rhodococcus ukmp-5m, an endogenous lipase producing actinomycete from peninsular malaysia

Biologia 69/2: 123—132, 2014Section Cellular and Molecular BiologyDOI: 10.2478/s11756-013-0308-x

Rhodococcus UKMP-5M, an endogenous lipase producingactinomycete from Peninsular Malaysia

Jayesree Nagarajan*, Norazah Mohammad Nawawi & Abdul Latif Ibrahim

Institute of Bio-IT Selangor, Universiti Selangor, Jalan Zirkon A/7A, Seksyen 7, 40 000 Shah Alam, Selangor Darul Ehsan,Malaysia; e-mail: jayesree [email protected]

Abstract: Escalation in food industries unctuous wastes has led to serious anthropogenic problems to the environment.Parallel to “green strategy”, growing awareness in biological treatment emphasizes efficacy of enzymatic technology forbioremediation. Pertinently, researchers are in search for new lipase-lipid interaction for improved outcome. Rhodococcusspecies have documented inadequate evidences on lipase enzyme production. Consequent assessments on Rhodococcus iso-lates from Peninsular Malaysia have identified twelve promising strains as lipase producer. Interestingly, apart from usuallipolytic behaviour, Rhodococcus sp. exhibited significant level of lipase endogenously, while cryogenic grinding methodeffectively ruptured the cell. An isolate from petroleum-contaminated site, namely Rhodococcus UKMP-5M, projected thehighest level of lipase specificity and has further been optimized. It was found out that the best specificity was apparent inacidic condition (pH 5) with 6% inoculum at 30◦C for 72 hours of incubation. Due to high level of mycolic cell-surfactantdeveloped in triacylglycerol supplements, cell lysis was employed with Triton X-100 detergent solubilisation. As a result, oilblend composed of various carbon-chain length fatty acids (composite 2) induces enzyme production extensively. Remark-ably, R. UKMP-5M found to cater enzyme production without aid of inducer by nature, but additional carbon source likeglucose represses lipase production. Further ability for biological treatment was revealed when the optimized R. UKMP-5Mwhole cell degraded waste cooking oil significantly by solubilizing fatty acids and commencing conversion into biomass.These qualities resemble practical new lipid-lipase biological lipid rich on-site treatment.

Key words: Rhodococcus UKMP-5M; lipase; endogenous; cryogenic grinding; Triton X-100.

Abbreviations: DCW, dry cell weight; FA, fatty acid; MSM, minimal salt medium; TAG, triacylglycerol; WCO, wastecooking oil.

Introduction

Volatilizing industrial advancements have resulted innumerous environmental threats especially in assuringclean and safe water to people. Lipids – fat, oil, greasesand fatty acids (FAs) – hold largest portion of or-ganic components in municipal and industrial wastewa-ter particularly from food processing industries (Prasad& Manjanuth 2011). The deposited lipids have led toserious anthropogenic damages, such as clogging pipes,congest treatment filters, odour problems, oil film for-mation, depleting oxygen hence damaging aquatic liv-ings (Cipinyte et al. 2009; Fadile et al. 2011). This sit-uation gets even worse in presence of great numbers offood industries, restaurants, and slaughterhouses wherehigh amount of fat, oil and greases are being disposedto the waste streams continuously (Bhumibamon et al.2002).Therefore, removal and remediation of lipid con-

taminated water is a great necessary due to its associ-ated negative impacts. Commonly, partial recovery oflipid residues will be done through air floatation where

floated lipid is discarded by sanitary landfill dumpingprior to normal water treatment (Mongkolthanaruk &Dharmsthiti 2002). Anaerobic water treatment on theother hand, imposed number of limitations includingclogging on digesters, mass transfer defects for solublesubstrates, floatation of biomass and severe lipid toxic-ity against both methanogenic and acetogenic microbes(Cirne et al. 2007; Dors et al. 2013). Meanwhile, otherapplicable alternative treatment includes thermochem-ical, alkaline and enzymatic pre-treatment.Considering the necessity of improvement in bio-

logical treatment, enzymatic technology especially withregard to lipase is gaining increasing attention (White-ley & Lee 2006; Dors et al. 2013). Aerobic biodegrada-tion processes by active lipolytic strains are regardedas slower and less viable on highly polluted fat contam-inants. However, this strategy was proven to be feasiblewith many other added advantages compared to otherstrategy (Cipinyte et al. 2009). Several studies on enzy-matic aerobic treatment have annotated that this strat-egy promises better effluent quality and does not de-velop odour problems (Chan et al. 2008). Interestingly,

* Corresponding author

c©2013 Institute of Molecular Biology, Slovak Academy of Sciences

124 J. Nagarajan et al.

a recent study by Abeynayaka & Visvanathan (2011)found out that aerobic biological treatment was ableto support organic loading rate up to 5–30 kg chemi-cal oxygen demand per m3 compared to aerobic con-ventional method. It was understood that aerobic bi-ological treatment also releases a substantial amountof energy which can be utilized by bacteria for maxi-mal synthesis and growth of new cells (Eckenfelder etal. 1988; Chan et al. 2008). Enzymatic pre-treatmenthave been labelled as efficient due to stringent environ-mental regulations, clean and eco-friendly applicationswhich effectually decreases chemical oxygen demand,colour, and suspended lipid solids (Dors et al. 2013).Few studies on lipase are effective for food waste treat-ment, such as study by Prasad & Manjunath (2011)who reported high removal of lipid content and reducedlevel of biochemical oxygen demand by several lipaseproducing bacteria on dairy effluents and slaughter-houses wastewater aerobically. Corresponding to these,numerous microbial lipases have been studied for lipiddegradation, including Enterobacter sp., Arthrobactersp., Pseudomonas sp., Bacillus sp., Lactobacillus sp.,Staphylococcus sp., Burkholderia sp. and Peniclliumchrysogenum (Cipinyte et al. 2009; Matsuoka et al.2009; Kumar et al. 2012; Facchin et al. 2013).In order to diversify lipid-lipase interaction, re-

searchers are now in search for new lipases with bet-ter characteristics. The gram positive Actinomycetales,Rhodococcus sp., fascinatingly exhibited metabolismrelated to many lipids. As a well-known “oil eating”bacteria, Rhodococcus sp. are extensively studied forvarious hydrocarbon degradation (Peng et al. 2007;Dass & Chandran 2011). In the study by Kis et al.(2013), Rhodococcus sp. MK1 has been suspected forhigh level of lipase activity due to its excellent levelof degradation of waste from food industry and do-mestic wastewater. However, this actinomycete is mini-mally assessed for lipase production although the pres-ence of lipase in mobilizing accumulated triacylglyc-erol (TAG) has been recognized significantly (Hernan-dez et al. 2008). Therefore, in the present study we re-port lipase production by a local strain ofRhodococcusUKMP-5M as a potential strain in degrading lipid richwastewater.

Material and methods

ChemicalsChemical reagents and media ingredients were purchased,respectively, from Sigma (USA), Fisher Scientific (Singa-pore) or Merck (Germany).

Bacteria and seed culture preparationIn total, twenty three Rhodococcus strains isolated through-out Peninsular Malaysia were kindly supplied by CultureCollection Unit, Institute Bio-IT Selangor. Cultures fromglycerol stock were streaked on nutrient agar plate and in-cubated at 30◦C. As for seed culture, a loop of bacteria(from routinely maintained nutrient agar plate) was inoc-ulated into 50 mL of nutrient broth medium and left forshaking in incubator shaker (Jeio Tech SI-600R, Korea) at30◦C at 160 rpm for overnight.

Lipase activity detection via preliminary screeningComposition of the agar plate consisted of 1% (w/v) pep-tone, 0.5% (w/v) NaCl, 0.01% (w/v) CaCl2 · .H20, 1.5%(w/v) agar and 1% (v/v) of Tween 80 (Rajan et al. 2011).

Lipase production mediumTen percentage of pre-prepared inoculum (OD600 0.6–0.8)were cultivated into 50 mL of production medium. Pro-duction medium contained minimal salt medium (MSM)of 0.1% (w/v) (NH4)2SO4, 0.09% (w/v) K2HPO4, 0.06%(w/v) KH2PO4, 0.02% (w/v) MgSO4.7H20, 0.01% (w/v)yeast extract and 1% (v/v) olive oil.

Enzyme extractionExtracellular crude. The cells free supernatant or extracel-lular crude was collected by centrifugation at 14,000 rpmfor 30 min at 4◦C (Tomy MX 350, Japan) and stored at–20◦C (Fisher & Paykel, United States) to be assayed lateron (Amara & Salem 2009).

Intracellular crude. Upon spinning, the remaining su-pernatant was discarded gently leaving the cell pellets. Thecells were washed twice with 0.05 M potassium phosphate(pH 7). Two mL of buffer were added onto washed cell pelletbefore subjecting for chosen mechanical disruptions.

Method 1 (Cryogenic grinding). The sample was frozenwith liquid nitrogen for 40 s before grinding. Later, thefrozen cells were scrapped and transferred into pre-cooledmortar placed in an ice-box. Sample was grinded with pes-tle constantly for 10 min (Benov & Al-Ibraheem 2002).

Method 2 (Glass bead milling). 0.5 g of acid washedglass beads from Sigma Aldrich (size: 425–600 µm) wasadded to the sample. The sample was vortexed for 10 mincontinuously using FinePCR, Finevortex Korea (Taskova etal. 2006).

The lysed samples were centrifuged for 15 min at 14,000rpm and yielded supernatant was used as intracellular crudefor both methods. Protein of the crude was measured withBradford assay using bovine serum albumin for standardcurve (Bradford 1976).

Spectrophotometry assay for lipase activity quantificationOne mL fresh reaction substrate was prepared by mixingwith 1:1 ratio of 0.1 M Tris-HCl buffer (pH 8.2) and 420 µMp-nitrophenyl substrate into an eppendorf tube. Onto it,200 µL of crude sample were added into reaction mixturewhile same amount of distilled water added for blank andvortexed briefly to initiate reaction (Pinsirodom & Parkin2001; Gupta et al. 2002). The reaction mixture was incu-bated for 10 min. Absorbance reading at 410 nm was mea-sured and released p-nitrophenol calculated based on pre-pared standard curve earlier spectrophotometrically (Bio-mate 3, Thermo Scientific, USA). One unit (U) of enzymeactivity represents amount of lipase releasing 1 µmol p-nitrophenyl per minute. In reference to the total proteincontent (mg/mL) reacted in the assay, relative lipase speci-ficity was calculated by regarding maximum specificity ob-tained at referring condition as 100% (Han et al. 2004).

Optimization of growing conditionsThe effect of pH was monitored by pH ranging from 5 to 9adjusted using 1 M NaOH and 1 M HCl respectively. Theeffect of the inoculum size was determined by inoculum from2%, 4%, 6%, 8% and 10%. With regard to effects of tem-perature and incubation time, various temperatures (20◦C,27◦C, 30◦C, 40◦C, 45◦C and 65◦C) and incubation times(24 h, 48 h, 72 h, 96 h and 120 h) were used.

Optimization of lipase producing Rhodococcus sp. from Peninsular Malaysia 125

Table 1. FA composition in natural oil substrate.

FA length in high contentOil Main composition

C chain length Symbola

Palm oil Palmitic acid (44%), oleic acid (39.1%) C16, C18 16:0, 18:1Coconut oil Lauric acid (47.5%), mystric acid (19%) C12, C14 12:0, 14:0Sunflower oil Linoleic acid (66.2%), oleic acid (21.3%) C18 18:2, 18:1Olive oil Oleic acid (90%) C18 18:1

a Symbol means a:b, i.e. carbon chain length : saturation ratio; if b = 0 it is saturated, b = 1 mono unsaturated, and b>2 poly-unsaturated.

Optimization of medium compositionTriton X-100 detergent-lysis treatment. Due to the nature ofclumpy cells growth of Rhodococcus sp. in various oil media,the cells were subjected for Triton X-100 detergent treat-ment before cryogenic grinding to soften and solubilize thecell membrane despite its different viscosity and properties(London & Brown 2000). The washed cells were incubatedwith 0.05 M potassium phosphate buffer containing 3 mLof 0.1% Triton X-100 for 30 min at room temperature aftera brief vortexing (Lashkarian et al. 2010).

Effect of different inducers. Various inducers (Table 1)such as oil from sunflower, sesame, mustard, coconut, palm,corn, canola, and olive were evaluated. In addition, compos-ite oil from sunflower with canola (Composite 1) and oleinpalm, peanut and sesame (Composite 2) were also tested forinducing lipase production.

Effect of different broth. Lipase production was furtherevaluated on different media with optimised growing con-ditions as follows: Medium 1 (M1): nutrient broth (NB);Medium 2 (M2): nutrient broth + glucose; Medium 3 (M3):nutrient broth + inducer; Medium 4 (M4): nutrient broth+ glucose + inducer; Medium 5 (M5): MSM; Medium 6(M6): MSM + glucose; Medium 7 (M7): MSM + inducer;and Medium 8 (M8): MSM + glucose + inducer.

Biodegradation on natural fatA reaction mixture consisting of 5% (w/v) oil emulsifiedwith gum arabic substrate was incubated at 37◦C in wa-ter bath for homogenization. Five mL of the substrate werelater on reacted with 1 mL of lipase crude with known pro-tein concentration. The reaction mixture was incubated byshaking with 120 rpm at 30◦C for 2 h. Hydrolysis reactionwas later quenched by adding 10 mL of 95% (v/v) ethanolin the mixture. The liberated FAs were estimated based ontitration using 0.05 M NaOH against 2–3 drops phenolph-thalein indicator in the reaction mixture (Rajan et al. 2011).In reference to the total protein content (mg/mL) reactedin the assay, relative lipase specificity was calculated by re-garding maximum specificity obtained at referring conditionas 100% (Han et al. 2004).

Feasibility of lipid degradation by optimized whole cellSubstrate 1 was the suitable natural fat substrate analysedin previous analysis. Concerning the substrate 2, the fol-lowing strategy was used: local restaurants at Shah Alamcity serve diversified Malaysian dishes including fried foodand curry dishes popularly. Hence, waste cooking oil (WCO)which would eventually turn into drainage disposal were col-lected and used as substrate for biological treatment. WCOwas filtered to remove solid food particles before furtheranalysis.

Prior to treatment, mixture of substrate and distilledwater in the ration of 1:1 was homogenized as substrate

using magnetic stirrer. Optimized whole cell after 72 h ofincubation corresponding to 3 g/L dry cell weight (DCW)were washed twice with 0.05 M potassium phosphate and re-suspended in 10 mL of the same buffer and stored in 4◦C.For lipid degradation, 2.5 mL of the cell suspension werecultivated onto the substrate and left shaking at 160 rpmat 30◦C (Maniyam et al. 2011).

The sample was centrifuged to separate the whole cellfrom the oil. The DCW (g/L) per mL and liberated FAs(µmol/mL) in 5 mL sample were titrated for every 12 hinterval (Jin & Bierma 2010).

Statistical analysisThe reading was obtained in triplicates and was graphedwith error bar representing the experimental errors. Thedata were further analysed using SPSS version 20.0. Evalu-ation on differences between groups was based on the one-way ANOVA test at 95% confidence interval with Duncananalysis. Value of p < 0.05 was regarded as statistically sig-nificant.

Molecular weight determinationLipase molecular weight was estimated based on SDS pageprotein separation performed using Bio-Rad electrophoresisdevice. Electrophoresis was carried out where 12% (w/v)polyacrylamide gel was used as resolving gel with 4% (w/v)stacking gel (Laemmli 1970).

Results and discussion

Lipase activity detection via preliminary screeningIn the presence of lipase, Tween or FAs of polyoxyethy-lene sorbitan will be hydrolysed into smaller free FAwhich eventually binds with calcium salt incorporatedin the agar. Thus, crystal-salt white like precipitate ap-pears around the bacteria colony (Plou et al. 1998). Fas-cinatingly, among twenty three screened strains, twelveRhodococcus strains exhibited prominent hydrolysiszone. From Table 2, it was evident that R.NAM319produced the largest hydrolysis zone (6.1 ± 0.46) in 5days of incubation followed by R.NAM350 (5.04± 0.51)in 6 days, R.SeAG1 (4.82 ± 0.24) in 7 days, R.NAM81(4.55 ± 0.21) in 7 days and R.UKMP-5M (4.39 ± 0.18)in 6 days of incubation.

Lipase assay analysis on extracellular crudeAnalysis on cell-free supernatant revealed almost neg-ligible amount of protein content (data not shown).Hence, lipase enzyme from this small protein quan-tity was regarded to contribute insignificant outcome


Table 2. Preliminary analysis of hydrolysis zone formed on Tween 80 plate assay.

Strains Preliminary hydrolysis zonea Time of incubation (day)

R.NAM502 3.17g ± 0.29 7R.NAM81* 4.55b,c,d ± 0.21 7R.NAM160 3.7f ± 0.14 7R.NAM178 3.28f,g ± 0.06 7R.NAM180 4.23d,e ± 0.23 7R.NAM319* 6.1a ± 0.46 5R.NAM342 4.23d,e ± 0.1 7R.NAM349 4.33c,d ± 0.14 7R.NAM350* 5.04b ± 0.51 6R.UKMP-5M* 4.39c,d ± 0.18 6R.AQ5NOL2 3.77e,f ± 0.39 7R. SeAG1* 4.82b,c ± 0.24 6

a Value depicts means of triplicate hydrolysis zone measurement ± standard deviation (n = 3). The asterisks signify the best fivestrains which were chosen for subsequent analysis.

Fig. 1. Comparisons between cryogenic grinding and glass bead milling in terms of total protein content in mg/mL (a) and relativelipase specificity in % (b).

for bioremediation. This scenario indicates cytoplasmiclocalization of the enzyme. Literally, protein of inter-est is produced by the cell destined to be free in cy-tosol, forming a portion of an organelle rather thanbeing expressed out of the membrane extracellularly(Arnoys & Wong 2006). Upon cultivation on a hy-drophobic substrate, nature of Rhodococcus sp. in de-veloping surfactant-like substance which eventually en-coats cell envelope tends to be an additional barrierfor enzyme secretion (Pirog et al. 2004). Similar out-come was reported earlier where carboxylesterase ofRhodococcus sp. Was proven to display a lower enzy-matic activity in extracellular crude than the cell ex-tract (Falchocchio et al. 2005). Therefore, lipase quan-tification was focused on intracellular activity through-out this research.

Lipase assay analysis of intracellular crude for two dif-ferent tested methodsPeculiar nature of Rhodococcus sp. by producing sur-

factant on hydrophobic substrate demonstrated highresistance towards cell lysing. There is even a pub-lished report previously stating that protein extractionusing ultrasonification method of Rhodococcus strainseems to be less effective due to high rigidity and oddchemical composition of the cell membrane (Prasad etal. 2009). Considering these, a comparison study todetermine effective cell lysing method was esteemedto be necessary. Figure 1a clearly indicates that dif-ferent strains of Rhodococcus sp. exhibited differentlevel of permeability towards protein extraction. Ev-idently, cryogenic grinding yielded 27% higher totalprotein from R.NAM350 and likewise 26% higher inR.NAM319, R.NAM81 with 28% and R.SeAG1 with36% and last but not least 70% higher protein contentfrom R.UKMP-5M cell lysate. This is an interestingoutcome since cryogenic grinding is usually applied onmolds and tissue samples lysing indicating competentlevel of cell wall rigidity exhibited by Rhodococcus sp.As shown in Figure 1b, crude for cryogenic grinding


seemingly shows higher specificity for all tested strains.It was understood that constant non-stop force fromgrinding will overcome barrier like “capsular slime”properties produced by these bacteria. Lipase structuremade of an α/β hydrolase fold is also recognised as astrong robust catalyst with greater capacity to with-stand harsh conditions (Hefner & Norin 1999). Hence,possibilities for denaturation or damage of this speci-fied protein during consistent grinding can be least ex-pected. In regards to relative lipase specificity (Fig. 1b),crude of R.UKMP-5M depicted the highest lipase speci-ficity significantly (p < 0.05) with the maximum valueof 93%. Thus, it can be concluded that although pro-tein extraction for R.UKMP-5M is laborious, each mil-ligram of extracted protein displayed high efficacy oflipolytic activity, promoting the strain for further opti-mizations. This value was followed by R.SeAG1 andR.NAM81 with relative specificity of 72% and 68%,respectively. Though highest protein content obtainedfrom R.NAM350, the lipase protein contained in thecrude is comparably low with the specificity of 35% forcryogenic grinding crude.

Optimization of growing condition for best lipase pro-ductionEffect of pH. Rhodococcus sp. are aerobic actino-mycetes displaying high preferences towards alkalinecondition for cell proliferation (Sharma & Pant 2001).A contradictory behaviour was documented in Table 3,displaying that cultivated cell exhibited higher lipasespecificity from acidic medium to alkaline medium indecelerating pattern. Precisely, greatest specificity wasattained in pH 5 with 96.59% ± 3.11 while pH 6 subse-quently followed with 28.96% lower value. In agreementto Sharma & Pant (2011), the amount of cells yieldedin pH 5 is substantially low, while no growth of cell ob-served in pH lower than 5. Therefore, an indirect con-nectivity can be concluded by correlating cell growthand lipase specificity rate. This acclimatization occursdue to the scarcity of energy for cell metabolism. Li-pase secretion in cytoplasm has been vastly induced towithstand acidic condition, the unfavourable environ-ment for actinomycete growth. It was understood thatexcessive secretion of endogenous lipase is crucial atthis moment to trigger the breakdown of accumulatedTAG, which is responsible for survivability as energyprecursor. This unique characteristic is an added ad-vantage since most of the disposed food waste will belater deposited as acidic disposal (Liu et al. 2007). Thisis a quality which is compatible with pronounced lipoly-tic fungi like Candida rugosa and Penicillium notatum(Rehman et al. 2011; Rekha et al. 2012).Effect of inoculum size. The amount of bacterial

load in the medium essentially influences the adapta-tion of the cell and the level of enzyme synthesized toserve cell metabolism. Drastic escalation of relative li-pase specificity was observed from 2% to 6% inoculumsize, however, this trend eventually depleted in higherpercentage of inoculum. As presented, R.UKMP-5Mdemonstrated an excellent level of acceleration from

Table 3. Relative lipase specificity (%) of R.UKMP-5M in regardto various aspects.

Parameter Intracellular crudea

pH 5 96.59a ± 3.116 67.63b ± 1.777 25.73c ± 1.768 23.37c ± 1.189 16.68d ± 0.36

Size of inoculum (%) 2 59.12d ± 0.814 60.14c ± 1.056 99.29a ± 0.618 88.24b ± 1.3410 63.63c ± 2.31

Temperature (◦C) 20 69.29c ± 1.5827 86.28b ± 2.8130 97.80a ± 2.4040 64.46c ± 1.9145 065 0

Incubation time (hour) 24 56.67d ± 0.3548 86.64b ± 1.5872 99.45a ± 0.5196 61.56c ± 2.09120 14.11e ± 1.76

Triacylglycerol supplement Sunflower 13.13e ± 1.20Sesame 26.06e ± 1.57Mustard 18.42d,e ± 1.57Canola 36.78c,d ± 1.87Olive 15.30d,e ± 1.85Corn 64.04b ± 1.93Palm 19.27d,e ± 0.16Coconut 66.82b ± 1.11Composite 1 54.96c ± 0.96Composite 2 99.11a ± 1.53

Broth type M1 14.34e ± 2.43M2 61.22b ± 1.17M3 52.08c ± 3.45M4 9.13f ± 0.05M5 40.24d ± 2.48M6 63.14b ± 4.83M7 98.69a ± 1.19M8 5.85f ± 0.44

a Value depicts means of % pNP liberation in regard to maximumspecificity ± standard deviation (n = 3).

59.12% ± 0.81 to 99.29% ± 0.61 in lipase specificity.Nevertheless, the enzyme acclimatisation reduced to11% and 35.7% specificity at 8% and 10% inoculum, re-spectively. In the present case, 6% inoculum was notedto be aptly suitable for Rhodococcus sp. lipase pro-duction corresponding to the characteristic exhibitedby Pseudomonas sp. in the finding of Baharum et al.(2003). At higher amount from optimum inoculum size,the enzyme production decreases signifying the possi-bility of nutrient depletion and insufficiency of totaldissolved oxygen encountered by the cells (Sharma &Rathore 2002). Meanwhile, a lower inoculum decreasesthe biomass thus reduces the level of enzyme secretion.Effect of temperature. Enzyme specificity increased

relatively from 20 ◦C to 30◦C from 69.29% ± 1.58to 97.80% ± 2.40. Specificity of the enzyme eventu-


ally drops 33.34% in specificity at 40◦C in relative tothe maximum lipase specificity. Meanwhile, at highertemperature of 45◦C and 65◦C, no significant growthwere recorded for R.UKMP-5M thus causing no pro-tein extraction and lipase activity measurement atthese ranges of temperature. Since the evaluated bac-teria have been isolated from Peninsular Malaysia, cli-mate of this tropical country encourages the growthof mesophilic bacteria. Equivalently, Rhodococcus sp.prefers optimum growth in a moderate temperature.This explains the reason for no growth of R.UKMP-5Mwhich led to no lipase detection at temperature 45◦Cand above. As discussed previously, lipase enzyme se-cretion is triggered to mobilize stored TAG to generateenergy for the growth (Hernandez et al. 2008). Thus,competent temperature for cell growth, 30◦C has beennoted to effectively maximize the enzyme production.This finding was in agreement with the study of Tem-burkhar et al. (2012) who concluded best lipase pro-duction for Staphylococcus sp. at 30◦C.Effect of incubation time. Incubation time is an-

other crucial factor to project the longevity of the en-zyme secretion in order to support the bacterial growth.Accurately, optimum incubation time is monitored byculture characteristic and growth rate at different timeintervals (Veerapagu et al. 2013). As seen, enzymespecificity peaked relatively on 72 h of incubation timewith 99.45% ± 0.51 specificity but gradually drops to14.11%± 1.76 at 120 h of incubation. In this case, as anenergy precursor, 72 h of time is the best time periodfor the cell to uptake lipid substrate actively and fur-ther initiate lipase production aggressively to aid andsupport the cell growth for survivability. The enzyme’sefficacy eventually depletes at longer days of incubationcausing cell death due to scarcity of energy. Similar casewas encountered by other lipolytic strains like Acineto-bacter sp., Bacillus sp. and Pseudomonas sp. (Jagtapet al. 2010; Mukesh et al. 2012; Tembhurkhar et al.2012).

Optimization of medium compositionTriton X-100 detergent-lysis treatment. For subsequentoptimization, the culture was supplemented with vari-ous TAG supplements. However, cell lysates from solecryogenic grinding extracted insignificant amount ofprotein content except for cells grown in olive oil and co-conut oil media. Meticulous reviews on Rhodococcus sp.behaviour revealed that type of carbon source greatlyinfluences the mycolic acid compositions of cell enve-lope. These formed mycolic acids or α-alkyl, β-hydroxyFAs are different in chain length and extensively mon-itor permeability of cell surface depending on the car-bon source (Sokolovska et al. 2003). Additionally, thesestrains synthesizes and accumulates wide range of high-molecular weight lipids in their DCW which intercalateswith formed mycolic acid, acting as firm and rigid pro-tective barrier for cell lysis (Sriwongchai et al. 2012).Thus, these phenomena cause different degree of cellmembrane rigidity upon cultivation in various lipid sub-strates. All these mentioned factors eventually lead to

imbalance amount of protein extraction from sole cryo-genic grinding.To troubleshoot these, detergent-lipid interaction

can be applied to solubilize or soften the thick cell mem-brane prior to cryogenic grinding. Triton X-100 is a non-ionic detergent proven to commence effective interac-tion with tightly packed lipid bilayers of cell membrane(London & Brown 2000). It is an excellent mitochon-drial inner-membrane degrader since it preserves theenzymatic content efficiently (Gurtubay et al. 1980).Additionally, when no protein identified, the detergentis expected to soften the capsulated membrane caus-ing the cell to be fewer firm with high permeability(Lashkarian et al. 2010). Impact of Triton X-100 mem-brane solubilisation was visible when substantial con-tent of protein can be analysed with sizeable amountof lipase detected later. Triton X-100 also suitably actsas an activator of the lipase gene and further induceslipase expression by removing repressor molecule (Bishtet al. 2012).

Optimization of medium composition for best lipaseproductionEffect of different inducers.An excellent level of enzymespecificity was evidently stimulated by the Composite 2with highest relative specificity of 99.11% ± 1.53, whilegradually tracked by coconut oil and corn oil in reduc-tion from 33-36% comparatively. As per current discov-ery, lipase specificity of R.UKMP-5M has been widelyinfluenced by the Composite 2, oil blend from oleinpalm, peanut and sesame. Various range of FA com-positions present in this blend ranging from saturatedmystric acids (C14:0) to polyunsaturated arachidonicacid (C20:4). Availability of these broad range FAs ascarbon source allows various type of TAG accumula-tion stored endogenously, which in return causes bothhigher and lower induction for lipase expression. Pres-ence of various types of FAs is known to induce diversebinding affinity towards the active site of surfactant-coated lipases (Kamiya & Goto 1997). Adding surfac-tants directly like Triton X-100, Tween 20, Tween 80and Span 80 into the production medium would repressthe growth of the cell (data not shown).Effect of broth compositions. In the present study,

cell lysates from the Medium M7 (MSM + inducer) re-sulted in relatively highest lipase specificity of98.69% ± 1.19 indicating higher preference of enzymetowards sole inducer. Meanwhile, cell lysates from theM6 (nutrient broth + glucose) and M2 (MSM + glu-cose) depicted the same level of specificity in relative36–37% approximately lower than maximum outcome.Therefore, medium containing only glucose as the car-bon source was also considerably capable of lipase pro-duction although it was not yielding maximum speci-ficity. This emphasizes the nativity of Rhodococcusstrain to display lipolytic activity even without acti-vation by any inducers.Another conclusion can be made from results

achieved with the Media M4 (nutrient broth + glu-cose + inducer) and M8 (MSM + inducer + glucose)


that incorporating both inducer and glucose simul-taneously inhibits the enzyme production with dras-tic drop to 9.13% ± 0.05 and 5.85% ± 0.44, respec-tively. Similar outcome was encountered by Lakshmi etal. (1999) where lipase production by Candida rugosaseverely dropped with the combination of glucose inthe medium. Combination of glucose and inducer even-tually represses lipase production suggesting the occur-rence of carbon catabolite repression. It was apparentthat R.UKMP-5M has selectively utilized carbon sourcefrom a mixture of two different carbon sources, namelyglucose and Composite 2 oil. Being the most preferablecarbon source, the Composite 2 indirectly represses theexpression and catabolic system activity that triggersthe consumption of glucose. This scenario eventuallymodifies the regulatory mechanism of the cultivated cell(Gurke & Stulke 2008).

Biodegradation on natural fatLipase specificity was monitored by molecular proper-ties of the enzyme, substrate size and affinity of enzyme-substrate binding. According to Figure 2, R.UKMP-5M lipase annotates the highest level of hydrolysis incoconut oil, meanwhile subsequent order followed bypalm oil, olive oil and sunflower oil. This shows thatthe enzyme is prone to hydrolyze medium carbon chainlength like lauric acid (C12) and mystric acid (C14)compared to higher chain length like palmitic acid,oleic acid and linoleic acid in other oil substrates. Thisfinding was noted to be compatible with the conclu-sion from Bassegoda et al. (2012) who stated thatthe lipase LipR from Rhodococcus CR-53 exhibitedhigher selectivity against medium chain length sub-strate (C10) compared to insoluble long chain lengthfatty substances attributable to more than 18 carbonchain length.

Fig. 2. Relative lipase specificity of R.UKMP-5M on differentnatural oils.

Feasibility of lipid degradation by optimized whole cellDirect use of intracellular crude is evidently feasible byemploying whole cell or naturally immobilized enzymefor application (Bana et al. 2001; Pazauki et al. 2010).Optimized whole cell of R.UKMP-5M favourably de-graded both coconut oil and WCO as demonstrated inFigure 3. In agreement to previous analysis, R.UKMP-5M exhibited higher growth and FA liberation in co-conut oil compared to WCO. Degradation rates of FSsare related to solubility of the FA; this eventually hints

Fig. 3. Behaviour of optimized R.UKMP-5M whole cell in terms of growth pattern (g/L) and FA liberation (µmol/mL) in coconut oiland WCO. FA liberated in WCO and degraded in coconut oil are expressed in µmol/mL. DCW in both coconut oil and in WCO isexpressed in g/L.


Fig. 4. SDS gel image showing protein bands separation of crudecell lysates of R.UKMP-5M.

that mystric acid is more soluble than modified FA inWCO (Chipasa & Medrzycka 2006).A successive exponential growth was depicted until

the incubation of 72 h in WCO, while 96 h in coconutoil. Chemically, waste cooking or “dead oil” or oil nolonger retains its original properties as the lipid hasbeen chemically modified upon repeated heating. Fry-ing process causes that from lipids, especially from thepolyunsaturated FAs, volatile and non-volatile prod-ucts are developed that undergo oxidation, hydrolysisand polymerization actions (Nazrun et al. 2007). Thismight be the reason for considerably lower growth ratefor cells in WCO although the equal level of growthwas observed until 24 h of incubation (0.14–0.16 g/L).A correlation of the growth level and FA liberation canbe evidently seen from the demonstrated pattern. Ascell proliferation occurs rapidly, activity of lipase en-zyme increases significantly, resulting in higher FA lib-eration.Effective digestion of lipid into smaller FA with

lower saturation increases solubility of the compoundin water, and hence it appears to be less harmful tothe environment (Loehr & Roth 1968). In regards toboth substrates, an escalation of lipid degradation upto level of 77% was observed during the early stageof exponential phase (12 h). As mentioned earlier, lipidsubstrates serve as the sole energy for the optimized cul-tivated cell. Thus during the early stage of cultivationthe cells secretes lipase extensively to adapt to the newenvironment and withstand its successive survivability.This causes the hiking value of FA liberation. A promi-nent fall in % of FA liberation was noticed after 84 hof incubation with a slight decrease approaching 120 h.This scenario aptly demonstrates the start of stationaryphase where the cells lose the ability to further prolif-erate and produce substantial enzyme to withstand thecurrent cell survivability. However, decrement in FAlater indicated that the cell has reached death phasewhile the viable cell mass drops correspondingly.

Molecular weight determinationA mere dark significant protein band was visible at

Fig. 5. Hydrolysis zone formation on Tween 80 plate assay dueto lipase action.

the range of 43–44 kDa indicating the expression ofintended lipase by the bacteria (Figure 4). From theliterature survey focused on rhodococcal lipase, a fewstudies have highlighted the endogenous lipase produc-tion from the inclusion body of this bacterium. Thereis a report which recognizes the molecular weight ofcarboxylesterase gene from Rhodococcus soil isolate at60 kDA (Falcocchio et al. 2005). However, another te-dious study discussed that the new putative lipase LipRfrom Rhodococcus CR-53 falls at the size of 43 kDa(Bassegoda et al. 2005), which was very similar to thepresent finding validating presence of lipase gene.

ConclusionsIn conclusion, we report here for the first time on thestrain Rhodococcus UKMP-5M, an isolate from Penin-sular Malaysia, which may be considered a promising li-pase producing strain aptly suitable for bioremediation.Successive analysis of enzyme purification, molecular-biology works and detailed assessment for feasible bi-ological treatment employing R.UKMP-5M lipase willbe further evaluated in future.

Acknowledgements

The authors would like to express gratitude to Min-istry of Science, Technology and Innovation (MOSTI),Malaysia (3090104000(G)) and the Selangor State Govern-ment, Malaysia for the financial assistance. We would like toacknowledge Institute Bio-IT, Universiti Selangor, for pro-viding us necessary lab facilities throughout this investiga-tion.

References

Abeynayaka A. & Visvanathan C. 2011. Performance comparisonof mesophilic and thermophilic aerobic sidestream membranebioreactors treating high strength wastewater. Biores. Tech-nol. 102: 5345–5352.


Amara A.A. & Salem S.R. 2009. Degradation of castor oil andlipase production by Pseudomonas aeruginosa. J. Agric. En-viron. Sci. 5: 556–563.

Arnoys E.J. & Wang J.L. 2006. Dual localization: proteins in ex-tracellular and intracellular compartments. Acta Histochem.109: 89–110.

Baharum S.N., Salleh A.B., Razak C.N.A., Basri M., RahmanM.B.A. & Rahman R.N.Z.R.A. 2003. Organic solvent tolerantlipase by Pseudomonas sp. strain S5: stability of enzyme inorganic solvent and physical factors affecting its production.Ann. Microbiol. 53: 75–83.

Bana K., Kaieda M., Matsumoto T., Kondo A. & Fukuda H. 2001.Whole cell biocatalysts for biodiesel fuel production utilizingRhizopus oryzae cells immobilized within biomass supportparticles. Biochem. Eng. J. 8: 39–43.

Bassegoda A., Pastor F.I.J. & Diaz P. 2012. Rhodococcus sp. CR-53 LipR is the first member of a new bacterial lipase family(Family X) displaying an unusual Y-type oxyanion hole, sim-ilar to Candida antarctica lipase clan. Appl. Environ. Micro-biol. 78: 1724–1732.

Benov L. & Al-Ibraheem J. 2002. Disrupting Escherichia coli:comparisons method. J. Biochem. Microbiol. 35: 428–431.

Bhumibamon O., Koprasartsek A. & Funthong S. 2002. Biotreat-ment of high fat and oil wastewater by lipase producing mi-croorganisms. Kasetsart J. (Nat. Sci.) 36: 261–267.

Bisht D., Yadav S.K. & Darmwal N.S. 2012. Enhanced produc-tion of extracellular alkaline lipase by an improved strain ofPseudomonas aeruginosa MTCC 10,055. Amer. J. Appl. Sci.9: 158–162.

Bradford M.M. 1976. A rapid and sensitive method for the quan-titation of microgram quantities of protein utilizing the prin-ciple of protein-dye binding. Anal. Biochem. 72: 248–254.

Chan Y.J., Chong M.F., Law C.L. & Hassell D.G. 2009. A reviewon anaerobic-aerobic treatment of industrial and municipalwastewater. Chem. Eng. J. 155: 1–18.

Chipasa K.B. & Medrzycka K. 2006. Behaviour of lipids in bi-ological wastewater treatment processes. J. Ind. Microbiol.Biotechnol. 33: 635–645.

Cipinyte V., Grigiskis S. & Baskys E. 2009. Selection of fat-degrading microorganisms for the treatment of lipid contam-inated environment. Biologija 55: 84–92.

Cirne D.G., Paloumet X., Bjornsson L., Alves M.M. &MattiassonB. 2007. Anaerobic digestion of lipid rich waste- effects of lipidconcentration. Renew. Energ. 36: 965–972.

Das N. & Chandran P. 2011. Microbial degradation of petroleumhydrocarbon contaminants: an overview. Biotechnol. Res. Int.2011: 941810.

Dors G., Mendes A.A., Pereira E.B., Castro H. & Furigo A. 2013.Simultaneous enzymatic hydrolysis and anaerobic biodegra-dation of lipid-rich wastewater from poultry industry. Appl.Water ScI. 3: 343–349.

Eckenfelder W.W., Patoczka J.B. & Pulliam G.W. 1988. Anaero-bic versus aerobic waste treatment in the U.S.A. In Proceed-ings of 5th International Symposium on Anaerobic Digestion,Bologna, Italy, pp. 105–114.

Efremenkor E., Senko O., Zubaerova D., Podorozhko E. & Lozin-sky V. 2008. New biocatalyst with multiple enzymatic activ-ities for treatment of complex food wastewaters. Food Tech-nol. Biotechnol. 46: 208–212.

Facchin S., Alves P.D.D., Siquiera F.D.F., Barocca T.M., VictoriaJ.M.N. & Kalapothakis E. 2013. Biodiversity and secretion ofenzymes with potential utility in wastewater treatment. OpenJ. Ecology 3: 34–47.

Fadile A., Hassani F., Aisam H., Merzouki M. & Benlemih M.2011. Aerobic treatment of lipid-rich wastewater by a bacte-rial consortium. Afr. J. Microbiol. 5: 5333–5342.

Falcocchio S., Ruiz C., Pastor F.J., Saso L. & Diaz P. 2005. Iden-tification of a carboxylesterase-producing Rhodococcus soilisolate. Can. J. Microbiol. 51: 753–758.

Gorke B. & Stulke J. 2008. Carbon catabolite repression in bacte-ria: many ways to make the most out of nutrients. Nat. Rev.Microbiol. 6: 613–624.

Gupta N., Rathi P. & Gupta R. 2002. Simplified p-nitrophenylpalmitate assay for lipase and esterase. Ann. Biochem. 311:98–99.

Gurtubay J.I.G., Goni F.M., Gomez-Fernandej J.C., OtamendiJ. & Macarulla J.M. 1980. Triton X-100 solubilization ofmitochondrial inner and outer membranes. J. Bioenerg.Biomembr. 12: 47–70.

Han S.J., Chung J.H., Cheong C.S., Chung Y.I. & Han Y.S. 2004.Enhancement of lipase activity from Acinetobacter speciesSY-01 by random mutagenesis and the role of lipase specificchaperone. Enzyme Microb. Technol. 35: 377–384.

Hefner F. & Norin T. 1999. Molecular modelling of lipasecatalysed reactions prediction of enantioselectivities. Chem.Pharm. Bull. 47: 591–600.

Hernandez M.A. , Mohn W.W., Martinenz E., Rost E., AlvarezA.F. & Alvarez H.M. 2008. Biosynthesis of storage com-pounds by Rhodococcus jostii RHA1 and global identifica-tion of genes involved in their metabolism. BMC Genomics9: 600.

Jagtap S., Gore S., Yavankar S., Pardesi K. & Chopade B. 2010.Optimization of medium for lipase production by Acineto-bacter haemolyticus from healthy human skin. Indian J. Exp.Biol. 48: 936–941.

Jin G. & Bierma T.G. 2010. Whole cell biocatalysts for producingbiodiesel from waste greases; http://www.istc.illinois.edu/info/library docs/rr/rr-117.pdf/ (accessed 7.9.2013).

Kamiya N. & Goto T. 1997. How is enzymatic selectivity of men-thol esterification catalysed by surfactant coated lipase in de-termined in organic media? Biotechnol. Progr. 13: 488–492.

Kis A., Laczi K., Hajdu A., Szilagyi A., Rakhley G. & Perei K.2013. Efficient removal of unctuous of wastes from wastewa-ter. Int. J. Biosci. Biochem. Bioinform. 3: 395–397.

Kumar S., Mathur A., Singh V. & Nandy K., Khare S.K. & NegiS. 2012. Bioremediation of waste cooking oil using a novellipase produced by Penicillium chrysogenum SNP5 grown insolid medium containing waste grease. Bioresour. Technol.120: 300–304.

Laemmli U.K. 1970. Cleavage of structure proteins during theassembly of head. Nature 227: 680–685.

Lakshmi B., Kangueane P., Abraham B. & Pennatheu G. 1999.Effect of vegetable oils in secretion of lipase from Candidarugosa 9DSM2031. Lett. Appl. Microbiol. 29: 66–70.

Lashkarian H., Raheb J., Shahzamani K., Shahbani H. &Shamsara M. 2010. Extracellular cholesterol oxidase fromRhodococcus sp.: isolation and molecular characterization.Iran. Biomed. J. 14: 49–57.

Liu C.H., Chen W.M. & Chang J.S. 2007. Methods for rapidscreening and isolation of bacteria producing acidic lipase:feasibility studies and novel activity assay protocols. WorldJ. Microbiol. Biotechnol. 23: 633–640.

Loehr R.C. & Roth J.C. 1968. Aerobic degradation of long chainfatty acid salts. J. Water Pollut. Control Fed 40: 385–403.

London E. & Brown D.A. 2000. Insolubility of lipids inTriton X-100: physical origin and relationship to sphin-golipid/cholesterol membrane domains (rafts). Biochim. Bio-phys. Acta 1508: 182–195.

Maniyam M.N., Sjahrir F. & Ibrahim A.L. 2011. Bioremediationof cyanide by optimized resting cells of Rhodococcus strainsisolated from Peninsular Malaysia. Int. J. Biosci. Biochem.Bioinfor. 1: 98–102.

Matsuoka J., Miora A. & Hori K. 2009. Symbiotic effects of alipase-secreting bacterium, Burkholderia arboris SL1B1, anda glycerol-assimilating yeast, Candida cylindracea SL1B2, ontriacylglycerol degradation. J Biosci. Bioeng. 107: 401–408.

Mongkholtanaruk W. & Dharmsthiti S. 2002. Biodegradation bylipid-rich wastewater by a mixed bacterial consortium. Int.Biodeterior. 50: 101–105.

Mukesh K.D.J., Rejitha R., Devika S., Balakumaran M.D.,Rebecca A.I.N. & Kalaichelvan P.T. 2012. Production,optimization and purification of lipase from Bacillus sp.MPTK912 isolated from oil mill effluent. Adv. Appl. Sci. Res.3: 930–938.

Nazrun A.S., Chew C.M., Norazlina M., Kamsiah J. & NirwanaI.S. 2007. Repeatedly heated frying oil and high cholesteroldiet are detrimental to the bone structure of ovariectomisedrats. Int. J. Pharm. 3: 160–167.

Pazouki M., Zamani F., Zamzamian A.H., Fahar M. & Najaf-pour G. 2010. Esterification of free fatty acids by Rhizopus


oryzae as cell-catalyzed from used cooking oil for biodieselproduction. World Appl. Sci. J. 8: 719–724.

Peng F., Liu Z., Wang L. & Shao Z. 2007. An oil-degrading bac-terium Rhodococcus erythropolis strain 3C-9 and its biosur-factant. J. Appl. Microbiol. 102: 1603–1611.

Pinsirodom P. & Parkin K.L. (eds) 2003. Current Protocols inFood Analytical Chemistry, C3.1. John Willey & Sons, NewYork.

Pirog T.P., Shevchuk T.A., Voloshina I.N. & Karpenko E.V.2004. Production of surfactants by Rhodococcus erythropo-lis strain EK-1, grown on hydrophilic and hydrophobic sub-strates. Prikl. Biokhim. Mikrobiol. 40: 544–550.

Plou F.J., Ferrer M., Nuero M.O., Calvo M.V., Alcalde M., ReyesF. & Ballesteros A. 1998. Analysis of Tween 80 as an es-terase/lipase substrate for lipolytic activity assay. BioTech-niques 12: 183–186.

Prasad M.P. & Manjunath K. 2011. Comparative study onbiodegradation of lipid rich wastewater using lipase produc-ing bacterial species. Indian. J. Biotechnol. 10: 121–124.

Prasad S., Raj J. & Bhalla T.C. 2009. Purification of a hy-peractive nitrile hydratase from resting cells of Rhodococcusrhodochrous PA-34. Indian J. Microbiol. 49: 237–242.

Rajan A., Kumar S. & Nair A.J. 2011. Isolation of novel al-kaline lipase producing fungusAspergillus fumigatus MTCC9657 from aged and crude rice bran oil and quantification byHPLTC. Int. J. Biol. Chem. 5: 116–126.

Rehman S., Bhatti H.N., Bhatti I.A. & Asgher M. 2011. Opti-mization of processed parameters for enhanced production oflipase by Penicilium notatum using agricultural wastes. Afr.J. Biotechnol. 10: 19580–19589.

Rekha K.S.S., Lakshmi M.V.V.C., Devi V.S. & Kumar S. 2012.Production and optimization of lipase from Candida rugosausing groundnut oil cake under solid state fermentation. Int.J. Res. Eng. Technol. 1: 571–577.

Sharma K. & Rathore M. 2010. Characterization of physical fac-tors for optimum lipase activity from some bacterial isolates.Inter. J. Curr. Trends Sci. Tech. 1: 51–62.

Sharma S.L. & Pant A. 2001. Crude oil degradation by a marineactinomycete Rhodococcus sp. Indian J. Mar. Sci. 30: 146–150.

Sokolovska I., Rozenberg R., Riez C., Rouxhet P.G., AgathosS.N. & Wattiau P. 2003. Carbon source-induced modifica-tions in the mycolic acid content and cell wall permeabilityof Rhodococcus erythropolis E1. Appl. Environ. Microbiol.69: 7019–7027.

Sriwongchai S., Pokethitiyook P., Pugkaew W., Kruatrachue M.& Lee H. 2012. Optimization of lipid production in the oleagi-nous bacterium Rhodococcus erythropolis growing on glyc-erol as the sole carbon source. Afr. J. Biotechnol. 11: 14400–14447.

Taskova R.M., Zorn H., Krings U., Bouws H. & Berger R.G. 2006.Cell wall disruption techniques for the isolation of intracel-lular metabolites from Pleurotus and Lepista sp. Z. Natur-forsch. 61C: 347–350.

Tembhurkhar V.R., Dama L.B., Attarde N.P. & Zope P.S. 2012.Production and characterization of extracellular lipases ofStaphylococcus sp. isolated from contaminated soil. TrendsBiotechnol. 19: 36–41.

Tembhurkhar V.R., Kulkarni M.B. & Peshwe S.A. 2012. Opti-mization of lipase production by Pseudomonas spp. in sub-merged batch process in shake flask culture. Sci. Res. Rep. 2:46–50.

Veerapagu M., Narayanan A.S., Ponmurugan K. & Jeya K.R.2013. Screening selection identification production and op-timization of bacteria lipase from oil spilled soil. Asian J.Pharm. Clin. Res. 3: 62–67.

Whiteley C.G. & Lee D.J. 2006. Enzyme technology and biolog-ical remediation. Enzyme Microb. Technol. 38: 291–316.

Received May 11, 2013Accepted October 26, 2013

rhodococcus ukmp-5m, an endogenous lipase producing actinomycete from peninsular malaysia

Documents