Article pubs.acs.org/IECR
Development of an Industrial Microbial System for Chitinolytic Enzymes Production F. Moscoso, L. Ferreira, M.A. Fernández de Dios, F.J. Deive, M.A. Longo, and M.A. Sanromán* Department of Chemical Engineering, University of Vigo, Isaac Newton Building, Campus As Lagoas, Marcosende 36310 Vigo, Spain ABSTRACT: Two endochitinases from Lactococcus lactis and Bacillus halodurans were expressed in E. coli, and the protein production process was performed at a bioreactor scale. The effect of engineering variables such as aeration and agitation on the performance of the biological process was determined; maximum levels of activity were achieved at an aeration rate of 0.33 vvm and an agitation rate of 300 rpm when using isopropyl-β-D-thiogalactopyranoside (IPTG) as an inducer. The use of lactose as an inducer in cultures of L. lactis generated maximum levels of chitinolytic activity (2.25 U/L) in just 3 h of cultivation, a level much higher than for other reported strains. The production process was kinetically analyzed using unstructured models, and the Luedeking and Piret equation was used to characterize the enzyme produced.
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INTRODUCTION Chitin, a β-(1,4)-linked polymer of N-acetyl D-glucosamine, is the second most abundant biopolymer on Earth and is present in the exoskeleton of arthropods, crustacean shells, and fungal cell walls in concentrations of up to 44%.1,2 It is a highly insoluble material with low chemical reactivity. Over the past few years, chitin and chitin derivatives have been used for various applications, such as cosmetics, food, nutrition, and biotechnology, due to the practical properties of this natural polymer, including biocompatibility and biodegradability.3 Various organisms that are capable of using chitin as a carbon and nitrogen source have been identified as producers of chitinolytic enzymes. These biocatalysts are hydrolases that are widely appreciated for their potential application as fungicides and plague control agents as well as for the treatment of chitinous waste or the preparation of chitooligosaccharides (chitohexaose and chitoheptaose) and N-acetyl D-glucosamine.4−9 Recently, chitinolytic enzymes were reclassified by the Enzyme Commission into two types: endochitinases (EC 3.2.1.14), which catalyze the cleavage of chitin at an internal site into chitotetraose, chitotriose, and chitobiose; and exochitinases or chitobiosidases and chitobiases (EC 3.2.1.52), which split chitobiose in a stepwise way without the formation of mono- or oligosaccharides. In addition, chitobiases break chitobiose, chitotriose, and chitotetraose into N-acetyl glucosamine (G1cNac) monomers in an exotype fashion.5 Bacterial chitinases have been identified in five genera of bacteria: Pseudomonas,10 Serratia,11 Streptomyces,11 Lactococcus12, and Bacillus.7,8,13,14 However, identifying novel organisms with chitinolytic activity is still an area of active research.4,5,13 The expression of chitinolytic enzymes in mesophilic hosts could help develop cost-effective production processes. However, the number of developed systems is very limited because not all chitinases can be expressed efficiently in E. coli (Billman-Jacobe, 1996).15 There is a scarcity of information concerning the industrial production of chitinases and only two bacteria have been used to date (Serratia marcescens and Streptomyces griseus).11 One of the production bottlenecks for these enzymes at an industrial scale is a lack of knowledge concerning bioreactor operation.16 Among the critical engineer© 2013 American Chemical Society
ing variables governing this biological reaction, aeration and agitation play a key role in the scale-up and understanding of the operation prior to production at an industrial scale.16,17 In this study, the production of chitinolytic enzymes by the E. coli expression of genes from Bacillus and Lactococcus strains was investigated. The efficiency of chitinase production was assessed using a logistic-type model, and the biological process was operated at the bioreactor scale. In addition, the operating conditions and the effect of two different inducers (isopropyl-βD-thiogalactopyranoside (IPTG) and lactose) were evaluated.
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EXPERIMENTAL SECTION Chemicals and Culture Medium. Luria−Bertani culture medium (10 g/L bacteriological peptone, 5 g/L yeast extract, and 5 g/L NaCl) was used for liquid culture in these chitinase production experiments and also for solid plates, together with 20 g/L of agar. The antibiotic ampicillin was supplied by Fisher Bioreagents and added at a final concentration of 100 μg/mL. A stock solution of 50 mg/mL was prepared and stored at −20 °C. Fifty millimolar potassium phosphate buffer containing 25 mM EDTA and 25 mM NaCl at pH 8 for Bacillus halodurans or pH 7.5 for Lactococcus lactis was used to resuspend cell pellets. Isopropyl-β-D-thiogalactopyranoside (IPTG) from SigmaAldrich was added at a final concentration of 0.5 mM. Lactose was purchased from Merck and prepared at a final concentration of 15 mM. Both chemicals were used as inducers. 4-Nitrophenyl-N-acetyl-β-D-glucosaminide (Bioline) was used as a specific substrate in the enzymatic activity assay. A stock solution of 5 mM was prepared and stored at −20 °C. All other solvents and chemicals used were at least reagent grade. Microorganisms. The protein was produced using E. coli BL21 as the host strain. The recombinant strains expressed a Received: Revised: Accepted: Published: 10046
March 2, 2013 June 12, 2013 July 4, 2013 July 4, 2013 dx.doi.org/10.1021/ie400687n | Ind. Eng. Chem. Res. 2013, 52, 10046−10051
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Table 1. Summary of Chitinase Production Procedure step
description
1 2 3
growth of recombinant strain on a solid agar plate growth in liquid medium at tube scale growth in liquid medium at flask scale
4 5 6 7
growth and induction for 5−7 h harvest induced cells resuspension in buffer and sonication purification and enzymatic assay
purpose
aditional information
overexpresion in E. coli host
briefly described in material and methods
to reach proper optical density used as inoculum in the scaleup chitinase activity production remove supernatant lysis of cells determination of chitinase activity
3 mL of LB medium with 100 μg/mL ampicillin 50 mL of LB medium with 100 μg/mL ampicillin IPTG or lactose used as inducers when D.O. is between 1.0 and 1.2 nm centrifuge, 5900g 4 °C for 10 min sonicated in 10 cycles of 30 pulses for 30 s in ice bath. after purifing by means of dialysis, the enzymatic reaction takes place for 1 h at 37 °C
Sodium Dodecyl Sulfate Polyacrylamide Gel Electrophoresis (SDS-PAGE). SDS−PAGE was performed with a 10% SDS polyacrylamide separating gel on a vertical slab mini gel (Pharmacia Biotech) as described by Laemmli.19 Following migration, gels were stained for protein detection with Coomassie Blue R-250 following standard procedures. The molecular mass of the enzyme was determined on a scale calibrated with a standard protein mixture that contained myosin, 200 kDa; β-galactosidase, 116.25 kDa; phosphorylase b, 97.4 kDa; BSA, 66.2 kDa; ovalbumin, 45 kDa; carbonic anhydrase, 31 kDa; trypsin inhibitor, 21.5 kDa; lysozyme, 14.4 kDa and aprotinin, 6.5 kDa.
Bacillus chitinase gene cloned into the pGEX and pET100 vectors, while the Lactococcus chiA1 gene was only cloned into the pGEX vector. Expression was controlled using either the lac (pGEX) or T7 (pET100) promoters. Chitinase Production at Bioreactor Scale. A 5-L stirredtank bioreactor (Biostat B, Braun, Germany) was used. It was filled with 3 L of medium containing ampicillin at a final concentration of 100 μg/mL. The bioreactor was inoculated at 3% v/v with actively growing cells in the late exponential phase (12 h) and incubated at 37 °C on a rotatory shaker at 200 rpm. When the optical density at 600 nm reached 1−1.2 units after approximately two hours, enzyme synthesis was induced with IPTG (0.5 mM) or lactose (15 mM), and growth was continued for 5 h. Samples (50 mL) were withdrawn at regular intervals to monitor expression. The cells were harvested by centrifugation at 5900g for 10 min at 4 °C and resuspended in potassium phosphate buffer (pH 7.5 or 8.0). The pH and extent of cell growth were monitored throughout. Random fermentations were repeated to evaluate the reproducibility of the scale-up. All samples were analyzed in triplicate. The values in the figures and tables correspond to mean values with a standard deviation (S.D.) lower than 15%. Analytical Methods. Sample Preparation. Cells were harvested by centrifugation (10 min, 5900g) and suspended in 3 mL of potassium phosphate buffer (pH 7.5 or 8.0 depending on the selected strain). The cell suspension was sonicated in 10 cycles of 30 pulses of 30 s each at 50% maximum power (Branson Sonifier, model 250) for the extraction of crude enzyme. Sonication was performed in an ice bath, and a 1 min cooling interval was allowed between cycles. Then, the lysed cells were centrifuged for 10 min at 4 °C and 5900g to remove insoluble cell debris and dialyzed for 24 h in buffer. The supernatant was retained for the measurement of chitinase activity. More details about sample preparation are provided in Table 1. Cell Growth Determination. Biomass concentration was measured by turbidimetry at 600 nm and the obtained values were converted to g cell dry wt/L using a previously determined calibration curve. Chitinolytic Enzyme Assay. Chitinase activity in the expressed recombinant endochitinase was determined spectrophotometrically by colorimetric reaction with 4-nitrophenylN-acetyl-β-D-glucosaminide (0.5 mM) at 37 °C for 1 h. The absorption of liberated p-nitrophenol was measured at 410 nm. One unit of activity was defined as the amount of enzyme required to release 1 μmol of p-nitrophenol per minute under the described conditions.18
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RESULTS AND DISCUSSION The efficient industrial production of chitinases requires the investigation of novel strains able to efficiently synthesize these enzymes at a bioreactor scale. Different species in the genus Bacillus, including B. atrophaeus, B. licheniformis, B. thuringiensis, and B. cereus, are well documented in the literature to be chitinase producers.2,8,9,20 Therefore, one strain from this genus was selected as a model microorganism, and one recombinant chitinase each from B. halodurans and L. lactis were expressed in E. coli in a stirred tank bioreactor (STB). Various critical variables affecting the performance of the expression process, such as the composition of the culture medium, the agitation, and the aeration, were screened to identify optimal expression conditions. Effects of Aeration and Agitation. First, the effect of the aeration was checked at different flow rates between 0.2 and 1 vvm. We observed that aeration levels of 0.33 and 0.66 vvm created a stable system without significant problems such as foaming. Both bacterial growth and the chitinase activity were monitored, and the results are shown in Figure 1. Maximum biomass values were reached after less than 9 h of cultivation under both levels of aeration (3.11 g/L and 3.85 g/L for L. lactis and 2.54 and 4.42 for B. halodurans at low and high aeration rates, respectively). The analysis of chitinase activity data revealed a 47% reduction in cultures of L. lactis, and a 58% reduction for B. halodurans at a flow rate of 0.66 vvm relative to 0.33 vvm. Therefore, a flow rate 0.33 vvm was selected for further studies. This aeration level was lower than levels proposed by other authors. While Fleuriet et al.21 found that 1.5 vvm promoted the highest levels of chitinase production by Cellulosimicrobium cellulans, Felse and Panda22 concluded that the optimum aeration rate was 0.75 vvm. These authors employed a culture medium containing chitin, which may create rheology issues. The effect of chitin on the fluid dynamics of their growth media may have motivated their selection of high levels of aeration. Our results were more 10047
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activity values, μm is the maximum specific growth rate, and μA is the maximum specific activity rate. X max
X=
((
1 + exp ln
X max X0
)
)
(1)
)
)
(2)
− 1 − μm t
A max
A=
((
1 + exp ln
A max A0
− 1 − μA t
The experimental data were adjusted to the models using an iterative procedure (applying the SOLVER function in Microsoft EXCEL), which seeks the parameter values that minimize the sum of the squared differences between the observed and predicted values of the dependent variable. The model parameter values are listed in Tables 2 and 3. The most suitable model for cell growth and enzyme activity was consistent with the experimental results from the exponential phase to the stationary phase (correlation coefficients > 0.91), demonstrating that this model was suitable for describing both variables under the given conditions. Taking these data into account, a preliminary experiment to characterize chitinase production was performed. The application of a suitable kinetic model of enzymatic production and microbial growth could help quantify the effect of the agitation and aeration. We applied a Luedeking and Piret type model, such as the integrated form reported by Marques et al.,25 for the first time to this type of enzyme. ⎧ ⎫ ⎪ ⎪ e μt A = A 0 + mX 0⎨ − 1⎬ X 0 ⎪ ⎪1 − (1 − e μt ) X max ⎭ ⎩ ⎤ ⎛ X ⎞ ⎛X ⎞ ⎡ + n⎜ max ⎟ ln⎢1 − ⎜ 0 ⎟(1 − e μt )⎥ ⎥⎦ ⎝ μ ⎠ ⎢⎣ ⎝ X max ⎠
Figure 1. (a) Biomass concentration and (b) chitinase activity profiles of L. lactis (filled symbols) and B. halodurans (open symbols) at different levels of aeration: (●) 0.33 vvm and (■) 0.66 vvm. Experimental data are represented by symbols and the logistic model by solid lines.
( )
(3)
This equation allows the determination of a range of chitinase production, including entirely growth-associated product (m ≠ 0 and n = 0), cases in which enzyme synthesis is partially growth-associated (m and n ≠ 0), and cases in which product formation is an alternative to growth (m = 0 and n ≠ 0). The values obtained from this equation are presented in Tables 2 and 3. Data analysis for both bacteria revealed that operation at lower aeration rates dramatically modified the kinetic behavior because the parameter m at low aeration rates was 60 and 40 times higher than parameter n for L. lactis and B. halodurans, respectively. In fact, the operation at higher aeration rates reduced the difference between both parameters (m was 3
economical because low levels of aeration require lower costs of compression. The microbial process was then modeled to elucidate the microbial response to the selected environmental conditions. This theoretical study provided a useful tool for the control of the biological process and characterization of bacterial behavior.23,24 The performance of the biological process, shown in Figure 1, was adequately modeled by fitting the data to the logistic eqs 1 and 2, in which X and A are the biomass and the chitinase activity at any specific moment (t), respectively, X0 and Xmax are the initial and maximum biomass concentrations, A0 and Amax are the initial and maximum
Table 2. Growth and Chitinase Activity Kinetic Parameters Defining the Logistic and Luedeking and Piret Type Models in Cultures of L. lactis at Different Aeration Flow Rates and Agitation Rates in a Stirred Tank Bioreactor engineering variables bacteria
agitation (rpm)
aeration (vvm)
L. lactis
200
B. halodurans
300 200
0.66 0.33 0.33 0.66 0.33 0.33
300
biomass parameters
chitinase activity
Xmax (g/L)
μm (h−1)
R2
± ± ± ± ± ±
0.83 0.72 1.56 0.88 1.44 1.59
0.97 0.96 0.99 0.99 0.98 0.91
3.85 3.11 3.97 4.42 2.54 3.81
0.01 0.03 0.01 0.04 0.01 0.02
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Luedeking and Piret parameters
Amax (U/L)
μA (h−1)
R2
m (U/g)
n (U/g/h)
R2
± ± ± ± ± ±
1.61 1.45 3.92 2.56 2.06 1.94
0.97 0.99 0.99 0.97 0.96 0.95
1.23 1.23 0.15 0.38 0.12 0.19
0.5 0.07 0.01 0.05 0.00 0.00
0.98 0.99 0.92 0.97 0.98 0.99
0.69 1.51 0.57 0.36 0.69 0.70
0.04 0.03 0.06 0.01 0.01 0.01
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Table 3. Growth and Chitinase Activity Kinetic Parameters Defining the Logistic and Luedeking and Piret Type Models in Cultures of L. lactis at 300 rpm and 0.33 vvm in Different Culture Media culture medium
biomass parameters
chitinase activity
peptone
inducer
Xmax (g/L)
μm (h−1)
R2
bacteriological bacteriological casein casein
IPTG Lactose IPTG Lactose
3.97 4.11 4.51 4.57
± ± ± ±
1.56 1.81 1.79 1.01
0.99 0.99 0.97 0.97
0.01 0.01 0.02 0.01
Luedeking and Piret parameters
Amax(U/L)
μA (h−1)
R2
m (U/g)
n (U/g/h)
R2
± ± ± ±
3.92 3.72 3.28 2.32
0.99 0.99 0.99 0.98
0.15 0.33 0.15 0.19
0.01 0.11 0.03 0.05
0.92 0.98 0.95 0.93
0.57 2.25 0.72 0.30
times higher than n). Therefore, the biological process was more dependent on biomass production in each case, but the relationship between m and n depended strongly on the hydrodynamic setting inside the bioreactor. In agreement with our findings, Bankar and Singhal26 have recently demonstrated the important effects that the hydrodynamic conditions have on the kinetic behavior of poly-ε-lysine biosynthesis by Streptomyces noursei NRRL 5126. They concluded a change occurred in the ratio between the parameters m and n when the scale was changed from a shaken flask to a stirred tank bioreactor (from 3.18 to 23.66), demonstrating the greater influence of the growth-associated parameter. Once the suitable aeration level was determined, different agitation rates were analyzed in a range between 100 and 400 rpm in search of a compromise between the minimum agitation rate that ensures sufficient mass transfer and the highest agitation rate that prevents excessive foaming.27,28 The influence of this variable on the kinetic behavior of enzyme biosynthesis was ascertained, and the data obtained for both bacterial strains are presented in Figures 2 and 3 and also in Table 2. From this data, two different trends were observed. (i) High stirring rates did not create significant variations in either the biomass or enzyme activity of recombinant E. coli cultures carrying the chitinase synthesis genes from B. halodurans. (ii) The cultivation of mutant E. coli containing the L. lactis gene attained similar levels of enzyme activity at high agitation rates after just 3 h of IPTG induction. This improvement was mirrored in an increase in the specific growth rate and a 2-fold increase in specific production. The difference between the responses of both strains confirmed the importance of developing ad hoc strategies for each microbiological process. Data from the literature indicates that the time required to reach maximum levels of chitinase production by the fungus Trichoderma harzianum in a stirred tank bioreactor is much higher than was observed in the present study (112 h vs 3 h).22 Analysis of the Luedeking and Piret parameters confirms the different trends between microorganisms. While no important changes were detected in the kinetic behavior of chitinases production by B. halodurans, the rate of m:n for L. lactis increased up to 2 orders of magnitude, similar to behavior obtained when aeration was decreased. Effect of Medium Composition. Once the viable operating conditions were determined (0.33 vvm and 300 rpm), we then evaluated different media additions. More specifically, we analyze the concomitant effect of two chitinase inducers (lactose and IPTG) and two peptones (bacteriological and casein peptone) as nitrogen sources. Biomass and chitinase biosynthesis data were monitored for 24 h to ensure that the stationary phase was reached in all cases (Figure 4). The logistic models presented previously were applied, and the values of the resulting parameters are listed in Table 3. The experimental data demonstrated a positive effect of lactose (15 mM) in the cultures of L. lactis, while no
0.06 0.02 0.01 0.01
Figure 2. (a) Biomass concentration and (b) chitinase activity profiles of L. lactis at different level of agitation: (●) 200 rpm and (▲) 300 rpm. Experimental data are represented by symbols and the logistic model by solid lines.
significant differences were detected when B. halodurans was used (data not shown). More specifically, the concomitant use of bacteriological peptone and lactose produced a significant increase in the levels of enzyme activity up to 2.25 U/L, an increase of approximately 4-fold. The effectiveness of lactose in this particular case may have been because L. lactis species are one of the most important groups of lactic acid bacteria that transform lactose in lactic acid. Lien et al.29 have also demonstrated the beneficial effect of different mono and disaccharides for maximizing biomass and chitinolytic activity in cultures of Aeromonas spp. The use of lactose was also found to be beneficial by Kopparapu et al.30 in cultures of Paecilomyces thermophile. It is notable that the introduction of casein peptone in the culture medium not only does improve in activity but also causes lower levels of enzyme biosynthesis. One difference between casein and bacteriological peptone is the total amount of nitrogen, which can vary between 12 and 10049
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Figure 3. (a) Biomass concentration and (b) chitinase activity profiles of B. halodurans at different levels of agitation; (○) 200 rpm and ( × ) 300 rpm. Experimental data are represented by symbols and the logistic model by solid lines.
Figure 4. (a) Biomass concentration and (b) chitinase activity profiles of L. lactis in different culture medium; (▲) IPTG + bacteriological peptone, (△) lactose + casein peptone, (◆) lactose + bacteriological peptone and (◇) IPTG + casein peptone. Experimental data are represented by symbols and the logistic model by solid lines.
17%. Therefore, the productive interaction between the inducer and the nitrogen source is notable. The kinetic characterization using the Luedeking and Piret equation revealed that the use of casein peptone and lactose created a partial dependence on the biomass-associated parameter, demonstrating a higher dependence on the growth parameter m in all of the media used. The production of recombinant chitinase from L. lactis presents an economic benefit because both the specific growth rate and the specific production rate are much higher (approximately 2 and 4 times, respectively) than those reported previously by Deive et al.31 for the production of bacterial lipolytic enzymes in a batch bioreactor. Regarding the time required to reach maximum expression, comparisons with other reported strains are even more promising. Thus, Fleuri et al. and Felse and Panda21,22 needed 112−168 h to reach lower levels of chitinase activity, while a recent work published by Fenice et al.16 achieved optimum values after 72 h of cultivation. Finally, complementary SDS-PAGE carried out at different culture times revealed a band with varied intensity for both strains, as shown in Figure 5. Gel comparisons with standard proteins revealed a molecular mass of approximately 65 kDa, which corresponds with the typical mass of bacterial chitinases (approximately 20−65 kDa) reported by Bhattacharya et al.4
Figure 5. SDS−PAGE analysis of chitinase produced by L. lactis (lanes 1−3) and B. halodurans (lanes 4−6). Lane P: molecular markers.
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CONCLUSIONS In the present investigation, we concluded that the production of chitinases from a recombinant E. coli carrying chitinase synthesis genes from L. lactis is a promising alternative to the existing processes. The process was successfully performed in a 10050
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(14) Fenice, M.; Barghini, P.; Selbmann, L.; Federici, F. Combined effects of agitation and aeration on the chitinolytic enzymes production by the Antarctic fungus Lecanicillium muscarium CCFEE 5003. Microb. Cell Fact. 2012, 11, 1−10. (15) Da Silva, A. F.; García-Fraga, B.; López-Seijas, J. ; Sieiro, C. Optimization of the expression of a chitinase gene from Lactococcus lactis. MicroBiotec11, Dec 1−3, 2011, Braga, Portugal. (16) Da Silva, A. F.; Gracía-Fraga, B.; Sieiro, C. Cloning and characterization of a gene encoding a family 18 chitinase from an alkaliphilic Bacillus. 5th Congress of European Microbiologists (FEMS) July 21−25, 2013, Leipzig, Germany. (17) Wang, S. Y.; Moyne, A. L.; Thottappilly, G.; Wu, S. J.; Locy, R. D.; Singh, N. K. Purification and characterization of a Bacillus cereus exochitinase. Enzyme Microb. Technol. 2001, 28, 492−49. (18) Laemmli, U. K. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 1970, 227, 680− 685. (19) Lee, Y. S.; Park, I. H.; Yoo, J. S.; Chung, S. Y.; Lee, Y. C.; Cho, Y. S.; Ahn, S. C.; Kim, C. M.; Choi, Y. L. Cloning, purification, and characterization of chitinase from Bacillus sp. DAU101. Bioresour. Technol. 2007, 98, 2734−2741. (20) Fleuri, L. F.; Kawaguti, H. Y.; Sato, H. H. Production, purification and application of extracellular chitinase from Cellulosimicrobium cellulans 191. Braz. J. Microbiol. 2009, 40, 623−630. (21) Felse, P. A.; Panda, T. Submerged culture production of chitinase by Trichoderma harzianum in stirred tank bioreactorsThe influence of agitator speed. Biochem. Eng. J. 2000, 4, 115−120. (22) Deive, F. J.; Á lvarez, M. S.; Morán, P.; Sanromán, M. A.; Longo, M. A. A process for extracelular termostable lipase production by a novel Bacillus thermoamylovorans strain. Bioprocess Biosyst. Eng. 2012, 35, 931−941. (23) Moscoso, F.; Deive, F. J.; Longo, M. A.; Sanromán, M. A. Technoeconomic assessment of phenanthrene degradation by Pseudomonas stutzeri CECT 930 in a batch bioreactor. Bioresour. Technol. 2012, 104, 81−89. (24) Marqués, A. M.; Estañol, I.; Alsina, J. M.; Fusté, C.; SimonPujol, D.; Guinea, J.; Congregado, F. Production and rheological properties of the extracelular polysaccharide synthesized by Pseudomonas sp. strain EPS-5028. Appl. Environ. Microbiol. 1986, 52, 1221−1223. (25) Bankar, S. B.; Singhal, R. S. Fermentation kinetics makeover in poly-e-lysine biosynthesis by Streptomyces noursei NRRL 5126. J. Biochem. Tech. 2012, 3, 1−5. (26) Moscoso, F.; Teijiz, I.; Deive, F. J.; Sanromán, M. A. Efficient PAHs biodegradation by a bacterial consortium at flask and bioreactor scale. Bioresour. Technol. 2012, 119, 270−276. (27) Domínguez, A.; Pastrana, L.; Rúa, M. A.; Longo, M. A.; Sanromán , M. A. Lipolytic enzyme production by Thermus thermophilus HB27 in a stirred tank bioreactor. Biochem. Eng. J. 2005, 26, 95−99. (28) Lien, T. S.; Yu, S. T.; Wu, S. T.; Too, J. R. Induction and purification of a thermophilic chitinase produced by Aeromonas sp. DYU-Too7 using glucosamine. Biotechnol. Bioprocess Eng. 2007, 12, 610−617. (29) Kopparapu, N. K.; Zhou, P.; Zhang, S.; Yam, Q.; Liu, Z.; Jiang, Z. Purification and characterization of a novel chitinase gene from Paecilomyces thermophile expressed in Escherichia coli. Carbohydr. Res. 2012, 347, 155−160. (30) Deive, F. J.; Carvalho, E.; Pastrana, L.; Rúa, M. L.; Longo, M. A.; Sanromán, M. A. Assessment of relevant factors influencing lipolytic enzyme production by Thermus thermophilus HB27 in laboratory-scale bioreactors. Chem. Eng. Technol. 2009, 32, 606−612. (31) Deive, F. J.; Carvalho, E.; Pastrana, L.; Rúa, M. L.; Longo, M. A.; Sanromán, M. A. Strategies for improving extracellular lipolytic enzyme production by Thermus thermophilus HB27. Bioresour. Technol. 2009, 100, 3630−3637.
bench scale stirred tank bioreactor. Aeration at 0.33 vvm and agitation at 300 rpm were optimum settings for producing high levels of enzyme expression. Replacing the more commonly used IPTG with lactose as the inducer dramatically improved yields (more than 2 U/L) in just 3 h of culture, a major advantage for bioprocess economy. The suitability of the Logistic and Luedeking and Piret models was revealed in very high regression coefficients, confirming that the chitinase product was partially a growth-associated metabolite with a stronger dependence on the biomass.
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AUTHOR INFORMATION
Corresponding Author
*Tel.: +34 986 812383. Fax: +34 986 812380. E-mail:
[email protected]. Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS This work was financed by Xunta de Galicia, Spain, project PXIB310278PR. F.J. Deive acknowleges Xunta de Galicia for funding through Isidro Parga Pondal program
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REFERENCES
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dx.doi.org/10.1021/ie400687n | Ind. Eng. Chem. Res. 2013, 52, 10046−10051
