Title: Production and characterization of a trehalolipid biosurfactant produced by the novel marine bacterium <i>Rhodococcus</i> sp., strain PML026
Abstract: Journal of Applied MicrobiologyVolume 115, Issue 3 p. 744-755 Original ArticleFree Access Production and characterization of a trehalolipid biosurfactant produced by the novel marine bacterium Rhodococcus sp., strain PML026 D.A. White, Corresponding Author D.A. White Plymouth Marine Laboratory, Plymouth, Devon, UK Correspondence Daniel A. White, Plymouth Marine Laboratory, Prospect Place, The Hoe, Plymouth, Devon, PL1 3DH, UK. E-mail: [email protected] for more papers by this authorL.C. Hird, L.C. Hird PML Applications Ltd., Plymouth, Devon, UKSearch for more papers by this authorS.T. Ali, S.T. Ali Plymouth Marine Laboratory, Plymouth, Devon, UKSearch for more papers by this author D.A. White, Corresponding Author D.A. White Plymouth Marine Laboratory, Plymouth, Devon, UK Correspondence Daniel A. White, Plymouth Marine Laboratory, Prospect Place, The Hoe, Plymouth, Devon, PL1 3DH, UK. E-mail: [email protected] for more papers by this authorL.C. Hird, L.C. Hird PML Applications Ltd., Plymouth, Devon, UKSearch for more papers by this authorS.T. Ali, S.T. Ali Plymouth Marine Laboratory, Plymouth, Devon, UKSearch for more papers by this author First published: 24 June 2013 https://doi.org/10.1111/jam.12287Citations: 73AboutSectionsPDF ToolsRequest permissionExport citationAdd to favoritesTrack citation ShareShare Give accessShare full text accessShare full-text accessPlease review our Terms and Conditions of Use and check box below to share full-text version of article.I have read and accept the Wiley Online Library Terms and Conditions of UseShareable LinkUse the link below to share a full-text version of this article with your friends and colleagues. Learn more.Copy URL Share a linkShare onFacebookTwitterLinked InRedditWechat Abstract Aims The aim of this study was to evaluate biosurfactant production by a novel marine Rhodococcus sp., strain PML026 and characterize the chemical nature and properties of the biosurfactant. Methods and Results A novel marine bacterium (Rhodococcus species; strain PML026) was shown to produce biosurfactant in the presence of hydrophobic substrate (sunflower oil). Biosurfactant production (identified as a trehalolipid) was monitored in whole-batch cultures (oil layer and aqueous phase), aqueous phase (no oil layer) and filtered (0·2 μm) aqueous phase (no oil or cells; extracellular) and was shown to be closely associated with growth/biomass production. Extracellular trehalolipid levels increased postonset of stationary growth phase. Purified trehalolipid was able to reduce the surface tension of water to 29 mN m−1 at Critical Micellar Concentration (CMC) of c. 250 mg l−1 and produced emulsions that were stable to a wide range of conditions (pH 2–10, temperatures of 20–100°C and NaCl concentrations of 5–25% w/v). Separate chemical analyses of the intact trehalolipid and its constituents demonstrated the compound was in fact a mixture of homologues (>1180 MW) consisting of a trehalose moiety esterified to a series of straight chain and hydroxylated fatty acids. Conclusions The trehalolipid biosurfactant produced by the novel marine strain Rhodococcus sp. PML026 was characterized and exhibited high surfactant activity under a wide range of conditions. Significance and Impact of Study Strain PML026 of Rhodococcus sp. is a potential candidate for bioremediation or biosurfactant production for various applications. Introduction Biosurfactants are a heterogeneous group of amphiphilic surface-active compounds that are produced by a multitude of micro-organisms including bacteria, yeast and fungi. Biosurfactants can generally be classified into low molecular weight (glycolipids, fatty acids, phospholipids and lipopeptides) and high molecular weight (polysaccharides, proteins, lipoproteins or polymeric compounds) groupings (Rosenberg and Ron 1997, 1999). In many cases, organisms produce biosurfactants to utilize water-insoluble hydrocarbons or other hydrophobic substrates such as fatty acids and vegetable oils (Haba et al. 2000; Ferraz et al. 2002; Qiao and Shao 2010). Bacteria species that produce biosurfactants have been the subject of detailed investigation. Most notable of these are Pseudomonas sp. (rhamnolipids; glycolipid); Rhodococcus sp. (trehalolipids; glycolipid) and Bacillus subtilus (surfactin; lipopeptide). Trehalolipids are excellent emulsifying compounds with application in microbial-enhanced oil recovery and oil spill treatment (Pacheco et al. 2010; Liu and Liu 2011) and potentially exhibit interesting bioactivities arising from interactions with phospholipid membranes and proteins, haemolytic activity and effects on cell differentiation (Isoda et al. 1995; Ortiz et al. 2008, 2011; Zaragoza et al. 2009, 2010, 2012). Rhodococci generally produce trehalolipid biosurfactants in the presence of hydrophobic substrates with both cell-bound and extracellular production demonstrated (Shulga et al. 1990; Lang and Philp 1998; Yakimov et al. 1999). However, production on soluble substrates has also been reported (Pirog et al. 2004; Ciapina et al. 2006). Further, the chemical diversity of trehalolipids produced by Rhodococci is vast and includes trehalose monomycolates, dimycolates and trimycolates (Kretschmer and Wagner 1983; Kurane et al. 1995; Niescher et al. 2006), and various nonionic acylated derivatives of trehalose (Singer et al. 1990; Philp et al. 2002) and anionic trehalose tetraesters and succinoyl trehalolipids (Uchida et al. 1989; Espuny et al. 1996; Tuleva et al. 2008; Tokumoto et al. 2009). Given the diversity of trehalolipids studied thus far, it is plausible that novel Rhodococcus strains from different environments may yield additional trehalolipids with further interesting properties. There is growing interest in the application of biosurfactants as natural replacements for synthetic surfactants in a range of industries (Muthusam et al. 2008; Banat et al. 2010; Franzetti et al. 2010). This coupled with growing evidence to suggest that the marine environment may yield micro-organisms that produce novel biosurfactant compounds with interesting properties (Maneerat 2005; Satpute et al. 2010) warrants the discovery, evaluation and reporting of novel marine biosurfactant producers. An in-house bacterial collection was previously screened for the production of biosurfactants when grown on glucose, hexadecane or sunflower oil as carbon source (data not shown). When grown on sunflower oil or hexadecane (but not water-soluble substrates), strain PML026 shows reductions in surface tension in the aqueous phase of cultures and in filtered supernatants and appears to form emulsions in situ towards the end of growth. Subsequently, this strain and the biosurfactants it produces were characterized in more detail in this study. Because Rhodococcus strains demonstrate heterogeneous phase distribution between oil and aqueous phases in water–oil cultures (Bredholt et al. 2002), attempts were made to distinguish between biomass and biosurfactant production associated with the oil and/or aqueous phases of cultures and extracellular production throughout an extended growth period in parallel experiments. Further, the biosurfactant produced was purified and characterized, to determine its biosurfactant activity under a range of conditions and its chemical composition. Materials and methods Strain PML026 identification (DNA extraction and 16S rRNA sequencing) Strain PML026 was originally isolated and purified from a locally collected seawater sample and was identified as a very promising biosurfactant producer in the presence of hydrophobic substrates (hexadecane, sunflower and olive oil), but not water-soluble carbon sources (glucose and glycerol) when assessed in a qualitative screening study of an in-house marine bacteria repository (>1200 strains; data not shown). Subsequently, this strain was identified and tested further for biosurfactant production. For identification, a single colony of PML026 was harvested from an agar plate and emulsified in 26·5 μl of sterile, nuclease-free water. Amplification of the 16S rRNA gene was achieved using standard conditions and primers. Amplification was performed using a G-Storm GS1 thermal cycler (Gene Technologies Ltd, Braintree, Essex, UK). The amplification product was purified using a Promega Wizard SV Gel and PCR Clean-Up System (www.promega.com) following the manufacturers' protocol. Direct sequencing of the purified DNA was completed at DNA Sequencing & Services (University of Dundee, Scotland, UK). DNA sequence analysis was performed using BLAST (http://blast.ncbi.nlm.nih.gov/Blast.cgi). Related sequences were identified from the GenBank database using BLAST software. PML026 growth and biosurfactant (trehalolipid) production in batch cultures Growth and biosurfactant production by strain PML026 were monitored in two parallel experiments where either (i) whole cultures (aqueous phase and oil layer) were sacrificed and analysed at each time point over a 20-day culture period or (ii) aqueous phase only (no oil layer) was sampled from batch cultures at each time point during the experiment; (iii) aqueous-phase samples were filtered (2 × 0·2 μm) to remove cells and residual oil to determine the levels of extracellular compounds. Comparative differences between the three types of samples allow interpretation of specific phase/layer-related growth, that is, biomass accumulation strictly associated with oil layer/phase vs just the aqueous phase, and assessment of true extracellular production of compounds. In the first experiment (whole cultures), 20 ml of culture in 100-ml flasks was utilized, and in the second experiment (aqueous-phase sampling only), 500 ml of culture in 2-l flasks was used. In both cases, media used were autoclaved basal seawater media (30 KDa filtered sea water; 0·1% w/v yeast extract) plus 2% v/v sunflower oil with cultures kept at 19°C in a shaking incubator (n = 2). All cultures were inoculated with a stock culture (1% v/v inoculum) of strain PML026 maintained on basal seawater media with 2% glucose as a carbon source on day 0 of the experiment. In the second experiment, at the different sampling points, cultures were removed from the shaking incubator and allowed to stand for 10 min to allow clear separation of the hydrophobic oil layer (determined visually) prior to sampling of aqueous phase (15 ml) via careful aspiration with a graduated Pasteur pipette. Biosurfactant (later characterized as a trehalolipid; see below) was recovered from whole cultures or aqueous-phase samples through addition of an equal volume (1 : 1) of methyl tert-butyl ether, MTBE (Kuyukina et al. 2001). Samples were vortexed vigorously and then centrifuged to obtain an upper organic layer containing trehalolipid. This layer was collected and the procedure repeated. Organic extracts were pooled, rotary evaporated and residues resuspended in 1 ml of MTBE before storage at −80°C prior to analyses. Trehalolipids were quantified after alkaline hydrolysis of MTBE extracts using the orcinol assay due to their carbohydrate content (Koch et al. 1991). The MTBE extracts (500 μl) were freed from solvent and then hydrolysed for 2 h in 1 ml of 1 M NaOH at 90°C in a sealed glass tube with regular mixing. After cooling, an equimolar amount of 1 M HCl was added followed by the addition of 1 ml of hexane, vortexing and centrifugation. The upper hexane layer was discarded, and an aliquot of the lower aqueous phase containing trehalose (100 μl) was added to orcinol reagent (900 μl; 0·4 w/v in 50% H2SO4) prior to incubation at 80°C for 2 h before measuring absorbance of the solution at 421 nm. Results were compared with a calibration curve plotted as a trehalose standard (0–100 μg ml−1) and are reported as trehalose equivalents. Trehalolipids in MTBE extracts were also chemically characterized using different analytical techniques. Biomass formation in whole cultures and aqueous phase of cultures was measured indirectly via quantification of total protein. The aqueous layer from the MTBE extractions was lyophilized, and the residues containing proteins were dissolved in 2% sodium dodecyl sulfate (SDS) solution with sonication and heating at 80°C. Protein content in the SDS solutions was then determined using the bicinchoninic assay (Smith et al. 1985). In addition to quantification of trehalolipids and protein (biomass), biosurfactant production was also indirectly measured in aqueous-phase samples (15 ml) from cultures using the Du Nouy ring method on a surface tensiometer (K6, Kruss, Germany). Physico-chemical characterization of trehalolipid biosurfactant produced by strain PML026 Thin-Layer Chromatography purification of trehalolipids Methyl tert-butyl ether extracts generated from cultures were resolved on Thin-Layer Chromatography (TLC) by spotting small aliquots (5 μl) on silica plates (Silica gel 60A on aluminium foil 20 cm × 20 cm, FLUKA, Sigma-Aldrich, Gillingham, Dorset, UK) and using a mobile phase of chloroform–methanol–20% acetic acid (65 : 15 : 2). Several reagents were tested to confirm biosurfactant compound type (ninhydrin, orcinol, iodine vapour), and a single major band (Rf 0·51) with biosurfactant activity (confirmed postband isolation, resuspension and testing via surface tensiometry) was found to be a glycolipid (confirmed by orcinol staining; 0·1% w/v in 5% H2SO4). Further extracts were run under the TLC conditions described, and the same band was visualized with iodine vapour, scraped from the plate and eluted with chloroform–methanol (2 : 1) for further physico-chemical characterization. Critical Micellar Concentration and emulsifying properties of isolated trehalolipid Purified glycolipid was dissolved in distilled water at various concentrations (0–1000 mg l−1), and the surface tension of solutions was measured using a surface tensiometer to determine the Critical Micellar Concentration (CMC). Postdetermination, pure glycolipid solutions (250 mg l−1) and emulsions produced by vortexing (1 min) glycolipid solutions (250 mg l−1; 10 ml) with sunflower oil (5 ml) were assessed for surface tension and emulsion stability (E24), respectively, over a range of temperatures (20–100°C) by heating for 1 h; pH (2–12) via addition of NaOH or HCl (1 mol l−1); and salt (NaCl) concentrations (5–25% w/v). Liquid Chromatography–Mass Spectrometry (LC-MS) analyses of intact glycolipid biosurfactant The purified glycolipid biosurfactant fraction generated from TLC and MTBE extracts from whole cultures was dried under a gentle stream of nitrogen and then resuspended in methanol before being analysed by LC-ESI-MS. Samples (5 μl) were injected onto a reversed-phase C18 HPLC column (150 mm × 4·6 mm × 5 μm particle size; C18 Alltima, Alltech, Deerfield, IL, USA) and resolved using a 4 mmol l−1 sodium acetate–acetonitrile gradient running from 5% to 100% acetonitrile over 55 min using a flow of 1 ml min−1 with compounds detected by an ESI-MS ion trap system (Agilent 6330 ion trap, Agilent Technologies Ltd., Edinburgh, UK) set in negative ionization mode. Spectra of the intact compound and fragments of the major ions (MS2n) were recorded. MS conditions were as follows: nebulizer pressure = 55 psi; dry gas flow = 12 l min−1; drying gas temperature = 350°C; capillary voltage = 3500 V; scan range 100–2000 m/z. Further trap control settings, for example, ionization voltage, etc., were controlled by the instrument 'Smart' function. Fourier transform infrared (FT-IR) spectroscopy on intact glycolipid biosurfactant FT-IR analyses of the purified glycolipid fraction were carried out on a Bruker IFS66 with Hyperion 1000 microscope fitted with a liquid nitrogen MCT detector, transmission spectra being the result of 32 accumulated scans at 4 cm−1 resolution within the wavenumber range of 4000–600 cm−1, using a Specac DC-2 diamond compression cell. LC-MS analyses of glycolipid saccharide composition Mono and disaccharide composition of the purified glycolipid biosurfactant (c. 10 mg) was determined after alkaline hydrolysis to free the intact sugar moiety from esterified acyl groups and after acid digestion of the purified glycolipid (1 ml of 2 mol l−1 HCl at 90°C for 2 h) to yield monosaccharides only. A LC-MS method incorporating atmospheric pressure chemical ionization (APCI) in negative ionization mode and chloride adduct formation of the respective sugars was utilized to determine individual monosaccharides and disaccharides in the samples (Richochon et al. 2011). In brief, samples (20 μl) were resolved on a Zorbax carbohydrate column (4·6 mm × 250 mm; 5 μm particle size; Agilent Technologies Ltd.) using a mobile phase of 25 : 75 water–acetonitrile (containing 1% chloroform) at flow rate 1 ml min−1. Saccharide identification was confirmed based on the retention time and negative ionization of chlorine adduct ions [M + 35]− and ion fragmentation spectra (MSn) using an ion trap mass spectrometer (Agilent 6330 ion trap, Agilent Technologies Ltd.) and authentic standards (Sigma, Gillingham, Dorset, UK) for identification based on retention times and mass spectra confirmation. MS conditions were as follows: nebulizer pressure = 60 psi; dry gas flow = 6 l min−1; drying gas temperature = 350°C; capillary voltage = 3500 V; scan range 100–2000 m/z. Further trap control settings, for example, ionization voltage, etc., were controlled by the instrument smart™ function. GC-MS analyses of glycolipid fatty acids Fatty acid composition of purified biosurfactant was determined after conversion to fatty acid methyl esters (FAMEs) and analysed by GC-MS (Agilent 7890A GC and 5975C inert MSD, Agilent Technologies Ltd.). Nonadecanoic acid (C19 : 0) was added as an internal standard to lipid extracts from acid digestion, and fatty acids were converted directly to FAMEs by adding 1 ml of transesterification mix (95 : 5 v/v 3N methanolic HCl; 2,2-dimethoxypropane) followed by incubation at 90°C for 1 h. After cooling, FAMEs were recovered by the addition of 1% w/v NaCl solution (1 ml) and n-hexane (1 ml) followed by vortexing. The upper hexane layer was injected directly onto the GC-MS system, and FAMEs were separated on a fused silica capillary column (15 m × 0·1 mm × 0·1 μm; Omegawax™ 100, Supelco, Sigma-Aldrich, Gillingham, Dorset, UK) using an oven temperature gradient of 70–280°C at 40°C min−1 followed by 3-min hold time at 280°C. Helium was used as a carrier gas (0·4 ml min−1), and the injector and detector inlet temperatures were maintained at 280 and 230°C, respectively. FAMEs were identified using a mass spectra library supplied with the GC-MS system (NIST/EPA/NIH Mass Spectral Library, version 2.0) and by comparison of retention times and mass spectra of a FAME standard mix (Supelco, Sigma-Aldrich, Gillingham, Dorset, UK). Results Screening, identification, growth and production of biosurfactant by strain PML026 PML026 is a Gram-positive, nonmotile bacterium. BLAST analysis of 16S rDNA sequence (Genbank accession no. KF211479) of PML026 firmly identifies it as a member of the Rhodococcus species with maximum identity (100%) to the environmental isolate Rhodococcus sp. AMA25 (accession no. JQ977388). Overall, the strongest identity is to environmental strains isolated from a range of diverse environments, both terrestrial and aquatic, and none of these strains have been previously described for biosurfactant production. More targeted BLAST analysis vs strains known to include biosurfactant producers such as Rhodococcus erythropolis (taxid: 1833) and Rhodococcus wratislaviensis (taxid: 44752) generated a maximum identity of 97 and 96%, respectively. Rapid biomass accumulation in whole cultures (exponential growth) was recorded following a lag period of c. 1 day, followed by the onset of stationary growth phase after 2 days of culture with final biomass levels at c. 600 mg l−1 protein (Fig. 1). In the aqueous phase of cultures, the lag period extended to c. 2 days before a rapid exponential rise in biomass followed by the onset of stationary phase after 3 days of culture, with similar protein levels as recorded in whole cultures (600–700 mg l−1). This was mirrored by an increase in the turbidity of the aqueous phase (data not shown). In filtered (<0·2 μm) aqueous-phase samples, protein levels (considered to be dissolved and extracellular) began to increase significantly after c. 5 days from background levels in a comparatively slower linear fashion with final levels of c. 500 mg l−1 at the end of the trial. Similarly, biosurfactant levels in whole cultures significantly increased in an exponential fashion after a lag period of 1 day and reached maximum levels (c. 320 mg l−1 trehalose eq.) after 3 days of culture as did biosurfactant levels (maximum 350–400 mg l−1) in aqueous phase of cultures, which were significantly higher than those in whole cultures. By comparison, extracellular levels of biosurfactant began to increase significantly after 2 days of culture at a much slower linear rate with maximum levels (c. 300 mg l−1) recorded at the end of the experiment (day 20). Figure 1Open in figure viewerPowerPoint (a) Growth (protein) and (b) biosurfactant production (trehalose eq.) in batch cultures of strain PML026 grown using 2% w/v sunflower oil as carbon substrate (n = 2). Measurements were taken from whole cultures (▲); aqueous phase of cultures (●) and filtered (0·2 μm) aqueous phase (○) (see 2 for detailed descriptions). In all parallel experiments, the levels of biosurfactant were in excess of 250 mg l−1 (trehalose eq.) at the end of the investigation. Measurement of surface tension in experimental samples was only conducted on the aqueous-phase samples as the oil layer present in whole cultures creates spurious results when using the Du Nouy ring tensiometer (Fig. 2a). Surface tension in unfiltered aqueous phase remained constant for 1 day at c. 50 mN m−1 and then rapidly dropped to c. 35 mN m−1 and remained fairly constant at that level from 2 days onwards. In filtered aqueous phases from cultures, surface tension again remained constant for 1 day at c. 65 mN m−1 and then dropped to 47 mN m−1 on day 3 and did not change significantly throughout the experiment. Figure 2Open in figure viewerPowerPoint Surface tension of (a) aqueous phase (●) and filtered aqueous phase (○) in batch cultures of strain PML026 grown using 2% w/v sunflower oil as carbon substrate (n = 2) and (b) aqueous solutions of purified trehalolipid (see 2) isolated from cultures of strain PML026 grown using 2% w/v sunflower oil. The CMC of the trehalolipid biosurfactant was c. 250 mg l−1. Isolation, purification and chemical characterization of biosurfactant Biosurfactant extracted from whole cultures of strain PML026 was identified and purified using TLC in the first instance. Production of biosurfactant in whole-culture experiments was demonstrated using TLC also (Fig. 3). Only orcinol reagent produced a positive staining result on TLC plates (positive for glycolipids), and only one significant band was detected in MTBE extracts throughout the experiment from day 2 onwards with an Rf of 0·51. TLC analyses of extracts from aqueous phase (pre- and postfiltration) throughout the experiment also demonstrated the occurrence of only one glycolipid band from day 2 onwards with the same Rf value (data not shown). Figure 3Open in figure viewerPowerPoint Thin-layer chromatogram of biosurfactant extracted (MTBE extracts; see 2) from whole cultures of strain PML026 grown on 2% w/v sunflower oil. Biosurfactant band (Rf value 0·50) was visualized using orcinol spray reagent and recovered from equivalent TLC plates for chemical characterization. Postpurification by TLC, the glycolipid biosurfactant was shown to reduce the surface tension of water to 29 mN m−1 at CMC of c. 250 mg l−1 (Fig. 2b), and solutions of glycolipid at this concentration were shown to retain surfactant activity over a broad range of temperatures (20–100°C), pH (2–10) and salt (NaCl) concentrations (5–25% w/v; Table 1). Similarly, emulsions formed from similar solution were comparatively stable under these conditions. Table 1. Effect of pH, temperature and salt (NaCl) concentrations on surface tension (ST) of aqueous solutions of purified trehalolipid (250 mg l−1) isolated from PML026 cultures and stability of emulsions (E24) prepared by mixing solutions of purified trehalolipid (250 mg l−1; 10 ml) and sunflower oil (5 ml) pH E24 (%) ST (mN m−1) Temp (°C) E24 (%) ST (mN m−1) NaCl (%) E24 (%) ST (mN m−1) 2 34 26 20 44 29 5 36 28 4 34 31 30 44 30 7·5 36 30 6 34 28 40 44 29 10 36 27 8 46 28 60 42 29 15 37 31 10 43 27 80 44 29 20 38 31 12 42 50 100 42 29 25 38 27 ESI-MS analyses of the intact glycolipid (negative ionization mode) showed the presence of a series of ions of nominal m/z 1182, 1210, 1240 and 1268 ([M − H]−; Fig. 4) with percentage areas of the integrated signal being 39, 33, 16 and 12%, respectively. This suggests the actual molecular weight (MW) of the two major homologues was 1183 and 1211 and was supported by positive ionization ESI-MS of the intact glycolipid which showed equivalent water and sodium adducts ([MH + 18]+ and [MH + 22]+, respectively) of 1202, 1206, m/z and 1230, 1234, m/z, respectively (data not shown). Fragmentation analyses (MS2n) of the two most abundant homologues (1182 and 1210 m/z) demonstrated similar fragmentation patterns with losses of 117 m/z, 217 m/z and 333 m/z and similar ions at 485, 503, 583, 601, 609 and 725 m/z, suggesting homologues with similar chemical structures. Figure 4Open in figure viewerPowerPoint ESI-MS spectra (negative ionization mode) of (a) the purified trehalolipid biosurfactant showing four major molecular species at 1182 m/z, 1210 m/z, 1240 m/z and 1268 m/z (b) MSn2 fragmentation spectra of the major homologue 1182 m/z and (c) MSn2 fragmentation spectra of 1210 m/z. FT-IR analyses of the intact glycolipid were undertaken in the range 4000–600 cm−1 (Fig. 5). Several C-H stretching bands of CH2 and CH3 were observed at 2860, 2929 and 2956 cm−1 demonstrating the presence of a saturated hydrocarbon, with a broad O-H stretching band at 3400 cm−1 indicating the presence of an alcohol group. Characteristic of esters, a carbonyl (C=O) stretching band was found around 1735 cm−1 plus confirmation peaks at around 1080–1300 cm−1 corresponding to C-O deformation vibrations associated with carboxylic acids, esters or alcohols. Figure 5Open in figure viewerPowerPoint Fourier transform infrared analyses of purified trehalolipid biosurfactant from cultures of strain PML026 grown using 2% w/v sunflower oil as a carbon source. Saccharide analyses of the isolated glycolipid by LC-MSn demonstrated the intact sugar moiety of alkali hydrolysates of the purified glycolipid biosurfactant to be trehalose based on retention time, mass spectra and fragmentation pattern (377 m/z; [M + Cl]−) with no other sugars (monosaccharides or disaccharides) detected (Fig. 6). Further acid hydrolysis of the purified glycolipid yielded monosaccharides, which demonstrated the presence of glucose as the only monosaccharide constituent of the glycolipid (215 m/z = [M + Cl]−). Figure 6Open in figure viewerPowerPoint LC-APCI-MS of purified biosurfactant saccharide constituents showing (a) chromatogram of glycolipid hydrolysates with two major peaks at 8·6 min (glucose) and 12·9 min (trehalose); (b) mass spectra of peak 1 (glucose) with major ion at 215 m/z [M+CL]−; (c) mass spectra of peak 2 (trehalose) with major ion at 377 m/z [M + Cl]−. Analyses of the fatty acids (FAMEs) present in the purified glycolipid (Table 2) showed there to be a mixture of straight chain and hydroxylated fatty acids ranging from C6 to C14 with the most abundant fatty acids (accounting for >75% based on signal intensity) being 3-hydroxy-octanoic acid (44%; C8H16O3, MW = 160·2) and 3-hydroxy-decanoic acid (34%; C10H20O3, MW = 188·3). Another hydroxylated fatty acid, 3-hydroxytetradecanoic acid (C14H28O3, MW = 244·4), was recorded at much lower abundances based on signal intensity (c. 2%). Table 2. Composition, retention time and percentage area contribution of fatty acid methyl esters (FAMEs) derived from purified trehalolipid biosurfactant FAME Chemical formula of respective fatty acid (MW) Retention time (min) Area contribution (%) Methyl hexanoate C6H12O2 (116·2) 3·1 3·7 Methyl octanoate C8H16O2 (144·21) 4·2 4·8 Methyl decanoate C10H20O2 (172·3) 5·0 7·7 Methyl 3-hydroxy-octanoate C8H16O3 (160·2) 5·8 43·8 Methyl 3-hydroxy-decanoate C10H20O3 (188·3) 6·4 33·7 Methyl hexadecanoate C16H32O2 (256·4) 6·7 3·3 Methyl 3-hydroxytetradecanoate C14H28O3 (244·4) 6·9 2·0 Discussion The marine environment and the diverse array of micro-organisms within it represent a huge source of natural products that can be potentially utilized for commercial applications ranging from therapeutics to biofuels. Biosurfactants derived from marine micro-organisms have been attracting greater interest, yet remain poorly studied in comparison with biosurfactant producers isolated from the terrestrial environment (Maneerat 2005; Satpute et al. 2010). In recent years owing in part to increasing consumer awareness, there has been a growing interest in the large-scale production and recovery of biosurfactants from micro-organisms to replace petrochemical-derived synthetic surfactant compounds. Naturally occurring biosurfactants are not easy to chemically synthesize and often convey novel/enhanced activity; they are nontoxic and biodegradable, are more stable to extremes in temperature, pH and ionic concentration and can be produced from renewable feedstocks (Makkar et al. 2011). Comparison of the bacterial biomass accumulation between whole cultures (oil + aqueous phase) and in the aqueous-phase only demonstrated a di
Publication Year: 2013
Publication Date: 2013-06-24
Language: en
Type: article
Indexed In: ['crossref', 'pubmed']
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