Table of Contents
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PREFACE
Introduction
Characteristics of a New Fungus Isolated for Dye Decolorization
Abstract
1. ISOLATION OF A NEW DECOLORIZING FUNGUS
1.1. Methods
1.1.1. Isolation of Dye-decolorizing Microorganisms
1.1.2. Structures of the Dyes Used
1.1.3. Decolorization Using the Purified Dec 1 on PDA Medium
1.1.4. Decolorization with the Purified Dec 1 in Liquid Medium
1.1.5. Extracellular or Intracellular Crude Enzyme Solutions Prepared from Dec 1
1.2. Results
1.2.1. Decolorization by the Isolated Fungus, Dec 1, on Solid Medium
1.2.2. Decolorization by the Isolated Dec 1 in Liquid Medium
1.2.3. Decolorization by Crude Enzyme Solution
1.3. Discussion
CONCLUSION
Characterization of Multi-Enzymes Produced by the Fungus, Dec 1, Responsible for Dye Decolorization
Abstract
1. NEW PEROXIDASE, DYP
1.1. DyP Purification
1.2. Spectral Characteristics of DyP
1.3. Substrate Specificity of DyP
1.4. Sugar Analysis
1.5. Effect of Temperature on DyP Activity
1.6. H2O2 Inhibition of DyP Activity
1.7. Comparison of Decolorization Rates of DyP and HRP
1.8. DyP Activity Analyzed by Second-order Kinetics
1.9. DyP Production
1.10. Effect of Molasses on DyP Production
1.10.1. Methods
1.10.1.1. Cultivation of Dec 1
1.10.1.2. Determination of Decolorization Degree of Culture Broth
1.10.1.3. Decolorization Activity of DyP Toward RB5 Dye
1.10.1.4. DyP Purification
1.10.1.5. Effect of Molasses Concentration on Decolorization Rate of RB5 by DyP
1.10.1.6. Fractionation of Molasses using Ultrafiltration
1.10.1.7. Gel Chromatography Fractionation of Molasses
1.10.1.8. The Dilution Effect of Molasses
1.10.2. Results
2. ARYL ALCOHOL OXIDASE (AAO)
2.1. Methods
2.1.1. Medium
2.1.2. AAO Purification
2.1.3. Detection of Maltose Concentration
2.1.4. Determination of Oxidization Activity of AAO in Cell-free Culture Broth
2.1.5. AAO Oxidizing Activity Toward Veratryl or Benzyl Alcohols
2.1.6. DyP Decolorization Toward RB5 in the Mixture of DyP and AAO
2.1.7. Determination of H2O2 Concentration in the Culture Broth of Dec l
2.1.8. Benzyl Alcohol and Aldehyde Derivatives Detection from Culture Broth of Dec l
2.1.9. Prevention of Polymerization of Dye-degraded Products by AAO
2.2. Results
2.2.1. Change in Activities of DyP and AAO
2.2.2. Purification of AAO
2.2.3. Effects of pH and Temperature on AAO Activity
2.2.4. Substrate Specificity of AAO
2.2.5. Detection of Benzyl Alcohol, Aldehyde Derivatives, and H2O2 from the Culture Broth of Dec l
2.2.6. Dye Decolorization by the Mixture of DyP and AAO
2.2.7. Prevention of Polymerization by AAO
3. VERSATILE PEROXIDASE OF DEC 1, TCVP1
3.1. Methods and Results
3.1.1. Cultivation of Dec 1
3.1.2. Enzyme Purification
3.1.3. N-terminal Amino Acid Sequencing of TcVP1
3.1.4. Effects of pH and Temperature on the Activity of Versatile Peroxidase, TcVP1
3.1.5. Substrate Specificity
3.1.6. Complete Decolorization of RB5 by rDyP and TcVP1 In Vitro
CONCLUSION
Enhanced Productivity of a New Peroxidase DyP by Genetic Manipulation and by Cultivation Methods
Abstract
1. CLONING OF DYP GENE FROM DEC 1
2. CHARACTERISTICS OF DYP AS A PEROXIDASE
3. EXPRESSION OF DYP IN A. ORYZAE
3.1. Characteristics of Recombinant DyP, rDyP
3.2. Crystallization and X-ray Analysis of DyP
4. Enhanced PRODUCTION OF DYP OR RDYP
4.1. Repeated-batch and Fed-batch Cultures by Recombinant A. oryzae
4.1.1. Methods
4.1.1.1. Strain
4.1.1.2. Batch Culture
4.1.1.3. Repeated-batch Culture
4.1.1.4. Fed-batch Culture
4.1.1.5. Media
4.1.1.6. Determination of rDyP Activity
4.1.2. Results
4.1.2.1. Repeated-batch Culture Using Medium-1
4.1.2.2. Repeated-batch Culture Using Medium-3
4.1.2.3. Fed-batch Culture Using Medium-5
4.1.2.4. Fed-batch Culture Using Medium-6
4.1.3. Discussion
4.2. Repeated-batch Liquid Cultivation by Recombinant A. oryzae in Synthetic Medium
4.2.1. Methods
4.2.2. Results
4.3. Solid-state Cultivation of Recombinant A. oryzae
4.3.1. Methods
4.3.1.1. Medium and Strain used
4.3.1.2. Solid-state Culture (SSC)
4.3.1.3. rDyP Activity Analysis
4.3.1.4. Liquid Culture
4.3.2. Results
4.4. Air Membrane Surface (AMS) Reactor Culture of Dec 1
4.4.1. Methods
4.4.1.1. Liquid Culture
4.4.1.2. AMS Reactor
4.4.1.3. Protein and Enzyme Analysis
4.4.1.4. Dye Decolorization
4.4.2. Results
4.4.2.1. Secreted Protein and DyP Production in Liquid Culture
4.4.2.2. Secreted Protein and DyP Production by AMS Culture
4.4.2.3. Decolorization of Dyes by Employing AMS Culture
4.4.2.4. Effects of Water Content on Enzyme Activities
4.4.2.5. Scanning Electron Micrographs of Mycelia
4.4.2.6. Enzyme Production by AMS and Liquid Cultures
4.4.3. Discussion
4.5. Production of Two Isozymes of DyP in AMS Culture
4.5.1. Methods
4.5.2. Results
4.6. Productivity Enhancement of DyP
CONCLUSION
Dye Decolorization by Immobilized Recombinant rDyP and Turnover Capacity of rDyP
Abstract
1. IMMOBILIZATION OF RDYP ON MESOPOROUS MATERIALS
1.1. Properties of Newly Synthesized Mesoporous Materials
1.2. Effect of pH on Adsorption and Activity Efficiency of rDyP Immobilized on the Mesoporous Materials
1.3. Leaching of Immobilized rDyP from Mesoporous Materials
1.4. Stability of Free and Immobilized rDyP When Exposed to H2O2
1.5. Effect of pH on Stability of Free rDyP
1.6. Decolorization of Dye by rDyP Immobilized on Mesoporous Materials in Repeated-batch Mode
1.7. Discussion
2. CHANGE IN TURNOVER CAPACITY OF RDYP
2.1. Methods
2.2. Experimental Conditions
2.3. Results
2.3.1. C1. Batch Addition of Equimolar H2O2 and Dye
2.3.2. C2. Stepwise Fed-batch Addition of H2O2 and Dye
2.3.3. C3. Stepwise Fed-batch Addition of H2O2 at a Higher Concentration of Dye
2.3.4. C4. Fed-batch Addition of Dye and Continuous Fed-batch Addition of H2O2
2.4. Discussion
CONCLUSION
Application of Dec 1 for Decolorization of Other Colored Substances
Abstract
1. DECOLORIZATION OF MOLASSES BY DEC 1
1.1. Methods
1.1.1. Decolorization Methods
1.1.2. Determination of the Activity of DyP
1.2. Results
2. SIMULTANEOUS DECOLORIZATION OF MOLASSES AND A DYE
2.1. Methods
2.1.1. Inhibitory Effect of a Fraction of the Molasses Medium on the Activity of Purified Peroxidase, DyP
2.1.2. Decolorization of Ultrafiltered Fractions of Molasses by Dec 1
2.1.3. Decolorization of Molasses and the Dye in Invertase-treated Molasses Medium
2.2. Results
2.2.1. Growth of Dec 1 in Molasses
2.2.2. Decolorization of Molasses and a Dye in a Molasses Medium
2.2.3. Change in DyP Activity During the Growth of Dec 1 in Molasses Medium
2.2.4. Decolorization of Molasses by Purified DyP
2.2.5. Decolorization of Molasses and the Dye in the Invertase Hydrolyzed Medium
2.3. Discussion
3. SIMULTANEOUS DECOLORIZATION OF MOLASSES AND A DYE BY SUSPENDED AND IMMOBILIZED DEC 1 CELLS
3.1. Methods
3.1.1. Decolorization Rate of the Dye, RB5
3.1.2. Immobilization of Dec 1
3.2. Results
3.2.1. Decolorization of Molasses by Suspended Dec 1
3.2.2. Decolorization of Molasses by Immobilized Dec 1
4. DECOLORIZATION OF KRAFT PULP BLEACHING EFFLUENT BY DEC 1
4.1. Methods
4.1.1. Strain and Preparation
4.1.2. Lignin Decolorizing Assay
4.1.3. Effect of pH on the Kraft Pulp Bleaching Effluent, E-effluent
4.1.4. Cultivation of Dec 1
4.1.5. Color Removal Method
4.1.6. Color Adsorption to Dec 1 Cells
4.1.7. Absorbable Organic Halogens (AOX)
4.1.8. Gel Filtration of E-effluent
4.1.9. Enzyme Assay
4.1.9.1. DyP
4.1.9.2. Manganese Peroxidase (MnP)
4.1.9.3. Lignin Peroxidase (LiP)
4.1.9.4. Laccase (Lac)
4.2. Results
4.2.1. Decolorization of Lignin by Dec 1
4.2.2. Effect of pH on E-effluent
4.2.3. Decolorization of the E-effluent
4.2.4. Color Adsorption to Dec 1 Cells
4.2.5. Reduction of AOX
4.2.6. Molecular Weight Distribution of the E-effluent Determined by Gel Filtration
4.2.7. Enzyme Activity and Color Removal
4.3. Discussion
5. DECOLORIZATION OF OXYGEN-DELIGNIFIED BLEACHING EFFLUENT (OBE) AND BIOBLEACHING OF OXYGEN-DELIGNIFIED KRAFT PULP (OKP) BY DEC 1
5.1. Methods
5.1.1. Strain
5.1.2. OBE and Oxygen-delignified Kraft Pulp (OKP)
5.1.3. Medium
5.1.4. Decolorization of OBE by Dec 1
5.1.4.1. Decolorization
5.1.4.2. Color Removal
5.1.5. Biobleaching of OKP by Dec 1 Grown in OGY30 Medium
5.1.5.1. Cultivation of Dec 1
5.1.5.2. Biobleaching of OKP by Dec 1
5.1.6. Biobleaching of OKP by the Culture after the Decolorization of OBE by Dec 1
5.1.6.1. Decolorization of OBE by Dec 1
5.1.6.2. Biobleaching of OKP by the Culture Filtrate
5.1.7. Measurement of Brightness and the Kappa Value of Pulp
5.1.8. Enzyme Assay
5.2. Results
5.2.1. Decolorization of OBE by Dec 1
5.2.2. Biobleaching of OKP by Dec 1 Cultured in OGY30 Medium
5.2.3. Biobleaching of OKP by the Culture Filtrate After the Decolorization of OBE by Dec 1
CONCLUSION
REFERENCES
Decolorization by Thanatephorus Cucumeris Dec 1
Authored by
Makoto Shoda
Emeritus of Tokyo Institute of Technology
316,1-4-2 Shin-Ishikawa, Aobaku
Yokohama, Japan
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PREFACE
Makoto ShodaThis book summarizes the results of more than 20 years of research on a fungus, Dec 1, which was isolated in my laboratory, as a candidate to decolorize colored substances that are visible xenobiotic pollutants and cause serious environmental problems.
Various studies have attempted to improve or change environments using biological activity, but practical examples of this technique are rare. This is mainly because of the lack of basic information and the lack of optimization in reactor systems. In this book, one possibility of using a single microorganism to resolve environmental issues related to colored substances is demonstrated.
This book is composed of six chapters, and various aspects of isolated Dec 1, including its scientific analysis and the optimization of the engineered production of enzymes responsible for decolorization, are provided in it.
The features and characteristics of Dec 1 included in this book are as follows:
1. Dec 1 has been isolated as a fungus for dye decolorization. It has been finally identified as the non-white rot fungus, Thanatephorus cucmerins Dec 1.
2. The decolorization spectrum of Dec 1 is vast, indicating that Dec 1 produces multiple enzymes that are responsible for decolorization.
3. New peroxidase DyP, Aryl Alcohol Oxidase (AAO), Manganese Peroxidase (MnP), and Versatile Peroxidase (TcVP1) have been purified from Dec 1 and characterized.
4. The gene, dyp, has been cloned from Dec 1 and expressed in the fungus, Aspergillus oryzae.
5. Recombinant DyP (rDyP) has been shown to be almost identical to DyP.
6. The unique characteristics of DyP have been verified using crystallization and X-ray analysis.
7. The immobilization of rDyP has been found to be successful only when using a new mesoporous material as a carrier.
8. The enhanced production of DyP has been attempted using liquid culture, Solid-state Culture (SSC), repeated-batch culture, and fed-batch culture.
9. To overcome the problems involved in these culture methods, an Air Membrane Surface (AMS) reactor has been introduced, and increased rDyP production has been confirmed.
10. The final level of rDyP production has been found to be more than a half million-fold higher than the original level of DyP production by T. cucumeris Dec 1.
11. Dec 1 has demonstrated efficient decolorization of molasses waste, kraft pulp bleaching effluent, and oxygen-delignified bleaching effluent.
12. As substrates for Dec 1 growth, complex media, including rice bran powder, wheat bran powder, and molasses, have been used, and the advantages of each medium have been described.
I believe that this book will provide researchers in this field with a useful resource to support current knowledge on biological decolorization and will provide students with a logical and practical scheme for approaching colored substance treatment. During my time as the Director of the Resources Recycling Process Laboratory, at the Chemical Resources Laboratory, Tokyo Institute of Technology, many researchers and graduate and undergraduate students at the campus and in private companies have helped to accomplish these results. I am indebted to the following researchers for giving me the opportunity to study Dec 1 and for offering many valuable suggestions: Drs. Y. Sugano, M. Hirai, T. Sato, M. Iwamoto, T.H. Lee, N. Uematsu, and J. Sugiura. l am also grateful to the following graduate and undergraduate students: S.J. Kim, M. Shakeri, N. Shintani, T. Shimokawa, K. Sasaki, Y. Matsushima, A. Ichiyanagi, R. Muramatu, R. Sasaki, N. Suzuki, and C. Matsuo.
I especially thank Professors T. Imanaka, Professor Emeritus of Kyoto University, and J. Takahashi, Professor Emeritus of Obihiro University of Agriculture and Veterinary Medicine, for their cooperation on this book.
Makoto Shoda
Emeritus of Tokyo Institute of Technology
316,1-4-2 Shin-Ishikawa, Aobaku
Yokohama, Japan
Introduction
Makoto Shoda1,*
1 Emeritus of Tokyo Institute of Technology 316,1-4-2 Shin-Ishikawa, Aobaku, Yokohama, Japan 225-0003
*Corresponding author Makoto Shoda: Emeritus of Tokyo Institute of Technology 316,1-4-2 Shin-Ishikawa, Aobaku, Yokohama, Japan 225-0003; E-mail:
[email protected]Several colored effluents, including dyes, molasses, and pulp bleaching effluents, are released into environments, and efficient treatments of these colored substances are urgent because severe regulation has been established worldwide. Conventional treatment methods have several disadvantages, such as energy consumption or the risk of treated products. On the other hand, biological treatment has advantages over conventional chemical and physical treatment because the products after treatment are relatively safer and the utilization of energy is smaller than other methods. However, in biological methods, finding efficient microorganisms and utilizing them under optimal conditions are key challenges.
A fungus, strain Dec 1 was isolated in our laboratory, exhibiting a wide spectrum of degradability for colored substances [1]. This fungus was initially identified as Geotrichum candidum Dec 1, but finally reidentified as Thanatephorus cucumeris Dec 1. In this book, the characteristics of Dec 1 have been described not only from the basic biochemical points, but also from an engineering point of view associated with its application for the decolorization of dyes, molasses, and pulp bleaching effluents.
More than 5×104 tons of dyes are used annually in Japan [2], and 10 to 15% of these dyes are estimated to be discarded into the environment [3]. As colored effluents from major textile and dyestuff industries are recalcitrant to biodegradation, the treatment of these effluents in wastewater treatment systems is mainly based on physical and/or chemical procedures, such as adsorption, concentration, chemical transformation, and incineration. Although these treatment methods are effective, they have several shortcomings, such as high cost, formation of hazardous byproducts, and intensive energy requirements. Therefore, biological degradation methods are receiving attention as better alternatives. Several strains as dye-degrading microorganisms have been reported, such as white-rot fungi Phanerochaete chrysosporium [4, 5], Pleurotus ostreatus
[6], Coriolus versicolor [7], and Streptomyces spp [4]. The effectiveness of microbial treatment depends on the survival, adaptability, and stable activity of the selected microorganisms in the treatment environment.
The newly isolated strain, Dec 1 decolorized various reactive dyes, including azo and anthraquinone dyes, as shown in Chapter 2. The involvement of several extracellular enzymes, such as Lignin Peroxidase (LiP) and Manganese Peroxidase (MnP), was suggested by the broad decolorization spectrum of this strain. LiP, MnP, Laccase (Lac), and Horseradish Peroxidase (HRP) have also been reported to have the ability to decolorize various dyes [8-14]. Among them, MnP from P. chrysosporium is a representative enzyme and plays a major decolorizing role in the presence of manganese ions [15, 16]. However, Mn-oxidizing peroxidases isolated from Bjerkandera adusta and Pleurotus eryngii decolorized with multiple azo dyes, regardless of the presence of manganese ions [17]. Those enzymes were able to oxidize Mn2+ to Mn3+ at pH 5 and also oxidized aromatic compounds, such as Veratryl Alcohol (VA), a typical substrate of LiP, at pH 3, regardless of the presence of Mn2+ [18, 19]. Therefore, these enzymes that expressed both LiP-like and MnP-like characteristics were named MnP–LiP hybrid peroxidase or manganese-independent peroxidase or Versatile Peroxidases (VPs).
The enzymes responsible for the dye-decolorizing activity of Dec 1 were purified. Their characteristics are clarified in Chapter 3. One of them is a new peroxidase, DyP that is a glycoprotein with a molecular mass of 60 kDa, showing a high ability to decolorize anthraquinone dyes [20].
The culture broth of Dec l showed the ability to oxidize Veratryl Alcohol (VA), but DyP did not degrade VA [21], suggesting Dec 1 to produce Aryl Alcohol Oxidase (AAO) [22]. Therefore, a veratryl alcohol-oxidizing enzyme was purified from the culture broth of Dec 1, and its enzymatic characteristics and roles in dye decolorization have been characterized in vivo, as elucidated in Chapter 3.
Dec 1 showed complete decolorization of anthraquinone dye, Reactive Blue 5 (RB 5) in vivo. However, this phenomenon was not observed in vitro by DyP alone. DyP changed the color of RB5 from dark blue to a light reddish-brown colored substance composed of an azo complex mixture. Based on this observation, for the complete decolorization of Dec 1, the involvement of other enzymes in addition to DyP has been suggested. Then, a novel Versatile Peroxidase (VP) from Dec 1, named TcVP1, was isolated and characterized. The first complete in vitro decolorization of an anthraquinone dye using DyP and TcVP1 [23] is described in Chapter 3.
DyP has two specific characteristics. The first characteristic is its high ability to decolorize anthraquinone dye at around pH 3 and it lacks an important histidine residue that is involved in other fungal peroxidase tertiary structures. Instead, DyP includes aspartic acid and arginine [24], as described in Chapter 4.
The second characteristic is that DyP belongs to a novel DyP-type peroxidase family.
Peroxidases are classified into two types: animal and plant peroxidase superfamilies. The plant peroxidase superfamily is further categorized into three classes according to the origin [25]. Class I peroxidases are prokaryotic, and the representatives are Cytochrome C peroxidase (CCP) and Escherichia coli Peroxidase (ECP) [26, 27]. Class II peroxidases are secretory fungal peroxidases and the representatives are Arthromyces ramosus Peroxidase (ARP), Lignin Peroxidase (LiP), and Manganese Peroxidase (MnP) [28-30]. Class III peroxidases are classical, secretory plant peroxidases, and the representatives are Horseradish Peroxidase (HRP) and Turnip Peroxidase (TP) [31, 32]. According to this classification, DyP belongs to class II. However, the characteristics of DyP are different from those of class II peroxidases. Thus, to clarify the classification of DyP, the gene encoding DyP was cloned from the cDNA library of Dec 1, and the primary structure of DyP was compared with those of other peroxidases. This analysis revealed DyP as a unique peroxidase among previously reported peroxidases. The detail is provided in Chapter 4.
Cloning the cDNA of the dyp gene was successful, and the gene encoding DyP was transformed into the host, Aspergillus oryzae, under the control of the amyB promotor. The massive production of rDyP using recombinant A. oryzae is of primary interest in order to use crude rDyP directly for decolorization. This aspect is described in Chapter 4.
A. oryzae was selected as a host because this fungus is recognized as a safe host and it has a high growth rate and can secrete gram-per-liter quantities of heterologous proteins [33].
As the productivity of original peroxidase, DyP, by Dec 1 was extremely low, the enhanced productivity of rDyP was tried. First, the production of rDyP by recombinant A. oryzae was enhanced, but it was still not found to be satisfactory [34, 35]. Most common bioprocesses for large-scale production of chemicals use batch culture, which has the advantages of stable nongrowth-associated product formation, maintenance of genetic stability, and a relatively low risk of contamination [36]. Especially when natural substrates are chosen, complex solid particles are involved in the substrates, and continuous cultivation complicates the operation. Thus, the production of the enzyme, rDyP, using different batch cultivation methods and complex substrates is demonstrated in Chapter 4.
As batch culture methods, repeated-batch and fed-batch cultures are the conventional methods of achieving high enzyme production. In repeated-batch culture, the medium is periodically sedimented to draw off the liquid broth, and fresh medium is supplied. This method is effective because the grown microorganisms can be reused and the lag phase associated with at the start of cultivation, and reactor clean-up after cultivation, preculture preparation, and bioreactor sterilization between each batch are unnecessary. For example, in the LiP production by P. chrysosporium, repeated batch cultures with 3-4 days cycles were conducted for more than 30 days [37, 38].
In fed-batch culture, fresh medium is supplied intermittently until the culture volume reaches its upper limit. This method is useful in that the by-product formation is minimal, and the energy source is efficiently utilized. Fed-batch culture is worldwide applied when high cell masses are desired, like in the production of baker’s yeast [39, 40]. As no fed-batch culture has been reported for the production of fungal enzymes, this method could be more time and cost-effective than repeated-batch culture because it requires less operation time. The comparison of fed-batch and repeated-batch cultures for the production of rDyP is provided in Chapter 4.
Dec 1 requires carbon sources to exhibit decolorization ability for a wide range of colored substances. Molasses is a practically available carbon source because it is widely used in the microbial industry. However, molasses contains highly colored substances, like melanoidins, and the color is deepened by the Maillard reaction after sterilization and remains in Molasses Wastewater (MWW) after its use [41]. Therefore, it is important to decolorize molasses after its use as a carbon source. Several microorganisms have been reported to be effective for the microbial decolorization of MWW [42-44], but the decolorizing efficiency is still not satisfactory.
First, molasses was used as a carbon source for the production of DyP, and the effect of molasses on its activity was investigated using a jar fermenter with different sufficient oxygen supply modes [45, 46], as shown in Chapter 3.
Next, the simultaneous decolorization of molasses and anthraquinone dye, RB5, in the molasses medium by Dec 1 was examined to assess the factors involved in the decolorization process [21]. The immobilized cells of Dec 1 on polyurethane foam and suspended Dec 1 cells were compared by repeated-batch culture for long-term simultaneous decolorization of molasses and dye, RB5 [47]. The result is demonstrated in Chapter 6.
The choice of substrate is an important factor in optimizing enzyme productivity. The optimization involves the evaluation of the cost and the availability of nutrients and the reactor system. Repeated-batch culture and fed-batch culture were selected to compare rDyP production using wheat bran powder and rice bran powder as complex substrates, as shown in Chapter 6.
As mentioned above, inexpensive complex media, such as rice bran powder and wheat bran powder, have been found effective for the high production of DyP in the two batch cultures. On the other hand, defined synthetic media are more expensive than complex media, but have more advantages over complex media in that reproducible cultivation performance is possible, process control and monitoring are easier, sterilization conditions are stable, and thus, the scaling-up problem is minimized [48]. The rDyP production in the repeated-batch process of A. oryzae pellets using a defined synthetic medium containing maltose as a carbon source increased the number of cycles and avoided the drawback of unstable rDyP activity caused by complex media, as described in Chapter 4. Maltose induced Taka-amylase A production and A. oryzae was transformed with the dyp gene from Dec 1 using the amyB promoter, and thus, high enzyme production became possible [49]. Then, the decolorization of an anthraquinone dye, Remazol Brilliant Blue R, abbreviated as RBBR, using a crude enzyme solution was conducted, as described in Chapter 4.
Among several cultivation methods, liquid cultivation is generally a convenient method for the growth of microorganisms and the isolation of products. The control of cultivation conditions is easy in liquid cultivation because of the uniform culture broth. However, enzyme production hardly occurs in liquid cultivation by the fungus. The liquid cultivation for enzyme production and in vivo wastewater treatment is not applicable because large amounts of suspended solids, including fungal flocs, metabolites, and solid particles derived from complex medium, disturb the smooth flow and oxygen transfer. On the other side, Solid-state Fermentation (SSF) or Solid-state Cultivation (SSC) is widely applied in the fungal industry for the production of metabolites from fungi. In Japan, traditional industrial cultivation with A. oryzae has been conducted using SSC.
The authors showed that even as a bacterium, Bacillus subtills produced lipopeptide antibiotics 10 times higher using SSC compared to liquid culture [50]. One of the advantages of SSC is that fungal surface growth under SSC conditions leads to high enzyme productivity. Thus, protein productivity per unit of dry cell mass in SSC has been reported to be higher than that in liquid culture [51]. The production of β-fructofuranosidase, feruloyl esterase, and lipase from SSC by Aspergillus niger was higher than that from liquid culture [52-54]. However, some problems remain to be solved in SSC. The extraction and isolation of secreted enzymes in SSC are difficult because of the lack of an aqueous phase and the use of rich suspended solids. In SSC, it is difficult to control incubation conditions, such as temperature, pH, and moisture content, because of heterogeneous solid culture broth.
To resolve these disadvantages of SSC and liquid culture in fungal cultivation, an Air Membrane Surface (AMS) bioreactor was introduced [55, 56]. This reactor system is a hybrid of liquid and solid-state culture. The production of a neutral protease from A. oryzae was 10-fold higher than using liquid culture [55], and smooth repeated-batch cultures were successful in accelerating enzyme production because the solid and aqueous phases are separated in AMS [57, 58]. Therefore, the AMS reactor allows easy isolation of soluble secreted proteins in the aqueous phase without contamination of suspended solids. Additionally, this method saves energy. Therefore, Dec 1 cultures using AMS bioreactor have been reported to be focused on the effect of enzyme productivity on in vivo dye decolorization and the relation between biofilm formation and water activity. Using the AMS reactor, two new isozymes of rDyP were produced and characterized, as detailed in Chapter 4.
The direct use of enzymes for water treatment is attractive due to the following advantages, such as the compatibility of enzymes, ease of use, simplicity of process control, and wide applicability to different contaminants and pHs [59, 60]. However, despite the potential industrial use of enzymes, their practical application is difficult because enzymes are available in limited amounts, and enzyme stability under unstable environments is low [61]. The dyp gene encoding DyP was transformed into A. oryzae to produce rDyP, and the enhanced production of rDyP by more than 100-fold compared to the production of Dec 1 was demonstrated. Furthermore, the selection of proper cultivation methods can increase the rDyP production level by nearly half a million times than the original level of DyP, leading to the possibility of using DyP practically, as described in Chapter 4.
When rDyP is applied in industrial processes, some constraints, such as maintenance of its stability, reuse of rDyP, and inactivation by H2O2, have to be considered [62]. To overcome these constraints and enhance the catalytic efficiency of rDyP, immobilization of rDyP is a potential method. Several enzyme immobilizations have been attempted. They include adsorption to activated carbon and glass beads, cross-linking by glutaraldehyde, and entrapping by photosensitive resins, but these methods for immobilization of rDyP have not been successful, presumably because of the completely new tertiary structures of rDyP relative to well-known peroxidases, as highlighted in Chapters 3 and 4 [23, 63]. In addition, ordered mesoporous materials have been applied for rDyP because they are new candidates for the immobilization of enzymes due to their uniform and adjustable pore size in the range of 1-10 nm, large surface area, pore volume, mild immobilization conditions, and efficient physical adsorption [64-66]. Mesoporous materials, FSM-16 and AlSBA-15, were synthesized using cationic and non-ionic surfactants, respectively, and immobilization of rDyP was attempted on these materials. RBBR was decolorized by immobilized rDyP on these materials using repeated-batch mode, as presented in Chapter 5.
When rDyP is used for dye decolorization, the decolorization activities of rDyP are affected due to the presence of several factors, such as organic acids, pH, temperature, and salts. Among them, inactivation by excess H2O2 is the most critical factor [67].
In order to obtain a quantitative evaluation of rDyP inactivation by H2O2, the potential turnover capacity of crude rDyP was introduced in batch and fed-batch cultures [62]. The turnover capacity is defined as the unit concentration of reducing substrate removed from the solution per unit concentration of inactivated enzyme [68]. In order to determine the potential of rDyP when the enzyme is subjected to H2O2 in different ways, the turnover capacity is useful, as described in Chapter 5. The capacity can compare the ability of rDyP with other dye decolorization methods, such as adsorption, oxidative processes, and microbial treatment.
Another possible application for Dec 1 is the treatment of kraft pulp bleaching effluent that contains colored substances and chlorinated organic compounds, causing environmental pollution because of their poor biodegradability and toxicity [69, 70]. Several methods, such as lime coagulation, rapid land filtration, membrane processes, and polymeric adsorbents, have been adopted to remove these waste products [71]. However, in general, these processes have several problems, such as high energy costs and the generation of hazardous by-products. As microbial treatments, white rot fungi, P. chrysosporium, C. versicolor, Pleurotus ostreatus, and Streptomyces spp. have been reported [72-75]. Although the microbial methods are energy-saving, some fungi are plant pathogens, have limited substrate specificity, and have a slow growth rate.
On the other hand, non-pathogenic Dec 1 secretes beneficial enzymes and shows a faster growth rate than other white rot fungi. Therefore, Dec 1 was applied to decolorize kraft pulp bleaching effluent to evaluate its effectiveness for the removal of colored substances, as described in Chapter 6.
The multistage bleaching process using chlorine-based chemicals was employed to remove existing lignin in Unbleached Kraft Pulp (UKP) in the paper industry. However, environmental pollutants, such as dioxins, are included in effluent drained from chlorine-based bleaching process [70, 76]. The oxygen bleaching process is a developed process to reduce the consumption of chlorine-based chemicals [77]. However, the removal of colored substances, including lignin, is still unsatisfactory in Oxygen-delignified Bleaching Effluent (OBE) drained from the oxygen bleaching process. As the content of residual lignin of Oxygen-delignified Kraft Pulp (OKP) is half of UKP, biobleaching is applicable. For the biobleaching, Trametes versicolor [78], a fungus IZU-154 [79], Bjerkandera sp. strain BOS55 [80], and a fungus SKB-1152 [81] showed the possibility of removing the lignin of pulp. Here, a decolorizing non-white rot fungus, Dec 1, was applied to the decolorization of OBE and the bleaching of OKP, as described in Chapter 6.
Dec 1 was initially identified as Geotrichum candidum. This fungus was introduced to have biotechnological importance [82] and has been used in mixed culture with other fungi for reducing environmental issues [83].
Since DyP was purified from fungus Dec 1 and proposed as a new DyP-type peroxidase [21, 24], the enzyme DyP was classified as a new family of heme peroxidase. DyP-type peroxidases were then isolated from various microorganisms and characterized. Bacterial genes of P class DyP and a phylogenetic tree with all available sequences of DyP genes offered an overview of their presence in the bacteria kingdom [84]. Ligninolytic bacteria, Raoultella ornithinolytica OKOH-1, showed DyP-type peroxidase degradation of congo red and melanin [85]. A salt-tolerant yeast, Galactomyces geotricum, degraded red azo dyes using two lignin peroxidase and laccase [86]. A new DyP from thermophilic actinomycete, Thermomonospora curvata (TcDyP), was then identified and characterized. TcDyP is highly close to fungal DyPs with the unique catalytic property of A-type DyPs [87]. Streptomyces albidoflavus BSII#1 oxidized seventeen phenolic substrates with actinobacterial DyP type peroxidase [88].
We also isolated a new versatile peroxidase VP from Dec 1 named TcVP1 and characterized it, and then, the first complete in vitro decolorization of an anthraquinone dye using DyP and TcVP1 was demonstrated [23]. Dye-decolorizing Peroxidase (DyP) from Auricularia auriculajudae and Versatile Peroxidase (VP) from Pleurotus eryngii oxidized mononitrophenol isomers, and the oxidation site was located at residue in both DyP (Trp377) and VP (Trp164) [89].
DyP produced from Vibrio cholerae catalyzed the anthraquinone dyes with acidic optimal pH and identified the decisive factor of the low pH [90]. Fungal DyPs showed efficient lignin biodegradation [91].
On the other hand, immobilization of DyP was found effective in enhancing the decolorization rate and efficiency, as described in detail in Chapters 3 and 4. The agarose-immobilized MnP exhibited a remarkable bioremediation potential as a biocatalyst using a packed bed reactor system [92]. Some books and papers have summarized the latest advancements in colored wastewater treatment, and introduced DyP to have potential application as a biocatalyst in advanced biological processes [93-96].
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