Isolation of an organic solvent-tolerant bacterium Bacillus licheniformis PAL05 that is able to secrete solvent-stable lipase
∗
Periasamy Anbu Byung Ki Hur
Department of Biological Engineering, Inha University, Incheon, South Korea
Abstract In this study, seven lipase-producing bacterial strains were isolated from salt-enriched and cattle farm soil samples after incubation in toluene- and benzene-enriched media. One strain (PAL05) showed significantly greater lipase activity on spirit blue agar medium and stability in organic solvents. The positive strain (PAL05) was identified as Bacillus licheniformis by 16S rRNA gene sequencing. Lipase production was optimized in a medium containing glycerol as the carbon source and Tween 80 as an inducer (0.5% glycerol + 0.5% Tween 80) at pH 8.0 and a temperature of 30 ◦ C. In addition, the enzyme was moderately halotolerant as it exhibited increased activity in the presence of 2.5% NaCl. Optimized
conditions increased the lipase production threefold. Crude lipase retained its activity for 14 days of incubation in the presence of various organic solvents at a level of 25% and 50%. The enzyme was stable at 25% in most solvents; some of the solvents such as hexane, benzene, and ethanol actually stimulated enzyme activity. The organic solvent stability of the lipase produced by the strain PAL05 enables the enzyme to be used as a potential biocatalyst for ester synthesis and other C 2014 International applications in nonaqueous conditions. Union of Biochemistry and Molecular Biology, Inc. Volume 61, Number 5, Pages 528–534, 2014
Keywords: organic solvent tolerance, lipase, Bacillus, optimization
1. Introduction Organic solvents are toxic to microorganisms and cause cell lysis because of disruptive effects on the cell membrane [1], as well as the internal structural and functional integrity of the cell, once it penetrates the cell [2]. However, some bacteria develop stability toward organic solvents by various adaptations such as solvent efflux pumps, rapid membrane repair, lower cell membrane permeability, increased membrane rigidity, and decreased cell surface hydrophobicity [3]. Recently, Segura et al. [4] have isolated organic solvent-stable bacteria, which are stable at low concentrations of organic solvents. Enzymes used for ester synthesis in organic solvents should be stable at high solvent concentrations to obtain high yields. Many solventstable lipases have been reported to be produced by several
Abbreviations: pNPP, 4-nitrophenyl palmitate; DMF, dimethyl formamide. ∗ Address
for correspondence: Dr. Periasamy Anbu, PhD, Department of Biological Engineering, Inha University, Incheon, 402-751, South Korea. Tel.: +82-32-860-7510; Fax: +82-32-872-4046; e-mail: [email protected]. Received 29 October 2013; accepted 27 December 2013 DOI: 10.1002/bab.1202 Published online 25 March 2014 in Wiley Online Library (wileyonlinelibrary.com)
528
Pseudomonas species [5, 6]; however, because of their high tolerance for organic solvents, Bacillus species have captured the attention of many researchers searching for a source of solvent-stable lipases and other enzymes useful for industrial applications. Lipases are triacylglycerol acylhydrolases (EC 3.1.1.3) that catalyze the hydrolysis of fats and oils to yield glycerol and free fatty acids under aqueous conditions [7]. In addition, the lipases are also involved in conversion reactions such as esterification, interesterification, transesterification, alcoholysis, acidolysis, and aminolysis [8]. Microbial lipases have many industrial and biotechnological applications as product components and manufacturing tools in areas such as synthesis of cosmetics, detergents, food, pharmaceuticals, esters, and biosurfactants [9–11]. Industrial demand for lipase stimulates the search for new enzyme sources. Two advantages of enzymatic synthesis in organic solvents or aqueous solution containing organic solvents are increased solubility of nonpolar substrates and elimination of microbial contamination in the reaction mixture [12]. Methods such as chemical modification, immobilization, entrapment, protein engineering, and directed evolution have been employed to stabilize the enzymes in organic solvents [13]. However, naturally stable enzymes with high activity in organic solvents are preferred for industrial applications. In this study,
we isolated a potent organic solvent-tolerant strain of Bacillus licheniformis and analyzed the secretion of a solvent-stable lipase. Furthermore, we optimized the conditions to maximize the lipase production and evaluated the solvent stability of lipase using various organic solvents. B. licheniformis has been reported to produce a variety of extracellular enzymes such as protease [14], amylase [15], lipase [16], and xylanase [17]. However, no reports are available on an organic solvent-stable lipase from a solvent-tolerant strain of B. licheniformis.
2. Materials and Methods 2.1. Isolation and screening of organic solvent-stable microorganisms The organic solvent-stable microorganisms were isolated according to the method of Ogino et al. [18]. Soil samples were collected from salt-enriched areas (Incheon) and cattle farms (Yongin-Si) in South Korea. About 1 g of the soil sample was suspended in 10 mL of sterile water. After shaking, 5 mL of the suspension was added to a 250 mL bottle containing 25 mL of LB broth supplemented with 2.5% toluene and 2.5% benzene (1:1 mixture). The cultivation vessels were sealed with chloroprene rubber stoppers to prevent evaporation of the organic solvents. The bottles were incubated at 37 ◦ C on a shaker at 180 rpm for 3 days. Then, 5 mL aliquots were transferred into the same medium and cultured again under the same conditions for 3 days. Aliquots of the culture suspension were diluted and spread on spirit blue agar (Tryptone, 10 g/L; yeast extract, 5 g/L; agar, 20 g/L; spirit blue, 0.15 g/L; lipid reagents containing a mixture of tributyrin and Tween 80, 30 mL/L; pH 6.8) without any organic solvent [19]. After incubation at 37 ◦ C for 36 H, the enzyme-producing strains were purified and subjected to another round of screening on spirit blue agar. Spectrophotometric analysis further confirmed the enzyme-producing activity of these strains.
2.2. Selection of a highly potent solvent-stable strain The bacteria were inoculated into 25 mL of LB medium and incubated at 30 ◦ C at 180 rpm for 4 H. About 0.5 mL of the culture suspension was transferred into 50 mL of lipase production medium composed of dipotassium hydrogen orthophosphate, 1.5 g/L; MgSO4, 0.05 g/L; CaCl2, 0.025 g/L; FeSO4, 0.015 g/L; and ZnSO4, 0.005 g/L. The inoculated flasks were incubated at 37 ◦ C at 180 rpm for 48 H. After incubation, the cultures were centrifuged at 11,100g at 4 ◦ C for 10 Min. Bacterial growth was determined at 600 nm. To obtain a highly organic solvent-stable lipase-producing strain, the strains selected by plate screening were further screened using organic solvents with various log P values. Each solvent was mixed with 1 mL of the supernatant to reach a concentration of about 20%. The tubes were covered with aluminum foil and incubated at 37 ◦ C with shaking at 100 rpm for 24 H. The residual lipase activity was measured. Based on the results of agar-based screening, spectrophotometric analysis, and organic solvent stabilities, the strain with highest activity
Biotechnology and Applied Biochemistry
and solvent stability was designated as PAL05 and was further identified by morphological, physiological, and 16S rRNA gene-sequencing methods. For 16S rRNA gene sequencing, the genomic DNA was extracted using a genomic purification kit following its manual instructions (Promega, Madison, WI, USA). The DNA was then amplified by PCR using the universal 16S rRNA gene primers: 8-27F (5 -AGAGTTTGATCCTGGCTCAG-3 ) and 1472R (5 -TACGGYTACCTTGTTACGACTT-3 ). The PCR product was electrophoresed on agarose gel electrophoresis, and the PCR product (16S rRNA band) was purified by SV gel and the PCR Clean up system (Promega). The 16S rRNA gene sequence was then compared with other sequences available in the nucleotide database by using the BLAST algorithm, and the phylogenetic tree was constructed using the neighbor-joining method (www.ncbi.nlm.nih.gov/blast).
2.3. Optimization of solvent-stable lipase production 2.3.1. Effect of incubation period on lipase production The effect of incubation time on lipase production was studied at 24 H intervals up to 72 H. Lipase activity of the cellfree supernatants obtained by centrifugation (11,100g for 10 Min at 4 ◦ C) was measured every 24 H intervals.
2.3.2. Effect of different carbon sources on lipase production Carbon sources composed of carbohydrates and lipids that were used at a concentration of 1%: glucose, lactose, glycerol, sucrose, maltose, olive oil, Tween 80, and tributyrin. Different concentrations of selected carbon sources were tested. The carbon sources were separately sterilized and then aseptically added to autoclaved medium.
2.3.3. Effect of pH, temperature, and NaCl on lipase production Lipase activity was determined at different pH values ranging from 6.0 to 10.0 and at temperatures ranging from 20 to 60 ◦ C. The pH of the medium was adjusted prior to autoclaving. The NaCl concentrations used varied from 0.25% to 7.5%.
2.4. Lipase assay Lipase activity was quantitatively assayed using 4-nitrophenyl palmitate (pNPP) as the substrate according to the method described by Winkler and Stuckmann [20]. Briefly, 10 mL of isopropanol containing 30 mg of pNPP was mixed with 90 mL of 0.05 M phosphate buffer (pH 8.0) containing 207 mg of sodium deoxycholate and 100 mg of gum arabic. A total of 2.4 mL of freshly prepared substrate solution was prewarmed to 37 ºC and mixed with 0.1 mL of enzyme solution. After incubation of this reaction mixture at 37 ºC for 15 Min, the absorbance was measured at 410 nm against an enzyme-free control. One enzyme unit was defined as the amount of enzyme that liberated 1 μmol of 4-nitrophenol/Min under the assay conditions.
529
Biotechnology and Applied Biochemistry
TABLE 1
Screening for solvent-stable lipase produced by positive strains on spirit blue agar medium
Source Salt-enriched soil
Cattle farm soil
Isolate no.
Lipase activity
PAL01
++
PAL02
+++
PAL03
++
PAL04
+
PAL05
+++
PAL06
+
PAL07
+
Symbol: +++, high activity; ++, moderate activity; +, low activity
2.5. Effect of organic solvents on the stability of crude enzyme Crude lipase was filtered through a 0.22 μm filter membrane. The organic solvents tested were benzene, chloroform, dimethyl formamide (DMF), ethanol, hexane, methanol, propanol, and toluene. Filtered crude lipase was mixed with each solvent at 25% and 50% (v/v). The mixtures were incubated at 30 ◦ C with shaking at 100 rpm for 2 weeks. After incubation, an aliquot was carefully withdrawn from the solution or from the aqueous phase in case of water-immiscible solvents. Residual lipase activity was measured. The control contained enzyme solution without any organic solvents (0% v/v). Enzyme stability was expressed as the remaining lipase activity relative to the control.
FIG. 1
Effect of incubation period on the growth (A) and lipase production (B) by positive strains. Each point represents the mean ± SEM of three independent experiments.
3. Results and Discussion The soil samples (collected from salt-enriched areas and cattle farms) were mixed with equal concentrations of tolueneand benzene-enriched medium (5% solvent). Solvent-tolerant bacteria were able to adapt to this condition. After incubation, enriched samples were spread on spirit blue agar medium. About 15 morphologically different organic solvent-tolerant bacterial colonies grew; however, only seven colonies (three from the salt-enriched soil sample and four from the cattle farm soil sample) produced a clear zone of substrate hydrolysis around the bacterial colonies. The seven positive colonies were isolated from the original spirit blue agar plate and screened again on the same medium for confirmation. Of the seven lipase producers, PAL02 and PAL05 produced the largest zones of clearance on spirit blue agar medium (Table 1). A spectrophotometric method was used to determine enzyme activity, and the highest enzyme activity was observed for PAL05; the other strains, including PAL02, showed low levels of enzyme activity. The growth rate and enzyme activity correlated for PAL02, but not for PAL05. For PAL02, growth rate and enzyme activity were maximized at 24 H. However,
530
PAL05 showed maximum growth on day 3 and maximum enzyme activity on day 2 (Fig. 1A and B). Among the seven positive strains, PAL05 was the most efficient, with the highest enzyme activity and growth rate. To choose the more efficient solvent-stable lipaseproducing strain, these two strains (PAL02 and PAL05) were further tested in various hydrophilic and hydrophobic organic solvents, including benzene, ethanol, hexane, and toluene. After mixing the supernatants with 20% (v/v) organic solvent, residual lipase activity was determined. The strain PAL05 demonstrated a wide range of stability in all solvents. PAL02 was moderately stable in a few organic solvents, but significantly less than PAL05 (Table 2). Therefore, PAL05 was selected for further study based on the spectrophotometric and solvent stability data. The selected potent strain was identified by morphological, physiological, and 16S rRNA gene sequence methods. PAL05 is a Gram-positive, aerobic, and rod-shaped bacterium. Phylogenetic analysis confirmed that the strain belongs to the Bacillus species (Fig. 2) and is highly homologous to B. licheniformis (99%). The 16S rRNA sequence (1,469 bp)
Solvent-Stable Lipase from B. licheniformis
TABLE 2
Screening for solvent stability of two potent strains in various organic solvents
Organic solvent
PAL02
PAL05
100
100
20% Ethanol
69
126
20% Hexane
96
121
20% Toluene
61
100
20% Benzene
77
93
Control (without solvent)
of this strain was deposited in GenBank (NCBI accession no. KC437094).
3.1. Organic solvent-stable lipase production Lipase production for commercial applications can be improved by optimizing the nutritional (carbon sources) and physical (pH and temperature) conditions for production. Many researchers have previously reported the optimization of lipase production from various microorganisms [21–24], but studies specific
FIG. 2
to organic solvent-stable lipases are very limited [25, 26]. Of the various carbon sources tested, a significant level of lipase production was obtained in glycerol (1%) and Tween 80 (1%) after 48 H of incubation (Fig. 3A). Low levels of enzyme activity were observed in case of other carbon sources. In contrast to this result, Volpato et al. [26] reported that glycerol acts as an inducer for the synthesis of organic solventtolerant lipase by Staphylococcus caseolyticus EX17. Our results confirm that the carbon source is a critical factor for enhancing enzyme activity. The enzyme activity was almost nil in the case of olive oil. Using glycerol and Tween 80 at different concentrations showed that lipase production was higher in a mixture of 0.5% glycerol and 0.5% Tween 80 compared with a concentration of 1%, individually or as a mixture (Fig. 3B). This result confirmed that glycerol and Tween 80 act as the carbon source and inducer, respectively. Many microorganisms prefer Tween 80 as an inducer of lipase synthesis, including Acinetobacter radioresistens [27], Serratia marcescens [28], and Acinetobacter sp. BK43 [24]. Contrary to our result, Shariff et al. [22] reported that Tween 80 inhibits lipase synthesis by Bacillus sp. L12. To observe the effect of pH on lipase production, the culture medium was prepared at various pH values. B. licheniformis
Phylogenetic tree based on the 16S rRNA gene sequence generated using the neighbor-joining method showing the relationships between the strain PAL05 and other related Bacillus species.
Biotechnology and Applied Biochemistry
531
Biotechnology and Applied Biochemistry
FIG. 4 FIG. 3
(A) Effect of carbon sources on lipase production by B. licheniformis. (B) Effect of different concentrations of select mixed carbon sources on lipase production by B. licheniformis. Each point represents the mean ± SEM of three independent experiments.
Effect of pH (A) and temperature (B) on lipase production by B. licheniformis. Each point represents the mean ± SEM of three independent experiments.
in an optimized medium was about threefold (Fig. 5) higher than that in a nonoptimized medium (Fig. 1B).
3.2. Effect of organic solvents on the stability of crude enzyme was shown to produce lipase across a wide pH range from 6.0 to 9.0, with the highest activity noted at pH 8.0 (Fig. 4A). A similar pH optimum for lipase production was reported for Acinetobacter sp. C9 and Pseudomonas spp. [29, 30]. Assessment of temperature dependence showed that the highest enzyme activity was obtained at 30 ◦ C (Fig. 4B). A low level of enzyme activity was observed even at 60 ◦ C. The optimum temperature for lipase production in B. licheniformis is similar to that noted for Pseudomonas sp. MSI057 and Acinetobacter sp. BK43 [23, 24]. In general, most microbial lipases are produced under a temperature range from 20 to 45 ◦ C [31]. The effect of salt concentration was also assessed at pH 8.0 and temperature 30 ◦ C. The highest activity was noted in the presence of 2.5% NaCl, with moderate levels of activity seen up to 7.5% NaCl (Fig. 5). At high NaCl concentrations, bacterial growth was affected (data not shown); however, the strain is considered to be moderately halotolerant. Lipase yield
532
Organic solvent-tolerant lipases are attractive for many industrial applications. Therefore, lipase stability was determined in the presence of various organic solvents (at 25% and 50%) with log P values ranging from –0.24 to 3.6. Thumar and Singh [32] reported that organic solvents with a log P value between 1.5 and 4 are extremely toxic to living organisms because the solvents preferentially partition into the cytoplasmic membrane, disorganizing its structure and impairing vital functions. Enzyme stability was high or low depending on whether the solvent was more hydrophobic or hydrophilic. Table 3 shows the effect of organic solvents on enzyme stability. The enzyme was stable in most solvents at 25% concentration; however, the enzyme activity dramatically reduced in a few solvents at 50% concentration. At 25% concentration, some solvents stimulated enzyme synthesis. Generally, organic solvents are toxic to microorganisms even at low concentrations, but some microorganisms are
Solvent-Stable Lipase from B. licheniformis
FIG. 5
Effect of different concentrations of NaCl on lipase production by B. licheniformis. Each point represents the mean ± SEM of three independent experiments.
stable at high concentrations of organic solvents as well [33]. Many microorganisms normally secrete extracellular lipases [16, 24]. However, only a few microbes such as Pseudomonas, Bacillus, and Staphylococcus can tolerate toxic solvents to grow and secrete solvent-stable lipase. Lipases are more sensitive to organic solvents; however, polar solvents are considered to be more destabilizing than nonpolar solvents [34], because polar solvents have a greater tendency to strip tightly bound water from the enzyme compared to nonpolar solvents [35]. Table 3 shows data consistent with the less disruptive nature of nonpolar solvents. The hydrophobic solvents n-hexane and benzene stimulated enzyme activity at a concentration of 25%, although at 50% concentration the enzyme activity
decreased by 30% and 34.2%, respectively. The stimulatory effect seen with n-hexane may be due to a thin layer of water molecules remaining tightly bound to the active site of the enzyme and acting as a protective sheath along with the hydrophilic surfaces of the enzyme, thereby maintaining the native conformation [34, 36]. Polar solvents such as ethanol stimulated enzyme activity by 27%; however, at 50% the enzyme activity decreased by 26.6%. It is interesting to note that at 25%, propanol and methanol retained 98% and 84.5% of the activity, respectively, and at 50%, they caused a loss of only 17.3% and 6.5% of the activity, respectively when compared with 25% concentrations. A few industrial enzymes such as lipase and cellulase are stable in both polar and nonpolar solvents at concentrations of 50% [39–42]. Solvent stabilities are based on the nature, structure, and amino acid sequence of the active site of an enzyme [43]. Normally, at the high solvent concentrations employed during ester synthesis, unwanted hydrolysis can be reduced [13, 35]. In conclusion, an organic solvent-stable lipase-producing bacterium was isolated from cattle farm soil and identified as B. licheniformis. The enzyme production process was optimized, leading to approximately threefold higher yields compared with the nonoptimized medium and related process. We established optimum conditions for lipase production: time (48 H), glycerol (0.5%), Tween 80 (0.5%), pH (8.0), and temperature (30 ◦ C). Assessing lipase stability in various organic solvents showed that the crude enzyme was stable in most organic solvents at 25%, but that enzyme activity was enhanced only in a few organic solvents. These results may make this solvent-stable
Effect of organic solvents on the stability of crude lipase
TABLE 3 Relative activity of lipase (%) Organic solvent
log P
25% concentration
50% concentration
–
100
100
−1.0
71.4 ± 1.96
53.3 ± 2.47
0.05
98.0 ± 2.55
80.7 ± 3.53
Ethanol
−0.24
127 ± 4.61
73.4 ± 4.5
Hexane
3.6
133 ± 6.4
70.0 ± 1.82
Toluene
2.5
91.4 ± 0.434
ND
Benzene
2.0
115 ± 5.76
65.8 ± 4.84
Chloroform
2.0
79.3 ± 0.73
54.4 ± 0.98
−0.76
84.5 ± 3.0
78.0 ± 6.91
Control (without solvent) DMF Propanol
Methanol
The data represent the mean ± SEM of three independent experiments. ND: Not detected.
Biotechnology and Applied Biochemistry
533
Biotechnology and Applied Biochemistry potent lipase from B. licheniformis PAL05 useful for industrial synthesis of esters and other applications.
4. Acknowledgements This work was supported by an Inha University Research Grant from Inha University, Republic of Korea.
References [1] [2] [3] [4] [5] [6] [7] [8] [9] [10] [11] [12] [13] [14] [15] [16] [17] [18] [19] [20]
534
Sikkema, J., de Bont, J., and Poolman, B. (1995) Microbiol. Rev. 59, 201–222. Inoue, A., and Horikoshi, K. (1989) Nature 338, 264–266. Sardessai, Y. N., and Bhosle, S. (2004) Biotechnol. Prog. 20, 655–660. Segura, A., Hurtado, B., Rivera, B., and Lazaroaie, M. M. (2008) J. Appl. Microbiol. 104, 1408–1416. Cadirci, B. H., and Yasa, I. (2010) J. Mol. Catal. B Enzym. 64, 155–161. Ogino, H., Miyamoto, K., Yasuda, M., Ishimi, K., and Ishikawa. (1999) Biochem. Eng. J. 4, 1–6. Singh, A. K., and Mukhopadhyay, M. (2012) Appl. Biochem. Biotechnol. 166, 486–520. Savitha, J., Srividya, S., Jagat, R., Payal, P., Priyanki, S., Rashmi, G. W., and Roshini, K. T. (2007) Afr. J. Biotechnol. 6, 564–568. Chopineau, J., McCfferty, F. D., Therisod, M., and Klibanov, M. (1988) Biotechnol. Bioeng. 31, 208–214. Jaeger, K. E., and Reetz, M. T. (1998) Trends Biotechnol. 16, 396–403. Saxena, R. K., Ghosh, P. K., Gupta, R., Davidson, W. S., Bradoo, S., and Gulati, R. (1999) Curr. Sci. 77, 101–115. Ogino, H., and Ishikawa, H. (2001) J. Biosci. Bioeng. 91, 109–116. Gupta, M. N., and Roy, I. (2004) Eur. J. Biochem. 271, 2575–2583. Pathak, A. P., and Deshmukh, K. B. (2012) Indian J. Exp. Biol. 50, 569–576. Ikram-Ul-Hak, Ashraf, H., Iqbal, J., and Qadeer, M. A. (2003) Bioresour. Technol. 87, 57–61. Khyami-Horani, H. (1996) World J. Microbiol. Biotechnol. 12, 399–401. Damiano, V. B., Ward, R., Gomes, E., Alves-Prado, H. F., and Da Silva, R. (2006) Appl. Biochem. Biotechnol. 129, 289–302. Ogino, H., Yasui, K., Shiotani, T., Ishihara, T., and Ishikawa, H. (1995) Appl. Environ. Microbiol. 61, 4258–4262. Marshall, R. T. (1992) Standard Methods for the Examination of Dairy Products, 16th ed. American Public Health Association, Washington, DC. Winkler, U. K., and Stuckmann, M. (1979) J. Bacteriol. 138, 663–670.
[21] Bora, L., and Kalita, M. C. (2007) Internet J. Microbiol. 4, 1–9. [22] Shariff, F. M., Leow, T. C., Mukred, A. D., Salleh, A. B., Basri, M., and Rahman, R. N. Z. R. A. (2007) J. Basic Microbiol. 47, 406–412. [23] Kiran, G. S., Shanmughapriya, S., Jayalakshmi, J., Selvin, J., Gandhimathi, R., Sivaramakrishnan, S., Arunkumar, M., Thangavelu, T., and Natarajaseenivasan, K. (2008) Bioprocess Biosys. Eng. 31, 483–492. [24] Anbu, P., Noh, M. J., Kim, D. H., Seo, J. S., Hur, B. K., and Min, K. H. (2011) Afr. J. Biotechnol. 10, 4147–4156. [25] Ruchi, G., Anshu, G., and Khare, S. K. (2008) Bioresour. Technol. 99, 4796–4802. [26] Volpato, G., Rodrigues, R. C., Heck, J. X., and Ayub, M. A. Z. (2008) J. Chem. Technol. Biotechnol. 83, 821–828. [27] Li, C. Y., Cheng, C. Y., and Chen, T. L. (2001) Enzym. Microb. Technol. 29, 258–263. [28] Gao, L., Xu, J. H., Li, X. J., and Liu, Z. Z. (2004) J. Ind. Microbiol. Biotechnol. 31, 525–530. [29] Kasana, R. C., Kaur, B., and Yadav, S. K. (2008) J. Basic Microbiol. 48, 207–212. [30] Tembhurkar, V. R., Kulkarni, M. B., and Peshwe, S. A. (2012) Sci. Res. Rep. 2, 46–50. [31] Gupta, R., Gupta, N., and Rathi, P. (2004) Appl. Microbiol. Biotechnol. 64, 763–781. [32] Thumar, J. T., and Singh, S. P. (2009) J. Ind. Microbiol. Biotechnol. 36, 211–218. [33] Kongpol, A., Pongtharangkul, T., Kato, J., Honda, K., Ohtake, H., and Vangnai, A. S. (2009) FEMS Microbiol. Lett. 297, 225–233. [34] Castro-Ochoa, L. D., Rodriguez-Gomez, C., Valerio-Alfaro, G., and Ros, R. O. (2005) Enzym. Microb. Technol. 37, 648–654. [35] Klibanov, A. M. (2001) Nature 409, 241–246. [36] Ahmed, E. H., Raghavendra, T., and Madamwar, D. (2010) Appl. Biochem. Biotechnol. 160, 2102–2113. [37] Fang, Y., Lu, Z., Lv, F., Bie, X., Liu, S., Ding, Z., and Xu, W. (2006) Curr. Microbiol. 53, 510–515. [38] Li, X., and Yu, H. Y. (2012) J. Ind. Microbiol. Biotechnol. 39, 1117–1124. [39] Anbu, P. (2013) Int. J. Biol. Macromol. 56, 162–168. [40] Li, X., and Yu, H. Y. (2012) FEMS Microbiol. Lett. 329, 204–211. [41] Tang, X. Y., Pan, Y., Li, S., and He, B. F. (2008) Bioresour. Technol. 99, 7388–7392. [42] Xu, J., Jiang, M., Sun, H., and He, B. (2010) Bioresour. Technol. 101, 7991–7994. [43] Kawata, T., and Ogino, H. (2010) Biochem. Biophys. Res. Commun. 400, 384–388.
Solvent-Stable Lipase from B. licheniformis
∗
Periasamy Anbu Byung Ki Hur
Department of Biological Engineering, Inha University, Incheon, South Korea
Abstract In this study, seven lipase-producing bacterial strains were isolated from salt-enriched and cattle farm soil samples after incubation in toluene- and benzene-enriched media. One strain (PAL05) showed significantly greater lipase activity on spirit blue agar medium and stability in organic solvents. The positive strain (PAL05) was identified as Bacillus licheniformis by 16S rRNA gene sequencing. Lipase production was optimized in a medium containing glycerol as the carbon source and Tween 80 as an inducer (0.5% glycerol + 0.5% Tween 80) at pH 8.0 and a temperature of 30 ◦ C. In addition, the enzyme was moderately halotolerant as it exhibited increased activity in the presence of 2.5% NaCl. Optimized
conditions increased the lipase production threefold. Crude lipase retained its activity for 14 days of incubation in the presence of various organic solvents at a level of 25% and 50%. The enzyme was stable at 25% in most solvents; some of the solvents such as hexane, benzene, and ethanol actually stimulated enzyme activity. The organic solvent stability of the lipase produced by the strain PAL05 enables the enzyme to be used as a potential biocatalyst for ester synthesis and other C 2014 International applications in nonaqueous conditions. Union of Biochemistry and Molecular Biology, Inc. Volume 61, Number 5, Pages 528–534, 2014
Keywords: organic solvent tolerance, lipase, Bacillus, optimization
1. Introduction Organic solvents are toxic to microorganisms and cause cell lysis because of disruptive effects on the cell membrane [1], as well as the internal structural and functional integrity of the cell, once it penetrates the cell [2]. However, some bacteria develop stability toward organic solvents by various adaptations such as solvent efflux pumps, rapid membrane repair, lower cell membrane permeability, increased membrane rigidity, and decreased cell surface hydrophobicity [3]. Recently, Segura et al. [4] have isolated organic solvent-stable bacteria, which are stable at low concentrations of organic solvents. Enzymes used for ester synthesis in organic solvents should be stable at high solvent concentrations to obtain high yields. Many solventstable lipases have been reported to be produced by several
Abbreviations: pNPP, 4-nitrophenyl palmitate; DMF, dimethyl formamide. ∗ Address
for correspondence: Dr. Periasamy Anbu, PhD, Department of Biological Engineering, Inha University, Incheon, 402-751, South Korea. Tel.: +82-32-860-7510; Fax: +82-32-872-4046; e-mail: [email protected]. Received 29 October 2013; accepted 27 December 2013 DOI: 10.1002/bab.1202 Published online 25 March 2014 in Wiley Online Library (wileyonlinelibrary.com)
528
Pseudomonas species [5, 6]; however, because of their high tolerance for organic solvents, Bacillus species have captured the attention of many researchers searching for a source of solvent-stable lipases and other enzymes useful for industrial applications. Lipases are triacylglycerol acylhydrolases (EC 3.1.1.3) that catalyze the hydrolysis of fats and oils to yield glycerol and free fatty acids under aqueous conditions [7]. In addition, the lipases are also involved in conversion reactions such as esterification, interesterification, transesterification, alcoholysis, acidolysis, and aminolysis [8]. Microbial lipases have many industrial and biotechnological applications as product components and manufacturing tools in areas such as synthesis of cosmetics, detergents, food, pharmaceuticals, esters, and biosurfactants [9–11]. Industrial demand for lipase stimulates the search for new enzyme sources. Two advantages of enzymatic synthesis in organic solvents or aqueous solution containing organic solvents are increased solubility of nonpolar substrates and elimination of microbial contamination in the reaction mixture [12]. Methods such as chemical modification, immobilization, entrapment, protein engineering, and directed evolution have been employed to stabilize the enzymes in organic solvents [13]. However, naturally stable enzymes with high activity in organic solvents are preferred for industrial applications. In this study,
we isolated a potent organic solvent-tolerant strain of Bacillus licheniformis and analyzed the secretion of a solvent-stable lipase. Furthermore, we optimized the conditions to maximize the lipase production and evaluated the solvent stability of lipase using various organic solvents. B. licheniformis has been reported to produce a variety of extracellular enzymes such as protease [14], amylase [15], lipase [16], and xylanase [17]. However, no reports are available on an organic solvent-stable lipase from a solvent-tolerant strain of B. licheniformis.
2. Materials and Methods 2.1. Isolation and screening of organic solvent-stable microorganisms The organic solvent-stable microorganisms were isolated according to the method of Ogino et al. [18]. Soil samples were collected from salt-enriched areas (Incheon) and cattle farms (Yongin-Si) in South Korea. About 1 g of the soil sample was suspended in 10 mL of sterile water. After shaking, 5 mL of the suspension was added to a 250 mL bottle containing 25 mL of LB broth supplemented with 2.5% toluene and 2.5% benzene (1:1 mixture). The cultivation vessels were sealed with chloroprene rubber stoppers to prevent evaporation of the organic solvents. The bottles were incubated at 37 ◦ C on a shaker at 180 rpm for 3 days. Then, 5 mL aliquots were transferred into the same medium and cultured again under the same conditions for 3 days. Aliquots of the culture suspension were diluted and spread on spirit blue agar (Tryptone, 10 g/L; yeast extract, 5 g/L; agar, 20 g/L; spirit blue, 0.15 g/L; lipid reagents containing a mixture of tributyrin and Tween 80, 30 mL/L; pH 6.8) without any organic solvent [19]. After incubation at 37 ◦ C for 36 H, the enzyme-producing strains were purified and subjected to another round of screening on spirit blue agar. Spectrophotometric analysis further confirmed the enzyme-producing activity of these strains.
2.2. Selection of a highly potent solvent-stable strain The bacteria were inoculated into 25 mL of LB medium and incubated at 30 ◦ C at 180 rpm for 4 H. About 0.5 mL of the culture suspension was transferred into 50 mL of lipase production medium composed of dipotassium hydrogen orthophosphate, 1.5 g/L; MgSO4, 0.05 g/L; CaCl2, 0.025 g/L; FeSO4, 0.015 g/L; and ZnSO4, 0.005 g/L. The inoculated flasks were incubated at 37 ◦ C at 180 rpm for 48 H. After incubation, the cultures were centrifuged at 11,100g at 4 ◦ C for 10 Min. Bacterial growth was determined at 600 nm. To obtain a highly organic solvent-stable lipase-producing strain, the strains selected by plate screening were further screened using organic solvents with various log P values. Each solvent was mixed with 1 mL of the supernatant to reach a concentration of about 20%. The tubes were covered with aluminum foil and incubated at 37 ◦ C with shaking at 100 rpm for 24 H. The residual lipase activity was measured. Based on the results of agar-based screening, spectrophotometric analysis, and organic solvent stabilities, the strain with highest activity
Biotechnology and Applied Biochemistry
and solvent stability was designated as PAL05 and was further identified by morphological, physiological, and 16S rRNA gene-sequencing methods. For 16S rRNA gene sequencing, the genomic DNA was extracted using a genomic purification kit following its manual instructions (Promega, Madison, WI, USA). The DNA was then amplified by PCR using the universal 16S rRNA gene primers: 8-27F (5 -AGAGTTTGATCCTGGCTCAG-3 ) and 1472R (5 -TACGGYTACCTTGTTACGACTT-3 ). The PCR product was electrophoresed on agarose gel electrophoresis, and the PCR product (16S rRNA band) was purified by SV gel and the PCR Clean up system (Promega). The 16S rRNA gene sequence was then compared with other sequences available in the nucleotide database by using the BLAST algorithm, and the phylogenetic tree was constructed using the neighbor-joining method (www.ncbi.nlm.nih.gov/blast).
2.3. Optimization of solvent-stable lipase production 2.3.1. Effect of incubation period on lipase production The effect of incubation time on lipase production was studied at 24 H intervals up to 72 H. Lipase activity of the cellfree supernatants obtained by centrifugation (11,100g for 10 Min at 4 ◦ C) was measured every 24 H intervals.
2.3.2. Effect of different carbon sources on lipase production Carbon sources composed of carbohydrates and lipids that were used at a concentration of 1%: glucose, lactose, glycerol, sucrose, maltose, olive oil, Tween 80, and tributyrin. Different concentrations of selected carbon sources were tested. The carbon sources were separately sterilized and then aseptically added to autoclaved medium.
2.3.3. Effect of pH, temperature, and NaCl on lipase production Lipase activity was determined at different pH values ranging from 6.0 to 10.0 and at temperatures ranging from 20 to 60 ◦ C. The pH of the medium was adjusted prior to autoclaving. The NaCl concentrations used varied from 0.25% to 7.5%.
2.4. Lipase assay Lipase activity was quantitatively assayed using 4-nitrophenyl palmitate (pNPP) as the substrate according to the method described by Winkler and Stuckmann [20]. Briefly, 10 mL of isopropanol containing 30 mg of pNPP was mixed with 90 mL of 0.05 M phosphate buffer (pH 8.0) containing 207 mg of sodium deoxycholate and 100 mg of gum arabic. A total of 2.4 mL of freshly prepared substrate solution was prewarmed to 37 ºC and mixed with 0.1 mL of enzyme solution. After incubation of this reaction mixture at 37 ºC for 15 Min, the absorbance was measured at 410 nm against an enzyme-free control. One enzyme unit was defined as the amount of enzyme that liberated 1 μmol of 4-nitrophenol/Min under the assay conditions.
529
Biotechnology and Applied Biochemistry
TABLE 1
Screening for solvent-stable lipase produced by positive strains on spirit blue agar medium
Source Salt-enriched soil
Cattle farm soil
Isolate no.
Lipase activity
PAL01
++
PAL02
+++
PAL03
++
PAL04
+
PAL05
+++
PAL06
+
PAL07
+
Symbol: +++, high activity; ++, moderate activity; +, low activity
2.5. Effect of organic solvents on the stability of crude enzyme Crude lipase was filtered through a 0.22 μm filter membrane. The organic solvents tested were benzene, chloroform, dimethyl formamide (DMF), ethanol, hexane, methanol, propanol, and toluene. Filtered crude lipase was mixed with each solvent at 25% and 50% (v/v). The mixtures were incubated at 30 ◦ C with shaking at 100 rpm for 2 weeks. After incubation, an aliquot was carefully withdrawn from the solution or from the aqueous phase in case of water-immiscible solvents. Residual lipase activity was measured. The control contained enzyme solution without any organic solvents (0% v/v). Enzyme stability was expressed as the remaining lipase activity relative to the control.
FIG. 1
Effect of incubation period on the growth (A) and lipase production (B) by positive strains. Each point represents the mean ± SEM of three independent experiments.
3. Results and Discussion The soil samples (collected from salt-enriched areas and cattle farms) were mixed with equal concentrations of tolueneand benzene-enriched medium (5% solvent). Solvent-tolerant bacteria were able to adapt to this condition. After incubation, enriched samples were spread on spirit blue agar medium. About 15 morphologically different organic solvent-tolerant bacterial colonies grew; however, only seven colonies (three from the salt-enriched soil sample and four from the cattle farm soil sample) produced a clear zone of substrate hydrolysis around the bacterial colonies. The seven positive colonies were isolated from the original spirit blue agar plate and screened again on the same medium for confirmation. Of the seven lipase producers, PAL02 and PAL05 produced the largest zones of clearance on spirit blue agar medium (Table 1). A spectrophotometric method was used to determine enzyme activity, and the highest enzyme activity was observed for PAL05; the other strains, including PAL02, showed low levels of enzyme activity. The growth rate and enzyme activity correlated for PAL02, but not for PAL05. For PAL02, growth rate and enzyme activity were maximized at 24 H. However,
530
PAL05 showed maximum growth on day 3 and maximum enzyme activity on day 2 (Fig. 1A and B). Among the seven positive strains, PAL05 was the most efficient, with the highest enzyme activity and growth rate. To choose the more efficient solvent-stable lipaseproducing strain, these two strains (PAL02 and PAL05) were further tested in various hydrophilic and hydrophobic organic solvents, including benzene, ethanol, hexane, and toluene. After mixing the supernatants with 20% (v/v) organic solvent, residual lipase activity was determined. The strain PAL05 demonstrated a wide range of stability in all solvents. PAL02 was moderately stable in a few organic solvents, but significantly less than PAL05 (Table 2). Therefore, PAL05 was selected for further study based on the spectrophotometric and solvent stability data. The selected potent strain was identified by morphological, physiological, and 16S rRNA gene sequence methods. PAL05 is a Gram-positive, aerobic, and rod-shaped bacterium. Phylogenetic analysis confirmed that the strain belongs to the Bacillus species (Fig. 2) and is highly homologous to B. licheniformis (99%). The 16S rRNA sequence (1,469 bp)
Solvent-Stable Lipase from B. licheniformis
TABLE 2
Screening for solvent stability of two potent strains in various organic solvents
Organic solvent
PAL02
PAL05
100
100
20% Ethanol
69
126
20% Hexane
96
121
20% Toluene
61
100
20% Benzene
77
93
Control (without solvent)
of this strain was deposited in GenBank (NCBI accession no. KC437094).
3.1. Organic solvent-stable lipase production Lipase production for commercial applications can be improved by optimizing the nutritional (carbon sources) and physical (pH and temperature) conditions for production. Many researchers have previously reported the optimization of lipase production from various microorganisms [21–24], but studies specific
FIG. 2
to organic solvent-stable lipases are very limited [25, 26]. Of the various carbon sources tested, a significant level of lipase production was obtained in glycerol (1%) and Tween 80 (1%) after 48 H of incubation (Fig. 3A). Low levels of enzyme activity were observed in case of other carbon sources. In contrast to this result, Volpato et al. [26] reported that glycerol acts as an inducer for the synthesis of organic solventtolerant lipase by Staphylococcus caseolyticus EX17. Our results confirm that the carbon source is a critical factor for enhancing enzyme activity. The enzyme activity was almost nil in the case of olive oil. Using glycerol and Tween 80 at different concentrations showed that lipase production was higher in a mixture of 0.5% glycerol and 0.5% Tween 80 compared with a concentration of 1%, individually or as a mixture (Fig. 3B). This result confirmed that glycerol and Tween 80 act as the carbon source and inducer, respectively. Many microorganisms prefer Tween 80 as an inducer of lipase synthesis, including Acinetobacter radioresistens [27], Serratia marcescens [28], and Acinetobacter sp. BK43 [24]. Contrary to our result, Shariff et al. [22] reported that Tween 80 inhibits lipase synthesis by Bacillus sp. L12. To observe the effect of pH on lipase production, the culture medium was prepared at various pH values. B. licheniformis
Phylogenetic tree based on the 16S rRNA gene sequence generated using the neighbor-joining method showing the relationships between the strain PAL05 and other related Bacillus species.
Biotechnology and Applied Biochemistry
531
Biotechnology and Applied Biochemistry
FIG. 4 FIG. 3
(A) Effect of carbon sources on lipase production by B. licheniformis. (B) Effect of different concentrations of select mixed carbon sources on lipase production by B. licheniformis. Each point represents the mean ± SEM of three independent experiments.
Effect of pH (A) and temperature (B) on lipase production by B. licheniformis. Each point represents the mean ± SEM of three independent experiments.
in an optimized medium was about threefold (Fig. 5) higher than that in a nonoptimized medium (Fig. 1B).
3.2. Effect of organic solvents on the stability of crude enzyme was shown to produce lipase across a wide pH range from 6.0 to 9.0, with the highest activity noted at pH 8.0 (Fig. 4A). A similar pH optimum for lipase production was reported for Acinetobacter sp. C9 and Pseudomonas spp. [29, 30]. Assessment of temperature dependence showed that the highest enzyme activity was obtained at 30 ◦ C (Fig. 4B). A low level of enzyme activity was observed even at 60 ◦ C. The optimum temperature for lipase production in B. licheniformis is similar to that noted for Pseudomonas sp. MSI057 and Acinetobacter sp. BK43 [23, 24]. In general, most microbial lipases are produced under a temperature range from 20 to 45 ◦ C [31]. The effect of salt concentration was also assessed at pH 8.0 and temperature 30 ◦ C. The highest activity was noted in the presence of 2.5% NaCl, with moderate levels of activity seen up to 7.5% NaCl (Fig. 5). At high NaCl concentrations, bacterial growth was affected (data not shown); however, the strain is considered to be moderately halotolerant. Lipase yield
532
Organic solvent-tolerant lipases are attractive for many industrial applications. Therefore, lipase stability was determined in the presence of various organic solvents (at 25% and 50%) with log P values ranging from –0.24 to 3.6. Thumar and Singh [32] reported that organic solvents with a log P value between 1.5 and 4 are extremely toxic to living organisms because the solvents preferentially partition into the cytoplasmic membrane, disorganizing its structure and impairing vital functions. Enzyme stability was high or low depending on whether the solvent was more hydrophobic or hydrophilic. Table 3 shows the effect of organic solvents on enzyme stability. The enzyme was stable in most solvents at 25% concentration; however, the enzyme activity dramatically reduced in a few solvents at 50% concentration. At 25% concentration, some solvents stimulated enzyme synthesis. Generally, organic solvents are toxic to microorganisms even at low concentrations, but some microorganisms are
Solvent-Stable Lipase from B. licheniformis
FIG. 5
Effect of different concentrations of NaCl on lipase production by B. licheniformis. Each point represents the mean ± SEM of three independent experiments.
stable at high concentrations of organic solvents as well [33]. Many microorganisms normally secrete extracellular lipases [16, 24]. However, only a few microbes such as Pseudomonas, Bacillus, and Staphylococcus can tolerate toxic solvents to grow and secrete solvent-stable lipase. Lipases are more sensitive to organic solvents; however, polar solvents are considered to be more destabilizing than nonpolar solvents [34], because polar solvents have a greater tendency to strip tightly bound water from the enzyme compared to nonpolar solvents [35]. Table 3 shows data consistent with the less disruptive nature of nonpolar solvents. The hydrophobic solvents n-hexane and benzene stimulated enzyme activity at a concentration of 25%, although at 50% concentration the enzyme activity
decreased by 30% and 34.2%, respectively. The stimulatory effect seen with n-hexane may be due to a thin layer of water molecules remaining tightly bound to the active site of the enzyme and acting as a protective sheath along with the hydrophilic surfaces of the enzyme, thereby maintaining the native conformation [34, 36]. Polar solvents such as ethanol stimulated enzyme activity by 27%; however, at 50% the enzyme activity decreased by 26.6%. It is interesting to note that at 25%, propanol and methanol retained 98% and 84.5% of the activity, respectively, and at 50%, they caused a loss of only 17.3% and 6.5% of the activity, respectively when compared with 25% concentrations. A few industrial enzymes such as lipase and cellulase are stable in both polar and nonpolar solvents at concentrations of 50% [39–42]. Solvent stabilities are based on the nature, structure, and amino acid sequence of the active site of an enzyme [43]. Normally, at the high solvent concentrations employed during ester synthesis, unwanted hydrolysis can be reduced [13, 35]. In conclusion, an organic solvent-stable lipase-producing bacterium was isolated from cattle farm soil and identified as B. licheniformis. The enzyme production process was optimized, leading to approximately threefold higher yields compared with the nonoptimized medium and related process. We established optimum conditions for lipase production: time (48 H), glycerol (0.5%), Tween 80 (0.5%), pH (8.0), and temperature (30 ◦ C). Assessing lipase stability in various organic solvents showed that the crude enzyme was stable in most organic solvents at 25%, but that enzyme activity was enhanced only in a few organic solvents. These results may make this solvent-stable
Effect of organic solvents on the stability of crude lipase
TABLE 3 Relative activity of lipase (%) Organic solvent
log P
25% concentration
50% concentration
–
100
100
−1.0
71.4 ± 1.96
53.3 ± 2.47
0.05
98.0 ± 2.55
80.7 ± 3.53
Ethanol
−0.24
127 ± 4.61
73.4 ± 4.5
Hexane
3.6
133 ± 6.4
70.0 ± 1.82
Toluene
2.5
91.4 ± 0.434
ND
Benzene
2.0
115 ± 5.76
65.8 ± 4.84
Chloroform
2.0
79.3 ± 0.73
54.4 ± 0.98
−0.76
84.5 ± 3.0
78.0 ± 6.91
Control (without solvent) DMF Propanol
Methanol
The data represent the mean ± SEM of three independent experiments. ND: Not detected.
Biotechnology and Applied Biochemistry
533
Biotechnology and Applied Biochemistry potent lipase from B. licheniformis PAL05 useful for industrial synthesis of esters and other applications.
4. Acknowledgements This work was supported by an Inha University Research Grant from Inha University, Republic of Korea.
References [1] [2] [3] [4] [5] [6] [7] [8] [9] [10] [11] [12] [13] [14] [15] [16] [17] [18] [19] [20]
534
Sikkema, J., de Bont, J., and Poolman, B. (1995) Microbiol. Rev. 59, 201–222. Inoue, A., and Horikoshi, K. (1989) Nature 338, 264–266. Sardessai, Y. N., and Bhosle, S. (2004) Biotechnol. Prog. 20, 655–660. Segura, A., Hurtado, B., Rivera, B., and Lazaroaie, M. M. (2008) J. Appl. Microbiol. 104, 1408–1416. Cadirci, B. H., and Yasa, I. (2010) J. Mol. Catal. B Enzym. 64, 155–161. Ogino, H., Miyamoto, K., Yasuda, M., Ishimi, K., and Ishikawa. (1999) Biochem. Eng. J. 4, 1–6. Singh, A. K., and Mukhopadhyay, M. (2012) Appl. Biochem. Biotechnol. 166, 486–520. Savitha, J., Srividya, S., Jagat, R., Payal, P., Priyanki, S., Rashmi, G. W., and Roshini, K. T. (2007) Afr. J. Biotechnol. 6, 564–568. Chopineau, J., McCfferty, F. D., Therisod, M., and Klibanov, M. (1988) Biotechnol. Bioeng. 31, 208–214. Jaeger, K. E., and Reetz, M. T. (1998) Trends Biotechnol. 16, 396–403. Saxena, R. K., Ghosh, P. K., Gupta, R., Davidson, W. S., Bradoo, S., and Gulati, R. (1999) Curr. Sci. 77, 101–115. Ogino, H., and Ishikawa, H. (2001) J. Biosci. Bioeng. 91, 109–116. Gupta, M. N., and Roy, I. (2004) Eur. J. Biochem. 271, 2575–2583. Pathak, A. P., and Deshmukh, K. B. (2012) Indian J. Exp. Biol. 50, 569–576. Ikram-Ul-Hak, Ashraf, H., Iqbal, J., and Qadeer, M. A. (2003) Bioresour. Technol. 87, 57–61. Khyami-Horani, H. (1996) World J. Microbiol. Biotechnol. 12, 399–401. Damiano, V. B., Ward, R., Gomes, E., Alves-Prado, H. F., and Da Silva, R. (2006) Appl. Biochem. Biotechnol. 129, 289–302. Ogino, H., Yasui, K., Shiotani, T., Ishihara, T., and Ishikawa, H. (1995) Appl. Environ. Microbiol. 61, 4258–4262. Marshall, R. T. (1992) Standard Methods for the Examination of Dairy Products, 16th ed. American Public Health Association, Washington, DC. Winkler, U. K., and Stuckmann, M. (1979) J. Bacteriol. 138, 663–670.
[21] Bora, L., and Kalita, M. C. (2007) Internet J. Microbiol. 4, 1–9. [22] Shariff, F. M., Leow, T. C., Mukred, A. D., Salleh, A. B., Basri, M., and Rahman, R. N. Z. R. A. (2007) J. Basic Microbiol. 47, 406–412. [23] Kiran, G. S., Shanmughapriya, S., Jayalakshmi, J., Selvin, J., Gandhimathi, R., Sivaramakrishnan, S., Arunkumar, M., Thangavelu, T., and Natarajaseenivasan, K. (2008) Bioprocess Biosys. Eng. 31, 483–492. [24] Anbu, P., Noh, M. J., Kim, D. H., Seo, J. S., Hur, B. K., and Min, K. H. (2011) Afr. J. Biotechnol. 10, 4147–4156. [25] Ruchi, G., Anshu, G., and Khare, S. K. (2008) Bioresour. Technol. 99, 4796–4802. [26] Volpato, G., Rodrigues, R. C., Heck, J. X., and Ayub, M. A. Z. (2008) J. Chem. Technol. Biotechnol. 83, 821–828. [27] Li, C. Y., Cheng, C. Y., and Chen, T. L. (2001) Enzym. Microb. Technol. 29, 258–263. [28] Gao, L., Xu, J. H., Li, X. J., and Liu, Z. Z. (2004) J. Ind. Microbiol. Biotechnol. 31, 525–530. [29] Kasana, R. C., Kaur, B., and Yadav, S. K. (2008) J. Basic Microbiol. 48, 207–212. [30] Tembhurkar, V. R., Kulkarni, M. B., and Peshwe, S. A. (2012) Sci. Res. Rep. 2, 46–50. [31] Gupta, R., Gupta, N., and Rathi, P. (2004) Appl. Microbiol. Biotechnol. 64, 763–781. [32] Thumar, J. T., and Singh, S. P. (2009) J. Ind. Microbiol. Biotechnol. 36, 211–218. [33] Kongpol, A., Pongtharangkul, T., Kato, J., Honda, K., Ohtake, H., and Vangnai, A. S. (2009) FEMS Microbiol. Lett. 297, 225–233. [34] Castro-Ochoa, L. D., Rodriguez-Gomez, C., Valerio-Alfaro, G., and Ros, R. O. (2005) Enzym. Microb. Technol. 37, 648–654. [35] Klibanov, A. M. (2001) Nature 409, 241–246. [36] Ahmed, E. H., Raghavendra, T., and Madamwar, D. (2010) Appl. Biochem. Biotechnol. 160, 2102–2113. [37] Fang, Y., Lu, Z., Lv, F., Bie, X., Liu, S., Ding, Z., and Xu, W. (2006) Curr. Microbiol. 53, 510–515. [38] Li, X., and Yu, H. Y. (2012) J. Ind. Microbiol. Biotechnol. 39, 1117–1124. [39] Anbu, P. (2013) Int. J. Biol. Macromol. 56, 162–168. [40] Li, X., and Yu, H. Y. (2012) FEMS Microbiol. Lett. 329, 204–211. [41] Tang, X. Y., Pan, Y., Li, S., and He, B. F. (2008) Bioresour. Technol. 99, 7388–7392. [42] Xu, J., Jiang, M., Sun, H., and He, B. (2010) Bioresour. Technol. 101, 7991–7994. [43] Kawata, T., and Ogino, H. (2010) Biochem. Biophys. Res. Commun. 400, 384–388.
Solvent-Stable Lipase from B. licheniformis
