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Peroxidase  was  extracted  from  cabbage  and  was  purified  in  three  different purification processes. It was first purified by ammonium sulphate precipitation and highest peroxidase activity was observed at 80% saturation.  Hence, 80% saturation was used to mass produce the enzyme. The enzyme was  again purified by dialysis which tends to remove salt as impurity from the  precipitated  enzyme. The enzyme was further purified by gel filtration which further removed salts and other proteins as impurities. The resulting enzyme was characterized to determine the optimum pH and temperature. The optimum pH and temperature were respectively 5.0 and 45oC. The Km and Vmax obtained from Lineweaver-Burk plot of initial velocities at different concentration of H2O2  were found to be 3.68mM and 37.04U/ml respectively. Also, Km  and  Vmax  of  o-dianisidine   were   found  to   be   9.89mM   and   28.57U/ml respectively.  The  enzymatic  activity  of  this  cabbage  peroxidase  with  hydrogen peroxide on synthetic dyes was investigated and was found to be very effective in the treatment  and  decolorization  of these  dyes.  This  partially  purified  enzyme  could decolorize many synthetic dyes; Azo Brilliant Black, Azo Trypan Blue, Azo Blue 5, Azo Citrus Red 2, Azo Yellow 6, Azo Pink, Azo Purple, Vat Green 11 and  Vat Orange 9. Azo Trypan Blue and Vat Orange 11 had the highest and least percentage decolorization  of  88.62  and  12%  respectively  after  contact  time  of  1  hour.  The cabbage peroxidase was found to decolorize Azo dyes more and  had little effect on Vat dyes. This peroxidase  could be an important  source for  dye and waste water decolorization.



Large  amounts  of  chemically  different  dyes  are  used  for  various  industrial applications such as textile dyeing, paper and pulp, leather and plastics (Park et al.,

2007). Textile dyes represent a major class of organic pollutants that are found in the waste  effluent  discharged  by  these  different  industries  (Kalsoom  et  al.,  2013). Approximately  20% of the dye  load  is lost in  the dyeing  residues  during  textile processing which represents one of the greatest environmental problems faced by the sector (Guarantini and Zanoni, 2000). These dyes are designed to be resistant to light, water  and  oxidizing  agents  and  are  therefore  the  most  problematic  groups  of pollutants,  considered  as xenobiotics  that are  not easily biodegradable  (Ong et al.,

2011). The dye effluent contains chemicals that are toxic, carcinogenic, mutagenic, or teratogenic to various aquatic species and humans (Celebi et al., 2012). Among the textile dyes, azo dyes account for 60-70% of all textile dyestuffs used and show the largest spectrum of colours (Bae and Freeman, 2007). They are the most common group of synthetic colorants released into the environment (Saratale et al., 2011). The discharge of azo dyes into water bodies presents  human and ecological risks, since both the original dyes and their  biotransformation  products can show toxic effects, mainly causing DNA damage. Therefore, the development of non-genotoxic dyes and investment in research to find effective treatments for effluents and drinking water is required, in order to  avoid environmental and human exposure to these compounds and prevent the deleterious effects they can have on humans and aquatic organisms.

The treatment of dye wastewater involves chemical and physical methods such as adsorption, coagulation, oxidation, filtration and ionizing radiation. All these methods have different  decolorization  capabilities,  operating speed  and proven  to be costly while  producing  large  amounts  of  sludge  (Leelakriangsak  and   Borisut,  2012). Biological processes have received increasing interest as a viable alternative owing to their cost effectiveness, ability to produce less sludge and environmental friendliness (Banat et al., 1996). However, synthetic dyes containing various substituents such as

nitro  and  sulfonic  groups  are  not  uniformly  susceptible  to  bio-decolorization  in conventional aerobic processes. Enzymatic approach has gained considerable interest in  the  decolorization/degradation  of  textile  and  other  industrially  important  dyes present  in  wastewater.  This  strategy  is  ecofriendly  and  useful  in  comparison  to conventional  chemical,  physical  and  biological  treatments,  which  have  inherent serious   limitations.   Stability,   activity   and   specificity   of   an   enzyme   are   the fundamental  parameters  that  control  the  development  of an  industrial  application (Torres and Ayala, 2010).

Many studies  have demonstrated  that fungi are  able to degrade  dyes and  this capability  to  degrade  dye  is  due  to  the  extracellular,   non-specific   and  non- stereoselective enzyme system (Bezalel et al., 1997). Peroxidases have been reported as  excellent  oxidant  agents  to  degrade  dyes  (Kirby  et al.,  1995).  Husain  (2010) reported  that  many  aromatic  dyes  could  be  decolorized  by  peroxidase  through precipitation or breaking of the aromatic ring structure. Several bacterial, fungal and plant peroxidases have been used for decolorization of synthetic textile dyes. Fungal extracted peroxidases have been mostly studied in dye removal processes (Novotny et al., 2001). Decolorization of different azo dyes by Phanerochaete chrysosporium RP

78  under  optimized  conditions  was  studied  by  reaction  mechanism  via  azo  dye (Ghasemi et al., 2010). Bacterial lignin peroxidases from Pseudomonas aeruginosa and Serratia marcescens have been shown to give 50% to 58% decolourization effect on textile dye-based effluent (Bholay et al., 2012). However, using peroxidases from microorganisms  to  decolorize  dyes  involves  high  cost  and  therefore  alternative sources such as plants are now considered (Chanwun et al., 2013). Among the plant peroxidases, the most studied are native or recombinant horseradish peroxidases, HRP (Shrivastav, 2003 and Tiirola et al., 2006). HRP has been shown to have the ability to precipitate and degrade aromatic azo compounds in the presence of H2O2 (Bhunia et al,  2001).  It  has  been   utilized   for  the  removal  of  halogenated   phenols  and pentachlorophenol (Meizler et al., 2011; Li et al., 2011). Plant peroxidases have been extracted from African oil bean seeds, sorghum, tea leaf, wheat germ, green pea and papaya fruit oil (Lee and Klein, 1990; Silva et al., 1990; Converso and Fernandez,

1995; Kvaratskhelia  et al., 1997; Eze et al., 2000; Eze, 2012;). Other  peroxidases, such  as  peroxidases  from  Allium  sativum,  Ipomoea  batatas,  Raphanus  sativus, Sorghum bicolor and soybean peroxidase have also been applied to phenol removal

(Al-Ansari et al., 2010 and Diao et al., 2011). Peroxidase has been extracted from red cabbage as reported by Ghahfarrokhi et al. (2013) but peroxidase from green cabbage is   poorly   studied.   This   research   is   therefore   focused   on   the    extraction, characterization, purification of peroxidase from green cabbage and its application on decolorization of industrial synthetic dyes.

1.1 Peroxidase

The name peroxidase was first used by Linossier,  who isolated  it from pus in

1898. They are one of the most extensively studied groups of enzymes (Azevedo et al.,  2003).     They  are  widely  distributed   in  nature  and  are  found  in   plants, microorganisms and animals where they catalyze the reduction of hydrogen peroxide (H2O2) to water (Bania and Mahanta, 2012). They use various peroxides (ROOH) as electron acceptors to catalyze a number of oxidative reactions. In mammals, they are implicated   in  biological   processes  as  various   as  immune  system  or  hormone regulations.  In plants,  they are  involved  in  auxin metabolism,  lignin  and  suberin formation, cross-linking of cell wall  components, defense against pathogens or cell elongation. They also show bad effect on the quality of vegetables during post-harvest senescence,  oxidation  of  phenolic  substances,  starch-sugar  conversion  and  post- harvest demethylation of pectic substances leading to softening of plant tissues during ripening  (Ghahfarrokhi  et  al.,  2013).  Humans  contain  more  than  30  peroxidases whereas  Arabidopsis  thaliana  has  about  130  peroxidases  that  are  grouped  in  13 different families and nine subfamilies (Koua et al., 2009). Peroxidase families from prokaryotic  organisms,  protists  and  fungi  have  been  shown  to  promote  virulence (Brenot et al., 2004; Missall et al., 2005 and Pineyro et al.,  2008). Commercially, peroxidases  find  application  in  biotransformations,  bioremediation,  in  Analytical Biochemistry  and  as  specific  reagents  such  as  bleaching  agents.  Peroxidases  are classified  as haem peroxidases  and non-haem  peroxidases  and distributed  between thirteen superfamilies and fifty subfamilies (Passardi et al., 2007).

1.1.1         Enzyme Commission Classification of peroxidase

Peroxidases   can  be  found   under   the   same   enzyme   classification   number EC.1.11.1.x,  donor:  hydrogenperoxide  oxidoreductase  (Fleischmann  et  al.,  2004). Currently,  15  different  EC  numbers  have  been  ascribed  to  peroxidase,  from  EC to EC, excluding EC (Passardi et al., 2007). Due to the presence of dual enzymatic domains, other peroxidase families were classified with

the following numbers: EC, EC, EC and EC To date, certain peroxidases do not possess an EC number and can only be classified in EC  Two  particular  cases  are  also  observed  for  EC  (NADPH peroxidase)  and EC (fatty acid peroxidase).  NADPH  peroxidase  activities have been observed  in different  studies  (Hochman  and Goldberg,  1991). However there is no known peroxidase  sequence that has been assigned to this EC number, probably due to the fact that none of the peroxidases known so far have a predominant NADPH    peroxidase    activity.    Peroxidasins,    peroxinectins,    other    non-animal peroxidases,  dyptype  peroxidases,  hybrid  ascorbate  cytochrome  c  peroxidase  and other  class  II peroxidases do not possess an EC number. The two independent EC numbers ( and both correspond to glutathione peroxidase and are based on the electron acceptor (hydrogen peroxide or lipid peroxide, respectively).

Table 1: The International Union of Biochemistry classification of peroxidases

EC numberRecommended nameAbbreviation in PeroxiBase
EC peroxidaseNadprx
  EC  NADPH peroxidase  No sequence available
  EC  Fatty acid peroxidase  No sequence available
  EC  Cytochrome C peroxidase  CcP, DiHCcP
  EC  Catalase  Kat, Cp
  EC  Peroxidase  POX
  EC  Iodide peroxidase  TPO
  EC  Glutathione peroxidase  GPx
  EC  Chloride peroxidase  Halprx, HalNprx, HalVprx
  EC  1-ascorbate Superoxide  APX
  EC  Phospholipidhydroperoxi  GPX
de glutathione peroxidase
  EC  Manganese peroxidase  MnP
  EC  Lignin peroxidase  Lip
  EC  Versatile peroxidase  VP
  EC  Linoleate diol synthase  LDS
  EC  Prostaglandinendoperoxi  PGHS
de synthase

EC                NAD(P)H oxidase                      DuOx

EC              4-carboxymuconolactone


AhpD, CMD, CMDn, HCMD,HCMDn, DCMD, DCMDn, Alkyprx, Alkyprxn

(Feischman et al., 2004).

1.1.2         Haem-Based and non-Haem based Classification

An important number of haem and non-haem peroxidase sequences are annotated and  classified  in the  peroxidase  database,  PeroxiBase.  PeroxiBase  contains  about

5800 peroxidase sequences classified as haem peroxidases and non-haem peroxidases and distributed between thirteen superfamilies and fifty subfamilies, (Passardi et al.,

2007). Haem and non-haem peroxidases are found in all kingdoms.

Figure 1: Schematic representation of the phylogenic relationships between the different protein classes and families found in PeroxiBase (Koua et al., 2009).       Haem based peroxidase

Haem peroxidase is found in plants, animals and microorganisms.  They contain ferriprotoporphyrin  IX (haematin  or  haem)  as a prosthetic  group  (Rodrigo  et al.,

1996). Out of 6,861 known peroxidase sequences collected in PeroxiBase, more than

73% of them code for haem-containing peroxidases. In the majority of cases, haem b is the prosthetic group and its evolutionary highly conserved amino acid surroundings influence its reactivity (Torres and Ayala, 2010). Haem peroxidases tend to promote rather than inhibit oxidative damage. Genes encoding haem peroxidases can be found in almost all kingdoms of life. They are grouped in  two major superfamilies:  one mainly found in bacteria, fungi and plants, Passardi et al. (2007) and a second mainly found in animals, fungi and bacteria (Daiyasu and Toh, 2000 and Furtmuller et al.,

2006). Members of the superfamily of plant/fungal/bacterial peroxidases (non-animal

peroxidases)  have  been  identified  in the  majority  of  the  living  organisms  except animals.  The  second  superfamily  described  as “animal  peroxidases”  comprises  a

group  of  homologous  proteins  mainly  found  in  animals.  The  mammalian  haem peroxidase  plays  a major  role in both disease  prevention  and  human  pathologies (Koua et al., 2009). Some mammalian haem peroxidases use H2O2  to generate more aggressive oxidants to fight intruding microorganisms (Flohe and Ursini, 2008).

In addition to these two large superfamilies, smaller protein families are classified as capable to reduce peroxide molecules. Examples are Catalase (Kat) that can also oxidize  hydrogen peroxide,  dihaem  cytochrome  C peroxidases  (DiHCcP),  dyptype peroxidases (DypPrx), haloperoxidases with (HalPrx) or without (HalNPrx, HalVPrx) haem. Non haem peroxidase

Non-haem peroxidases  are not evolutionarily  linked and form five  independent superfamilies.   These   are   alkylhydroperoxidase,    NADH   peroxidase   (NadPrx), manganese  catalases  (MnCat)  and  thiol  peroxidases.  The  largest  one  is the  thiol peroxidase,  which  currently  contains  more  than  1000  members  grouped  in  two different subfamilies (Glutathione peroxidases and Peroxiredoxines).

1.1.3. Plant Peroxidases

Plant Peroxidases (PODs) are haem peroxidases. In the presence of peroxide, they oxidize  a  wide  range  of  phenolic  compounds,  such  as  guaiacol,  o-dianisidine, pyrogallol,  chlorogenic  acid, catechin,  and catechol (Onsa et al.,  2004).  They are divided  into  three  classes  based  on  their  structural  and  catalytic  properties.  The overall primary sequences and the 3-dimentional structure of these three peroxidases are quite different, implying that these subfamily genes evolve from distinct ancestral genes (Taurog, 1999). The amino  acid sequences were found to be highly variable among the members of the plant peroxidase superfamily with less than 20% identity in the most divergent cases (Hiraga et al., 2001).

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