- Email: [email protected]
Isolation, optimization and molecular characterization of lipase producing bacteria from contaminated soil
Journal Pre-proof
ISOLATION, OPTIMIZATION AND MOLECULAR CHARACTERIZATION OF LIPASE PRODUCING BACTERIA FROM CONTAMINATED SOIL O.I. Ilesanmi , E.A. Adekunle , J.A. Omolaiye , E.M. Olorode , L.A. Ogunkanmi PII: DOI: Reference:
S2468-2276(20)30017-X https://doi.org/10.1016/j.sciaf.2020.e00279 SCIAF 279
To appear in:
Scientific African
Received date: Revised date: Accepted date:
30 July 2019 14 November 2019 23 January 2020
Please cite this article as: O.I. Ilesanmi , E.A. Adekunle , J.A. Omolaiye , E.M. Olorode , L.A. Ogunkanmi , ISOLATION, OPTIMIZATION AND MOLECULAR CHARACTERIZATION OF LIPASE PRODUCING BACTERIA FROM CONTAMINATED SOIL, Scientific African (2020), doi: https://doi.org/10.1016/j.sciaf.2020.e00279
This is a PDF file of an article that has undergone enhancements after acceptance, such as the addition of a cover page and metadata, and formatting for readability, but it is not yet the definitive version of record. This version will undergo additional copyediting, typesetting and review before it is published in its final form, but we are providing this version to give early visibility of the article. Please note that, during the production process, errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain. © 2020 Published by Elsevier B.V. on behalf of African Institute of Mathematical Sciences / Next Einstein Initiative. This is an open access article under the CC BY-NC-ND license. (http://creativecommons.org/licenses/by-nc-nd/4.0/)
Article for Scientific African
ISOLATION, OPTIMIZATION AND MOLECULAR CHARACTERIZATION OF LIPASE PRODUCING BACTERIA FROM CONTAMINATED SOIL Ilesanmi O.I1, Adekunle E.A2*, Omolaiye J.A2, Olorode E.M2, Ogunkanmi L.A1 1
Department of Cell Biology and Genetics, Faculty of Science, University of Lagos, Lagos, Nigeria.
2
Biotechnology Section, Bioscience, Department, Forestry Research Institute of Nigeria. P.M.B 5054, Ibadan, Nigeria.
*Corresponding author Adekunle Abiodun E (PhD). Biotechnology Laboratory, Forestry Research Institute of Nigeria, P.M.B 5054, Ibadan, Nigeria. Tel: +234-706-229-5251 E-mail: [email protected], [email protected]
ABSTRACT Lipase enzymes have application in different industrial processes. Screening different habitats for bacteria with lipase activity and optimizing its fermentation parameters will facilitate its effective production and use in industries. In this study, soil sample from a mechanic’s workshop in Jericho G.R.A, Ibadan and water sample from Iyake lake, Ado-awaye both in Oyo-state were screened for presence of lipase positive bacterial strains. Culture medium parameters such as carbon source, nitrogen source, pH, inoculum volume and incubation time were varied for the purpose of optimization. Maximum lipase activity was recorded at 12 h incubation. This activity occurred in media with olive oil and yeast which were the best carbon and nitrogen source respectively. Inoculum volume had an effect of direct proportionality on lipase activity while the isolated organism was determine to be alkalophilic in nature. Under optimized conditions, the organism in the soil sample produced 528.54 U/L lipase activity; showing a 5.3-fold increase in the activity recorded from innate activity. The nucleotide sequence obtained with 16S rRNA gene identified the isolates with highest lipase activity to be Pseudomonas aeuriginosa when subjected to homology search using the Basic Local Alignment Search Tool (BLAST) program. Results obtained showed that lipase obtained from this source could be of potential use in different industrial processes
Keywords: lipase, bacteria, optimization, Pseudomonas aeuriginosa, alkalophilic
Graphical abstract
Introduction The industrial revolution of the 19th century boosted activities that greatly contributed to environmental depletion (Narayanan, 2014). The present state of the environment and the need to salvage it has led manufacturing industries to come up with resolutions that embrace sustainable production processes, one of which is enzyme catalysis (Sarmah, et al., 2018). Enzyme catalysis embraces functionality under relatively milder conditions of temperatures, pH and pressure resulting in less energy consumption and minimal production of unwanted by-products (Waltes et al., 2001). Lipases (Triacyclglycerol acylhydrolases EC3.1.1.3) belong to a class of hydrolases that are specific for the hydrolysis of fats into fatty acids and glycerol at water-lipid interface. They are also capable of reversing the reaction in non aqueous media and they are abundantly present in nature (Singh et al., 2017). Lipase was determine theed by Clade Bernad, 1856 in pancreatic juice as an enzyme that hydrolyzed insoluble oil droplets and converts them to soluble products (Sangeetha et al., 2010). Different species of fungi, yeast, bacteria, animal and plants have been reported to be sources to lipase. However, microbial lipases have become cynosure of all eyes as a result of its numerous industrial application potentials, ease of culture handling and the ease for scale-up during production (Pratush et al., 2013). The 3-D structure of lipases from different microbial sources are not exactly alike, they exhibit high sequence diversity (Gupta et al., 2015). As a result, these enzymes are unique and specific to the type of bioconversion processes they catalyze, finding relevance in a wide array of industrial processes (Mehta et al., 2017). They are found to be useful in catalyzing various reactions synonymous to food, pharmaceuticals, medical and diagnosis, dairy, fatty acid, leather, cosmetic, detergent, beverage and paper industries. (Sharma et al., 2017).
The implication that microorganisms are ubiquitous is that they would be found in almost any natural habitat (soil, water, air, leaves and tree trunks), soil being the most preferred source as it serves as reservoir for diverse microorganisms (Okafor, 2007). Nevertheless, new habitats especially marine environments have now been included in the list to be studied (Metin and Bakir, 2017). Lipase producing microorganisms are not left out in the microbial population found almost everywhere as they have been reported to be isolated from soil (Sharma et al., 2017; Colla et al., 2016; Alhamdan and Alkabbi, 2016), marine environment (Lodha, 2018), wastewater from fish industry (Suharjono, 2015), agro-industrial waste (Maldonado et al., 2014), waste volatile substances (Muthumari et al., 2016), air (Abada, 2008), palm-oil mill effluent (Nwuche and Ogbonna, 2011), intestine of silkworm (Feng et al., 2011) and on human skin (Jagtap and Chopade, 2015). Cultures isolated ab initio from natural sources are usually identified and classified based on morphological characteristics and cell types. However, due to the advanced techniques in genomics and proteomics, isolates can be classified based on sequence of ribosomal RNA in the 16S of the small subunit (SSU) of the prokaryotic ribosome and 18S ribosomal units of eukaryotes (Boonmahome and Mongkolthanaruk, 2013; Alhamdani and Alkabbi, 2016; Sharma et al.,2017;). Harnessing microorganisms for their potential metabolic activities is largely dependent on the culture media composition. The approach for optimization process is usually carried out one variable at a time and the media composition factors subjected to variation include; nutritional factors (carbon and nitrogen sources) and physicochemical factors (incubation time, temperature,
pH, presence of lipid as inducer, cofactors, inhibitors) (Soleyman et al., 2017). Therefore, different media have different stimulatory effect on lipase production (Dhiman and Chapadgaonkar, 2013). This study seeks to evaluate the lipase producing potentials of organisms from two sources, determine the optimal conditions for scale-up production, isolate and identify the organism with better lipase producing ability
Materials and Methods Collection of sample Soil sample was taken from a mechanic’s workshop opposite Forestry College, Jericho, G.R.A, Ibadan and water sample taken from Ado-awaye Lake. Samples collected were kept in a sterile container and taken to the Biotechnology Laboratory, Forestry Research Institute of Nigeria (FRIN) for analysis. Isolation of bacteria Isolation of microorganism from the water and soil samples was carried out by serial dilution (1:10), using the spread plate method. Aliquots of 0.1 ml were inoculated on nutrient agar that was previously sterilized by autoclaving at 15 lb pressure (121 ) for 15min (Salle, 1974). Plates were incubated at 37
for 24 h in an inverted position. Isolated colonies were purified by
picking and streaking repeatedly on nutrient agar plates to obtain pure cultures. Pure bacterial cultures were kept in slants and stored at 4
for further use.
Primary Screening for lipase producing Bacteria Pure isolated bacterial cultures were screened for lipase activity using tributyrin agar medium (Ahmed et al., 2010). The tributyrin agar medium pH 7.5 was prepared with 5g/L peptone, 3g/L
yeast extract, 15g/L agar, 10 ml/L tributyrin in distilled water. The medium was sterilized for 15min by autoclaving with 15 lb pressure at 121°C and cooled. The aliquot was transferred to petri dishes and allowed to solidify. A loopful of each pure culture was streaked each onto tributyrin agar plates separately and incubated at 37°C for 24 h. After incubation, the clear zone of hydrolysis around the colonies indicated the presence of lipase activity. Positive cultures that showed maximum zone of hydrolysis were selected for further studies. Rhodamine olive oil agar medium pH7.0 was prepared with 5g/L peptone, 3g/L yeast extract, 15g/L agar, 4g/L NaCl in distilled water, sterilized for 15 min by autoclaving with 15 lb pressure at 121
and cooled to 60 . 31.25 ml/L olive oil, 10ml/L sterilized filtered rhodamine dye
(1mg/ml) was added. Aliquots were transferred to petri dishes and allowed to solidify. A loopful of each pure culture was streaked onto rhodamine olive oil agar plates separately and incubated the inoculated plates at 37
for 48 h. After incubation, formation of orange flourescent halos
around bacterial colonies visible upon UV irradiation indicates lipase production. Phenol red agar pH 7.4 was prepared with 5g/L peptone, 3g/L yeast extract, 15g/L agar,1g/L CaCl2 , 0.1M NaOH to adjust pH to 7.4 in distilled water, sterilized for 15 min by autoclaving with 15 lb pressure at 121
and cooled to 60
10ml/L phenol red dye (1mg/ml) and10ml/L
substrate (olive oil) were added. Aliquots were transferred to petri dishes and allowed to solidify. A loopful of each pure culture was streaked onto phenol red agar and incubated at 37
for 48h.
After incubation, a change of color from orange to pink was observed. This indicated the release of fatty acids due to lipolysis. Tween80 agar medium pH7.0 was prepared with 10g/L peptone,20g/L agar, 5g/L NaCl, 0.1g/L CaCl2.2H2O, in distilled water, sterilized for 15 min by autoclaving with 15 lb pressure at 121⁰C
and cooled to 45 . 10 ml/L of Tween 80 was added to the cooled media. Aliquots were transferred to petri dishes and allowed to solidify. A loopful of each pure culture was streaked onto Tween80 agar plates separately and incubated the inoculated plates at 37
for 48h. After
incubation, a white precipitate around colony indicates lipase activity.
Secondary Screening for Lipase Production All the selected lipase producing bacterial cultures were screened for production of lipase. The production media (pH7.0) was prepared with 5g/L peptone, 5g/L beef extract in distilled water, autoclaved for 15 min at 15lb pressure (121°C) and cooled to about 60°C before the addition of 10 ml/L olive oil. 1.0 ml of overnight grown selected lipase producing bacterial cultures was inoculated in 100 ml of production medium in 250ml Erlenmeyer flasks separately and incubated at 35
for 24h. Culture was centrifuged at 10,000rpm for 10min after incubation, supernatant
was used to assay for lipase. Lipase activity assay Lipase activity was assayed for using p-nitrophenyl laureate (p-NPL) as substrate. Reaction mixture contained 2µL of 0.03g/L p-NPL, 6µL of 0.05M phosphate buffer and 2µL of enzyme extract (lipase) incubated at 30oC for 10 min. Reaction was terminated by adding 30µL of absolute ethanol. Absorbance was measured using Nanodrop spectrophotometer at wavelength of 380 nm for lipase reaction. One unit of lipase activity is defined as an enzyme releasing 1µmol of free p-nitrophenol per minute. Optimization of Conditions for Lipase Production Determination of optimum incubation time
The organism was grown in nutrient broth containing appropriate 1% oil (olive oil); pH of the medium to was adjusted to 9.0. Cultures were incubated at 30
and samples were taken at 3 h
interval over a period of 12 h for lipase assay. Determination of a suitable carbon source Different kinds of carbon sources (Tween20, Tween80, Olive oil Glucose and Fructose) were used as they might induce lipase production. Bacterial culture was inoculated into nutrient broth containing each kind of carbon source. Cultures were incubated at 30
for 12 h taking samples
for lipase assay every 3 h. Optimum pH The organism was grown in nutrient broth containing appropriate 1% oil (olive oil); media were adjusted to different pH (3, 5, 7, 9 and 11) and incubated at 30
for 12 h.
Determination of optimum inoculum volume Cell cultures of different volume (1000, 2000, 3000 and 4000 µL) were inoculated into nutrient broth containing appropriate 1% oil (olive oil). Cultures were incubated at 30°C for 12 h taking samples for lipase assay every 3h.
Determination of suitable nitrogen source Different kinds of nitrogen sources (Yeast extract, Peptone, Casein, Ammonium Nitrate and Potassium Nitrate) were used in production media as they might affect lipase production.
Bacterial culture was inoculated into nutrient broth containing each kind of nitrogen source. Cultures were incubated at 30
for 12 h taking samples for lipase assay every 3h.
Molecular Characterization Molecular characterization was studied based on 16S rRNA gene sequencing, BLAST search analysis and phylogenetic tree construction. 16S rRNA gene sequencing Genomic DNA of the lipase positive strain was extracted using procedure as stated in commercially available kit, Quick DNA TM Fecal/Soil Microbe Microprep Kit (Zymo research Corp).The 16S rRNA gene was amplified via PCR, and then the amplicon was sequenced After the isolation, qualitative and quantitative analysis was carried out using 50ng of DNA for DNA amplification. For the amplification of 16S region, the following two degenerate primers were designed Table 1. Table 1: Primer sequence for amplification Primer
Sequence
Forward
5´-AGAGTTTGATCCTGGCTCAG-3´
Reverse
3´-ACGGCTACCTTGTTACGACTT-5´
Cycling conditions for PCR: initial denaturation at 96 at 96
for 1 min, annealing at 53
extension at 68
for 5 min; and 35 cycles of denaturation
for 45 s and extension at 68°C for 90 s; followed by a final
for 10 min. 5μl of PCR product was mixed with equal volume of gel loading
dye and was loaded onto agarose gel. Electrophoresis the samples on1% agarose gel. PCR
products were purified using the gel elution kit, and subjected the product obtained to DNA sequencing process using Big Dye® Terminatorv3.1 Cycle sequencing kit.
BLAST search analysis of 16S rRNA gene sequence The nucleotide sequence obtained with 16S rRNA gene was subjected to homology search using the Basic Local Alignment Search Tool (BLAST) program of the National Centre for Biotechnology Information (NCBI) with default parameters.
Phylogenetic tree construction Representative sequences of similar neighbors in BLAST analysis was retrieved and aligned using multiple alignment program. The multiple alignment file was used to create neighborjoining tree using MAFFT (Multiple Alignment using Fast Fourier Transform).
RESULTS ISOLATION AND IDENTIFICATION OF A LIPASE POSITIVE BACTERIUM FROM SOIL AND WATER SAMPLES.
Soil sample and water sample were examined for the presence of lipase positive strains using a screening method suitable for detection of lipase producers as described in materials and methods. Lipase producer strains in the samples were identified by the formation of orange fluorescent halos around the colonies when olive oil-rhodamine B spread plates incubated at 37°C were exposed to UV light at 350 nm (Plate 1). Olive oil is used as lipase substrate and rhodamine B is the indicator of lipase activity.
Plate 1: UV determination at 350 nm of lipase activity on rhodamine agar plate. Plate A and B from soil, C from water and D as control. All plates were incubated at 37 .
Plate 2: Lipase activity on phenol red agar plate. Plate A and B from soil, C from water and D as control. All plates were incubated at 37 .
Lipase producer strains in the samples were identified by the formation of pink halos around the colonies when olive oil-phenol red spread plates were incubated at 37°C. (Plate 2).
4.2. Determination of lipase activity from soil and water samples. Nutrient broth supplemented with 1% olive oil was used as lipase production media. To determine the organism with lipase activity, cultures were grown at 30
for a period of 18 h and
supernatant was assayed for lipase activity using spectrophotometric method (Table 2).
Table 2: Determination of lipase activity in Water and Soil samples
Isolate
Lipase activity (U/L)
Soil
99.69
Water
45.55
OPTIMIZATION OF GROWTH CONDITIONS
Effect of incubation time Nutrient broth supplemented with 1% olive oil was used as lipase production media. To determine the effect of incubation time on lipase activity and select organisms with higher lipase
activity, cultures were grown at pH 9, 30oC for a period of 12 h and supernatant was taken every 3 h to assay for lipase activity using spectrophotometric method. Lipase activity was prominent in soil as highest activity was valued 364.82 U/L at 12 h, whereas that of water was valued to 162.85 U/L at 9 h (Figure 1).
Lipase activity (U/L)
500
water Soil
400 300 200 100 0 0
2
4
6
8
10
12
Time (h) Figure 1: Determination of lipase activity from water and soil sample Effect of carbon source Nutrient broth supplemented with different carbon sources were used as lipase production media. To determine the effect of different carbon sources ( Tween 20, Tween 80, Glucose and Fructose) on
lipase
production
in organism with higher lipase activity from previous
experiment, cultures were grown at 30oC in each production medium over a period of 12 h and supernatant was taken to assay for lipase ativity using spectrophotometric method.The medium containing olive oil had the highest lipase activity at 364.81 U/L while Tween 20, Tween 80 ,
Glucose and Fructose had activity at 202.39, 313.69, 61.8 and 106.57 U/L respectively (Figure 2).
600
Lipase activity (U/L)
500 400 300 200 100 0 Tween 20 Tween 80 Glucose
Olive oil
Fructose
Carbon Source ( 1%) Figure 2: Effect of different carbon sources on isolate from soil Effect of pH Nutrient broth of different pH supplemented with 1% olive oil was used as lipase production media. To determine the effect of different pH (3,5,7,9 and 11) on lipase production, cultures were grown in production media of different pH at 30OC over a period of 12 h and supernatant was taken to assay for lipase ativity using spectrophotometric method.The least lipase activity, 84.66 U/L was recorded at pH 3, while pH 5, 7, 9 had 158.99, 230.75, 360.62 U/L respectively and the highest activity of 432.28U/L was at pH11 (Figure 3).
Lipase activity (U/L)
600 500 400 300 200 100 0 3
5
7
9
11
pH Figure 3: pH optimization of isolate from soil Effect of inoculum volume Nutrient broth inoculated with different volume of cell cultures supplemented with 1% olive oil was used as lipase production media. To determine the effect of different inoculum volume ( 1000, 2000, 3000, 4000 µL) on lipase production, cultures were grown in production media of different inoculum volume at pH 11, 30OC over a period of 12 h and supernatant was taken to assay for lipase ativity using spectrophotometric method. The highest lipase activity recorded was 528.54 U/L obtained from the medium containing 4000µL inoculum volume (Figure 4).
Volume
600
Lipase Activity (U/L)
500 400 300 200 100 0 1000
2000
3000
4000
Volume (µL)
Figure 4: Inoculum volume optimization of isolate from soil Effect of Nitrogen sources Medium containing different nitrogen sources supplemented with 1% olive oil was used as lipase production media. To determine the effect of different nitrogen sources (Yeast Extract,Peptone, Casein, Ammonium Nitrate and Potasssium Nitrate) on lipase production, cultures media were set at pH 11, temperature 30
over a period of 12 h. The supernatant was taken to assay for
lipase ativity using spectrophotometric method. Lipase activity of 490.72, 445, 225.6, 482 U/L was recorded for Yeast Extract, Peptone, Casein, Ammonium Nitrate respectively while no activity was detected in the medium that had Potassium Nitrate as its nitrogen source (Figure 5).
600
Lipase activity (U/L)
500
400
300
200
100
0
Yeast Extract
Peptone
Casein
Ammonium Potassium Nitrate Nitrate
Nitrogen source (1%) Figure 5: Effect of different nitrogen sources on isolate from soil DNA Sequence Analysis/ Molecular Identification of Bacterial Isolates BLAST analysis revealed that all the amplified DNA sequences were bacteria in the phyla Proteobacteria and Firmicutes and belonged to two different genera. The 16S rRNA sequence of the isolate B_27-F_C05_08 illustrated a high similarity to the members of the genus Pseudomonas, showing 98% similarity with Pseudomonas sp. RH-7 (GenBank Accession no. KT715742), Pseudomonas sp. strain GJY (GenBank Accession no. MG585077), Pseudomonas aeruginosa strain GHJ15 (GenBank Accession no. MG396976) and Pseudomonas sp. strain S2 (GenBank Accession no. MF425660). Isolate C_27-F_D05_11 belongs to the genus Bacillus showing 78% similarity with Bacillus sp. strain WP7 (GenBank Accession no. KY347914), Bacillus cereus strain M-1 (GenBank Accession no. EF633995), Bacillus thuringiensis strain WS 2625 (GenBank Accession no. Z84587) and Bacillus cereus isolate DRG3 (GenBank Accession no. AM419192).
Table 3: Molecular Characterization of Bacterial Isolates Sample ID.
Organisms identified by BLAST
IDENT (%)
Sequence length (Bp)
B_27-F_C05_08
Pseudomonas sp. RH-7
98
1101
78
1180
Pseudomonas sp. strain GJY Pseudomonas aeruginosa strain GHJ15 Pseudomonas sp. strain S2 C_27-F_D05_11
Bacillus sp. strain WP7 Bacillus cereus strain M-1 Bacillus thuringiensisstrain WS 2625 Bacillus cereus isolate DRG3
NOTE: B_27-F_C05_08, isolate from soil; C_27-F_D05_11, isolate from water
Figure 6: Phylogenetic tree of the isolate B_27-F_C05_08 and C_27-F_D05_11
DISCUSSION Lipases are enzymes capable of hydrolyzing triacylglycerol. They are obtained from different sources, bacteria being one of them are found in almost every habitats. Screening samples collected from different habitats as described in materials and methods indicated the presence of lipase positive strains from both sources. Strains found in soil sample from a mechanic’s workshop exhibited higher lipase activity when compared to those found in water sample from Iyake lake. This result supports the evidence that the environment influences the status and behaviour of the living system (Verma and Agarwal, 2005). This is explainable that the organisms isolated from the soil are accustomed to the presence of oils and its analogues whereas organisms from water are not.
Upon identifying and isolating organisms with highest lipase activity, different growth conditions that influence lipase production were varied using one variable at a time approach for the optimization of lipase production Starting with incubation time, highest lipase activity was recorded at 12h after which a noticeable decline occurred thereafter. The decline could be related to consumption of nutritional elements. Gupta et al., 2004 reported that maximum lipase activity occur from few hours to several days. As a result of this finding, incubation time for subsequent cultures did not exceed 12 h period. Lipase has been reported to be an inducible enzyme and carbon has been a major factor for the expression of lipase activity (Bakir and Metin, 2017). The effect of different carbon source showed higher activity on lipidic substrate (olive oil) and its analogue (Tween20 and Tween80) with olive oil being the carbon source with the highest lipase activity. Glucose showed the least lipase activity. Catabolic repression may be reponsible for this as sugar molecules act as repressors in lipase synthesis (Soleymani et al., 2017). After determining olive oil as the carbon source with highest lipase activity, pH of the production medium was varied (pH 3, 5, 7, 9 and 11) in order to determine the optimum pH. High lipase activity was exhibited in the pH range of 7-11 while optimum activity was observed at pH 11. In the acidic, reduction of enzyme activity was observed. The present report shows that the organism is an alkalophilic bacteria. Bacteria with lipase producing ability have been reported to prefer pH around 7. However, maximum activity has also been reported in pH higher than 7 (Gupta et al., 2004).
Salihu et al., 2011 reported that inoculum volume has direct proportional effect on lipase activity. Our findings supported this claim as the medium inoculated with the highest culture
volume, 4000µL showed highest lipase activity. However, Thakur et al., 2014 reported inoculum volume is inversely proportional to lipase activity because cells are struggling for nutrients. The last factor that was varied was nitrogen source. Both organic and inorganic nitrogen sources have reported to supply nitogen, cell growth factors and amino acids necessary for enzyme synthesis (Thakur et al., 2014). Three (Yeast Extract, Peptone and Ammonium Nitrate) out of five tested nitrogen sources showed really high lipase activity. Potassium Nitrate did not show any lipase activity. In conlusion, the lipase producing bacteria strain, B_27-F_C05_08 isolated from the mechanic’s workshop was identified as Pseudomonas aeruginosa whose preferred carbon source, nitrogen source, pH and incubation time are olive oil, ammonium nitrate, pH 11 and 12 h respectively.
Conclusion Utilization of enzymes synthesized from biological organism for industrial application is gaining global attention due to their numerous industrial and biotechnology applications. Increased enzyme production from biological sources can be achieved by culture media optimization. Different variables such as substrates concentration, inducers, nitrogen and carbon sources which facilitated increased lipase production have been subjects of focus to researchers. In this work, improved lipase production from Pseudomonas aeruginosa was achieved via optimization of growth and physiological parameters. The high yield of lipase obtained from this process is an indication of the possibility for its large scale production and effective application for industrial/biotechnological purposes. Producing this important enzyme from innate producing organisms such as Pseudomonas aeruginosa is economic and environmentally advantageous.
Declaration of interest statement The authors declare no conflict of interest.
Funding: This research did not receive any specific grant from funding agencies in the public, commercial, or not-for-profit sectors.
REFERENCES
Abada, E. 2008. Production and Characterization of a Mesophilic Lipase isolated from Bacillus stearothermophilus AB-10. Pakistan Journal of Biological Sciences. 11(8): 1100-1106. Alhamdani, M. and Alkabbi, H. 2016. Isolation and Identification of lipase producing bacteria from oil contaminant soil. Journal of Biology, Agriculture and Healthcare. 6(2):1-7. Alhamdani, M. and Alkabbi, H. 2016. Isolation and Identification of lipase producing bacteria from oil contaminant soil. Journal of Biology, Agriculture and Healthcare. 6(2):1-7. Boonmahome, P. and Mongkolthanaruk, W. 2013. Lipase producing Bacterium and its Enzyme Characterization. Journal of Life Sciences and Technologies.1(4):196-200. Colla, L., Primaz, A., Benedetti, S., Loss, R., Lima, M. And Reinehr, C. 2016. Surface response method for the optimization of lipase production under submerged fermentation by filamentous fungi. Brazilian journal of microbiology.47: 461-467. Dhiman, S. and Chapadgaonkar, S. 2013. Optimization of Lipase Production Medium for a Bacterial Isolate. International Journal of ChemTech Research.5(6):2837-2843.
Feng, W., Wang, X., Zhou, W., Liu, G. and Wan, J. 2011. Isolation and Characterization of lipase-producing bacteria in the intestine of the Silkworm, Bombyxmori, reared on different forage. Journal of Insect Science. 11 (135): 1-10. Gupta, R., Kumari, A., Syal, P. and Singh, Y. 2015. Molecular and functional diversity of yeasts and fungal lipases: Their role in biotechnology and cellular physiology. Progress in Lipid Research 57:40-54. Jagtap, S. and Chopade, B. 2015. Purification and Characterization of Lipase from Acinetobacter haemolyticus TA-106 Isolated from human skin. Songklanakarin Journal of Science and Technology.37 (1): 7-13. Lodha, K. 2018. Isolation and Screening of multiple enzyme producing Alkaphilic Bacillus Species from Lonar lake. International journal of Biology Research.3(1):164-168. Maldonado, R., Macedo, G. and Rodrigues, M. 2014. Lipase production using Microorganisms from different Agro-Industrial By-products. International Journal of Applied Science of Technology.4(1):1-8. Mehta, A., Bodhu, U. and Gupta, R. 2017. Fungal lipases: a review. Journal of Biotechnology Research 8:58-77. Metin, K. and Bakir, Z. 2017. Production and Characterization of an Alkaline lipase from Thermophilic anoxybacillus sp. HBB16. Chemical and Biochemical Engineering Quarterly 31(3):303-312. Muthumari, G., Thilagarathi, S. and Hariram, N. 2016. Industrial enzymes: Lipase Produicing Microbes from waste volatile substances. International Journal of Pharaceutical Sciences and Research.7(5): 2201-2208.
Narayanan, P. 2014. Enviromental Pollution: Principles, Analysis and Control. CBS Publishers & Distributors.New Delhi. 671pp. Nwuche, C. and Ogbonna, J. 2011. Isolation of Lipase Producing Fungi from Palm Oil Mill Effluent (POME) Dumpsites at Nsukka. Brazilian Archives of Biology and Technology. 54(1) :113-116. Okafor, N. 2007. Modern Industrial Microbiology & Biotechnology. Enfield, New Hampshire: Science Publishers. 551pp. Pratush, A., Gupta, A., Vyas, G. and Sharma, P. 2013. Microbiology application.Bhalla Publishers. Dehradun, India. pp64-83. Sangeetha, R., Geetha, A. and Arulpandi, I. 2010.Concomitant production of protease and lipase by Bacillus licheniformis VSG1: Production, purification and characterization.Brazilian Journal of Microbiology. 41: 179-185. Sarmah, N., Revathi, D., Sheelu, G., Rani, Y., Sridhar, S. and Mehtab, V. 2018. Recent Advances on Sources and Industrial Applications of Lipases. Biotechnology progress.34(1): 1-24. Sharma, P., Sharma, N., Pathania, S. and Handa, S. 2017. Purification and characterization of lipase by Bacillus methylotrophicus PS3 under submerged fermentation and its application
in
detergent
industry.
Journal
of
genetic
engineering
and
biotechnology.15:369-377. Sharma, P., Sharma, N., Pathania, S. and Handa, S. 2017. Purification and Characterization of Lipase by Bacillus methylotrophicus PS3 under submerged fermentation and its application
in
detergent
Biotechnology.15:369-377.
industry.
Journal
of
Genetic
Engineering
and
Singh, M., Chandraveer. and Tripathi, A. 2017. Isolation and Screen ing of Lipases producing Microorganisms from Natural Sources. Indian journal of ecology. 44(1): 19-23. Soleyman, S., Alizadeh, H., Mohammadian, H., Rabbani, E., Moazen, F. and Sadeghi, H. 2017. Efficient Media for High Lipase Production: One Variable at a Time Approach. Avicenna Journal of Medical B iotechnology. 9(2): 82-86. Suharjono, N. 2015. Activity Assay and Identification of Lipolytic Bacteria from Wastewater FishIndustry at Mucar, Banyuwang, Indonesia. International Journal of ChemTech Research.8(11): 377-383. Waltes, M., Morgan, N., Rockey, J. and Higton, G. 2001. Industrial Microbiology: An Introduction.Blackwell Science Ltd. London.302pp.
ISOLATION, OPTIMIZATION AND MOLECULAR CHARACTERIZATION OF LIPASE PRODUCING BACTERIA FROM CONTAMINATED SOIL O.I. Ilesanmi , E.A. Adekunle , J.A. Omolaiye , E.M. Olorode , L.A. Ogunkanmi PII: DOI: Reference:
S2468-2276(20)30017-X https://doi.org/10.1016/j.sciaf.2020.e00279 SCIAF 279
To appear in:
Scientific African
Received date: Revised date: Accepted date:
30 July 2019 14 November 2019 23 January 2020
Please cite this article as: O.I. Ilesanmi , E.A. Adekunle , J.A. Omolaiye , E.M. Olorode , L.A. Ogunkanmi , ISOLATION, OPTIMIZATION AND MOLECULAR CHARACTERIZATION OF LIPASE PRODUCING BACTERIA FROM CONTAMINATED SOIL, Scientific African (2020), doi: https://doi.org/10.1016/j.sciaf.2020.e00279
This is a PDF file of an article that has undergone enhancements after acceptance, such as the addition of a cover page and metadata, and formatting for readability, but it is not yet the definitive version of record. This version will undergo additional copyediting, typesetting and review before it is published in its final form, but we are providing this version to give early visibility of the article. Please note that, during the production process, errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain. © 2020 Published by Elsevier B.V. on behalf of African Institute of Mathematical Sciences / Next Einstein Initiative. This is an open access article under the CC BY-NC-ND license. (http://creativecommons.org/licenses/by-nc-nd/4.0/)
Article for Scientific African
ISOLATION, OPTIMIZATION AND MOLECULAR CHARACTERIZATION OF LIPASE PRODUCING BACTERIA FROM CONTAMINATED SOIL Ilesanmi O.I1, Adekunle E.A2*, Omolaiye J.A2, Olorode E.M2, Ogunkanmi L.A1 1
Department of Cell Biology and Genetics, Faculty of Science, University of Lagos, Lagos, Nigeria.
2
Biotechnology Section, Bioscience, Department, Forestry Research Institute of Nigeria. P.M.B 5054, Ibadan, Nigeria.
*Corresponding author Adekunle Abiodun E (PhD). Biotechnology Laboratory, Forestry Research Institute of Nigeria, P.M.B 5054, Ibadan, Nigeria. Tel: +234-706-229-5251 E-mail: [email protected], [email protected]
ABSTRACT Lipase enzymes have application in different industrial processes. Screening different habitats for bacteria with lipase activity and optimizing its fermentation parameters will facilitate its effective production and use in industries. In this study, soil sample from a mechanic’s workshop in Jericho G.R.A, Ibadan and water sample from Iyake lake, Ado-awaye both in Oyo-state were screened for presence of lipase positive bacterial strains. Culture medium parameters such as carbon source, nitrogen source, pH, inoculum volume and incubation time were varied for the purpose of optimization. Maximum lipase activity was recorded at 12 h incubation. This activity occurred in media with olive oil and yeast which were the best carbon and nitrogen source respectively. Inoculum volume had an effect of direct proportionality on lipase activity while the isolated organism was determine to be alkalophilic in nature. Under optimized conditions, the organism in the soil sample produced 528.54 U/L lipase activity; showing a 5.3-fold increase in the activity recorded from innate activity. The nucleotide sequence obtained with 16S rRNA gene identified the isolates with highest lipase activity to be Pseudomonas aeuriginosa when subjected to homology search using the Basic Local Alignment Search Tool (BLAST) program. Results obtained showed that lipase obtained from this source could be of potential use in different industrial processes
Keywords: lipase, bacteria, optimization, Pseudomonas aeuriginosa, alkalophilic
Graphical abstract
Introduction The industrial revolution of the 19th century boosted activities that greatly contributed to environmental depletion (Narayanan, 2014). The present state of the environment and the need to salvage it has led manufacturing industries to come up with resolutions that embrace sustainable production processes, one of which is enzyme catalysis (Sarmah, et al., 2018). Enzyme catalysis embraces functionality under relatively milder conditions of temperatures, pH and pressure resulting in less energy consumption and minimal production of unwanted by-products (Waltes et al., 2001). Lipases (Triacyclglycerol acylhydrolases EC3.1.1.3) belong to a class of hydrolases that are specific for the hydrolysis of fats into fatty acids and glycerol at water-lipid interface. They are also capable of reversing the reaction in non aqueous media and they are abundantly present in nature (Singh et al., 2017). Lipase was determine theed by Clade Bernad, 1856 in pancreatic juice as an enzyme that hydrolyzed insoluble oil droplets and converts them to soluble products (Sangeetha et al., 2010). Different species of fungi, yeast, bacteria, animal and plants have been reported to be sources to lipase. However, microbial lipases have become cynosure of all eyes as a result of its numerous industrial application potentials, ease of culture handling and the ease for scale-up during production (Pratush et al., 2013). The 3-D structure of lipases from different microbial sources are not exactly alike, they exhibit high sequence diversity (Gupta et al., 2015). As a result, these enzymes are unique and specific to the type of bioconversion processes they catalyze, finding relevance in a wide array of industrial processes (Mehta et al., 2017). They are found to be useful in catalyzing various reactions synonymous to food, pharmaceuticals, medical and diagnosis, dairy, fatty acid, leather, cosmetic, detergent, beverage and paper industries. (Sharma et al., 2017).
The implication that microorganisms are ubiquitous is that they would be found in almost any natural habitat (soil, water, air, leaves and tree trunks), soil being the most preferred source as it serves as reservoir for diverse microorganisms (Okafor, 2007). Nevertheless, new habitats especially marine environments have now been included in the list to be studied (Metin and Bakir, 2017). Lipase producing microorganisms are not left out in the microbial population found almost everywhere as they have been reported to be isolated from soil (Sharma et al., 2017; Colla et al., 2016; Alhamdan and Alkabbi, 2016), marine environment (Lodha, 2018), wastewater from fish industry (Suharjono, 2015), agro-industrial waste (Maldonado et al., 2014), waste volatile substances (Muthumari et al., 2016), air (Abada, 2008), palm-oil mill effluent (Nwuche and Ogbonna, 2011), intestine of silkworm (Feng et al., 2011) and on human skin (Jagtap and Chopade, 2015). Cultures isolated ab initio from natural sources are usually identified and classified based on morphological characteristics and cell types. However, due to the advanced techniques in genomics and proteomics, isolates can be classified based on sequence of ribosomal RNA in the 16S of the small subunit (SSU) of the prokaryotic ribosome and 18S ribosomal units of eukaryotes (Boonmahome and Mongkolthanaruk, 2013; Alhamdani and Alkabbi, 2016; Sharma et al.,2017;). Harnessing microorganisms for their potential metabolic activities is largely dependent on the culture media composition. The approach for optimization process is usually carried out one variable at a time and the media composition factors subjected to variation include; nutritional factors (carbon and nitrogen sources) and physicochemical factors (incubation time, temperature,
pH, presence of lipid as inducer, cofactors, inhibitors) (Soleyman et al., 2017). Therefore, different media have different stimulatory effect on lipase production (Dhiman and Chapadgaonkar, 2013). This study seeks to evaluate the lipase producing potentials of organisms from two sources, determine the optimal conditions for scale-up production, isolate and identify the organism with better lipase producing ability
Materials and Methods Collection of sample Soil sample was taken from a mechanic’s workshop opposite Forestry College, Jericho, G.R.A, Ibadan and water sample taken from Ado-awaye Lake. Samples collected were kept in a sterile container and taken to the Biotechnology Laboratory, Forestry Research Institute of Nigeria (FRIN) for analysis. Isolation of bacteria Isolation of microorganism from the water and soil samples was carried out by serial dilution (1:10), using the spread plate method. Aliquots of 0.1 ml were inoculated on nutrient agar that was previously sterilized by autoclaving at 15 lb pressure (121 ) for 15min (Salle, 1974). Plates were incubated at 37
for 24 h in an inverted position. Isolated colonies were purified by
picking and streaking repeatedly on nutrient agar plates to obtain pure cultures. Pure bacterial cultures were kept in slants and stored at 4
for further use.
Primary Screening for lipase producing Bacteria Pure isolated bacterial cultures were screened for lipase activity using tributyrin agar medium (Ahmed et al., 2010). The tributyrin agar medium pH 7.5 was prepared with 5g/L peptone, 3g/L
yeast extract, 15g/L agar, 10 ml/L tributyrin in distilled water. The medium was sterilized for 15min by autoclaving with 15 lb pressure at 121°C and cooled. The aliquot was transferred to petri dishes and allowed to solidify. A loopful of each pure culture was streaked each onto tributyrin agar plates separately and incubated at 37°C for 24 h. After incubation, the clear zone of hydrolysis around the colonies indicated the presence of lipase activity. Positive cultures that showed maximum zone of hydrolysis were selected for further studies. Rhodamine olive oil agar medium pH7.0 was prepared with 5g/L peptone, 3g/L yeast extract, 15g/L agar, 4g/L NaCl in distilled water, sterilized for 15 min by autoclaving with 15 lb pressure at 121
and cooled to 60 . 31.25 ml/L olive oil, 10ml/L sterilized filtered rhodamine dye
(1mg/ml) was added. Aliquots were transferred to petri dishes and allowed to solidify. A loopful of each pure culture was streaked onto rhodamine olive oil agar plates separately and incubated the inoculated plates at 37
for 48 h. After incubation, formation of orange flourescent halos
around bacterial colonies visible upon UV irradiation indicates lipase production. Phenol red agar pH 7.4 was prepared with 5g/L peptone, 3g/L yeast extract, 15g/L agar,1g/L CaCl2 , 0.1M NaOH to adjust pH to 7.4 in distilled water, sterilized for 15 min by autoclaving with 15 lb pressure at 121
and cooled to 60
10ml/L phenol red dye (1mg/ml) and10ml/L
substrate (olive oil) were added. Aliquots were transferred to petri dishes and allowed to solidify. A loopful of each pure culture was streaked onto phenol red agar and incubated at 37
for 48h.
After incubation, a change of color from orange to pink was observed. This indicated the release of fatty acids due to lipolysis. Tween80 agar medium pH7.0 was prepared with 10g/L peptone,20g/L agar, 5g/L NaCl, 0.1g/L CaCl2.2H2O, in distilled water, sterilized for 15 min by autoclaving with 15 lb pressure at 121⁰C
and cooled to 45 . 10 ml/L of Tween 80 was added to the cooled media. Aliquots were transferred to petri dishes and allowed to solidify. A loopful of each pure culture was streaked onto Tween80 agar plates separately and incubated the inoculated plates at 37
for 48h. After
incubation, a white precipitate around colony indicates lipase activity.
Secondary Screening for Lipase Production All the selected lipase producing bacterial cultures were screened for production of lipase. The production media (pH7.0) was prepared with 5g/L peptone, 5g/L beef extract in distilled water, autoclaved for 15 min at 15lb pressure (121°C) and cooled to about 60°C before the addition of 10 ml/L olive oil. 1.0 ml of overnight grown selected lipase producing bacterial cultures was inoculated in 100 ml of production medium in 250ml Erlenmeyer flasks separately and incubated at 35
for 24h. Culture was centrifuged at 10,000rpm for 10min after incubation, supernatant
was used to assay for lipase. Lipase activity assay Lipase activity was assayed for using p-nitrophenyl laureate (p-NPL) as substrate. Reaction mixture contained 2µL of 0.03g/L p-NPL, 6µL of 0.05M phosphate buffer and 2µL of enzyme extract (lipase) incubated at 30oC for 10 min. Reaction was terminated by adding 30µL of absolute ethanol. Absorbance was measured using Nanodrop spectrophotometer at wavelength of 380 nm for lipase reaction. One unit of lipase activity is defined as an enzyme releasing 1µmol of free p-nitrophenol per minute. Optimization of Conditions for Lipase Production Determination of optimum incubation time
The organism was grown in nutrient broth containing appropriate 1% oil (olive oil); pH of the medium to was adjusted to 9.0. Cultures were incubated at 30
and samples were taken at 3 h
interval over a period of 12 h for lipase assay. Determination of a suitable carbon source Different kinds of carbon sources (Tween20, Tween80, Olive oil Glucose and Fructose) were used as they might induce lipase production. Bacterial culture was inoculated into nutrient broth containing each kind of carbon source. Cultures were incubated at 30
for 12 h taking samples
for lipase assay every 3 h. Optimum pH The organism was grown in nutrient broth containing appropriate 1% oil (olive oil); media were adjusted to different pH (3, 5, 7, 9 and 11) and incubated at 30
for 12 h.
Determination of optimum inoculum volume Cell cultures of different volume (1000, 2000, 3000 and 4000 µL) were inoculated into nutrient broth containing appropriate 1% oil (olive oil). Cultures were incubated at 30°C for 12 h taking samples for lipase assay every 3h.
Determination of suitable nitrogen source Different kinds of nitrogen sources (Yeast extract, Peptone, Casein, Ammonium Nitrate and Potassium Nitrate) were used in production media as they might affect lipase production.
Bacterial culture was inoculated into nutrient broth containing each kind of nitrogen source. Cultures were incubated at 30
for 12 h taking samples for lipase assay every 3h.
Molecular Characterization Molecular characterization was studied based on 16S rRNA gene sequencing, BLAST search analysis and phylogenetic tree construction. 16S rRNA gene sequencing Genomic DNA of the lipase positive strain was extracted using procedure as stated in commercially available kit, Quick DNA TM Fecal/Soil Microbe Microprep Kit (Zymo research Corp).The 16S rRNA gene was amplified via PCR, and then the amplicon was sequenced After the isolation, qualitative and quantitative analysis was carried out using 50ng of DNA for DNA amplification. For the amplification of 16S region, the following two degenerate primers were designed Table 1. Table 1: Primer sequence for amplification Primer
Sequence
Forward
5´-AGAGTTTGATCCTGGCTCAG-3´
Reverse
3´-ACGGCTACCTTGTTACGACTT-5´
Cycling conditions for PCR: initial denaturation at 96 at 96
for 1 min, annealing at 53
extension at 68
for 5 min; and 35 cycles of denaturation
for 45 s and extension at 68°C for 90 s; followed by a final
for 10 min. 5μl of PCR product was mixed with equal volume of gel loading
dye and was loaded onto agarose gel. Electrophoresis the samples on1% agarose gel. PCR
products were purified using the gel elution kit, and subjected the product obtained to DNA sequencing process using Big Dye® Terminatorv3.1 Cycle sequencing kit.
BLAST search analysis of 16S rRNA gene sequence The nucleotide sequence obtained with 16S rRNA gene was subjected to homology search using the Basic Local Alignment Search Tool (BLAST) program of the National Centre for Biotechnology Information (NCBI) with default parameters.
Phylogenetic tree construction Representative sequences of similar neighbors in BLAST analysis was retrieved and aligned using multiple alignment program. The multiple alignment file was used to create neighborjoining tree using MAFFT (Multiple Alignment using Fast Fourier Transform).
RESULTS ISOLATION AND IDENTIFICATION OF A LIPASE POSITIVE BACTERIUM FROM SOIL AND WATER SAMPLES.
Soil sample and water sample were examined for the presence of lipase positive strains using a screening method suitable for detection of lipase producers as described in materials and methods. Lipase producer strains in the samples were identified by the formation of orange fluorescent halos around the colonies when olive oil-rhodamine B spread plates incubated at 37°C were exposed to UV light at 350 nm (Plate 1). Olive oil is used as lipase substrate and rhodamine B is the indicator of lipase activity.
Plate 1: UV determination at 350 nm of lipase activity on rhodamine agar plate. Plate A and B from soil, C from water and D as control. All plates were incubated at 37 .
Plate 2: Lipase activity on phenol red agar plate. Plate A and B from soil, C from water and D as control. All plates were incubated at 37 .
Lipase producer strains in the samples were identified by the formation of pink halos around the colonies when olive oil-phenol red spread plates were incubated at 37°C. (Plate 2).
4.2. Determination of lipase activity from soil and water samples. Nutrient broth supplemented with 1% olive oil was used as lipase production media. To determine the organism with lipase activity, cultures were grown at 30
for a period of 18 h and
supernatant was assayed for lipase activity using spectrophotometric method (Table 2).
Table 2: Determination of lipase activity in Water and Soil samples
Isolate
Lipase activity (U/L)
Soil
99.69
Water
45.55
OPTIMIZATION OF GROWTH CONDITIONS
Effect of incubation time Nutrient broth supplemented with 1% olive oil was used as lipase production media. To determine the effect of incubation time on lipase activity and select organisms with higher lipase
activity, cultures were grown at pH 9, 30oC for a period of 12 h and supernatant was taken every 3 h to assay for lipase activity using spectrophotometric method. Lipase activity was prominent in soil as highest activity was valued 364.82 U/L at 12 h, whereas that of water was valued to 162.85 U/L at 9 h (Figure 1).
Lipase activity (U/L)
500
water Soil
400 300 200 100 0 0
2
4
6
8
10
12
Time (h) Figure 1: Determination of lipase activity from water and soil sample Effect of carbon source Nutrient broth supplemented with different carbon sources were used as lipase production media. To determine the effect of different carbon sources ( Tween 20, Tween 80, Glucose and Fructose) on
lipase
production
in organism with higher lipase activity from previous
experiment, cultures were grown at 30oC in each production medium over a period of 12 h and supernatant was taken to assay for lipase ativity using spectrophotometric method.The medium containing olive oil had the highest lipase activity at 364.81 U/L while Tween 20, Tween 80 ,
Glucose and Fructose had activity at 202.39, 313.69, 61.8 and 106.57 U/L respectively (Figure 2).
600
Lipase activity (U/L)
500 400 300 200 100 0 Tween 20 Tween 80 Glucose
Olive oil
Fructose
Carbon Source ( 1%) Figure 2: Effect of different carbon sources on isolate from soil Effect of pH Nutrient broth of different pH supplemented with 1% olive oil was used as lipase production media. To determine the effect of different pH (3,5,7,9 and 11) on lipase production, cultures were grown in production media of different pH at 30OC over a period of 12 h and supernatant was taken to assay for lipase ativity using spectrophotometric method.The least lipase activity, 84.66 U/L was recorded at pH 3, while pH 5, 7, 9 had 158.99, 230.75, 360.62 U/L respectively and the highest activity of 432.28U/L was at pH11 (Figure 3).
Lipase activity (U/L)
600 500 400 300 200 100 0 3
5
7
9
11
pH Figure 3: pH optimization of isolate from soil Effect of inoculum volume Nutrient broth inoculated with different volume of cell cultures supplemented with 1% olive oil was used as lipase production media. To determine the effect of different inoculum volume ( 1000, 2000, 3000, 4000 µL) on lipase production, cultures were grown in production media of different inoculum volume at pH 11, 30OC over a period of 12 h and supernatant was taken to assay for lipase ativity using spectrophotometric method. The highest lipase activity recorded was 528.54 U/L obtained from the medium containing 4000µL inoculum volume (Figure 4).
Volume
600
Lipase Activity (U/L)
500 400 300 200 100 0 1000
2000
3000
4000
Volume (µL)
Figure 4: Inoculum volume optimization of isolate from soil Effect of Nitrogen sources Medium containing different nitrogen sources supplemented with 1% olive oil was used as lipase production media. To determine the effect of different nitrogen sources (Yeast Extract,Peptone, Casein, Ammonium Nitrate and Potasssium Nitrate) on lipase production, cultures media were set at pH 11, temperature 30
over a period of 12 h. The supernatant was taken to assay for
lipase ativity using spectrophotometric method. Lipase activity of 490.72, 445, 225.6, 482 U/L was recorded for Yeast Extract, Peptone, Casein, Ammonium Nitrate respectively while no activity was detected in the medium that had Potassium Nitrate as its nitrogen source (Figure 5).
600
Lipase activity (U/L)
500
400
300
200
100
0
Yeast Extract
Peptone
Casein
Ammonium Potassium Nitrate Nitrate
Nitrogen source (1%) Figure 5: Effect of different nitrogen sources on isolate from soil DNA Sequence Analysis/ Molecular Identification of Bacterial Isolates BLAST analysis revealed that all the amplified DNA sequences were bacteria in the phyla Proteobacteria and Firmicutes and belonged to two different genera. The 16S rRNA sequence of the isolate B_27-F_C05_08 illustrated a high similarity to the members of the genus Pseudomonas, showing 98% similarity with Pseudomonas sp. RH-7 (GenBank Accession no. KT715742), Pseudomonas sp. strain GJY (GenBank Accession no. MG585077), Pseudomonas aeruginosa strain GHJ15 (GenBank Accession no. MG396976) and Pseudomonas sp. strain S2 (GenBank Accession no. MF425660). Isolate C_27-F_D05_11 belongs to the genus Bacillus showing 78% similarity with Bacillus sp. strain WP7 (GenBank Accession no. KY347914), Bacillus cereus strain M-1 (GenBank Accession no. EF633995), Bacillus thuringiensis strain WS 2625 (GenBank Accession no. Z84587) and Bacillus cereus isolate DRG3 (GenBank Accession no. AM419192).
Table 3: Molecular Characterization of Bacterial Isolates Sample ID.
Organisms identified by BLAST
IDENT (%)
Sequence length (Bp)
B_27-F_C05_08
Pseudomonas sp. RH-7
98
1101
78
1180
Pseudomonas sp. strain GJY Pseudomonas aeruginosa strain GHJ15 Pseudomonas sp. strain S2 C_27-F_D05_11
Bacillus sp. strain WP7 Bacillus cereus strain M-1 Bacillus thuringiensisstrain WS 2625 Bacillus cereus isolate DRG3
NOTE: B_27-F_C05_08, isolate from soil; C_27-F_D05_11, isolate from water
Figure 6: Phylogenetic tree of the isolate B_27-F_C05_08 and C_27-F_D05_11
DISCUSSION Lipases are enzymes capable of hydrolyzing triacylglycerol. They are obtained from different sources, bacteria being one of them are found in almost every habitats. Screening samples collected from different habitats as described in materials and methods indicated the presence of lipase positive strains from both sources. Strains found in soil sample from a mechanic’s workshop exhibited higher lipase activity when compared to those found in water sample from Iyake lake. This result supports the evidence that the environment influences the status and behaviour of the living system (Verma and Agarwal, 2005). This is explainable that the organisms isolated from the soil are accustomed to the presence of oils and its analogues whereas organisms from water are not.
Upon identifying and isolating organisms with highest lipase activity, different growth conditions that influence lipase production were varied using one variable at a time approach for the optimization of lipase production Starting with incubation time, highest lipase activity was recorded at 12h after which a noticeable decline occurred thereafter. The decline could be related to consumption of nutritional elements. Gupta et al., 2004 reported that maximum lipase activity occur from few hours to several days. As a result of this finding, incubation time for subsequent cultures did not exceed 12 h period. Lipase has been reported to be an inducible enzyme and carbon has been a major factor for the expression of lipase activity (Bakir and Metin, 2017). The effect of different carbon source showed higher activity on lipidic substrate (olive oil) and its analogue (Tween20 and Tween80) with olive oil being the carbon source with the highest lipase activity. Glucose showed the least lipase activity. Catabolic repression may be reponsible for this as sugar molecules act as repressors in lipase synthesis (Soleymani et al., 2017). After determining olive oil as the carbon source with highest lipase activity, pH of the production medium was varied (pH 3, 5, 7, 9 and 11) in order to determine the optimum pH. High lipase activity was exhibited in the pH range of 7-11 while optimum activity was observed at pH 11. In the acidic, reduction of enzyme activity was observed. The present report shows that the organism is an alkalophilic bacteria. Bacteria with lipase producing ability have been reported to prefer pH around 7. However, maximum activity has also been reported in pH higher than 7 (Gupta et al., 2004).
Salihu et al., 2011 reported that inoculum volume has direct proportional effect on lipase activity. Our findings supported this claim as the medium inoculated with the highest culture
volume, 4000µL showed highest lipase activity. However, Thakur et al., 2014 reported inoculum volume is inversely proportional to lipase activity because cells are struggling for nutrients. The last factor that was varied was nitrogen source. Both organic and inorganic nitrogen sources have reported to supply nitogen, cell growth factors and amino acids necessary for enzyme synthesis (Thakur et al., 2014). Three (Yeast Extract, Peptone and Ammonium Nitrate) out of five tested nitrogen sources showed really high lipase activity. Potassium Nitrate did not show any lipase activity. In conlusion, the lipase producing bacteria strain, B_27-F_C05_08 isolated from the mechanic’s workshop was identified as Pseudomonas aeruginosa whose preferred carbon source, nitrogen source, pH and incubation time are olive oil, ammonium nitrate, pH 11 and 12 h respectively.
Conclusion Utilization of enzymes synthesized from biological organism for industrial application is gaining global attention due to their numerous industrial and biotechnology applications. Increased enzyme production from biological sources can be achieved by culture media optimization. Different variables such as substrates concentration, inducers, nitrogen and carbon sources which facilitated increased lipase production have been subjects of focus to researchers. In this work, improved lipase production from Pseudomonas aeruginosa was achieved via optimization of growth and physiological parameters. The high yield of lipase obtained from this process is an indication of the possibility for its large scale production and effective application for industrial/biotechnological purposes. Producing this important enzyme from innate producing organisms such as Pseudomonas aeruginosa is economic and environmentally advantageous.
Declaration of interest statement The authors declare no conflict of interest.
Funding: This research did not receive any specific grant from funding agencies in the public, commercial, or not-for-profit sectors.
REFERENCES
Abada, E. 2008. Production and Characterization of a Mesophilic Lipase isolated from Bacillus stearothermophilus AB-10. Pakistan Journal of Biological Sciences. 11(8): 1100-1106. Alhamdani, M. and Alkabbi, H. 2016. Isolation and Identification of lipase producing bacteria from oil contaminant soil. Journal of Biology, Agriculture and Healthcare. 6(2):1-7. Alhamdani, M. and Alkabbi, H. 2016. Isolation and Identification of lipase producing bacteria from oil contaminant soil. Journal of Biology, Agriculture and Healthcare. 6(2):1-7. Boonmahome, P. and Mongkolthanaruk, W. 2013. Lipase producing Bacterium and its Enzyme Characterization. Journal of Life Sciences and Technologies.1(4):196-200. Colla, L., Primaz, A., Benedetti, S., Loss, R., Lima, M. And Reinehr, C. 2016. Surface response method for the optimization of lipase production under submerged fermentation by filamentous fungi. Brazilian journal of microbiology.47: 461-467. Dhiman, S. and Chapadgaonkar, S. 2013. Optimization of Lipase Production Medium for a Bacterial Isolate. International Journal of ChemTech Research.5(6):2837-2843.
Feng, W., Wang, X., Zhou, W., Liu, G. and Wan, J. 2011. Isolation and Characterization of lipase-producing bacteria in the intestine of the Silkworm, Bombyxmori, reared on different forage. Journal of Insect Science. 11 (135): 1-10. Gupta, R., Kumari, A., Syal, P. and Singh, Y. 2015. Molecular and functional diversity of yeasts and fungal lipases: Their role in biotechnology and cellular physiology. Progress in Lipid Research 57:40-54. Jagtap, S. and Chopade, B. 2015. Purification and Characterization of Lipase from Acinetobacter haemolyticus TA-106 Isolated from human skin. Songklanakarin Journal of Science and Technology.37 (1): 7-13. Lodha, K. 2018. Isolation and Screening of multiple enzyme producing Alkaphilic Bacillus Species from Lonar lake. International journal of Biology Research.3(1):164-168. Maldonado, R., Macedo, G. and Rodrigues, M. 2014. Lipase production using Microorganisms from different Agro-Industrial By-products. International Journal of Applied Science of Technology.4(1):1-8. Mehta, A., Bodhu, U. and Gupta, R. 2017. Fungal lipases: a review. Journal of Biotechnology Research 8:58-77. Metin, K. and Bakir, Z. 2017. Production and Characterization of an Alkaline lipase from Thermophilic anoxybacillus sp. HBB16. Chemical and Biochemical Engineering Quarterly 31(3):303-312. Muthumari, G., Thilagarathi, S. and Hariram, N. 2016. Industrial enzymes: Lipase Produicing Microbes from waste volatile substances. International Journal of Pharaceutical Sciences and Research.7(5): 2201-2208.
Narayanan, P. 2014. Enviromental Pollution: Principles, Analysis and Control. CBS Publishers & Distributors.New Delhi. 671pp. Nwuche, C. and Ogbonna, J. 2011. Isolation of Lipase Producing Fungi from Palm Oil Mill Effluent (POME) Dumpsites at Nsukka. Brazilian Archives of Biology and Technology. 54(1) :113-116. Okafor, N. 2007. Modern Industrial Microbiology & Biotechnology. Enfield, New Hampshire: Science Publishers. 551pp. Pratush, A., Gupta, A., Vyas, G. and Sharma, P. 2013. Microbiology application.Bhalla Publishers. Dehradun, India. pp64-83. Sangeetha, R., Geetha, A. and Arulpandi, I. 2010.Concomitant production of protease and lipase by Bacillus licheniformis VSG1: Production, purification and characterization.Brazilian Journal of Microbiology. 41: 179-185. Sarmah, N., Revathi, D., Sheelu, G., Rani, Y., Sridhar, S. and Mehtab, V. 2018. Recent Advances on Sources and Industrial Applications of Lipases. Biotechnology progress.34(1): 1-24. Sharma, P., Sharma, N., Pathania, S. and Handa, S. 2017. Purification and characterization of lipase by Bacillus methylotrophicus PS3 under submerged fermentation and its application
in
detergent
industry.
Journal
of
genetic
engineering
and
biotechnology.15:369-377. Sharma, P., Sharma, N., Pathania, S. and Handa, S. 2017. Purification and Characterization of Lipase by Bacillus methylotrophicus PS3 under submerged fermentation and its application
in
detergent
Biotechnology.15:369-377.
industry.
Journal
of
Genetic
Engineering
and
Singh, M., Chandraveer. and Tripathi, A. 2017. Isolation and Screen ing of Lipases producing Microorganisms from Natural Sources. Indian journal of ecology. 44(1): 19-23. Soleyman, S., Alizadeh, H., Mohammadian, H., Rabbani, E., Moazen, F. and Sadeghi, H. 2017. Efficient Media for High Lipase Production: One Variable at a Time Approach. Avicenna Journal of Medical B iotechnology. 9(2): 82-86. Suharjono, N. 2015. Activity Assay and Identification of Lipolytic Bacteria from Wastewater FishIndustry at Mucar, Banyuwang, Indonesia. International Journal of ChemTech Research.8(11): 377-383. Waltes, M., Morgan, N., Rockey, J. and Higton, G. 2001. Industrial Microbiology: An Introduction.Blackwell Science Ltd. London.302pp.