, Coleraine BT52 1SA, Northern Ireland, UK b

a r t i c l e

i n f o

Article history: Received 12 October 2010 Received in revised form 16 November 2010 Accepted 17 November 2010 Available online 21 November 2010 Keywords: Biosurfactant Peanut oil cake Bioremediation Emulsification Lactobacillus delbrueckii

a b s t r a c t Lactobacillus delbrueckii cultured with peanut oil cake as the carbon source yielded 5.35 mg ml1 of biosurfactant production. Five sets of microcosm biodegradation experiments were carried out with crude oil as follows: set 1 – bacterial cells + crude oil, set 2 – bacterial cells + crude oil + fertilizer, set 3 – bacterial cells + crude oil + biosurfactant, set 4 – bacterial cells + crude oil + biosurfactant + fertilizer, set 5 – with no bacterial cells, fertilizer and biosurfactant (control). Maximum degradation of crude oil was observed in set 4 (75%). Interestingly, when biosurfactant and bacterial cells were used (set 3), significant oil biodegradation activity occurred and the difference between this treatment and that in set 4 was 7% higher degradation level in microcosm experiments. It is evident from the results that biosurfactants alone is capable of promoting biodegradation to a large extent without added fertilizers. Ó 2010 Elsevier Ltd. All rights reserved.

1. Introduction Pollution caused by petroleum hydrocarbons in terrestrial and aquatic environment is a common phenomenon that causes significant ecological and social problems. Physical and chemical cleaning processes used to decontaminate the oil polluted areas have been limited in their application (Perfumo et al., 2010a). Physical collection methods such as booms, skimmers, and adsorbents typically recover no more than 10–15% of the spilled oil and the use of chemical surfactants as remediating agents is not favorable due to their toxic effects on the existing biota in the polluted area. Therefore, despite decades of research, successful bioremediation of oil contaminated environment still remains a challenge (Perfumo et al., 2010b). Biodegradation in aquatic environment is limited by the availability of nutrients such as nitrogen and phosphorous, which are necessary for initial microbial cell growth. The use of water-soluble salts containing nitrogen and phosphorus is effective under laboratory conditions, but are readily washed away by surface agitation and mixing in the aquatic environment. A possible alternate method could be using biosurfactants along with nutrients in the form of fertilizers. Because biosurfactants rapidly emulsify the oil and therefore facilitate fast microbial growth and once the cell number

⇑ Corresponding author. Tel.: +1 718 260 3960; fax: +1 718 260 3075. E-mail address: [email protected] (R. Thavasi). 0960-8524/$ - see front matter Ó 2010 Elsevier Ltd. All rights reserved. doi:10.1016/j.biortech.2010.11.071

reaches its peak, nutrient limitation would not be a problem to precede the biodegradation process. Biosurfactants are surface active compounds produced by microorganisms. There are many types of biosurfactants based on their chemical composition such as glycolipids, lipopolysaccharides, oligosaccharides, and lipopeptides that have been reported to be produced by diverse bacterial genera (Franzetti et al., 2010; Banat et al., 2010). Biosurfactants received considerable attention in the field of environmental remediation processes such as bioremediation, soil washing, and soil flushing. Biosurfactants influence these processes because of their efficacy as dispersion and remediation agents and their environmentally friendly characteristics such as low toxicity and high biodegradability (Sivapathasekaran et al., 2010; Kiran et al., 2010; Satpute et al., 2010). Although biosurfactants exhibit such important advantages, they have not yet been employed extensively in industry because of relatively high production costs. One possible strategy for reducing costs is the utilization of alternative substrates such as agroindustrial wastes. The main problem related to the use of alternative substrates as carbon source is to find a waste with the right balance of nutrients that permits cell growth and product accumulation (Makkar and Cameotra, 1999). Peat hydrolysate (Sheppard and Mulligan, 1987), molasses (Makkar and Cameotra, 1999), potato processing effluents (Fox and Bala, 2000), cheese whey and molasses (Rodrigues et al., 2006c), and agriculture residues (Moldes et al., 2007) are few examples of alternative substrates that have been used for biosurfactant production. The establishment of

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waste-based medium for biosurfactant production also faces another problem – the kind and the properties of final product are dependent on the composition of the culture medium. Hence, in the present study peanut oil cake was tried as a cheaper carbon source for biosurfactant production. Peanut oil cake is a carbohydrate, protein and lipid rich residue generated in large quantities during the production of peanut oil and the cost of this cake is very low as compared to other carbon sources like glucose, fructose, crude oil, and other hydrocarbons. In this study Lactobacillus delbrueckii was used to investigate its biosurfactant production and biodegradation potentials. The reason behind selecting this strain for this work is that among many bacterial genera, genus Lactobacillus is known for its benevolent uses to humans in many ways such as production of lactic acid (Wee et al., 2005), yogurt (Omogbai et al., 2005), cheese (Blaiotta et al., 2001), bacteriocin (Vuyst and Leroy, 2007), and in some cases the bacteria is used as probiotics, prebiotics (Teitelbaum and Walker, 2002), and biotherapeutics (Buddington, 2009). Studies on biosurfactant production by Lactobacillus are scarce and there are only few reports available on biosurfactant production by the lactic acid bacteria Lactobacillus (Reid et al., 2002; Rodrigues et al., 2006a; Moldes et al., 2007; Golek et al., 2007; Gudina et al., 2010) and only two reports on biosurfactant production using agriculture residues by Lactobacillus pentosus (Moldes et al., 2007; Portilla-Rivera et al., 2008). So, as an addition to above reports on beneficial uses of Lactobacillus, L. delbrueckii used in this study was reported as a strain with biosurfactant production and crude oil biodegradation potentials (Thavasi, 2006). Hence, the present study was undertaken with two objectives: (i) Evaluation of biosurfactant producing potential of L. delbrueckii using peanut oil cake, and (ii) Effect of biosurfactant and fertilizer addition on biodegradation of crude oil by L. delbrueckii.

2. Methods 2.1. Culture conditions and biosurfactant production L. delbrueckii used in this study was isolated and characterized as a biosurfactant producing strain and reported earlier (Thavasi, 2006). Optimization of culture conditions such as temperature, pH, and substrate concentration (carbon source) in shake flask experiments reported in the above study were used for biosurfactant production and biodegradation experiments in this study. Biosurfactant production was carried out in a 3 l fermentor with working volume of 2.1 l (Scigenics India Pvt. Ltd., Chennai). L. delbrueckii was cultured in mineral salts medium containing (g l1) 1.0 K2HPO4, 0.2 MgSO47H2O, 0.05 FeSO47H2O, 0.1 CaCl22H2O, 0.001 Na2MoO42H2O, 5.0 NaCl, and peanut oil cake (2.0%, w/v) as the carbon source. Sterilized culture medium was inoculated with 1% (v/v) inoculum containing 105 bacterial cells ml1 and the culture was maintained at 34 °C temperature, pH 7.5, 350 rpm of agitation and 1.5 l min1 of air flow (8.5 mg ml1 dissolved oxygen) for 168 h. Five millilitre samples of culture broth were collected at 24 h intervals for a period of 168 h. Bacterial cell growth was estimated using viable cell count method. Briefly, 1 ml of sample drawn from the culture broth was serially diluted and plated on nutrient agar plates and incubated for 24 h at 34 °C, colonies developed on the nutrient agar were counted and expressed as Log CFU ml1. Concentration of extracellular biosurfactant in the culture broth was estimated according to the procedure described by Thavasi et al. (2007) and the biosurfactant concentration was expressed as mg ml1 (dry weight). Briefly, culture broth was centrifuged at 6000 rpm for 20 min at 4 °C and the cell free culture broth was extracted twice with chloroform and methanol (2:1, v/v). Solvents in

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the extracts were removed by rotary evaporation and the residue was partially purified in silica gel (60–120 mesh) column eluted with chloroform and methanol ranging from 20:1 to 2:1 (v/v) in a gradient manner. The fractions were pooled and solvents were evaporated. The resulting residue was dialysed against distilled water and freeze dried. Freeze dried partially purified biosurfactant was used for further analysis and biodegradation experiments. 2.2. Characterization of biosurfactant 2.2.1. Biochemical composition of biosurfactant Carbohydrate content of the biosurfactant was determined by the phenol–sulfuric acid method (Dubois et al., 1956) using D-glucose as a standard. Protein content was determined by the method of Lowry et al. (1951) using bovine serum albumin as a standard and lipid content was estimated by following the procedure of Folch et al. (1956). 2.2.2. Fourier transform infrared spectroscopy Fourier transform infrared spectroscopy (FTIR) is most useful for identifying types of chemical bonds (functional groups), therefore can be used to elucidate some components in an unknown mixture. Freeze dried biosurfactant (10 mg) was mixed with 100 mg of potassium bromide and pressed with 7500 kg for 30 s to obtain translucent pellets. Infrared absorption spectra were recorded on a Thermo Niocolet, AVATAR 330 FTIR system with a spectral resolution and wave number accuracy of 4 and 0.01 cm1, respectively. All measurements consisted of 500 scans, and a potassium bromide pellet was used as background reference. 2.2.3. Mass spectrometric analysis of biosurfactant Biosurfactant was dissolved in methanol and mixed thoroughly. The mass spectrometric analysis of the biosurfactant was carried out in an LCQ™ quadrupole ion-trap mass spectrometer (Finnigan MAT, San Jose, CA, USA) which utilizes electrospray ionization (ESI). Standard solutions and samples under investigation were infused into the mass spectrometer at a flow rate of 10 ll min1. In the ESI, nitrogen and auxiliary gas flows were maintained at 50 and 5 ml min1, respectively and referred to arbitrary values set by the software. The heated capillary temperature was 250 °C and the spray voltage was set to 5 kV. Negative ion mode was used and scanning was performed at 50–2000m/z range. 2.3. Shake flask crude oil biodegradation experiments with different concentration of fertilizer and biosurfactant Shake flask biodegradation experiments were carried out in 500 ml Erlenmeyer flasks with 100 ml of mineral salt medium (same medium used for biosurfactant production) with 2.0% (w/v) of crude oil. Crude oil used in this study was obtained from Chennai Refineries Ltd., Chennai, India with a specific gravity of 0.844 at 25 °C. Sterilized culture medium was inoculated with 1% (v/v) inoculum containing 105 bacterial cells ml1 and the culture flasks were maintained in a shaker at 200 rpm for 168 h. Other culture conditions used for this experiment were same as mentioned in biosurfactant production experiments. The effect of fertilizer and biosurfactant concentration on biodegradation of crude oil was evaluated with different concentrations of fertilizer and biosurfactant (0.1%, 0.5%, 1.0%, 1.5%, and 2.0%, w/v). Urea and K2HPO4 (1:1, w/w) were used as fertilizers. Critical micelle concentration (CMC) obtained for the biosurfactant used in this study was 2 mg ml1 (unpublished data), based on which above concentrations were used to check the effect biosurfactant on biodegradation. The biosurfactant used in this study for biodegradation experiments was produced by the same strain that produces the

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biosurfactant, i.e., L. delbrueckii. All the experiments were carried out in duplicate and the mean values were used as results. 2.4. Estimation of growth and crude oil degradation Crude oil degradation was estimated fluorometrically as described in Intergovernmental Oceanographic Commission (IOC) Manuals and Guide No. 13 (1982). Five millilitres of culture medium was drawn from each above experimental sets and centrifuged at 6000 rpm to remove the bacterial cells. Crude oil residues from the cell free culture medium were extracted with n-hexane and total volume of the extract was made up to 10 ml. Crude oil content in the n-hexane extract was measured in a Varian, Cary Eclipse fluorescence spectrophotometer at 310 nm excitation and at 374 nm emission wavelengths. Values were compared with a standard graph prepared with different concentration of crude oil using Cary Eclipse software and the degradation levels were expressed as percentage values. Bacterial cell growth was estimated using viable cell count method as described in (Section 2.1). 2.5. Laboratory scale microcosm experiment on biodegradation of crude oil This experiment was carried out to investigate the influence of biosurfactant and fertilizers on biodegradation of crude oil in natural sea water. Seventy five litres plastic tanks were filled with 50 l of filtered and UV treated sea water having 30‰ (‰ – parts per thousand) salt content, and pH 8.0. Biodegradation experiments were conducted in five different sets and they are as follows: set 1 – Bacterial cells + mineral salts medium + crude oil (normal), set 2 – Bacterial cells + mineral salts medium + fertilizer + crude oil, set 3 – Bacterial cells + mineral salts medium + biosurfactant + crude oil, set 4 – Bacterial cells + mineral salts medium + fertilizer + biosurfactant + crude oil, and set 5 – Crude oil + mineral salts medium (control-with no bacterial cells, fertilizer and biosurfactant). To the above experimental sets, 2.0% (w/v) of crude oil, 0.1% (w/v) of fertilizer (Urea and K2HPO4), and biosurfactant were added and inoculated with 1% (v/v) of 24 h old bacterial culture containing 105 bacterial cells ml1. Continuous aeration was provided (1.5 l min1) using an oil free aerator and the experimental sets were maintained at room temperature for a period of 168 h (Thavasi et al., 2007). Bacterial cell growth and biodegradation of crude oil was estimated as described above in Sections 2.1 and 2.4, respectively. An uninoculated control (set 5) was maintained to assess the natural weathering of crude oil and it was estimated as 6.5% and the value was subtracted from the results obtained in other experimental sets (sets 1–4). 2.6. Gas chromatographic analysis of degraded residual crude oil Crude oil extract from culture broth was made up to 10 ml using n-hexane. A capillary column (30 m Fused Silica column, Restek Corporation, USA) and GC (Perkin–Elmer 8310) with flame ionization detector were used for the analysis. The injection and detector temperature was 250 °C, and column temperature was programmed as 50 °C/4 min which was then increased at the rate of 10 °C/min to attain 330 °C and maintained at that temperature for 20 min. Florida Total Recoverable Petroleum Hydrocarbon (TRPH) standard (Restek, USA) was used to identify the compounds in the crude oil (Thavasi et al., 2011). 2.7. Bacterial adhesion to hydrocarbons (BATH assay) Cell hydrophobicity was measured by bacterial adherence to hydrocarbons (BATH) as described by Rosenberg et al. (1980). L. delbrueckii cells obtained from centrifuged culture broth were

washed twice and suspended in a buffer salt solution (g l1 16.9 K2HPO4 and 7.3 KH2PO4) to give an OD of 0.5 at 600 nm. The cell suspension (2 ml) with 100 ll crude oil was vortex-shaken for 3 min in 5 ml screw capped test tube. After shaking, crude oil and aqueous phases were allowed to separate for 1 h. The optical density (OD) of the aqueous phase was then measured at 600 nm in a spectrophotometer (Varian, Cary Eclipse Spectrophotometer). For a given sample, three independent determinations were made and the mean value was calculated. Cells adhering to oil droplet were visualized by following a method described by Betts et al. (1989). Briefly, few drops of 2-(4-iodophenyl)-3-(4-nitrophenyl)-5-phenyltetrazolium chloride (INT) solution was added to the BATH assay culture broth and observed under the microscope. The INT turned red if it was reduced inside the cells, indicating the viability and visualized the adherence of cells with crude oil droplets. Hydrophobicity was expressed as the percentage of cell adherence to crude oil and calculated as follows:

% of bacterial adherence ¼ ð1  ðODshaken with oil =ODoriginal ÞÞ  100 where ODshaken with oil is OD of the mixture containing cells and crude oil and ODoriginal is the OD of the cell suspension in the buffer solution (before mixing with crude oil). 2.8. Emulsification assay Partially purified biosurfactant (1 mg ml1) was dissolved in 5 ml of tris buffer (pH 8.0) in 30 ml test tubes. Hydrocarbons such as waste motor lubricant oil, crude oil, diesel, kerosene, naphthalene, anthracene, xylene, and peanut oil were tested for emulsification activity. Five milligrams of hydrocarbon was added to the above solution and vortexed for 1 min and the mixture was allowed to stand for 20 min. OD of the emulsified mixture was measured at 610 nm and the results were expressed as D610. A control was maintained with above buffer solution and hydrocarbons (no biosurfactant added), and the values obtained for this control was subtracted from emulsification values obtained with biosurfactant. A standard chemical surfactant, Triton X-100 was used to compare the emulsification of biosurfactant, and the reaction conditions and procedure used were same as described above for biosurfactant. 3. Results and discussion 3.1. Bacterial cell growth and biosurfactant production In order to economize the biosurfactant production, the cheaper carbon source – peanut oil cake was used. Fig. 1 shows the time course of biosurfactant production by L. delbrueckii with peanut oil cake as the substrate. Maximum biosurfactant concentration of 5.35 mg ml1 was observed at 144 h of incubation, when the cells reached their early stationary phase. Maximum biomass was observed at 120 h (9.04 Log CFU ml1). Higher biosurfactant concentration after the offset of growth may be because of the release of cell-bound biosurfactant at the early stationary phase (144 h), which leads to an increase in extracellular biosurfactant concentration in the culture medium. Similar observation was made by Thavasi et al. (2007, 2008) while culturing Bacillus megaterium and Corynebacterium kutscheri with peanut oil cake for biosurfactant production. Biosurfactant production reported in this study for peanut oil cake (5.35 g l1) was higher than earlier reports on use of cheese whey and molasses as carbon sources for biosurfactant production (Rodrigues et al., 2006a) using four Lactobacillus strains, in which they obtained 1.45 and 1.7 g l1, respectively for above carbon sources. This indicated the potential

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Biosurfactant

10

Log CFU/ml

9 5

8 7

4

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Biosurfactant concentration (mg/ml)

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1 0

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Incubation time (h) Fig. 1. Growth and biosurfactant production by L. delbrueckii using peanut oil cake in fermentor.

use of peanut oil cake as a renewable cheaper carbon source for biosurfactant production. 3.2. Characterization of biosurfactant Biosurfactant produced by L. delbrueckii in this study was classified as a glycolipid with carbohydrate and lipid combination of 30%:70% (w/w). The molecular composition of the biosurfactant evaluated by FTIR revealed that the most important bands were located at 2962, 2924, and 2854 cm1 (for the CH aliphatic stretching), 1793 cm1 (for the C@O ester bond), 1061 cm1 (PII band:polysaccharides) and 766, 700 cm1 (for the CH2 group) and 3388 and 3696 cm1 (for OAH bonds) confirming the presence of glycolipid moieties (figure not shown here, see Supplementary data) (Thavasi et al., 2008; Rodrigues et al. 2006b). In addition, the mass spectrometric analysis of the biosurfactant (figure not shown here, see Supplementary data) also confirmed the above FTIR results with peaks observed at m/z = 326.5 and at 663.4 for lipid and glycolipid moieties, respectively (Thavasi et al., 2008). However, a detailed structural analysis of the biosurfactant produced by L. delbrueckii is needed. 3.3. Growth and biodegradation of crude oil with different concentration of fertilizer and biosurfactant Experiments conducted with different concentrations of fertilizer revealed that, 0.1% of fertilizer concentration resulted maximum degradation of crude oil (61.25%), and cell growth (9.04 Log CFU ml1) (Fig.2a and c). Beyond 0.1% there was no significant change observed in bacterial cell growth and biodegradation rate. As observations made in this experiment, Vyas and Dave (2010) reported that addition of excess nutrients beyond certain limit in bioremediation would have no impact on cell growth and biodegradation process and excess nutrient content can be toxic to cell growth. Thus, 0.1% fertilizer concentration was used as the optimum level in microcosm experiments carried out in this study. Among the different biosurfactant concentrations used in biodegradation experiments, maximum crude oil degradation and cell growth was observed with 0.1% biosurfactant concentration (Fig.2b and c). There was no significant increase in degradation activity observed at concentrations above 0.1%, which indicates that at this concentration sufficient emulsification of the crude oil occurred making it bioavailable for degradation. Biosurfactants are known for their potential emulsification activity even at very low concentration (Raza et al., 2009). Bacterial strain used in this study is a

biosurfactant producer and the additional initial biosurfactant added to the culture medium could be enough to facilitate the cell access to the crude oil followed with more biosurfactant production. In this study, we used the biosurfactant produced by the same strain in its biodegradation experiments; however a detailed investigation is needed to determine the effect of biosurfactant produced by one bacterium on other bacteria to select the potential biosurfactant, most useful for all biodegrading microbes. Bacterial cell growth in above experiments conducted with different concentrations of fertilizer and biosurfactant revealed that cell growth was in its maximum at 120 h of incubation (Fig 2a and b), which is similar to the observation made in this study for this strain during biosurfactant production (Fig. 1). Similar growth trend was reported by Thavasi et al. (2011) for three marine bacterial isolates, B. megaterium, C. kutscheri and Pseudomonas aeruginosa while culturing with different concentration fertilizer and biosurfactant in biodegradation experiments. 3.4. Biodegradation of crude oil in laboratory scale microcosm experiments Biodegradation of crude oil in laboratory scale microcosm experiment (Fig. 3a and b) showed maximum growth and biodegradation in set 4 (bacterial cells + crude oil + biosurfactant + fertilizer). The difference in biodegradation levels between sets 1 (bacterial cells + crude oil) and 4 was 20.25%. However there was not much difference between sets 3 (bacterial cells + crude oil + biosurfactant) and 4 (7%). The difference between sets 4 and 2 (bacterial cells + crude oil + fertilizer) was 12.5%. Interestingly, the difference in biodegradation rate between experiments with sets 1 (normal, bacterial cells + crude oil) and 3 (bacterial cells + crude oil + biosurfactant) was 13.25%, which is higher than the values obtained for sets 1 and 2 (7.15%). This indicated that the main factor promoting oil degradation was the presence of biosurfactant rather than the fertilizer. The improved biodegradation levels obtained with biosurfactant indicated that they represent the most efficient accelerators for hydrocarbon biodegradation through increasing the bioavailability of oil (Perfumo et al., 2010a, b). The use of biosurfactant in combination with fertilizer could reduce the actual amount of fertilizer to be added to polluted sites. In some studies, water soluble fertilizers encountered problem such as washing away and rapid dilution in aquatic environment. Maki et al. (2003) reported that fertilizers only stimulate the early stage degradation rate of the oil and that the final degradation efficiencies with fertilizers were not significantly

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different from those where no fertilizers were used. It is important however to keep in mind that nutrients or fertilizer use may be essential in some environments with insufficient nutrient levels. Gas chromatographic analysis of the degraded crude oil extracted from the culture medium commensurate with the degradation values obtained by quantitative fluorometric analysis.

Bacterium used in this study was able to break the compounds present in crude oil. Preferential degradation of compounds by L. delbrueckii revealed that compounds with carbon numbers C22 and C23 were completely degraded (data not shown). Similar selective degradation of compounds was reported by Thavasi et al. (2011) for three marine bacterial isolates – B. megaterium,

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Fig. 3. (a) Growth and (b) biodegradation of crude oil by L. delbrueckii in lab scale microcosm experiments – normal (without biosurfactant and fertilizer) (set 1), fertilizer (set 2), biosurfactant (set 3), and biosurfactant + fertilizer (set 4).

C. kutscheri and P. aeruginosa, while culturing them for crude oil degradation.

Table 1 Emulsification of hydrocarbons by biosurfactant isolated from L. delbrueckii. Hydrocarbons

3.5. BATH assay BATH assay results revealed that a high cell adherence of 93.2 ± 1.2% was found for L. delbrueckii cells with crude oil, which directly correlated with the biodegradation potential observed in this study for this strain. Similar high cell hydrophobicity and degradation reported by Thavasi et al. (2011) for P. aeruginosa support the results obtained in this study. Cell surface properties are important factors that determine the rate of degradation of hydrophobic substrates. Therefore, isolates with high hydrophobicity are likely to be more efficient degraders, as reported in this study for L. delbrueckii. Cell hydrophobicity is also an indication of biosurfactant production (Franzetti et al., 2009). 3.6. Emulsification assay Emulsification activity of biosurfactant isolated from L. delbrueckii in this study was comparatively less than the emulsification activity recorded with the standard chemical surfactant Triton X-100 (Table 1). However, when considering the advantages of biosurfactants over chemically synthesized surfactants such as

Waste motor lubricant oil Crude oil Peanut oil Kerosene Diesel Xylene Anthracene Naphthalene a

Emulsification activity (D610) Biosurfactanta

Triton X-100

1.85 ± 0.14 1.75 ± 0.07 1.45 ± 0.08 1.05 0.85 ± 0.05 0.55 ± 0.04 0.45 ± 0.11 0.40 ± 0.01

1.93 ± 0.08 1.85 ± 0.07 1.56 ± 0.04 1.12 ± 0.10 0.94 ± 0.02 0.78 ± 0.12 0.58 ± 0.01 0.63

Biosurfactant isolated from L. delbrueckii.

lower toxicity, biodegradability and ecological acceptability, their use in bioremediation activity becomes more favorable. Emulsification of different hydrocarbons by the biosurfactant was in the order of waste motor lubricant oil > crude oil > peanut oil > kerosene > diesel > xylene > anthracene > naphthalene. Crude oil therefore was the second compound that has been highly emulsified by biosurfactant used in this study, which coincided with the enhanced biodegradation of crude oil obtained in sets 3 and 4 in this study. It is evident from the above results that emulsification is an essential process in biodegradation.

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Influence on biodegradation and emulsification activity of the biosurfactant isolated in this study was comparatively less than earlier reports on emulsification and biodegradation of crude oil with biosurfactants isolated from marine bacterial isolates (Thavasi et al., 2010). But the ability to improve the degradation process and emulsification of hydrocarbons are still comparable with above report. A detailed study on influence of biosurfactant isolated from L. delbrueckii on biodegradation of crude oil with other potential oil degrading bacteria and its toxicity on those bacteria could make this biosurfactant more economically and environmentally viable. 4. Conclusion Biosurfactant produced by L. delbrueckii using peanut oil cake in this study showed its potential to be used in bioremediation process. Unlike medicinal applications, environmental application of biosurfactants needs comparatively less purity and high activity. In this study peanut oil cake was used as the carbon source for biosurfactant production. Even though the biosurfactant was not purified to its purest form and structurally not well characterized but the results on emulsification and biodegradation experiments revealed the potential use of this biosurfactant in bioremediation of hydrocarbon pollution. Which emphasize that for environmental applications the biosurfactants need not be pure and could be synthesized from a mixed cheaper carbon source like peanut oil cake used in this study. All these approaches will make the bioremediation process an economically and environmentally viable mitigation technology. Acknowledgements We thank the authorities of Annamalai University for providing the facilities and Department of Ocean Development (DOD) and Council of Scientific and Industrial Research (CSIR), Government of India for providing financial support. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.biortech.2010.11.071. References Banat, I.M., Franzetti, A., Gandolfi, I., Bestetti, G., Martinotti, M.G., Fracchia, L., Smyth, T.J., Marchant, R., 2010. Microbial biosurfactants production, applications and future potential. Appl. Microbiol. Biotechnol. 87, 427–444. Betts, R.P., Bankers, P., Banks, J.G., 1989. Rapid enumeration of viable microorganisms by staining and direct microscopy. Lett. Appl. Microbiol. 9, 199–202. Blaiotta, G., Moschetti, G., Simeoli, E., Andolfi, R., Villani, F., Coppola, S., 2001. Monitoring lactic acid bacteria strains during ‘Cacioricotta’ cheese production by restriction endonuclease analysis and pulsed-field gel electrophoresis. J. Dairy Res. 68, 139–144. Buddington, R., 2009. Using probiotics and prebiotics to manage the gastrointestinal tract ecosystem. In: Charalampopoulos, D., Rastall, R.A. (Eds.), Prebiotics and Probiotics Science and Technology. Springer Science + Business Media, New York, pp. 1–32. Dubois, M., Gilles, K.A., Hamilton, J.K., Rebers, P.A., Smith, F., 1956. Colorimetric method for determination of sugars and related substances. Anal. Chem. 28, 350–356. Folch, J.M., Lees, M., Stanly, H.S., 1956. A simple method for the isolation and quantification of total lipids from animal tissues. J. Biol. Chem. 226, 497–509. Fox, S.L., Bala, G.A., 2000. Production of surfactant from Bacillus subtilis ATCC 21332 using potato substrates. Bioresour. Technol. 75, 235–240. Franzetti, A., Caredda, P., Colla, P.L., Pintus, M., Tamburini, E., Papacchini, M., Bestetti, G., 2009. Cultural factors affecting biosurfactant production by Gordonia sp. BS29. Int. Biodeterioration Biodegrad. 63, 943–947. Franzetti, A., Tamburini, E., Banat, I.M., 2010. Application of biological surface active compounds in remediation technologies. In: Sen, R. (Ed.), Biosurfactants: Advances in Experimental Medicine and Biology, vol. 672. Springer-Verlag, Berlin Heidelberg, pp. 121–134.

Gołek, P., Bednarskil, W., Lewandowska, M., 2007. Characteristics of adhesive properties of Lactobacillus strains synthesising biosurfactants. Pol. J. Natur. Sc. 22, 333–342. Gudina, E.J., Teixeira, J.A., Rodrigues, L.R., 2010. Isolation and functional characterization of a biosurfactant produced by Lactobacillus paracasei. Colloids Surf. B. Biointerfaces 76, 298–304. Intergovernmental Oceanographic Commission Manuals and Guide No.13, 1982. Manual for Monitoring Oil and Dissolved/Dispersed Petroleum Hydrocarbons in Marine Waters and Beaches. UNESCO. Kiran, S.G., Thomas, T.A., Selvin, J., Sabarathnam, B., Lipton, A.P., 2010. Optimization and characterization of a new lipopeptide biosurfactant produced by marine Brevibacterium aureum MSA13 in solid state culture. Bioresour. Technol. 101, 2389–2396. Lowry, O.H., Rosebrough, N.J., Farr, A.L., Randall, R.J., 1951. Protein measurement with the Folin phenol reagent. J. Biol. Chem. 193, 265–275. Maki, H., Hirayama, N., Hiwatari, T., Kohata, K., Uchiyama, H., Watanabe, M., Yamasaki, F., Furuki, M., 2003. Crude oil bioremediation field experiment in the Sea of Japan. Mar. Poll. Bull. 47, 74–77. Makkar, R.S., Cameotra, S.S., 1999. Biosurfactant production by microorganisms on unconventional carbon sources – a review. J. Surf. Det. 2, 237–241. Moldes, A.B., Torrado, A.M., Barral, M.T., Domianguez, J.M., 2007. Evaluation of biosurfactant production from various agricultural residues by Lactobacillus pentosus. J. Agric. Food Chem. 55, 4481–4486. Omogbai, B.A., Ikenebomeh, M.J., Ojeaburu, S.I., 2005. Microbial utilization of stachyose in soymilk yogurt production. Afr. J. Biotechnol. 4, 905–908. Perfumo, A., Rancich, I., Banat, I.M., 2010a. Possibilities and challenges for biosurfactants use in petroleum industry. In: Sen, R. (Ed.), Biosurfactants: Advances in Experimental Medicine and Biology, vol. 672. Springer, Berlin, pp. 135–157. Perfumo, A., Smyth, T.J.P., Marchant, R., Banat, I.M., 2010b. Production and roles of biosurfactants and bioemulsifiers in accessing hydrophobic substrates. In: Timmis, K.N. (Ed.), Handbook of Hydrocarbon and Lipid Microbiology. SpringerVerlag, Berlin Heidelberg, pp. 1501–1512. Portilla-Rivera, O., Torrado, A., Dominguez, J.M., Moldes, A.B., 2008. Stability and emulsifying capacity of biosurfactants obtained from lignocellulosic sources using Lactobacillus pentosus. J. Agric. Food Chem. 56, 8074–8080. Raza, Z.A., Khalid, Z.M., Banat, I.M., 2009. Characterization of rhamnolipids produced by a Pseudomonas aeruginosa mutant strain grown on waste oils. J. Environ. Sci. Health Part A. 44, 1367–1373. Reid, G., Gan, B.S., She, Y.-M., Ens, W., Weinberger, S., Howard, J.C., 2002. Rapid identification of probiotic Lactobacillus biosurfactant proteins by proteinchip tandem mass spectrometry tryptic peptide sequencing. Appl. Environ. Microbiol. 68, 977–980. Rodrigues, L., Moldes, A., Teixeira, J., Oliveira, R., 2006a. Kinetic study of fermentative biosurfactant production by Lactobacillus strains. Biochem. Eng. J. 28, 109–116. Rodrigues, L.R., Teixeira, J.A., van der Mei, H.C., Oliveira, R., 2006b. Isolation and partial characterization of a biosurfactant produced by Streptococcus thermophilus A.. Colloids Surf. B. Biointerfaces 53, 105–112. Rodrigues, L.R., Teixeira, J.A., Oliveira, R., 2006c. Low-cost fermentative medium for biosurfactant production by probiotic bacteria. Biochem. Eng. J. 32, 135–142. Rosenberg, M., Gutnick, D., Rosenberg, E., 1980. Adherence to bacteria to hydrocarbons: a simple method for measuring cell-surface hydrophobicity. FEMS Microbiol. Lett. 9, 29–33. Satpute, S.K., Banat, I.M., Dhakephalkar, P.K., Banpurkar, A.G., Chopade, B.A., 2010. Biosurfactants, bioemulsifiers and exopolysaccharides from marine microorganisms. Biotechnol. Adv. 28, 436–450. Sheppard, J.D., Mulligan, C.N., 1987. The production of surfactin by Bacillus subtilis grown on peat hydrolysate. Appl. Microbiol. Biotechnol. 27, 110–116. Sivapathasekaran, C., Mukherjee, S., Ray, A., Gupta, A., Sen, R., 2010. Artificial neural network modeling and genetic algorithm based medium optimization for the improved production of marine biosurfactant. Bioresour. Technol. 101, 2884–2887. Teitelbaum, J.E., Walker, W.A., 2002. Nutritional impact of pre- and probiotics as protective gastrointestinal organisms. Annu. Rev. Nutr. 22, 107–138. Thavasi, R., 2006. Biosurfactants from marine hydrocarbonoclastic bacteria and their application in marine oil pollution abatement. Ph.D Thesis, Annamalai University, India. p. 162. Thavasi, R., Jayalakshmi, S., Balasubramanian, T., Banat, I.M., 2007. Biosurfactant production by Corynebacterium kutscheri from waste motor lubricant oil and peanut oil cake. Lett. Appl. Microbiol. 45, 686–691. Thavasi, R., Jayalakshmi, S., Balasubramanian, T., Banat, I.M., 2008. Production and characterization of a glycolipid biosurfactant from Bacillus megaterium. World J. Microbiol. Biotechnol. 24 (7), 917–925. Thavasi, R., Jayalakshmi, S., Banat, I.M., 2011. Effect of biosurfactant and fertilizer on biodegradation of crude oil by marine isolates of Bacillus megaterium, Corynebacterium kutscheri and Pseudomonas aeruginosa. Bioresour. Technol. 102 (2), 772–778. Vuyst, L.D., Leroy, F., 2007. Bacteriocins from lactic acid bacteria: production, purification, and food applications. J. Mol. Microbiol. Biotechnol. 13, 194–199. Vyas, T.K., Dave, B.P., 2010. Effect of addition of nitrogen, phosphorus and potassium fertilizers on biodegradation of crude oil by marine bacteria. Ind. J. Mar. Sci. 39, 143–150. Wee, Y.-J., Kim, J.-N., Yun, J.-S., Ryu, H.-W., 2005. Optimum conditions for the biological production of lactic acid by newly isolated lactic acid bacterium Lactobacillus sp. RKY2. Biotechnol. Bioprocess Eng. 10, 23–28.

Mar Biotechnol (2009) 11:551–556 DOI 10.1007/s10126-008-9162-1

SHORT COMMUNICATION

Biosurfactant Production by Azotobacter chroococcum Isolated from the Marine Environment R. Thavasi & V. R. M. Subramanyam Nambaru & S. Jayalakshmi & T. Balasubramanian & Ibrahim M. Banat

Received: 19 October 2007 / Accepted: 29 October 2008 / Published online: 26 November 2008 # Springer Science + Business Media, LLC 2008

Abstract Preliminary characterization of a biosurfactantproducing Azotobacter chroococcum isolated from marine environment showed maximum biomass and biosurfactant production at 120 and 132 h, respectively, at pH 8.0, 38°C, and 30‰ salinity utilizing a 2% carbon substrate. It grew and produced biosurfactant on crude oil, waste motor lubricant oil, and peanut oil cake. Peanut oil cake gave the highest biosurfactant production (4.6 mg/mL) under fermentation conditions. The biosurfactant product emulsified waste motor lubricant oil, crude oil, diesel, kerosene, naphthalene, anthracene, and xylene. Preliminary characterization of the biosurfactant using biochemical, Fourier transform infrared spectroscopy, and mass spectral analysis indicated that the biosurfactant was a lipopeptide with percentage lipid and protein proportion of 31.3:68.7. R. Thavasi (*) Department of Chemical and Biological Sciences, Polytechnic Institute of New York University, Six Metrotech Center, Brooklyn, NY 11201,, USA e-mail: [email protected] V. R. M. Subramanyam Nambaru : S. Jayalakshmi : T. Balasubramanian CAS in Marine Biology, Annamalai University, Parangipettai 608 502, Tamil Nadu, India V. R. M. Subramanyam Nambaru e-mail: [email protected] S. Jayalakshmi e-mail: [email protected] T. Balasubramanian e-mail: [email protected] I. M. Banat School of Biomedical Sciences, University of Ulster, Coleraine BT52 1SA, Northern Ireland, UK e-mail: [email protected]

Keywords Biosurfactant . Emulsification . Hydrocarbon . Crude oil . Waste motor lubricant oil . Peanut oil cake

Introduction Microbial biosurfactants are extracellular products containing both hydrophilic and hydrophobic moieties capable of reducing surface tension and facilitating hydrocarbon uptake and emulsification/dispersion. They can improve the bioavailability of hydrocarbons to the microbial cells by increasing the area of contact at the aqueous–hydrocarbon interface. This increases the rate of hydrocarbon dissolution and their utilization by microorganisms (Rahaman et al. 2002a; Vasileva-Tonkova and Gesheva 2005; Perfumo et al. 2006). Biosurfactants can be used in many processes involving emulsification, foaming, detergency, wetting, dispersion, and solubilization of hydrophobic compounds (Desai and Banat 1997). They have several advantages over the chemical surfactants, such as lower toxicity, higher biodegradability (Zajic et al. 1977), environmental compatibility (Georgiou et al. 1992), higher foaming (Razafindralambo et al. 1996), selectivity and specific activity at extreme temperatures, pH, and salinity (Velikonja and Kosaric 1993), and the ability to be synthesized from renewable feedstock (Desai and Banat 1997; Nitschke and Pastore 2004). Although biosurfactants have many interesting properties, their industrial importance depends on the cost and ease of production (Banat et al. 2000). Low yields are a major limitation for profitable industrial production and commercialization. Hence, in the present study, peanut oil cake and waste motor lubricant oil were selected as cheaper carbon sources for production. Peanut oil cake is a cheap carbohydrate, protein, and lipid-rich residue generated in large

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amounts during the production of peanut oil. Waste motor lubricant oil is waste oil drained from geared motor vehicles containing weathered hydrocarbon fractions. The aims of this study were to optimize biosurfactant production under fermentation condition using cheaper carbon sources and characterize the biosurfactant using biochemical, Fourier transform infrared spectroscopy (FTIR) and mass spectral analysis.

Materials and Methods

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Na2MoO4·2H2O 0.001, and NaCl 5.0 (Benson 1990) supplemented with crude oil or waste motor lubricant oil or peanut oil cake was used for optimization and production in a fermentor. The strain was cultured at different temperatures (30°C to 46°C), substrate concentrations (0.5% to 2.5%, w/v of crude oil, waste motor lubricant oil, and peanut oil cake), pH (5.0 to 9.0), and salinities (NaCl 0‰ to 40‰). All the experiments were carried out in 500-mL conical flasks containing 100 mL mineral salt medium. The culture was maintained in a water bath shaker at 150 rpm. Statistical analysis (analysis of variance, ANOVA) was carried out for all experiments.

Microorganism Biosurfactant Production in Fermentor Azotobacter chroococcum was isolated from water sample collected at Tuticorin harbor, Tamil Nadu, India (08°45′ N, 78°13′ E) and characterized by Thavasi et al. (2006). Optimization of Biosurfactant Production in Shake Flasks Mineral salt medium containing (g/L) K2HPO4 1.0, MgSO4·7H2O 0.2, FeSO4·7H2O 0.05, CaCl2·2H2O 0.1,

A 3-L laboratory fermentor (Scigenics, India Pvt. Ltd., Chennai) with a 2.1-L working volume was used. The culture conditions were: pH 8.0, temperature 38°C, salinity 30‰ and 2.0% substrate (crude oil, peanut oil cake, and waste motor lubricant oil), stirring at 350 rpm, 8.0 mg/L dissolved oxygen concentration, and 0.5-kg/cm2 pressure.

Fig. 1 Growth and biosurfactant production of A. chroococcum at different pH (a, b), temperatures (c, d), and salinities (e, f)

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Growth, Biochemical Analysis, and Emulsification Measurements Five-milliliter culture broth samples were collected at 12-h intervals for a period of 168 h for gravimetrical biomass measurements in milligrams per milliliter dry weight as described by Thavasi et al. (2006). Carbohydrate content of the biosurfactant was determined by the phenol sulfuric acid method (Dubois et al. 1956) using D-glucose as a standard. Protein content was determined by the method of Lowry et al. (1951) using bovine serum albumin as a standard, and lipid content was estimated adopting the procedure of Folch et al. (1956). To estimate emulsification activity, purified biosurfactant (1 mg/mL) was dissolved in 5 mL of Tris buffer (pH 8.0) in a 30-mL screw-capped test tube. Five milligrams of hydrocarbon (waste motor lubricant oil, crude oil, diesel, kerosene, naphthalene, anthracene, or xylene) was added to the above solution which was shaken well for 20 min and allowed to stand for 20 min. The optical density of the mixture was then measured at 610 nm, and the results were expressed as OD610 (Rosenberg et al. 1979).

Fig. 2 Growth and biosurfactant production by A. chroococcum at different concentrations of crude oil (a, b), waste motor lubricant oil (c, d), and peanut oil cake (e, f)

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Purification of Biosurfactant The culture broth was centrifuged at 6,000 rpm for 20 min and extracted with chloroform and methanol (2:1, v/v). The solvents were removed by rotary evaporation and the residue purified on a silica gel (60–120 mesh) column eluting with a chloroform/methanol gradient ranging from 20:1 to 2:1 (v/v), collecting ten fractions. The fractions were pooled and the solvents evaporated; the resulting residue was dialyzed against distilled water and lyophilized as reported by Li et al. (1984). Weight of the biosurfactant was expressed in terms of milligrams per milliliter (dry weight). Fourier Transform Infrared Spectroscopy and Mass Spectrometric Analysis FTIR is most useful for identifying types of chemical bonds (functional groups) and therefore can be used to elucidate some components of an unknown mixture. Ten milligrams of freeze-dried pure biosurfactant was ground with 100 mg of KBr and pressed with 7,500 kg for 30 s to obtain translucent pellets. Infrared absorption spectra were

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Fig. 3 Biosurfactant production (a) and growth (b) of A. chrooroccum in a fermentor

recorded on a Thermo Niocolet, AVATAR 330 FTIR system with a spectral resolution and wavenumber accuracy of 4 and 0.01 cm−1, respectively. All measurements consisted of 500 scans, and a KBr pellet was used as background reference. Mass spectrometric analysis was carried out as described by Rahaman et al. (2002b). The purified biosurfactant was dissolved in methanol and mixed thoroughly. The mass spectrometric analysis of the biosurfactant was carried out in a LCQTM quadrupole ion trap mass spectrometer (Finnigan MAT, San Jose, CA, USA) utilizing electrospray ionization (ESI). Standard solutions and samples under investigation were infused into the mass spectrometer at a flow rate of 10 μL/min. In the ESI, nitrogen and auxiliary gas flows were maintained at 50 and 5 mL/min, respectively, and referred to arbitrary values set by the software. The heated capillary temperature was 250°C and the spray voltage set to 5 kV. Negative ion mode was used and scanning was done at 50–2,000 m/z range.

Fig. 4 FTIR spectrum of biosurfactant produced by A. chroococcum

Results and Discussion Growth and biosurfactant production followed similar patterns on crude oil with maximum detected at pH 8.0 (Fig. 1a,b), at a temperature of 38°C (Fig. 1c,d), and a salinity of 30‰ (Fig. 1e,f). Biomass and biosurfactant production were in the range of 1.26 to 3.12 and 0.98 to 2.97 mg/mL, respectively. The strain tolerated 30‰ NaCl, which is higher than the results reported by Page (1986) at 23‰ for such a strain. Growth and biosurfactant production on crude oil (Fig. 2a,b), waste motor lubricant oil (Fig. 2c,d), and peanut oil cake (Fig. 2e,f) showed a maximum at 2.0% substrate concentration for both the substrates tested (1.9 and 3.6 mg/mL, respectively). Biosurfactant production on peanut oil cake both in flask and in fermentor conditions was higher than on crude oil and waste motor lubricant oil, which indicated its suitability as a cheaper substrate for large-scale biosurfactant production. In all culture condi-

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tions tested, biosurfactant concentration was highest at the early stationary phase, 120–132 h. Higher concentration of biosurfactant at the early stationary phase may be due to the release of cell-bound biosurfactant into the culture broth which led to a rise in extracellular biosurfactant concentration (Goldman et al. 1982). Statistical analysis (ANOVA) of the influence of pH, temperature, salinity, and substrate concentration on biosurfactant production showed a high significance (p=0.05). Biomass and biosurfactant production were higher under fermentation conditions ranging from 4.86 to 2.15 mg/mL for crude oil, 5.12 to 2.82 mg/mL for waste motor lubricant oil, and 8.7 to 4.6 mg/mL for peanut oil cake, respectively (Fig. 3a,b). These results indicated the possibility of biosurfactant production at industrial scale using a cheaper cost substrate such as peanut oil cake. In other studies, vegetable oils had been used as carbon sources for biosurfactant production (Rahaman et al. 2002b; Bednarski et al. 2004), whereas in the present study, peanut oil cake is used, which may be more economically viable for largescale production. Emulsification of different hydrocarbons by the biosurfactant was in the order of waste motor lubricant oil > crude oil > kerosene > diesel > naphthalene > anthracene > xylene, and the emulsification activity (OD610) was 1.51, 1.43, 0.67, 0.36, 0.33, and 0.42, respectively. FernandezLinares et al. (1996) reported similar emulsification results by two marine strains, Pseudomonas nautical and Marinobacter hydrocarboclasticus, and concluded that emulsification is a major essential process in alkane biodegradation. Emulsification of various hydrocarbons by the biosurfactant used in the present study indicated the possibility of their application in the remediation of different types of hydrocarbon pollution either as a means of their direct removal or as a promoter of biodegradation (Thavasi et al. 2007). The biosurfactant component of A. chroococcum is a lipopeptide with a lipid and protein combination of 31.30:68.69%. The FTIR analysis of biosurfactant revealed that wavenumbers 2,852, 2,923, 1,421, and 1,465 cm−1 resulting from the C–H stretching mode suggested the presence of aliphatic chain. The presence of N–H and CO– N bonds was indicated by wavenumbers 3,383 and 1,647 cm−1, respectively. The C–O bonds were observed at 1,058 cm−1. The obtained wavenumbers are consistent with the lipopeptide moieties of the biosurfactant (Fig. 4). The FTIR analysis of the biosurfactant was similar to the results obtained by earlier workers (Onwurah 1999; Kuiper et al. 2004; Das and Mukherjee 2007) who used FTIR as an analytical tool for the characterization of biosurfactants. The mass spectral analysis of the biosurfactant showed ionized compounds with molecular weights of m/z=326.5, 663.4, and 1,347.3 which may represent the lipid and

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protein molecules, respectively. Similar results were obtained by Kuiper et al. (2004) using Pseudomonas putida with a lipopeptide biosurfactant and by Kalinovskaya et al. (1995) for surfactin, a lipopeptide biosurfactant produced by Bacillus pumilus. In conclusion, biosurfactants can be produced growing A. chroococcum on economically cheaper carbon source such as peanut oil cake or waste motor lubricant oil for application in oil pollution removal or bioremediation. Acknowledgments We thank the authorities of Annamalai University for providing the facilities and DOD and CSIR, Government of India for financial support.

References Banat IM, Makkar RS, Cameotra SS (2000) Potential commercial applications of microbial surfactants. Appl Microbiol Biotechnol 53:495–508 Bednarski W, Adamczak M, Tomasik J, Plaszczyk M (2004) Application of oil refinery waste in the biosynthesis of glycolipids by yeast. Bioresour Technol 95:15–18 Benson JH (1990) Microbial applications: a laboratory manual in general microbiology. WM.C. Brown, Dubuque Das K, Mukherjee AK (2007) Comparison of lipopeptide biosurfactants production by Bacillus subtilis strains in submerged and solid state fermentation systems using a cheap carbon source: some industrial applications of biosurfactants. Process Biochem 42:1191–1199 Desai JD, Banat IM (1997) Microbial production of surfactants and their commercial potential. Microbiol Mol Biol Rev 61:47–64 Dubois M, Gilles KA, Hamilton JK, Rebers PA, Smith F (1956) Colorimetric method for determination of sugars and related substances. Anal Chem 28:350–356 Fernandez-Linares L, Acquaviva M, Bertrand J-C, Gauthier M (1996) Effect of sodium chloride concentration on growth and degradation of eicosane by marine halotolerent bacterium Marinobacter hydrocarbonoclastieus. Appl Microbiol 19:113–121 Folch JM, Lees M, Stanly HS (1956) A simple method for the isolation and quantification of total lipids from animal tissues. J Biol Chem 226:497–509 Georgiou G, Lin SC, Sharma MM (1992) Surface-active compounds from microorganism. Biotechnology 10:60–65 Goldman S, Shabtai Y, Rubinovitz C, Rosenberg E, Gutnick DL (1982) Emulsan in Acinetobacter calcoaceticus RAG-I: distribution of cell-free and cell associated cross-reacting materials. Appl Environ Microbiol 44:165–170 Kalinovskaya N, Kuznetsova T, Rashkes Ya, Mil’grom Yu, Mil’grom E, Willis R, Wood A, Kurtz H, Carabedian C, Murphy P, Elyakov G (1995) Surfactin-like structures of five cyclic despsipeptises from the marine isolates of Bacillus pumilus. Russ Chem Bull 44:951–955 (English translation) Kuiper I, Lagendijk EL, Pickford R, Derrick JP, Lamers GEM, Thomas-Oates JE, Lugtenberg BJJ, Bloemberg GV (2004) Characterization of two Pseudomonas putida lipopeptide biosurfactants, putisolvin I and II, which inhibit biofilm formation and break down existing biofilms. Mol Microbiol 51:97–113 Li Z-Y, Lang S, Wagner F, Witte L, Wary V (1984) Formation and identification of interfacial-active glycolipids from resting microbial cells. Appl Environ Microbiol 48:610–617

556 Lowry OH, Rosebrough NJ, Farr AL, Randall RJ (1951) Protein measurement with the Folin phenol reagent. J Biol Chem 193:265–275 Nitschke M, Pastore GM (2004) Biosurfactant production by Bacillus subtilis using cassava processing effluent. Appl Biochem Biotechnol 112:163–172 Onwurah INE (1999) Role of diazotropic bacteria in the bioremediation of crude oil-polluted soil. J Chem Technol Biotechnol 74:957–964 Page WJ (1986) Sodium-dependent growth of Azotobactor chroococcum. Appl Environ Microbiol 51:510–514 Perfumo A, Banat IM, Canganella F, Marchant R (2006) Rhamnolipid production by a novel thermotolerant hydrocarbon-degrading Pseudomonas aeruginosa AP02-1. Appl Microbiol Biotechnol 72:132–138 Rahaman KSM, Banat IM, Thahira J, Thayumanavan T, Lakshmanaperumalsamy P (2002a) Bioremediation of gasoline contaminated soil by bacterial consortium with poultry litter, coir pith and rhamnolipid biosurfactant. Bioresour Technol 81:25–32 Rahaman KSM, Rahman TJ, McClean S, Marchant R, Banat IM (2002b) Rhamnolipid biosurfactant production by strains of Pseudomonas aeruginosa using low-cost raw materials. Biotechnol Prog 18:1277–1281

Mar Biotechnol (2009) 11:551–556 Razafindralambo H, Paquot M, Baniel A, Popineau Y, Hbid C, Jacques P, Thonart P (1996) Foaming properties of surfactin, a lipopeptide biosurfactant from Bacillus subtilis. J Am Oil Chem Soc 73:149–151 Rosenberg E, Zuckerberg A, Rubinovitz C, Gutnick DL (1979) Emulsifier of Arthrobacter RAG-I: isolation and emulsifying properties. Appl Environ Microbiol 37:402–408 Thavasi R, Jayalakshmi S, Balasubramanian T, Banat IM (2006) Biodegradation of crude oil by nitrogen fixing marine bacteria Azotobacter chroococcum. Res J Microbiol 1:401–408 Thavasi R, Jayalakshmi S, Balasubramanian T, Banat IM (2007) Biosurfactant production by Corynebacterium kutscheri from waste motor lubricant oil and peanut oil cake. Lett Appl Microbiol 45:686–691 Vasileva-Tonkova E, Gesheva V (2005) Glycolipids produced by Antarctic Nocardioides sp. during growth on n-paraffin. Process Biochem 40:2387–2391 Velikonja J, Kosaric N (1993) Biosurfactant in food application. In: Kosaric N (ed) Biosurfactants: production, properties, applications. Marcel Dekker, New York Zajic JE, Gignard H, Gerson DF (1977) Properties and biodegradation of a bioemulsifier from Corynebacterium hydrocarbonoclasticus. Biotechnol Bioeng 91:1303–1320

World J Microbiol Biotechnol (2008) 24:917–925 DOI 10.1007/s11274-007-9609-y

ORIGINAL PAPER

Production and characterization of a glycolipid biosurfactant from Bacillus megaterium using economically cheaper sources R. Thavasi Æ S. Jayalakshmi Æ T. Balasubramanian Æ Ibrahim M. Banat

Received: 7 May 2007 / Accepted: 27 August 2007 / Published online: 21 November 2007
Springer Science+Business Media B.V. 2007

Abstract Criteria selected for screening of biosurfactant production by Bacillus megaterium were hemolytic assay, bacterial cell hydrophobicity and the drop-collapse test. The data on hemolytic activity, bacterial cell adherence with crude oil and the drop-collapse test confirmed the biosurfactant-producing ability of the strain. Accordingly, the strain was cultured at different temperatures, pH values, salinity and substrate (crude oil) concentration in mineral salt medium to establish the optimum culture conditions, and it was shown that 38°C, 2.0% of substrate concentration, pH 8.0 and 30% of salt concentration were optimal for maximum growth and biosurfactant production. Laboratory scale biosurfactant production in a fermentor was done with crude oil and cheaper carbon sources like waste motor lubricant oil and peanut oil cake, and the highest biosurfactant production was found with peanut oil cake. Characterization of partially purified biosurfactant inferred that it was a glycolipid with emulsification potential of waste motor lubricant oil, crude oil, peanut oil, diesel, kerosene, naphthalene, anthracene and xylene.

R. Thavasi
S. Jayalakshmi (&)
T. Balasubramanian CAS in Marine Biology, Annamalai University, Parangipettai 608 502, Tamil Nadu, India e-mail: [email protected]; [email protected] T. Balasubramanian e-mail: [email protected] I. M. Banat School of Biomedical Sciences, University of Ulster, Coleraine BT52 1SA, Northern Ireland, UK e-mail: [email protected]

Keywords Biosurfactants
Emulsification
Biodegradation
Crude oil
Waste lubricant oil
Peanut oil

Introduction Many microorganisms produce extracellular or membraneassociated surface-active compounds (biosurfactants). Most microbial surfactants are complex molecules, comprising different structures that include lipopeptides, glycolipids, polysaccharide protein complexes, fatty acids and phospholipids. In the past few decades, biosurfactants have gained attention because of their advantages such as biodegradability, low toxicity, ecological acceptability and ability to be produced from renewable and cheaper substrates (Desai and Banat 1997; Nitschke and Pastore 2004). The range of industrial applications of biosurfactants includes enhanced oil recovery, crude oil drilling, lubricants, bioremediation of pollutants, health care and food processing (Banat et al. 2000). Among the many classes of biosurfactants, lipopeptides from Bacillus subtilis are particularly interesting because of their high surface activity and therapeutic potential (Besson and Michel 1992; Sandrin et al. 1990). Although biosurfactants exhibit such important advantages, they have not been yet employed extensively in industry because of relatively high production costs. One possible strategy for reducing costs is the utilization of alternative substrates such as agroindustrial wastes (Mercade and Manresa 1994). The main problem related to use of alternative substrates as culture medium is to find a waste with the right balance of nutrients that permits cell growth and product accumulation (Makkar and Cameotra 1999). Molasses (Makkar and Cameotra 1999), peat

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hydrolysate (Sheppard and Mulligan 1987) and potato processing effluents (Fox and Bala 2000) are examples of alternative substrates that have been suggested for biosurfactant production by Bacillus megaterium. The establishment of waste-based medium for biosurfactant production also faces another problem, once the kind and the properties of final product are dependent on the composition of the culture medium (Besson and Michel 1992). Although biosurfactants have many interesting properties, their industrial importance depends upon the ease of production (Cooper and Goldenberg 1987). Low yield of biosurfactant is a major limitation to influencing its commercialization. Hence, in the present study peanut oil cake and waste motor lubricant oil were tried as cheaper carbon sources for biosurfactant production. Peanut oil cake is a carbohydrate, protein and lipid rich residue generated in large amounts during the production of peanut oil, and the cost of this cake is very low when compared to other carbon sources like glucose, fructose, crude oil and other hydrocarbons. Waste motor lubricant oil is waste oil drained from geared motor vehicles after a long run, which contains weathered hydrocarbon fractions and may be useful for biosurfactant production. The present investigation was conducted with the following objectives: screening of B. megaterium for biosurfactant production, optimization of biosurfactant production, laboratory scale production of biosurfactant using crude oil, waste motor lubricant oil and peanut oil cake, characterization of biosurfactant and estimation of emulsification activity.

Materials and methods

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around the colony. The diameter of the zone of clearance is a qualitative method used as an indicator of biosurfactant production (Mulligan et al. 1984; Rodrigues et al. 2006).

Bacterial adhesion to hydrocarbons (BATH) Cell hydrophobicity was measured by bacterial adherence to hydrocarbons (BATH) according to a method similar to that described by Rosenberg et al. (1980). The cells were washed twice and suspended in a buffer salt solution (g/l 16.9 K2HPO4, 7.3 KH2PO4) to give an OD at 600 nm of *0.5. The cell suspension (2 ml) with 100 ll crude oil added was vortex-shaken for 3 min in test tubes (10 9 100 mm). After shaking, crude oil and aqueous phase were allowed to separate for 1 h. The OD of the aqueous phase was then measured at 600 nm in a spectrophotometer (Varian, Cary Eclipse, Spectrophotometer, Australia). Hydrophobicity is expressed as the percentage of cell adherence to crude oil calculated as follows: 100 9 (1 - OD of the aqueous phase/OD of the initial cell suspension). For a given sample, three independent determinations were made and the mean value was accounted.

Visualization of bacteria in the oil droplets A few drops of 2-(4-iodophenyl)-3-(4-nitrophenyl)-5-phenyltetrazolium chloride (INT) solution was added to the BATH assay culture broth and observed under the microscope. The INT turned red if it was reduced inside the cells, indicating the viability and adherence of cells with crude oil droplets (Betts et al. 1989).

Microorganism B. megaterium was isolated from water sample collected at Tuticorin harbor (08°450 N; 78°130 E) South East coast of India using Bushnell–Haas agar with 0.1% of crude oil and identified to the species level by Thavasi and Jayalakshmi (2003) following Bergey’s Manual of determinative bacteriology (Buchanan et al. 1974).

Screening for biosurfactant production Hemolytic activity Hemolytic assay was performed in blood agar plates (Mulligan et al. 1984). Fifty microliter broth culture of B. megaterium was spot-inoculated on to blood agar plates and incubated for 48 h at 37°C. The plates were visually inspected for zone of clearance (hemolysis)

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Drop-collapse test B. megaterium was cultured in mineral medium with 0.1% crude oil for 48 h. Screening of biosurfactant production was performed using the qualitative drop-collapse test described by Jain et al. (1991) as modified by Bodour and Maier (1998). Crude oil was used in this test, 2 ll of oil was applied to the well regions delimited on the covers of 96-well microplates and these were left to equilibrate for 24 h. Five microliters of the 48 h culture, before and after centrifugation at 12,000g for 5 min to remove cells, was transferred to the oil-coated well regions and drop size was observed 1 min later with the aid of a magnifying glass. A result was considered positive for biosurfactant production when the drop diameter was at least 1 mm larger than that produced by deionized water (negative control).

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Optimization of biosurfactant production To find the optimum culture conditions for biosurfactant production, the strain was cultured at different temperature (28–46°C), substrate concentrations (crude oil 0.1–4.5% v/ v) and pH values (5.0–9.5). The impact of salinity on biosurfactant production was evaluated using various concentration of NaCl (0–40% w/v). All the experiments were carried out in 500-ml conical flasks containing 100 ml of mineral salt medium (g/l) NaNO3 14, KH2PO4 2, K2HPO4 4, KCl 0.2, MgSO4
7H2O 1, CaCl2
2H2O 0.02, FeSO4
7H2O 0.024, NaCl 5.0 and 0.5 ml of trace element solution containing (g/l) H3BO3 0.26, CuSO4
5H2O 0.5, MnSO4
H2O 0.5, MoNaO4
2H2O 0.06, ZnSO4
7H2O 0.7. The culture flasks were maintained in a water bath shaker at 150 rev min-1.

Biosurfactant production in fermentor Laboratory scale biosurfactant production was performed in a 3 l laboratory fermentor (Scigenics, India Pvt. Ltd. Chennai) with a working volume of 2.1 l. The culture conditions were as follows: pH 8.0, temperature 38°C, salinity (NaCl) 30% and 2.0% substrate concentration, 350 rev min-1 of agitation and 6.0 mg l-1 dissolved oxygen (DO) concentration. Substrates used were crude oil, peanut oil cake and waste motor lubricant oil.

Estimation of growth Two milliliters of culture broth was collected at 12 h intervals for a period of 168 h and the biomass was estimated gravimetrically. For gravimetric estimation of biomass 1 ml of broth culture was taken and allowed to stand for 20 min. When the oil phase separated, the bottom phase with cells was siphoned out and filtered through a 0.45lm sized Millipore filter. The filter with cells was dried at 120°C in a hot air oven for a period of 24 h and weighed; a control was maintained to exclude the weight of crude oil adhered to the filter. Biomass was quoted in terms of mg/ml (dry weight).

Purification of biosurfactant Five milliliters of culture broth was centrifuged at 6,000 rev min-1 for 20 min and extracted with chloroform and methanol (2:1 v/v). The solvents were removed by rotary evaporation and the residue was purified in a silica gel (60–120 mesh) column and the elutions were made with chloroform and methanol ranging from 20:1 to 2:1 v/v in a gradient manner and 10 fractions were obtained. The

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fractions were pooled and the solvents were evaporated, the resulting residue was dialysed against distilled water and lyophilized (Li et al. 1984). Weight of the biosurfactant was expressed in terms of mg/ml (dry weight).

Estimation of emulsification activity Hydrocarbons used for testing emulsification ability of biosurfactant were waste motor lubricant oil, crude oil, diesel, kerosene, naphthalene, anthracene, xylene and vegetable oil like peanut oil. Purified biosurfactant (1 mg/ ml) was dissolved in 5 ml of Tris buffer (pH 8.0) in a 30 ml screw-capped test tube. Five milligram of hydrocarbon was added to the above solution and vortex-shaken for 20 min and the mixture was allowed to stand for 20 min. The optical density of the mixture was measured at 610 nm in a spectrophotometer (Varian, Cary Eclipse, Spectrophotometer, Australia) and the results were expressed as D610 (Rosenberg et al. 1979). A control was maintained with buffer and crude oil and the results (D610) were subtracted from the control.

Characterization of biosurfactant Biochemical composition of biosurfactant Chemical composition of the biosurfactant was analysed following standard methods. Carbohydrate content of the biosurfactant was determined by the phenol–sulfuric acid method (Dubois et al. 1956) using D-glucose as a standard. Protein content was determined by the Lowry et al. (1951) method using bovine serum albumin as a standard and lipid content was estimated adopting the procedure of Folch et al. (1956).

Fourier transform infrared spectroscopy Fourier transform infrared spectroscopy (FTIR) is most useful for identifying types of chemical bonds (functional groups), therefore can be used to elucidate some components of an unknown mixture. The molecular characterization was performed using biosurfactant. One milligram of freeze-dried partially purified biosurfactant was ground with 100 mg of KBr and pressed with 7500 kg for 30 s to obtain translucent pellets. Infrared absorption spectra were recorded on a Thermo Niocolet, AVATAR 330 FTIR system with a spectral resolution and wave number accuracy of 4 and 0.01 cm-1, respectively. All measurements consisted of 500 scans, and a KBr pellet was used as background reference.

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Results and discussion Screening for biosurfactant production All the three tests performed to screen the strain for biosurfactant production revealed: a 1 cm zone of clearance (hemolysis) around the colony in blood agar plate, 42.3% of cell adherence with crude oil in the BATH assay and positive results in the drop collapse test, thereby confirming the biosurfactant-producing ability of the strain. Hemolysis was included in this study since it is widely used to screen biosurfactant production and in some cases, it is the sole method used (Banat 1993; Yonebayashi et al. 2000). Mulligan et al. (1984) recommended blood agar lysis as a preliminary screening method for biosurfactant production. The hemolytic activity of biosurfactants was first discovered when Bernheimer and Avigad (1970) reported that the biosurfactant produced by B. subtilis, surfactin, lysed red blood cells. Blood agar lysis has been used to quantify surfactin (Moran et al. 2002) and rhamnolipids (Johnson and Boese-Marrazzo 1980) and has been used to screen biosurfactant production by new isolates (Carrillo et al. 1996; Banat 1993). Carrillo et al. (1996) found an association between hemolytic activity and surfactant production, and they recommended the use of blood agar lysis as a primary method to screen biosurfactant production. None of the studies reported in the literature (Moran et al. 2002; Johnson and Boese-Marrazzo 1980; Carrillo et al. 1996; Banat 1993) mention the possibility of biosurfactant production without a hemolytic activity. However, in some cases hemolytic assay excluded many good biosurfactant producers (Youssef et al. 2004); hence in the present investigation the BATH assay and drop collapse test with crude oil were also done to confirm biosurfactant production. The BATH assay was confirmed by visualization of cells adhered to crude oil confirmed the affinity of cells

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Optimization of biosurfactant production The strain was able to grow and produce biosurfactant at all tested temperature, pH, substrate and salt concentrations, but optimum biosurfactant production was observed at 38°C temperature (Fig. 1a and b) 2.0% of substrate concentration (Fig. 2a and b), pH 8.0 (Fig. 3a and b) and 30% of salt concentration (Fig. 4a and b). The biomass and biosurfactant production were in the range of 3.24–3.58 and 1.1–1.4 mg/ml respectively. Growth curve and biosurfactant production in all the culture conditions revealed that maximum biomass and biosurfactant production occurred at 120 and 132 h respectively. Biosurfactant concentration was higher at early stationary phase (i.e.) at 132 h, which may be due to the release of cell-bound

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Biosurfactant was dissolved in methanol and mixed thoroughly. The mass spectrometric analysis of the biosurfactant was carried out in a LCQTM quadrupole iontrap mass spectrometer (Finnigan MAT, San Jose, California, USA) utilizing electrospray ionization (ESI). Standard solutions and samples under investigation were infused into the mass spectrometer at a flow rate of 10 ll/ min. In the ESI, nitrogen and auxiliary gas flows were maintained at 50 and 5 ml/min respectively and refer to arbitrary values set by the software. The heated capillary temperature was 250°C and the spray voltage was set to 5 kV. Negative ion mode was used and scanning was done at 50–2,000 m/z range.

towards crude oil facilitated by producing biosurfactant. Both the drop-collapse and visualization of cells adhered to crude oil have several advantages in requiring a small volume of samples, are rapid and easy to carry out, and do not require specialized equipment.

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biosurfactant into the culture broth which led to a rise in extracellular biosurfactant concentration (Goldman et al. 1982; Deziel et al. 1999). It shows the growth-related pattern of biosurfactant production. In all the experiments, below and above the optimum culture conditions, reduction in biosurfactant concentration was observed. Statistical analysis (ANOVA) of the influence of pH, temperature, salinity and crude oil concentration on biosurfactant production showed high significance (P = 0.05).

Biosurfactant production in fermentor In order to economize the biosurfactant production, cheaper carbon sources like waste motor oil and peanut oil cake were used in fermentor. In the fermentor similar results were obtained as in optimization, i.e. biomass and biosurfactant production were high at 120 and 132 h respectively. Among the three substrates used, namely crude oil, waste motor lubricant oil and peanut oil cake (Fig. 5a and b), biomass and biosurfactant production was highest with peanut oil cake (10.5 and 7.8 mg/ml). Waste motor lubricant oil and peanut oil cake used in the present

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study were economically more viable than crude oil. Youssef et al. (2007) reported a maximum biosurfactant production of 90 mg/l by B. subtilis using glucose as carbon source. Cooper and Goldenberg (1987) reported 1.6 g/l of biosurfactant production by B. cereus using sucrose as a carbon source. Where as in the present study we have used waste motor oil and peanut oil cake as carbon sources, biosurfactant production also found higher than their study (7.8 mg/ml) and economically cheaper. The possibility of biosurfactant production using cheaper carbon sources was already reported by earlier workers; Mercade et al. (1993) used olive oil mill effluent and animal fat, frying oil by Deshpande and Daniels (1995), molasses by Benincasa et al. (2002), and starch-rich wastes by Nitschke and Pastore (2004) supporting the present study on use of renewable carbon sources for biosurfactant production. Biosurfactant produced in the present study using waste motor lubricant oil and peanut oil cake showed good emulsification activity against seven different hydrocarbons and peanut oil. Further it encouraged the aim of the present study to produce biosurfactants from cheaper carbon sources with the emulsification property of seven different hydrocarbons.

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Time (h) 0‰ 25‰

5‰ 30‰

10‰ 35‰

60

72

84

96 108 120 132 144 156 168

Time (h) 15‰ 40‰

20‰

Fig. 4 (a) Growth and (b) Biosurfactant production of B. megaterium at different NaCl concentration (%) using crude oil

Estimation of emulsification activity Biosurfactant isolated from B. megaterium showed maximum emulsification activity against waste motor lubricant oil. Emulsification of hydrocarbons and peanut oil were in the order of waste motor lubricant oil [ crude oil [ peanut oil [ kerosene [ diesel [ xylene [ naphthalene [ anthracene and the emulsification activity (D610) was 1.86, 1.72, 1.42, 1.01, 0.85, 0.53, 0.46 and 0.42 respectively. These results showed that, biosurfactant produced from a hydrocarbon substrate can emulsify different hydrocarbons to a greater extent which confirmed its applicability against different hydrocarbon pollution. Emulsification enhances the biodegradation of hydrocarbons by increasing their bioavailability to the microbes involved. Juwarkar and Khirsagar (1991) also reported the emulsification of crude oil, xylene, pristine, n-octane, ndecane, n-hexadecane and n-dodecane by an unidentified marine bacterium. Fernandez-Linares et al. (1996) reported similar emulsification results by two marine strains, Pseudomonas nautica and Marinobacter hydrocarboclasticus and concluded that emulsification is a major essential process in alkane biodegradation. Emulsification of crude

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48

Crude oil

Waste m otor lubricant oil

Peanut oil cake

Fig. 5 (a) Biosurfactant production and (b) Growth of B. megaterium using crude oil, waste motor oil and peanut oil cake in fermentor

oil by an alkane-oxidizing Rhodococcus species isolated from seawater reported by Bredholt et al. (2002) also supported the present study on emulsification of different hydrocarbons.

Characterization of the biosurfactant The biosurfactant produced by B. megaterium was classified as a glycolipid with carbohydrate and lipid combination of 28:70%. The molecular composition of the biosurfactant was evaluated by FTIR. Figure 6 represents the spectra of the freeze-dried samples. The most important bands were located at 2929 cm-1 (for the CH aliphatic stretching), 1700 cm-1 (for the C=O ester bond), 1066 cm-1 (PII band: polysaccharides) and 764, 699 cm-1 (for the CH2 group) and 3342 cm-1 (for O–H bonds) confirming the presence of glycolipid moieties (Tuleva et al. 2001; Tahzibi et al. 2004; Rodrigues et al. 2006). In addition, the mass spectrometric analysis of the biosurfactant (Fig. 7) also confirmed the above results with peaks observed at m/z = 326.5, 413.3, 429.3 for lipids and at 663.4 for carbohydrate moieties (Deziel et al. 1999; Rahman et al. 2002). Earlier reports on chemical composition

World J Microbiol Biotechnol (2008) 24:917–925

923

Fig. 6 FTIR spectrum of biosurfactant produced by B. megaterium

india1#1-13 RT:0.01-0.41 AV: 13 NL:2.04E5 F: + c Full ms [ 300.00-2000.00] 663.3

100 95 90 85 80

Relative Abundance

75 70 65 60 55 50 45

664.4

40 35 30 326.5 25 20

551.5 607.4

369.3

15

413.3

537.4 495.4

10

677.5

805.6 881.6 943.9 992.0

1125.9 1174.9

5

300

400

500

600

700

800

900

1000

1100

1200

1243.4 1371.8

1300

1400

1462.7

1500

1602.7

1600

1741.5 1826.3 1897.6 1993.6

1700

1800

1900

2000

m/z

Fig. 7 Mass spectrum of biosurfactant produced by B. megaterium

of the biosurfactant produced by Bacillus spp. were lipopeptides (Arima et al. 1968; Bernheimer and Avigad 1970; Marahiel et al. 1977; Neu and Poralla 1990; Mukherjee and Das 2005) but in the present study it is a glycolipid. The

chemical nature of the biosurfactant thus varies with both species and strains within the genus Bacillus. This paper aims to contribute to the biosurfactant production using economically cheaper carbon sources.

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Microbial surfactants are not yet competitive with chemical surfactants, but efforts should be made on different aspects of production to find a suitable and economically viable substrate. Peanut oil cake proved to be a suitable substrate for biosurfactant biosynthesis, providing not only bacterial growth and product accumulation but also a surfactant that has interesting and useful properties with potential of many industrial applications. Further research on structural characterization, gene regulation of biosurfactant and cost of production are in progress. Acknowledgement The authors thank the authorities of Annamalai University for providing the facilities and Department of Ocean Development and Council of Scientific and Industrial Research Govt. of India for their financial support to carryout the work in a successful way.

References Arima K, Kakinuma A, Tamura G (1968) Surfactin, a crystalline peptide surfactant produced by Bacillus subtilis: isolation, characterization and its inhibition of fibrin clot formation. Biochem Biophys Res Commun 31:488–494 Banat IM (1993) The isolation of a thermophilic biosurfactant producing Bacillus sp. Biotechnol Lett 15:591–594 Banat IM, Makkar RS, Cameotra SS (2000) Potential commercial applications of microbial surfactants. Appl Microbiol Biotechnol 53:495–508 Benincasa M, Contiero J, Manresa MA et al (2002) Rhamnolipid production by Pseudomonas aeruginosa LBI growing on soapstock as the sole carbon source. J Food Eng 54:283–288 Bernheimer AW, Avigad LS (1970) Nature and properties of a cytological agent produced by Bacillus subtilis. J Gen Microbiol 61:361–369 Besson F, Michel G, (1992) Biosynthesis of iturin and surfactin by Bacillus subtilis. Evidence for amino acid activating enzymes. Biotechnol Let 14:1013–1018 Betts RP, Bankers P, Banks JG (1989) Rapid enumeration of viable microorganisms by staining and direct microscopy. Lett Appl Microbiol 9:199–202 Bodour AA, Maier RM (1998) Application of a modified dropcollapse technique for surfactant quantification and screening of biosurfactant-producing microorganisms. J Microbiol Methods 32:273–280 Bredholt H, Bruheim P, Potocky M et al (2002) Hydrophobicity development, alkane oxidation and crude-oil emulsification in a Rhodococcus species. Can J Microbiol 48:295–304 Buchanan RE, Gibbons NE, Cowan ST et al (1974) Bergey’s manual of determinative bacteriology. Williams and Wilkinns Co, Baltimore Carrillo P, Mardaraz C, Pitta-Alvarez S et al (1996) Isolation and selection of biosurfactant-producing bacteria. World J Microbiol Biotechnol 12:82–84 Cooper DG, Goldenberg BG (1987) Surface-active agents from two Bacillus species. Appl Environ Microbiol 53:224–229 Desai JD, Banat IM (1997) Microbial production of surfactants and their commercial potential. Microbiol Mol Biol Rev 61:47–64 Deshpande M, Daniels L (1995) Evaluation of sophorolipid biosurfactant production by Candida bombicola using animal fat. Bioresour Technol 54:143–150 Deziel E, Paquette G, Villemur R et al (1999) Biosurfactant production by a soil Pseudomonas strains growing on poly aromatic hydrocarbons. Appl Environ Microbiol 62:1908–1912

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World J Microbiol Biotechnol (2008) 24:917–925 Dubois M, Gilles KA, Hamilton JK et al (1956) Colorimetric method for determination of sugars and related substances. Anal Chem 28:350–356 Fernandez-Linares L, Acquaviva M, Bertrand J-C et al (1996) Effect of sodium chloride concentration on growth and degradation of eicosane by marine halotolerent bacterium Marinobacter hydrocarbonoclastieus. Appl Microbiol 19:113–121 Folch JM, Lees M, Stanly HS (1956) A simple method for the isolation and quantification of total lipids from animal tissues. J Biol Chem 226:497–509 Fox SL, Bala GA (2000) Production of surfactant from Bacillus subtilis ATCC 21332 using potato substrates. Bioresour Technol 75:235–240 Goldman S, Shabtai Y, Rubinovitz C, Rosenberg E et al (1982) Emulsan in Acinetobacter calcoaceticus RAG-I: distribution of cell-free and cell associated cross-reacting materials. Appl Environ Microbiol 44:165–170 Jain DK, Collins-Thompson DL, Lee H et al (1991) A dropcollapsing test for screening surfactant-producing microorganisms. J Microbiol Methods 13:271–279 Johnson M, Boese-Marrazzo D (1980) Production and properties of heat stable extracellular hemolysin from Pseudomonas aeruginosa. Infect Immun 29:1028–1033 Juwarkar A, Khirsagar DG (1991) Emulsification and oil degradation by marine bacteria. Indian J Mar Sci 20:78–79 Li Z-Y, Lang S, Wagner F et al (1984) Formation and identification of interfacial-active glycolipids from resting microbial cells. Appl Environ Microbiol 48:610–617 Makkar RS, Cameotra SS (1999) Biosurfactant production by microorganisms on unconventional carbon sources-a review. J Surf Det 2:237–241 Marahiel M, Denders W, Krause M et al (1977) Biological role of gramicidin S in spore function. Studies on gramicidinc-S negative mutants of Bacillus brevis 9999. Eur J Microbiol 99:49–52 Mercade ME, Manresa MA (1994) The use of agroindustrial byproducts for biosurfactant production. J Am Oil Chem Soc 71:61–64 Mercade ME, Manresa MA, Robert M et al (1993) Olive oil mill effluent (OOME). New substrate for biosurfactant production. Bioresour Technol 43:1–6 Moran A, Alejandra M, Martinez F et al (2002) Quantification of surfactin in culture supernatant by hemolytic activity. Biotechnol Lett 24:177–180 Mukherjee AK, Das K (2005) Correlation between diverse cyclic lipopeptides production and regulation of growth and substrate utilization by Bacillus subtilis strains in a particular habitat. FEMS Microbiol Ecol 54:479–489 Mulligan CN, Cooper DG, Neufeld RJ (1984) Selection of microbes producing biosurfactants in media without hydrocarbons. J Ferment Technol 62:311–314 Neu TR, Poralla K (1990) Emulsifying agent from bacteria isolated during screening for cells with hydrophobic surfaces. Appl Microbiol Biotechnol 32:521–525 Nitschke M, Pastore GM (2004) Biosurfactant production by Bacillus subtilis using cassava processing effluent. Appl Biochem Biotechnol 112:163–172 Rahman KSM, Rahman TJ, McClean S et al (2002) Rhamnolipid biosurfactant production by strains of Pseudomonas aeruginosa using low-cost raw materials. Biotechnol Prog 18:1277–1281 Rodrigues LR, Teixeira JA, van der Mei HC, Oliveira R (2006) Isolation and partial characterization of a biosurfactant produced by Streptococcus thermophilus A. Colloids Surf B Biointerfaces 53:105–112 Rosenberg E, Zuckerberg A, Rubinovitz C et al (1979) Emulsifier of Arthrobacter RAG-I: isolation and emulsifying properties. Appl Environ Microbiol 37:402–408

World J Microbiol Biotechnol (2008) 24:917–925 Rosenberg M, Gutnick DL, Rosenberg E (1980) Adherence of bacteria to hydrocarbons: a simple method for measuring cell-surface hydrophobicity. FEMS Microbiol Lett 9: 29–33 Sandrin C, Peypoux F, Michel G (1990) Coproduction of surfactin and iturin A lipopeptides with surfactant and antifungal properties by Bacillus subtilis. Biotechnol Appl Biochem 12:370–375 Sheppard JD, Mulligan CN (1987) The production of surfactin by Bacillus subtilis grown on peat hydrolysate. Appl Microbiol Biotechnol 27:110–116 Tahzibi A, Kamal F, Assadi MM (2004) Improved production of rhamnolipids by a Pseudomonas aeruginosa mutant. Iran Biomed J 8:25–31

925 Thavasi R, Jayalakshmi S (2003) Bioremediation potential of hydrocarbonoclastic bacteria in Cuddalore harbour waters (India). Res J Chem Environ 7:17–22 Tuleva BK, Ivanov RG, Christova NE (2001) Biosurfactant production by an new Pseudomonas putida strain. Z Naturforsch 57:356–360 Yonebayashi H, Yoshida S, Ono K et al (2000) Screening of microorganisms for microbial enhanced oil recovery process. Sekiyu Gakkaishi 43:59–69 Youssef NH, Duncan KE, Nagle DP et al (2004) Comparison of methods to detect biosurfactant production by diverse microorganisms. J Microbiol Methods 56:339–347 Youssef N, Simpson DR, Duncan KE et al (2007) In situ biosurfactant production by Bacillus strains injected into a limestone petroleum reservoir. Appl Environ Microbiol 73:1239–1247

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Letters in Applied Microbiology ISSN 0266-8254

ORIGINAL ARTICLE

Biosurfactant production by Corynebacterium kutscheri from waste motor lubricant oil and peanut oil cake R. Thavasi1, S. Jayalakshmi1, T. Balasubramanian1 and Ibrahim M. Banat2 1 CAS in Marine Biology, Annamalai University, Parangipettai, Tamil Nadu, India 2 School of Biomedical Sciences, University of Ulster, Coleraine, Northern Ireland, UK

Keywords biodegradation, bioremediation, biosurfactant, emulsification, glycolipopeptide. Correspondence Rengathavasi Thavasi, CAS in Marine Biology, Annamalai University, Parangipettai 608 502, Tamil Nadu, India. E-mail: [email protected]

2007 ⁄ 1837: received 13 January 2007, revised 15 August 2007 and accepted 20 August 2007 doi:10.1111/j.1472-765X.2007.02256.x

Abstract Aim: Production and characterization of biosurfactant from renewable sources. Methods and Results: Biosurfactant production was carried out in 3-l fermentor using waste motor lubricant oil and peanut oil cake. Maximum biomass (9Æ8 mg ml–l) and biosurfactant production (6Æ4 mg ml–l) occurred with peanut oil cake at 120 and 132 h, respectively. Chemical characterization of the biosurfactant revealed that it is a glycolipopeptide with chemical composition of carbohydrate (40%), lipid (27%) and protein (29%). The biosurfactant is able to emulsify waste motor lubricant oil, crude oil, peanut oil, kerosene, diesel, xylene, naphthalene and anthracene; the emulsification activity was comparatively higher than the activity found with Triton X-100. Conclusion: This study indicates the possibility of biosurfactant production using renewable, relatively inexpensive and easily available resources like waste motor lubricant oil and peanut oil cake. Emulsification activity found with the biosurfactant against different hydrocarbons showed the possibility of the application of biosurfactants against diverse hydrocarbon pollution. Significance and Impact of the Study: The data obtained from the study could be useful for large-scale biosurfactant production using economically cheaper substrates. Information obtained in emulsification activity and laboratory-scale experiment on bioremediation inferred that bioremediation of hydrocarbon-polluted sites may be treated with biosurfactants or the bacteria that produces it.

Introduction Biosurfactants are organic compounds belonging to various classes including glycolipids, lipopeptides, fatty acids, phospholipids, neutral lipids and lipopolysaccharides (Rosenberg 1986). The properties ⁄ applications of biosurfactants includes excellent detergency, emulsification, foaming, dispersing traits, wetting, penetrating, thickening, microbial growth enhancement, metal sequestering and resource recovering (oil) which make surfactants replace some of the most versatile process chemicals. Biosurfactants are promising natural surfactants that offer several advantages over chemically synthesized surfactants, such as lower toxicity, biodegradability and ecological acceptability. In order to spread the application of microbial surfactants, methods of possible cost reductions have been sought. Currently, their prices range between 2 and 686

3US$ per kg and are 20–30% more expensive than their synthetic equivalents (Banat 1995). The reduction of the production cost of microbial biosurfactants requires enhancement of biosynthesis efficiency and the selection of inexpensive culture media components as they constitute 50% of the total production costs. The objective of this study was to demonstrate the production of biosurfactant by Corynebacterium kutscheri using waste motor lubricant oil and peanut oil cake and the emulsification capacities of the biosurfactant against hydrocarbons. Materials and methods Organism Corynebacterium. kutscheri was isolated from sea water sample from Tuticorin harbor (08o45’N; 78o13’E) using

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R. Thavasi et al.

Bushnell Haas agar supplemented with 0Æ1% of crude oil and identified to the species level by following Bergey’s Manual of Determinative Bacteriology (Buchanan et al. 1974). Media and growth conditions Biosurfactant production was carried out in a 3-l laboratory fermentor with working volume of 2Æ1 l (Scigenics India Pvt. Ltd., Chennai). The strain was cultured in mineral medium (in g l)1; K2HPO4, 1Æ0; MgSO4.7H2O, 0Æ2; FeSO4.7H2O, 0Æ05; CaCl2.2H2O, 0Æ1; Na2MoO4.2H2O, 0Æ001; NaCl, 30). Optimization of the culture conditions was carried out elsewhere (by Dr R. Thavasi during his PhD work for this strain; unpublished data). The culture conditions are as follows: pH 8Æ0, temperature 38C, salinity 30& and 2Æ0% substrate concentration (waste motor lubricant oil ⁄ peanut oil cake) and 8Æ0 mg l)1of dissolved oxygen (DO). Estimation of growth and biosurfactant production Five-millilitre samples of culture broth were collected at 12-h intervals for a period of 168 h. Biomass was estimated gravimetrically, broth culture was filtered through a Millipore filter paper (0Æ45 lm) and dried at 80C in hot air oven and weighed. Biomass was quoted in terms of mg ml)1 (dry weight). Biosurfactant concentration in the culture broth was estimated according to the procedure described by Li et al. (1984) and the weight was expressed as mg ml)1. The culture broth was centrifuged at 6000 rev min)1 for 20 min at 4C and extracted twice with chloroform and methanol (2 : 1 v ⁄ v). The solvents were removed by rotary evaporation and the residue was partially purified in silica gel (60–120 mesh) column eluted with chloroform and methanol ranging from 20 : 1 to 2 : 1 (v ⁄ v) in a gradient manner. The fractions were pooled and the solvents were evaporated and the resulting residue was dialysed against distilled water and lyophilized. Estimation of emulsification activity Partially purified biosurfactant (1 mg ml)1) was dissolved in 5 ml of Tris buffer (pH 8Æ0) in 30-ml test tubes. Hydrocarbons like waste motor lubricant oil, crude oil, peanut oil, diesel, kerosene, naphthalene, anthracene and xylene were tested for emulsification activity. Five milligrams of the hydrocarbon was added to this solution and shaken well for 20 min and the mixture was allowed to stand for 20 min. The optical density (OD) of the emulsified mixture was measured at 610 nm and the results were expressed as D610 (Rosenberg et al. 1979). Emulsification activity of the

Biosurfactant production by Corynebacterium kutscheri

biosurfactant was compared with Triton X-100, concentration and conditions for emulsification study were maintained similar to that of the biosurfactant. Laboratory experiment on biodegradation of crude oil with biosurfactant The experiment was conducted to study the impact of the biosurfactant isolated from C. kutscheri on biodegradation of crude oil in natural sea water. Crude oil used in this study was obtained from Chennai Refineries Limited, Chennai, India. Its specific gravity was reported by them as 0Æ844 at 25C. Seventy-five-litre plastic tanks were filled with 50 l of filtered and ultraviolet (UV)-treated sea water with 30& salinity, and pH 8Æ0. The experiment was conducted with four different sets: (i) bacterial cells alone; (ii) with fertilizer and cells; (iii) with cells and biosurfactants (0Æ1%w ⁄ v); and (iv) with fertilizer and biosurfactant. Exactly 2Æ0% (w ⁄ v) of crude oil was added to the filtered sea water, inoculation was performed with 24-hold culture at the rate of 1% (v ⁄ v, 103–104 CFU ml)1) concentration. Continuous aeration was provided at 1Æ5 l min)1 with an oil-free aerator and the set up was maintained at room temperature for a period of 168 h. Biodegradation of crud oil was estimated fluorimeterically as described in IOC Manuals and Guide No. 11 (1982). An uninoculated control was kept to asses the natural weathering of crude oil and degradation. Characterization of biosurfactant Biochemical composition of biosurfactant Carbohydrate content of the biosurfactant was determined by the phenol-sulfuric acid method (Dubois et al. 1956) using d-glucose as a standard. Protein content was determined by the method of Lowry et al. (1951) using bovine serum albumin as a standard and lipid content was estimated by following the procedure of Folch et al. (1956). Fourier transform infrared spectroscopy Fourier transform infrared spectroscopy (FTIR) is most useful for identifying types of chemical bonds (functional groups), therefore can be used to elucidate some components of an unknown mixture. The molecular characterization was performed using biosurfactant. Freeze-dried crude biosurfactant (10 mg) was ground with 100 mg of potassium bromide and pressed with 7500 kg for 30 s to obtain translucent pellets. Infrared absorption spectra were recorded on a Thermo Niocolet, AVATAR 330 FTIR system with a spectral resolution and wave number accuracy of 4 and 0Æ01 cm)1, respectively. All measurements consisted of 500 scans, and a potassium bromide pellet was used as background reference.

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687

Physiological and biochemical characteristics of C. kutscheri are illustrated in Table 1. Biosurfactant production was studied using 2Æ0% of waste motor oil and peanut oil cake. Figure 1 shows the time course of biosurfactant production by C. kutscheri with waste motor oil as the substrate. Maximum biosurfactant concentration of 3Æ85 mg ml)1 occurred at 132 h of incubation, when the cells reached their early stationary phase. Maximum biomass was observed at 120 h (5Æ89 mg ml)1). Biosurfactant production with peanut oil cake also shows similar trend, but with higher biosurfactant production (6.4 mg ml)1) than waste motor lubricant oil (Fig. 2). Biosurfactant isolated from C. kutscheri and Triton X-100 showed maximum emulsification activity against Table 1 Characteristics of Corynebacterium kutscheri Name of the test

Result

Gram reaction Shape of the cell Spore formation Utilization of carbohydrates Arabinose Mannitol Sucrose Xylose Hydrolysis Starch Gelatin Fat Casein Catalase Urease Nitrate reduction Citrate

+ Rod –

+, positive; –, negative.

688

– – + – + – + – + + – –

Biosurfactant

0

Biomass

12 24 36 48 60 72 84 96 108 120 132 144 156 168

7 6 5 4 3 2 1 0

Figure 1 Growth and biosurfactant production by Corynebacterium kutscheri using waste motor lubricant oil.

7 6 5 4 3 2 1 0

Biosurfactant

Biomass

12 10 8 6 4 2

0

12 24 36 48 60 72 84 96 108 120 132 144 156 168

Biomass (mg ml–1)

RESULTS

4·5 4 3·5 3 2·5 2 1·5 1 0·5 0

Time (h)

Biosurfactant (mg ml–1)

Mass spectrometric analysis of biosurfactant Biosurfactant was dissolved in methanol and mixed thoroughly. The mass spectrometric analysis of the biosurfactant was carried out in an LCQTM quadrupole ion-trap mass spectrometer (Finnigan MAT, San Jose, CA, USA) which utilizes electrospray ionization (ESI). Standard solutions and samples under investigation were infused into the mass spectrometer at a flow rate of 10 ll min)1. In the ESI, nitrogen and auxiliary gas flows were maintained at 50 and 5 ml min)1, respectively and refer to arbitrary values set by the software. The heated capillary temperature was 250C and the spray voltage was set to 5 kV. Negative ion mode was used and scanning was performed at 50–2000 m ⁄ z range. This analysis was carried out at the School of Biomedical Sciences, University of Ulster, Coleraine BT52 1SA, Northern Ireland, UK.

Biomass (mg ml–1)

R. Thavasi et al.

Biosurfactant (mg ml–1)

Biosurfactant production by Corynebacterium kutscheri

0

Time (h) Figure 2 Growth and biosurfactant production by Corynebacterium kutscheri using peanut oil cake.

waste motor lubricant oil. Emulsification of different hydrocarbons were in the order of waste motor lubricant oil > crude oil > peanut oil > kerosene > diesel > xylene > naphthalene > anthracene. Emulsification of eight different hydrocarbons by the biosurfactant inferred the possibility of its applicability against different hydrocarbon pollution. Biodegradation of crude oil in the laboratory scale experiment inferred that, maximum biodegradation was found with biosurfactant and fertilizer added set (70.7%) followed by biosurfactant (67.2%), fertilizer (61%) and 56% in normal setup. Biochemical composition of the biosurfactant revealed that it is a mixture of carbohydrate, lipid and protein in a combination of 40% : 27% : 29%, respectively. FTIR spectral analysis of the biosurfactant inferred that wave number 3513 cm)1 indicated the presence of carboxylic acids and 3444 cm)1 for N–H ⁄ C–H bonds of protein. CH2 ⁄ C–H asymmetric vibrations were found at 2918 and 2858 cm)1 which confirmed the presence of alkanes (C–H). CH and CH2 deformation was found at 880 and 856 cm)1. Presence of C–O bond was found at 1113, 1022, 797 and 711 cm)1 (Fig. 3). This information from the respective wave numbers confirmed the glycolipopeptide nature of the biosurfactant. The mass spectrometric analysis of the biosurfactant also complements the biochemical and FTIR results that the peaks observed at m ⁄ z = 326, 413, 663 and 1075 corresponded to the carbohydrate, lipid and protein moieties (Fig. 4).

ª 2007 The Authors Journal compilation ª 2007 The Society for Applied Microbiology, Letters in Applied Microbiology 45 (2007) 686–691

Figure 3 Fourier transform infrared spectrum of the biosurfactant produced by Corynebacterium kutscheri.

1022·76 880·01 856·23 797·12 711·38 596·34 481·94 429·52

1113·28 1636·96

2918·36

2858·27

3748·71 3646·08

3500

3000

2500 2000 Wavenumbers (cm–1)

1425·13

1484·75

3444·70

3513·81

95 90 85 80 75 70 65 60 55 50 45 40 35 30 25 20 15 10 4000

2533·96

Biosurfactant production by Corynebacterium kutscheri

% Transmittance

R. Thavasi et al.

1500

1000

500

india pos mode2 #10 RT:0·27 AV: 1 NL: 2·68E6 F:+ c Full ms [ 250·00-2000·00] 413·3

326·5

Relative abundance

100 95 90 85 80 75 70 65 60 55 50 45 40 35 30 25 20 15 10 5

663·4

1075·2 829·7 327·6

414·4

282·3

827·7 825·6 551·5

607·4

787·6 757·6

880·5 905·5 960·4

1134·1

1253·7

429·6

400

600

800

1000

1200

1347·7

1449·3

1400

1572·7 1648·0

1600

1737·9 1822·0 1882·4

1800

2000

m/z Figure 4 Mass spectrum of the biosurfactant produced by Corynebacterium kutscheri.

Discussion This study addressed the possibility of biosurfactant production using cheaper carbon sources like waste motor lubricant oil and peanut oil cake. The possibility of biosurfactant production using cheaper carbon sources was already reported by earlier workers: olive oil mill effluent

was used by Mercade et al. (1993); animal fat and frying oil were used by Deshpande and Daniels (1995); molasses by Benincasa et al. (2002); and starch-rich wastes by Nitschke and Pastore (2004) supporting the present study on use of renewable carbon sources for biosurfactant production. Haba et al. (2000) reported that biosurfactant produced from firing oil did not have emulsifying properties,

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Biosurfactant production by Corynebacterium kutscheri

Table 2 Emulsification of hydrocarbons by biosurfactant isolated from Corynebacterium kutscheri and Triton X-100 Emulsification activity (D610)

Waste motor lubricant oil Crude oil Peanut oil Kerosene Diesel Xylene Naphthalene Anthracene

Biosurfactant

Triton X-100

1Æ72

1Æ93

1Æ69 1Æ31 0Æ95 0Æ81 0Æ51 0Æ41 0Æ41

1Æ85 1Æ56 1Æ12 0Æ94 0Æ78 0Æ63 0Æ58

R. Thavasi et al.

In conclusion, the present study is an attempt to find economically cheaper carbon sources for the large-scale production of microbial biosurfactants. Results obtained in biosurfactant production with waste motor lubricant oil and peanut oil cake suggested the possibility of industrial production of biosurfactants using economically cheaper carbon sources. Satisfactory emulsification activity of the biosurfactant against eight different hydrocarbons indicated its diverse applicability against different hydrocarbon pollution. Further purification, structural characterization of the biosurfactant and genetic regulation of the biosurfactant production are in progress. Acknowledgement

whereas biosurfactant produced in the present study using waste motor lubricant oil and peanut oil cake showed good emulsification activity against eight different hydrocarbons (Table 2). Further it encouraged the aim of the present study to produce biosurfactants from cheaper carbon sources with emulsification property. Parallel increases in biomass and biosurfactant were found from 12 h to 120 h, but maximum biosurfactant concentration was found at 132 h. Higher concentrations of the biosurfactant even after the offset of biomass may be because of the release of cell-bound biosurfactant at the early stationary phase (132 h), which leads to an increase in the extracellular biosurfactant concentration in the medium (Goldman et al. 1982). The emulsification activity of the biosurfactant used in this study was comparatively less than the emulsification activity recorded with Triton X-100. However, while considering the advantages of biosurfactant over chemically synthesized surfactants, such as lower toxicity, biodegradability and ecological acceptability the possibility of replacing the chemical surfactant in oil pollution with the biosurfactant is sought and needs further research with different kinds of experiments. Emulsification activity found with the biosurfactant produced by C. kutscheri in this study inferred that biosurfactant produced with one carbon source like waste motor oil or peanut oil cake could be used against different hydrocarbons. Glycolipopeptide nature of the biosurfactant produced in the present study was similar to the results obtained by earlier workers in Corynebacterium spp. (Zajic et al. 1977; Cooper et al. 1979; Akit et al. 1981). They have opined that biosurfactant produced by Corynebacterium spp. are most commonly a mixture of corynemycolic acids, which are usually composed of cell wall polysaccharides or polymers containing mixtures of carbohydrates, proteins and lipids. There was no variation in chemical composition between biosurfactants produced with waste motor lubricant oil and peanut oil cake. 690

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