Hamada Abdel-Sattar Ahmed Metwally Abou-Bakr ..... tannic acid concentration and nitrogen source concentration at the central ...... Dr. Eman Fathy Sharaf.
! " # # ! $ # ! " %& '!(!(!)*+ #
! " ! ,
! ! " !
!%)*(*+ - !. "!/ !-! 0 1 !, 1 2# ! ! %)*(*+! 0 !&
3/! !
!
! !"
##$
% & ' ( % " ) "*
! "
# $ !% ! & $ ' ' ( $ ' # % % ) %* %' $
' + " %
& ' ! #
$, ( $ - $$$ . /"/"#012" )*3- +) * !4 !& 5!6%55787& % 2 9:;567!%2 ?9:;567!; 0 @ ! " A" ?B A%8>7> * C & D +,-./ 0 /1/1 -203 4 & " 5 )* A 2 ! "A ( % 2
% % % % $ $ $A A
AA - $$$ /"/"#012" )*3- +) * !4 !& 5!6%55787& %) A 9:;567!%E?9:;567!; 0 @ ! B & " B + AC
D +,-./ 0 /1/1 -203 4 - A F8>78A /"/"#012" )*3- +)
"
& 8>78
ISOLATION AND SCREENING OF TANNASE- PRODUCING FUNGI AND OPTIMIZING THE ENZYME PRODUCTION BY THE PROMISING ISOLATE
A Thesis Presented to the Graduate School Faculty of Agriculture, Alexandria University In Partial fulfillment of the Requirements for the Degree Of
MASTER OF SCIENCE In
Food Science and Technology (Biotechnology)
By
Hamada Abdel-Sattar Ahmed Metwally Abou-Bakr
2010
ISOLATION AND SCREENING OF TANNASE- PRODUCING FUNGI AND OPTIMIZING THE ENZYME PRODUCTION BY THE PROMISING ISOLATE Presented by Hamada Abdel-Sattar Ahmed Metwally Abou-Bakr For the Degree of MASTER OF SCIENCE In Food Science and Technology (Biotechnology)
Examiners’ Committee
Approved
Prof. Dr. Ahmed Yousef Gibriel Professor “Emeritus” of Food Science, Food Science Department, Faculty of Agriculture, Ain Shams University.
Prof. Dr. Hassan Ali Ibrahim Siliha Professor of Food Science and Technology, and Dean of Faculty of Agriculture, Zagazeg University.
Prof. Dr. Malak Ahmed El-Sahn Professor of Food Science and Technology, Food Science and Technology Department, Faculty of Agriculture, Alexandria University.
Prof. Dr. Amr Abdel-Rahman El-Banna Professor “Emeritus” of Food Science and Technology, Food Science and Technology Department, Faculty of Agriculture, Alexandria University.
Date: 10/8/2010
Advisors’ Committee Prof. Dr. Malak Ahmed El-Sahn Professor of Food Science and Technology, Food Science and Technology Department, Faculty of Agriculture, Alexandria University.
Prof. Dr. Amr Abdel-Rahman El-Banna Professor “Emeritus” of Food Science and Technology, Food Science and Technology Department, Faculty of Agriculture, Alexandria University.
DEDICATION Tothe spirit of my dear father To my great mother To my beloved wife Walaa To my Children Haneen, El-Baraa & Omar
ACKNOWLEDGEMENT First, cordial thanks to ALLAH who enabled me to complete this work, solved all problems which faced me and gave me the opportunity to seek knowledge, which is one of the greatest works in our life, as our Prophet Muhammad (peace be upon him) told us.
I would like to express my sincere thank, deep appreciation and heartful gratitude to Prof. Dr. Amr Abdel-Rahman El-Banna, Professor of Food Science and Technology, Faculty of Agriculture, Alexandria University, for his moral support, constant assistance, useful advices, valuable criticism, encouragement , supervision and guidance throughout the course of the work as well as during the preparation of manuscript.
My deep appreciation, thanks and sincere gratitude are also due to Prof. Dr. Malak Ahmed El-Sahn Professor of Food Science and Technology, Faculty of Agriculture, Alexandria University, for her supervision, interest, guidance, useful advice, suggestion, and great encouragement throughout the course of the work as well as during the preparation of manuscript.
May Special appreciation is also to be attributed to Prof. Dr. Ahmed Rafik ElMahdy, Professor of Food Science and Technology, Faculty of Agriculture, Alexandria University, for his support, interesting and unlimited help, scientific consultations, laboratory facilities, great encouragement, useful advice and assistance.
I am grateful and indebted to Prof. Dr. Mamdoh El-Rouby, Professor of crop production and statistics, Faculty of Agriculture, Alexandria University, for his unlimited help, valuable consultations and guidance throughout the course of the work during designing the statistical experiments and the analysis of data.
Thanks to all members of the Department of Food Science and Technology, Faculty of Agriculture, Alexandria University, for their encouragement and help.
I also wish to express my deepest thanks to my family for their constant and unlimited support.
TABLE OF CONTENTS Page 1. INTRODUCTION................................................................................................... 2. AIM OF INVESTIGATION.................................................................................. 3. REVIEW OF LITERATURE................................................................................ 3.1. Action of tannase……………………………………………………………... 3.2. Methods of tannase determination………………………..…………………... 3.3. Microorganisms producing tannase………………………....………….…….. 3.3.1. Bacterial tannase……………………………………………………….... 3.3.2. Fungal tannase……………………………………………….………….. 3.4. Isolation and screening of microorganisms for tannase production…………... 3.4.1. Isolation and screening of tannase-producing bacteria………………….. 3.4.2. Isolation and screening of tannase-producing fungi…………….............. 3.5. Fermentation systems and production media for production of tannase from fungi…………………………………………………………………………… 3.6. Biosynthesis of tannase and regulation of its production……………………... 3.7. Factors affecting tannase production by fungi………………………………… 3.7.1. Medium constituents…………………………………………………….. 3.7.1.1.The effect of carbon source……………………………………… 3.7.1.2.The effect of nitrogen sources…………………………………… 3.7.2. Environmental conditions…………………….…………………………. 3.7.2.1.The effect of fermentation time…………………………………. 3.7.2.2.The effect of temperature……………………………………….. 3.7.2.3.The effect of initial pH of production medium…………………. 3.7.2.4.The effect of inoculum size……………………………………... 3.7.2.5.The effect of agitation…………………………………………… 3.7.2.6.Interactions between factors affecting tannase production……… 4. MATERIALS AND METHODS........................................................................... 4.1. Materials ……………………………………………………………………… 4.1.1. Chemicals ………………………………………………………………. 4.1.2. Media and media constituents …………………….……………………. 4.1.3. Solid supports for solid state fermentation (SSF)……………………….. 4.1.4. Sources for isolation of tannase-producing microorganisms……………. 4.1.5. Media ……………………………………………………………............ 4.1.5.1.Media for isolation and primary screening ……………………... 4.1.5.2.Medium for secondary and final screening of fungi..…………… 4.1.5.3.Media for tannase production …………….…………………...... 4.1.5.4.Media for maintenance of isolates ……………………………… 4.2. Methods ………………………………………………………………………. 4.2.1. Isolation of tannase-producing microorganisms ……………………….. 4.2.2. Purification of tannase-producing isolates ……………………………... 4.2.3. Maintenance of tannase-producing isolates …………………………….. 4.2.4. Screening and selection of tannase-producing fungal cultures …………. 4.2.4.1.Primary screening ……………………………………………..... 4.2.4.2.Secondary and final screening …………………………………..
I
1 5 7 7 9 12 12 16 17 17 17 18 26 27 27 27 30 32 32 35 35 38 39 39 41 41 41 41 41 42 42 42 42 45 45 46 46 46 46 47 47 47
Page 4.2.5. Inoculum preparation …………………………………………………… 47 4.2.6. Harvesting the enzyme …………………………………………………. 47 4.2.7. Biomass determination ………………………......................................... 48 4.2.8. Identification of the most promising fungal isolate …………………….. 48 4.2.9. Optimization of conditions controlling tannase production…………….. 48 4.2.9.1.Fermentation techniques ……………………………................... 49 4.2.9.1.1. Liquid surface fermentation (LSF) …………………............. 49 4.2.9.1.2. Submerged fermentation (SmF) ……………………………. 49 4.2.9.1.3. Solid state fermentation (SSF)………………………............. 49 4.2.9.2.Fermentation temperature and fermentation time ……………… 50 4.2.9.3.Concentrations of tannic acid and nitrogen source, initial pH and inoculum size ………..…………………………………………… 50 4.2.10. Analytical methods …………………………………………………..... 56 4.2.10.1. Estimation of tannic acid ……………………………………. 56 4.2.10.2. Enzyme assay ………………………………………………... 56 4.2.10.3. Calibration curve for gallic acid estimation……...................... 57 4.2.10.4. Determination of nitrogen source in the medium……………. 57 4.2.10.5. Measurements of pH ………………………………………… 57 4.2.10.6. Paper chromatographic analysis of fermented broth................. 57 5. RESULTS AND DISCUSSION…………………………………………………. 59 5.1. Isolation of tannase-producing microorganisms ………….…………………. 59 5.2. Characteristics of the obtained fungal isolates……………………………….. 61 5.3. Screening of tannase-producing microorganisms………………………......... 61 5.3.1. Primary screening of tannase producers………….……………..………. 65 5.3.1.1.Primary screening of fungal cultures……………………………. 65 5.3.1.2.Primary screening of bacterial cultures …………………………. 65 5.3.2. Secondary screening of selected fungal isolates…...…………..……….. 70 5.3.3. Final screening of promising fungal isolates …………………………… 73 5.4. Identification of the most promising tannase-producing fungal isolate…........ 73 5.5. Studying the nature of Aspergillus niger Van Tieghem tannase….………….. 83 5.6. Optimization of tannase production by Aspergillus niger Van Tieghem…….. 86 5.6.1. Effect of fermentation technique and agitation condition………………. 86 5.6.2. Effect of tannins source ………………………………………................ 90 5.6.3. Effect of addition carbohydrates incorporation with tannic acid……….. 94 5.6.4. Effect of glucose concentration as additive carbon source …………….. 98 5.6.5. Effect of nitrogen source type……………………....………….……….. 101 5.6.6. Effect of divalent cations……………………….……………..………… 106 5.6.7. Effect of fermentation temperature and fermentation time……………… 109 5.6.8. Effect of tannic acid and nitrogen source concentrations, initial pH and inoculum size on extracellular tannase production…….....………….. 119 5.7. Changes occurred during fermentation process of tannase production………. 135 6. SUMMARY……………………………………………………………………….. 141 7. LITERATURE CITED…………………………………………………………... 145
II
LIST OF TABLES Table
Title
Page
1.1
Enzyme markets based on application sectors ($ Million)………………….
2
3.1
Tannase -producing microorganisms in literature…………………………..
13
3.2
Fermentation systems and microbial tannase production media used for production of tannnase from fungi……………...…………………………...
19
3.3
The optimum fermentation time for tannase production by various fungi….
33
3.4
The optimum temperatures for tannase production by various fungi ...…….
36
4.1
Isolation sources of tannase-producing microorganisms……………………
43
4.2
Coded levels and combinations between two variables, fermentation temperature (T) and fermentation time (D) in split plot experiment…..........
51
Coded and actual levels of the variables in rotatable central composite design (RCCD)……………………………………………………………...
54
The full experimental plan with respect to their values in coded and actual forms…………………………………………………………..…………….
55
5.1
Sources, types and number of tannase-producing isolates………………….
60
5.2
Morphological characterization of fungal isolates…………………………..
62
5.3
List of the provided cultures…………………………………………...……
64
5.4
Diameters of fungal colonies and clear zones measured after 72 and 96 h of growth on TAA medium….…………………………………………………
66
Secondary screening of promising fungal isolates selected from the primary screening step ……………………………………………………...
71
5.6
Final screening of promising fungal isolates..………………………………
74
5.7
Cultural and morphological characteristics of the most promising tannasproducing fungus (isolate No. I5)..........................................................…….
76
5.8
Extracellular tannase production in the presence of tannic acid or glucose..
84
5.9
Effect of fermentation technique and agitation condition on intracellular, extracellular and total tannase production by Aspergillus niger Van Tieghem……………………………………………………………………..
87
4.3
4.4
5.5
III
Table
Title
Page
5.10
Tannins contents in natural tannins sources and calculated amounts needed to give 1% tannins in each flask………………………………….…………
91
Effect of tannins source on tannase production by Aspergillus niger Van Tieghem using SmF and SSF techniques………………...………………....
92
Effect of addition carbohydrates incorporation with tannic acid on tannase and biomass production by Aspergillus niger Van Tieghem………………..
95
Effect of glucose concentration as additive carbon source on tannase and biomass production by Aspergillus niger Van Tieghem………………...….
99
Nitrogen content of used nitrogen sources and required amounts of each source………………………...………………………………….…….…….
102
Effect of nitrogen source type on tannase and biomass production by Aspergillus niger Van Teighem…….……………………………………….
104
Effect of divalent cations on tannase and biomass production by Aspergillus niger Van Tieghem …………………………………………….
107
Effect of fermentation temperature and fermentation time on tannase production…………………………………………………………………...
110
Analysis of variance (ANOVA) for split plot experiment studying the effect of fermentation time and temperature on tannase production………..
111
5.19
The main effect of fermentation temperature on tannase roduction………..
113
5.20
The main effect of fermentation time on tannase production……………….
115
5.21
The effect of interaction between fermentation temperature and fermentation time levels on tannase production and comparison between means using Fisher LSD0.05 test…..………………………………………...
118
RCCD experimental layout and extracellular tannase yields (Y) obtained from each treatment against the predicted values…………………………...
120
5.23
Regression coefficients of the full second order polynomial model……….
121
5.24
Analysis of variance of RCCD for the fitted model of appropriate degree of multinomial………………………………………………………………….
122
5.11
5.12
5.13
5.14
5.15
5.16
5.17
5.18
5.22
IV
Table
Title
Page
5.25
Validation of reduced second order polynomial model (Equation 2) using different levels of tannic acid (X1), sodium nitrate as nitrogen source (X2), initial pH of the medium (X3) and inoculum size (X4) for tannase production…………………………………………………………………...
125
Changes in tannic acid concentration, nitrogen source concentration, fungal biomass, extracellular tannase activity and pH during fermentation process under optimized conditions………………………………………………….
136
5.26
V
LIST OF FIGURES Figure
Title
Page
1.1
Applications of microbial enzymes……………………………………….
3
3.1
Esterase and depsidase activities of tannase………………………………
8
3.2
Action of tannase on tannic acid…………………………………………..
9
3.3
Esertification of gallic acid in organic media using tannase……………...
10
4.1
Split plot design and random distribution of experimental units …………
52
5.1
Secondary screening of promising fungal isolates selected from the primary screening step…………………………………………………….
72
5.2
Final screening of promising fungal isolates……………………………...
5.3
Aspergillus niger Van Teighem grown on PDA medium at 30º C for 96 h…………………………………………………………………………...
77
Aspergillus niger Van Teighem colony surrounded by a hydrolytic clear zone………………………………………………………………………..
78
Microscopic photo showing the detailed structure of Aspergillus niger Van Teighem conidiophores………………………………………………
79
Microscopic photo of Aspergillus niger Van Teighem showing the arising condiophore from a thick walled foot cell………………………...
80
Microscopic photo of Aspergillus niger Van Teighem showing the shape and arrangement of conidia………………………………………………..
81
Microscopic photo of Aspergillus niger Van Teighem showing the septate hyphae……………………………………………………………..
82
5.9
Extracellular tannase production in the presence of tannic acid or glucose
85
5.10
Effect of fermentation technique and agitation condition on intracellular, extracellular and total tannase production………………………………...
88
Effect of tannins source on tannase production using SmF and SSF Techniques………………………………………………………………...
93
Effect of addition carbohydrates incorporation with tannic acid on tannase and biomass production by Aspergillus niger Van Tieghem……..
96
5.4
5.5
5.6
5.7
5.8
5.11
5.12
VI
75
Figure
Title
Page
5.13
Effect of glucose concentration as additive carbon source on tannase and biomass production by Aspergillus niger Van Tieghem………………….
100
5.14
Effect of nitrogen source type on tannase and biomass production by Aspergillus niger Van Teighem…………………………………………...
105
Effect of divalent cations on tannase and biomass production by Aspergillus niger Van Tieghem…………………………………………...
108
5.16
The main effect of fermentation temperature on tannase production……..
112
5.17
The main effect of fermentation time on tannase production……………..
114
5.18
The effect of interaction between fermentation temperature (T) and fermentation time (D) levels on tannase production ……………………...
117
Response surface and contour diagrams for tannase activity considering tannic acid concentration and nitrogen source concentration at the central levels of initial pH (5) and inoculums size (9×107 spores/flask)….……...
127
Response surface and contour diagrams for tannase activity considering tannic acid concentration and initial pH of the medium at the central levels of nitrogen source concentration (6 g/L) and inoculums size (9×107 spores/flask)………………………………………………………
129
Response surface and contour diagrams for tannase activity considering tannic acid concentration and inoculum size at the central levels of nitrogen source concentration (6 g/L) and medium initial pH (5)………...
130
Response surface and contour diagrams for tannase activity considering nitrogen source concentration and initial pH of the medium at the central levels of tannic acid concentration (1.5 %) and inoculums size (9×107 spores/flask)……………………………………………………………….
131
Response surface and contour diagrams for tannase activity considering nitrogen source concentration and inoculum size at the central levels of tannic acid concentration (1.5 %) and medium initial pH (5)…………….
133
Response surface and contour diagrams for tannase activity considering initial pH of the medium and nitrogen source concentration at the central levels of tannic acid concentration (1.5 %) and nitrogen source concentration (6 g/L)……………………………………………………...
134
5.15
5.19
5.20
5.21
5.22
5.23
5.24
VII
Figure
Title
Page
5.25
Changes in tannic acid concentration, nitrogen source concentration, fungal biomass, extracellular tannase activity and pH during fermentation process under optimized conditions………………………………………. 137
5.26
Paper chromatogram of hydrolytic product of tannic acid at different hours of fermentation process ……...…...………………………………... 138
VIII
1. INTRODUCTION Enzymes are highly efficient environment-friendly catalysts, synthesized by living systems. They have significant advantages over chemical catalysts, of which the most important are specificity, high catalytic activity, ability to work at moderate temperatures, and the ability to be produced in large amounts (Barredo, 2005). Enzymes production is a major and growing field of contemporary biotechnology. According to a study of Business Communications Co. Inc. (BCC, Table 1.1), the global market for industrial enzymes was $2.2 billion in 2006 while it should reach $2.7 billion by 2012, with a compound annual growth rate (CAGR) of 4% (Thakore, 2008). Moreover, the latest market study of industrial enzymes performed by Freedonia group (2009) reported that the world industrial enzymes trade in 2009 is 5.1 billion dollars and the global enzyme demand will rise 6.3 percent annually through 2013. Although many enzymes are obtained from animal and plant sources, microorganisms are the favoured source for production of industrial enzymes because of their biochemical and economic advantages. Microbial enzymes are more stable than analogous proteins obtained from plant and animal sources. Also, microorganisms can be subjected to genetic manipulation more readily (Hatamoto et al., 1996; Kar and Banerjee, 2000; Rout and Banerjee, 2006). The main industries, which utilize microbial enzymes (Figure 1.1), are detergents (35%), dairy processes (14%), starch (12%), textiles (11%), beverages and brewing (7%), animal feed (7%), baking (5%), and other uses (9%), (Viniegra-Gonzalez and FavelaTorres, 2004). Tannin acyl hydrolase (EC 3.1.1.20) which is commonly referred to as tannase is one of the important hydrolytic microbial enzymes (Bajpai and Patil, 1997) . Van Teighem accidentally discovered this unique enzyme in 1867. He was the first to demonstrate that the formation of gallic acid during fermentation of gall nut (which contains high levels of tannins) is due to the action of Aspergillus niger tannase (Knudson, 1913a; Albertse, 2002 and Aguilar et al., 2007). Tannase activity has been detected in bacteria, moulds and yeasts (Albertse, 2002; Purohit et al., 2006; Sabu et al., 2006; Banerjee and Pati, 2007; Rodriguez et al., 2008) Tannase is an industrially important enzyme and has several applications in various industries such as foods, animal feeds, cosmetic, pharmaceutical, chemical, leather industries etc. (Guo and Yang, 2000; Aguilar and Gutierrez-Sanchez, 2001; DNJHQDVet al., 2007a,b; Yu and Li, 2006; Aguilar et al., 2007; Jun et al., 2007; Noguchi et al., 2007). Concerning the applications of tannase in food industries, it is used as a clarifying agent in manufacture of instant tea (Takino, 1976; Tsai, 1987; Barmentlo et al., 1993; Agbo et al., 1995; Lehmberg et al., 1999; Boadi and Neufeld, 2001; Wright, 2005; Lu and Chen, 2007; Lu and Chen, 2008; Lu et al., 2009), the industrial processing of coffeflavored soft drinks (Aguilar and Gutierrez-Sanchez, 2001; Saxena and Saxena, 2004; Yu et al., 2004a) and as clarifying and debittring agent in fruit juices manufacture (Aguilar and Gutierrez-Sanchez, 2001 and Rout and Banerjee, 2006). In addition, it is also used for detannification of high tannins-contining foods such as lentils and pea flours and green tea
-1-
1,105
800
260
2,165
1,075
775
240
2,090
Technical Enzymes
Food Enzymes
Animal feed Enzymes
Total
Data were collected by BCC as mentioned by Thakore (2008)
2006
2005
Application sector
-2-
Table (1.1): Enzyme markets based on application sectors ($ Million).
2,250
280
8,30
1,140
2007
2,740
375
1,010
1,355
2012
4
6
4
3.5
CAGR% (2007-2012)
extract in order to increase its nutritive value and antioxidant activity (Mingshu et al., 2006; DNJHQDVet al., 2007a,b; Urbano et al., 2007; Battestin et al., 2008). Also, it is used in the enzymatic production of the antioxidant propyle gallate (Gaathon et al., 1989; Sharma and Gupta, 2003; Yu et al., 2004 a,b,c; Yu and Li, 2005; Yu and Li, 2006; Yu et al., 2007).
-4-
2. AIM OF INVESTIGATION
This study was carried out with the following objectives in mind: 1. Isolation of tannase-producing microorganisms from natural sources. 2. Screening isolates as well as cultures obtained from colleagues or culture collections for its capability to secrete tannase. 3. Identification of the best tannase producer. 4. Optimizing of conditions controlling tannase production applying suitable statistical designs. 5. Studying the changes occurring during enzyme production under optimized conditions.
-5-
-6-
3. REVIEW OF LITERATURE
3.1 Action of tannase Tannase (tannin acyl hydrolase; EC 3.1.1.20) is one of hydrolases and was known to catalyze the breakdown of ester and depside bonds from hydrolysable tannins and gallic acid esters (Bajpai and Patil, 1997; Sharma et al., 1999; Sharma et al., 2000; Aguilar and Gutierrez-Sanchez, 2001; Pinto et al., 2001; Chowdhury et al., 2004; Saxena and Saxena, 2004). This enzyme is known to display two different activities (Haslam and Stangroom, 1966). The first one is an esterase activity; by which it can hydrolyze ester bonds of gallic acid esters with glucose (galloyl-glucose) or alcohols (e.g. methyl gallate) as shown in Figure (3.1-A). The second activity is called depsidase activity; by which it can hydrolyze depside bonds of m-digallic acid (Figure 3.1-B). It is well known that tannase can hydrolyze the tannic acid although tannic acid is known to denaturate proteins. According to Aguilar and Gutierrez-Sanchez (2001) and Albertse (2002) tannase was shown to completely hydrolyze tannic acid to gallic acid and glucose (Figure 3.2), that lost tannic acid its propriety of binding and denaturating proteins. In addition, Aguilar and Gutierrez-Sanchez (2001) and Albertse (2002) reviewed also that when the reaction substrate is the methyl ester of gallic acid, tannase gives gallic acid and methanol as final products. Tewari et al. (1996) and Sharma and Gupta (2003) noticed that in aqueous solution; the direction of tannase reaction is predominantly toward the formation of gallic acid. On the contrary, tannase under non-aqueous solvent (organic media) can carry out the reversed reaction and can synthesize gallic acid esters such as methyle gallate and propyle gallate with a variety of alcohols such as methyle and n-propyle alcohols as shown in Figure (3.3).
3.2 Methods of tannase determination There are several methods (titrimetric, spectrophotmetric, colorimetric and chromatographic methods) that exist for determining tannase activity (Rhind and Smith, 1922; Parmentier, 1970; Haslam and Tanner, 1970; Jean et al., 1981; Katawa et al., 1981; Osawa and Walsh, 1993 Sharma et al., 2000; Mondal et al., 2001b; Nishitani and Osawa, 2003). Most of these methods have many limitations. The oldest method for tannase assay is based on the titration of gallic acid liberated by the action of tannase on tannic acid. This method did not give correct results due to the problem of accurately determining the end point (Sharma et al., 2000 and Mondal et al., 2001 b) because both substrates (tannic acid) and product (gallic acid) become brown by the addition of NaOH (Mondal et al., 2001 b).
-7-
-8-
Figure (3.1): Esterase and depsidase activities of tannase.
-9-
Figure (3.2): Action of tannase on tannic acid.
- 10 -
Figure (3.3): Esertification of gallic acid in organic media using tannase.
Sharma et al. (2000) and Ayed and Hamdi (2002) described the method published in 1967 by Libuchi, which is a spectrophotometeric method that has been used by many workers in its original and modified forms, this method is based on the decrease in absorbance of the substrate (tannic acid) at 310 nm. The shortcoming of this method is based on the extremely narrow difference between the absorbance optima of gallic acid (263 nm) and tannic acid (278 nm) (Haslam and Tanner, 1970; Sharma et al., 2000; Mondal et al., 2001 b). Haslam and Tanner (1970) developed a spectrophotometeric method which used pnitrophenol esters of gallic acid as substrates. But this method is not acceptable because of the non availability of the substrate (Iacazio et al., 2000 and Mondal et al., 2001 b). In addition to the colour instability of librated p-nitrophenol which affected by changes in substrate pH value (Mondal et al., 2001 b). After that, in the year of 1983, the method of Deschamps was reported. This method as described by Mondal et al. (2001 b) and Saxena and Saxena (2004) depends on measuring the gallic acid produced at 260 nm after precipitating the residual tannic acid using bovine serum albumin (BSA) solution. But Mondal et al. (2001 b) noticed that it is erroneous, as Absorbance (A) optima of gallic acid (263 nm), tannic acid (278 nm) and BSA (280 nm) are much closed to each other. At the same time all protein molecules could not bind with total tannic acid which interfere the A value of gallic acid. Therefore, Mondal et al. (2001 b) tried to avoid this drawback by using a method based on the changes in Absorbance at 530 nm of brown colour formed between FeCl3 and substrate (tannic acid) after enzymatic reaction that is also after precipitating the residual tannic acid using BSA solution. Sharma et al. (2000) invented a new spectrophotometric method to determine fungal tannase activity. This method is based on chromogen formation between gallic acid (released by the action of tannase on methyl gallate) and rhodanine (2-thio-4ketothiazolidine) and determining the enzyme activity spectrophotomctrically at 520 nm. But Mondal et al.(2001 b) noticed that the main drawback of this method is methyl gallate which is not specific substrate for tannase and tannic acid in the culture medium also interfere with colour formation. However, Jean et al. (1981) showed that methyl gallate is appropriate substrate because it is chemically defined and an easily available compound. Nishitani and Osawa (2003) suggested the new colorimetric method to assay bacterial tannase activity. This method used methyl gallate (a simple galloylester of methanol) as a substrate to be hydrolyzed by the bacterial tannase and then gallic acid released from methyl gallate was oxidized in alkaline condition by Na HCO 3 to give a green to brown colour that can be measured at 450 nm. The authors compared between this method and the method of Sharma et al. (2000).They found that this method successfully calibrated a range of tannase activities (0.63-10 mU/ml) but that of Sharma et al. (2000) failed to calibrate these activities. In contrast, the conventional method of Sharma et al. (2000) successfully estimated tannase at much higher activities (100-1000 mU /ml).
- 11 -
Concerning the chromatographic techniques, it was take place as methods to determine tannase activity either by gas chromatography (GC) or high performance liquid chromatography (HPLC) techniques. Jean et al. (1981) reported a method based on the quantitative determination of tannase activity by GC technique after silylation of gallic acid released from tannase action on methyl gallate as a substrate and silylated caffeic acid is used as internal standard. Ramirez-Coronel et al. (2003) used HPLC methodology to assay tannase activity. But these methods require more sophisticated instrumentation, more time consuming and are not suitable for routine assays (Mondal et al., 2001 b).
3.3 Microorganisms producing tannase
Microorganisms are considered as the most important and commercial sources of tannase. That is because the produced tannases are more stable than similar ones obtained from the other sources. Moreover, microorganisms can produce tannase in high quantities in a constant way. Microbial tannase is favoured also because the microbes can be subjected to genetic manipulation more readily than plants and animals, resulting in an increase in tannase production (Aguilar and Gutierrez-Sanchez, 2001; Kar et al., 2002; Purohit et al., 2006; Sabu et al., 2006). Mahadevan and Muthukumar (1980); Scalbert (1991); Bhardwaj et al. (2003); Saxena and Saxena (2004) reported that tannins display a high antibacterial and antifungal activity, these activities are due to the denaturation and precipitation of proteins and enzymes produced by these microorganisms. However, a few bacteria and fungi (Table 3.1) have been shown to exhibit resistance to tannins by the production of tannase. Saxena and Saxena (2004) noticed that condensed catechinic tannins are mainly depolymerized by bacteria and yeasts whereas, hydrolysable tannins, particularly gallotannins, are mostly hydrolyzed by mould species.
3.3.1 Bacterial tannase Usually, using bacterial isolates for tannase production is more favourable than moulds and yeasts because it can grow easily and produce a huge amount of tannase within a short period. Over past two decades, many tannase-producing bacterial genera have been isolated from different sources. Most of them are belonging to the following genera: Acromobacter, Bacillus, Citrobacter, Corynebacterium, Enterococcus, Herbaspirillum, Klebsiella, Lactobacillus, Lonepinella, Pseudomanas, Selenomonas, Staphylococcus, Stenotrophomonas and Streptococcus (Table 3.1).
- 12 -
Table (3.1): Tannase -producing microorganisms in literature. Microorganism
Reference(s)
Bacteria Acromobacter sp. Bacillus cereus. Bacillus licheniforms. Bacillus polymexa. Bacillus pumilus. Bacillus sp. Citrobacter freundii. Corynebacterium sp. Enterococcus faecalis. Herbarspirillum chlorophenolicum. Klebsiella oxytoca. Klebsiella pneumoniae. Lactbacillus animalis. Lactbacillus apodemi Lactobacillus murinns. Lactobacillus paraplantarum. Lactobacillus pentosus. Lactobacillus plantarum. Lonepinella koalarum. Pseudomonas aeruginosa Pseudomonas citronellolis. Pseudomanas putida. Selenomonas ruminantium. Staphylococcus lugdunensis. Stenotrophomonas maltoophilia. Streptococcus bovis. Streptococcus gallolyticus.
Bhat et al. (1998) Mondal et al. (2001 a) Mondal et al. (2000); Nishitani and Osawa (2003); Das Mohapatra et al. (2006); Mohapatra et al. (2007); Das Mohapatra et al. (2009) Bhat et al. (1998) and Belmares et al. (2004) Belmares et al. (2004) Mondal and Pati (2000); Mohapatra et al. (2007) Kumar et al. (1999) Belmares et al. (2004) Goel et al. (2007) Franco et al. (2005) Franco et al. (2005) Osawa et al. (2000); Franco et al. (2005) Sasaki et al. (2005) Osawa et al. (2006) Sasaki et al. (2005) Nishitani and Osawa (2003); Nishitani et al. (2004) Nishitani and Osawa (2003); Vaquero et al. (2004); Noguchi et al. (2007) Osawa et al. (2000); Ayed and Hamdi (2002); Nishitani and Osawa (2003); Vaquero et al. (2004); Noguchi et al. (2007) Nishitani and Osawa (2003) Selwal et al. (2010) Chowdhury et al. (2004) Franco et al. (2005) Belmares et al. (2004) Noguchi et al. (2007) Franco et al. (2005) Osawa (1990); Belmares et al. (2004) Nishitani and Osawa (2003); Sasaki et al. (2005); Noguchi et al. (2007)
Fungi A. Moulds Aspergillus acolumaria. Aspergillus aculeatus. DBF9
Rajakumar and Nandy (1983) Banerjee et al. (2001) and Banerjee et al. (2007)
- 13 -
Continued Aspergillus alliaceus. Aspergillus amstellodeni. Aspergillus awamori. Aspergillus caespitosum. Aspergillus carbonarius Aspergillus carneus. Aspergillus cescheri. Aspergillus fischerii. Aspergillus flavus. Aspergillus foetidus. Aspergillus fumigates. Aspergillus janus. Aspergillus japonicas. Aspergillus luchuensis. Aspergillus nidulans. Aspergillus niger. Aspergillus niger. Aa-20 Aspergillus niger. ATCC 16620 Aspergillus niger. HA37 Aspergillus niger. LCF8 Aspergillus niger. LLT25A5 Aspergillus niger. PKL104 Aspergillus niger. VanTieghem. Aspergillus oeavipes. Aspergillus oeavus. Aspergillus oryzae. Aspergillus penicilliformis. Aspergillus ruber. Aspergillus striatus. Aspergillus tamarii Fusarium solani. Paecilomyces variotii. Penicillium capsulatum.
Rajakumar and Nandy (1983) Rajakumar and Nandy (1983) Rajakumar and Nandy (1983); Chhokar et al. (2010) Rajakumar and Nandy (1983) Diepeningen et al. (2004) Rajakumar and Nandy (1983) Rajakumar and Nandy (1983) Bajpai and Patil (1997) Kachouri et al. (2005); Paranthaman et al. (2009) Mukherjee and Banerjee (2004); Purohit et al. (2006) Rajakumar and Nandy (1983); Manjit et al. (2008) Rajakumar and Nandy (1983) Bradoo et al. (1997) and Gupta et al. (1997) Nierenstein (1922) Rajakumar and Nandy (1983) Weetal (1985a,b); Bajpai and Patil (1997); Yu, et al. (2004a); Murugan et al. (2007); Sharma et al. (2007) Aguilar et al. (2002); Ramirez-Coronel et al. (2003); Trevino et al. (2007) Sabu et al. (2005 a,b) Aissam et al. (2005) Barthomeuf et al. (1994) Pinto (2001) Lekha and Lonsane (1994); Lekha et al. (1994) Knudson (1913a,b); Sharma et al. (1999); Sharma and Gupta (2003); Bhardwaj et al. (2003); Rana and Bhat (2005) Rajakumar and Nandy (1983) Rajakumar and Nandy (1983) Hatamoto et al. (1996); Rajakumar and Nandy (1983); Kar and Banerjee (2000); Garcia-conesa et al. (2001); Huang et al. (2007); Rodrigues et al. (2007); Paranthaman et al. (2008) Rajakumar and Nandy (1983) Kumar et al. (2007) Rajakumar and Nandy (1983) Enemuor and Odibo (2009) Bajpai and Patil (1997) Mahendran et al. (2006); Battestin and Macedo (2007); Battestin et al. (2008); Raaman et al. (2010). Rajakumar and Nandy (1983)
- 14 -
Continued Penicillium charlesii. Penicillium chrysogenum. Penicillium corylophilum. Penicillium crustosum. Penicillium digitatum. Penicillium expansum. Penicillium giganticans. Penicillium glabrum Penicillium granulatum. Penicillium restrictum. Penicillium rutgulosum. Penicillium spiculosporum. Penicillium vaiable. Rhizopus oryzae. Trametes versicolor. Trichoderma viride. Verticillium sp
Rajakumar and Nandy (1983) Nuero and Reyes (2002) and Batra and Saxena (2005) Rajakumar and Nandy (1983) Rajakumar and Nandy (1983) Rajakumar and Nandy (1983) Rajakumar and Nandy (1983) Rajakumar and Nandy (1983) Van de Lagemaat and Pyle (2005) Rajakumar and Nandy (1983) Rajakumar and Nandy (1983) Rajakumar and Nandy (1983) Rajakumar and Nandy (1983) Saxena and Saxena (2004);Sharma et al. (2008) Hadi et al. (1994); Chaterjee et al. (1996); Misro et al. (1997); Kar and Bnerjee (2000); Kar et al. (2002); Purohit et al. (2006) Archambault et al. (1996) Bajpai and Patil (1997) Kasieczka-Burnecka et al. (2007)
B. Yeasts Aureobasidium pullulans DBS66 Candida guillermondii. Candida sp. Candida tropicalis. Debaryomyces hansenii. Mycotorula japonica. *Pichia pastoris. *Saccharomyces cerevisia.
Banerjee and Pati (2007) Saxena and Saxena (2004) Belmares et al. (2004) Saxena and Saxena (2004) Bhat et al. (1998) Belmares et al. (2004) Zhong et al. (2004); Yu and Li (2008) Albertse (2002)
*Recombined strains
- 15 -
Generally, it is acceptable that the bacteria are very sensitive to the presence of tannic acid but the aforementioned bacteria were able to grow on this compound and degrade it (Osawa, 1990; Kumar et al., 1999; Mondal and Pati, 2000; Osawa et al., 2000; Ayed and Hamdi, 2002; Chowdhury et al., 2004; Franco et al., 2005; Noguchi et al., 2007)
3.3.2 Fungal tannase A-Tannase from moulds Moulds are the most studied microorganisms for tannase production. Moulds have the ability to degrade tannins as a sole carbon source (Aguilar and Gutierrez-Sanchez, 2001). The common mould genus used for tannase production either for research purposes or on industrial scale production was Aspergillus and the common Aspergillus species used for tannase production was Aspergillus niger. Many investigators have produced the enzyme tannase from the mould Aspergillus niger (Knudson, 1913 a,b; Weetal, 1985a,b; Barthomeuf et al., 1994; Lekha and Lonsane, 1994; Lekha et al., 1994; Bajpai and Patil, 1997; Sharma et al., 1999; Pinto et al., 2001; Aguilar et al., 2002; Bhardwaj et al., 2003; Ramirez-Coronel et al., 2003; Yu et al., 2004 a; Aissam et al., 2005; Rana and Bhat, 2005; Sabu et al., 2005 a,b; Murugan et al., 2007; Sharma et al., 2007; and Trevino et al., 2007). On the other hand, Bajpai and Patil (1997) compared tannase production by several fungal species; they found that Fusarium solani, Trichoderma viride and Aspergillus fischerii were better enzyme producers than Aspergillus niger which is the fungal strain widely studied and characterized as the best tannase producer. In addition, there are number of investigations about production of tannase from other different Aspergillus species such as A. acolumaria, A. aculeatus, A. alliaceus, A. amstellodeni, A. awamori, A. caespitosum, A. carbonarius, A. carneus, A. cescheri, A. fischerii, A. flavus, A. foetidus, A. fumigates, A. janus, A. japonicas, A. luchuensis, A. nidulans, A. oeavipes, A. oeavus, A. oryzae, A. penicilliformis, A. ruber, A. striatus and A. tamarii. Also, there are few articles about tannase production by several mould genera other than Aspergillus, such as Fusarium, Paecilomyces, Penicillium, Rhizopus, Trametes, Trichoderma and Verticillium (Table 3.1). Additionally, Lane et al. (1997) evaluated the safety of Aspergillus oryzae tannase for using it in food processing and reported that Aspergillus oryzae tannase can be regarded as safe for its intended use in processing tea. Sharma et al. (1999) reported that Aspergillus niger is an officially approved microorganism in France for tannase production to be used in the food industries. Also, tannase produced by A. niger is classified as generally regarded as safe (GRAS) by U.S. Food and Drug Administration. B- Tannase from yeasts: Saxena and Saxena (2004) reviewed that the yeasts Candida guilermondii and Candida tropicales have potential ability to produce tannase. Zhong et al. (2004) produced tannase from the recombinant yeast Pichia pastoris which is regarded as a good producer of tannase. Few other species of yeast are reported to produce tannase (Table 3.1).
- 16 -
3.4 Isolation and screening of microorganisms for tannase production Many microorganisms were isolated and screened for tannase production using various methods and media as well (Osawa, 1990; Bradoo et al., 1996; Pinto et al., 2001; Chowdhury et al., 2004; Batra and Saxena, 2005; Sasaki et al., 2005; Murugan et al., 2007).
Osawa (1990) used tannin-treated brain heart infusion agar to isolate tannaseproducing Streptococcus sp. from feces of koalas animals fed on forests with high content ration of tannins. Among the twelve obtained isolates, Streptococcus bovis biotype I was the promising one. In this technique, the surface of solidified brain heart infusion agar medium was overlaid aseptically by approximately 5 ml of 2 % tannic acid solution for 20 minutes then 0.1 ml of the sample suspension was spread onto the plates after draining of the excess tannic acid solution. The plates were incubated anaerobically at 37º C for 72 h. Colonies with distinct clear zones grew on the plate were selected as tannase producers. Also, Sasaki et al. (2005) successfully used tannin-treated brain heart infusion agar described by Osawa (1990) to isolate tannase-producing bacteria (cocci and bacilli) from the feces of the Japanese large wood mouse, Apodemus speciosus. The tannase-producing isolates were belonging to Streptococcus gallolyticus, Lactobacillus animalis and Lactobacillus murinus. Chowdhury et al. (2004) used a minimal medium containing 1% (w/v) tannic acid as sole carbon source to isolate tannin-degrading aerobic bacteria from tannery soil. Results showed that the most efficient isolate degrading tannic acid was belonging to Pseudomonas citronellolis.
3.4.2 Isolation and screening of tannase-producing fungi
Bradoo et al. (1996) applied a rapid and simple plate assay to isolate and screen tannase-producing fungi. In which, a medium consists of 0.01 M phosphate buffer (pH 6.0), 1% tannic acid as sole carbon source and 3% agar was point inoculated, incubated for 48 h and clear zone diameter (included colony diameter) was measured. The authors reported a high correlation coefficient (r = 0.93) between clear zone diameter (included colony diameter) and the quantitative enzyme production in broth medium. Fifty fungal isolates belonging to the genera Aspergillus, Cunnighamella, Fusarium, Helicostylum, Neurospora, Syncephalastrum and Trichoderma were successfully screened by this method. Among Aspergillus spp. the best tannase producers were Aspergillus japonicus ITCC 2620, Aspergillus awamori ITCC 924 and Aspergillus niger ITCC 595. Among Penicillium spp., Penicillium acrellanum and Penicillium digitatum were the best producers, while Fusarium solani and Trichoderma viride were the best producers among their genera.
- 17 -
The same previous technique was used by Pinto et al. (2001) in order to screen thirty strains of Aspergillus niger from EMBRAPA/Food Technology stock collection, Rio de Janeiro, Brazil. But using tannic acid agar medium (Czapek-Dox’s minimal medium containing 1% tannic acid as sole carbon source). They measured the diameters of colonies after 24 hours periods. They reported that because of the medium contained tannic acid as sole carbon source, the growth on the plate and determination of clear zones around the colonies suggesting tannase activity. They also mentioned that in case of difficulty to observe the clear zones, colony diameter can be used instead. Results showed that among the thirty tested strains of Aspergillus niger, Aspergillus niger 11T25A5 was the promising isolate.
3.4.1 Isolation and screening of tannase-producing bacteria
Batra and Saxena (2005) primary screened sixty isolates of fungi belong to Aspergillus and Penicillium for tannase activity using the simple plate technique adopted by Bradoo et al. (1996) then followed by a confirmatory quantitative secondary screening under submerged fermentation (SmF) technique using Czapek-Dox’s medium containing 1% tannic acid as sole carbon source. Results showed that twenty-five Aspergilli and twenty Penicillii were able to produce tannase and the potent tannase-producing Aspergilli were Aspergillus fumigatus (8.3 IU/ml), Aspergillus versicolor (7.0 IU/ml), Aspergillus flavus (4.95 IU/ml) and Aspergillus caespitosum (4.47 IU/ml). While amongst Penicillii, Penicillium charlesii (4.82 IU/ml) exhibited the highest tannase activity. Tannase-producing fungi were isolated from tannery effluent by Murugan et al. (2007) using the method described by Bradoo et al. (1996), then followed by a confirmatory secondary screening through SmF technique in a stirred tank bioreactor (Applikon B.V, The Netherlands) using tannic acid containing-mineral medium. The colony diameter on the solid surface media shows high correlation with quantitative production of tannase. Among the screened isolates, Aspergillus niger exhibited maximum production of extracellular (16.8 U/ml) and intracellular (3.6 U/ml) tannase.
3.5 Fermentation systems and production media for production of tannase from fungi
Several fermentation systems have been developed for the production of tannase from fungi using various production media. These systems can be divided into liquid surface fermentation (LSF), submerged fermentation (SmF), solid-state fermentation (SSF) and Modified solid state fermentation (MSSF). In which, several production media were used to grow the tannase-producing moulds (Table 3.2).
- 18 -
SmF
LSF
Fermentation system
Aspergillus awamori
Aspergillus aculeatus.
Aspergillus niger Van Tieghem.
Aspergillus niger PKL 104.
Aspergillus japonicas.
Aspergillus fischerii, Aspergillus niger., Fusarium solani and Trichoderma viride.
Fungi
- 19 -
Liquid medium consist of, (g/L): Tannic acid, 10; (NH 4 ) 2 HPO 4 , 3; KH 2 PO 4 , 0.5; MgSO 4 , 1; NH 4 Cl, 0.5; CaCl2 , 0.3. Czapek-Dox’s minimal medium containing 3.5% tannic acid.
Modified mildew test medium consist of, (g/L): NaNO 3 , 3.0; K 2 HPO 4 , 1.0; MgSO 4 .7H 2 O, 0.5; KCl, 0.5 + Tannic acid, 2%.
Mineral salt medium containing, (g/L): sucrose, 30; NH 4 NO 3 , 1.56; KNO 3, 1.90; MgSO 4 .7H 2 O, 6.37; CaCl2 .2H 2 O, 0.44; KH 2 PO 4 , 0.17 and (mg/L): H 3 Bo 3, 6.2; MnSO 4 .H 2 O, 16.9; ZnSO 4 .7H 2 O, 8.6; Na 2 MnO 4 .2H 2 O, 0.25; CuSO 4 .5H 2 O, 0.025; CoCl2 .6H 2 O, 0.025; FeSO 4 .7H 2 O, 5.5; NaEDTA, 7.6 + 0.1% appropriate tannins source (gallotannin, methyl gallate or gallic acid). Czapek-Dox’s minimal medium containing 2% glucose as sole carbon source. Tannic acid liquid medium containing, (g/L): KH 2 PO 4 ,1; NH 4 NO 3 , 2; MgSO 4 .7H 2 O,0.2; CaCl2 .2H 2 O, 0.02; MnCl.6H 2 O, 0.004; Na 2 MoO 4 .2H 2 O, 0.002; FeSO 4 .7H 2 O, 0.0025+ tannic acid, 20.
Medium composition
Reference
Seth and Chand (2000)
Banerjee et al. (2001)
Rana and Bhat (2005)
Lekha and Lonsane (1994)
Gupta et al. (1997)
Bajpai and Patil (1997)
Table (3.2): Fermentation systems and microbial tannase production media used for production of tannnase from fungi.
SmF
SmF
Continued
Aspergillus niger.
Aspergillus niger Aa-20.
Aspergillus niger Van Tieghem
Aspergillus niger PKL 104.
Aspergillus awamori nakazawa.
Ramirez-Coronel, et al. (2003)
Mineral salt medium containing, (g/L): KH 2 PO 4 , 5; NH 4 NO 3 , 10; CaCl2 .6H 2 O, 0.1; MgSO 4 .7H 2 O, 1; MnCl2 .6H 2 O, 0.02; NaMoO 4 .H 2 O, glucose, 2.5 + tannic acid, 0.1.
- 20 -
Aguilar et al. (2002)
Aguilar et al. (2001 a,b)
Tannic acid liquid medium containing, (g/L): KH 2 PO 4 , 1; NH 4 NO 3 , 2; MgSO 4 .7H 2 O, 0.2; CaCl2 .2H 2 O, 0.02; MnCl.6H 2 O, 0.004; Na 2 MoO 4 .2H 2 O, 0.002; FeSO 4 .7H 2 O, 0.0025 + tannic acid, 20. Modified mildew test medium consist of, (g/L): NaNO 3 , 3.0; K 2 HPO 4 , 1.0; MgSO 4 .7H 2 O, 0.5; KCl, 0.5 + tannic acid, 2%.
Sharma et al.(1999); Sharma et al. (2000); Rana and Bhat (2005); Sharma et al. (2007)
Bhat et al. (1997)
Lekha and Lonsane (1994)
Mahapatra et al. (2005)
Modified mildew test medium consist of, (g/L): NaNO 3 , 3.0; K 2 HPO 4 , 1.0; MgSO 4 .7H 2 O, 0.5; KCl, 0.5 + tannic acid, 2%.
Czapek-Dox’s minimal medium containing tannin rich substrates which consisted of myrobalan fruits and gallo seed cover in a fixed ratio of 0.25:1.5. Tannic acid liquid medium containing, (g/L): KH 2 PO 4 , 1; NH 4 NO 3 , 2; MgSO 4 .7H 2 O, 0.2; CaCl2 .2H 2 O, 0.02; MnCl.6H 2 O, 0.004; Na 2 MoO 4 .2H 2 O, 0.002; FeSO 4 .7H 2 O, 0.0025 + tannic acid, 20. Modified mildew test broth consist of, (g/L): NaNO 3 , 3.0; K 2 HPO 4 , 1.0; MgSO 4 .7H 2 O, 0.25; KCl, 0.25 + tannic acid, 150.
SSF
SmF
Continued
Manjit et al. (2008)
Various solid substrates such as Amla, Ber, Jamun, Jamoa and Keekar leaves powderes moistened with mineral salt solution (w/v): 0.25% NaNO 3 , 0.1% KH 2 PO 4 , 0.05% MgSO 4 .7H 2 O, 0.05% KCl, pH 5.0.
Aspergillus fumigates.
- 21 -
Hadi et al. (1994)
Modified Czapek's Dox medium (pH 6) containing, (g/L): NaNO 3 , 2.5; KCl, 0.5; MgSO 4 .7H 2 O, 0.5; KH 2 PO 4 , 1+ tannic acid 2 %.
Rhizopus oryzae.
Penicillium chrysogenum.
Saxena and Saxena (2004) and Sharma et al. (2008)
Rajakumar and Nandy (1983)
Czapek-Dox’s minimal medium containing 2% tannic acid as sole carbon source.
Paecilomyces variotii.
Penicillium variable.
Mahendran et al. (2006)
Minimal salt medium consisting of, (g/L):KH2PO4,1; NH 4 NO 3 , 2; MgSO 4 .7H 2 O,0.2 and, (mg/L) CaCl2 .2H 2 O,20; MnCl2 .4H 2 O,4; NaMoO 4 .2H 2 O,2; FeSO 4 .7H 2 O,2.5 + tannic acid 2%.
Modified Czapek's Dox medium (pH 6) containing, (g/L): NaNO 3 , 6; KCl, 0.52; MgSO 4 .7H 2 O, 0.52; KH 2 PO 4 , 1.52; glucose, 2; powdered Chebulic myrobalan, 1.16; Cu(NO 3 ) 2 .3H 2 O, FeSO 4 .7H 2 O, and ZnSO 4 .7H 2 O were added as trace elements.
Huang et al. (2005)
Inorganic salt medium containing 0.5%Valonia tannin. The medium consists of, (g/L): KH 2 PO 4 , 0.5; K 2 HPO 4 , 0.5; NaNO 3 , 2.0; KCl, 0.5; MgSO 4 , 0.5; and FeSO 4 , 0.01.
Aspergillus niger SHL6.
SSF
Continued
Aspergillus niger ATCC 16620.
Aspergillus niger Van Tieghem.
Aspergillus niger.
Aspergillus niger Aa-20.
Aspergillus niger PKL 104.
- 22 -
Sugar cane pith bagasse impregnated with mineral salt medium containing 1 % tannic acid. Sugar cane pith bagasse impregnated with mineral salt medium containing 1 % tannic acid. Wheat bran medium supplemented with 4% tannic acid (w/w). Polyurethane foam (PUF) as a solid support impregnated with tannic acid liquid medium containing, (g/L): KH 2 PO 4 , 1; NH 4 NO 3 , 2; MgSO 4 .7H 2 O, 0.2; CaCl2 .2H 2 O, 0.02; MnCl.6H 2 O, 0.004; Na 2 MoO 4 .2H 2 O, 0.002; FeSO 4 .7H 2 O, 0.0025 + tannic acid, 20. Gobernadora (Larrea trientata cov.) powder impregnated with Czapek-Dox’s modium in 3:7 ratio. Wheat bran enriched with 0.8% (w/w) tannic acid and moistened with 0.91 % w/v (NH 4 ) 2 SO 4 solution. Polyurethan foam (PUF) as solid support moistened with liquid medium containing, (g/L): KH 2 PO 4 , 5; NH 4 NO 3 , 10; CaCl2 .6H 2 O, 0.1; MgSO 4 .7H 2 O, 1; MnCl2 .6H 2 O, 0.02; NaMoO 4 .H 2 O, 0.002; glucose, 2.5 + tannic acid, 0.1. PUF as solid support moistened with slightly modified Mildew test medium. The basal medium containing, (g/L): NaNO 3 , 3.0; K 2 HPO 4 , 1.0; MgSO 4 .7H 2 O, 0.5; KCl, 0.5 6 + tannic acid, 2%. Palm kernel cake and tamarind seed powder as solid supports and carbon sources, moistened with a salt solution composed of 0.5% w/v NH 4 NO 3 , 0.1% w/v MgSO 4 .7H 2 O and 0.1% w/v NaCl.
Sabu et al. (2005 b)
Rana and Bhat (2005)
Ramirez-Coronel et al. (2003)
Pinto et al. (2001)
Trevino-Cueto et al. (2007)
Aguilar et al. (2001 a,b); Aguilar et al. (2002); Montalvo et al. (2005)
Lekha et al. (1994)
Lekha and Lonsane (1994)
Caesalpinia digyna (Teri pod shrub cover powder) 2% as a substrate in float coms of modified Czapek-Dox's medium.
Rhizopus oryzae
- 23 -
SmF: Submerged Fermentation. MSSF: Modified Solid State Fermentation.
Termenalia chebula (myrobalan) powder and Caesalpinia digyna (Teri pod shrub cover powder) 2% as a substrate in float coms of modified Czapek-Dox's medium.
Powdered Caesalpinia digyna seed cover as solid support and substrate moistened with modified Czapek-Dox's medium.
Cashew apple bagasse enriched with 2% tannic acid and moistened by water. Rice straw powder and sugarcane baggase powder (1:1 ratio) enriched with 0.1% gallic acid moistened with 5 ml of mineral salt solution consist of: NH 4 NO 3 , 0.5 %; NaCl, 0.1 %; MgSO 4 .7H 2 O, 0.1 % + Gallic acid 4% Jamun leaves moistened with mineral salt solution (KNO 3 , 0.2%; KH 2 PO 4 , 0.1%; MgSO 4 .7H 2 O, 0.05%; KCl, 0.05%). PUF as solid support moistened with liquid medium containing (w/v): 5% tannic acid, 1.0% NH 4 NO 3 , 0.5% KH 2 PO 4 , 0.25% glucose, 0.1% MgSO 4 ·7H 2 O, 0.01% CaCl2 , 0.002%MnCl2 ·4H 2 O, 0.001% FeSO 4 ·7H 2 O and 0.001% Na 2 MoO 4 ·2H 2 O in 0.15M phthalate buffer (pH 5.5). Wheat bran moistened by Czapek-Dox’s medium contained 2.5% tannic acid.
Aspergillus foetidus and Rhizopus oryzae.
Rhizopus oryzae.
Penicillium glabrum.
Aspergillus ruber.
Aspergillus oryzae.
LSF: Liquid Surface Fermentation. SSF: Solid State Fermentation.
MSSF
SSF
Continued
Kar and Banerjee (2000), Kar et al. (2002) and Kar et al. (2003) Mukherjee and Banerjee, (2004); Banerjee et al. (2005); Mukherjee and Banerjee, (2006); Purohit et al. (2006) Kar and Banerjee (2000) and Kar et al. (2002)
Chaterjee et al. (1996)
Van de Lagemaat and Pyle (2001) and (2004)
Kumar et al. (2007)
Paranthaman et al. (2008)
Rodrigues et al. (2007); Rodrigues et al. (2008)
Belmares et al. (2004) mentioned in a comprehensive review that tannase production has been carried out mainly under submerged (SmF) and solid-state fermentation (SSF) techniques, depending on the strain and culture conditions. Concerning submerged culture fermentation (SmF), Viniegra-Gonzalez et al. (2003) reported that most enzymes manufacturers produce enzymes using SmF technique with tannase titers in the range of grams per liters. However, recently there is a significant interest with using SSF technique to produce tannase from fungi as indicated by the growing number of research papers (Lekha and Lonsane, 1994; Chatterjee et al., 1996; Kar and Banerjee, 2000; Aguilar et al., 2001a,b; Pinto et al., 2001; Aguilar et al., 2002; Van de Lagemaat and Pyle, 2004; Montalvo et al., 2005; Rana and Bhat, 2005; Sabu et al., 2005b; Kumar et al., 2007; Rodrigues et al., 2007; Trevino-Cueto et al., 2007; Manjit et al., 2008). Several studies have reported interesting advantages of tannase produced by SSF over that produced by SmF. All of these studies showed that the first advantage of SSF technique is the higher enzyme titers than in SmF, when comparing the production of the same strain and fermentation broth (Lekha and Lonsan, 1994; Kar and Banerjee, 2000; Aguilar et al., 2002; Ramirez-Coronel et al., 2003; Viniegra-Gonzalez et al., 2003; Montalvo et al., 2005; Trevino-Cueto et al., 2007). A comparative study was carried out by Lekha and Lonsane (1994), in which the effect of fermentation system on tannase titers produced by Aspergillus niger PKL104 was studied. The results showed that tannase production was 2.5 and 4.8 times higher in the SSF system as compared to those in SmF and LSF techniques, respectively. Moreover, the fermentation time required to obtain the maximum enzyme titers was shorter in the SSF process as compared to the other two fermentation techniques. Aguilar et al. (2001a) reviewed that tannase production at high concentration of sugars was induced in SSF technique whereas in SmF technique tannase production was repressed. They also reviewed that there are several hypotheses have been proposed to explain this phenomenon. One of these hypotheses suggested that the SSF technique minimizes the catabolic repression phenomenon. Additionally, Aguilar et al. (2002) compared activity titers of Aspergillus niger Aa-20 tannase in each of SmF and SSF techniques. The results showed that the titer for enzyme produced by SSF was higher (up to six times) than those obtained by SmF. They also reported that using SSF can minimize unwanted proteolytic activities which can considerably reduce final titer and the stability of desired enzyme tannase. Viniegra-Gonzalez et al. (2003) observed that tannase productivity by Aspergillus niger Aa-20 in SSF using polyurethane foam as solid support was higher than that in SmF. They reported that SSF cultures have much longer air to liquid interphase than conventional SmF cultures, about 400 times higher. Moreover, SSF cultures don’t need mechanical energy expenditures. Montalvo et al. (2005) mentioned that the higher tannase productivity in SSF technique is due to the lower catabolic repression effect of glucose (the product of tannic hydrolysis by tannase) in SSF technique than the other techniques. This is because of the
24
fact that the repressor molecule (glucose) uptake rate on solid matrix (polyurethane foam) is higher than the repressor molecule diffusion rate on solid matrix. In addition to previous advantages of SSF over SmF, Lekha and Lonsane (1994); Chatterjee et al. (1996); Pinto et al. (2001) and Ramirez-Coronel et al. (2003) reported that tannase produced by SSF was wholly extracellular while it was exclusively intracellular in SmF and LSF techniques. They also mentioned that the small size reactor required, the simple technology, reduced cost of dewatering are also among the advantages of SSF over SmF technique. Rana and Bhat, (2005) found that tannase produced by SSF exhibited activity over a wider pH range and more thermostablity than the same produced by SmF or LSF techniques. Moreover, during SSF most of supports are also substrates. Lekha and Lonsane (1994) used sugar cane pith bagasse as inert solid support for tannase production under SSF by Aspergillus niger PKL 104. Chatterjee et al. (1996) used wheat bran as solid support to produce tannase by Rhyzopus oryzae under SSF technique. Kar and Banerjee (2000) used powdered Caesalpinia digyna seeds covers as a substrate and solid support to produce tannase by Rhizopus oryzae under SSF technique. Sabu et al. (2005 b) used tamarind seed powder and Palm kernel cake as substrates and solid supports in SSF to produce tannase by Aspergillus niger ATCC 16620. Kumar et al. (2007) used jamun leaves as natural substrate and solid support for tannase production by Aspergillus ruber under SSF. Rodrigues et al. (2007) used Cashew apple bagasse for tannase production by Rhizopus oryzae. Also, gobernadora (Larra tridentate cov.) powder has been used as a solid support and natural substrate for production of Aspergillus niger Aa-20 tannase (Trevino-Cueto et al., 2007). Recently, extracellular tannase of Aspergillus fumigatus has been produced under SSF using different agro forest residues such as Amla leaves (Phyllanthus emblica), Ber leaves (Zyzyphus mauritiana), Jamun leaves (Syzygium cumini), Jamoa leaves (Syzygium sp.) and Keekar leaves (Acacia nilotica) (Manjit et al., 2008). Although tannase production using SSF technique is more advantageous over SmF or LSF techniques as mentioned above, a large quantity of heat is generated in solid media due to the microbial metabolic activity in solid process leading to rapid rise in temperature of fermenting solid bed. The removal of this heat doesn’t occur in SSF due to lack of heatexchange surface and poor heat transfer through the solid bed causing large moisture losses and drying of the solid substrate. To overcome the harmful effects on microbial growth and activity due to this heat, modified solid-state fermentation (MSSF) has been introduced as a new fermentation technique for tannase production (Kar and Banerjee, 2000; Mukherjee and Banerjee, 2004; Banerjee et al., 2005; Mukherjee and Banerjee, 2006; Purohit et al., 2006) in which the solid substrates placed on its float comes in continuous contact with the liquid medium (Czapek-Dox’s medium) to avoid this disadvantage of traditional solid-state fermentation using a special bioreactor called GROWTEK bioreactor. The bioreactor design used in MSSF was described by Mukherjee and Banerjee, (2004) it is a cylindrical vessel having height (16 cm) and diameter (11.3 cm) with a spout near the base on the wall (inclined at 15° to the vertical axis). Medium can be removed through the spout without disturbing fermentation. A perforated float is present inside the vessel consisting of a base of glass wool cloth (72 cm2 area) and a periphery made of polypropylene having a hollow structure.
25
3.6 Biosynthesis of tannase and regulation of its production
Many enzymes are produced continuously irrespective of environmental conditions. These are called constitutive enzymes, and are formed at constant rates and in constant amounts, regardless of the metabolic state of the organism. The others are of inducible enzymes which are either completely absent or present as traces. Their concentration can quickly increase a thousand-fold or more when their substrate is present in the medium, particularly when the substrate is the only carbon source of the cell (Rastogi, 2003). There are very few studies about the mechanism of tannase biosynthesis and regulation of its production in the cell. Knudson (1913 a) reported that Aspergillus niger tannase is inducible enzyme as it is induced only when the Aspergillus niger grown in the presence of tannic acid , giving gallic acid and glucose as final products. This observation is contradictory to that published by Gupta et al. (1997). They reported that tannase is a constitutive enzyme as its activity is expressed when the Aspergillus japonicus is grown on glucose only and in the absence of tannic acid. Also Bradoo et al. (1997) demonstrated the constitutive nature of tannase produced by Aspergillus japonicus as the enzyme could be expressed even in the absence of tannic acid. They recorded end-product repression of the enzyme by gallic acid. Bhat et al. (1998) noticed that tannase was produced constitutively on simple and complex sugar substrates but its activity was doubled in the presence of tannic acid as a sole carbon source. They also referred to strong end-product inhibition of the enzyme by gallic acid. Bradoo et al. (1997) suggested that one of intermediates that formed during biodegradation of tannic acid could be the true inducer because of the low reactivity of tannic acid molecule and its great size. On this aspect, it has been proposed that the tannase induction process is ejected similarly to the induction process of cellulases where it requires certain basal levels of the enzyme (constitutive) needed to start the hydrolysis of the substrate and to give conditions to obtain the true inducer activity (Aguilar and Gutierrez-Sanchez, 2001). Aguilar et al. (2001 a) showed that it is important to test the induction or repression of tannase by gallic acid. They studied the effect of gallic acid on tannase production by Aspergillus niger Aa-20. Results of cultures with gallic acid as sole carbon source showed that it did not induce tannase activity. Moreover, this compound repressed tannase activity. Generally, there is a lack of studies related to the mechanisms involved in the biosynthesis of tannase and its natural inducers.
26
3.7 Factors affecting tannase production by fungi
Enzyme production by fungi is influenced by various factors, especially the cultural conditions (carbon and nitrogen sources) and environmental conditions (temperature, pH, inoculum size, and agitation) (Hadi et al., 1994; Bajpai and Patil, 1997; Kumar et al., 2007).
3.7.1 Medium constituents There are many reports on the influence of medium components on the production of tannase by fungi. Many authors have used various fungi and culture media for tannase production (Chatterjee et al., 1996; Aguilar et al,. 2001 b; Aguilar et al., 2002 and Banerjee et al., 2005).
3.7.1.1 The effect of carbon source The synthesis of tannase by fungi is not only affected by the concentration of carbon source but also affected by the kind of certain carbon sources. Therefore, the effect of many carbon sources on the synthesis of tannase by fungi has been subjected to several studies. A-Tannic acid concentration Knudson (1913 b) observed a progressive increase in tannase synthesis by Aspergillus niger and Penicillium sp. in Czapek-Dox’s medium with the increase of tannic acid concentration up to 10%. Hadi et al. (1994) studied the effect of tannic acid concentration on the production of tannase by Rhizopus oryzae using modified Czapek-Dox’s medium under SmF technique. They reported that 2% tannic acid was the optimum concentration. Bradoo et al. (1997) noted that using 2% tannic acid as carbon source in Czapek-Dox's minimal medium gave the optimum production of tannase by Aspergillus japonicus. Chatterjee et al. (1996) studied the effect of tannic acid concentration on tannase synthesis by Rhizopus oryzae under SSF using wheat bran as solid support. They found that the maximum production was obtained at 2.5 % tannic acid. Aguilar et al. (2001 b) studied the effect of different tannic acid concentrations (12.5, 25, 50 and 100 g/L) on Aspergillus niger Aa-20 tannase production by SmF and SSF techniques. They found that the increase in tannic acid concentration increased tannase production in both fermentation techniques. Banerjee et al. (2001) studied the effect of tannic acid and glucose concentrations on tannase production by Aspergillus aculeatus DBF9. The results revealed that maximum intracellular tannase production occurred in the culture broth containing 1% (w/v) tannic
27
acid whereas, 2% (w/v) tannic acid favoured maximum growth and extracellular tannase production. Aissam et al. (2005) noted that optimum tannase synthesis by Aspergillus niger HA37 was attained when the medium of diluted olive mill waste water was supplemented with 1% tannic acid as carbon source. While, Sharma et al. (2007) reported that the statistical optimization of tannase production in modified Czapek-Dox’s medium by Aspergillus niger showed that the maximum tannase was yielded at 5% tannic acid. Banerjee and Pati (2007) reported that maximum growth and tannase production by Aureobasidium pullulans was noticed in the medium containing 1% tannic acid under SmF. Battestin and Macedo (2007) confirmed the significance effect of tannic acid on the secretion of tannase by Paecilomyces variotii under SSF technique using wheat bran as solid support. They found that 8.5-14% (w/w) tannic acid was the optimum concentrations range. Raaman et al. (2010) revealed that Paecilomyces variotii tannase production under SmF technique using tannic acid medium increased with increasing substrate concentration up to 1.5% beyond which it decreased.
B- Effect of addition carbon source incorporation with tannic Knudson (1913 b) observed that the addition of cane sugar to Czapek-Dox’s medium containing 2% tannic acid as sole carbon source decreased tannase production by Aspergillus niger and Penicillium sp. Hadi et al. (1994) studied the addition effect of 1% (w/v) different sugars (glucose, fructose, galactose, maltose and sucrose) incorporation with 2% tannic acid on Rhizopus oryzae tannase production in modified Czapek-Dox’s minimal medium. The results showed that the maximum tannase production was attained with tannic acid without sugar addition. They also studied the effect of glucose concentration (1, 3 and 5 % w/v) on tannase production. They found that tannase production was decreased with the increase in glucose concentration and 1% glucose was the optimum for tannase production. The addition of various mono and polysaccharides (arabinose, fructose, galactose, xylose, manitol, lactose, sucrose, starch, and carboxy methyle cellulose) to Czapek-Dox’s minimal medium containing 2% tannic acid had no positive effect on tannase production by Aspergillus japonicus except in case of glucose which maximized tannase activity at 0.2% concentration (Bradoo et al., 1997) Aguilar et al. (2001b) studied the effect of different glucose concentrations (6.25, 12.5, 25, 50 and 200 g/L) incorporation with tannic acid (25 g/L) on Aspergillus niger Aa20 tannase production by SmF and SSF. The results showed that the addition of glucose decreased tannase activity in SSF. Whereas, in SmF tannase activity increased when glucose concentration increased up to 2.5%, but strong catabolic repression of tannase synthesis was observed when glucose concentration of 5% was used.
28
Banerjee et al. (2001) studied the effect of glucose concentrations on tannase production by Aspergillus aculeatus DBF9 incorporation with tannic acid. The results showed that glucose at higher concentration (1% w/v) repressed extracellular and intracellular tannase synthesis while the lower concentrations (0.05 and 0.1% w/v) were not repressive for intracellular and extracellular, respectively. Battestin and Macedo (2007) reported that the addition of starch (0.4, 0.8, 1.2 % w/v) as additive carbon source incorporation with tannic acid had no significant effect on Paecilomyces variotii tannase production. The effect of sucrose addition (0.2%) as external carbon source on Aspergillus rubber tannase production under SSF was studied by Kumar et al. (2007). They found that there was a decrease in tannase production with sucrose as additive carbon source beside tannic acid. Banerjee and Pati (2007) found that using 0.1% additional carbon sources (maltose, glucose, sucrose, arabinos, lactose, mannitol, and xylose) incorporation with tannic acid (1%) in the mineral medium exhibited a negative effect on tannase formation by Aureobasidium pullulans, except glucose, maltose and sucrose and the maximum was for 0.1% glucose. Rodrigues et al. (2008) studied the effect of using 1% (w/v) additional carbon sources (glucose, maltose, starch, sucrose and glycerol) incorporation with tannic acid (2.5%) in cashew apple bagasse solid state medium on tannase production by Aspergillus oryzae. They found that supplementation with maltose and glycerol inhibited tannase synthesis whereas, supplementation with starch and sucrose increased enzyme production. Raaman et al. (2010) revealed that Paecilomyces variotii tannase production affected by the addition of 1% additional carbohydrates (arabinose, fructose, glucose, lactose, maltose, mannitol, raffinose, rhamnose, sucrose, sorbose, starch and xylose) incorporation with tannic acid (1.5%) under SmF technique using mineral medium. They found that all studied carbohydrates had adverse effect on tannase production except glucose which increased tannase production. In addition, they found that maximum tannase production was attained at 1% glucose concentration beyond which tannase production decreased. C-Natural carbon sources The effect of natural carbon source (Caesalpina digyna seed cover powder) as sole carbon source for the production of tannase by Rhizopus oryzae was carried out using submerged fermentation (SmF), solid state fermentation (SSF) and modified solid state fermentation (MSSF) techniques. The results showed that 2 grams of substrate/ Petri dish (moistened with 2 ml of modified Czapek-Dox’s medium) in SSF, 10% (w/v) in SmF and 20 g substrate/50 ml medium in MSSF gave optimum tannase production. Further increase in the natural substrate decreased the production (Kar and Banerjee, 2000). Belmares-Cerda et al. (2003) used several carbon sources (glucose, gallic acid, tannic acid, catechin, and infusions from sorghum, pecan peels and creosote bush) as
29
carbon sources to produce tannase by Aspergillus niger PSH. Results obtained revealed that the highest tannase activity was obtained with the infusion of creosote bush Saxena and Saxena (2004) used the dried powder of Chebulic myrobalan fruit (which contain about 32 % gallotannins) as a natural substrate for the production of Penicillium variable tannase. They found that the optimum concentration of substrate was 5.8 g/50 ml of the medium. Huang et al. (2005) used valonia tannins as a sole carbon source in inorganic salt medium to produce tannase by the fungus Aspergillus SHL6. The results indicated that valonia is not an effective carbon source for tannase production. Macedo et al. (2005) studied the effect of using coffee and grape residues which are rich in tannin on tannase synthesis using SSF, the best results were obtained with coffee residues.
3.7.1.2 The effect of nitrogen sources Different inorganic and organic nitrogen sources were tested to enhance tannase synthesis by fungi (Hadi et al., 1994; Bradoo et al., 1997; Huang et al., 2005; Sharma et al., 2007; Manjit et al., 2008). Belmares et al. (2004) reviewed that the nitrogen requirements for tannase production can be supplied by different inorganic and organic sources. Inorganic source can be supplemented as ammonium salts (sulphate, carbonate, chloride, nitrate, and monohydrated phosphate) or nitrate salts (sodium, potassium or ammonium) while organic sources including urea, aspartic acid, palmetic acid, peptone, brew's cream, yeast extract, malt extract and corn steep liquor. A-Inorganic sources Hadi et al. (1994) tested the effect of inorganic nitrogen sources (ammonium sulphate, ammonium nitrate, sodium nitrate, sodium nitrite, and urea) incorporation into Czapek-Dox's medium on Rhizopus oryzae tannase productivity. Results revealed that among the various tested inorganic nitrogen sources, sodium nitrate was found to be the best, and the highest tannase production was attained at 0.05% sodium nitrate. In addition, Sharma et al. (2007) reported that sodium nitrate concentration in modified Czapek-Dox's medium was effective for tannase production by Aspergillus niger. They observed that increasing sodium nitrate concentration from 0.6 to 0.8 w/v was accompanied by an increase in tannase activity from 9.7 U/ml to 19.4 U/ml. Banerjee and Pati (2007) determined the specificity of nitrogen sources (ammonium chloride, sodium nitrate, ammonium nitrate, ammonium di-hydrogen phosphate, di-ammonium hydrogen phosphate, potassium nitrate and ammonium sulphate) on tannase production by Aureobasidium pullulans under submerged culture technique using Czapek-Dox’s medium containing 1% tannic acid as sole carbon source. They indicated that most of the used nitrogen sources were effective for tannase production and the maximum tannase synthesis was obtained in the presence of 0.3% di-ammonium hydrogen phosphate. Moreover, they arranged the enzyme production by fungus in relation
30
to nitrogen sources in the following descending order: di-ammonium hydrogen phosphate, ammonium di-hydrogen phosphate, ammonium chloride, ammonium nitrate.
B-Inorganic and organic sources Bradoo et al. (1997) studied the use of various inorganic and organic nitrogen sources (sodium nitrate, potassium nitrate, ammonium nitrate, ammonium chloride, urea, aspartic acid, palmetic acid) into Czapek-Dox’s medium on the production of tannase by Aspergillus japonicus under submerged fermentation technique. They noted that the sodium nitrate was the ideal nitrogen source for both growth and enzyme production and the maximum tannase was obtained at 0.2% sodium nitrate. The effect of various nitrogen sources (peptone, brew's cream, urea, potassium nitrate and sodium nitrate) on the Aspergillus SH6 tannase production was studied in a medium containing valonia tannins (ellagitannins) as a substrate. Results indicated that organic nitrogen sources were favorable to the degradation of tannins, especially peptone as nitrogen source, which delivered high tannase activity (Huang et al., 2005). Sabu et al. (2005 b) investigated the effect of supplementation of different inorganic and organic nitrogen sources on tannase production by Aspergillus niger 16620 under solid state fermentation (SSF) using palm kernel cake and tamarind seed powder as carbon sources. Results showed that all tested compounds (peptone, yeast extract, malt extract, corn steep liquor, ammonium nitrate, ammonium chloride, sodium nitrate and potassium nitrate compared with control) exerted harmful impact on tannase production in case of palm kernel cake medium. They suggested that palm kernel cake had enough nitrogen required for the activity of the culture and supplementation of additional nitrogen sources resulted in imbalanced C/N ratio, affecting the cultures activity harmfully. However, in case of tamarind seed powder medium it was the opposite observation. All tested compounds irrespective of their organic or inorganic nature resulted in marginal increase in tannase activity and potassium nitrate was the most effective among all at 1% concentration. Battestin and Macedo (2007) investigated the impact of supplementing the fermentation medium (50:50, coffee husk: wheat bran) with different external nitrogen sources at 0.4-1.2 % concentrations on Paecilomyces variotii tannase production using solid state fermentation. The results revealed that among the tested nitrogen sources (ammonium nitrate, sodium nitrate, and yeast extract), ammonium nitrate was the most suitable as it resulted in highest enzyme activity while other nitrogen sources inhibited enzyme formation by the fungal culture. According to Kumar et al. (2007) the addition of ammonium sulphate (0.2%) as external nitrogen source decreased Aspergillus rubber tannase production under SSF using jamun leaves as solid medium. Recently, Manjit et al. (2008) found that the supplementation of the Jamun leave medium with 0.2% ammonium sulfate as nitrogen source increased Aspergillus fumigatus tannase titer under SSF conditions.
31
Rodrigues et al. (2007) studied the effect of supplementation with different organic (peptone and yeast extract) and inorganic (ammonium nitrate and ammonium sulfate) nitrogen sources on Aspergillus oryzae tannase production by solid state fermentation using cashew apple bagasse as solid support. They observed that ammonium nitrate, peptone, and yeast extract exerted no influence on tannase production while ammonium sulphate at a concentration of 1% improved the enzyme production in 1.43-fold compared with control.
3.7.2 Environmental conditions The growth of fungi and their tannase yield are highly depended on environmental conditions (Kar et al., 2002). The environmental conditions (fermentation time, temperature, pH, inoculum size and agitation) have been studied to optimize the production of tannase by various fungi (Hadi et al., 1994; Chaterjee et al,. 1996; Kar and Banerjee, 2000; Kar et al., 2002; Belmares-Cerda et al., 2003; Banerjee et al., 2005; Sabu et al., 2006).
3.7.2.1 The effect of fermentation time The fermentation time needed for obtaining maximum tannase production by fungi varied from fungal strain to another as shown in Table (3.3). In general, the optimum fermentation time ranged from 24 to 150 h. The type of tannase produced by the same fungus affects the suitable fermentation time for maximum production. Banerjee et al. (2001) found that the maximum production of Aspergillus aculeatus intracellular tannase was obtained after 24 h of incubation under SmF using tannic acid broth medium while 36 h was the optimum for extracellular tannase under the same conditions. Rana and Bhat (2005) revealed that maximum yield of Aspergillus niger Van Tieghem tannase was obtained after 48 and 120 h for extracellular and intracellular tannase, respectively under SmF using modified mildew test medium containing 2% tannic acid. Moreover, Kar and Banerjee (2000) confirmed that the optimum fermentation period depended on the applied fermentation technique. They found that maximum Rhizopus oryzae tannase production was attained under SmF technique after 48 h of incubation while, it was attained after 72 h when the production was carried out under SSF and MSSF techniques. In addition, many investigations revealed that fungal tannase production was peaked at the optimum incubation period. Afterwards tannase production decreased gradually (Hadi et al., 1994; Huang et al., 2005; Kumar et al., 2007; Rodrigues et al., 2007). This might be due to catabolic repression and substrate scarcity in the medium (Hadi et al., 1994) or due to scarcity in nitrogen source (Huang et al., 2005). Additionally, it may be owing to accumulation of toxic metabolites secreted during fermentation (Hadi et al., 1994; Kumar et al., 2007; Rodrigues et al., 2007)
32
48 (extracellular)
96 (extracellular)
48-72 (extracellular)
Aspergillus oryzae
Aspergillus ruber
Aspergillus SH6
120 for intacellular
48 (extracellular) 48 (Intracellular) and 96 (extracellular) 120 (Intra) and 48 (extra)
Aspergillus japonicus
Aspergillus niger Van Tieghem
72(extracellular)
The optimum fermentation time (h) 24 (intracellular) and 36 (extracellular)
Aspergillus foetidus
Aspergillus aculeatus
Microbial source of tannase
33
Huang et al. (2005)
Kumar et al. (2007)
SSF using different tannin rich substrates like ber leaves (Zyzyphus mauritiana), jamun leaves (Syzygium cumini), amla leaves (Phyllanthus emblica) and jawar leaves (Sorghum vulgaris). SmF using buffered inorganic salt solution medium containing valonia tannins as carbon source.
Rodrigues et al. (2007)
Rana and Bhat (2005)
SSF using cashew apple bagasse.
LSF using Mildew Test (MT)-basal medium containing 2% tannic acid.
SmF using mildew test medium containing 2% tannic acid.
SSF using PUF impregnated with Mildew Test (MT)-basal medium.
Bradoo et al. (1997)
Mukherjee and Banerjee (2004)
MSSF using powdered fruits of Terminalia chebula (myrobalan) and Caesalpinia digyna (teri pod) cover as solid supports and substrates. SmF using Czapek-Dox's minimal medium
Banerjee et al. (2001)
Reference
SmF using tannic acid broth medium
Fermentation technique and medium
Table (3.3): The optimum fermentation time for tannase production by various fungi.
Co culture of Aspergillus foetidus and Rhizpous oryzae
Rhizopus oryzae
Aureobasidium pullulans
Continued
48 (extracellular)
34
MSSF using powdered fruits of Terminalia chebula and powdered pod cover of Caesalpinia digyna.
MSSF using powdered fruits of Terminalia chebula (myrobalan) and Caesalpinia digyna (teri pod) cover as solid supports and substrates.
SmF using modified Czapek-Dox’s minimal medium with 2% Caesalpinia digyna seed cover powder.
48 (extracellular)
60 (extracellular)
SSF conditions using wheat bran and tannic acid as substrate.
150 (extracellular)
SSF and MSSF using Caesalpinia digyna seed cover powder as substrate.
SmF using Czapek-Dox’s minimal medium.
120 (extracellular)
72 (extracellular)
SmF using tannic acid broth medium.
36 (extracellular)
Banerjee et al. (2005)
Mukherjee and Banerjee (2004)
Kar and Banerjee (2000)
Chatterjee et al. (1996)
Hadi et al. (1994)
Banerjee et al. (2007) and Banerjee and Pati (2007)
3.7.2.2 The effect of temperature Fermentation temperature affects microbial cellular growth, spore formation, germination, microbial physiology, and thus product formation. Therefore, maintenance of an optimal process temperature is considered one of major factors which affect the economics of the microbial enzymes production process (Banerjee et al., 2005). Different optimum incubation temperatures were used by researchers for the production of tannase by various fungi as shown in Table (3.4). The used temperatures ranged from 25°C to 42°C. The optimum temperature for tannase production by the same microorganism differed according to the applied fermentation technique as reported by Kar and Banerjee (2000). They found that the optimum temperature of Rhizopus oryzae tannase production under solid state fermentation (SSF) and modified solid state fermentation (MSSF) was 32°C but under submerged fermentation (SmF) the optimum temperature was 37°C.
3.7.2.3 The effect of initial pH of production medium: The initial pH of the medium is of great significance since it affects the rate of tannase production (Chaterjee et al., 1996). Different pH values were used for various fungi by several investigators. The used pH values were ranged from 3.8 to 6.6 (Hadi et al., 1994; Lekha et al.,1994; Chaterjee et al., 1996; Bradoo et al., 1997; Kar et al., 2002; Belmares-Cerda et al., 2003; Mukherjee and Banerjee, 2004; Sharma et al., 2007; Manjit et al., 2008; Enemuor and Odibo, 2009).
Aspergillus spp. Mukherjee and Banerjee (2004) reported that pH 5.0 was the optimum for tannase production by Aspergillus foetidus isolated from soil of acidic nature, under modified solid state fermentation using Czapek-Dox’s medium containing various tannin-rich natural substrates. Manjit et al. (2008) mentioned that Aspergillus fumigatus produced maximum tannase activity at pH 5.0 under solid state fermentation using different tannin-rich agro forest residues moistened by mineral salts solution. Bradoo et al. (1997) noticed that the highest Aspergillus japonicus tannase secretion in Czapek-Dox’s minimal medium under liquid surface fermentation was attained at pH 6.6.
35
Table (3.4): The optimum temperatures for tannase production by various fungi.
Microbial source of tannase
The optimum temperature (°C )
Reference
Aspergillus fumigatus MA
25
Manjit et al. (2008)
Aspergillus japonicus
30
Bradoo et al. (1997)
Aspergillus niger PKL 104
28
Lekha et al. (1994)
30
Bhat et al. (1997)
30
Belmares-Cerda et al. (2003)
Aspergillus niger ATCC 16620
30
Sabu et al. (2005 b)
Aspergillus SHL6
35
Huang et al. (2005)
Aspergillus ruber
30
Kumar et al. (2007)
Aspergillus niger
Paecilomyces variotii
29-34
Battestin and Macedo (2007)
42
Chaterjee et al. (1996)
32 on SSF and MSSF
Rhizopus oryzae
Kar and Banerjee (2000) 37 on SmF
Co-culture of Rhizopus oryzae and Aspergillus foetidus
36
32
Kar et al. (2002)
30
Mukherjee and Banerjee (2004) and Banerjee et al. (2005)
Pourrat et al. (1982) studied the effect of initial pH of the medium (3% tannic acidcontaining mineral medium) on Aspergillus niger tannase production. They revealed that no significant tannase activity was found in the culture medium where initial pH was less than 4 whereas, between pH 4 and 7 the activity rose with increasing pH up to pH 7. Lekha et al. (1994) studied the effect of initial pH of production medium (wheat bran supplemented with tannic acid) on tannase production by Aspergillus niger. They found that maximum yield of tannase was attained at initial pH 6.5. Bhat et al. (1997) tested pH range of 3.5-6.5 for the elaboration of tannase by Aspergillus niger Van Tieghem in Czapek yeast extract medium. They found that pH 5.0 was the optimum. Belmares-Cerda et al. (2003) noticed that Aspergillus niger PSH produced its maximum tannase at pH 5.5 under submerged fermentation technique using minimal medium containing tannin-rich natural substrates. Sharma et al. (2007) reported that the initial pH of the fermentation medium was identified as one of the important process parameters affecting cell growth and tannase synthesis by Aspergillus niger in modified Czapek-Dox’s medium. They mentioned that the highest enzyme synthesis was obtained at pH 5.0. Kumar et al. (2007) investigated the effect of pH on tannase production by Asergillus ruber under solid state fermentation using different tannin-rich substrates. The results revealed that the maximum enzyme productivity was attained at initial pH of 5.5. Enemuor and Odibo (2009) reported that pH 3.8 was the optimum for tannase production by Aspergillus tamarii under SmF technique using tannic-acid containing mineral medium.
Penicillium variable Statistical optimization of extracellular tannase production by Penicillium variable using Modified Czapek-Dox’s medium containing powdered Chebulic myrobalan as a substrate were carried out under submerged culture fermentation (Saxena and Saxena, 2004). Results revealed that the pH in the tested range (2.0-8.0) had no statistical significant effect on tannase production and the maximum tannase production was attained at pH 5.0. Rhizopus oryzae Hadi et al. (1994) studied the production of Rhizopus oryzae tannase in modified Czapek-Dox’s medium at different acidic pH. They found that the maximum synthesis of the tannase was observed at initial pH 5.0. Chaterjee et al. (1996) investigated the effect of pH on synthesis of Rhizopus oryzae tannase under solid state fermentation using CzapekDox’s medidum containing tannic acid as sole carbon source. They observed that tannase synthesis was negligible at pH 4.5, but the maximum synthesis of the enzyme was recorded at pH 5.0. Thereafter the decline trend was obtained. Misro et al. (1997) tested the influence of the initial pH of modified Czapek-Dox's medium in the range of 3.0-6.0 on the productivity of tannase by immobilized Rhizopus oryzae. The results showed that at pH 5.0 the enzyme productivity was found to be the maximum and beyond it the secretion of the enzyme stopped. Kar et al. (2002); Mukherjee and Banerjee (2004) mentioned that
37
pH 4.5 was the optimum for the production Rhizopus oryzae tannase and gallic acid under modified solid state fermentation using modified Czapek-Dox’s medium containing various plant materials. Banerjee et al. (2005) studied the effect of initial pH of fermentation medium on tannase production by co-culture (Rhizopus oryzae and Aspergillus foetidus) using solid state fermentation (SSF). The results showed that the optimum pH of the medium was 5.0, up to which there was an increase in enzyme activity followed by a decrease.
3.7.2.4 The effect of inoculum size Belmares-Cerda et al. (2003) studied the tannase production kinetics in submerged cultures of Aspergillus niger PSH. They reported that the optimum inoculum density for maximum extracellular and intracellular tannase activity was 3×107 spores per reactor containing 30 ml mineral salts medium containing tannin-rich natural substrates under submerged fermentation technique. Ramirez-Coronel et al. (2003) inoculated the liquid medium (mineral salts solution) to be absorbed by SSF support (PUF) with 5×107 spores per g of carbon source as the optimum inoculum size for maximum production of Aspergillus niger Aa-20 tannase. Aissam et al. (2005) mentioned that the highest tannase synthesis (0.65 U/ml) by Aspergillus niger HA37 using olive mill waste water under submerged culture was achieved at 107 conidiospors/flask (50ml). Sabu et al. (2005 b) investigated the effect of inoculum size on Aspergillus niger ATCC 16620 tannase production under solid state fermentation using palm kernel cake and tamarind seed powder as substrates. Results revealed that in palm kernel cake medium, the yield of tannase increased with the increase in inoculum size and maximal tannase activity (3.94 U/g of dry substrate) was recorded with an inoculum level of 33×109 spores/5g of substrate. Whereas, in tamarind seed powder medium, there was an increase in tannase secretion with increasing inoculum level up to spore concentration of 11×109 /5g of substrate, and further increase in inoculum size reduce the enzyme production. A statistical optimization of extracellular tannase production from Penicillium variable using a natural source of tannins (chebulic myrobalan fruits powder) was carried out under SmF. Results showed that the inoculum size within the range of 5×106- 5×108 spores/50 ml medium had no significant effect on tannase production and the maximum tannase secretion was obtained at 5×107 spores/50 ml medium (Saxena and Saxena, 2004). Rodrigues et al. (2008) studied the effect of inoculum concentrations (104 to 107 spores/g) on tannase production by Aspergillus oryzae under SSF technique using cashew apple bagasse medium. They found that the yield of tannase increased as the inoculum size increased, and the maximal activity and productivity attained when the bagasse was inoculated with 107 spores/g.
38
3.7.2.5 The effect of agitation Hadi et al. (1994) studied the effect of oxygenation on Rhizopus oryzae tannase production in modified Czapek-Dox’s medium under various conditions (shaking, intermittent shaking, and stationary). Results showed that tannase production was at maximum under shaking condition using SmF technique. On the other hand, Kar and Banerjee (2000) investigated the effect of agitation on Rhizopus oryzae tannase production under modified solid state fermentation using medium of the Caesalpania digyna powder impregnated with modified Czapek-Dox’s medium. The results revealed that agitation is unfavorable for tannase synthesis and that maximum tannase secretion was obtained after 72 h of incubation under stationary condition using MSSF. Aureobasidium pullulans tannase production using submerged culture was studied under shaking and static conditions. It was found that the shaking was suitable for growth and tannase production and the maximum tannase production attained at speed of 120 r.p.m. using a mineral medium contained tannic acid as sole carbon source (Banerjee and Pati, 2007).
3.7.2.6 Interactions between factors affecting tannase production
It is of importance to note that the independent factors may show only a main influence on tannase production without depending on any other factor effect. In fact, it would be best and simple if all the independent factors influence the production of tannase in their own ways, and irrespective of the level at which the other factors may be present. But in reality, the effect of each on the tannase production may be influenced by the change in the levels of one or more factors. This phenomenon falls under the category of interactions. Although the importance of studying the interactions between factors influencing tannase production, there are few researches about this point (Lekha et al., 1994). Lekha et al. (1994) studied the effects of five independent factors [initial pH (3.57.5), fermentation temperature (20-35 °C), initial moisture content (30-70%), inoculum ratio (5-20%) and fermentation time (1-7 days)] on tannase production by Aspergillus niger PKL 104 under solid state fermentation using response surface methodology. They found a positive interaction between the initial pH and initial moisture level but no interaction between initial pH and other factors. Also they confirmed that initial moisture level and inoculum ratio showed negative interaction in contrast to positive interaction between inoculum ratio and fermentation period. Also Sharma et al. (2007) studied the interaction between four independent factors (%tannic acid; %sodium nitrate; agitation rate, and Incubation time) affecting production of tannase by Aspergillus niger under SmF. The statistical study was carried out using the rotatable central composite design (RCCD) method. Results confirmed the presence of only an interaction between agitation rate and % tannic acid at constant sodium nitrate concentration (0.8 %).
39
40
4. MATERIALS AND METHODS
4.1 Materials
4.1.1 Chemicals The chemicals used throughout the study were of analytical reagent grade and obtained from Sigma, Aldrich, Merck and VEB Laborchemie Apolda (Germany); BDH and JUDEX chemicals (England); Biolife (Italy); Chemapol (Czechoslovakia); El-Nasr Pharmaceutical Chemicals Co. “ADWIC” and United Company for Chemical and Medical Preparations “UCCMA” (Egypt).
4.1.2 Media and media constituents Peptone, yeast extract and beef extract were obtained from Merck (Germany). Agar agar (No. 1) was obtained from LAB MTM (England). Brain heart infusion agar was purchased from Oxoid (England). Distilled water was used throughout the work. Several plant raw materials as well as industrial wastes which incorporated in the media for enzyme production were obtained from the following sources: Peels of pomegranate (Punica granatum) were kindly supplied by the post harvest Lab., Faculty of Agriculture, Alexandria. Wheat bran, persimmon (Diospyrus kaki) fruits, Soybeans, cotton seeds and flax seeds were purchased from Alexandria local market. River-Red-gum (Eucalyptus camaldulensis Dehn) leaves and unripe dates were obtained from Abees farm, Alexandria. Wheat bran was kept directly in tightly closed Kilner jars and maintained at room temperature in dry place. While, pomegranate peels, persimmon fruits, unripe date fruits and river-red-gum leaves were washed with distilled water twice. After cutting or slicing they were dried at 60º C in hot air oven for 48 hours. The dried materials were then finely milled and sieved in 80 mesh screen. The obtained powder was then kept in tightly closed Kilner jars and maintained at room temperature in dry place. Soybean and seeds of cotton and flax were ground, defatted by hexane and kept in Kilner jars.
4.1.3 Solid supports for solid state fermentation (SSF) Two synthetic solid supports were used in the solid state fermentation technique. Polyurethane foam (PUF), bulk density 15 kg/m3 was kindly supplied by Foam Industrial Company, Egypt, and synthetic sponge was obtained from Taki Vita Company, Egypt.
41
4.1.4 Sources for isolation of tannase-producing microorganisms Nineteen samples collected from various natural sources (9 samples of mouldy tannins-rich plants, 8 samples of soil down tannin-rich plants, 1 sample of tannery soil and one sample of tannery effluent) were used to isolate tannase-producing microorganisms (Table 4.1). Mouldy pomegranate fruits, persimmon fruits and tea (7, 8 and 13, respectively, Table 4.1) were obtained from local market, Alexandria. Mouldy bengal fig fruits (14) were obtained from Antoniadis Park, Alexandria, Egypt. Tannery effluent as well as tannery soil (1, 9) were obtained from El-Max tannery region, El- Max, Alexandria, Egypt. Soil samples (2, 3, 4, 5, 12, 15, 17, and 19) were collected from different locations in Alexandria and El-Beheira Governorates. Mouldy leaves of pomegranate and guava trees (16 and 18 respectively) were gathered from El-Kilo 59Alexandri-Cairo desert road. Mouldy river-red-gum leaves and mouldy dates (6 and 10) were obtained from Abees farm-Alexandria. All samples were collected and transported to laboratory either for direct use or preserved in refrigerator at 4º C until use. Date, source and location of isolation as well as the codes of isolates are presented in Table (4.1).
4.1.5 Media 4.1.5.1 Media for isolation and primary screening Tannic acid agar medium (TAA): TAA was used for the isolation of tannase-producing microorganisms. The pH of the medium was adjusted at 4.5±0.2 or 7±0.2 for isolation of fungi or bacteria, respectively. It was prepared from the following components (g/L): tannic acid (10), NaNO 3 (3), KCl (0.5), MgSO 4 .7H 2 O (0.5), KH 2 PO 4 (1.0), FeSO 4 .7H 2 O (0.01), agar (30) (Pinto et al., 2001). TAA at pH 4.5± 0.2 was also used for the primary screening of fungal isolates. Brain-heart infusion agar: Brain heart infusion agar (Oxoid) was used to carry out the primary screening of tannic acid-degrading bacterial cultures. Solidified plates of Brain heart infusion agar were overlaid by 2% filter sterilized tannic acid solution as described by Osawa (1990).
4.1.5.2 Medium for secondary and final screening of fungi The modified Czapek-Dox’s minimal medium (basal medium) was used for secondary and final screening of fungi (Bradoo et al., 1996). It was prepared from the following ingredients (g/L): tannic acid (10), NaNO 3 (6), KCl (0.52), MgSO 4 .7H 2 O (0.52), KH 2 PO 4 (1.52), FeSO 4 .7H 2 O (0.01), ZnSO 4 .7H 2 O (0.01), Cu(NO 3 ) 2 .3H 2 O (0.01), pH 4.5± 0.2.
42
Table (4.1): Isolation sources of tannase-producing microorganisms. Serial No.
Date of isolation
1
15/2/2005
Tannery effluent.
2
24/2/2005
Muddy soil sample down palm trees.
3
26/2/2005
Muddy soil sample down river-red-gum trees.
The entertainment forest gardens, Alexandria, Egypt.
4
5/3/2005
Sandy soil sample down palm trees.
5
5/3/2005
Sandy soil sample down palm trees.
Edko city, Beheira Governorate, Egypt. Borg Rosetta, Beheira Governorate, Egypt.
Mouldy river-redgum (Eucalyptus camaldulensis Dehn) leaves. Mouldy pomegranate (Punica granatum) fruits.
Location El-Max tannery region, El- Max, Alexandria, Egypt. Boselly region, Beheira Governorate, Egypt.
Code No. of isolation source A
B
C
D
E
Abees farm, Alexandria-Egypt
F
Alexandria local market.
G
Mouldy persimmon (Diospyrus kaki) fruits.
Alexandria local market.
H
10/12/2005
Tannery soil sample.
El-Max tannery region, El- Max, Alexandria, Egypt.
I
25/12/2005
Mouldy date fruits sample.
Abees farm, Alexandria, Egypt.
J
Mouldy peels of pomegranate fruits.
Postharvest lab, Faculty of Agriculture, Alexandria University, Egypt.
K
6
15/3/2005
7
30/11/2005
8
2/12/2005
9
10
11
Isolation source
28/12/2005
43
Continue Faculty of Agriculture, Alexandria University, Egypt.
L
Alexandria local market.
M
12
13/3/2006
Soil sample from plant pots.
13
15/3/2006
Mouldy tea
14
17/3/2006
Mouldy bengal fig (Ficus benghalensis) fruits.
Antoniadis parks, Alexandria- Egypt.
15
17/3/2006
Soil sample down Bengal fig trees.
Antoniadis parks, Alexandria, Egypt.
16
10/4/2006
17
10/4/2006
18
10/4/2006
19
10/4/2006
El-Kilo 59Alexandri-Cairo desert road. El-Kilo 59Soil sample down riverAlexandri-Cairo red-gum trees. desert road. El-Kilo 59Mouldy guava (Psidium Alexandri-Cairo guajava) leaves. desert road. El-Kilo 59Soil sample down Alexandri-Cairo guava trees; desert road. Mouldy pomegranate leaves.
44
N
O
P
Q
R
S
4.1.5.3 Media for tannase production Modefied Czapek-Dox’s minimal medium was used for tannase production under submerged fermentation (SmF) and solid-state fermentation (SSF) techniques using various tannins sources [pure tannic acid (basal medium) or tannins-containing plant materials] (Bradoo et al., 1996 and Aguilar et al., 2002).
4.1.5.4 Media for maintenance of isolates
Potato dextrose agar (PDA) supplemented with 0.01% tannic acid: PDA supplemented with 0.01% tannic acid was used for maintaining fungal isolates. It was prepared from the following ingredients (g/L): glucose (20), infusion of 200 g potatoes; agar (15), tannic acid (0.1), pH 5.6± 0.2 (Bajpai and Patil, 1997 and Atlas, 2006).
MRS Broth (De Man, Rogosa, Sharpe Broth) with 20% glycerol:
MRS broth was used for maintaining bacterial isolates. It contained (g/L): glucose, (20), peptone (10), beef extract (8), sodium acetate.3H 2 O (5), yeast extract (4), K2 HPO 4 (2), tri-ammonium citrate (2), MgSO 4 ·7H 2 O (0.2), MnSO 4 ·4H 2 O (0.05), tween 80 (1 ml), glycerol (200), pH 6.2 ± 0.2. Glycerol was added as a cryogenic protecting agent (De Man et al., 1960; Stanbury et al., 2003; Moldenhauer, 2008).
45
4.2 Methods
4.2.1 Isolation of tannase-producing microorganisms Each isolation source sample (Table 4.1) was serially diluted (10-1 to 10-4) with sterile distilled water. One milliliter from each dilution was plated into tannic acid agar medium (TAA) using pour plate technique. Incubation was carried out at 30º C for 96 h under aerobic conditions for isolation of fungi and anaerobic conditions for isolation of bacteria. Microorganisms capable to grow and form clearing zone around its colonies due to the hydrolysis of tannin were selected and purified (Murugan et al., 2007). The pH of the modified Czapek-Dox’s minimal medium as well as 1% tannic acid solution were separately adjusted to 4.5±0.2 for isolation of fungi and 7.0±0.2 for isolation of bacteria. The medium was sterilized separately without tannic acid at 121º C for 15 min. Then mixed with filter sterilized tannic acid solution using MILLEX®-OR membrane filter (33mm diameter, 0.22µm pore size, Millipore, France).
4.2.2 Purification of tannase-producing isolates Single colony of each positive isolate was picked up, transferred to a test tube containing 3 ml of sterile distilled water and mixed for 5 min using vortex. A loopfull of this suspension was streaked on a plate containing TAA medium before incubation at 30º C for 4 days. These steps were repeated until pure culture was obtained, i.e. all colonies on a plate had the same cultural characteristics and the same results of microscopic examination of the original picked colony. Subsequently, these steps were repeated again at least 3 times to confirm the purity of the isolate (El-Banna, 1976).
4.2.3 Maintenance of tannase-producing isolates Maintenance of fungal cultures Single colony from the pure culture plate was grown on slant of potato dextrose agar (PDA) supplemented with 0.01% tannic acid and stored at 4°C under sterilized mineral oil (paraffin oil) as stock culture and sub-cultured every year (Stanbury et al., 2003). While working culture was stored at 4°C without adding mineral oil and subcultured every alternate month (Bajpai and Patil, 1997).
Maintenance of bacterial cultures Single colony, from the pure culture plate, was propagate in MRS medium containing 20% (v/v) of glycerol and stored at -20º C. Working and stock cultures were sub-cultured every month and year, respectively (De Man et al., 1960; Stanbury et al., 2003; Moldenhauer, 2008).
46
4.2.4 Screening and selection of tannase-producing fungal cultures 4.2.4.1 Primary screening Primary screening for highest tannase producers was carried out on tannic acid agar plates (TAA) as described by Bradoo et al. (1996). The plates were point inoculated with the isolate and incubated at 30ºC. The diameter of Clear zones (including the colonies diameters) formed due to hydrolysis of tannic acid around the fungal colonies were measured after 72 and 96 h of incubation, then compared in order to choose the highest tannase producers.
4.2.4.2 Secondary and final screening Fungal cultures which exhibited high tannase production in the primary screening were subjected to secondary screening as described by Bradoo et al. (1996) and Batra and Saxena (2005). Approximately 5 × 107 spores of each of potent tannase-producing fungi were inoculated into 250 ml-Erlenmeyer flasks containing 50 ml of sterilized modified CzapekDox’s minimal medium (pH 4.5± 0.2). Filter-sterilized tannic acid solution (adjusted at pH 4.5± 0.2) was added to the autoclaved medium (pH 4.5± 0.2) to provide a final concentration of 1% (w/v) tannic acid in the medium. Cultures were grown at 30º C for 96 h and shaked intermittently three times a day at 200 rpm for 2 min each time in an orbital incubator (Model INR-200, Gallenkamp, UK). At the end of cultivation time the extracellular and intracellular tannase activity per flask were assayed. The promising fungal cultures that selected from the secondary screening were subjected to final screening using above mentioned method. Samples were withdrawn at regular intervals of 24 h and assayed for extracellular and intracellular tannase activities.
4.2.5 Inoculum preparation Inoculum was prepared by the method described by Ramirez-Coronel et al. (2003). The fungal isolate was cultivated on PDA slant then incubated for 4 days at 30°C until a good sporulation was obtained. Spores were then scraped into a sterile 0.02% Tween 80 solution and counted in a Neubauer chamber (Hemocytometer slide) by the method described by Harisha, (2007). The suitable volume which contains the needed number of spores was calculated.
4.2.6 Harvesting the enzyme Harvesting the extracellular tannase The fermentation medium was filtered through Whatman No.1 filter paper. The obtained filtrate was used for extracellular tannase determination (Gupta et al., 1997).
47
Harvesting the intracellular tannase The method described by Sharma, et al. (2000) was adopted. In which a 10% suspension of the mycelial mass was made in 0.05 M citrate buffer (pH 5.0) and then frozen overnight at -20°C. Acid washed sand was added (the amount of sand was four times of the weight of the mycelium), and the mixture was manually ground for about 5 min in a chilled pestle-mortar kept in an ice bath. The homogenate was centrifuged at 5000 ×g for 30 min at 4°C using KK centrifuge (Gemmy Industrial Corporation, Taiwan). The supernatant was used for the intracellular tannase assay.
4.2.7 Biomass determination Biomass was separated by filtration through Whatman No.1 filter paper, and quantified by dry weight after drying in hot air oven at 105 ºC for 4 h according to the method described by Aguilar et al. (2002).
4.2.8 Identification of the most promising fungal isolate The cultural characteristics of the selected fungal isolates on PDA plate (the form, elevation and margin shapes of the fungal colony) were recorded and compared with descriptions, illustrations and photos of genera reported by microbial database of doctorfungus website (http://www.doctorfungus.org/ thefungi/ Aspergillus_niger.htm). The selected fungal isolate was also examined microscopically. The slide agar method described by Benson (2001) was used for preparation of tested slides. Slides were examined microscopically under low and high power after staining with lactophenol cotton blue stain. The morphological characteristics (type and appearance of hyphae, long, diameter, and appearance of conidiophores; colour, shape, diameter and type of spores) then compared with descriptions, illustrations and photos of fungal genera reported by Barnett and Hunter (1987), Benson (2001) and microbial database of doctorfungus website (http://www.doctorfungus.org/thefungi/ Aspergillus_niger.htm). For confirmation, the fungal isolate was kindly re-identified by the mycological center (Faculty of science, Assiut University, Egypt).
4.2.9 Optimization of conditions controlling tannase production In an attempt to optimize the conditions controlling tannase production by the selected and most promising fungal isolate, several experiments were considered. One variable at a time (O.V.A.T) method was used in the first step of optimization to select among each of the following qualitative variables: fermentation technique [liquid surface fermentation (LSF), submerged fermentation (SmF) and solid state fermentation (SSF)], agitation condition (continuous shaking, intermittent shaking and static condition), type of tannins source (pure tannic acid and dried powders of each of : pomegranate peels, persimmon fruits, river-red-gum leaves, and unripe date fruits), type of nitrogen source
48
(organic and inorganic sources) and the effect of divalent cations (Hg2+, Mg2+, Cd2+, Cu2+ , Ca2+, Zn2+, Fe2+, Pb2+). Quantitative variables were analyzed using split plot design and response surface methodology using rotatable central composite design (RCCD). Split plot design was used in the second step of optimization, which involved screening of fermentation temperature and fermentation time. The last step of optimization was carried out to determine the optimal level of the following variables: tannic acid concentration, nitrogen source concentration, pH value, and inoculum size using rotatable central composite design.
4.2.9.1 Fermentation techniques 4.2.9.1.1 Liquid surface fermentation (LSF) Sterilized modified Czapek-Dox’s minimal medium (pH 4.5±0.2) containing 1% filter sterilized tannic acid was distributed in 50 ml portions into 250 ml-Erlenmeyer flaskes. Each flask was inoculated with approximately 5 × 107 spores of Aspergillus niger Van Tieghem. Incubation of flasks was carried out statically at 30º C for 96 hours (Bradoo et al., 1997). 4.2.9.1.2 Submerged fermentation (SmF) The aforementioned cultivation conditions were adopted except the incubation was carried out by shaking in an orbital incubator (Model INR-200, Gallenkamp, UK) either continuously at 120 rpm or by intermittent shaking three times a day at 200 rpm for 2 min each time. Incubation of flasks was carried out at 30º C for 96 h (Hadi et al., 1994 and Saxena and Saxena, 2004).
4.2.9.1.3 Solid state fermentation (SSF) Wheat bran and two synthetic supports [polyurethane foam (PUF) and sponge] were used separately. Solid state fermentation using wheat bran as solid support was carried out according to the procedure of Pinto (2001) with some modifications. Sterilized 250 ml-conical flasks containing 20 g of wheat bran moistened with 20 ml of sterilized modified Cazpek-Dox’s minimal medium (pH, 4.5±0.2) containing 1% filter sterilized tannic acid were inoculated with 5 × 107 fungal spores and then incubated at 30° C for 96 h without shaking. In case of synthetic solid supports, the procedure described by Ramirez-Coronel et al. (2003) was adopted. Polyurethane foam and sponge were cut into 0.5 cm cubes, washed once with cold water followed by warm water and dried at 60°C. Erlenmeyer flasks (500 ml-capacity) containing either 5 gm of PUF or sponge impregnated with 50 ml of sterilized modified Cazpek-Dox’s minimal medium (pH 4.5±0.2) containing 1% filter sterilized tannic acid and inoculated with 5 × 107 fungal spores. The flasks were incubated at 30°C for 96 h under static conditions.
49
The extracellular and intracellular enzymes produced in LSF and SmF were harvested by the methods mentioned in sections (4.2.6). While, the enzyme produced by SSF using either PUF or sponge as solid supports was obtained by compressing PUF or sponge separately in a Buchner funnel. The extract was then centrifuged (5000 ×g, 15 min) then the supernatant was used for extracellular tannase assay (Ramirez-Coronel et al., 2003). Enzyme produced by SSF using wheat bran was extracted from the fermented matter by adding 50 ml of 0.05 M citrate buffer (pH 5.0). The flasks were then shaked in an orbital shaker (200 rpm) for one h at 30°C. The crude enzyme was separated by filtration over Whatman No.1 filter paper. The filtrate was used for extracellular tannase assay (Pinto et al., 2001).
4.2.9.2 Fermentation temperature and fermentation time The effect of five fermentation temperatures (T) and five fermentation times (D) were studied simultaneously by a full factorial experiment using split plot design in order to study the main effect of each variable separately and the interaction between these two variables. Coded levels of each variable are shown in Table (4.2). Five incubators were adjusted at the studied temperatures (the temperature of each incubator was chosen randomly). At each incubator five experimental units (flasks) were incubated, one experimental unit for each fermentation time (Figure 4.1). This factorial experiment was repeated three times and the average of extracellular tannase activity was measured for each experimental unit. Each experimental unit was a 250 ml-Erlenmeyer flask containing 50 ml of autoclaved modified Czapek-Dox’s minimal medium (initial pH 4.5±0.2) containing 1% tannic acid (filter sterilized separately). Each flask was inoculated with approximately 5 × 107 of Aspergillus niger VanTieghem spores then incubated at the studied temperature (20, 25, 30, 35 or 40°C) with intermittent shaking in orbital shaker three times a day at 200 rpm for 2 min each time.
4.2.9.3 Concentrations of tannic acid and nitrogen source, initial pH and inoculum size The last step of optimization was carried out statistically for obtaining the optimum levels of the following four quantitative variables: Tannic acid and nitrogen source concentrations, initial pH value and inoculum size. The fermentation process was carried out in 250 ml-Erlenmeyer flasks each containing 50 ml of the autoclaved modified Czapek-Dox’s minimal medium. After inoculation with Aspergillus niger VanTieghem spores, flasks were incubated at 35º C for 96 h under intermittent shaking as described before. Flasks containing media with (X 1 ) of filter sterilized tannic acid, (X 2 ) of nitrogen source (sodium nitrate), (X 3 ) of pH value and (X 4 ) inoculum size were all illustrated.
50
Variables
(T 2 ,D 3 ) (T 2 ,D 4 ) (T 2 ,D 5 )
(T 1 ,D 3 ) (T 1 ,D 4 ) (T 1 ,D 5 )
D3
D4
D5
51
(T 2 ,D 2 )
(T 1 ,D 2 )
D2
T 1, T 2, T 3, T 4 and T 5 were 20,25,30,35 and 40º C respectively. D 1 , D 2 , D 3 , D 4 and D 5 were 24, 48, 72, 96 and 120 h respectively.
(T 2 ,D 1 )
(T 1 ,D 1 )
T2
D1
T1
(T 3 ,D 5 )
(T 3 ,D 4 )
(T 3 ,D 3 )
(T 3 ,D 2 )
(T 3 ,D 1 )
T3
(T 4 ,D 5 )
(T 4 ,D 4 )
(T 4 ,D 3 )
(T 4 ,D 2 )
(T 4 ,D 1 )
T4
Coded levels of Fermentation temperatures (º C)
(T 5 ,D 5 )
(T 5 ,D 4 )
(T 5 ,D 3 )
(T 5 ,D 2 )
(T 5 ,D 1 )
T5
Table (4.2): Coded levels and combinations between two variables, fermentation temperature (T) and fermentation time (D) in split plot experiment.
Coded levels of fermentation time (h)
Experimental design and Data analysis Response surface methodology (RSM) is a powerful technique for testing multiple process variables because fewer experimental trials are needed compared to the study of one variable at a time. Also, significant interactions between variables can be identified and quantified by this technique. Rotatable Central Composite Design (RCCD), which falls under RSM, was used to study the main effects (linear and quadratic) and interactions of these factors (Petersen, 1985). The statistical software package "STATESTICA 7.0", StatSoft, Inc., USA was used to analyze the experimental design. Each factor in this design was studied at five levels (–Į – Į $OO WKH YDULDEOHV ZHUH WDNHQ DW D FHQWUDO coded value considered as zero. For statistical calculations, the relation between the coded values and real values were as described in the following equation:
Where X i is the independent variable coded value, U i the real value of the independent variable, U o WKHUHDO YDOXHRIWKH LQGHSHQGHQWYDULDEOHRQWKH FHQWHUSRLQW DQG ǻU is the step change. The variables and levels are shown in Table (4.3). A five levels-four factors rotatable central composite design (RCCD) was adopted in this study, requiring 31 experiments, which included sixteen factorial points, eight star (axial) points, and seven central points to provide information about the interior of the experiment region, allowing evaluation for curvature. The minimum and maximum ranges of variables investigated and the full experimental plan with respect to their values in coded and actual forms are listed in Table (4.4). Upon completion of experiment, the tannase production was taken as dependent variable or response (Y).
Statistical analysis and modeling: The data of tannase production obtained was subjected to analysis of variance (ANOVA), appropriate to the design of experiments (Petersen, 1985) using the statistical software package "STATESTICA 7.0", StatSoft, Inc., USA. The mathematical relationship of the independent variables and the responses (tannase Units/flask) were calculated by the second-order polynomial equation i.e. Y=
ȕ 0 + ȕ 1 X 1 + ȕ 2 X 2 + ȕ 3 X 3 + ȕ 4 X 4 + ȕ 11 X 1 2 + ȕ 22 X 2 2 + ȕ 33 X 3 2 + ȕ 44 X 4 2 + ȕ 12 X 1 X 2 + ȕ 13 X 1 X 3 + ȕ 14 X 1 X 4 + ȕ 23 X 2 X 3 + ȕ 24 X 2 X 4 + ȕ 34 X 3 X 4
Where Y = predicted response; ȕ 0 = intercept; ȕ 1 ȕ 2 ȕ 3 ȕ 4 = linear coefficients; ȕ 11 ȕ 22 , ȕ 33 ȕ 44 = quadratic coefficients; ȕ 12 , ȕ 13 ȕ 14 ȕ 23 ȕ 24 ȕ 34 = interaction coefficients.
53
Inoculum size
spores / 50 ml
X3
˰˰
pH value
X4
X2
g/L
Nitrogen source concentration
X1
Coded value
%
Units
Tannic acid concentration
Factors
54
5x107
107
4
4
1
-1
3
2
0.5
(-2)
Į˰
9x107
5
6
1.5
0
Table (4.3): Coded and actual levels of the variables in rotatable central composite design (RCCD).
1.3x108
6
8
2
+1
+Į
1.7x108
7
10
2.5
(+2)
Table (4.4): The full experimental plan with respect to their values in coded and actual forms. Coded combinations
-1 +1 -1 +1 -1 +1 -1 +1 -1 +1 -1 +1 -1 +1 -1
-1 -1 +1 +1 -1 -1 +1 +1 -1 -1 +1 +1 -1 -1 +1
-1 -1 -1 -1 +1 +1 +1 +1 -1 -1 -1 -1 +1 +1 +1
-1 -1 -1 -1 -1 -1 -1 -1 +1 +1 +1 +1 +1 +1 +1
16 17 18 19 20 21 22 23 24
+1 +1 +1 +1 -2 0 0 0 +2 0 0 0 0 -2 0 0 0 +2 0 0 0 0 -2 0 0 0 +2 0 0 0 0 -2 0 0 0 +2
25 26 27 28 29 30
0 0 0 0 0 0
0 0 0 0 0 0
0 0 0 0 0 0
0 0 0 0 0 0
31
0
0
0
0
Star points =2x4=8
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15
Factorial points =2 4 =16
X1 X2 X3 X4
Central points =7
Exp.unit No.
Actual combinations Tannic acid conc. (%) 1 2 1 2 1 2 1 2 1 2 1 2 1 2 1
Nitrogen source pH conc. value (g/L) 4 4 4 4 8 4 8 4 4 6 4 6 8 6 8 6 4 4 4 4 8 4 8 4 4 6 4 6 8 6
Inoculum size (spores/50ml) 5×107 5×107 5×107 5×107 5×107 5×107 5×107 5×107 1.3×108 1.3×108 1.3×108 1.3×108 1.3×108 1.3×108 1.3×108
2 0.5 2.5 1.5 1.5 1.5 1.5 1.5 1.5
8 6 6 2 10 6 6 6 6
6 5 5 5 5 3 7 5 5
1.3×108
1.5 1.5 1.5 1.5 1.5 1.5
6 6 6 6 6 6
5 5 5 5 5 5
1.5
6
5
9×107 9×107 9×107 9×107 9×107 9×107 9×107
55
9×107 9×107 9×107 9×107 9×107 9×107 107 1.7×108
The stepwise elimination procedure was applied to eliminate the non significant effects from the full model in order to generate the best describing model. The response surface three dimensional curves were used to describe the interaction effects of each two variables at zero levels of the other two variables.
4.2.10 Analytical methods
4.2.10.1 Estimation of tannic acid Protein precipitation method described by Hagerman and Butler (1978) was used for determination of either tannic acid in the medium or hydrolysable tannins in natural tannin sources. Two ml of bovine serum albumin (BSA) solution (1 mg/ml prepared in 0.2 M acetate buffer, pH 5.0 contains 0.17 M NaCl), were added to one ml of culture filtrate or alcoholic extract of natural substrate tannins (about 5 g of substrate powder extracted by 50 ml ethyl alcohol acidified by 0.1% HCl). After 10 min, the content of the tube was centrifuged at 5000 ×g for 10 min. The pellet was washed with acetate buffer, and dissolved in 4 ml of SDS/TEA solution {1% (w/v) sodium dodecyl sulphate and 5% (v/v) triethanol amine} and one ml of FeCl3 (0.01 M in 0.01 N HCl) was added. The mixture was incubated at 30 °C for 15 min and the absorbance was measured at 510 nm. Gradient concentrations of tannic acid (from 0.2 to 1 mg) were used for standard curve.
4.2.10.2 Enzyme assay
Tannase activity was assayed using the spectrophotometric method of Sharma et al. (2000) with some modifications. The method is based on the formation of chromogen between gallic acid (released by the action of tannase on methyl gallate) and rhodanine (2thio-4-ketothiazolidine). The reaction mixture in the blank, test, and control tubes contained 0.25 ml of substrate solution to which 0.25 ml of citrate buffer (pH 5) and 0.25 ml of the enzyme were added to the blank and test, respectively. The tubes were incubated at 30°C for 5 min, and 0.3 ml of methanolic Rhodanine (0.667% w/v) was added to all the tubes in order to stop the enzymatic reaction as well as formation of chromgen. Tubes were then kept at 30°C for 5 min. After this 0.2 ml of 1 N potassium hydroxide solution was added to each tube and these were incubated at 30°C for 5 min. This was followed by addition of the enzyme (0.25 ml) to the reaction mixture in the control tube only. Finally, each tube was diluted with 4 ml distilled water and incubated at 30°C for 10 min and the absorbance was recorded against water at 520 nm using SPEKOL 11 spectrophotometer (Carl Zeiss, Jena, Germany). The enzyme activity was calculated from the change in absorbance: ǻ A520 = (A test – A blank ) - (A control – A blank ). All the assays were carried out in triplicate.
One unit of the enzyme was defined as micromoles of gallic acid formed per minute by one ml of enzyme extract under optimum conditions of tannase activity.
56
4.2.10.3 Calibration curve for gallic acid estimation Gallic acid (0.5 mM) in citrate buffer (0.05 M, pH=5.0) was freshly prepared and used as a standard solution. Aliquots of gallic standard solution containing 5-100 nM gallic acid were taken and volume were made up to 0.5 ml with citrate buffer. Then, 0.3 ml of methanolic rhodanine solution (0.667%, w/v) was added to the tubes. After that, 0.2 ml of potassium hydroxide solution (1N) was added followed by incubation for 5 min at 30°C. Then 4 ml of distilled water was added to all tubes. Absorbance was measured at 520 nm, after 10 minutes (Sharma et al., 2000).
4.2.10.4 Determination of nitrogen source in the medium Improved Kjeldahl method No 955.04 of AOAC international (2000) for nitratecontaining samples was used with some modifications to follow-up the consumption of nitrogen source through the fermentation time. Half ml of 2N sodium hydroxide solution and 0.28 g of mercuric oxide were added to 20 ml of the medium filtrate then the volume was made up to 50 ml with distilled water in a 50 ml-volumetric flask. Five milliliters of the previous solution was placed in digestion flask and 4 ml of concentrated sulfuric acid and 0.2 g salicylic acid were added followed by occasionally shaking for 30 min. Then, 0.5 g of sodium thiosulphate and 0.2 g of zinc dust were added followed by shaking for 5 min. The mixture was heated over low flame until frothing ceases. This step was followed by addition of 0.07 g HgO and 1.5 g of anhydrous sodium sulphate. The digestion was continued for approximately 2 h until a clear solution was obtained. Then the distillation of nitrogen was carried out in a micro-Kjeldahl distillation unit to receive the distilled ammonia in 4% boric acid in the presence of Tashiro indicator then titration was carried out with a standard HCl solution.
4.2.10.5 Measurements of pH Measurements and adjustment of pH were carried out by a pH meter (WPA LITON, Cambridge, UK) standardized by potassium phosphate buffers pH 4 and 7.
4.2.10.6 Paper chromatographic analysis of fermented broth During enzyme production, periodically fermented broth (10 µl) was spotted on a sheet of cellulose (WHATMAN – 1 mm). Separation of compounds was carried out by ascending technique using 6% acetic acid as mobile phase. Visualization was carried out using 0.1% ferric chloride in 30% methanol as visualizing agent (Bradoo et al., 1997; Mondal and Pati, 2000).
57
58
5. RESULTS AND DISCUSSION
5.1 Isolation of tannase-producing microorganisms One hundred and seven isolates of tannase-producing fungi were isolated from nineteen samples collected from local environment at Alexandria and Beheira Governorates. Sources, types and numbers of isolates are presented in Table (5-1). Selection of the isolation sources was based on the fact that the presence of tannaseproducing microorganisms may fairly exist in tannins containing environments. 1. Tannery effluent and soils from a local tannery region contain remarkable amount of tannins. 2. Soils down high tannins-containing trees (river-red-trees, persimmon trees, pomegranate trees, Bengal fig trees, guava trees and palm trees). These soils are rich sources of tannins due to the leaching of tannins from fallen tannins-containing fruits.
3. Mouldy tannins-containing fruits (persimmon, pomegranate, unripe date and bengal fig fruits) and plant residues (leaves of river-red-gum, persimmon pomegranate and guava trees) are good sources of tannase-producing microorganisms. Many authors used similar isolation sources for isolating tannase-producing microorganisms. Batra and Saxena (2005) used tannery effluent samples for isolating tannase-producing Aspergilli and Penicillii. Murugan et al. (2007) and Manjit et al. (2008) used tannery effluent samples for isolating tannase-producing fungi. Chowdhury et al. (2004) used soil samples from a tannery region for isolating tannic acid-degrading bacteria. Franco et al. (2005) Isolated tannic acid-degrading bacteria from the bacteria colonizing the rhizosphere of plants growing in the area of discharge of tannery effluent. Enemuor and Odibo (2009) isolated tannase-producing Aspergillus tamari from soil inundated by effluent of a tannery at Oji River local Government Area of Enugu State, Nigeria. Hadi et al. (1994) isolated tannase-producing Rhizopus oryzae from soil sample of Indian Institute of Technology campus. Banerjee et al. (2001) isolated sixteen potent tannase-producing fungi from different forest soils of Midnapore District; West Bengal, India. Kachouri et al. (2005) used soil samples of Tunisian olive oil mill. All isolates are belonging to moulds. No yeast or bacterial isolates were obtained under both aerobic and anaerobic incubation (Table 5.1). These results reflect the domination of moulds as tannase-producing microorganisms in the examined samples. There are tremendous numbers of studies about tannase production by filamentous fungi (Rajakumar and Nandy, 1983; Hadi et al., 1994; Chaterjee et al., 1996; Misro et al., 1997; Kar and Bnerjee, 2000; Nuero and Reyes, 2002; Kar et al., 2002; Ramiez-coronel et al., 2003; Mukherjee and Banerjee, 2004; Batra and Saxena, 2005; Purohit et al., 2006; Battestin and Macedo, 2007; Sharma et al., 2007; Trevino-Cueto et al., 2007; Battestin et al., 2008; Sharma et al., 2008) in comparison with the few studies on isolating tannaseproducing bacteria (Kumar et al., 1999; Mondal et al., 2000; Nishitani and Osawa, 2003;
59
Table (5.1): Sources, types and number of tannase-producing isolates. Isolation source code
Isolation source Tannery effluent from El-Max tannery region, El-
Max, Alexandria, Egypt. Muddy soil sample down palm trees from Boselly
region, Beheira Governorate, Egypt. Muddy soil sample sample down river-red-gum (Eucalyptus camaldulensis Dehn) trees from the entertainment forest gardens, Alexandria, Egypt. Sandy soil sample down palm trees from Edko city, Beheira Governorate, Egypt. Sandy soil sample down Palm trees from Borg Rosetta, Beheira Governorate, Egypt. Mouldy river-red-gum (Eucalyptus camaldulensis Dehn) leaves from Abees farm, Alexandria, Egypt. Mouldy Pomegranate (Punica granatum) fruits from local market, Alexandria, Egypt. Mouldy persimmon (Diospyrus kaki) fruits from local market, Alexandria, Egypt. Tannery soil sample from El-Max tannery region,
El- Max, Alexandria, Egypt. Mouldy dates sample from Abees farm, Alexandria, Egypt. Mouldy pomegranate (Punica granatum) peels from Postharvest Lab, Faculty of Agriculture, Alexandria University, Egypt. Soil sample from plant pots from Faculty of
Agriculture, Alexandria University, Egypt. Mouldy tea from local market, Alexandria, Egypt. Mouldy bengal fig (ficus benghalensis ) fruits from
Antoniadis park, Alexandria, Egypt. Soil sample down bengal fig tree from Antoniadis park, Alexandria, Egypt. Mouldy pomegranate (Punica granatum) leaves from El-Kilo 59-Alexandri-Cairo desert road,
Numbers of tannaseproducing isolates Moulds Yeasts Bacteria
A
6
0
0
B
5
0
0
C
6
0
0
D
3
0
0
E
12
0
0
F
4
0
0
G
4
0
0
H
7
0
0
I
7
0
0
J
6
0
0
K
4
0
0
L
6
0
0
M
3
0
0
N
5
0
0
O
6
0
0
P
5
0
0
Q
5
0
0
R
5
0
0
S
8
0
0
Alexandria, Egypt. Soil sample down pomegranate (Punica granatum) trees from El-Kilo 59-Alexandri-Cairo desert
road, Alexandria, Egypt. Mouldy guava leaves from El-Kilo 59-AlexandriCairo desert road, Alexandria, Egypt. Soil sample down guava (Psidium guajava) trees from El-Kilo 59-Alexandri-Cairo desert road,
Alexandria, Egypt.
60
Belmares et al., 2004; Mahapatra et al., 2005; Noguchi et al., 2007).Aguilar and GutierrezSanchez (2001) also confirmed that the filamentous fungi are the most studied microorganisms for tannase production. The isolation of tannase-producing moulds was carried out using tannic acid agar medium (TAA). TAA was used by many investigations as selective medium for tannaseproducing isolates. TAA is a minimal agar medium containing only tannic acid as sole carbon source (Bradoo et al., 1996; Pinto et al., 2001; Batra and Saxena, 2005 and Murugan et al., 2007). Consequently, the growth and formation of hydrolytic clear zone surrounding the colony of the microorganism are sufficient indications for the tannase production capability of this microorganism.
5.2 Characteristics of the obtained fungal isolates The tannase-hydrolyzing fungal isolates were purified on TTA plates. Microscopic characteristics of the purified fungal isolates were described and data are presented in Table (5.2). Forty eight fungal isolates (A2, A4, A5, B1, B2, B3, C2, C4, D1, E4, E5, E6, E9, F2, G3, G4, H4, H5, H6, I5, I6, I7, J3, J4, J5, J6, K2, L5, L6, M2, M3, N1, N5, O2, O3, O6, P2, P3, P4, Q1, Q2, Q4, R1, R3, S2, S3, S6, S7) were belonging to Aspergillus niger, Fourteen fungal isolates (A6, C3, E7, F1, G1, H2, I1, K3, L4, N2, O1, Q5, S1, S5) were belonging to Aspergillus flavus, Five fungal isolates (A3, E3, I3, J2, L3) were belonging to Aspergillus sp., three fungal isolates (C5, E8, R5) were belonging to Fusarium spp., twenty three (A1, B4, B5, C1, D2, E2, E12, F4, G4, H1, H7, I2, K1, L1, M1, N3, N4, O5, P1, Q3, R4, S4, S8) were belonging to Penicillium spp., twelve isolates (C6, D3, E10, F3, H3, I4, J1, K4, L2, O4, P5, R2) were belonging to Trichoderma spp. while, two fungal isolates, (E1, E11) couldn’t be identified. Many investigators have isolated tannase-producing Aspergillii (Knudson 1913 a,b; Pinto et al., 2001; Batra and Saxena, 2005; Murugan et al., 2007) and Penicillii (Batra and Saxena, 2005 and Murugan et al., 2007).
5.3 Screening of tannase-producing microorganisms One hundred and thirty three fungal cultures in addition to forty eight bacterial cultures were screened in three stages (primary, secondary and final screening) to select the most promising producer of tannase. One hundred and seven mould cultures of the 133 studied mould cultures were isolated from the local environment and twenty six identified mould cultures were kindly provided by five microbiologists in Egyptian universities as shown in Table (5.3). Bacterial cultures were kindly provided by two microbiologists form Egyptian universities. These cultures included forty eight unidentified rumen’s bacterial cultures and ten identified lactic acid bacteria.
61
A2, A4, A5, B1, B2, B3, C2, C4, D1, E4, E5, E6, E9, F2, G3, G4, H4, H5, H6, I5, I6, I7, J3, J4, J5, J6, K2, L5, L6, M2, M3, N1, N5, O2, O3, O6, P2, P3, P4, Q1, Q2, Q4, R1, R3, S2, S3, S6, S7 A6, C3, E7, F1, G1, H2, I1, K3, L4, N2, O1, Q5, S1, S5
Isolate code or name
Very fine
Single-celled round arranged in unbranching chains
Lime green or olive green
Conidia
Conidia
Size
Fine, small or big
Shape
Single-celled spherical or oval in unbranching chains
*Colour
Dark black, dark brown
Type
Spores
Very long
short, moderate, or long
Length
62
Nonseptate
Nonseptate
Septation
Unbranched
Unbranched
Branching
Conidiophores
Microscopic examination
Table (5.2): Morphological characterisation of fungal isolates
Metulae covering nearly the entire vesicle
Metulae and phialides radiating from the entire vesicle surface
Other characterstics
Septate
Septate
Type
Hyaline
Hyaline
Appearance
Hyphae
Aspergillus flavus
Aspergillus niger
Scientific name of isolates
White to pink
Bluish green
Unsporula -ted
Conidia
Macro conidia (multicellular)
Conidia
Conidia
Unsporula -ted
C5, E8, R5
A1, B4, B5, C1, D2, E2, E12, F4, G4, H1, H7, I2, K1, L1, M1, N3, N4, O5, P1, Q3, R4, S4, S8
C6, D3, E10, F3, H3, I4, J1, K4, L2, O4, P5, R2
E1, E11
Unsporulated
Big
One-celled, ellipsoidal in shape and grouped in balls Unsporulated
Small or big
Big
Small
Singlecelled round arranged in unbranchin g chains
Crescent shaped individually
Singlecelled oval in unbranchin g chains
* Color of spores of colonies isolates grown on PDA plates for 96 h
Bluish green; dark green
Brownish yellow
A3, E3, I3, J2, L3
Continued
63
Didn’t present
Short
Short or long
Short
Short
Didn’t present
Septate
Septate
Nonseptate
Nonseptate
Solitary phialides
Didn’t present
Didn’t present
The metulae form brushlike conidial head
Branched monophialides
Irregularly branched and display a pyramidal arrangement
Branched
Unbranched
Unbranched
Metulae and phialides radiating from the entire vesicle surface
Septate
Septate
Septate
Septate
Septate
Hyaline
Hyaline
Hyaline
Hyaline
Hyaline
Couldn’t identified
Trichoderma spp.
Penicillium spp.
Fusarium spp.
Aspergillus spp.
Table (5.3): List of the provided cultures. Microorganism
Code Number
Moulds Alternaria alternate. Aspergillus flavus. Aspergillus flavus var. columnaris. Aspergillus niger. Curvularia geniculata. Fusarium moniliforme. Mucor hiemalis. Penicillium coryophillum. Rhizopus stolonifer. Trichoderma harizianum. Aspergillus niger. NRRL 337
ESC1 ESC2 ESC3 ESC4 ESC5 ESC6 ESC7 ESC8 ESC9 ESC10 EGA1
Aspergillus oryzae. NRRL 9362 Aspergillus phoenicis. CAIM 62068 Aspergillus terreus. CAIM 151
EGA2 EGA3 EGA4
Aspergillus flavus.
ISA 1
Aspergillus niger. Aspergillus flavus. Aspergillus niger. penicillium sp. Fusarium culmorum. Fusarium gramin. Fusarium moniliforme. Fusarium oxysporum. Fusarium solani. Trichoderma harizianum. Trichoderma viride. Bacteria Befedobacterium sp. Lactobacillus acidofilus. Lactococcus cazi. J1 Leuconostoc lactis. Lactococcus lactis. 1990 Lactococcus lactis. 19904 Leuconostoc mesintroids sub dectrinicum. Leuconostoc mesintroids sub mesintroides Lactobacillus volgaricus. Streptococcus lactis sub diacetilactis.
ISA2 WSA1 WSA2 WSA3 EAA1 EAA2 EAA3 EAA4 EAA5 EAA6 EAA7
RMS9 RMS10
48 unidentified rumen’s bacterial isolates
YGA1 to YGA48
RMS1 RMS2 RMS3 RMS4 RMS5 RMS6 RMS7
Provider
Prof. Dr. Eman Fathy Sharaf. Department of microbiology- Faculty of Science, Cairo Univerity, Egypt.
Prof. Dr. Emad El-Dein Gomaa. Food science and technology dept.Faculty of Agriculture, Alexandria University, Egypt. Mrs. Islam Othman; assistant lecturer in Phytopathology Department, Faculty of Agriculture, Alexandria University, Egypt. Dr. Waiel Sabrah, Faculty of Science, Alexandria University, Egypt.
Prof. Dr. Eid Abou-Taleb; Phytopathology Department, Faculty of Agriculture, Alexandria University, Egypt.
Dr. Rabab Medany; Faculty of Agriculture, Suez Canal University, Egypt.
RMS8
64
Prof.Dr Yousry Gohar; Faculty of Science, Alexandria University, Egypt.
5.3.1 Primary screening of tannase producers
5.3.1.1 Primary screening of fungal cultures: The first screening step of fungal cultures for tannase production was carried out by determining the sum of colony and hydrolytic clear zone diameters of each fungal isolates after point inoculation on TAA plates and incubation at 30º C for 72 and 96 h. Results in Table (5.4) showed that the sum of colony and clear zone diameters vary among fungal cultures and ranged from 0.0 (no growth) to 35 mm and 0.0 to 53 mm. after 72 and 96 h, respectively. Out of one hundred and thirty three fungal cultures, only fifteen cultures were not able to produce tannase under the used experimental conditions. Among all fungal cultures tested, E8 was the poorest producer of tannase. The mould cultures coded B2, I7> I5> Q2> P4> Q4> R3> H6> P2> E6 and J4 had the largest sum of colonies and hydrolytic clear zone diameters either after 72 or 96 h of incubation. Consequently, they chosen as the best tannase- producers and were subjected to secondary screening. Bradoo et al. (1996) reported a high correlation coefficient (r = 0.93) between clear zone diameter (included colony diameter) and the quantitative enzyme production in broth medium. Pinto et al. (2001) preferred determination of the colonies diameter instead of diameter of the clear zones due to their observation of the clear zones was very difficult. Murugan et al. (2007) also confirmed the high correlation between the colony diameter and tannase production on tannic acid agar (TAA) plates. They also mentioned that in the case of potential tannase producers though the mycelial spread was scanty but clear zones were produced along with colony growth.
5.3.1.2 Primary screening of bacterial cultures:
Fifty eight bacterial cultures (see Table 5.3, page 64), (ten lactic acid bacteria and forty eight unidentified rumen’s bacterial isolates) were screened for tannase production using plates of tannins-treated brain heart infusion agar as described by Osawa (1990). The growth of all tested bacterial cultures was restricted and no distinct clear zones were formed by any tested bacterial isolates. This result indicated that no culture of the tested bacteria was tannase producer. Therefore, the second screening step and the further work were carried out only on the selected fungal cultures from primary screening step.
65
Table (5.4): Diameters of fungal* colonies and clear zones measured after 72 and 96 h of growth on TAA medium. Fungal culture The sum of colony and hydrolytic clear zone diameters** (mm.) ± SD code 72 h Incubation time 96 h Incubation time 6±0 9±0 A1 28 ± 0 42 ± 0 A2 24 ± 0.6 39 ± 0.6 A3 30 ± 1 46 ± 0.6 A4 24 ± 0.6 36 ± 0 A5 A6 17 ± 1 24 ± 0.6 25 ± 0 38 ± 0 B1 35 ± 0 53 ± 0 B2 26 ± 0 40 ± 0 B3 7 ± 0.6 9 ± 0.6 B4 B5 7±0 10 ± 0 8±0 13 ± 0.6 C1 23 ± 0.6 35 ± 0.6 C2 17 ± 0 26 ± 0 C3 11 ± 0 16 ± 0 C4 3±0 7 ± 0.6 C5 C6 7±0 10 ± 0 24 ± 1.2 36 ± 1.2 D1 7 ± 0.6 12 ± 0.6 D2 D3 6±0 10 ± 0 17 ± 0 25 ± 0 E1 6 ± 0.6 11 ± 0 E2 16 ± 0 25 ± 0 E3 30 ± 0.6 44 ± 0.6 E4 24 ± 0.6 35 ± 1 E5 33 ± 0.6 49 ± 0 E6 8 ± 0.6 10 ± 0 E7 ng 4±0 E8 26 ± 0.6 40 ± 1.2 E9 6±0 10 ± 0 E10 3±0 6±0 E11 E12 26 ± 0.6 34 ± 0.6 18 ± 0 25 ± 0.6 F1 27 ± 0.6 40 ± 0.6 F2 19 ± 0.6 25 ± 0.6 F3 F4 18 ± 0.6 26 ± 0
66
Continued Fungal culture code G1 G2 G3 G4 H1 H2 H3 H4 H5 H6 H7 I1 I2 I3 I4 I5 I6 I7 J1 J2 J3 J4 J5 J6 K1 K2 K3 K4 L1 L2 L3 L4 L5 L6 M1 M2 M3
The sum of colony and hydrolytic clear zone diameters** (mm.) ± SD 72 h Incubation time 96 h Incubation time 16 ± 0.6 21 ± 0.6 19 ± 0.6 26 ± 0.6 30 ± 0.6 42 ± 0.6 32 ± 0.6 47 ± 1 20 ± 0 29 ± 0 19 ± 0 27 ± 0 5±0 7±0 30 ± 0 42 ± 0 27 ± 0 39 ± 0 33 ± 2.9 50 ± 2.3 9±0 11 ± 0 6±0 11 ± 0 28 ± 0.6 40 ± 0 30 ± 1.4 44 ± 1.4 32 ± 0 47 ± 0.6 34 ± 0 52 ± 0 32 ± 0.7 45 ± 1.4 35 ± 0 53 ± 0 3±0 6±0 11 ± 0 15 ± 0 29 ± 0 42 ± 0.6 33 ± 0 49 ± 0.6 31 ± 0 41 ± 0 30 ± 0 43 ± 0 14 ± 1.5 21 ± 2.3 26 ± 0 39 ± 0 13 ± 0 20 ± 0 4±0 5±0 12 ± 0 18 ± 0 19 ± 0.6 27 ± 0 10 ± 0.6 14 ± 0.6 7±0 8±0 33 ± 0.6 47 ± 1 30 ± 0.6 44 ± 1.2 5±0 10 ± 0 31 ± 0.6 45 ± 0 28 ± 0.6 40 ± 0.6
67
Continued Fungal The sum of colony and hydrolytic clear zone diameters** (mm.) ± SD culture code 72 h Incubation time 96 h Incubation time 29 ± 0 42 ± 0 N1 21 ± 0 29 ± 0 N2 11 ± 0 18 ± 0 N3 6±0 9±0 N4 26 ± 0.6 38 ± 0.6 N5 19 ± 0 28 ± 0 O1 27 ± 0.6 42 ± 0 O2 30 ± 0.6 44 ± 1 O3 10 ± 0.6 16 ± 0 O4 7 ± 0.6 10 ± 0 O5 29 ± 0.6 45 ± 1 O6 9 ± 0.6 11 ± 0 P1 34 ± 0.6 49 ± 0.6 P2 29 ± 0 44 ± 0 P3 34 ± 0 51 ± 0 P4 9 ± 1.2 12 ± 1.2 P5 31 ± 0 48 ± 0.6 Q1 36 ± 0 52 ± 0 Q2 Q3 10 ± 0 13 ± 0.6 Q4 33 ± 0.6 50 ± 0 Q5 18 ± 0 27 ± 0.6 R1 26 ± 0.6 39 ± 0 R2 12 ± 0 18 ± 0 R3 34 ± 0 50 ± 0 R4 10 ± 0 15 ± 0.6 R5 10 ± 0 16 ± 0.6 S1 16 ± 0.6 27 ± 0 S2 27 ± 0.6 40 ± 0.6 S3 29 ± 0.6 47 ±1.2 S4 8±0 10 ± 0 S5 19 ± 0 28 ± 0 S6 26 ± 0 39 ± 0.6 S7 32 ± 0 48 ± 1.5 S8 7±0 9±0 ESC1 ng ng ESC2 17 ± 0.6 20 ± 0.6 ESC3 9 ± 0.6 14 ± 0.6 ESC4 33 ± 0 45 ± 1 ng ESC5 ng ng ESC6 ng
68
Continued The sum of colony and hydrolytic clear zone diameters** (mm.) ± SD Fungal culture code 72 h Incubation time 96 h Incubation time ng ESC7 Ng ESC8 7 ± 0.6 10 ± 0 ng ESC9 ng ng ESC10 ng EGA1 27 ± 0.6 40 ± 0.6 EGA2 18 ± 0 27 ± 0.6 EGA3 26 ± 0 36 ± 0 EGA4 ng ng ISA 1 12 ± 0.6 23 ± 0.6 ISA2 22 ± 0.6 37 ± 0.6 WSA1 WSA2 WSA3
15 ± 0.6 24 ± 0 ng
27 ± 0.6 39 ± 0.6 ng
EAA1 EAA2 EAA3 EAA4 EAA5 EAA6 EAA7
ng ng ng ng ng ng ng
ng ng ng ng ng ng ng
* **
: Isolated and collected cultures : Means of triplicates. SD : The standard deviation ng : No growth. Highlighted rows: Indicate cultures with large sum of colony and clear zone diameters.
69
5.3.2 Secondary screening of selected fungal isolates The eleven selected mould cultures from the primary screening step were subjected to secondary screening for tannase production. This screen was carried out using submerged fermentation (SmF) technique in 250 ml-conical flasks containing 50 ml of the modified Czapek-Dox’s minimal medium containing 1% tannic acid as sole carbon source. Flasks were incubated at 30º C for 96 h with intermittent shaking three times a day at 200 rpm for 2 min each time. Extracellular, intracellular and total tannase activities per flask were measured in triplicates. Comparison between the means of total tannase activities (extracellular + intracellular) for each tested culture was carried out statistically. The analysis of variance (one way ANOVA) was carried out by STATESTICA 7.0 software (StatSoft, Inc., USA) and differences among the means were determined for significance at p
