Open access peer-reviewed chapter
Abstract
Oxidative enzymes are often considered for use in industrial processes because of the variety of reactions they are able to catalyse. In the past, most of these oxidative enzymes were obtained from fungi. However, in recent years, it has become evident that these enzymes are also produced by bacteria, including actinobacterial strains, which can therefore be considered as an underexploited resource of oxidative enzymes with potential for application in various industries. This chapter will focus on selected oxidative enzymes found in actinobacteria, their potential for application in industrial processes and how we can access and improve these enzymes to suit the required bioprocess conditions.
Keywords
- Actinobacteria
- oxidative enzymes
1. Introduction
Dating back over 100 years, organisms belonging to the class
Over the past 20 years, there has been an increased interest in the production of certain oxidative enzymes by actinobacteria, especially in the field of lignin degradation, detoxification of organic pollutants such as polynuclear aromatic hydrocarbons (PAHs), organophosphate pesticides and azo dyes [2]. Oxidative enzymes or oxidoreductases (EC 1.x.x.x) catalyse oxidation-reduction reactions. These enzymes are further subdivided depending on the target donor molecule and the final electron acceptor, for example, the EC 1.1 grouping refers to enzymes acting on the CH-OH group of donors, while EC 1.1.3 further defines the type of electron acceptor, which in this case is oxygen (http://www.chem.qmul.ac.uk/iubmb/enzyme/). Based on the information obtained from BRENDA, the comprehensive enzyme information system (http://www.brenda-enzymes.org/), there have been more than 1,500 reports of oxidoreductase production by actinobacteria. This represents a vast number of oxidoreductases, and therefore this chapter will focus only on enzymes grouped within the EC 1.x.3.x, EC 1.11.x.x, EC 1.13.x.x and EC 1.14.x.x classes (where oxygen acts as an electron acceptor or is incorporated during the reaction). Based on the literature reported in BRENDA, the following actinobacterial genera are represented in the EC 1.x.3.x, EC 1.11.x.x, EC 1.13.x.x and EC 1.14.x.x classes:
Figure 1.
The class
2. Oxidative enzymes produced by actinobacterial strains
The development of practical biocatalytic oxidation/reduction (redox) processes is very important because many chemical and biochemical transformations involve redox processes [4]. It is therefore not surprising that oxidative enzymes have been applied to a wide range of industrial processes: in the beverage industry to remove phenolics (which causes turbidity) in drinks and to remove/diminish the cork smell in wines [5]; in the pulp paper industry to delignify wood for the bleaching process [6]; in the textile industry to decolourise the effluents from dyes [7]; to bleach textiles and to synthesise dyes [8]; in the nanotechnology industry as biosensors [9-11]; for clinical and environmental analysis and for cosmetics, for use in hair dyes and for skin lightening creams [12]. Peroxidases, laccases and tyrosinases are among the oxidative enzymes that have been widely researched and much information on their applications and mechanistic roles has been published, with a particular interest directed towards the fungal producers. In this chapter, we wish to draw attention to selected oxidative enzymes that are produced by the actinobacteria, including the well-known peroxidases, laccases and tyrosinases.
2.1. Cholesterol oxidase (EC 1.1.3.6)
Many actinobacteria carry out useful biotransformations, which allow for the production of a wide range of substances of clinical and commercial interest [13]. Of these compounds, steroids are among the most important pharmaceutical products used for the treatment of various diseases [14]. In eukaryotic organisms, cholesterol is an essential component for the maintenance of cell membrane structure and the synthesis of a number of compounds. Cholesterol oxidase (CO) is a prokaryotic enzyme that has been very useful for biotechnological applications, where it has been applied in the detection and conversion of cholesterol [15-17].
CO is a flavoenzyme that catalyses the oxidation and isomerisation of cholesterol to cholest-4-en-3-one, with the reduction of oxygen at C3 to hydrogen peroxide (Figure 2) [18-19]. Despite having a broad substrate range, the presence of a 3β-hydroxyl group is essential for the CO activity. Whilst most microorganisms produce cell-bound CO, the actinobacteria are prolific producers of high levels of extracellular CO [18].
Figure 2.
Mechanism of action of cholesterol oxidase on cholesterol (adapted from [
Actinobacterial COs have been isolated from
2.2. Laccases (EC 1.10.3.2)
Laccases are multicopper oxidases that catalyse the one electron oxidation of four reducing substrate molecules, whilst simultaneously reducing molecular oxygen to water [24] (Figure 3). These enzymes have been extensively studied and are ubiquitous in nature, found in higher plants, fungi, insects and prokaryotes, including the actinobacteria [24, 25]. For decades, laccases of fungal origin have been at the centre of research efforts. Bacterial laccases, or laccase-like enzymes, are becoming increasingly prominent. As of 2014, less than 10 bacterial laccases from actinobacteria have been fully characterised, all of which belong to the genus
Figure 3.
The typical laccase reaction mechanism [
Whilst typical laccases consist of three cupredoxin domains, several laccases of actinobacterial origin have only two domains, which were first elucidated for the structure of the small laccase (SLAC) from
Given the broad substrate range that laccases exhibit they can be applied in a wide variety of industries. Laccases have been employed in the following industrial applications: degradation of dyes in the textile industries, in the food industry where the consumption of oxygen in the packaged food is removed to avoid spoilage, in the paper and pulp industries for decolourisation of ink and the breakdown of lignocellulosic compounds, bioremediation for the treatment of toxic environmental pollutants including PAHs, pesticides, dyes from the textile industries and endocrine-disrupting chemicals [12, 35-40].
2.3. Peroxidases (EC 1.11.1.X)
Peroxidases are a large group of oxidoreductases that catalyse the oxidation of substrate molecules using hydrogen peroxide (H2O2) as the electron acceptor, with the majority of peroxidases using haem as a cofactor [41]. Haem peroxidases are typically grouped into two superfamilies: the first consists of bacterial, fungal and plant peroxidases and the second contains peroxidases from animals, fungi and bacteria [42]. The production of peroxidases from actinobacteria have been well described and are a good potential source of novel industrially relevant peroxidases, especially in a market that is largely dominated by the plant horseradish peroxidase (HRP) [42-44].
Antonopoulos et al. [45] investigated the biotechnological potential of the extracellular peroxidase from
2.4. Catechol 1,2-dioxygenase (EC 1.13.11.1) / Catechol 2,3-dioxygenase (EC 1.13.11.2)
Aromatic compounds are typically broken down by bacteria. However, recalcitrant varieties, such as polycyclic aromatic compounds do exist [47]. Bacterial populations often possess genes which code for enzymes that are able to degrade such toxins into protocatechuate and catechol [48]. The catechol in question is further broken down by one of two processes: (1) the action of catechol 1,2-dioxygenase (C12O) via an
Figure 4.
The breakdown of catechol through the action of (1) catechol 1,2-dioxygenase and (2) catechol 2,3-dioxygenase.
These dioxygenases are widely distributed among actinobacteria. Molecular analysis of catechol-degrading bacteria has shown that the
Many studies have demonstrated the use of C12O and C23O in the degradation of environmental contaminants using actinobacterial strains. Sutherland et al. [52] demonstrated C12O activity in four thermophilic
2.5. Baeyer-Villiger monooxygenases (EC 1.14.13.X)
The interest in using Baeyer-Villiger monooxygenases (BVMOs) as biocatalysts has increased over recent years. BVMOs are flavin-dependent enzymes that are used to efficiently perform not only regio-, chemo- and/or enantioselective Baeyer-Villiger oxygenation reactions (Figure 5) using stoichiometric quantities of O2 as an oxidant and NADPH as an electron donor, but also selected sulphoxidations and epoxidations [58-62].
Figure 5.
The Baeyer-Villiger oxidation reaction in which BVMOs convert ketones to their corresponding esters [
BMVOs are widely distributed among bacteria, being especially prevalent among the actinobacteria, with an average of one BVMO per genome [64]. In contrast, no Type 1 BVMOs (based on sequence similarity) have been found in plant, human or animal genomes [63]. As with many oxidases, the presence of a highly conserved protein sequence motif can be used to identify BVMOs [65]. Using this motif, Fraaije et al
In
2.6. Cytochrome P450 monooxygenase (EC 1.14.14.1)
Cytochrome P450 monooxygenases (CYPs) are perhaps one of the most widely studied enzymes. CYPs are haem
CYPs, with their ubiquitous nature, have been observed in a number of actinobacteria. An environmental
2.7. Tyrosinases (EC 1.14.18.1)
Tyrosinases (polyphenol oxidases) are copper-dependent oxidases that catalyse the
Figure 6.
Enzymatic activity of tyrosinase on a monophenol (adapted from [
These enzymes are ubiquitous in nature, and serve a multitude of biological functions [80]. Most notably, tyrosinases play a key role in the production of melanin. For example, in plants tyrosinases are responsible for the browning of open surfaces in fruits [77], while in microbes melanin plays a key role in the defence of DNA against radiation and reactive oxygen species (ROS) and binds to toxic heavy metals [81-82]. Biologically active melanin has been shown to have many advantages, including anti-tumour activity and providing protection against UV radiation [83-86].
The most commonly used tyrosinase for commercial purposes is that of the fungus,
The cresolase and catecholase activities of tyrosinase are advantageous for many industrial processes, including the production of pharmaceutically important compounds such as the
2.8. Other oxidases
2.8.1. L-amino acid oxidases (EC 1.4.3.2)
L-amino acid oxidases (L-AAO) are oxidoreductases that catalyse the oxidative deamination of L-amino acids to yield keto-acids, ammonia and hydrogen peroxide [96]. These enzymes exhibit broad substrate ranges, and as such are commonly applied for the resolution of racemic mixtures. For example, an L-AAO isolated from
2.8.2. Putrescine oxidase (EC 1.4.3.10)
Putrescine is a low molecular weight diamine that belongs to a group of compounds that are termed biogenic amines. The accumulation of biogenic amines in foods, such as putrescine, can be used as a marker for food spoilage caused by microbes such as
2.8.3. L-glutamate oxidases (EC 1.4.3.11)
In contrast to the broad substrate range of L-AAOs, there are strict substrate-specific amino acid oxidases, such as the L-glutamate oxidases [105], the first of which was isolated from the actinobacterium,
2.8.4. Sarcosine oxidases (EC 1.5.3.1)
Sarcosine oxidases (SOs) catalyse the hydrolysis of sarcosine and formaldehyde, while simultaneously yielding hydrogen peroxide [109]. It is predominantly being exploited in clinical assays for the determination of creatinine in serum. The SO from
3. Industrial relevance of actinobacterial oxidases: how to access them and improve their functionality
With oxidising enzymes, as with most other enzyme groups, the emphasis in the discovery and development of new enzymes for industrial processes is increasingly focused on the properties of the new enzymes that need to match the stringent conditions imposed by the industrial setting [117]. Thus, while the existence of large numbers of novel enzymes is demonstrated constantly, via a host of modern gene discovery technologies, this evidence of their existence is not sufficient to guarantee our capacity to provide the enzymes that industry demands. A further consideration in the development and application of new oxidizing biocatalysts is the requirement to demonstrate the novelty which will provide market advantage [118]. Thus, as new oxidizing enzymes are discovered, they need to be characterised in terms of substrate selectivity, product scope and stability in the presence of process constraints (e.g. the presence of organic solvents, temperature conditions, pH conditions).
The majority of oxidising enzymes are co-factor-dependent, which leads to requirements for co-factor recycle or replacement in the industrial processes utilising these enzymes. While this can be overcome by application of whole-cell biocatalysts, the search for non-co-factor-dependent oxidases which can catalyse equivalent reactions is a useful goal [117-118]. The three most commonly used screening methods to screen for novel enzyme activity include: (1) screening environmental samples for organisms whose enzymes have the ability to catalyse certain reactions; (2) the use of protein engineering to manipulate an existing biocatalyst and (3) looking for novel functionality/substrate specificity in existing biocatalysts [119]. These processes along with others can further be grouped into molecular-based (
Figure 7.
Summary of standard methods employed to access novel enzymes and to improve enzymes.
3.1. Isolated strains
Based on information obtained from BRENDA, the majority of the reports on oxidative enzymes from actinobacterial strains have originated from studies based on isolated strains and yet only a fraction of the genera within the order
3.2. Metagenomics - Accessing the genetic information of the ‘unculturables’
Over the past two decades, there has been a dramatic increase in the number of metagenomics-based studies. The belief that the majority of bacteria in the environment (>99%) are unculturable or that culture techniques for their isolation have not yet been developed have necessitated the analysis of the metagenome [123]. With the advent of metagenomics in the 1990s, numerous novel genes have been discovered [124-126]. Metagenomics allows for the cloning and expression or screening of multiple genomic DNA extracts from any given sample [127]. Accessing the gene or enzyme of interest is, however, dependent on various factors, but typically involve sequence-based screening and/or function-based screening, some of which are also applicable to isolated strains.
3.2.1. Sequence-based screening
Sequence-based screening relies heavily on prior knowledge of the enzyme of interest, e.g. the polymerase chain reaction (PCR) primers and hybridisation probes need to be designed based on conserved regions in sequences that are currently available in databases. This approach is therefore not suited for the discovery of novel protein classes [128]. In two metagenomic studies, a sequence-based screening approach allowed researchers to identify the presence of actinobacterial two-domain laccases in lignin-rich environments. Ausec et al. [129] made use of previously published primer sets, while Lu et al. [130] designed their own primers that were designed based on sequence alignments of known two-domain laccases. Both of these studies showed the vast biodiversity of these laccases in the respective environments analysed: drained peat soils [129] and compost prepared from agricultural waste [130].
Similarly, sequence-based screening has been shown to be a valuable tool for determining the diversity of
3.2.1.1. In silico screening of genome sequences
Of the nearly 6,400 bacterial genomes available on the NCBI ‘Microbial genomes’ page, (http://www.ncbi.nlm.nih.gov/genomes/MICROBES/microbial_taxtree.html; accessed 24 June 2015), 871 represent the class
In addition, access to various genomes allows for comparative genomic analysis of the diversity of specific enzyme groups/classes among the genomes analysed. For example, Li et al. [137] performed comparative genomic analyses on 18
3.2.1.2. Reverse genetics
Recently, a strategy known as ‘reverse genetics’ has been developed for the identification of numerous gene clusters [138]. The first step in this strategy entails the deduction of a biosynthesis hypothesis for the corresponding substance. On the basis of the chemical structures, radiolabelling studies and other available data about the substances, the enzyme systems required for backbone-synthesis and tailoring enzymes can be predicted. In the second step, characteristic target genes and enzymes are selected and compared in multiple sequence alignments with homologous genes and gene products retrieved from public and in-house databases. Subsequently, conserved protein motifs can be determined and used for primer design. In the last step, internal fragments of the selected biosynthesis genes can be amplified by PCR and used as probes to identify these genes in genome libraries [138]. This method has been successfully used in the identification of genes from
3.2.2. Function-based screening
Function-based screening is often preferred to sequence-based screening because it allows for access to a wider range of enzymes or biological activities [128, 141]. In metagenomics, the success of function-based screening is dependent on various factors: (1) abundance of the target gene; (2) the size of the gene; (3) the presence of a full-length sequence; (4) the expression host; (5) the expression system and (6) the assay or means of detection of the enzyme activity [123]. In the majority of metagenomic studies,
There are three main approaches in function-based screening: the detection of a phenotypic trait (most commonly used), production of the enzyme or compound due to substrate/product/metabolite-induced gene expression (SIGEX/PIGEX/METREX) and modulated detection where the expressed product is linked to a reporter gene allowing for detection by fluorescence or luminescence [142]. In the next few paragraphs, methods used for the detection of oxidases in actinobacteria are described. For more detail on the different function-based screening approaches mentioned above, see Ekkers et al. [142] for a complete review.
3.2.2.1. Selective isolation of oxidase-producing strains directly from an environmental sample
Kiiskinen
Isolated strains can also be screened for enzyme activity on solid media containing dyes. McMullan et al. [145] gives a short review on published work regarding the degradation of dyes by filamentous actinomycetes. The extracellular enzymes that are involved in the degradation of dyes are typically the enzymes involved in lignin degradation: lignin peroxidase, laccase and manganese-dependent peroxidase [146-147]. In filamentous actinomycetes, however, it was found that there is no correlation between the degradation of the polymeric dye, Poly R478, and that the enzymatic process involved is still unexplained [145]. The degradation of the azo dye, Remazol Brilliant Blue R (RBBR), was, however, found to be linked to the action of peroxidases which are similar in structure and function to the fungal manganese-dependent peroxidases [145, 147].
3.2.2.2. High-throughput screening with liquid-based enzyme assays
Oxidoreductases produced by actinobacteria can also be accessed by using various high-throughput assays. With the increased interest in finding novel enzymes for use in biocatalysis, the number of assays currently available is quite vast and most are based on the use of chromogenic or fluorogenic substrates [148]. Most of these assays employ the use of a standard UV/visible spectrophotometer or microtiter plate readers, but for certain assays, the screening process involves the use of mass spectrometry, nuclear magnetic resonance (NMR), Fourier-transform infrared (FT-IR), thin layer chromatography (TLC), capillary array electrophoresis and enzyme-linked immunosorbent assays (ELISAs) to detect the changes in the catalysed reaction [148-149]. Enzyme fingerprinting and the use of microarrays are increasingly becoming powerful tools in the high-throughput screening of enzymes [148]. In addition, fluorescence-activated cell sorting (FACS) is a powerful high-throughput screening method that allows for the screening of large (109) clone libraries in a relatively short time period [123]. Zhu and Fang [150] recently also reviewed the potential of droplet microfluidics as a high-throughput technique for the screening of enzyme activities.
3.2.2.3. Protein engineering
Protein engineering is an alternative approach to obtain an enzyme with novel activities and biochemical properties. Random mutagenesis through the use of UV radiation or chemical mutagenesis is often favoured when the enzyme structure or sequence is not known. Even though this approach is typically used to generate an enzyme with improved properties, it is often limited to a change in a single property [123]. Random mutagenesis, however, was successfully used by Fujii et al. [151] to improve the vitamin D3 hydroxylase activity of
For more complicated changes, such as changes to specific amino acids, knowledge is required on sequence-structure information, allowing for a more rational or semi-rational design. Liu et al. [153] made use of a combination of site-directed mutagenesis and error-prone PCR to generate 7,800 variants of a cytochrome P450 monooxygenase from
The de novo synthesis of enzymes allows for the design of enzymes with specific properties. This, however, requires a deep understanding of sequence-structure-function [160]. A similar approach is the production of chimeric proteins, where different enzymes are combined so that the properties from the different enzymes can be harnessed in one reaction setup. These fusion enzymes are designed to enhance the biocatalytic activity/function of the enzymes. They can consist of enzymes that act synergistically (e.g. enzymes involved in lignin degradation) [161] or as in the case of the BVMO from
4. In conclusion
Members of the order
References
- 1.
Stackebrandt E, Schumann P. Introduction to the taxonomy of actinobacteria. In: Dworkin M, Falkow S, Rosenberg E, Schleifer K-H, Stackebrandt E. (eds.) The Prokaryotes, Volume 3: Archaea. Bacteria: Firmicutes, Actinomycetes Springer Science+Business Media, New York, USA, 2006. pp. 297-321. - 2.
Torres E, Bustos-Jaimes I, Le Borgne S. Potential use of oxidative enzymes for the detoxification of organic pollutants. Appl Catalysis B: Environ 2003;46:1-15. - 3.
Stackebrandt E, Rainey FA, Ward-Rainey NL. Proposal for a new hierarchic classification system, Actinobacteria classis nov. Int J Systematic Bacteriol 1997;47:479-91. - 4.
May SW. Applications of oxidoreductases. Biotechnology 1999;10:370-5. - 5.
Conrad LS, Sponholz WR, Berker O. Treatment of cork with a phenol oxidizing enzyme. US patent US6152966. 2000. - 6.
Bajpai P, Anand A, Bajpai PK. Bleaching with lignin-oxidizing enzymes. Biotechnol Annu Rev 2006;12:349-78. - 7.
López MJ, Guisado G, Vargas-García MC, Suárez-Estrella F, Moreno J. Decolourization of industrial dyes by lignolytic microorganisms isolated from composting environment. Enzyme Microbial Technol 2006;40:42-5. - 8.
Setti L, Giuliani S, Spinozzi G, Pitteri PG. Laccase catalyzed-oxidative coupling of 3-methyl 2-benzothiazolinone hydrazone and methoxyphenols. Enzyme Microbial Technol 1999;25:285-9. - 9.
Carredero V, Mena ML, Gonzalez-Cortés A, Yáñez-Sedeño P, Pingarrón JM. Development of a high analytical performance tyrosinase biosensor based on a composite graphite-Teflon electrode modified with gold nanoparticles. Biosensors Bioelectron 2006;22:730-6. - 10.
Torriero AAJ, Salinas E, Marchevsky EJ, Raba J, Silber JJ. Penicillamine determination using a tyrosinase micro-rotating biosensor. Analy Chim Acta 2006;580:136-42. - 11.
Luppa PB, Sokoll LJ, Chan DW. Immunosensors - principles and applications to clinical chemistry. Clinic Chim Acta 2001;314:1-26. - 12.
Couto SR, Herrera JLT. Industrial and biotechnological applications of laccases: a review. Biotechnol Adv 2006;24:500-13. - 13.
Sojo M, Bru R, Lopez-Molina D, Garcia-Carmona F, Argüelles JC. Cell-linked and extracellular cholesterol oxidase activities from Rhodococcus erythropolis. Isolation and physiological characterization. Appl Microbiol Biotechnol 1997;47:583-9. - 14.
Donova MV. Transformation of steroids by actinobacteria: a review. Appl Biochem Microbiol 2007;43:1-14. - 15.
Richmond W. Preparation and properties of a cholesterol oxidase from Nocardia sp. and its application to the enzymatic assay of total cholesterol in serum. Clin Chem 1973;19:1350-6. - 16.
Allain CC, Poon LS, Chan CSG, Richmond W, Fu PC. Enzymatic determination of total serum cholesterol. Clin Chem 1974;20:470-5. - 17.
Pollegioni L, Piubelli L, Molla G. Cholesterol oxidase: biotechnological applications. FEBS J 2009;276:6857-70. - 18.
Yazdi MT, Zahraei M, Aghaepour K, Kamranpour N. Purification and partial characterization of a cholesterol oxidase from Streptomyces fradiae . Enzyme Microbial Technol 2001;28:410-4. - 19.
Kreit J, Sampson NS. Cholesterol oxidase: physiological functions. FEBS J 2009;276:6844-56. - 20.
MacLachlan J, Wotherspoon ATL, Ansell RO, Brooks CJW. Cholesterol oxidase: sources, physical properties and analytical applications. J Steroid Biochem Molecul Biol 2000;72:169-95. - 21.
Ivshina IB, Grishko VV, Nogovitsina EM, Kukina TP, Tolstikov GA. Bioconversion of β-sitosterol and its esters by actinobacteria of the genus Rhodococcus . Appl Biochem Microbiol 2005; 41:551-7. - 22.
Kumari L, Kanwar SS. Cholesterol oxidase and its applications. Adv Microbiol 2012;2:49-65. - 23.
Brzostek A, Dziadek B, Rumijowska-Galewicz A, Pawelczyk J, Dziadek J. Cholesterol oxidase is required for virulence of Mycobacterium tuberculosis . FEMS Microbiol Lett 2007;275:106-12. - 24.
Giardina P, Faraco V, Pezzella C, Piscitelli A, Vanhulle S, Sannia, G. Laccases: a never-ending story. Cell Molecul Life Sci 2010:67:369-85. - 25.
Riva S. Laccases: Blue enzymes for green chemistry. Trends Biotechnol 2006;24:219-26. - 26.
Fernandes TAR, da Silveira WB, Passos FML, Zucchi TD Laccases from Actinobacteria - what we have and what to expect. Adv Microbiol. 2014a:4:285-96. - 27.
Skálová T, Dohńalek J, Østergaard LH, Østergaard PR, Kolenko P, Dušková J, Štěpánková A, Hašek J. The structure of the small laccase from Streptomyces coelicolor reveals a link between laccases and nitrite reductases. J Molecul Biol 2009;385:1165-78. - 28.
Reiss R, Ihssen J, Thöny-Meyer L. Bacillus pumilus laccase: a heat stable enzyme with a wide substrate spectrum. BMC Biotechnol 2011;11:9. - 29.
Fernandes TAR, da Silveira WB, Passos FML, Zucchi, TD. Characterization of a thermotolerant laccase produced by Streptomyces sp. SB086. Annal Microbiol 2014b;64:1363-9. - 30.
Prins A. The site-directed mutagenesis of the small laccase from Streptomyces coelicolor to increase its redox potential. Bachelor of Science (Honours) Thesis, University of the Western Cape; 2012. - 31.
Machczynski MC, Vijgenboom E, Samyn B, Canters GW. Characterization of SLAC: a small laccase from Streptomyces coelicolor with unprecedented activity. Protein Sci 2004;13:2388-97. - 32.
Molina-Guijarro JM, Pérez J, Muñoz-Dorado J, Guillén F, Moya R, Hernández M, Arias ME. Detoxification of azo dyes by a novel pH-versatile, salt-resistant laccase from Streptomyces ipomoea . Int Microbiol 2009;12:13-21. - 33.
Gunne M, Urlacher VB. Characterization of the alkaline laccase Ssl1 from Streptomyces sviceus with unusual properties discovered by genome mining. PLoS ONE 2012;7:e52360. - 34.
Lu L, Zeng G, Fan C, Ren X, Wang C, Zhao Q, Zhang J, Chen M, Chen A, Jiang M. Characterization of a laccase-like multicopper oxidase from newly isolated Streptomyces sp. C1 in agricultural waste compost and enzymatic decolorization of azo dyes. Biochem Eng J 2013;72:70-6. - 35.
Gianfreda L, Xu F, Bollag, JM. Laccases: A useful group of oxidoreductive enzymes. Bioremed J 2009;3:1-26. - 36.
Kim YJ, Nicell JA. Impact of reaction conditions on the laccase-catalyzed conversion of bisphenol A. Biores Technol 2006;97:1431-42. - 37.
Alcade M. Laccases: Biological functions, molecular structure and industrial applications. In: Polaina, J, MacCabe AP. (eds.) Industrial Enzymes. Springer Netherlands, Netherlands, 2007. pp. 461-476. - 38.
Madhavi V, Lele SS. Laccase: properties and applications. Bioresources 2009;4:1694-717. - 39.
Strong PJ, Claus H. Laccase: A review of its past and its future in bioremediation. Crit Rev Environ Sci Technol 2011;41:373-434. - 40.
Margot J, Bennati-Granier C, Maillard J, Blánquez P, Barry DA, Holliger C. Bacterial versus fungal laccase: potential for micropollutant degradation. AMB Express 2013;3:63. - 41.
Van Bloois E, Torres-Pazmiño DE, Winter RT, Fraaije MW. A robust and extracellular heme-containing peroxidase from Thermobifida fusca as prototype of a bacterial peroxidase superfamily. Appl Microbiol Biotechnol 2010;86:1419-30. - 42.
Le Roes-Hill M, Khan N, Burton SG. Actinobacterial peroxidases: an unexplored resource for biocatalysis. Appl Biochem Biotechnol 2011a;164:681-713. - 43.
Mercer DK, Iqbal M, Miller PGG, McCarthy, AJ. Screening actinomycetes for extracellular peroxidase activity. Appl Environ Microbiol 1996;62:2186-90. - 44.
Tuncer M, Ball AS, Rob A, Wilson MT. Optimisation of extracellular lignocellulolytic enzyme production by a thermophilic actinomycete Thermomonospora fusca BD25. Enzyme Microbial Technol 1999;25:38-47. - 45.
Antonopoulos VT, Hernandez M, Arias ME, Mavrakos E, Ball AS. The use of extracellular enzymes from Streptomyces albus ATCC 3005 for the bleaching of eucalyptus kraft pulp. Appl Microbiol Biotechnol 2001;57:92-7. - 46.
Jaouadi B, Rekik H, Badis A, Jaouadi NZ, Belhoul M, Hmidi M, Kourdali S, Fodil D, Bejar S. Production, purification, and characterization of a highly thermostable and humic acid biodegrading peroxidase from a decolorizing Streptomyces albidoflavus strain TN644 isolated from a Tunisian off-shore oil field. Int Biodeterior Biodegrad 2014;90:36-44. - 47.
Nair CI, Jayachandran K, Shashidhar S. Biodegradation of phenol. Afr J Biotechnol 2006;25:4951-8. - 48.
El Azhari N, Devers-Lamrani M, Chatagnier G, Rouard N, Martin-Laurent F. Molecular analysis of the catechol-degrading bacterial community in a coal wasteland heavily contaminated with PAHs. J Hazard Mater 2010;177:593-601. - 49.
Harwood CS, Parales RE. The beta-ketoadipate pathway and the biology of self-identity. Annu Rev Microbiol 1995;50:553-90. - 50.
Hamzah RY, al-Bahama BS. Catechol ring-cleavage in Pseudomonas cepacia : the simultaneous induction ofortho andmeta pathways. Appl Microbiol Biotechnol 1994;41:250-6. - 51.
Shen F-T, Lin J-L, Huang C-C, Ho Y-N, Arun AB, Young LS, Young, C-C. Molecular detection and phylogenetic analysis of the catechol 1,2-dioxygenase gene from Gordonia spp. Systemat Appl Microbiol 2009;32:291-300. - 52.
Sutherland JB, Crawford DL, Pometto III AL. Catabolism of substituted benzoic acids by Streptomyces species. Appl Environ Microbiol 1981;41:442-8. - 53.
Nadaf NH, Ghosh JS. Purification and characterization of catechol 1,2-dioxygenase from Rhodococcus sp. NCIM 2891. Res J Environ Earth Sci 2011;3:608-13. - 54.
An H-R, Park H-J, Kim E-S. Cloning and expression of thermophilic catechol 1,2-dioxygenase gene ( catA ) fromStreptomyces setonii . FEMS Microbiol Lett 2001;195:17-22. - 55.
Kaminski U, Janke D, Prauser H, Fritsche W. Degradation of aniline and monochloroanilines by Rhodococcus sp. An 117 and a pseudomonad: A comparative study. Zeitschrift für Allgemeine Mikrobiologie 1983;23:235-46. - 56.
Field JA, Sierra-Alvarez R. Microbial degradation of chlorinated benzenes. Biodegradation 2008;19:463-80. - 57.
Silva AS, Camargo FADO, Andreazza R, Jacques RJS, Baldoni DB, Bento FM. Enzymatic activity of catechol 1,2-dioxygenase and catechol 2,3-dioxygenase produced by Gordonia polyisoprenivorans . Quimica Nova 2012;35:1587-92. - 58.
Roberts SM, Wan PWH. Enzyme-catalysed Baeyer-Villiger oxidations. J Molecul Catalysis B: Enzym 1998;4:111-36. - 59.
Kelly DR. Enantioselective Baeyer-Villiger reactions. Chimica Oggi 2000;18:333-7. - 60.
Flitsch S, Grogan G. Baeyer-Villiger monooxygenases. In: Drauz K, Waldmann H. (eds.) Enzyme catalysis in Organic Synthesis: A Comprehensive Handbook, Wiley-VCH, Weinheimm, 2002. pp. 1202-1245. - 61.
Mihovilovic MD, Müller B, Stanetty P. Monooxygenase-mediated Baeyer-Villiger oxidations. Eur J Org Chem 2002:3711-30. - 62.
Riebel A, Dudek HM, de Gonzalo G, Stepniak P, Rychlewski L, Fraaije MW. Expanding the set of rhodococcal Baeyer-Villiger monooxygenases by high-throughput cloning, expression and substrate screening. Appl Microbiol Biotechnol 2012;95:1479-89. - 63.
Torres-Pazmiño DE, Fraaije MW. Discovery, redesign and applications of Baeyer-Villiger monooxygenases. In: Matsuda T. (ed.) Future Directions in Biocatalysis 2007. pp. 107-128. - 64.
Torres-Pazmiño DE, Dudek HM, Fraaije MW. Baeyer-Villiger monooxygenases: recent advances and future challenges. Curr Opin Chem Biol 2010;14:138-44. - 65.
Fraaije MW, Kamerbeek NM, van Berkel WJH, Janssen DB. Identification of a Baeyer-Villiger monooxygenase sequence motif. FEBS Lett 2002;518:43-7. - 66.
Fraaije MW, Wu J, Heutes DPHM, van Hellemond EW, Spelberg JHL, Janssen, DB. Discovery of a thermostable Baeyer-Villiger monooxygenase by genome mining. Appl Microbiol Biotechnol 2005;66:393-400. - 67.
Jiang J, Tetzlaff CN, Takamatsu S, Iwatsuki M, Komatsu M, Ikeda H, Cane DE. Genome mining in Streptomyces avermitilis: a biochemical Baeyer-Villiger reaction and discovery of a new branch of the pentalenolactone family tree. Biochemistry 2009;48:6431-40. - 68.
Kotani T, Yurimoto H, Kato N, Sakai, Y. Novel acetone metabolism in a propane-utilizing bacterium Gordonia sp. strain TY-5. J Bacteriol 2007;189:886-93. - 69.
de Gonzalo G, Mihovilovic MD, Fraaije MW. Recent developments in the application of Baeyer-Villiger monooxygenases as biocatalysts. ChemBioChem 2010;11:2208-31. - 70.
Urlacher VB, Girhard M. Cytochrome P450 monooxygenases: an update on perspectives for synthetic application. Trends Biotechnol 2012;30:26-36. - 71.
Rendic S, DiCarlo FJ. Human cytochrome P450 enzymes: a status report summarizing their reactions, substrates, inducers, and inhibitors. Drug Metabolism Rev 1997;29:413-580. - 72.
Sasaki T. The 50th anniversary and new horizons of cytochrome P450 research: expanding knowledge on the multiplicity and versatility of P450 and its industrial applications: practical application of cytochrome P450. Biol Pharm Bull 2012;35:844-9. - 73.
Grogan G. Cytochromes P450: exploiting diversity and enabling application as biocatalysts. Curr Opin Chem Biol 2011;15:241-8. - 74.
Poupin P, Truffaut N, Combourieu B, Besse P, Sancelme M, Veschambre H., Delort AM. Degradation of Morpholine by an environmental Mycobacterium strain involves a cytochrome P-450. Appl Environ Microbiol 1998;64:159-65. - 75.
Lamb DC, Skaug T, Song H-L, Jackson CJ, Podust LM, Waterman MR, Kelly, DB, Kelly DE, Kelly SL. The cytochrome P450 complement (CYPome) of Streptomyces coelicolor A3(2). J Biol Chem 2002;277:24000-5. - 76.
Shresta P, Oh T-J, Liou K, Sohng JK. Cytochrome P450 (CYP105F2) from Streptomyces peucetius and its activity with oleandomycin. Appl Microbiol Biotechnol 2008; 79:555-62. - 77.
Schmid A, Hollmann F, Bühler B. Oxidation of phenols. In: Drauz K, Waldmann H. (eds.) Enzyme Catalysis in Organic Synthesis: A Comprehensive Handbook. Wiley-VCH, Weinhemm, 2002a. pp. 1170-93. - 78.
Claus H, Decker H. Bacterial tyrosinases. Systematic Appl Microbiol 2006;29:3-14. - 79.
Matoba Y, Kumagai T, Yamamoto A, Yoshitsu H, Sugiyama M. Crystallographic evidence that the dinuclear copper centre of tyrosinase is flexible during catalysis. J Biol Chem 2006;281:8981-90. - 80.
Manivasagan P, Venkatesan J, Sivakumar K, Kim S-K. Actinobacterial melanins: current status and perspective for the future. World J Microbiol Biotechnol 2013;29:1737-50. - 81.
Geng J, Yu S-B, Wan X, Wang X-J, Shen P, Zhou P, Chen X-D. Protective action of bacterial melanin against DNA damage in full UV spectrums by a sensitive plasmid-based noncellular system. J Biochem Biophys Methods 2008;70:1151-5. - 82.
Geng J, Yuan P, Shao C, Yu S-B, Zhou B, Zhou P, Chen, X-D. Bacterial melanin interacts with double-stranded DNA with high affinity and may inhibit cell metabolism in vivo. Arch Microbiol 2010;192:321-9. - 83.
Montefiori DC, Zhou J. Selective antiviral activity of synthetic soluble L-tyrosine and L-Dopa melanins against human immunodeficiency virus in vitro. Antiviral Res 1991;15:11-25. - 84.
Hung Y-C, Sava V, Hong M-Y, Huang GS. Inhibitory effects on phospholipase A2 and antivenin activity of melanin extracted from Thea sinensis Linn. Life Sci 2004;74:2037-47. - 85.
Goncalves RDCR, Pombeiro-Sponchiado SR. Antioxidant activity of the melanin pigment extracted from Aspergillus nidulans . Biol Pharm Bull 2005;28:1129-31. - 86.
Dadachova E, Bryan RA, Huang X, Moadel T, Schweitzer AD, Aisen P, Nosanchuk JD, Casadevall, A. Ionizing radiation changes the electronic properties of melanin and enhances the growth of melanized fungi. PLoS ONE 2007;2:e457. - 87.
Sambasiva Rao KRS, Mahalaxmi Y. Laccase-and peroxidase-free tyrosinase production by isolated microbial strain. J Microbiol Biotechnol 2012;22:207-14. - 88.
Kohasi PY, Kumagai T, Matoba Y, Yamamoto A, Maruyama M, Sugiyama M. An efficient method for the overexpression and purification of active tyrosinase from Streptomyces castaneoglobisporus . Protein Expression Purification 2004;34:202-7. - 89.
Della-Cioppa GR, Garger Jr. SJ, Holtz RB, McCulloch MJ, Sverlow GG. Method for making stable extracellular tyrosinase and synthesis of polyphenolic polymers therefrom. US PATENT 5801047 A, 1998. - 90.
Hirsch EC, Faucheux B, Damier P, Mouatt-Prigent A, Agid Y. Neuronal vulnerability in Parkinson’s disease. In: Riederer et al. (eds.) Advances in Research on Neurodegeneration. Springer-Verlag Wien, Vienna, 1987. pp. 80-88. - 91.
Fasano M, Giraudo S, Coha S, Bergamasco B, Lopiano L. Residual substantia nigra neuromelanin in Parkinson’s disease is cross-linked to α-synuclein. Neurochem Int 2003;42:603-6. - 92.
Madhusudhan DN, Mazhari BZ, Dastager SG, Agsar, D. Production and cytotoxicity of extracellular insoluble and droplets of soluble melanin by Streptomyces lusitanus DMZ-3. BioMed Res Int 2014, Article ID 306895, 11 pages, 2014. doi:10.1155/2014/306895 - 93.
Fernandez A, Picouet P, Lloret E. Cellulose-silver nanoparticle hybrid materials to control spoilage-related microflora in absorbent pads located in trays of fresh-cut melon. Int J Food Microbiol 2010;142:222-8. - 94.
Kiran GS, Dhasayan A, Lipton AN, Selvin J, Arasu MV, Al-Dhabi NA. Melanin-templated rapid synthesis of silver nanostructures. J Nanobiotechnol 2014;12:18. - 95.
Roy S, Das I, Munjal M, Karthik L, Kumar G, Kumar S, Rao KVB. Isolation and characterization of tyrosinase produced by marine actinobacteria and its application in the removal of phenol from aqueous environment. Frontiers Biol 2014;9:306-16. - 96.
Geueke B, Hummel W. A new bacterial L-amino acid oxidase with a broad substrate specificity: purification and characterization. Enzyme Microbial Technol 2002;31:77-87. - 97.
Shalaby AR. Significance of biogenic amines to food safety and human health. Food Res Int 1996;29:675-90. - 98.
Santos MHS. Biogenic amines: their importance in foods. Int J Food Microbiol 1996;29:213-31. - 99.
Agostinelli E, Arancia G, Vedova LD, Bell F, Marra M, Salvi M, Toninello A, The biological functions of polyamine oxidation products by amine oxidases: perspectives of clinical applications. Amino Acids 2004;27:347-58. - 100.
Adachi O, Yamada H, Ogata K. Purification and properties of putrescine oxidase of Micrococcus rubens . Agri Biol Chem 1966; 30:1202-10. - 101.
Van Hellemond EW, van Dijk M, Heuts DPHM, Janssen DB, Fraaije MW. Discovery and characterization of a putrescine oxidase from Rhodococcus erythropolis NCIMB 11540. Appl Microbiol Biotechnol 2008;78:455-63. - 102.
Dadakova E, Križek M, Pelikánová T. Determination of biogenic amines in foods using ultra-performance liquid chromatography. Food Chem 2009;116:365-70. - 103.
Bóka B, Adányi N, Szamos J, Virág D, Kiss A. Putrescine biosensor based on putrescine oxidase from Kocuria rosea. Enzyme Microbial Technol 2012;51:258-62. - 104.
Rochette J-F, Sacher E, Meunier M, Luong HT. A mediatorless biosensor for putrescine using multiwalled carbon nanotubes. Analyt Biochem 2005;336:305-11. - 105.
Schmid A, Hollmann F, Bühler, B. Oxidation of C-N bonds. In: Drauz K, Waldmann H. (eds.) Enzyme Catalysis in Organic Synthesis: A Comprehensive Handbook Wiley-VCH, Weinhemm, 2002b. pp. 1250-61. - 106.
Kamei T, Asano K, Suzuki H, Matsuzaki M, Nakamura S. L-glutamate oxidase from Streptomyces violascens . I Production, isolation and some properties. Chem Pharm Bull 1983;31:1307-14. - 107.
Patel RN. Enzymatic synthesis of chiral intermediates for Omapatrilat, an antihypertensive drug. Biomolecul Eng 2001a;17:167-82. - 108.
Patel RN. Biocatalytic synthesis of intermediates for the synthesis of chiral drug substances. Curr Opin Biotechnol 2001b;12:587-604. - 109.
Inouye Y, Nishimura M, Matsuda Y, Hosika H, Iwasaki H, Hujimura K, Asano K, Nakamura S. Purification and characterization of sarcosine oxidase of Streptomyces origin. Chem Pharm Bull 1987;35:4194-202. - 110.
Nishiya Y, Imanaka T. Cloning and sequencing of the sarcosine oxidase gene from Arthrobacter sp. TE1826. J Fermentation Bioeng 1993;75:239-44. - 111.
Meškys R, Rudomanskis R, Leipuviené R. Cloning of genes encoding heterotetrameric sarcosine oxidase from Arthrobacter sp. Biotechnol Lett 1997;18:781-6. - 112.
Harris RJ, Meškys R, Sutcliffe MJ, Scrutton NS. Kinetic studies of the mechanisms of carbon-hydrogen bond breakage by the heterotetrameric sarcosine oxidase of Athrobacter sp. 1-IN. Biochemistry 2000;39:1189-98. - 113.
Suzuki K, Ogishima M, Sugiyama M, Inouye Y, Nakamura S, Imamura S. Molecular cloning and expression of a Streptomyces sarcosine oxidase gene inStreptomyces lividans . Biosci Biotechnol Biochem 1992;56:432-6. - 114.
Chouaia B, Crotti E, Brusetti L, Daffonchio D, Essoussi I, Nouioui I, Sbissi I, Ghodbane-Gtari F , et al. Genome sequence ofBlastococcus saxobsidens DD2, a stone-inhabiting bacterium. J Bacteriol 2012;194:2752-3. - 115.
Normand P, Gury J, Pujic P, Chouaia B, Crotti E, Brusetti L, Daffonchio D, Vacherie B, et al . Genome sequence of radiation resistantModestobacter marinus strain BC501, a representative actinobacterium that thrives on calcareous stone surfaces. J Bacteriol 2012;194:4773-4. - 116.
Niewerth H, Schuldes J, Parschet K, Kiefer P, Vorholt JA, Daniel R, Fetzner S. Complete genome sequence and metabolic potential of the quinoldine-degrading bacterium Athrobacter sp. Rue61a. BMC Genomics 2012;13:534. - 117.
Burton S, Cowan DA, Woodley JM. The search for the ideal biocatalyst. Nat Biotechnol 2002;20:35-46. - 118.
Dordick J.S. An introduction to industrial biocatalysis. In : Dordick JS. (eds.) Biocatalysts for Industry, Springer Science+Business Media, New York, USA, 1991. pp. 3-20. - 119.
Bommarus AS, Polizzi KM. Novel biocatalysts: Recent developments. Chem Eng Sci 2006;61:1004-16. - 120.
Kurtböke Dİ. Biodiscovery from rare Actinomycetes: an ecotaxonomical perspective. Appl Microbiol Biotechnol 2012;93:1843-52. - 121.
Goodfellow M, Fiedler H-P. A guide to successful bioprospecting: informed by actinobacterial systematics. Antonie van Leeuwenhoek 2010;98:119-42. - 122.
Le Roes-Hill M, Rohland J, Burton SG. Actinobacteria isolated from termite guts as a source of novel oxidative enzymes. Antonie van Leeuwenhoek 2011b;100:589-605. - 123.
Van Rossum T, Kengen SWM, van der Oost J. Reporter-based screening and selection of enzymes. FEBS J 2013;280:2979-96. - 124.
Yun J, Ryu S. Review: Screening for novel enzymes from metagenome and SIGEX, as a way to improve it. Microbial Cell Factories 2005;4:8-12. - 125.
Cowan D, Meyer Q, Stafford W, Muyanga S, Cameron R, Wittwer P. Metagenomic gene discovery: past, present and future. Trends Biotechnol 2005;23:321-9. - 126.
Cowan DA. Microbial genomes - the untapped resource. Trends Biotechnol 2000;18:14-6. - 127.
Cowan DA, Arslanoglu A, Burton SG, Baker GC, Cameron RA, Smith JJ, Meyer Q. Metagenomics, gene discovery and the ideal biocatalyst. Biochem Soc Trans 2004;32:298-302. - 128.
Kennedy J, O’Leary ND, Kiran GS, Morrissey JP, O’Gara F, Selvin J, Dobson ADW. Functional metagenomic strategies for the discovery of novel enzymes and biosurfactants with biotechnological applications from marine ecosystems. J Appl Microbiol 2011;111:787-99. - 129.
Ausec L, van Elsas JD, Mandic-Mulec I. Two- and three-domain bacterial laccase-like genes are present in drained peat soils. Soil Biol Biochem 2011;43:975-83. - 130.
Lu L, Zeng G, Fan C, Zhang J, Chen A, Chen M, Jiang M, Yuan Y, Wu H, Lai M, He Y. Diversity of two-domain laccase-like multicopper oxidase genes in Streptomyces spp.: identification of genes potentially involved in extracellular activities and lignocellulose degradation during composting of agriculture waste. Appl Environ Microbiol 2014;80:3305-14. - 131.
Wright F, Bibb MJ. Codon usage in the G+C rich Streptomyces genome. Gene 1992;113:55565. - 132.
Decker H, Gaisser S, Pelzer S, Schneider P, Westrich L, Wohlleben W, Bechthold A. A general approach for cloning and characterizing dNDP-glucose dehydratase genes from actinomycetes. FEMS Microbiol Lett 1996;141:195-201. - 133.
Adrio JL, Demain AL Microbial enzymes: tools for biotechnological processes. Biomolecules 2014;4:117-39; doi:10.3390/biom4010117 - 134.
de Gonzalo G, Rodríguez C, Rioz-Martínez A, Gotor V. Improvement of the biocatalytic properties of one phenylacetone monooxygenase mutant in hydrophilic organic solvents. Enzyme Microbial Technol 2012;50:43-9. - 135.
Ricklefs E, Winkler N, Koschorreck K, Urlacher VB. Expanding the laccase-toolbox: a laccase from Corynebacterium glutamicum with phenol coupling and cuprous oxidase activity. J Biotechnol 2014;191:46-53. - 136.
Yao K, Wang F-O, Zhang H-C, Wei D-Z. Identification and engineering of cholesterol oxidases involved in the initial step of sterols catabolism in Mycobacterium neoaurum . Metabolic Eng 2013;15:75-87. - 137.
Li H-W, Zhi X-Y, Yao J-C, Zhou Y, Tang S-K, Klenk H-P, Zhao J, Li W-J. Comparative genomic analysis of the genus Nocardiopsis provides new insight into its genetic mechanisms of environmental adaptability. PLoS ONE 2013;8:e61528. DOI: 10.1371/journal.pone.0061528 - 138.
Weber T, Welzel K, Pelzer S, Vente A, Wohlleben W. Exploiting the genetic potential of polyketide producing streptomycetes. J Biotechnol 2003;106:221-32. - 139.
Chiu HT, Hubbard BK, Shah AN, Eide J, Fredenburg RA, Walsh CT, Khosla C. Molecular cloning and sequence analysis of the complestatin biosynthetic gene cluster. Proc Nat Acad Sci USA 2001;98:8548-53. - 140.
Piraee M, Vining LC. Use of degenerate primers and touchdown PCR to amplify a halogenase gene fragment from Streptomyces venezuelae ISP5230. J Ind Microbiol Biotechnol 2002;29:1-5. - 141.
McMahon MD, Guan C, Handelsman J, Thomas MG. Metagenomic analysis of Streptomyces lividans reveals host-dependent functional expression. Appl Environ Microbiol 2012;78:3622-9. - 142.
Ekkers DM, Cretoíu MS, Kielak AM, Van Elsas JD. The great screen anomaly - a new frontier in product discovery through functional metagenomics. Appl Microbiol Biotechnol 2012;93:1005-20. - 143.
Kiiskinen L-L, Rättö M, Kraus K. Screening for novel laccase-producing microbes. J Appl Microbiol 2004;97:640-6. - 144.
Bordeleau LM, Bartha R. Rapid technique for enumeration and isolation of peroxidase-producing microorganisms. Appl Microbiol 1969;18:274-5. - 145.
McMullan G, Meehan C, Connely A, Kirby N, Robinson T, Nigam P, Banat IM, Marchant R, Smyth WF. Microbial decolourisation and degradation of textile dyes. Appl Microbiol Biotechnol 2001;56:81-7. - 146.
Tekere M, Mswaka AY, Zvauya R, Read JS. Growth, dye degradation and ligninolytic activity studies on Zimbabwean white rot fungi. Enzyme Microbial Technol 2001;28:420-6. - 147.
Pasti-Grigsby MB, Paszczynski A, Goszczynski S, Crawford DL, Crawford RL. Use of dyes in assaying Phanerochaete chrysosporium Mn(II)-peroxidase and ligninase. Proceedings of the University of Idaho, Institute for Molecular and Agricultural Genetic Engineering 1. (http://www.uidaho.edu), 1994. - 148.
Goddard J, Reymond J. Enzyme assays for high-throughput screening. Curr Opin Biotechnol 2004;15:314-22. - 149.
Wahler D, Reymond J-L. Novel methods for biocatalyst screening. Curr Opin Chem Biol 2001;5:152-8. - 150.
Zhu Y, Fang Q. Analytical detection techniques for droplet microfluidics - a review. Analyt Chim Acta 2013;787:24-35. - 151.
Fujii Y, Kabumoto H, Nishimura K, Fujii T, Tanai S, Takeda K, Tamura N, Arisawa A, Tamura T. Purification, characterisation, and directed evolution study of a vitamin D3 hydroxylase from Pseudonocardia autotrophica . Biochem Biophys Res Commun 2009;385:170-5. - 152.
Dudek HM, Fink MJ, Shivange AV, Dennig A, Mihovilovic MD, Schwaneberg U, Fraaije MW. Extending the substrate scope of a Baeyer-Villiger monooxygenase by multiple-site directed mutagenesis. Appl Microbiol Biotechnol 2014;98:4009-20. - 153.
Liu L, Schmid RD, Urlacher VB. Engineering cytochrome P450 monooxygenase CYP 116B3 for high dealkylation activity. Biotechnol Lett 2010;32:841-5. - 154.
Li G, Ren J, Iwaki H, Zhang D, Hasegawa Y, Wu Q, Feng J, Lau PCK, Zhu D. Substrate profiling of cyclohexylamine oxidase and its mutants reveals new biocatalytic potential in deracemization of racemic amines. Appl Microbiol Biotechnol 2014;98:1681-9. - 155.
Gunne M, Höppner A, Hagedoorn P-L, Urlacher VB. Structural and redox properties of the small laccase Ssl1 from Streptomyces sviceus . FEBS J 2014;281:4307-18. - 156.
Sherif M, Waung D, Korbeci B, Mavisakalyan V, Flick R, Brown G, Abou-Zaid M, Yakunin AF, Master ER. Biochemical studies of the multicopper oxidase (small laccase) from Streptomyces coelicolor using bioactive phytochemicals and site-directed mutagenesis. Microbial Biotechnol 2013; doi: 10.1111/1751-7915.12068 - 157.
Toscano MD, De María L, Lobedanz S and Østergaard LH. Optimization of a small laccase by active site design. ChemBioChem 2013, doi: 10.1002/cbic.201300256 - 158.
Prins A, Kleinsmidt L, Khan N, Kirby B, Kudanga T, Vollmer J, Pleiss J, Burton S, Le Roes-Hill M. The effect of mutagenesis near the T1 copper site on the biochemical characteristics of the small laccase from Streptomyces coelicolor . Enzyme Microbial Technol 2015;68:23-32. - 159.
Jing G, Zhiming R, Taowei Y, Zaiwei M, Meijuan X, Xian Z, Shang-Tian Y. Enhancement of the thermostability of Streptomyces kathirae SC-1 tyrosinase by rational design and empirical mutation. Enzyme Microbial Technol 2015, http://dx.doi.org/10.1016/j.enzmictec.2015.06.002 - 160.
Davids T, Schmidt M, Böttcher D, Bornscheuer, UT. Strategies for the discovery and engineering of enzymes for biocatalysis. Curr Opin Chemi Biology. 2013;17:215-20. - 161.
Elleuche S. Bringing functions together with fusion enzymes - from nature’s inventions to biotechnological applications. Appl Microbiol Biotechnol 2014;99:1545-56; doi: 10.1007/s00253-014-6315-1 - 162.
Reetz MT, Wu S. Laboratory evolution of robust and enantioselective Baeyer-Villiger monooxygenases for asymmetric catalysis. J Am Chemical Soc 2009;131:15424-32.