CHAPTER 2

LITERATURE REVIEW

2.1 Tannin

Tannins are widespread in the plant kingdom, widely distributed in different

parts (bark, needles, heartwood, grasses, seeds and flowers) of vascular plants, and

can accumulate in large amounts in particular organs or tissues of the plant (Haslam,

1989). They are a group of complex oligomeric chains substances characterized by

the presence of polyphenolic compounds. They have molecular weight higher than

500 kDa, reaching values above 20000 kDa. One of the major characteristic of

tannins is its ability to form strong complexes with protein and to a lesser extent with

other macromolecules such as starch, cellulose and minerals (Spencer et al., 1988;

Lekha and Lonsane, 1997; Aguilar and Gutiérrez-Sánchez, 2001; Mueller-Harvey

et al., 1987).

Tannins can be divided into hydrolysable and nonhydrolysable tannins or

condensed tannins based on their structure and properties.

4

2.1.1 Hydrolysable tannins

Hydrolysable tannins are polyphenolic plant, composed of esters of gallic

acid (gallotannins) or ellagic acid (ellagitannins) with a sugar core which is usually

glucose (Figure 2.1). Hydrolysable tannins can be easily hydrolysed under mild acid

or alkaline conditions; with hot water or enzyme (López-Ríos, 1984).

Figure 2.1 Hydrolysable tannins and some of their constituents. (A) Gallotannin, (B)

Ellagitannin, (C) Ellagic acid, (D) Hexahydroxyphenic acid and (E) Gallic acid

(Mueller-Harvey, 2001).

5

2.1.2 Condensed tannins

Condensed tannins, also known as polymeric proanthocyanidins, are

composed of phenols of the flavonoid units. They are also called flavolans because

they are polymers of flavan-3-ols such as catechin or flavan-3,4-diols known as

leucocyanidins (Figure 2.2). A very interesting difference between condensed tannins

and hydrolysable tannins is the fact that condensed tannins do not contain any sugar

moieties (Porter, 1994).

Figure 2.2 Structure of (A) catechin, (B) epicatechin, and (C) 4,8 linked proantho-

cyanidins or condensed tannin (Porter, 1994).

6

An intermediate group also exists that combines both characteristics of

hydrolysable tannins and condensed tannins. This family of tannins is called the

catechin tannins. The catechin tannins are most abundant in tea leaves (Graham,

1992). Table 2.1 summarizes the different types of natural occurring tannins that can

serve as substrates for tannase.

Table 2.1 The major groups of tannins as substrates for tannase with their represent-

tative types and main sources

Hydrolysable Tannins Catechin Tannins Condensed Tannins

1. Gallotannins e.g. tannic acid (commercial name of Chinese gall tannins); yield gallic acid and glucose on hydrolysis. Sources: Tara pods (Caesal-pina spinosa), gall nuts (pathological excrescences) from Quercus infectoria (Tur-kish gall) and Rhus semialata (Chinese gall), sumac leaves (Rhus coriara).

Catechin and epi-catechin Gallates; yield catechin, epi-catechin and gallic acid on hydrolysis; have properties of hydrol-ysable and condensed tannins. Sources: Tropical shrub legu-mes, tea leaves.

Polymeric proanthocyani-dins; yield monomeric flavor-noids such as flavan-3,4-diols and flavan-3-ols on hydrolysis e.g., quebracho tannins from the wood of quebracho tree. Sources: Commonly found in fruits and seeds such as grapes, apple, olives, beans, sorghum grains, carob pods, cocoa and coffee, besides tree bark and heart wood.

2. Ellagitannins; yield ellagic acid and glucose on hydroly-sis. Sources: Wood of oak (Quer-cus spp.), chestnut (Castanea spp.) and myrobalan (Termin-alia chebula).

Common types are 1.Quebracho tannins from wood of Schinopsis spp., Loxoptery-gium spp. 2.Wattle tannins from Acacia spp. 3.Bark tannins from pine (Pinus spp.), oak (Quercus spp.) and gaboon wood (Aucoumea kleneana).

Source : Haslam and Tanner, 1970; Bhat et al., 1998

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2.2 Tannase

Tannin acyl hydrolase (E.C. 3.1.1.20) is commonly referred to as tannase.

Teighem accidentally discovered this unique enzyme in 1867 (Teighem, 1867). He

reported the formation of gallic acid when two fungal species were exposed to an

aqueous solution of tannins. The fungal species were later identified as Penicillium

glaucum and Aspergillus niger (Lekha and Lonsane, 1997).

Tannase catalyzes the hydrolysis of bonds present in the molecule of

hydrolyzable tannins and gallic acid esters to liberate gallic acid and glucose. (Lekha

and Lonsane, 1997).

2.2.1 Sources of tannase

Tannins inhibit the growth of a number of microorganisms, resist microbial

attack and are recalcitrant to biodegradation (Field and Lettinga, 1992). Condensed

tannins are more resistant to microbial attack than hydrolysable tannins and are toxic

to a variety of microorganisms. For this reason, tannins generally retard the rate of

decomposition of soil organic matter via inhibition of biodegradative enzymes of the

attacking organism (Scalbert, 1991). Biodegradation of soil is usually a very complex

process, the process usually involves degradation of organic matter by microorga-

nisms to utilize the broken down constituents as carbon, energy or nitrogen sources.

The large amounts of polyphenolic compounds on the tannin substrate structure can

form complexes with the extra and intracellular enzymes from the biodegradative

8

organisms. This complexation leads to inhibition of the biodegradative enzymes

(Scalbert, 1991), which in turn leads to a loss in the microbial growth and eventually

an increase in the bioconversion time taken for the decomposition of soil organic

matter. Despite the antimicrobial properties of tannins, many fungi, bacteria and

yeasts are quite resistant to tannins, and can grow and develop on them as a carbon

source (Deschamps, 1989; Deschamps et al., 1983).

2.2.1.1 Microbial tannase

Tannase can be obtained from plant, animal and microbial sources. From

plant sources, the enzyme is present in tannin-rich vegetables mainly in their fruits,

leaves, branches and barks of trees like konnam, mirobolano and badúl (Madha-

vakrishna et al., 1960; Pourrat, et al., 1985; Lekha and Lonsane, 1997). As for animal

sources, tannase can be extracted from bovine intestine and from the ruminal mucous

(Begovic and Duzic, 1977). Also, it has been reported that some insects produce the

enzyme during the larval state (Aguilar and Gutiérrez-Sánchez, 2001).

The most important source to obtain the enzyme is by microbial way, because

the produced enzymes are more stable than similar ones obtained from other sources

(Lekha and Lonsane, 1997). Likewise, microorganisms can produce tannase in high

quantities in a constant way. Additionally, microorganisms can undergo new tech-

niques, such as genetic manipulation, resulting in an increase in the tannase activity

titers (Aguilar and Gutiérrez-Sánchez, 2001).

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Most of all the microorganisms able to produce tannase, Aspergillus sp. were

commercially the most efficient producers of this enzyme (Table 2.2). Tannase is

produced as a membrane bound or intracellular enzyme. Not all tannase is equally

active against the different tannin substrates. Fungal tannases have a better activity in

degrading hydrolysable tannins, whereas yeast tannases degrade tannic acid better and

has a lower affinity for naturally occurring tannins (Deschamps et al., 1983). On the

other end of the spectrum, bacterial tannase can degrade and hydrolyse natural tannin

and tannic acid very efficiently (Deschamps et al., 1983; Lewis and Starkey, 1969).

Table 2.2 Microorganisms producing tannin acyl hydrolase (tannase)

Bacteria

Bacillus cereus Streptococcus bovis Streptococcus gallolyticus Lactobacillus plantarum Lactobacillus paraplantarum Lactobacillus pentosus Lactobacillus acidophilus Lactobacillus animalis Lactobacillus murinus Enterococcus faecalis Weissella paramesenteroides Leuconostoc fallax Leuconostoc mesenteroides Pediococcus acidilactici Pediococcus pentosaceus Citrobacter freundii Selenomonas ruminantium

Mondal et al.(2001) Belmares et al. (2004) Sasaki et al. (2005) Ayed and Hamdi (2002); Kostinek et al. (2007) Nishitani and Osawa (2003); Nishitani et al. (2004) Nishitani et al. (2004); Kostinek et al. (2007) Nishitani et al. (2004); Sabu et al. (2006) Nishitani et al. (2004) Nishitani et al. (2004) Goel et al. (2005) Kostinek et al. (2007) Kostinek et al. (2007) Kostinek et al. (2007) Nishitani et al. (2004) Nishitani et al. (2004) Belmares et al. (2004) Belmares et al. (2004)

10

Table 2.2 (Continued)

Yeasts

Candida sp. Pichia spp. Debaryomyces hansenii Saccharomyces cerevisiae Mycotorula japonica

Aoki et al. (1976) Deschamps et al. (1983) Deschamps et al. (1983) Zhong et al. (2004) Belmares et al. (2004)

Fungi

Aspergillus niger Aspergillus gallonyces Aspergillus awamori Aspergillus fumigatus Aspergillus versicolor Aspergillus flavus Aspergillus caespitosum Aspergillus aculeatus Aspergillus foetidus Penicillium charlessi Penicillium variable Penicillium crustosum Penicillium restrictum Penicillium glabrum Mucor sp. Paecilomyces variotii Rhizopus oryzae

Rana and Bhat (2005); Cruz-Hernández et al. (2006); Treviño-Cueto et al. (2007); Murugan et al. (2007) Belmares et al. (2004) Mahapatra et al. (2005) Batra and Saxena (2005) Batra and Saxena (2005) Batra and Saxena (2005) Batra and Saxena (2005) Banerjee et al. (2001) Banerjee et al. (2005) Batra and Saxena (2005) Batra and Saxena (2005) Batra and Saxena (2005) Batra and Saxena (2005) Van de Lagemaat and Pyle (2005) Belmares et al. (2004) Mahendran et al. (2005); Battestin and Macedo (2007) Purohit et al. (2006)

Source : Adapted from Aguilar et al., 2007

Some bacterial cultures have developed the ability to express extracellular

tannase to degrade tannins, thus releasing gallic acid and glucose. Deschamps et al.

(1983) showed that strains of Bacillus pumilus, B. polymyxia, and Klebsiella

planticola were able to produce extracellular tannase with chestnut bark as the sole

11

source of carbon. The most abundant group of bacteria able to degrade tannins is

found in the gastrointestinal track of ruminants (Deschamps et al., 1983).

Filamentous fungi also have the ability to degrade tannins as a sole source of

carbon (Lewis and Starkey, 1969; Hadi et al., 1994). Researchers revealed that

degradation of tannins increased with the addition of other metabolisable substances.

Ganga et al. (1977) found that A. niger and Penicillium spp. grew better on a medium

containing glucose and tannin (Bhat et al., 1997; 1998), which meant that the addition

of carbon and nitrogen sources favoured the production of tannase for the subsequent

cleavage of the tannin molecules to liberate a supply of carbon for growth.

Tannin degradation by yeasts has not been studied to its full potential. Aoki

et al. (1976) isolated and reported the enzymatic degradation of gallotannins by yeast

species belonging to Candida that was able to produce tannase. The tannase from this

yeast was able to hydrolyse the ester and depside linkages from tannic acid to liberate

gallic acid and glucose.

2.2.1.2 Plant tannase

Many tannin-rich plant materials have been isolated that contain tannase

activity, for example Myrobolan fruits (Terminalia chebula), divi-divi pods

(Caesalpinia coriaria) and from English oak (Quercus robur), Penduculate oak

(Quercus rubra) and from the leaves of the Karee (Rhus typhina) tree (Niehaus and

Gross, 1997; Madhavakrishna et al., 1960).

12

Cell free extracts from Quercus robur, Quercus rubra and Rhus typhina

revealed the pronounced hydrolysis of the substrate β-glucogallin (1-O-galloyl-β-D-

glucopyranose) in in vitro assays. The esterase purified from the leaves of the

Penduculate oak was shown to be an analogue to fungal tannase (Niehaus and Gross,

1997). It can be postulated that plant and microbial organisms have adapted a

mechanism to overcome the degradative resistance of tannins and in return utilize

them in their metabolism.

2.2.2 Properties of tannase

2.2.2.1 The optimum pH and pH stability

The optimum pH for tannase isolated from A. niger was shown to be between

5.0 and 6.0, with instability occurring at a pH above pH 6.0 (Iibuchi et al., 1968).

Barthomeuf et al. (1994) confirmed that the tannase from A. niger contained both

esterase and depsidase activity with the esterase and tannase activities peaking at a pH

of 5.0. The stability was also good over a wide pH range between a pH of 3.5 and 8.0.

Tannase isolated from the organism Chryphonectria parasitica had an optimum pH of

5.5 (Iibuchi et al., 1968). The plant tannase isolated from Penduculate oak was shown

to be active over a wide pH range with an optimum of approximately 5.0 (Niehaus

and Gross 1997). Good stability was maintained even if the enzyme was incubated

for 24 hours at a pH of 5.0 (Madhavakrishna and Bose, 1962).

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The tannase from Candida sp. K-1 showed an optimum activity at a pH value

of 6.0. The investigation also revealed that the enzyme was stable over a wide pH

range, from a pH of 3.5 to 7.5 (Aoki et al., 1976).

The fungal tannase from A. flavus has also been characterised extensively and

the authors showed that the enzyme could be preserved at a pH range of 5.0 and 5.5.

A rapid decrease in activity occurred outside this pH range. An interesting obser-

vation was that on surface cultures the mycelial tannase activity peaked at a pH of 3-7

but in culture media the tannase activity was active between a pH of 4-7, here

the activity increased with an increase in pH (Pourrat et al., 1982; Yamada et al.,

1968).

Iibuchi et al. (1968) purified a tannase enzyme from A. oryzae. The tannase

was shown to be stable at a pH range of 3-7.5 for 12h, but at narrower pH range of

4.5-6.0 the stability was maintained for 25 hours. The authors concluded that the

optimum pH for tannase from A. oryzae was pH of 5.5 (Iibuchi et al., 1968).

Tannase from Penicillium chrysogenum showed broad pH dependence with

optimum enzyme activity at a pH of 5.0-6.0, with the enzyme apparently stable at

16 °C in a pH range of 4.0 to 6.5 (Rajakumar and Nandy, 1983).

14

2.2.2.2 Iso–electric points of tannase

Not many pI values has been reported for tannase. The iso-electric points

reported to date are from the organism Chryphonectria parasitica with a pI value of

4.6-5.1 (Aoki et al., 1976), and for A. oryzae tannase a pI value of near to pH 4.0

(Iibuchi et al., 1968).

2.2.2.3 Optimum temperature and stability

The optimum temperature and stability values for tannase isolated from

various organisms are shown in Table 2.3. The optimum temperature for tannase

ranged between 30-50 °C, with a temperature stability ranging from as low as 0 °C to

as high as 80 °C in the case of A. oryzae.

Table 2.3 The optimum temperature and stability of tannase

Organism Optimum

temperature Temperature

stability Reference

Fungal tannase A. flavus 50-60 ºC ≤ 70 °C Yamada et al. (1968);

Pourrat et al. (1982) A. oryzae 30-40 ºC 55 ºC Beverini and Metche (1990);

Iibuchi et al. (1968) A. niger 35 ºC ≤ 50 ºC Haslam and Tanner (1970) Penicillium chrysogenum

30-40 ºC 45 ºC Rajakumar and Nandy (1983)

Chryphonectria parasitica

30 ºC 25-40 ºC Farias, et al. (1992); Iibuchi et al. (1968)

Plant tannase Penduculate oak 35 and 40 ºC ≤ 50 ºC Niehaus and Gross (1997) Yeast tannase Candida sp. K-1 50 ºC ≤ 50 ºC Aoki et al. (1976)

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2.2.2.4 Molecular mass and carbohydrate content

The molecular weight of tannase was shown to vary from 186,000 Da to

300,000 Da as shown in Table 2.4.

Table 2.4 Molecular weight and carbohydrate content of tannase

Organism Molecular

weight (Da) Carbohydrate content (%)

Reference

A. flavus

192,000

25.4%

Yamada et al. (1968); Adachi et al. (1971)

A. niger

186,000

43%

Barthomeuf et al. (1994); Parthasarathy and Bose (1976)

A. oryzae

300,000

22.7%

Hatamoto et al. (1996); Abdel-Naby et al. (1999)

Candida sp. K-1 250,000 61.9% Aoki et al. (1976) Chryphonectria parasitica

240,000 64% Aoki et al. (1976)

Penduculate oak 300,000 NA Niehaus and Gross (1997)

2.2.2.5 The effect of additives on tannase activity

The effect of metal ions and additives on tannase activity was studied recently

by Kar et al. (2003). One mM Mg+2 or Hg+ activated tannase activity. Ba+2, Ca+2,

Zn+2, Hg+2 and Ag+ inhibited tannase activity at 1.0 mM concentration and Fe+3 and

Co+2 completely inhibited tannase activity. Ag+, Ba+2, Zn+2 and Hg+2 competitively

inhibited tannase activity. Among the anions studied, 1 mM Br or S2O3-2 enhanced

tannase activity. Tween 40 and Tween 80 enhanced tannase activity whereas Tween

60 inhibited tannase activity. Sodium lauryl sulfate and Triton X-100 inhibited

16

tannase activity. Urea stimulated tannase activity at a concentration of 1.5 M. Among

the chelators chosen for the present study, 1mM EDTA or 1,10-o-phenanthrolein

inhibited tannase activity. Dimethyl sulphoxide and β-mercaptoethanol inhibited

tannase activity at 1 mM concentration whereas soybean extract inhibited tannase

activity at concentrations varying from 0.05% to 1.0% (w/v). Among the nitrogen

sources selected ammonium ferrous sulfate, ammonium sulfate, ammonium nitrate

and ammonium chloride enhanced tannase activity at 0.1% (w/v) concentration.

2.2.3 The specificity of tannase

2.2.3.1 The mode of hydrolytic action

It is known that tannase hydrolyses the ester bonds of tannic acid although

tannic acid is known to denature proteins. According to research done by Iibuchi

et al. (1972), tannase was shown to hydrolyse tannic acid (Figure 2.3. (I)) completely

to gallic acid and glucose through 2,3,4,6-tetragalloyl glucose (Figure 2.3. (III)) and

two kinds of monogalloyl glucose (Figure 2.3. (IV)). This is supported by the facts

that the same products were detected in the hydrolysate of 1,2,3,4,6-pentagalloyl

glucose, and that depsidic gallic acid of methyl-m-digallate was liberated first.

In affect this meant that the enzyme would react with any phenolic hydroxyl

group, but for a true enzyme substrate complex to form the substrate had to be an

ester compound of gallic acid (Iibuchi et al., 1972).

17

Figure 2.3 Hydrolysis pathway of tannic acid by tannase (Iibuchi et al., 1972)

Where R1 and R2 are gallate and digallate respectively.

2.2.3.2 Kinetic parameters of tannase catalytic activity

Barthomeuf et al. (1994) showed that the tannase enzyme from A. niger

contained esterase activity that catalyses the hydrolysis of the galloyl esters that are

attached to glucose moieties. Depsidase activity hydrolyses the depside linkages

between two galloyl residues (Haslam and Stangroom, 1966). Gallotannins are

exclusively poly-O-galloyl-D-glucose with varying complexity according to the plant

source. In gallotannins a certain proportion of the galloyl groups are bound in the

18

form of m-depsides. It is suggested that the depsidically linked gallolyl groups are not

randomly distributed but that they form one polygalloyl chain of variable length

linked to a carbohydrate nucleus at one specific position. Tannase was shown to

contain two separate activities containing esterase and depsidase activities with

specificity for methyl gallate (Figure 2.4, I) and m-digallic acid (Figure 2.4, II) ester

linkages (Haslam and Stangroom, 1966).

The tannase enzyme isolated from A. niger was subjected to a series of

experiments in which it was possible to vary the ratio of esterase/depsidase activities

of the enzyme; i.e. the activity against methyl gallate (Figure 2.4, I)/m-digallic acid

(Figure 2.4, II) ester linkages (Haslam and Stangroom, 1966).

The authors showed that when A. niger was grown on a depside-free media,

in this case methyl gallate, a tannase was yielded with an increase in the esterase/

depsidase ratio. This was in contrast with tannase yielded upon the growth of the

organism on gallotannin media. They reported that each of these enzymes were

capable of hydrolysing both esters and depsides of gallic acid (Figure 2.4, I and II)

and that each enzyme had a relative specificity, one for esters and the other for

depsides (Haslam and Stangroom, 1966).

19

Figure 2.4 A model showing the esterase and depsidase activities of tannase from

A. niger (Haslam and Stangroom, 1966)

2.2.4 Tannase extraction and purification

Tannase extraction strongly depends on the fermentation system used. Since

tannase is mostly extracellular when produced by SSF, it can be easily extracted with

water or a buffer. Two to three volumes of the agent extraction is well mixed with the

fermented mass and pressed to obtain the enzymatic extract. Tannase location during

its production by SmF depends on the cultivation time (Rajakumar and Nandy, 1983).

It is mainly intracellular at the beginning of the culture and it is further secreted to the

culture medium. The enzyme was purified to homogeneity from the cell-free culture

broth by preparative isoelectric focusing and by FPLC using anion-exchange and gel-

filtration chromatography. SDS-PAGE analysis as well as gel localization studies of

purified tannase indicated the presence of two enzyme forms (Ramirez-Coronel et al.,

2003).

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2.2.5 Evaluation of tannase Activity

The activity of tannase can be measured by estimating the residual tannic acid

or gallic acid formed due to enzyme action (Deschamp et al., 1983). Numerous

titrimetric method (Yamada et al., 1967), a photometric assay (Chen, 1969), a

colorimetric method (Haslam and Tanner, 1970), UV-spectrophotometric methods

(Iibuchi et al., 1968; Bajpai and Patil, 1996), and chromatographic method (Sharma

et al., 2000) have been developed to study tannase activity. However, it was report

that the colorimetric and photometric method is not specific procedure (Jean et al.,

1981). The spectrophotometric method developed by Iibuchi et al. (1968) has been

used by many researchers (Rajkumar and Nandy, 1983). This method is base on the

change in optical density of the substrate tannic acid at 310 nm. According to this

method, one unit of enzyme activity is define as the amount of enzyme that

hydrolyzes 1 μmol of ester bond in tannic acid per minute. Haslam and Tanner

(1970) was developed a spectrophotometric method which used p-nitrophenyl esters

of gallic acid as a substrates. However, this method did not find wide acceptance,

probably due to the nonavailability of substrate. Some workers have assayed tannase

activity by measuring gallic acid using such chromatographic techniques as gas

chromatography (Jean et al., 1981) or high-performance liquid chromatography

(Niehaus and Gross , 1997). These methods require more sophisticated instrument-

tation, are more time-consuming, and are not suitable for routine assays. Deschamps

et al. (1983) studies tannase activity by measuring the gallic acid produced at 260 nm

after precipitating the residual tannic acid using bovine serum albumin (BSA)

solution. Beside that, it was found that the optimum wavelength of gallic acid

21

(263 nm), tannic acid (278 nm) and BSA (280 nm) were very close to each other. In

the same time, all protein molecules which could not bind with total tannic acid could

interfere absorbance value of gallic acid. Sharma et al. (2000) reported a

spectrophotometric method for assaying microbial tannase by using methylgallate as a

substrate. This method based on the formation of chromogen between liberated gallic

acid and rhodanine. It was observed that this method was sensitive up to gallic acid

concentration of 5 nmol and has a precision of 1.7% (relative standard deviation).

The assay is complete in a short time, very convenient, and reproducible. Mondal et

al. (2001) developed the colorimetric assay method of tannase using its specific

substrate tannic acid. The method based on the changes in optical density of substrate

tannic acid after enzymatic reaction at 530 nm and the residual tannic acid was

measured by a modified BSA precipitation method. This assay is very simple,

reproducible, and very convenient, and with it tannase activity can be measured in

relation to the growth of the organism. However, precipitation of the residual tannic

acid with BSA solution is drawback of this method.

Each tannase assay method has its own limitations such as the cost of

chemical substrate, time consuming and specific instrument requirement.

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2.2.6 Applications and potential uses of tannase

2.2.6.1 Cold tea products

In producing the above-mentioned beverages the most important factor is to

produce a product with a high cold water solubility, which is a very large problem in

the manufacturing of instant tea, as tea-cream is formed when the tea is stored at or

below temperatures of 4°C (Powell et al., 1993). This haze formation is due to the

coacervation of tea flavonoids, consisting mainly of epicatechin, epicatechin gallate,

epigallocatechin and epigallocatechin gallate. Tea polyphenols also form hydrogen

bonds with caffeine, which leads to the cream formation. Consumers would prefer

clear products, thus the compounds forming the haze must be removed in order to

leave a product that is free of turbidity and chemicals used as clarifiers (Sanderson

et al., 1974). Methods used to prepare cold water-soluble teas, thus preventing the

haze formation, frequently affect the flavour quality of the beverage, tannase on the

other hand has the catalytic activity to remove gallic acid moieties from tannins

and the polyphenols from tea extract, resulting in cold water-soluble products.

Tannase catalyzes the hydrolysis of the ester linkages between galloyl group and

various compounds present in unconverted tea leaves (Sanderson et al., 1974). The

reaction is called deesterification (Figure 2.5), where R-OH group can be defined as

epicatechin and epigallocatechin.

23

Figure 2.5 Deesterification of tea polyphenols by tannase. Where ROH is epicatechin

or epigallocatechin (Sanderson et al., 1974).

This deesterification enhances the natural levels of gallic acid and epicatechin

in nonconverted green tea leaf material. This reaction favors the formation of large

amounts of epitheaflavic acid during the tea conversion process on the tea leaf

material, which has undergone preconversion tannase treatment. Epitheaflavic acid is

responsible for the bright reddish-black tea-like color and has very good cold-water

solubility. Further deesterification of green tea leaf constituents prevent the formation

of any gallated tea oxidation products by eliminating precursors of these compounds,

which are normally present in black tea infusion. Therefore, elimination of such

poorly soluble compounds is probably important for producing instant tea with good

color and solubility, and for obtaining a good yield when the green tea conversion

process is carried out after preconversion tannase treatment (Sanderson et al., 1974).

24

2.2.6.2 Beer, wine and fruits juices production

In the production of beer, tannase could be used to remove tannins, since they

are present in low quantities, especially as anthocyanidins. When the proteins of the

beer are in considerably high quantities, an undesirable turbidity is presented due to

the accomplished tannins. This problem could be resolved with the employment of

tannase (Aguilar and Gutiérrez-Sánchez, 2001).

Masschelein and Batum (1981) reported that tannase from a certain strain of

A. flavus has been shown to dramatically reduce the haze formation in beer after

storage. This implicates tannase in the hydrolysis of wort phenolics which complex

with the other chemicals in the beer mixture and results in the haze formation.

Giovanelli (1989) showed that upon treatment of the stored beer with tannase the

potential of haze formation was dramatically reduced.

In the case of wines, it is important to consider that the main tannins present

are catequins and epicatequins, which can create a complex with galactocatequins and

others galoyl-derivates. Fifty percent of the color of the wine is due to the presence of

tannins; however, if these compounds are oxidized to quinones by contact with the

air, they could form an undesirable turbidity, that causes severe problems in the

quality (Aguilar and Gutiérrez-Sánchez, 2001).

25

In the early days wine was treated chemically to remove the unfavoured

phenolics. Now tannase is being employed to hydrolyse chlorogenic acid to caffeic

acid and quinic acid, which influences the taste of the wine favourably (Chae et al.,

1983). Tannase is also used as a stabilizing and clarifying agent in some fruit juices

and in cold drinks with coffee flavor where its use applies to the removal of the

phenolic compounds present in the plant materials (Lekha and Lonsane, 1997;

Canterelli et al., 1989).

2.2.6.3 Waste treatment

The effluents contains high amounts of tannins, mainly polyphenols, which

are dangerous pollutants, for this reason the use of the tannase represents a cheap

treatment and cash for the removal of these compounds (Van de Lagemaat and Pyle,

2001).

2.2.6.4 Production of the gallic acid

One of the most important applications of tannase is the production of gallic

acid from plant byproducts rich in tannins (Pourrat et al., 1985; Kar et al., 2002).

Gallic acid has been synthesized chemically, but this chemical synthesis

has been known to be very expensive and not always very selective. Gallic acid is

one of the products liberated upon hydrolysis of tannic acid with tannase (Iibuchi

et al., 1972). It is used as a synthetic intermediate for the production of pyro-

26

gallols and gallic acid esters. Today gallic acid is mainly used for the synthesis of

trimethoprim, antibacterial drug, used in the pharmaceutical industry (Sittig, 1988). In

the food industry, it is a substrate for the chemical or enzymatic synthesis of pyrogalol

or ester galates, which are used as preservatives. The use of tannase in the production

of gallic acid also finds great application, because one can obtain the production of

the propyl-galate in an enzymatic way, which is used as an oxidant agent in fatty acids

and oils (Lekha and Lonsane, 1997; Sharma and Gupta, 2003). Now by employing

biotechnological means to synthesize gallic acid huge expenses can be saved with

better and more selective yields (Deschamps and Lebeault, 1984).

2.2.6.5 Animal feed additives

Tannins are present in large number of plant materials that are use as feed,

e.g., tree leaves, agro-industrial byproducts, agricultural wastes, and are one of the

most common antinutritional factors (Lekha and Lonsane, 1997). They characteris-

tically bind protein; the strength and nature of the binding depends on the chemical

nature of the reactive phenolic groups (Van Buren and Robinson, 1969). Formation

of complexes of tannins with nutrients, especially proteins, has both negative and

positive effects on their utilization (Reed, 1995). In small quantities, condensed

tannins are useful as they prevent bloat and protect proteins but when present in large

quantities, reduce animal feed quality. Tannins inhibit the activity of enzymes of

rumen microbes (Bae et al., 1993). For tannins present in plants can, in general,

adversely affect animal nutrition by reducing intake, protein digestibility, inhibiting

digestive enzymes or by direct systemic toxicity (Kumar and Singh, 1984). Other

27

deleterious effects of tannin include damage to the mucosal lining of the

gastrointestinal tract, alteration in excretion of certain cations, and increased excretion

of certain protein and essential amino acids (Lekha and Lonsane, 1997). This leads to

a reduction in their feed intake, adversely affects rumen fermentation and significantly

depresses digestibility of almost all the nutrients. Recent studies have focused on the

possible use of enzymes in animal feed is gaining in importance (Lekha and Lonsane,

1997). The use of tannase as an ingredient of animal feed would improve the

digestibility of the feed (Nuero and Reyes, 2002). Also, it can be used in cosmetology

to eliminate the turbidity of plant extracts and in the leather industry to homogenize

tannin preparation for high-grade leather tannins (Barthomeuf et al., 1994).

2.2.7 Microbial production of tannase

Studies on tannase production by microbial have been carried out on

submerged and solid state cultures. Depending on the strain and the culture

conditions, the enzyme can be constitutive or inducible, showing different production

patterns. Phenolic compounds such as gallic acid, pyrogallol, methyl gallate and

tannic acid induces tannase synthesis (Bajpai and Patil, 1997). However, the

induction mechanism has not been demonstrated and there is some controversy about

the role of some of the hydrolysable tannins constituents on the synthesis of tannase

(Deschamps et al., 1983). For instance, gallic acid, one of the structural constituents

of some hydrolysable tannins, such as tannic acid, has been reported as an inducer of

tannase synthesis under submerged fermentation, whilst it represses tannase synthesis

under solid state fermentation. Nevertheless, independently of the involved mecha-

28

nism, it has been well accepted that due to the complex composition of the hydroly-

sable tannins, some of their hydrolysis products induces tannase synthesis (Aguilar

et al., 2002).

Filamentous fungi of the Aspergillus genus have been widely used for tannase

production. Although tannase production by Aspergillus can occur in the absence of

tannic acid, this fungi tolerates tannic acid concentrations as high as 20% without

having a deleterious effect on both growth and enzyme production (Belmares et al.,

2004).

Addition of carbon sources such as glucose, fructose, sucrose, maltose,

arabinose to the culture medium at initial concentrations from 10 to 30 g/l improves

tannase production by Aspergillus niger (Belmares et al., 2004). Nitrogen require-

ments can be supplied by different organic and inorganic sources. Inorganic nitrogen

can be supplemented as ammonium salts (sulphate, carbonate, chloride, nitrate,

monohydrated phosphate) or nitrate salts (sodium, potassium or ammonium). Other

nutritional requirements such as potassium, magnesium, zinc, phosphate and sulphur

are supplied as salts (Belmares et al., 2004).

Tannase production has been mostly studied in submerged fermentation;

however, few studies have been also carried out under solid state fermentation

conditions. Types of strain, culture conditions, nature of the substrate and availability

of the nutrients are critical for selecting a particular production technique.

29

2.2.7.1 Production of tannase in submerged fermentation

Submerged culture fermentation (SmF) is generally used for commercial

production of microbial enzymes (Pandey et al., 1999). Presently, the commercial

production of tannase (tannin acyl hydrolase, TAH) is exclusively carried out in the

SmF system and a number of protocols based on this fermentation system have been

patented which shown in Table 2.5 (Lekha and Lonsane, 1997).

Presently, SmF is a preferred method for production of most of the

commercial enzymes like tannase, principally because sterilization and process

control are easier to handle in this system (Lekha and Lonsane, 1997). Submerged

fermentation involves the growth of the microorganism as a suspension in a liquid

medium in which various nutrients are either dissolved or suspended as particulate

solids in many commercial media (Lekha and Lonsane, 1997). Table 2.6 presents

detail as media, cultivation time, temperature and location of tannase produced by

different microorganisms, especially from filamentous fungi, mainly of Aspergillus

through processes of submerged culture.

Tannase production in submerged culture by Aspergillus sp. is improved at

high aeration rates. It is favoured at 30–33°C, initial pH of 3.5–6.5 and agitation

between 169 and 250 rpm (Adachi et al., 1968; Doi et al., 1973; Rajakumar and

Nandy, 1983; Barthomeuf et al., 1994; Lekha and Lonsane, 1994; Bajpai and Patil,

1997).

30

Aut

hors

/Ow

ner/

Yea

r

Fum

ihik

o, Y

. and

Kiy

oshi

, M.,

1975

Oka

mur

a, S

. (K

ikko

man

Cor

p.) a

nd Y

uasa

, K.

(Ina

bata

and

Co.

Ltd

.), 1

987

Oka

mur

a, S

., M

izus

awa,

K.,

Take

i, K

., Im

ai, Y

.

and

Ito, S

. (K

ikko

man

and

Inab

ata)

, 198

8

Van

dam

me,

E.,

Jero

me,

M.,

Ver

mie

ra, A

. and

Mar

ía, M

., 19

89

Sand

erso

n, G

., En

glew

ood,

N.,

Cog

gon,

P. a

nd

Ora

ngeb

urg,

N.,

1974

Cog

gon,

P.,

Gra

ham

, H. a

nd S

ande

rson

, G.,

1975

Cog

gon,

P.,

Gra

ham

, H.,

Hoe

fler,

A. a

nd

Sand

erso

n, G

. (U

nile

ver N

V),

1976

Num

ber

Jpn.

Pat.7

2,25

,786

Jpn.

Pat.6

2,27

2,97

3

Jpn.

Pat.6

3,30

4,98

1

Eur.P

at.3

39,0

11

U.S

.Pat

.3,8

12,2

66

U.K

.Pat

.1,2

80,1

35

Ger

.Pat

.2,6

10,5

33

Titl

e

Prod

uctio

n of

tann

ase

by A

sper

gillu

s

Man

ufac

turin

g of

tann

ase

with

Asp

ergi

llus

Elab

orat

ion

of ta

nnas

e by

ferm

enta

tion

Tann

ase

prod

uctio

n pr

oces

s by

acid

by

Asp

ergi

llus a

nd it

s app

licat

ion

to o

btai

n ga

llic

Con

vers

ion

of g

reen

tea

and

natu

ral t

ea le

aves

usin

g ta

nnas

e

Tea

solu

ble

in c

old

wat

er

Extra

ctio

n of

tea

in c

old

wat

er

Tab

le 2

.5 P

ublis

hed

pate

nts o

n pr

oduc

tion

and

som

e ap

plic

atio

ns o

f the

tann

ase

Typ

ea

I I I I II II II

a Ty

pe I:

Liq

uid

ferm

enta

tion

tann

ase

prod

uctio

n pa

tent

s. Ty

pe II

: Tan

nase

app

licat

ion

pate

nts.

Sour

ce :

Lek

ha a

nd L

onsa

ne, 1

997

31

Ref

.

Agu

ilar e

t al.

(200

1)

Mah

apat

ra e

t al

. (20

05)

Cru

z-H

erná

ndez

et

al. (

2006

)

Bra

doo

et a

l. (1

996)

Tan

nase

ac

tivity

(L

ocat

ion

of

enzy

me)

2.5

U/m

L (E

xtra

cellu

-la

r)

3.23

U/m

L (E

xtra

cellu

-la

r)

0.53

7 U

/mL

(Ext

race

llu-

lar)

33.0

6 U

/mL

(Ext

race

llu-

lar)

Tem

pera

ture

( °

C)

30

35

30

30

Tim

e (h

)

48

46

30

24

Initi

al

conc

entr

atio

n (%

)

5 2.5

0.6

1.5:

0.25

.25

0.43

8 0.

219

0.04

4 0.

0044

0.

006

2 2

Med

ia

Con

stitu

ent

Tann

ic a

cid

Glu

cose

(N

H4)

2SO

4 pH

5.5

Gal

lo se

ed :

myr

obal

an

frui

t in

Cza

pek

Dox

m

ediu

m

Tann

ic a

cid

(NH

4)2S

O4

KH

2PO

4 M

gSO

4.7H

2O

CaC

l 2.7H

2O

FeSO

4.7H

2O

pH 5

.5

Tann

ic a

cid

G

luco

se

in C

zape

k D

ox m

ediu

m

pH 6

.6

Tab

le 2

.6 F

erm

enta

tion

cond

ition

use

d fo

r tan

nase

pro

duct

ion

by S

mF

Mic

roor

gani

sms

A. n

iger

Aa-

20

A. a

wam

ori

naka

zaw

a

A. n

iger

GH

1

A. ja

poni

cus

32

Ref

.

Mur

ugan

et a

l. (2

007)

Had

i et a

l. (1

994)

Moh

apat

ra e

t al.

(200

6)

Tan

nase

ac

tivity

(L

ocat

ion

of

enzy

me)

16.7

7 U

/mL

(Ext

race

llu-

lar)

6.12

U/m

L (E

xtra

cellu

-la

r)

0.66

U/m

L (E

xtra

cellu

-la

r)

Tem

pera

ture

( °

C)

30

30

35

Tim

e (h

) - 96

18

Initi

al

conc

entr

atio

n (%

)

100 2 0.

05

0.1

0.05

0.

05

1 0.5

0.3

0.05

0.

05

0.05

Med

ia

Con

stitu

ent

Tann

ery

efflu

ent

pH 5

.5

Tann

ic a

cid

NaN

O3

KH

2PO

4 M

gSO

4.7H

2O

KC

l G

luco

se

pH 5

.0

Cru

de ta

nnin

ext

ract

N

H4C

l K

H2P

O4

K2H

PO4

MgS

O4.7

H2O

pH

5.0

Tab

le 2

.6 (

Con

tinue

d)

Mic

roor

gani

sms

A. n

iger

R. O

ryza

e

B. li

chen

iform

is

KB

R6

33

In SmF, tannase was constitutive when produced on simple or complex

substrates and the activity of enzyme was doubled in the presence of tannic acid as the

sole carbon (Bradoo et al., 1996). Besides that, the environmental control of SmF

was relatively simple because of the gomogeneity of the suspension of microbial cells

and of the solution of nutrients and products on the liquid phase (Raimbault, 1998).

Murugan et al. (2007) reported the production of tannase from Aspergillus

niger, Aspergillus Xavus, Trichoderma spp., Penicillium spp. and Fusarium spp. in

submerged fermentation using controlled bioreactor. Among the isolates the best

extracellular tannase producer is A. niger which produces 16.77 U/ ml. All the

isolates recorded only low tannase activity intracellularly. These results corroborates

with the reports of Lekha and Lonsane (1994) who reported that the whole

extracellular production of tannase during the entire fermentation period.

Lekha and Lonsane (1994) reported that tannase produced in SmF was

intracellular enzyme during the initial 48 h of incubation and was subsequently

excreted to an extent of 83% of the total enzyme titres at 144 h. Beside that, the

recovery of intracellular enzyme involves rather difficult and cost-intensive steps such

as centrifugation of the fermented medium for recovery of biomass, lysis of the cell

for releasing of enzyme and separation of cell debris after lysis.

34

2.2.7.2 Production of tannase in solid state fermentation

Tannase production has been mostly studied in submerged fermentation.

During the past decade, efforts have been intensified for production of tannase using

the SSF system (Kar and Banerjee, 2000; Aguilar et al., 2000; 2001; 2002; Van de

Lagemaat and Pyle, 2001; 2005; Ramirez-Coronel et al., 2003; Rana and Bhat, 2005).

SSF is defined as any fermentation process occurring in the absence or near

absence of free water, employing a natural substrate or an inert support; however,

substrate must possess enough moisture to support growth and metabolism of micro-

organism (Pandey, 2003; Pandey et al., 2001). The low moisture content means that

fermentation can only be carried out by a limited number of microorganisms, mainly

yeasts and fungi, although some bacteria have also been used (Pandey et al., 2000c).

SSF seems to have theoretical advantages over submerged substrate

fermentation (SmF). Nevertheless, SSF has several important limitations. Table 2.7

shows advantages and disadvantages of SSF compared to SmF. However, there are

few designs available in the literature for bioreactors operating in solid-state

conditions. This is principally due to several problems encountered in the control of

different parameters such as pH, temperature, aeration and oxygen transfer and

moisture. SSF lacks the sophisticated control mechanisms that are usually associated

with SmF. Control of the environment within the bioreactors is also difficult to

achieve, particularly temperature and moisture (Rodríguez Couto and Sanromán,

2006).

35

Table 2.7 Advantages and disadvantages of SSF

Advantages

Disadvantages

1. Higher productivity in a shorter time

period

2. Better oxygen circulation

3. Low-cost media

4. Less effort in downstream processing

5. Reduced energy and cost requirements

6. Simple technology

7. Scarce operational problems

8. It resembles the natural habitat for several

microrganisms

1. Difficulties on scale-up

2. Low mix effectively

3. Difficult control of process

parameters (pH, heat, moisture,

nutrient, conditions, etc.)

4. Problems with heat build-up

5. Higher impurity product,

increasing recovery product costs

6. Less knowledge of the SSF

process by west scientists

Source : Rodríguez Couto and Sanromán, 2005; 2006

For optimization of tannase production, Pinto et al. (2003) evaluated the

tannic acid/wheat bran ratio, different moisture levels, addition of supplementary

nitrogen sources, addition of supplementary phosphate, and concentration of

supplementary nitrogen and phosphate added to the medium. Their results showed

that the best medium was with 15% of tannic acid, 37.5% of initial moisture, 1.7

ammonium sulphate, and 2.0% of sodium phosphate. The presence of phosphate was

of great importance for optimization because it promoted the increase in the synthesis

level and a very expressive decrease in the maximum production time, from 72 to 24

h of fermentation. The optimized process promoted an increase of 861% in yield and

2783% in productivity.

36

2.3 Application of agricultural residues used as cultivation substrates

In recent years, there has been an increasing trend toward efficient use of

agricultural residues (Pandey et al., 2000b; 2000c; 2000d). Several processes have

been developed that use these as raw materials for the production of bulk chemicals

and value-added fine products such as ethanol, single-cell protein (SCP), mushrooms,

enzymes, organic acids, amino acid, biologically active secondary metabolites, etc.

(Pandey et al., 2000b).

2.3.1 Phenolic compounds from agro-industrial by-products

Plant food especially fruit are usually characterized by a large edible portion

and moderate amounts of waste material such as peels, seeds and stones. In contrast,

considerably higher ratios of by-products arise from tropical and subtropical fruit

processing. Due to increasing production, disposal represents a growing problem

since the plant material is usually prone to microbial spoilage, thus limiting further

exploitation. On the other hand, costs of drying, storage and shipment of by-products

are economically limiting factors. Therefore, agro-industrial waste is often utilized as

feed or as fertilizer. However, demand for feed may be varying and dependent on

agricultural yields. The problem of disposing by-products is further aggravated by

legal restrictions. Thus, efficient, inexpensive and environmentally sound utilization

of these materials is becoming more important especially since profitability and jobs

may suffer (Lowe and Buckmaster, 1995). However, the processing of plant foods

results in the production of by-products that are rich sources of bioactive compounds,

37

including phenolic compounds (Schieber et al., 2001) especially tannins. The

phenolic compounds content of several other agro-industrial by-products are as

illustrated in Table 2.8.

Thus, tannin containing agricultural residues would be considered to be a

substrate for tannase production because its composition has high tannin content, it’s

easy to find and inexpensive. Beside that, it would be benefit for the economic

tannase production and alternative method to convert agricultural residues into a

useful product. Furthermore, disposal agricultural residues would be reduced. Table

2.9 shows the tannin-rich materials employed in both culture systems (SmF and SSF)

for tannase production.

38

Ref

eren

ce

Take

oka

and

Dao

(200

2)

Wol

fe a

nd L

iu (2

003)

Shrik

hand

e (2

000)

; To

rres

and

Bob

et (2

001)

Wat

anab

e et

al.

(199

7)

Schi

eber

et a

l. (2

003)

Dey

et a

l. (2

003)

Lev

elsa

42.

52 m

g/10

0 g

fw

7.9

0 m

g/10

0 g

fw

3.0

4 m

g/10

0 g

fw

229

9 m

g C

E/10

0 g

dw

169

mg

CG

E/10

0 g

dw

5-8%

13.

4 m

g/10

0 m

g dw

6

.1 m

g/10

0 g

dw

5.0

mg/

100

g dw

4

.3 m

g/10

0 g

dw

2.5

mg/

100

g dw

6

73 m

g/kg

dw

3

18 m

g/kg

dw

8

61 m

g/kg

dw

5

62 m

g/kg

dw

13.

0 m

g ph

enol

ics/

g

dw

Phe

nolic

com

poun

ds

Chl

orog

enic

aci

d 4

-O-C

a.eo

ylqu

inic

aci

d 3

-O-C

a.eo

ylqu

inic

aci

d

Fla

vono

ids

Ant

hocy

anin

Mon

o-, o

ligo-

, and

pol

y-

mer

ic p

roan

thoc

yani

dins

Pro

toca

tech

uic

acid

3

,4-D

ihyd

roxy

benz

alde

- h

yde

Hyp

erin

R

utin

Q

uerc

etin

Fla

vono

ls

Fla

vano

ls

Dih

ydro

chal

cone

s H

ydro

xyci

nnam

ates

4-H

ydro

xybe

nzoi

c ac

id

fer

ulic

aci

d

Tab

le 2

.8 P

heno

lic c

ompo

unds

from

agr

icul

tura

l by-

prod

ucts

B

y-pr

oduc

t

Alm

ond

[Pru

nus d

ulci

s (M

ill.)

D.A

.

Web

b] h

ulls

App

le p

eels

G

rape

seed

s

Buc

kwhe

at (F

agop

yrum

esc

ulen

tum

M

őenc

h) h

ulls

D

ried

appl

e po

mac

e

Drie

d co

conu

t hus

k

39

Ref

eren

ce

Li e

t al.,

(200

6)

Som

eya

et a

l., (2

002)

Vis

ioli

and

Gal

li (2

003)

Llor

ach

et a

l. (2

002)

Geo

rge

et a

l., (2

004)

; To

or a

nd S

avag

e (2

005)

Lev

elsa

249.

4 ±

17.2

mg/

g 59

.1 ±

4.8

mg/

g 10

.9 ±

0.5

mg/

g

29.

6 m

g/ 1

00 g

dw

1.0-

1.8%

11.

3 g

phen

olic

s/10

0

mL

10.

4-40

.0 m

g

phe

nolic

s /10

0 g

fw

Phe

nolic

com

poun

ds

Phen

olic

s Fl

avon

oids

Pr

oant

hocy

anid

ins

Gal

loca

tech

in

Hyd

roxy

tyro

sol

Tyr

osol

O

leur

opei

n h

ydro

xyci

nnam

ic a

cids

Neo

chlo

roge

nic

acid

C

rypt

ochl

orog

enic

aci

d C

hlor

ogen

ic a

cid

Cyn

arin

C

affe

ic a

cid

deriv

ativ

es

Cat

echi

n G

allic

aci

d

Tab

le 2

.8 (

Con

tinue

d)

By-

prod

uct

Pom

egra

nate

pee

l B

anan

a pe

els (

Mus

a ca

vend

ish)

Oliv

e m

ill w

aste

wat

er

Arti

chok

e bl

anch

ing

wat

ers

Tom

atoe

s pee

ls a

nd se

eds

a E

xpre

ssed

on

fres

h w

eigh

t (fw

) or d

ry w

eigh

t (dw

) bas

is.

Sour

ce :

Ada

pted

from

Bal

asun

dram

et a

l., 2

006

40

Table 2.9 Tannin-rich materials used as enzyme inducer in SSF and SmF for tannase

production

Materials

Tannin content

Reference

Coffee husk 4.5-9.3% Battestin and Macedo

(2007); Sabu et al. (2006);

Pandey et al. (2000b)

Tamarind seed powder 94.5±4.9 mg /g dw Sabu et al. (2005),

Soong and Barlow (2004)

T. chebula (myrobalan)

powder

32% tannin Banerjee et al. (2005)

C. digyna (teri pod) cover

powder

45% tannin Banerjee et al. (2005)

Jamun leaves

(Syzygium cumini)

35.2 mg/g dry leaves Kumar et al. (2007)

Jawar leaves

(Sorghum vulgaris)

3.96 mg/g dry leaves Kumar et al. (2007)

Amla leaves

(Phyllanthus emblica)

45.5 mg/g dry leaves Kumar et al. (2007)

Ber leaves

(Zyzyphus mauritiana)

6.7 mg/g dry leaves Kumar et al. (2007)

Gobernadora Leaves

(Larrea tridentata Coville)

39.4% condensed tannins

and 22.8% hydrolysable

tannins

Treviño-Cueto et al.

(2007)

41

2.4 Longan

The commercial longan (Dimocarpus longan Lour.) is a highly esteemed

arilloid fruit species in Asia and belongs to the family of Sapindaceae. It grows and

crops satisfactorily in a range of tropical and subtropical countries but is exploited

commercially only in Thailand, China, Taiwan and recently, Vietnam. Other areas

which grow longan include Queensland in Australia and Florida and Hawaii in USA.

The longan resembles the lychee (Litchi chinensis) in that the tree is grown for its

fleshy, translucent, white aril which surrounds a red brown to black seed from which

it separates easily. Fruit can be eaten fresh, frozen, canned or dried. In many

countries where both the fruit species are grown, longan has not achieved the

importance of the lychee. However, in Thailand longan production is regarded to be

more economically important than lychee (Choo, 2000).

Under the family Sapindaceae, the genus Dimocarpus is reported to contain

six species of trees and shrubs (Leenhouts, 1971; 1973). Five of the species

(Dimocarpus longan, Dimocarpus dentatus, Dimocarpus gardneri, Dimocarpus

foveolatus, and Dimocarpus fumatus) are found in Asia from Sri Lanka and India to

eastern Malaysia; one (Dimocarpus australianus) exists in Queensland, Australia.

Among these species, the most commonly cultivated species is Dimocarpus longan

where the taxon Dimocarpus longan spp. longan var. longan is commonly known as

the commercial longan (Figure 2.6). The word ‘longan’ or ‘long yan’ or ‘lungngan’

comes from the Chinese and literally means ‘dragoneye’ which is an apt description

of the fruit after the skin has been removed. Other vernacular names for longan

42

include ‘lam-yai’ (Thailand), ‘leng-keng’ (Malaysia and Indonesia), ‘kyet mouk’

(Myanmar), ‘mien’ (Cambodia), ‘lam nhai’, ‘nam nhai’ (Laos), and ‘nhan’ (Vietnam)

(Choo, 2000).

Many other scientific names have been given to the longan. These include

Nephelium longana (Lam.) Cam. and Euphoria longana Lam. Beside lychee, other

related fruits under the Sapindaceae family include the ‘rambutan’ (Nephelium

lappaceum) and ‘pulasan’ (Nephelium mutabile) (Choo, 2000).

Figure 2.6 The commercial longan (Dimocarpus longan ssp. longan var. longan )

(Choo, 2000).

2.4.1 Origin

The origin of longan is disputed. Whereas some authors limit the area of

origin of longan to the mountain chain from Myanmar through Southern China, others

extend it to southwest India and Sri Lanka, including the lowlands. In China, it has

43

been suggested that the primary centre of origin of longan was Yunnan, and the

secondary centres were Guangdong, Guangxi and Hainan provinces (Guangwu et al.,

2000). This was based on studies made on the morphological characteristics of

pollens of longan cultivars and their wild species in five zones in China as well as the

analysis of botanical geography and evolution (Choo, 2000).

2.4.2 Botanical description

Longan is an evergreen tree which can grow up to 20 m and possesses a

spreading or erect habit, depending on the cultivars. The period from bloom to

harvest is 5 - 7 months, depending on cultivars and climate. In Thailand a panicle

may carry up to 80 individual fruits which vary in weight from 5 to 20g. The aril has

total soluble solid values ranging from 15 to 25 percent. It is translucent white to off

white and may constitute from 60 to 75 percent of the total fruit weight. The seed is

small, round to ovoid in shape and glossy reddish brown to black in colour and easily

detached from the aril. Only one seed is present in each fruit and in some cultivars

there are a certain percentage of small-seeded fruits (Choo, 2000).

2.4.3 Properties

The edible portion of export quality fruit ranges from 67 to 78 % of the whole

fruit. The energy value averages 458 kJ/100g. The sugar content is very high.

Composition of longan per 100g edible portion is presented in Table 2.10 (Wong and

Saichol, 1991).

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Table 2.10 Nutritional composition of longan fruit

Nutritional composition

Concentration

Calorie (Unit) Moisture (%) Protein (g) Fat (g) Carbohydrate (g) Fibre (g) Ca (mg) P (mg) Fe (mg) Vit. A (I.U.) Vit B1 (mg) Vit B2 (mg) Niacin (mg) Vit. C (mg)

109.0 72.4 1.0 0.5 25.2 0.4 2.0 6.0 0.3 28.0 0.04 0.07 0.6 8.0

Source : Wong and Saichol, 1991

2.4.4 Current world status and productivity

Currently only China, Thailand and Taiwan have exploited the commercial

growing of the longan, although Vietnam has recently started exporting longan to

other countries. Thailand is currently the biggest exporter of longan in the world.

Longan production is concentrated in the upper northern provinces with cultivation

recently extended to eastern and central regions. Major longan growing provinces

include Lamphun, Chiang Mai, Chiang Rai, Nan, Phra Yao, Lampang, Phrae and

Chanthaburi (Subhadrabandhu and Yapwattanaphun, 2000). Longan has contributed

more towards Thailand’s economy when compared to lychee. About 50 percent of

the total productions were exported (Table 2.11).

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Table 2.11 Productivity of longan in Thailand

Productivity

2004

2005

World’s productivity (Tons) 1,700,000 2,000,000 Thailand’s productivity (Tons) 597,300 712,200 Fresh longan (Tons) 100,000 143,700 Processing longan (Tons) -Dried -Canned

275,100 18,900

412,900 21,200

Export (Tons) -Dried -Canned -Fresh

71,562 11,323 116,200

94,700 21,200 134,400

Source : Department of Internal Trade (Agricultural products)

http://www.dit.go.th/agriculture/product/agri_6/agri_60650.htm (20 Jan

2008)

2.4.5 Longan seed

Thailand is the largest producer of longan in the world, contributing

approximately 35% of the world’s production (Subhadrabandhu and Yapwattanaphun,

2000). According to statistical data is presented in Table 2.11, its production reached

about 700 thousand tons. However, about 60 - 75 % of the longan pulp (total fruit

weight) constitute the portion produced as canned and dried longan; the remaining

25 - 40 % is obtained as by-products such as seed and peel. Beside that, processing of

canned longan pulp and dried longan pulp results in high proportion longan seed as

the residue (Rangkadilok et al., 2005). Although a small quantity of longan seed is

46

used as cattle feed, or in compost, it does not find any adequate application (Viet

et al., 2005). In recent years, some report have studied the composition of phenolic

compounds in longan seed. It was found that longan seed was a rich source of tannin

and polyphenols (Rangkadilok et al., 2005; Soong and Barlow, 2005).

Longan seed is byproducts of canned longan pulp and dried longan pulp

manufacture, there are several residue as longan seed are generated in large amount.

Therefore, this work to use these dried longan seed for producing valuable substances.