Microbial Pectinases and Ruminant Nutrition
Pectinases are widely distributed in higher plants and microorganisms. They are today one of the upcoming enzymes of the commercial sector. It has been estimated that microbial pectinases account for 25% of the global food enzymes sales. Microbial pectinases can be produced from bacteria including actinomycetes, yeast and fungi. Aspergillus niger is the most commonly used fungal species for industrial production of pectinolytic enzymes. Among industrial applications of pectinases are using these enzymes as an animal feed supplementation. This usage of pectinases for ruminant's feed production can reduces the feed viscosity, which increases absorption of nutrients, liberates nutrients, either by hydrolysis of non-biodegradable fibers or by liberating nutrients blocked by these fibers and reduces the amount of faeces. This study deals with types and classification of pectinolytic enzymes, their mode of action, production techniques and the methods of activity assay. Furthermore, it provides a birds eye view of the possible applications of these enzymes in ruminant nutrition.
Received: June 01, 2010;
Accepted: July 25, 2010;
Published: November 04, 2010
Pectinases or pectinolytic enzymes are today one of the upcoming enzymes of
the commercial sector. It has been reported that microbial pectinases account
for 25% of the global food enzymes sales (Jayani et al.,
2005). Primarily, these enzymes are a heterogeneous group of related enzymes
that hydrolyze the pectic substances, present mostly in plants.
Pectinolytic enzymes are widely distributed in higher plants and microorganisms
(Whitaker, 1990). They are of prime importance for plants
as they help in cell wall extension (Ward and Moo-Young,
1989) and softening of some plant tissues during maturation and storage
(Sakai, 1992; Aguilar and Huirton,
1990). They also aid in maintaining ecological balance by causing decomposition
and recycling of waste plant materials (Jayani et al.,
Microbial production of pectinases has been extensively studied during recent
years (Kashyap et al., 2001; Torres
et al., 2006). Pectinase production has been reported from bacteria
including actinomycetes (Cao et al., 1992; Bruhlmann
et al., 1994; Beg et al., 2000),
yeast (Blanco et al., 1999; Reid
and Ricard, 2000) and fungi (Elegado and Fujio, 1994;
Huang and Mahoney, 1999). However, almost all the commercial
preparations of pectinases are produced from fungal sources (Singh
et al., 1999). Aspergillus niger is the most commonly used
fungal species for industrial production of pectinolytic enzymes (Naidu
and Panda, 1998; Gummadi and Panda, 2003).
Higher cost of the production is perhaps the major constraint in commercialization
of new sources of enzymes. Though, using high yielding strains, optimal fermentation
conditions and cheap raw materials as a carbon source can reduce the cost of
enzyme production for subsequent applications in industrial processes. Among
industrial applications of pectinases are using these enzymes as an animal feed
supplementation. This usage of pectinases for ruminant's feed production can
reduces the feed viscosity, which increases absorption of nutrients, liberates
nutrients, either by hydrolysis of non-biodegradable fibers or by liberating
nutrients blocked by these fibers and reduces the amount of faeces (Hoondal
et al., 2002).
This study mainly concentrates on the types and classification of pectinolytic enzymes, their mode of action, production techniques and the methods of activity assay. It also provides a birds eye view of the possible applications of these enzymes in ruminant nutrition.
Pectic substance is the generic name used for the compounds that are acted
upon by the pectinolytic enzymes. They are high molecular weight, negatively
charged, acidic, complex glycosidic macromolecules (polysaccharides) that are
present in the plant kingdom. They are present as the major components of middle
lamella between the cells in the form of calcium pectate and magnesium pectate
(Jayani et al., 2005).
Chemically, pectic substances are essentially branched heteropolysaccharides
containing between a few hundred and about one thousand building blocks per
molecule, with a backbone consisting of galacturonic acid residues part of which
are methylesteritied. The pectin molecule (Fig. 1) is generally
agreed to consist of a chain structure of axial-axial E-(1, 4)-linked D-galacturonic
acid units, containing blocks of I.-rhamnosc rich regions, with mainly arabinose,
galactose and xylose as side chains. The carboxyl groups of the galacturonic
acid are partly esterified by methyl groups and partly or completely neutralized
by sodium, potassium or ammonium ions. Some of the hydroxyl groups on C2
and C3 may be acetylated (Pilnik and Voragen,
The American Chemical Society classified pectic substances into four main types
as reported by Alkorta et al. (1998) as follows:
||Protopectin is the water insoluble pectic substance present
in intact tissue. Protopectin on restricted hydrolysis yields pectin or
||Pectic acid is the soluble polymer of galacturonans that contains
negligible amount of methoxyl groups Normal or acid salts of pectic acid
are called pectates
||Pectinic acids is the polygalacturonan chain that contains
>0 and <75% methylated galacturonate units. Normal or acid salts of
pectinic acid are referred to as pectinates
||Pectin (Polymethyl galacturonate) is the polymeric material
in which, at least, 75% of the carboxyl groups of the galacturonate units
are esterified with methanol. It confers rigidity on cell wall when it is
bound to cellulose in the cell wall
The pectinases may be divided in three broader groups as reported by Sakai
(1992) and Palomaki and Saarilahti (1997) as follows:
||Protopectinases: Degrade the insoluble protopectin
and give rise to highly polymerized soluble pectin
||Esterases: Catalyze the de-esterification of pectin by the removal
of methoxy esters
||Depolymerases: Catalyze the hydrolytic cleavage of the α-(1-4)-glycosidic
bonds in the D-galacturonic acid moieties of the pectic substances
Depolymerases act on pectic substances by two different mechanisms, hydrolysis,
in which they catalyze the hydrolytic cleavage with the introduction of water
across the oxygen bridge and trans-elimination lysis, in which they break the
glycosidic bond by a trans-elimination reaction without any participation of
water molecule (Fig. 2) (Albersheim et
al., 1960; Codner, 2001). Depolymerases can
be subdivided into four different categories, depending on the preference of
enzyme for the substrate, the mechanism of cleavage and the splitting of the
glycosidic bonds (Rexova -Benkova and Markovic , 1976).
Polygalacturonase and polymethylgalacturonase breakdown pectate and pectin,
respectively by the mechanism of hydrolysis. However, polygalacturonate lyase
and polymethylgalacturonate lyase breakdown pectate and pectin by β elimination,
respectively. Depending upon the pattern of action, i.e., random or terminal,
these enzymes are termed as Endo or Exo enzymes, respectively. During the course
of investigations and research conducted in recent years, the above classification
of pectinolytic enzymes has slowly become obsolete. A more recent and elaborated
classification of these enzymes has been provided in Table 1
(Rexova -Benkova and Markovic , 1976; Ward
and Moo-Young, 1989; Whitaker, 1990; Kashyap
et al., 2001).
Protopectinases: The enzyme that catalyzes the solubilization of protopectin
was originally named protopectinase by Brinton et al.
(1927). Pectinosinase is also synonymous with protopectinase (PPase). Protopectinase
catalyzes the following reaction:
PPases are classified into two types, on the basis of their reaction mechanism
(Sakai and Okushima, 1982; Sakamoto
et al., 1994). A-type PPases react with the inner site, i.e., the
polygalacturonic acid region of protopectin, whereas B-type PPases react on
the outer site, i.e., on the polysaccharide chains that may connect the polygalacturonic
acid chain and cell wall constituents.
A-type PPase are found in the culture filtrates of yeast and yeast-like fungi.
They have been isolated from Kluyveromyces fragilis IFO 0288, Galactomyces
reesei L. and Trichosporon penicillatum SNO 3 and are referred to
as PPase-F, -L and -S, respectively (Whitaker,1990).
B-type PPases have been reported in Bacillus subtilis IFO 12113 (Sakai
and Ozaki, 1988), B. subtilis IFO3134 (Sakai
and Sakamoto, 1990) and Trametes sp. (Sakai et
al., 1993) and are referred to as PPase-B, -C and -T, respectively.
B-type PPases have also been found in the culture filtrate of a wide range of
Bacillus sp. (Sakai, 1992).
||Enzymatic mode of action of the most frequent depolymerases
on the pectin molecule. PMGL: Polymethylgalacturonate lyase (pectin lyase);
PMG: Polymethylgalacturonase (pectin hydrolase); PMGE: Polymethylgalacturonate
esterase (pectinestcrase); PGL: Polygalacturonate lyase (pectate lyase);
PC: Polygalacturonase (pectate hydrolase). (Serra et
Ppase activity is assayed by measuring the amount of pectic substance liberated
from protopectin by the carbazole-sulphuric acid method (Siebert
and Anto, 1946). The pectin concentration is measured as D-galacturonic
acid from its standard curve. One unit of PPase activity is defined as the enzyme
that liberates pectic substance corresponding to 1 mmol of Dgalacturonic acid
per millilitre of reaction mixture under assay conditions.
Polygalacturonases: Polygalacturonases (PGases) are the pectinolytic enzymes that catalyze the hydrolytic cleavage of the polygalacturonic acid chain with the introduction of water across the oxygen bridge. They are the most extensively studied among the family of pectinolytic enzymes. The PGases involved in the hydrolysis of pectic substances are endo-PGase (E.C. 22.214.171.124) and exo-PGase (E.C. 126.96.36.199) (Table 1). PGases have the biological, functional and technical applications in food processing and plant-fungal interactions.
Endo-PGases are widely distributed among fungi, bacteria and many yeasts (Luh
and Phaff, 1951). They are also found in higher plants and some plant parasitic
nematodes (Sakai et al., 1993). They have been
reported in many microorganisms, including Aureobasidium pullulans (Sakai
and Takaoka, 1984), Rhizoctonia solani Kuhn (Marcus
et al., 1986), Fusarium moniliforme (De
Lorenzo et al., 1987), Neurospora crassa (Polizeli
et al., 1991), Rhizopus stolonifer (Manachini
et al., 1987), Aspergillus sp. (Nagai
et al., 2000), Thermomyces lanuginosus (Kumar
and Palanivelu, 1999), Peacilomyces clavisporus (Souza
et al., 2003). Endo-PGases have also been cloned and genetically
studied in a large number of microbial species (Laing and
Pretorius, 1993; Naumov et al., 2001). In
contrast, exo-PGases occur less frequently. They have been reported in Erwinia
carotovora (Palomaki and Saarilahti, 1997),
Agrobacterium tumefaciens (Rodrigues-Palenzuela
et al., 1991), Bacteroides thetaiotamicron (Tierny
et al., 1994), E. chrysanthemi (Kobayashi,
et al., 2001), Alternaria mali (Nozaki
et al., 1997), Fusarium oxysporum (Maceira
et al., 1997), Ralstonia solanacearum (Huang
and Allen, 1997), Bacillus sp. (Kobayashi et
al., 2001). Exo-PGases can be distinguished into two types: fungal exo-PGases,
which produce monogalacturonic acid as the main end product and the bacterial
exo-PGases, which produce digalacturonic acid as the main end product (Sakai
et al., 1993). Occurrence of PGases in plants has also been reported
(Pressey and Avants, 1973; Alonso
et al., 2003).
PGase activity is determined on the basis of measuring, during the course of
the reaction: (1) the rate of increase in number of reducing groups and (2)
the decrease in viscosity of the substrate solution (Rexova
-Benkova and Markovic , 1976). The amount of reducing sugar can be readily
measured by colorimetric methods like 3, 5-dinitrosalicylate reagent method
(Miller, 1959) and the arsenomolybdate-copper reagent
method (Nelson, 1944; Collmer et
al., 1988). One unit of enzyme activity is defined as the enzyme that
releases 1 mmol/mL/min galacturonic acid under standard assay conditions.
Viscosity reduction measurements have also found widespread use in determining
the PGase activity (Roboz et al., 1952):
where, To, Tt and Tw represent the flow rate (in a capillary viscometer) in
seconds for the reaction mixture without enzyme, the test mixture and water,
respectively. The unit of enzyme activity is mostly selected as the amount of
enzyme required for attaining a certain decrease of viscosity per unit time.
However, this method has met with limited success. There is no direct correlation
between viscosity reduction and number of glycosidic bonds hydrolyzed. PGase
activity can also be determined by the cup-plate method (Dingle
et al., 1953). Cups are cut out from the solidified agar containing
the substrate and are filled with the enzyme solution. After lapse of a certain
period of time, the zones of degraded substrate are stained with iodine and
Lyases: Lyases (or transeliminases) perform non-hydrolytic breakdown
of pectates or pectinates, characterized by a trans-eliminative split of the
pectic polymer (Sakai et al., 1993). The lyases
break the glycosidic linkages at C-4 and simultaneously eliminate H from C-5,
producing a D 4:5 unsaturated products (Codner, 2001;
Albersheim et al., 1960).
Lyases can be classified into following types on the basis of the pattern of action and the substrate acted upon by them (Table 1):
(I) Endopolygalacturonate lyase (EndoPGL, E.C. 188.8.131.52); (II) exopolygalacturonate lyase (ExoPGL, E.C. 184.108.40.206); (III) endopolymethylgalacturonate lyase (EndoPMGL, E.C. 220.127.116.11); (IV) exopolymethylgalacturonate lyase (ExoPMGL).
The reactions catalyzed by lyases can be illustrated as follows:
Polygalacturonate lyases (Pectate lyases or PGLs) are produced by many bacteria
and some pathogenic fungi with endo-PGLs being more abundant than exo-PGLs.
PGLs have been isolated from bacteria and fungi associated with food spoilage
and soft rot. They have been reported in Colletotrichum lindemuthionum (Wijesundera
et al., 1984), Bacteroides thetaiotaomicron (McCarthy
et al., 1985), Erwinia carotovora (Kotoujansky,
1987), Amucala sp. (Bruhlmann, 1995), Pseudomonas
syringae pv. Glycinea (Margo et al., 1994),
Colletotrichum magna (Wattad et al., 1995),
E. chrysanthemi (Favey et al., 1992; Shevchik
et al., 1997), Bacillus sp. (Kobayashi
et al., 1999; Takao et al., 2001),
Bacillus sp. DT-7 (Kashyap et al., 2000),
C. gloeosporioides (Yakoby et al., 2000;
Drori et al., 2003)
Very few reports on the production of polymethylgalacturonate lyases (pectin
lyases or PMGLs) have been reported in literature. They have been reported to
be produced by Aspergillus japonicus (Ishii and Yokotsuka,
1975), Penicillium paxilli (Szajer and Szajer,
1982, 1985), Penicillium sp. (Diaz
Borras et al., 1987; Alana et al., 1990;
Sapunova et al., 1995), Pythium splendens
(Chen et al., 1998) Pichia pinus (Moharib
et al., 2000), Aspergillus sp. (Sunnotel
and Nigam, 2002), Thermoascus auratniacus (Martins
et al., 2002).
The most convenient method of following the activity of lyases is to measure
the increase in absorbance at 235 nm due to formation of the Δ4:5 double
bonds produced at the non-reducing ends of the unsaturated products (Albersheim,
1966; Whitaker, 1990; Liao et
al., 1999). The molar extinction coefficients for PGL and PMGL are 4.6x103
and 5.5x103/M/cm, respectively. One unit of enzyme activity is defined
as the amount of enzyme that releases 1 mmol of unsaturated product per minute
under assay conditions. Reducing group methods (Miller, 1959;
Collmer et al., 1988) are also useful in determining
the lyase activity. Viscosity reduction method (Roboz et
al., 1952), in conjunction with a reducing group method or along with
intermediate product analysis by HPLC or GC, can be used to distinguish between
endo-and exo-splitting enzymes (Albersheim, 1966).
Pectinesterase: Pectinesterase (PE, Pectin pectlylhydrolase, E.C. 18.104.22.168),
often referred to as pectinmethylesterase, pectase, pectin methoxylase, pectin
demethoxylase and pectolipase, is a carboxylic acid esterase and belongs to
the hydrolase group of enzymes (Whitaker, 1984). It
catalyzes the deesterification of methyl ester linkages of galacturonan backbone
of pectic substances to release acidic pectins and methanol (Cosgrove,
1997). The resulting pectin is then acted upon by polygalacturonases and
lyases (Sakai et al., 1993; Prade
et al., 1999). The mode of action of PE varies according to its origin
(Micheli, 2001). Fungal PEs act by a multi-chain mechanism,
removing the methyl groups at random. In contrast, plant PEs tends to act either
at the non-reducing end or next to a free carboxyl group and proceed along the
molecule by a single chain mechanism. The reaction catalyzed by PE can be represented
as follows (Forster, 1988):
PE activity is implicated in cell wall metabolism including cell growth, fruit
ripening, abscission, senescence and pathogenesis (Gaffe
et al., 1997; Dorokhov et al., 1999).
Commercially PE can be used for protecting and improving the texture and firmness
of several processed fruits and vegetables as well as in the extraction and
clarification of fruit juices (Fayyaz et al., 1993).
PE is found in plants, plant pathogenic bacteria and fungi (Hasunuma
et al., 2003). It has been reported in Rhodotorula sp. (Vaughan
et al., 1969), Phytophthora infestans (Forster,
1988), Erwinia chrysanthemi B341 (Pitkanen et
al., 1992), Saccharomyces cerevisiae (Gainvors
et al., 1994), Lachnospira pectinoschiza (Cornick
et al., 1994), Pseudomonas solanacearum (Schell
et al., 1994), Aspergillus niger (Maldonado
et al., 1994; Maldonado and de Saad,1998),
Lactobacillus lactis subsp. Cremoris (Karam and Belarbi,
1995), Penicillium frequentans (Kawano et
al., 1999), E. chrysanthemi 3604 (Laurent
et al., 2000), Penicillium occitanis (Hadj-Taieb
et al., 2002), A. japonicus (Semenova
et al., 2003) and others. There are many reports of occurrence of
PE in plants, viz., Carica papaya (Fayyaz et al.,
1993; Innocenzo and Lajalo, 2001), Lycopersicum
esculentum (Warrilow et al., 1994), Prunus
malus (Macdonald and Evans, 1996), Vitis vinifera
(Corredig et al., 2000), Citrus sp. (Arias
and Burns, 2002), Pouteria sapota (Arenas-Ocampo
et al., 2003) and Malpighia glabra L. (Assis
et al., 2004).
PE activity is most readily followed by gel diffusion assay, as described by
Downie et al. (1998). Increased binding of ruthenium
red to pectin, as the number of methyl esters attached to the pectin decreases,
is used as the basis of the assay. The unit of activity in nano-or picokatals
is calculated, based on the standard curve generated from the log-transformed
commercial enzyme activity versus stained zone diameter. The sensitivity, specificity
and simplicity of this PE assay are superior to all others. PE activity can
also be measured by using a pH stat because ionization of the carboxyl group
of the product releases a proton, which causes a change in pH (Whitaker,
MICROBIAL PECTINASES PRODUCTION
In developing nations, there is noticeable growth of food and feed processing industries, leading to an increase in the demand for pectic enzymes. Unfortunately, many of these nations depend largely on imported pectinase, which has become more expensive because of higher cost of foreign exchange. Therefore, with a view of improving the utilization of horticultural product in the depressed economy of a nation like Egypt, it is necessary to carry out research on the production of pectic enzymes locally.
Microbial pectinases production techniques: There were two fermentation
techniques we can use for pectinases production, as many other enzymes (Murad
and Foda, 1992) these techniques are Solid State Fermentation (SSF) and
submerged fermentation (SmF).
Solid state fermentation is defined as the cultivation of microorganisms on
moist solid supports, either on inert carriers or on insoluble substrates that
can be used as carbon and energy source. This process occurs in the absence
or near absence of free water in the space between substrate particles. In this
system, water is present in the solid substrate whose capacity for liquid retention
varies with the type of material (Lonsane et al.,
1985; Pandey et al., 2001). In contrast, in
submerged fermentation (SmF) the nutrients and microorganisms are both submerged
in water (Grigelmo-Migeul and Martin-Belloso, 1998).
Approximately 90% of all industrial enzymes are produced in SmF, frequently
using specifically optimized, genetically manipulated microorganisms. In this
respect SmF processing offers an insurmountable advantage over SSF. On the other
hand, almost all these enzymes could also be produced in SSF using wild-type
microorganisms (Filer, 2001; Pandey
et al., 2001). Interestingly, fungi, yeasts and bacteria that were
tested in SSF in recent decades exhibited different metabolic strategies under
conditions of solid state and submerged fermentation.
|| Biotechnological advantages of Solid State Fermentation (SSF)
against submerged technology (SmF) (Holker et al.,
The aim of SSF is to bring the cultivated fungi or bacteria into tight contact with the insoluble substrate and thus to achieve the highest substrate concentrations for fermentation. This technology results, only on a small scale, in several processing advantages of significant potential economic and ecological importance as compared with SmF (Table 2).
However, there are also several disadvantages of SSF, which have discouraged use of this technique for industrial production. The main obstructions are due mainly to the build-up of gradients of temperature, pH, moisture, substrate concentration or CO2 during cultivation, which are difficult to control under limited water availability. It has become clear (as mentioned in nearly every review cited) that the cost-factor for the production of bulk-ware enzymes in most cases favors SSF over SmF. The low estimated costs of SSF are due to the rather traditional preferential claim of SSF, viz. SSF utilises complex, heterogenous agricultural wastes as substrates and uses low-cost technology regarding sterility and regulation demands. However, attempts to reduce costs progress in SSF because of the strongly increased diversity in SSF research.
There is no consensus on the methods, the microorganisms or the substrates
used, that would allow comparison with other cultivation technologies. The broad
spectrum of substrates used represents an especially severe problem. As already
mentioned, one great advantage of SSF has always been the possibility of using
substrates that are abundant, cheap and not applicable to SmF. However, regardless
of the differences in process up-scaling, the scientific and technological impact
of research data is difficult to compare when results are obtained with different
microorganisms producing different products and using a vast variety of substrates
(Rafae et al., 2006; Abd
El-Salam et al., 1994). There are many substrates could be used as,
pineapple, mixed fruit, maosmi waste, orange bagasse, mango waste, corn tegument,
almond meal, apple pomace, molasses, banana peels, amaranth grain, eucalyptus
kraft pulp, coffee residues and some food industry wastes (Holker
et al., 2004; Murad, 1998).
To facilitate comparison of results, the use of inert substrates as solid supports
is becoming increasingly important (Gautam et al.,
2002; Ooijkaas et al., 2000). Surprisingly,
biological parameters, such as the stability of the produced enzymes at high
temperature or extreme pH, have also been reported to be better in SSF (Deschamps
and Huet, 1985; Acuna-Arguelles et al., 1995).
Capability repression or protein degradation by proteases severe problems in
SmF were often reduced or absent in SSF (Pereira et al.,
1993; Aguilar et al., 2001). In contrast,
much less research has been carried out to evaluate the metabolic differences
of microorganisms when cultivated in SSF or SmF.
The factors affecting microbial pectinases production: Environmental and nutritional factors are known to have marked effects on enzyme production by microorganisms. There are, therefore, variations in optimum conditions for pectic enzyme production. Some of the cultural factors that affect the production of pectic enzymes are presented in this study.
Initial pH of growth medium: According to Shoichi
et al. (1985) the initial pH of the medium has a great effect on
the growth of the organism, on the membrane permeability, also on the biosynthesis
and stability of the enzymes (Murad, 1998; Murad
and Salem, 2001). Optimum production of pectic enzymes from many moulds
has been reported to be within the acidic pH range (Zetelaki-Horvath,
1980; Shin et al., 1983). Zheng
and Shetty (1999), reported that, polygalacturonase produced from Lentinus
edodes has a relatively lower optimum pH (pH 5.0) in addition, Piccoli-Valle
et al. (2001) observed that a high polygalacturonase and pectin esterase
activity was showed by P. griseoroseum in more acid pH of 4.5 and 5. Also, Silva
et al. (2002) found that P. viridicatum showed maximum production
of polygalacturonase and pectinlyase at a pH of 4.5 and 5, respectively. Fawole
and Odunfa (2003) reported that the optimum pectolytic activity observed
was at pH 5. Phutela et al. (2005) concluded
that the thermophilic fungi A. fumigatus Fres expressed maximum pectinase
(1116 Ug-1) activity at pH 4.0 while polygalacturonase was active at pH 5.0
(1270 Ug-1). Also, Debing et al. (2005) found
that the pH 6.5 was the optimal pH for pectinase production from A. niger
by solid state fermentation. Reda et al. (2008)
found that the polygalacturonase productivity by Bacillus firmus-I-10104
reached its maximum at initial pH 6.0 and 6.2. Rasheedha
et al. (2010) found that P. chrysogenum exhibited maximum
polygalacturonase production at initial pH of 6.5. However, the mechanism by
which the pH acts on the production pectic enzyme is not known.
Incubation period: The time of fermentation had a profound effect on
microbial product formation (Murad and Foda, 1992; Murad,
1998; Murad and Salem, 2001). Maximum production
of pectic enzyme from different moulds varies from 1 to 6 days (Ghildyal
et al., 1981). Castilho et al. (2000)
reported that the highest polygalacturonase activities were obtained by A.
niger after 70 h of fermentation period. In addition, Fawole
and Odunfa (2003) reported that optimum production of pectinmethylesterase
was obtained after 4 days of fermentation under submerged fermentation condition.
Moreover, Sarvamangala and Dayanand (2006) observed
a gradual increase in the production of pectinase from deseeded sunflower head
by A. niger after 72 h of fermentation period in submerged and up to
96 h in solid-state conditions. Reda et al. (2008)
found that the level of polygalacturonase increased gradually with increasing
the incubation period up to a maximum of 96 h by Bacillus firmus-I-10104
under solid state fermentation conditions.
Nitrogen source: The effects of organic and inorganic nitrogen sources
on the production of pectinase were extensively studied. The observations of
Hours et al. (1988) suggested that lower levels
of (NH4)2SO4 (0.16%), or K2HPO4
(0.1%) added to the growth medium as inorganic nitrogen sources did not influence
pectinase yield. In addition Galiotou-Panayotou and Kapantai
(1993) observed that ammonium phosphate and ammonium sulphate did influence
production of pectinase positively but also recorded the inhibitory effects
of ammonium nitrate and potassium nitrate on pectinase production. Moreover,
Sarvamangala and Dayanand (2006) revealed that both
ammonium phosphate and ammonium sulphate did influence production of pectinase
positively in both submerged and solid-state conditions. In contrast, Sapunova
(1990) found that ammonium salts stimulated the pectinolytic enzyme production
in A. alliaceus BIM-83. Moreover, Sapunova et
al. (1997) has also observed that (NH4)2SO4
stimulated pectinase synthesis, as in its absence fungus displayed a slight
proteolytic activity and did not produce extracellular pectinases. In addition,
Fawole and Odunfa (2003) found that ammonium sulphate
and ammonium nitrate were good nitrogen sources for pectic enzyme production
from A. niger while glycine and tryptophan did not support enzyme production.
Also, Phutela et al. (2005) reported that (NH4)2SO4
stimulated pectinase production, as in its absence fungus displayed a
slight proteolytic activity and did not produce extracellular pectinases. Phutela
et al. (2005) found that the presence of yeast extract + (NH4)2
SO4 in growth medium supported maximal production of pectinase (925
Ug-1) followed by malt sprouts + (NH4)2SO4
(785 Ug-1), which also supported maximal polygalacturonase activity
(938 Ug-1). In addition, Rasheedha et al. (2010)
found that ammonium sulphate has enhanced the production of P. chrysogenum
On the other hand, report of Aguilar et al. (1991)
showed yeast extract (organic nitrogen source) as the best inducer of exopectinases
by Aspergillus sp. Moreover Kashyap et al.
(2003) found that, yeast extract, peptone and ammonium chloride were found
to enhance pectinase production up to 24% and addition of glycine, urea and
ammonium nitrate inhibited pectinase production. Also, Reda
et al. (2008) found that the maximum value of polygalacturonase productivity
by Bacillus firmus-I-10104 reached up to 350 U mL-1 in the
presence of peptone as a nitrogen source in the growth medium. In addition,
Vivek et al. (2010) found that organic nitrogen
sources showed higher endo, exo pectinases activities than inorganic nitrogen
sources. Also the increasing trend in the enzymes activity with the increase
in nitrogen source content was observed in the case of organic nitrogen sources
while decreasing trend observed for inorganic nitrogen sources. Soybean meal
(4%) showed the maximum Exopectinase activity of 5128 IU g-1 and
endo-pectinase activity of 793 IU g-1.
Carbon source: An adequate supply of carbon as energy source is critical
for optimum growth affecting the growth of organism and its metabolism. Aguilar
and Huitron (1987) reported that the production of pectic enzymes from many
moulds is known to be enhanced by the presence of pectic substrates in the medium.
Fawole and Odunfa (2003) found that pectin and polygalacturonic
acid promoted the production of pectic enzyme and they observed the lack of
pectolytic activity in cultures with glucose as sole carbon source reflects
the inducible nature of pectic enzyme from the strain of A. niger. However,
when different concentrations of glucose were added to the medium containing
pectin, production of pectic enzymes was inhibited at high glucose concentration
while low glucose concentrations (0.5% w/v) stimulated enzyme production Also,
the reducing sugar content of the culture filtrate increased with increase in
the amount of glucose added to the growth medium. The ability of high concentrations
of glucose in the medium to meet growth requirement of the organism probably
made the breakdown of pectin in the medium unnecessary or minimal and thus the
low pectic activities observed in cultures. Phutela et
al. (2005) stated that wheat bran supported maximum pectinase production
(589 U g-1) while pure pectin give the maximum production of polygalacturonase
(642 U g-1). Sarvamangala and Dayanand (2006)
reported that glucose (4-6%) increase the production of pectinase in submerged
condition whereas 6-8% sucrose gives better yield of pectinase in solid-state
condition. Reda et al. (2008) reported that Solanum
tuberosum (ST) peels was the best carbon source for polygalacturonase production
by Bacillus firmus-I-10104 under solid state condition.
PURIFICATION OF MICROBIAL PECTINASES
In order to characterize and study the properties of microbial pectinases the
enzymes must be purified. Important purification methods for the isolation of
different pectinases are briefly summarized in this section. Pectinases from
various sources of microorganisms have been purified to homogeneity. An exo-PG
has been separated from mycelial extracts of Aspergillus niger by eluting
from DEAE cellulose with 0.2 M sodium acetate buffer at pH 4.6. Purification
was efficient with 209-fold increase in specific activity with a recovery of
8.6% and the enzyme displayed its full activity only in the presence of Hg2_
ions (Mill, 1966a). A second PG was isolated with 205-fold
increase in specific activity with a recovery of 1%.These two PGs are differentiated
by their optimum pH and PG II was not inhibited by chelating agents and did
not require Hg2_ for activity (Mill, 1966b).
Benkova and Slezarik (1966) developed a purification
strategy for the isolation of extracellular PMG, PG and PE. The enzyme was salted
out with ammonium sulphate and precipitated with ethanol after gel filtration
through Sephadex G-25. Repeated chromatography on DEAE-cellulose column yielded
a homogeneous preparation of enzyme. Exo-PG, Endo-PG and pectinesterase have
been separated from the culture filtrate of Trichoderma reesei by Sephadex
chromatography (Markovic et al., 1985). Polygalacturonase
from Rhizopus stolonifer has been purified up to 10-fold by ethanol precipitation
followed by CM-Sepharose 6B ion exchange chromatography and gel filtration by
Sephadex G-100 (Manachini et al., 1987). PG and
PL (pectinlyase) from Aureobasidium pullulans LV10 have been separated
by CMSepharose 6B followed by column chromatography (DEAE-cellulose column)
and gel filtration on Sephadex G-100 (Manachini et al.,
1988). PG and PL (pectinlyase) have been separated into PG I and PG II and
PL I and PL II, respectively. Pectatelyase (PGL) was synthesized by Amycolata
species and the extracellular crude enzyme has been purified to homogeneity
by both cation and anion exchange columns and hydrophobic interaction chromatography
(Bruhlmann, 1995). It has been observed that purification
resulted in a 4-fold increase in specific activity with 37% recovery. Pectinases
from Clostridium aectobutylicum ID 91-36 a UV mutant, has been purified by cation
exchange chromatography on a Sepharose column by eluting with NaCl (Seethaler
and Hartmeier, 1992).
Endopectate lyase synthesized by Bacillus macerans has been purified by ammonium
sulphate precipitation followed by DEAE-Sephadex A-50 chromatography and CM-cellulofine
chromatography (Miyazaki, 1991). Similarly endopectate
lyase I_/IV have been isolated from the culture filtrate of Erwinia carotovora
by CM Sepharose CL 6B chromatography, Sephadex S-200 gel filtration and isoelectric
focusing (Tanabe et al., 1984). Kobayashi
et al. (2001) purified the first bacterial; exo-PG from Bacillus
sp. strain KSM-P443 to homogeneity. This enzyme releases exclusively mono-glacturonic
acid from polygalacturonic acid (PGA), di-, tri-, tetra-and penta-galacturonic
acids. They also determined the N-terminal sequence and concluded that no sequence
matched with other pectinases reported to-date.
An extracelluar endo-PG produced by Aspergillus awamori IFO 4033 was
purified homogeneity using cation-exchange and size-exclusion chromatographic
columns (Nagai et al., 2000). Sakamoto
et al. (1994) isolated protopectinase-N (PPN) and protopectinase-R
(PPR) from the culture filtrate of Bacillus subtilis IFO3134. These enzymes
have been purified by hydrophobic interaction chromatography on butyl-toyopearl
650 M, cation exchange chromatography on CMtoyopearl650 M and gel filtration
on sepharose 12HR. These enzymes have been found to be stable over a wide range
of pH and temperature. Endopectate lyase produced by Erwinia caratovara
FERM P-7576 has been selectively co-sedimented with an extracellularly produced
lipopolysaccharide lipid complex (Fukoka et al.,
1990). The cell free broth was precipitated and the enzyme separated by
gel chromatography with a specific activity of 710 U mg-1 of protein.
Co-sedimentation has been affected by pH and ionic strength. Denis
et al. (1990) studied the effect of shear stress on purification
of five isozymes of pectate lyase produced by Erwinia chrysanthemi 3937
in ultrafiltration equipment. Activity was not affected during 7 h of pumping
and 36% activity was lost after 25 000 passes.
New affinity matrices have been developed for the purification of pectinases,
which possess better mechanical and chemical stability than those cross-linked
one with pectic acid (Lobarzewski et al. 1985).
The culture filtrate was desalted on a Sephadex G-25 column. The supports used
were silanized controlled pore glass, silica gel silanized with 5-aminopropyl
triethoxysilane. All supports were activated with 3-(3-dimethylaminopropyl)
carbodiimide and best results were obtained with silanized controlled pore glass.
Gupta et al. (1996) developed an affinity precipitation
technique for separation of selective proteins using heterobifunctional ligands.
They used a soluble form of the ligand for affinity binding and then precipitation
was induced for separating the protein complex. Alginate was used as successful
ligand for pectinases. Wu et al. (2000) studied
the partitioning behavior of endo-PG and total protein from Kluveromyces marxianus
culture broth in polyethylene glycol (PEG) and PEG-potassium phosphate aqueous
two-phase systems (ATPS). Both enzyme and total protein partitioned into the
bottom phase in both systems. Since, the enzyme protein content in the total
protein of the fermentation broth was higher, they proposed that separating
endo-PG was concentration rather than separation.
An efficient concentration was achieved using ATPS with polymer recycling and
dialysis. Immobilized metal ion affinity polysulphone hollow-fiber membranes
with a high capacity for protein adsorption were tested for commercial pectic
enzyme fractionation (Camperi et al., 2000).
The flow through (unbound) fraction has higher activity for pectinlyase and
PE was retained on the column. Similarly, Savary (2001)
developed a rapid and simple method to separate PE from PG and other pectinases
in potato enzyme preparations using perfusion chromatography (Poros HS). PE
was eluted at low salt concentrations (-/80 mM NaCl) and PG at high salt concentrations
(-/300 mM NaCl). The development of such columns looks promising for economical
purification strategies for pectinases. A literature survey showed that extensive
work has been carried out on the purification of PG. PE and PGL and less attention
has been focused on PMG protopectinases and oligogalacturonases.
RUMINANT AND MICROBIAL PECTINASES
Ruminants have an extensive array of microbial enzymes produced in the rumen
and these enzymes play an important role in the ruminant digestive process.
Pectinolytic activity in the rumen arises from protozoal, fungal and bacterial
sources, one of the major pectinolytic bacterial species inhabiting the rumen,
Lachnospira multiparus, produces a pectin lyase and a pectin methylesterase
(Silley, 1985). Ruminal fungi and protozoa also express
one or more of these enzymes (Orpin, 1984; Chesson
and Forsberg, 1988; Bonhomme, 1990).
On the other hand, specific enzyme preparations have become valuable tools
for economically improving digestive processes in the ruminant (Gado
et al., 2009; Murad et al., 2009;
Azzaz, 2009). Use supplemental enzymes in the rumen may
be important since the digestibility of organic matter in the rumen does not
reach 100% and even small changes in digestibility can influence the efficiency
of ruminal fermentations. As a result, many strategies have been developed to
stimulate the digestion of the fibrous components in ruminant feeds. One particularly
promising technology is supplementation of fibrolytic enzymes (Cellulases, xylanases
and pectinases) to ruminant's diets (Selinger et al.,
1996). Although extensive studies were carried out to investigate the effects
of cellulases and xylanases supplementation to ruminant diets on their productivity,
study effects of pectinases supplementation to ruminant diets is lacking. Hoondal
et al. (2002) reported that use of pectinases as ruminant's feed
supplementation can reduces the feed viscosity, which increases absorption of
nutrients, liberates nutrients, either by hydrolysis of non-biodegradable fibers
or by liberating nutrients blocked by these fibers and reduces the amount of
faeces. In addition, Jacob et al. (2008) reported
that the reducing sugar level was increasing gradually for banana fiber treatment
with polygalacturonase indicating the effectiveness of the treatment and it
was evident from the photographs that the cells were separated after treatment
as a result of pectin hydrolysis. This may be reflecting the importance of use
pectinases as a fiber degrading enzymes in animal feed production.
To date, little is known about the way that exogenous fibrolytic enzymes improve
feed by rumen microorganisms. Several potential modes of action have been proposed.
These include: a) increase in microbial colonization of feed particles. Yang
et al. (1999), enhancing attachment and/or improve access to the
cell wall matrix by ruminal microorganisms and by doing so, accelerate the rate
of digestion (Nsereko et al., 2000) enhancing
the hydrolytic capacity of the rumen due to added enzyme activities and /or
synergy with rumen microbial enzymes (Morgavi et al.,
2000). Also to date, it has not been possible to predict the effectiveness
of enzyme products for ruminants based on their enzymatic activities because
the effectiveness of enzyme products is highly variable (Colombatto
et al., 2003). Part of this variability may be due to the types and
activities of enzyme, which can vary widely depending on the source organism,
growth substrate and culture conditions employed, type of diet fed to animals,
enzyme application method and even the level of animal productivity (Beauchemin
et al., 2003). Lineweaver and Jansen (1951)
study effect of low and high concentration of pectic substances (citrus pulp)
on polygalacturonase activity in steer's rumen fluid (in vitro). They
found that polygalacturonase activity in steer's rumen fluid increased by ten
folds compared with basal diet and there was no significant difference in polygalacturonase
activity between the low and high level of citrus pulp. Both levels reached
about the same maximum activity of 2,000 units mL-1) and returned
to about 200 units on the basal diet. This ten fold increase in activity of
polygalacturonase activity attributed to citrus pulp suggests that the formation
of extracellular polygalacturonase was induced by the increase in total pectic
substances in the diet.
Finally, we recommended in this study giving more attention to production of microbial pectinases and more attention to their application especially in animal feed production.
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