Comparative biochemical and structural ...

Biologia 63/1: 1—19, 2008 Section Cellular and Molecular Biology DOI: 10.2478/s11756-008-0018-y

Review

Comparative biochemical and structural characterizations of fungal polygalacturonases Suryakant K. Niture Division of Biochemical Sciences, National Chemical Laboratory, Pune, 411008, India; e-mail: [email protected]

Abstract: Endo- and exo-polygalacturonases produced by various fungi are involved in the degradation of pectic substances. They have found a wide range of applications in the food and textile industries. Several phyto-pathogenic fungi secrete polygalacturonases and they act as virulence factors during plant pathogenesis. The comparison of biochemical properties of different fungal polygalacturonases, their mechanism of actions, structural aspects and interactions with inhibitors/proteins could be used as a possible strategy for the fungal-crop disease management. This review focuses on fungal polygalacturonases, including their regulation, comparative biochemical and structural characterizations and their interactions with inhibitors. Key words: polygalacturonase; pectate lyase; polygalacturonase-inhibiting proteins; plant pathogenesis. Abbreviations: GH, glycoside hydrolase; PG, polygalacturonase; PL pectate lyase; PGIP, polygalacturonase-inhibiting protein.

Introduction The plant cell wall consists of four major groups of complex polysaccharides such as cellulose, hemicellulose, lignin and pectin. Pectin is a family of complex polysaccharides found in the middle lamella of primary cell walls of dicotyledonous and non-graminaceous monocotyledonous plants. Pectin and complex hemicellulose are responsible for the integrity and coherence of plant tissues as well as for the texture of vegetables and fruits (Pilnik & Voragen 1991). Several studies have shown that cell wall degrading enzymes, such as cellulases and pectinases, are not the primary determinants of softening of fruits although their participation in texture changes during the late stages of ripening seems evident (Fischer & Bennett 1991). Mostly pectin-degrading enzymes are used in food industries in the clarification of fruit juices, fermentation of coffee and tea, oil extractions, treatment of pectic wastewater and in textile and paper industry (Whitakar 1984; Kashyap et al. 2001). Generally, Aspergillus niger, a non-pathogenic fungus, is a good source and have considerable economic importance, since it is employed to produce extracellular pectinases used in the food industry. On the other hand, several phyto-pathogenic fungi secrete pectin-degrading enzymes during plant pathogenesis. Pectinases are the first enzymes secreted by fungal pathogens when they attack plant cell walls (Collmer & Keen 1986; Idnurm & Howlett 2001). These enzymes are essential for fungal pathogens that do not have specialized penetration structures as well as for necrotrophic pathogens during the late stages of the

invasion process (De Lorenzo et al. 1997). Among the pectinases, polygalacturonases (PGs) are the important virulence factor in Botrytis cinerea-tomato interaction (ten Have et al. 1998) and in the Claviceps purpurea-rye interaction (Oeser et al. 2002). PGs are also crucial for citrus black rot caused by Alternariacitri (Isshiki et al. 2001) and for lesion development on cotton caused by Aspergillus flavus (Shieh et al. 1997). The aim of this review is to present an overview of comparative biochemical characteristics of different fungal PGs, such as molecular mass, pI, kinetic constants, mode of action, catalytic conserved residues, structural aspects and their interactions with inhibitors. This information will be helpful to contribute to the knowledge on PGs, helpful for designing a selective inhibitor against fungal PGs and also useful from the point of view of various biotechnological applications. Pectin and pectinases Pectins are an extremely complex and structurally diverse group of polymers. The fine structures of pectins can be widely heterogeneous between plants, between tissues, and even within a single cell wall. The composition of pectin is different in different plant species and dependent on the age and maturity of the plant parts. Although researchers have reported several possible structures of pectin molecules, it is extremely difficult to predict its fine exact structure. Pectin occurs as constituent of higher-plant cell walls embedded in the cellulose fibrils. The term ‘pectin’ is a somewhat mislead-

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S. K. Niture

2 Galacturonic acid

PE

Neutral sugars side chains

Arabinose Galactose Methyl group Acetyl group Feruloyl

Homogalacturonan

Rhamnose Endo/exo PG Arabinases Endo/exo PL Galactanases

Rhamnogalacturonases

Rhamnogalacturonan

Fig. 1. Diagrammatic representation of different components of pectin molecule and associated different neutral sugars side-chains. The cleavage sites of pectin degrading enzymes are also represented (Vincken et al. 2003). PG, polygalacturonase; PL, pectate lyase; PE, pectinesterase.

ing because it is a family of polysaccharides that have common features, but are extremely diverse in their fine structures (Ridley et al. 2001). In plant cell wall, three major pectic polysaccharides are identified, all containing galacturonic acid to a greater or lesser extent. Homogalacturonan is a linear polymer consisting of 1,4linked α-D-galacturonic acid. The galacturonic acid residues of homogalacturonan can be methyl-esterified at C-6 and carry acetyl group on O-2 and O-3. Xylogalacturonan is a branched galacturonan consisting of 1,3-β-D-xylose side chains (Schols et al. 1996; Visser & Voragen 1996). The degree of xylosylation can vary between 25% (in watermelon; Citrullis vulgaris) and 75% (in apple; Malus domestica). The galacturonic acid residues of xylogalacturonan can be methyl-esterified as in homogalacturonan (Schols et al. 1995; Visser & Voragen 1996). Rhamnogalacturonan I consists of the repeating disaccharide unit [→4)-α-D-galacturonic acid(1→2)-α-L-rhamnnose-(1→] to which a variety of different glycan chains (principally arabinan and galactan) are attached to the rhamnose residues (Fig. 1). The confusingly named rhamnogalacturonan II has a backbone of homogalacturonan rather than rhamnogalacturonan, with complex side chains of arabinose, galactose, rhamnose and more rare monosaccharides such as apiose, aceric acid, deoxylyxo-heptulopyranosylaric acid and ketodeoxymanno-octulopyranosylonic acid attached to the galacturonic acid residues (Ridley et al. 2001; Willats et al. 2001). Until recently, it was accepted that rhamnogalacturonan and homogalacturonan domains constitute the ‘backbone’ of pectic polymers. However, an alternative structure has been recently proposed in which homogalacturonan is a long side chain of rhamnogalacturonan I (Vincken et al. 2003) (Fig. 1). Galacturonic acids of homogalacturonan may be both methyl-esterified and acetylated. Both the degree of methyl-esterification and degree of acetylation have

an impact on functional properties of pectin. When esterification is more than 50%, the homogalacturonan is termed as high-methoxyl pectin whereas degree of esterification less than 50% is termed as low-methoxyl pectin (Voragen et al. 1995; Rinaldo 1996). The methylesterification of homogalacturonan, in particular, has drawn the attention of many research groups, because it determines the industrial applicability of pectin. Not only the amount of methyl-esterification is important, but also the distribution of methyl groups on the homogalacturonan backbone. Blocks of more than 10 unesterified galacturonic acid residues generally yield pectin molecule, which are sensitive to Ca2+ cross-linking (Daas et al. 2001). Various physical and chemical factors such as temperature, pH, sugar and other solutes also affect the gelling properties of pectin. Generally, homogalacturonan regions of pectin are termed as ‘sooth-regions’ and rhamnogalacturonan regions as ‘hairy/branched region’. These two different regions of pectin can be cleaved by several specific enzymes, which are classified into two major classes such as esterases and depolymerases. Pectin methylesterase (EC 3.1.1.11) can remove methoxyl group from pectin or partially esterified homogalacturonan. The hairy regions of pectin can be cleaved by rhamnogalacturonases (EC 3.2.1.40), endo-xylogalacturonan hydrolases, arabinases, and galactanases. Partially esterified homogalacturonan or polygalacturonic acid can be cleaved by two distinct depolymerases such as hydrolase and lyase. The esterified homogalacturonan can be cleaved by pectin lyase (EC 4.2.2.10), whereas partially esterified homogalacturonan is the best substrate for pectate lyase (PL). Endo-PL (EC 4.2.2.2) causes random cleavage in polygalacturonic acid by a trans-elimination process, whereas exo-PL (EC 4.2.2.9) causes sequential cleavage in polygalacturonic acid by a trans-elimination process and produces unsaturated galacturonic acid. Further, homogalacturonan or polygalacturonic acid

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Fungal polygalacturonases

3

can be hydrolysed by two more enzymes such as endoPG (EC 3.2.1.15) which hydrolyses polygalacturonic acid in a random fashion and liberates saturated oligogalacturonides and galacturonic acid. Exo-PG (EC 3.2.1.67) catalyses the hydrolytic release of one saturated galacturonic acid residue from non-reducing end of homogalacturonan, whereas another exo-polyα-galacturonosidase (EC 3.2.1.82), mostly produced by bacterial species, liberates digalacturonic acid residues form the non-reducing end of galacturonan. With this brief introduction of pectin and pectinases, this review will focus on PGs specifically produced by fungal organisms. Regulation of fungal PGs production The production PGs from several fungal species has been reported by several researchers and extensively cited in the literature. Several researchers have isolated PGs from different fungal organisms, which are isolated from different sources, such as soil, decomposed plant parts, infected host tissue of plants and from mangrove/estuarine environments. Under laboratory conditions and in submerged culture, the production of PGs generally depends on medium compositions such as pectin source, nitrogen source, initial pH of the medium, temperature and agitation. Since fungi prefer to grow on semi-solid medium, semi-solid fermentation is the best system in which the moisture content, crude agricultural source and pH are the main factors. Various fungal species including Aspergillus niger, Aspergillus ustus, Aspergillus awamori (Kester & Visser 1990; Acuna-Arguelles et al. 1995; Rao et al. 1996; Blandino et al. 2002), Botrytis cinerea (Schejter & Marcus 1988; Cabanne & Doneche 2002), Colletorichum lindemuthianum (English et al. 1972; Herert et al. 2004), Corticium rolfsii (Kaji & Okada 1969), Fusarium oxysporum f. sp. melonis (Martinez et al. 1991), Fusarium moniliforme (Bonnin et al. 2001; Niture et al. 2001), Neurospora crassa (De Lourdes et al. 1991), Penicillium frequentans (Barense et al. 2001; dos Santos Cunha Chellegatti et al. 2002), Rhizopus stolonifer (Manachini et al. 1987), a non-pathogenic fungus Rhizoctonia AG-G (Machinandiarena et al. 2005) and yeast strain Saccharomyces cerevisiae (Blanco et al. 1994) produced either endo/exo or both PGs during the course of saprophytic growth or during plant pathogenesis. PGs are inducible enzymes (Nyiri 1968; Aguilar & Huitron 1987; Kawano et al. 1999), however, constitutive PG expression was also reported in Botrytis cinerea (Van der Cruyssen et al. 1994). PG production is induced by galacturonic acid and its polymer (pectin and polygalacturonic acid), galactose and structural relatives (mucic acid, tartonic acid and dulcitol) (Keen & Horton 1966; Polizeli et al. 1991; Maldonado & Strasser de Saad 1998; Malvessi & da Silveira 2004). The maximum production of PG activity (500 U/mL) was reported in Aspergillus japonicus when the fungus was grown of liquid medium containing pectin and glucose (Teixeira et al. 2000). Catabolite repression of PG production by glucose

during saprophytic growth conditions and during plant pathogenesis has been reported in numerous fungal species. PG production by Aspergillus niger is repressed by the addition of high concentration of glucose (10%) in semi-solid medium (Fontana et al. 2005). In contrast, the production of PG from Kluyveromyces marxianus does not affect when the fungus was grown in presence of 10% glucose in liquid medium (Serrat et al. 2002). Recently it has been shown that 0.2% glucose along with 1% pectin is essential for the maximum production of PG from Fusarium moniliforme NCIM1276 whereas higher glucose concentration decreased the production (Niture et al. 2006). Glucose repression is a common phenomenon in the regulation of fungal PGs; glucose acts as a catabolite repressor for PG production in numerous fungi including Fusarium oxysporum, Fisarium moniliforme, Botrytis cinerea and Saccharomyces cerevisiae (Blanco et al. 1994; Di Pietro & Roncero 1996a; Wubben et al. 2000; Niture et al. 2006). Naidu & Panda (2004) reported recently on the roles of different carbon sources and other fermentation parameters for the production of pectinases including PG. The majority of fungi secrete PG at acidic pH of the medium. For example, when induced with pectin Fusarium roseum (Perley & Page 1971) and Aspergillus nidulans (Dean & Timberlake 1989) produced maximum PG at acidic pH of the medium. During pathogenesis of tomato and cauliflower plants, a pathogenic strain of Fusarium moniliforme NCIM1276 produced more PG in tomato host tissue which having slightly acidic cell sap (pH 6.8) compared with cauliflower host tissue which having cell sap pH 7.7 (Niture et al. 2007). Moreover, Yakoby et al (2000) have suggested that, during ripening of Persea americana cv. Fuerte fruits, the pericarp pH regulates PL secretion and affects the pathogenicity of Colletotrichum gloeosporioides. These different evidences clearly suggested that extracellular medium or local host cell sap acidic pH regulates the pectinases production. Interestingly, several bacterial species like Bacillus and Streptomyces are known to produce PGs at alkaline pH (Kapoor & Kuhad 2002; Kuhad et al. 2004). Solid-state fermentation has been widely used for the production of PG by different fungal microorganisms. Solid-state fermentation is considered more suitable for fungal growth; this is justified in terms of the natural habitat conditions for fungi (mainly soil) (Pandey 2003). Majority of industrial production of PG was achieved by semi-solid fermentation because this fermentation gives higher activity of the enzyme than submerged fermentation. Studies have been conducted on comparative production of PG in systems of submerged and solid state fermentation (Solis-Pereira et al. 1993; Minjares-Carranco et al. 1997). When the fermentation medium is supplemented with different carbon sources like glucose, sucrose and galacturonic acid, PG production by Aspergillus niger CH4 is increased in solid state fermentation systems but decreased in liquid or submerged fermentation systems. The overall production of endo-PG and exo-PG is increased by 18.8


S. K. Niture

4 and 4.9 fold, respectively, in solid state fermentation compared with submerged fermentation systems (SolisPereira et al. 1993). Similarly, Maldonado & Strasser de Saad (1998) have reported that, PG production by Aspergillus niger is near about six times higher in solid state fermentation than in submerged fermentation. Agricultural by-products, such as wheat bran (Kavita & Umeshkumar 2000; Fontana et al. 2005), rice bran and sugar-cane bagasse (Solis-Pereira et al. 1993) with beet peel and citrus peel (Siessere & Said 1989) and orange peel (Niture & Pant 2004) as additional pectin sources, have been use in this process. Addition of pure sugars in the semi-solid medium also enhanced the production of enzyme, for example, the PG production by Aspergillus niger DMF27 was increased by the addition glucose (4%) and sucrose (6%) in the semi-solid medium containing deseeded sunflower head (Patil & Dayanand, 2006). In our previous study, we have also shown that not only PG but other different cell wall degrading enzymes (PL, xylanase, cellulase and amylase) production is enhanced several folds in semi-solid medium compared to submerged medium by Fusarium moniliforme (Niture & Pant 2007). Most recently Favela-Torres et al. (2006) reviewed a comparative production of PGs from different fungi in submerged culture and in solidstate fermentation. Genetic improvement for pectinase production by using protocols of mutagenesis or protoplast fusion has also been successful. Hadj-Taieb et al. (2002) have isolated a constitutive mutant Penicillium occitanis CT1, after mutagenesis with nitrous acid; the mutant produced 15 times more endo- and exo-PGs than the wild type strain of Penicillium occitanis. Protoplast fusion technique was used to obtain hybrid strains with improved characteristics for PG production. A prototrophic hybrid was obtained after protoplast fusion of auxotrophic mutants of Aspergillus sp. CH-Y-1043 (ade-) and Aspergillus flavipes ATCC 16795 (F7 lys-). This hybrid presented 1.5–fold endo-PG production, whilst exo-PG remained similar to that obtained by the wild strain of Aspergillus sp. (Solis-Pereira et al. 1997). Inter-specific protoplast fusion of Aspergillus niger and Aspergillus carbonarius was carried out to obtain hybrids with both, higher production of pectinases (6– fold more than the wild type strain) and improved growth on solid state fermentation with wheat bran as sole source of nutrients (Kavita & Umeshkumar 2000). Taken together, utilization of solid-state fermentation system and genetic improvements of fungal strains are the best strategies for PG production. Biochemical properties of fungal PGs Not only the production of PG is essential but also the knowledge about the biochemical properties of individual PG makes more sense regarding the fact that these enzymes are used in different biotechnological applications. Some biochemical properties of PGs produced by different fungi are shown in Table 1. The physicochemical properties show that the ma-

jority of fungal PGs have pH optimum between pH 3 and pH 6 (Kester & Visser 1990; De Lourdes et al. 1991; Waksman et al. 1991; Devi & AppuRao, 1996; Rao et al. 1996; Gainvors et al. 2000; Niture & Pant 2004). Interestingly, some PGs, for example, the PG of Corticium rolfsi shows maximum activity at pH 2.5 (Kaji & Okada 1969) and the one produced by Aspergillus kawachii shows maximum activity between pH 2–3 (Contreras Esquivel et al. 1999). The pH optimum of PG also depends on the size of the substrate. The optimum pH of Geotrichum candidum PG shifted from 4.5 to 3.5 on two different substrates such as polygalacturonic acid and tri-galacturonic acid, respectively (Brash & Eyal 1970). Several PGs show temperature optima between 40– 60 ◦C (Urbanek & Zelewska-Sobczak 1975; Gillespie & Coughlan 1989; Devi & AppuRao 1996). Acidic PG produced by Saccharomyces cerevisiae shows maximum activity at 25 ◦C (Gainvors et al. 2000) and several unusual temperature stable PGs have also been reported (Mill & Tuttobello 1961; Kumar & Palanivelu 1999). Thermophilic fungal strains such as Thermoascus aurantiacus, Aspergillus aculeatus and Fusarium oxysporum produce different PGs with optimal activity at temperatures between 60–65 ◦C (Foda et al. 1984; Vazquez et al. 1993; Martins et al. 2002). Fungal PGs are monomeric proteins comprising a single polypeptide covalently attached with different sugar moieties by post-translation modifications. Generally, most fungal PGs show 5–10% carbohydrate contents (Rao et al. 1996; Niture et al. 2001; Yan & Liou 2005). Endo-PG and exo-PG produced by Penicillium frequentans possess higher amounts of carbohydrate content: 20% and 81%, respectively (de Fatima Borin et al. 1996; dos Santos Cunha Chellegatti et al. 2002), whereas PG produced by Neurospora crassa contains 38% carbohydrate (Polizeli et al. 1991). Aspergillus niger PG exhibits at least two polypeptide chains with different glycosylation patterns. The component with the higher electrophoretic mobility is a high mannose or hybrid-type glycoprotein. The other component may contain O-glycosidically linked mannose, N-acetylglucosamine or glucose (Raab 1992). PG produced by a mutant strain of Saccharomyces cerevisiae shows several glycosylation sites at Asn294 and Asn306 (Hirose et al. 1999). Recently, the glycosylation sites of endo-PG I produced by Stereum purpureum was determined by tryptic digestion followed by LC-ESI-MS (liquid chromatographyelectrospray ionization mass spectrometry). The three purified compounds Ia, Ib and Ic of endo-PG I were highly glycosylated with high mannose type sugars chains containing mannose and N-acetylglucosamine residues (Man5GlcNAc2–Man10GlcNa2Man10GlcNa2) at different sites of the three components of this endoPG I [Ia: at Asn92 and Asn161 ; Ib: at Asn92 , Asn161 and Asn179 or Asn302 (3 sugar chains); and Ic: at Asn92 , Asn161 , Asn279 and Asn302 ] (Shimizu et al. 2000). Using different approach, MALDI-TOF (matrix-assisted laser desorption/ionization time-of-flight), LC-ESI-MS and



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Table 1. Some biochemical properties of fungal PGs. Fungal source

PG form

Aspergillus niger

II IV Exo I Exo II I I II III I II Exo PG I I Exo I II I & II I Exo PG2 I I I I I I I I II I I I II Exo I I I I I II

Aspergillus niger Aspergillus awamori Aspergillus carbonarius

Aspergillus japonicus Aspergillus tubingensis Aspergillus kawachii Aspergillus ustus Botrytis cinerea Fusarium oxysporum f. sp. lycopersici Fusarium oxysporum Fusarium oxysporum f. sp. lycopersici Fusarium moniliforme Kluyveromyces marxianus Neurospora crassa Penicillium frequentans Penicillium frequentans Postia placenta Phytophthora parasitica Rhizoctonia fragariae Rhizopus oryzae Sclerotinia borealis Sclerotinia sclerotiorum Sclerotinia sclerotiorum Saccharomyces cerevisiae Saccharomyces cerevisiae Thermomyces lanuginosus Thermoasus aurantiacus Trichoderma harzianum

pH optimum Mw (kDa) 3.8–4.3 3–4.6 – – – 4.0 4.1 4.3 4.0 5 4.2 2–3 5.0 5 5.2 5 – 5 5 4 4.6 3.9 4.0–4.7 3.2–3.9 – – – 4.5 4.5 – – 5 3–4.5 5.5 5.5 5.5 5.0

BEMAD (β-elimination followed by Michael addition with dithiothreitol), Woosley et al. (2006) have identified several glycosylation sites of recombinant endo-PG C produced by Aspergillus niger. The PG C possesses one N-linked high mannose (GlcNAc2 Mannose3−11 ) glycosylation site at Asn220 and seven O-linked glycosylation sites at different residues (Thr2 , Thr3 , Thr5 , Ser7 , Ser9 , Ser70 , and Ser114 ). The exact role of glycosylation is not clear, however, N-glycosylation of three isoforms produced by Fusarium oxysporum is important for secretion of these isoforms in the tomato host tissue during pathogenesis (Di Pietro & Roncero 1996b). Deglycosylation of endo-PG produced by Phytophthora parasitica and exo-PG produced by Aspergillus niger resulted in lost of catalytic activity suggesting that glycosylation sites may be present near to the active site of the enzymes (Sakamoto et al. 2002; Yan and Liou 2005). All Aspergillus niger PGs investigated have been identified as a glycoprotein (Archer and Peberdy 1997; Yang et al. 1997). Protein glycosylation has been shown to be important in maintaining protein structure and function and is also important during protein-protein interactions (Haltiwanger & Lowe 2004). Majority of purified fungal PGs have molecular mass in the range from 25 kDa to 82 kDa (Kester &

61 38 82 56 41 61 42 47 38 65 78 60 36 65 52 37 35 74 36 41.7 37 74 20 34 39.2 36 36 31 40 42 41.5 60 42 65 59 30 31.0

pI

Reference

– – – – 6.1 – – – 5.6 3.3 3.7–4.7 3.55 8.2 8.0 7.8 – 8.3 4.5 8.1 – – 4.2 5.6 3.3 5.2 6.76 7.08

Singh & AppuRao (2002)

7.5 4.8 4.8 – – – – – 4.5

Sakamota et al (2002) Nagai et al. (2000) Devi & AppuRao (1996)

Semenova et al. (2003) Kester et al. (1996) Contreras Esquivel & Voget (2004) Rao et al. (1996) Cabanne & Doneche (2002) Strand et al. (1976) Garcia-Maceira et al. (2001) Di Pietro & Roncero (1996A) Niture et al. (2001) Serrat et al. (2002) Polizeli et al. (1991) dos-Santos et al. (2002) De Fatima Borin et al. (1996) Clausen and Green (1996) Yan & Liou (2005) Cervone et al. (1977) Saito et al. (2004) Takasawa et al. (1997) Martel et al. (1998) Riou et al. (1992) Gainvors et al. (2000) Blanco et al. (1994) Kumar & Palanivelu (1999) Martins et al. (2007) Mohamed et al. (2006)

Visser 1990; Polizeli et al. 1991; Barense et al. 2001; Sakamoto et al. 2002). Penicillium frequentans PG had molecular weight 20 kDa as determined by gel filtration (De Fatima Borin et al. 1996), whereas the PG of Neurospora sitophila had a molecular mass of only 13 kDa (Fogarty & Kelly 1983). Researchers have used different methods, such as gel filtration, SDS-PAGE and HPLC for determining the molecular mass of fungal PGs. Due to the differential glycosylation, several fungal PGs show variation in their molecular mass, for example the endo-PG produced by Saccharomyces cerevisiae with a molecular mass of 39 kDa by gel filtration, however, the same protein shows molecular weight 65 kDa when determined by HPLC (Blanco et al. 1994). PG produced by Aspergillus kawachii shows molecular mass 60 kDa by SDS-PAGE but 40 kDa by gel filtration on Sephacryl S-100 column (Contreras Esquivel & Voget 2004). Therefore, to compare molecular mass of different PGs, it is essential to determine the molecular weight of the protein using the same method. Most of the molecular weights of different PGs determined by SDS-PAGE are listed in Table 1. Most fungal organisms produce multiple forms of the enzyme (Kester & Visser 1990; Devi & AppuRao 1996; Mikhailova et al. 2000) suggesting either that sev-


S. K. Niture

6 eral genes are involved, or that post-translational modification results in different physical properties of the enzymes (Caprari et al. 1993). Most PGs reported so far have acidic, neutral or alkaline pI (Polizeli et al. 1991; Clausen & Green 1996; Takasawa et al. 1997; Zhang et al. 1999; Nagai et al. 2000; Cabanne & Doneche 2002). Aspergillus niger and Sclerotinia sclerotiorum produced several forms of PG and all of them show acidic pI between 3.2 and 4.9 (Kester & Visser 1990; Waksman et al. 1991). Some PGs are basic molecules, for example, the PG of Aspergillus ustus has a pI of 8.1 (Rao et al. 1996), the one of Sclerotinia borealis has a pI of 7.5 (Takasawa et al. 1997), and the exo-PG of Botrytis cinerea shows a pI of 8.0 (Cabanne & Doneche 2002). Multiple forms produced by the same organism may also exhibit either acidic or basic pI, or the same molecular mass PGs exhibit also different pIs (Waksman et al. 1991; Cervone et al. 1977). Purified fungal PGs shows high variation in their specific activity, for example, PG produced by a strain of Botrytis cinerea E-200 Pers showing 2049 µmol/min/mg of specific activity on sodium polypectate or polygalacturonic acid (Urbanek & ZalewskaSobczak 1975). Two isozymes of PG produced by another strain of Botrytis cinerea show specific activity 1.25 µmol/min/mg of protein (Cabanne & Doneche 2002). Exo-PGs of Fusarium oxysporum and Aspergillur niger have 31.49 and 13.5 µmol/min/mg of specific activity, respectively (Vasquez et al. 1993; Sakamoto et al. 2002). Generally, exo-PGs show lower specific activities compared with their endo-counterparts on homogalacturonan. The specific activity of fungal PG is also dependent on the nature and affinity towards the substrate. Niture et al. (2001) have reported that a single PG produced by Fusarium moniliforme NCIM1276 shows very low Km value (Km 0.12 mg/mL) towards polygalacturonic acid and specific activity 178 U/mg. Interestingly, PG from another strain of Fusarium moniliforme also shows high affinity (Km 0.16 mg/mL) towards homogalacturonan (Bonnin et al. 2001), compared with PG1 from Fusarium solani – Km 1.34 mg/mL (Zhang et al. 1999), PG-E from Aspergillus niger – Km 2.5 mg/mL (Parenicova et al. 1998), and PG from Neurospora crassa – Km 5.0 mg/ml (Polizeli et al. 1991). Most fungal PGs have Km values between 0.19–20 mg/mL (Wang & Keen 1970; Manachini et al. 1987; Devi & AppuRao 1996; Rao et al. 1996). The hyphal fungus Aspergillus tubingensis produced an exo-PG which shows Km 3.2 mM for polygalacturonic acid (degree of polymerization 56) whereas, on trigalacturonate (degrees of polymerization 3), the Km value was decreased to 0.94 mM (Kester et al. 1996). However, endo-PG produced by Fusarium moniliforme shows Km 0.16 mg/mL for homogalacturonan; the same enzyme shows very low affinity towards hexa- or hepta-galacturonides (Km 0.023 nmol/mL and Km 0.016 nmol/mL, respectively) compared with trigalacturonide (Km 0.071 nmol/mL) suggesting that the enzyme have low affinity towards tri-galacturonide

(Bonnin et al. 2001). The same enzyme shows Vmax 2080 nkat/mg on homogalacturonan, which is 4–folds higher than the value of the endo-PG of Neurospora crassa (Vmax 500 nkat/mg) (Zhang et al. 1999). In the case of PG of Aspergillus niger N400, it has Vmax value in the range of µkat/mg (Benen et al. 1999; Parenicova et al. 2000). PG produced by Fusarium moniliforme NCIM 1276 also shows Vmax in the range of µmol (Vmax 111.11 µmol/min/mg) (Niture et al. 2001). Km values of different PGs decrease with increasing chain length of galacturonide (3–7) (Heinrichova et al. 1981; Kester et al. 1996; Bonnin et al. 2001). The Vmax value of exo-PG increases with chain length up to four, then reaches a plateau value, whereas, endo-PG shows increasing Vmax value with increasing chain length of galacturonides (3– 7) (Kester et al. 1996; Bonnin et al. 2001). Several fungal PGs have different kcat values, for example PG of Fusarium moniliforme show different kcat values 70/s and 104/s on polygalacturonic acid (Bonnin et al. 2001; Niture et al. 2001). Aspergillus japonicus PG1 and PG2 also show high variation in their turnover number (PG1 – 90/s, PG2 – 29/s) on polygalacturonide (Semenova et al. 2003). Interestingly, Kluyveromyces marxianus PG shows very low kcat value (6/s) on apple pectin having degree of esterification 75%, whereas the same enzyme shows the kcat value 1.83/s on polygalacturonic acid (Serrat et al. 2002). PG of Fusarium moniliforme shows increasing kcat values on increasing chain length of galacturonides similar to the Vmax values (Bonnin et al. 2001). In conclusion, from a commercial point of view, the best enzymes for biotechnological applications will be those with lower Km and high Vmax /specific activity. Mode of action of fungal PGs The mode of action of endo-PGs may be either a singlechain multiple attack mechanism in which end products appear rapidly or by a multi-chain attack where the monomers, dimers, and trimers accumulate only after further hydrolyses of higher oligogalacturonates. It has been shown that Colletotrichum lindemuthianum PG produced only di or trigalacturonic acids (English et al. 1972) whereas Saccharomyces fragilis enzyme hydrolyzed pectate through a series of higher oligogalacturonides subsequently further hydrolysed to mono and digalacturonic acids (Phaff 1966). Activity of PG on oligogalacturonates also decreases with decreasing chain length while some typical endo-PGs fail to hydrolyse digalacturonic acid and trigalacturonic acid (Rexova-Benkova & Markovic 1976; Bonnin et al. 2001). Recently in another study, Bonnin et al. (2002) have demonstrated that endo-PG produced by Fusarium moniliforme hydrolyses the linkages between two galacturonic acid residues by a multi-chain attack mechanism, at least at the early stage of the reaction. Prolonged incubation of enzyme with pectin having increasing degree of methylation resulted in the decreased percentage of hydrolysis of substrate. Random distribution of methyl groups in the pectin molecule affect the



7

hydrolysis, however, block-wise distribution of methyl groups is more favourable for hydrolysis of substrate by the enzyme. The detailed analysis of the structure of the released oligomers shows that the enzyme was able to accommodate methylated galacturonic acid in its active site and subsequently enzyme decreased its affinity (lowKm ) towards the substrate. Fungal endoPGs also show considerable differences in action patterns on oligogalacturonates. These differences depend on the nature of active site of the enzyme but more specifically on the size of the substrate binding site and position of catalytic group (Rexova-Benkova 1973). Benen et al. (1999) studied the kinetics of hydrolysis of oligogalacturonides by PG I, II and C produced by a recombinant strain of Aspergillus niger. All three different enzymes cleaved polygalacturonate by random cleavage pattern suggesting ‘endo’ type mode of action. The authors further confirmed these results by analysis of the mode of action of theses enzymes using oligogalacturonates. Processivity is observed when the degree of polymerization of the substrate exceeded 5 or 6 for endo-PG I and endo-PG C, respectively. All three enzymes cleaved these oligogalacturonides form reducing end with different rates. The bond-cleavage frequencies obtained for the hydrolysis of the oligogalacturonates were used to assess the subsite maps. The maps indicate that the minimum number of subsites is seven for all three enzymes. Additionally, the differences in substrate specificity of these three enzymes were also examined by hydrolysis of unsaturated oligogalacturonates (chain length 4–8) produced by the action of PL. Endo-PG I showed a strong preference for unsaturated oligogalacturonates generating the δ-4,5unsaturated dimmer, whereas the endo-PG II generated the δ-4,5-unsaturated trimer indicating the differences in substrate specificity. With regard to endo-PG C and E, both produced δ-4,5-unsaturated dimer and trimer in different ratios. Using pectins of various degrees of esterification, it has been shown that endo-PG II is the most sensitive to the presence of methyl esters, whereas the remaining PGs II, C and E preferred the non-esterified pectate. Using similar approach, Bonnin et al. (2001) have analyzed the mode of action of endoPG produced by Fusarium moniliforme. The enzyme was demonstrated to have a multi-chain attack mode of action. By studying the hydrolysis of oligomers, the authors have demonstrated that endo-PG have five subsites ranging form −3 to +2. The enzyme was unable to cleave the digalacturonide whereas it cleaved preferentially other oligogalacturonides (chain length 3–7) from reducing end rather than the non-reducing end. An asymmetry of binding site is observed in the most of the endo-PG already described in the literature. In fungal PGs the binding sites often include an odd numbers, for examples seven subsites are found in PG of Saccharomyces fragilis (McClendon 1979), seven subsites in Aspergillus niger PG I, II and C (Benen et al. 1999) and five in endo-PG of Fusarium moniliforme (Bonnin et al. 2001). Recently, Pages et al. (2000) have analyzed the subsite mapping of endo-PG II pro-

duced by Aspergillus niger by site-directed mutagenesis. The authors have studied the roles of key amino acid residues which are located on the different subsites of the enzyme by mutating each of residues which are located in the potential substrate binding cleft. The kinetic parameters and bond cleavage frequencies of wild and mutated proteins were examined using polygalacturonic acid and oligogalacturonides with defined chain length (n = 3–6) as a substrate. The mutation of enzyme at Asp183 →Asn183 and Met150 →Gln150 , both located at subsite -2, decreased the catalytic rate probably mediated via the sugar residue bound at subsite −1. The residues Asn186 and Asp282 are located at subsite −4 and +2, respectively. Mutation at Asn186 →Glu186 (which binds to the non-reducing end of substrate) and mutation at Asp282 →Lys282 resulted in decreasing the affinity of enzyme towards substrate by 3.5-fold and 2-fold, respectively, compared with wild type of enzyme. Tyr291 residue (which is highly conserved in different PGs) is located at subsite +1. Mutation of Tyr291 →Leu/Phe291 drastically decreased the specific activity and Vmax of the mutated enzymes, and increased Km by ∼7-fold compared with the wild type suggesting its major role in substrate binding. Other mutations at residues Glu252 →Ala252 and Gln288 →Glu288 , which are located at subsite +2, show only slight effects on catalysis and mode of action. Interestingly, mutation of Glu252 →Ala252 increased the affinity of enzyme towards partially methylesterified substrate, whereas Asn186 →Glu186 mutant protein decreased the affinity. Tyr326 is located at the imaginary subsite +3 and mutation of Tyr326 →Leu326 decreased the stability of the enzyme. In summary in most endo-PGs, the catalytic residues are situated on −1 or +1 subsites of the enzymes, and generally aspartate residues are involved the catalytic process whereas tyrosine and basic amino acids like lysine and arginine are important in the substrate recognition. Amino acid similarities of fungal PGs Based on sequence similarities the glycoside hydrolases (GHs) degrading pectin have been classified into the family GH28 (Henrissat 1991). This family consists of three types of PGs such as endo-PG (EC 3.2.1.15), exoPG (EC 3.2.1.67) and exo-poly-α-galacuronosidase (EC 3.2.1.82) along with rhamnogalacturonase and endoxylogalacturonan hydrolase. All the family GH28 members act with an inverting mechanism (Henrissat & Davis 1997) including endo- and exo-PGs (Biely et al. 1996). Markovic & Janecek (2001) have analyzed the amino acid sequence alignment of 115 PGs (including bacterial, fungal and plant enzymes) and confirmed that there are four strictly conserved sequence segments 178 NTD, 201 DD, 222 GHG and 256 RIK (according to the Aspergillus niger PG II sequence numbering). In the 178 NTD conserved segment, Asn178 and Thr179 are strictly conserved residues (the asparagine is substituted in one PG from Penicillium griseoroseum by a histidine, whereas threonine is 93% conserved; Ta-


S. K. Niture

8 Table 2. Conserved amino acid sequence alignment of some fungal PGs. Aspni2.pg AA #

30 32

45

74

80

85

114–120

128–131

177–183

200–203

Aspni2.pg Botfu6.pg Fusmo.pg Pengr2.pg Sclsc2.pg

SCTF ACTA XCSV SCTF SCTF

KCS PCV SCK SCS SCA

IFE TFA TFK IFK VFE

TFQ TFG TFA TLG TFG

EWA DSS DND EWT EWY

WWDGKG YWDGLG YWDGKG WWDGKG YWDGKG

KFFY HFIA HFIV KFFY XFFY

HNTDAFD HNTDGFD HNTDGFD HHTDAFD HNTDAFD

QDDC QDDC QDDC QDDC QDDC

216–224

228

255–259

272

283

291

329 334

337

353 362

GGTCIGGHG GLYCSGGHG MMYCSGGHG GGTCSCGHG GGTCSGGHG

SIGS SIGS SIGS SIGS SIGS

VRIKT VRIKS CRIKS VRIKT VRIKT

TYS TYS TYQ TPQ TYK

YGV YGV YGV QGI YCV

YED YLN YNL YLN YEN

WTW IKF FTF WTW WTW

CK CN CN CK CA

Aspni2.pg AA # Aspni2.pg Botfu6.pg Fusmo.pg Pengr2.pg Sclsc2.pg

LCG LCG LCG ECG LCA

SCS SCS SCS SCS KCS

SC GC TC TC KC

The amino acid sequence alignments of five endo-PGs from different fungi presented according to Aspni2.pg numbering (Markovic & Janecek 2001). The conserved amino acid residues are shown in bold letters, whereas the individual conserved sequence segments are bolded and underlined. Aspni2.pg – Aspergillus niger II PG (Swiss-Prot # P26214); Botfu6.pg – Botryotinia fuckeliana PG6 (Swiss-Prot # Q9Y7W2); Fusmo.pg – Fusarium moniliforme PG (Swiss-Prot # Q07181); Pengr2.pg – Penicillium griseoroseum PG2 (Swiss-Prot # Q9UR16); Sclsc2.pg – Sclerotinia sclerotiorum PG2 (Swiss-Prot # Q11134).

ble 2), the Asp180 being conserved invariantly. The second segment 201 DD is also conserved in all PGs with specific amino acid residues neighbouring at both sides of this dipeptide. The third segment 222 GHG is also highly conserved: 95% conserved Gly222 followed by two totally conserved His223 and Gly224 . The first Gly222 is replaced in one of the two fungal PGs produced by Colletotrichum lindemuthianum. The forth conserved segment 256 RIK contains a highly conserved (87%) Ile257 in addition to the strictly conserved Lys258 and Arg256 (replaced by a histidine in insect PG). The Ile257 is also not present in some fungal PGs and is replaced by a valine. In addition to this, Tyr291 residue was found as an invariantly conserved residue in all PGs (Markovic & Janecek 2001), which was also previously reported (Stratilová et al. 1996). Based on sequence alignments of forty-three fungal PGs, Markovic & Janecek (2001) have also suggested that PGs sequences are 8.9% identical and 17.4% similar with the average pair-wise similarity of about 60% ranging from lower than 20% to higher than 90%. In the amino acid sequence alignment several aromatic residues can be found as characteristic of these fungal PGs: Phe32 , Phe74 , Phe80 , Trp85 , Trp114 , Trp115 , Phe128 , Phe129 , Phe182 , Phe214 , Tyr272 , Tyr283 , Tyr326 , Trp337 and Trp339 [according to Aspergillus niger PG II sequence numbering] (Aspni2.pg in Table 2). Not all of them are conserved strictly except of Trp115 and Phe182 , but in most cases, there are conservative (aromatic→aromatic) substitutions. The interest in these residues is due to the fact that they (or at least some of them) may be involved in binding of substrate (Rao et al. 1996; Niture et al. 2001). Although few cysteine residues are present in the different fungal PGs, they are highly conserved. Shimizu et al. (2000) have identified seven cysteine residues in Stereum purpureum endo-PG I. The enzyme cotains three disulfide bridges Cys3 -Cys17 , Cys175 Cys191 and Cys300 -Cys303 and a free Cys167 . The conservation of these disulfide bridges in other fungal

PGs are presented by the amino acid sequence alignment of endo-PG I and other four fungal endo-PGs produced by Sclerotinia sclerotiorum (Reymond et al. 1994), Aspergillus oryzae (Kitamoto et al. 1993), Aspergillus niger (Bussink et al. 1990) and Cochliobolus corbonum (Scott-Craig et al. 1990). The alignment clearly suggested that in each endo-PG six cysteine residues are conserved (according to S. purpureum sequence numbering), although they have 40–43 % amino acid identity. Markovic & Janecek (2001) have performed the sequence alignments of forty-three fungal PGs and suggested that cysteine residues in fungal PGs are well conserved (Table 2). Aspergillus niger PG II has four disulfides: Cys30 -Cys45 , Cys203 -Cys219 , Cys329 Cys334 and Cys353 -Cys362 (van Santen et al. 1999). While the first two bridges are present in the most of the fungal PGs, the one corresponding with Cys329 Cys334 is missing in the PGs from yeasts (Cys→Val and Cys→Ala substitutions). With regard to the fourth Cys353 -Cys362 bridge, the corresponding cysteines are absent in both PGs from Claviceps purpurea and the one from Chondrostereum purpureum. Interestingly, there is only one cysteine residue (Cys45 ) conserved throughout all the fungal PGs [according to Aspergillus niger PG II (Aspni2.pg) sequence numbering]. The conservation of cysteines reflects taxonomy, i.e. the corresponding disulfides could be conserved only in the frames of the fungal groups. Catalytic mechanisms of fungal PGs Catalytic mechanisms of retaining and inverting glycosyl hydrolases involve in general two acidic residues (McCarter & Withers 1994; Davis & Henrissat 1995). In the case of retaining enzymes these residues are involved in a double displacement mechanism, via a covalent glycosyl-enzyme intermediate, and their carboxylate groups are spaced at approximately 5.5 ˚ A. In inverting enzymes, the distance between the side chains of the acidic residues is approximately 9.5 ˚ A and catalysis



9

Arg256 H+

His223 Lys258

Fig. 2. Catalytic mechanism proposed for endo-PG II of Aspergillus niger. The conserved Asp201 (Aspni2.pg numbering) acts as a proton donor, while Asp180 and Asp202 activate the hydrolytic water molecule. His223 donates the proton in the catalysis reaction. Arg180 and Lys201 are involved in substrate binding or maintain the proper ionization state of His223 . (Adapted from van Santen et al. 1999; Armand et al. 2000.)

proceeds via a single displacement mechanism. In the mechanism, one of the acidic residues acts as a general acid, donating a proton to the glycosidic oxygen of the scissile bond, whereas the second carboxylate acts as a general base, which activates a water molecule that performs a nucleophilic attack on the sugar anomeric carbon. Based on crystallographic data of the endo-PG II produced by Aspergillus niger van Santen et al. (1999) have suggested that three conserved aspartate residues (Asp180 , Asp201 and Asp202 ), present in the active-site cleft of the enzyme, are involved in the catalytic mechanism of the enzyme (Fig. 2). The distances between the conserved aspartates are 4.1 ˚ A (Asp180 -Asp201 ), 5.7 180 202 ˚ A (Asp201 -Asp202 ), respecA (Asp -Asp ) and 4.9 ˚ tively, and the conserved histidine residue (His223 ) is close by 3.5 ˚ A from Asp201 , 4.0 ˚ A from Asp180 , and 5.0 202 ˚ A from Asp , respectively. No conserved carboxylic acid residues are found at a distance of about 9.5 ˚ A from each other. This suggests that the family GH28 enzymes do not conform to the “standard” inverting mechanism. The authors also suggested that the catalytic mechanism of endo-PG II is somewhat similar to that of the phase 22 tailspike rhamnosidase enzymes (Steinbacher et al. 1994) which are not classified into GH28 family. It was also pointed out (van Santen et al. 1999; Armand et al. 2000) that a water molecule is bound between Asp180 and Asp202 , while the Asp201 carboxylic group is in a somewhat less solvent-exposed environment stabilizing its protonated state. This suggests that in endo-PG II, Asp180 and Asp202 activate the attacking nucleophilic water molecule, while Asp201 protonates the glycosidic oxygen of the scissile bond. Arg256 and Lys258 residues are involved in the substrate binding or maintain the His223 residue in the proper ionization state (Fig. 2).

Shimizu et al. (2002) have proposed another catalytic mechanism of endo-PG I produced by Stereum purpureum fungus. Endo-PG I of Stereum purpureum consists of three conserved aspartates residues (Asp153 , Asp173 and Asp174 ) which are present near to the active site of enzyme. The distances between the absolutely conserved three aspartates are 4.4 ˚ A (Asp153 -Asp173 ), 153 174 5.5 ˚ A (Asp -Asp ) and 4.8 ˚ A (Asp173 -Asp174 ). No conserved carboxylic acid residues are found at a distance of about 9.5 ˚ A from each other like in the Aspergillus niger endo-PG II. Shimizu et al (2002) have suggested that two aspartates (Asp153 and Asp174 ) act as general base in catalytic process. The Asp174 interacts with the basic residues, Lys228 and Arg226 , and this interaction would decrease the pKa of Asp174 , through making it possible for Asp174 to act as a general base. On the other hand, Asp153 interacts with Asp156 acidic residue, which would make the deprotonation of Asp153 unfavourable. However, the distance between Asp153 and Asp156 would facilitate low barrier hydrogen bonding (2.53 ˚ A). The low barring hydrogen bonding may affect the pKa of Asp153 to act as a general base. In both PGs of Aspergillus niger and Stereum purpureum, the distances between catalytic aspartate residues are not larger than 5.7 ˚ A and no conserved carboxylic acid residues are found at a distance of about 9.5 ˚ A from each other, suggesting that although both of them are classified in the family GH28 enzymes, they do not perform standard inverting catalytic mechanism. Structural aspects and active-site important amino acids residues of fungal PGs van-Santen et al. (1999) solved the first crystal structure of PG II from Aspergillus niger fungus. The crystal structure represents enzyme folded into a right handed parallel β-helix with 10 complete turns (Fig. 3a). The number of amino acids per turn varies from 22 to 39, averaging to 29 residues per turn. This variation is caused by the diversity of lengths of the loops connecting the β-strands. The average rise per turn is 4.8 ˚ A, a value typical for β-helical structures (Yoder & Jurnak 1995). The parallel β-sheets have been named PB1, PB2a, PB2b, and PB3. This naming was adopted to be consistent with the naming of the β-sheets in the PL structure, the first right-handed parallel β-helical structure solved (Yoder et al. 1993). In contrast to the three βsheets in PL, the endo-PG II β-helix is composed of four β-sheets. PB1, PB2b and PB3 are the endo-PG II’s counterparts of PB1, PB2 and PB3, respectively, of PL. The five-stranded PB2a sheet, located in β-helix turns 6 to 10, can be regarded as an N-terminal extension to sheet PB2b. Its strands are connected to those of PB2b via one residue in a left-handed α-helix conformation. The β-helix has one very small α-helix near to the N-terminus, which shields the enzyme’s hydrophobic core (Fig. 3a). Loop regions form a cleft on the exterior to the β-helix. The essential catalytic residues Asp180 , Asp201 , Asp202 , His223 , Arg256 and Lys258 are located in this cleft. Site-directed mutagenesis showed


S. K. Niture

10 (a)

(b)

(d)

(c)

Fig. 3. Schematic representation of crystal structures of some fungal PGs. The structures are viewed from the C-terminal side. (a) The endo-PG II from Aspergillus niger (PDB code: 1CZF). The active site cleft is formed by the loop regions T1 (left side) and T3 (right side). The active site residues present in the cleft are shown in different colors: Asp180 , Asp201 and Asp202 are in blue, His223 in red, Arg256 in orange, and Lys258 in purple (adapted from van Santen et al. 1999). (b) The PGA from Aspergillus aculeatus (PDB code: 1IB4). The N-glycosylated Asn219 residue is shown in purple color at the C terminus region (right side) whereas, Thr5 , Thr8 and several serine residues present in the N-terminus are shown in red color which are glycosylated by O-linked mannose residues (left side) (adapted from Cho et al. 2001). (c) The endo-PG I from Stereum purpureum (PDB code: 1KSC). A large active site cleft is formed by the parallel β-sheet, PB1, and the loops of both sides of PB1. The three aspartate residues, Asp153 , Asp173 and Asp174 , from the active site are shown in blue color, His195 in red, Arg226 in yellow and Lys228 in purple (adapted from Shimizu et al. 2002). (d) The endo-PG from Fusarium moniliforme (PDB code: 1HG8). Important active site residues are highlighted as follows: Asp191 , Asp213 and Asp212 in blue, His188 in red, Arg267 in yellow and Lys269 in turquoise (adapted from Federici et al. 2001).

that these residues are involved either in substrate binding or in the catalytic process (van-Santen et al. 1999). Recently van Pouderoyen et al. (2003) have solved the crystal structure of PG I produced by the same strain of Aspergillus niger. Endo-PG I folds into righthanded parallel β-helical structure comprising 10 complete turns and the structure is very similar (60% sequence identity) to Aspergillus niger PG II (van-Santen et al. 1999). The PG I and PG II structure shows an open active-site cleft from which substrate could easily diffuse away. Interestingly, the active-site cleft of PG I (6.4 ˚ A) is narrower than that of PG II (11.0 ˚ A). This narrowing is caused by insertion of an asparagine after The125 in a loop above the active site. In addition, the side-chain of Asn126 may play a role in hydrogen bonding the substrate. The Arg96 residue is essential for the processive behaviour of PG I at subsite −5 (Pages et al. 2001); site directed mutagenesis of Arg96 →Ser96 of PG I showed non-processive character like wild type of PG II suggesting that the Arg96 and narrowed substrate binding cleft contribute to the processive behaviour of PG I. Cho et al. (2001) solved another structure of PG produced by Aspergillus aculeatus fungus. Although the crystal structure is quite similar to that of Aspergillus

niger endo-PG II, there are also some dissimilarities. Aspergillus aculeatus PG A folds into a large righthanded parallel β-helix with overall dimensions of 65 ˚ A × 34 ˚ A ×36 ˚ A. There are 10 complete turns and the number of residues per turn varies from 22 to 29, excluding the residues in the loop regions (Fig. 3b) similar to PG II of Aspergillus niger. Superposition of the equivalent Cα positions for PG A with PG II gives the root mean square deviations of 0.76 ˚ A (330 out of 335 residues; sequence identity 67.7%). The structures of PG A and PGII show much resemblance to each other, however a notable difference between them occurs in a long T1 loop that forms the 9th complete turn and contains a catalytic tyrosine residue. The T1 loop of PG A comprises 15 residues from Gln268 whereas in PG II 17 residues are present from the equivalent Gln289 . The second major difference between PG A and PG II is the glycosylation pattern, because PG II is a recombinant protein. PG A has one N-glycosylation site and is highly O-glycosylated at the N-terminal site (Fig. 3b), whereas PG II has only one N-glycosylation site at Asn240 . As reported previously by Kester et al. (1996), there are 8 residues strictly conserved in the PG family of proteins. These include Asn157 , Asp159 , Asp180 , Asp181 , His202 , Gly203 , Arg235 , and Lys237 (ac-



11

cording to A. aculeatus PG A numbering) which are clustered at subsites −1 and +1 of PG A. The structure clearly suggested that conserved aspartate residues are directly involved in catalysis, the arginine and lysine residues in the substrate binding and histidine residue in maintaining the proper ionization state of the catalytic Asp180 . The complex model of PG A with octagalacturonic acid provides a clear explanation regarding some others amino acid residues which are involved in the catalytic process. All conserved non-glycine residues are involved in the intricate hydrogen bonding network between the catalytic residues that facilitate catalysis. More importantly, the modelling data suggested that amino group of conserved Asn157 is hydrogen-bounded to the hydroxyl group of conserved Tyr270 and the hydroxyl group of Tyr270 is close (3.2 ˚ A) to endo-cyclic O5 of galacturonopyranose. These interactions occurring on both sides of galacturonopyranose support the idea that these two residues facilitate the catalysis process by contributing considerable energy to transition state stabilization. Similarly, using molecular modelling approach Shimizu et al. (2002) have determined the structure of endo-PG I produced by Stereum purpureum with complex oligogalacturonide. Endo-PG I has also a righthanded β-helical structure (Fig. 3c). Structural superpositions of this endo-PG I with Aspergillus niger endoPG II show that 301 Cα atoms are in structurally equivalent positions, with the root mean square deviation of 1.4 ˚ A. Their comparison revealed that most of the secondary structural elements are in the same positions (Fig. 3a,c). Differences in the two structures are seen only in their C-terminal regions. The C-terminus of the Aspergillus niger endo-PG II is fixed by a disulfide bridge, whereas that of endo-PG I is fixed by an additional β-sheet structure, PB2c composed of two βstrands, 36 and 38. A large cleft is formed along the surface of the β-sheet PB1 and is surrounded by the three loops connecting β-strands 9 and 10, 12 and 13, and 15 and 16, as well as by the three loops on the other side, connecting the β-strand 23 and 24, 27 and 28, and 31 and 32. As reported earlier by Benen et al. (1996) and Bonnin et al. (2001) endo-PGs hydrolyse generally only the first glycosidic bond from the reducing end of tri or tetra-galacturonate. This type of reaction indicates that the enzymes form only one productive enzymesubstrate complex with this substrate and most likely recognize the reducing end galactofuranuronic acid residues in the +1 subsite. Shimizu et al. (2002) have provided the experimental data and suggested that in binary complex an electron density for one galactofuranuronic acid molecule was observed in the +1 subsite of endo-PG I. The tight binding of the substrate to subsite +1 was observed due to the electrostatic interactions between the carboxyl group of galactofuranuronic acid and basic amino acids present in the subsite +1. In the subsite +1, the caboxy group of galacturonate interacts with three conserved basic amino acid residues such as His195 , Arg226 and Lys228 , whereas in subsite

−1, the unique nonprolyl cis-peptide bond is believed to be involved in the binding the carboxy group of the substrate. The digalacturonate model structure suggested that three aspartate residues are located on the β-face of the −1 and +1 subsite pyranoside rings. Asp173 is at the appropriate position to be proton donor to the scissile glycosidic bond and the Asp153 and Asp174 seem to act as a general base to abstract a proton from the nucleophilic water. Further Federici et al. (2001) have solved another crystal structure of endo-PG produced by phytopathogenic fungus Fusarium moniliforme. The overall architecture consists of a right-handed parallel βhelix, resulting in the tandem repetition of 10 coils, each formed by three or four β-strands (Fig. 3d). The β-strands of consecutive turns line up to form parallel β-sheets indicated as PB1, PB2a, PB2b, and PB3. The β-sheet PB2a, starting from the 6th turn of the β-helix, is typical of PGs and is absent in bacterial PG sharing the same overall fold (Pickersgills et al. 1998). The length of the β-strands is generally short (3 to 5 residues); more variable is the length of the turns (T) between β-strands. The turns T1 (between PB1 and PB2b or PB2a) and T2 (between PB2b and PB3), and the PG-specific turns between PB2a and PB2b, are very short and often composed by only one residue in αL conformation; this residue is often an asparagine, which may contribute, through its H-bonding capability, to the significant change in the polypeptide backbone direction. The T3 turns (between PB3 and PB1) are more variable, and their length varies from 3 to 24 residues; these loops determine the formation of a deep cleft on one side of the β-helix, where the putative active site is located. Federici et al. (2001) have mutated several amino acids present in the active-site cleft of the enzyme and discussed their critical roles in complex formation with PG-inhibiting protein 2 (PGIP-2). They suggested that the residues Lys269 and Arg267 , located inside the active-site cleft, and His188 , at the edge of the active site cleft (Fig. 3d), are critical for the formation of the complex with PGIP 2. The replacement of His188 with a proline or the insertion of a tryptophan after position 270, interferes with the formation of PG-PGIP complex. Most recently, Abbott & Boraston (2007) have determined the crystal structure of native exo-PG (YeGH28) and digalacturonic acid complex with exoPG from Gram-negative enteric pathogen Yerisinia enterocolitica. The protein possessed 570 amino acids and adopts a conventional right-handed parallel β-helix topology, containing ten complete turns that comprise four discernable β-sheets, and fibronectin-type III domain grafted to its N-terminal. The protein shows significant sequence and structural similarities with other GH28 members. Eight conserved important catalytic residues which are present in other fungal endo-PGs are stringently conserved in this protein also. The fibronectin-type III domain is very common in bacterial carbohydrate-active proteins and it may be involved in a variety of molecular recognition processes, such as


S. K. Niture

12 cell adhesion, cell surface hormone and cytokine receptor. Interestingly, the authors have suggested that the fibronectin-type III domain present is in the opposite site of the enzyme than the active site of the protein therefore it may not be involved directly in catalysis of YeGH28 (Abbott & Boraston 2007). The architecture of the active site of YeGH28 was determined by soaking the protein molecule with tri-galacturonic acid ligand. There are several basic amino acids involved in stabilization of the substrate by electrostatic interactions. O-2 is in hydrogen bond contact with Arg440 and Asn406 , an amino acid that interacts with O-3. The C-5 uronate group, a definitive substituent in galacturonic acid, forms a hydrogen bond with His335 and a salt bridge with Arg240 . The Asp381 , Asp402 and Asp403 of the protein are all within 5 ˚ A away from one another. The digalacturonic product complex is orientated with its reducing end residue presenting its α-face towards the catalytic cluster, which is consistent with the PG I of S. purpureum tertiary structure. The scissile glycosidic oxygen is within 3.3 ˚ A of the catalytic acid Asp402 . Although there is no water molecule interacting with Asp381 and Asp403 in the product complex, overlapping the native structure with the complex one introduces a water molecule into the active site that is in hydrogen bond contact with Asp403 and has satisfactory geometry and position to attack the anomeric carbon (Abbott & Boraston 2007). In summary, in comparison to other right-handed parallel β-helix of PGs, endo-PG II of Aspergillus niger structure is most similar with that of the endo-PG produced by bacterial strain Erwinia carotovora (Pickersgill et al. 1998) and exo-PG of Yerisinia enterocolitica (Abbott & Boraston 2007). This is in agreement with their classification into the same family GH28 (Henrissat 1991). The Aspergillus niger and Erwinia carotovora endo-PGs have 10 complete turns. All have four β-sheets that form the β-helix, and their loop regions are also similar, in both size and location. The Erwinia carotovora PG shows only 19% sequence identity to the Aspergillus niger endo-PG II. Nevertheless, the two structures are very similar, with 265 equivalent Cα atoms (out of 335) superimposable with a root mean square deviation of 1.8 ˚ A. Similarly, endo-PG I produced by Stereum purpureum has a right-handed βhelical structure, which is common in the other pectinases, such as PLs (Yoder et al., 1993), rhamnogalacturonase (Petersen et al. 1997), and endo-PGs from Erwinia carotovora (Pickersgill et al. 1998), Aspergillus niger (van Santen et al. 1999), and Aspergillus aculeatus (Cho et al. 2001). The amino acid sequence of endoPG I has 25.8% and 40.6% identity with Erwinia carotovora endo-PG and Aspergillus niger endo-PG II, respectively. Fusarium moniliforme PG and PG II from Aspergillus niger have a sequence identity of 43.5%, and their structures are almost completely superimposable, with an average root mean square deviation between equivalent Cα atoms of 0.97 ˚ A. All secondary structure elements are conserved between the two proteins both in the β-helical region and in the region outside the β-

helix. Thirteen of the 15 additional residues of Fusarium moniliforme PG form two loops that are absent in Aspergillus niger PG II. More interestingly, the structures of PGs produced by bacterial species of Erwinia or fungal species such as Aspergillus niger, Fusarium moniliforme and Stereum purpureum showed that all the proteins exhibit a common β-helical structure and a “cleft” where catalytic residues are located (Fig. 3). Additionally, the fungal enzymes contain more structurally aligned residues; and the four disulfide bridges are conserved among them, whereas these are not conserved in the Erwinia carotovora endo-PG. This may indicate that the evolutionary divergence of the fungal β-helical proteins from the bacterial ones occurred before the divergence of the PGs. The molecular mass of Erwinia, Aspergillus and Fusarium PGs are 40 kDa, 35 kDa and 38 kDa, respectively. All these three different PGs show lower α-helix components between 1.9– 16% and 27–43% of β-sheet components suggesting that they have similar secondary structure, although they are produced by different organisms. Amino acid sequence similarity between fungal and plant PGs (Lycopesicum esculentum, Prunus persica) revealed that the similarity is seldom greater than 40% (Stratilova et al. 1993). In addition to this, Henrissat (1991) reported that amino acid sequences of different PGs do not share significant identity with other glycoside hydrolases as well as between the individual PGs, nevertheless the structure is well-conserved in different PGs. Inhibitors of fungal PG Majority of fungal PGs are directly involved in plant pathogenesis and they are termed as “virulent factors” of fungal pathogens. Therefore, it is extremely important to inhibit PGs catalytic activity either in vitro or in vivo. Several studies showed that purified PGs are inhibited by metal ions, oligogalacturonides and its substrates. Specific inhibition of PGs is the key for prevention of disease symptoms in plant caused by fungal organisms. Studies showed that in vitro the fungal PG activities are inhibited by metal ions, such as Ba2+ , Ca2+ , Co2+ , Zn2+ , Mn2+ and Hg2+ (Cananne & Doneche 2002; Mohamed et al. 2006; Martins et al. 2007). Nonspecific and non-proteinaceous inhibitors like polyphenols and tannins from American and Chinese chestnuts have been suggested to inhibit PG activities produced by Verticillium sp. and Cryphonectria parasitica, respectively (Patil & Dimond 1967; Gao & Shain 1995). More specifically, PG inhibition by substrate and its end-products were also reported. For example, the exoPG produced by Aspergillus tubingensis was competitively inhibited by galacturonic acid (Ki = 0.3 mM) as well as by reduced digalacturonate (Ki = 0.4 mM) (Kester et al. 1996). The purified endo-PG produced by phytopathogenic fungus Fusarium moniliforme was inhibited by gluconic acid D-lactone competitively (Ki = 18 mM) (Niture et al. 2001). Recently, we have also demonstrated that the same enzyme was inhibited also by pectin with Ki = 0.123 mg/ml (Niture et al. 2007).



13

The plant cell wall is resilient and structurally heterogeneous barrier composed of complex polysaccharides and diverse proteins (O’ Neill & York 2003). The role of plant cell polysaccharides in disease resistance has been reviewed recently by Vorwerk et al. (2004). In earlier studies, Albersheim & Anderson (1971) have been reported that proteins extracted from the cell wall of kidney bean hypocotyls, tomato stems and suspension-cultured sycamore cells were able to inhibit competitively the activity of PGs secreted by the fungal pathogens Colletotrichum lindimuthianum, Fusarium oxysporum and Sclerotium rolsfii. In 1980s PGIPs from different dicotyledonous and monocotyledonous (Allium porrum) plant species have been isolated and characterized (Stotz et al. 1994; Favoron et al. 1997). The biochemical properties of PGIPs isolated from different plant species and their inhibitory effects against different fungal PGs have been reviewed by Gomathi & Gnanamanickam (2004). PGIP belongs to the superfamily of leucine-rich repeat proteins; the motif being important in PGs recognition. The Phaseolus vulgaris PGIP consists of ten repeats of short (24 amino acid) leucine-rich repeat motif (Leech et al. 2000), having three domains, the central leucine-rich repeat region and two cysteine-rich flanking domains (Mattei et al. 2001). The most of plant PGIPs have a molecular mass between 34–55 kDa, and pI between 5–9 (Cervone et al. 1987; Stotz et al. 1994; Berger et al. 2000; Favaron et al. 2004). Several plants produced numerous isofoms of PGIPs, a monocotyledonous plant species Allium porrum produced near about 20 PGIP isoforms (Favaron et al. 1997; Mahalingam et al. 1999). PGIPs isolated from different plants show different specificities towards PGs produced by fungal pathogens. Most of the PGIPs are tightly bound with different PGs between pH 4.5–6. The association is stable at physiological salt concentration (150 mM) whereas the PG-PGIP complex dissociated above 500 mM salt concentration (Cervone et al. 1987). Phaseolus vulgaris PGIP-2 is the most effective inhibitor of many fungal PGs and shows different inhibition kinetics (D’Ovidio et al. 2004; Manfredini et al. 2005). For example, PGIP-2 inhibits competitively and non-competitively PGs activities produced by Fusarium moniliforme and Aspergillus niger (Federici et al. 2001; King et al. 2002) which suggests that the orientation of PGIP in the complex might differ depending on the PG ligand. These observations were supported by molecular docking studies presented by Federici el al. (2006). In Fusarium moniliforme PG-PGIP2 complex, the active-site cleft of PG is completely occupied by PGIP-2 whereas in the Aspergillus niger PG-PGIP2 complex the enzyme was still capable of binding the substrate. Site-directed mutagenesis of PG from Fusarium moniliforme at the His234 residue to lysine abolished the enzyme activity, but its binding with PGIP of Phaseolus vulgaris was not affected (Caprari et al. 1996). This indicates that the critical site for enzyme activity and binding site for PGIP are different. In short, PGIPs are versatile proteins and they are capable for recognizing different surface motifs of

functionally related but structurally different PGs. Recently, Spadoni et al. (2006) have analysed the interaction of two PGIP isoforms from Phaseolus vulgaris (PvPGIP1 and PvPGIP2) with both polygalacturonic acid and cell wall fractions containing uronic acids. They identified, in the three-dimensionalstructure of PvPGIP2, a motif of four clustered (RRKR) arginine and lysine residues (R183, R206, K230, and R252) responsible for this binding. The four residues were mutated and the protein variants were expressed in Pichia pastoris. The ability of both wild-type and mutated proteins to bind pectins was investigated by affinity chromatography. Single mutations (R183→Q183) reduced the binding by 78% and double mutations (R183→Q183 and K230→Q230) complete abolished the interaction, suggesting that the four clustered residues are essential for the pectin-binding site. Interestingly, the binding of PGIP to pectin is displaced in vitro by PGs, suggesting that PGIP interacts with pectin and PGs through overlapping although not identical regions. The author suggested (Spadoni et al. 2006) that the specific interaction of PGIP with polygalacturonic acid may be strategic to protect pectins from the degrading activity of fungal PGs. The best advantages of PGIPs are in that they do not inhibit other cell wall degrading enzymes and PGs of same plant origin (Abu-Goukh & Labavitch 1983; Cervone et al. 1990). Interestingly, pear PGIP inhibits only PGs secreted by pathogenic fungi in pear host tissue and do not inhibits PGs produced by other pathogen like Aspergillus niger and Fusarium oxysporum or PG of pear fruit (Abu-Goukh & Labavitch 1983). PGIPs of Phaseolus vulgaris does not inhibit PG of tomato, however, it inhibits exo-PG of corn pollen and sorghum (Cervone et al. 1990). PGIP is constitutive protein and differentially expressed in tomato, apple and raspberry fruits (Johnston et al. 1993; Stoz et al. 1994; Yao et al. 1999). In the decayed area of apple fruit incubated with fungal pathogens Penicillium expansum and Botrytis cinerea, there was observed a high level of PGIP gene expression, suggesting that PGIP gene is readily activated by fungal infection (Conwey et al. 1998). In Phaseolus vulgaris four PGIP genes were expressed constrictively, however, the PGIP-1 gene expression was stimulated by wounding only, the PGIP-2 was up-regulated by oligogalacturonides, salicylic acid and wounding, the PGIP-3 was induced by oligogalacturonides, and the PGIP-4 gene expression was not altered by any of these treatments (Devoto et al. 1998; D’Ovidio et al. 2004). The biological significance of PG-PGIP interaction is to limit the destructive potential of PGs and leads to accumulation of elicitor active oligogalacturonides which activate plant defence response, such as synthesis of phytoalexins (Davis et al. 1984), lignin and ethylene, expression of proteinase inhibitor I and β-1,3–glucanase and production of reactive oxygen species (Ridley et al. 2001). Most recently, Ferrari et al (2006) have suggested that biotic and abiotic stress is responsible for the accumulation of PGIP protein in Arabidopsis thaliana plant


S. K. Niture

14 and PGIP reduced the activity PG secreted by Botrytis cinerea and control its pathogenicity. Direct evidence of the involvement of PGIP in plant defence was demonstrated using plant transformation. The overexpression of pgip genes in tobacco (Nicotiana tabacum) (Manfredini et al. 2005), tomato (Lycopersicon esculentum) (Powell et al. 2000), grapevine (Vitis vinifera) (Aguero et al. 2005) and Arabidopsis (Ferrari et al. 2003) limits colonization by B. cinerea and reduces disease symptoms. In contrast, recently Katoh et al (2007) have over-expressed lemon (Citrus jambhiri) PGIP gene (RlemPGIP) in the pathogenic fungus Alternaria citri and suggested that, the RlemPGIP overexpression mutant AcOP16 has an ability to utilize pectin as a carbon source and produces both endo-PG and PGIP in culture filtrate. Interestingly, the pathogenicity of the mutant AcOP16 strain to cause a black rot symptom in citrus fruit was not changed (Katoh et al 2007). The different roles of PGIPs in the plant defence response is now clear; these proteins are induced early during fungal infection, inhibit PG function, prevent cell wall degradation, and control fungal growth and colonization. However, the active oligogalacturonides released from the cell wall of plant by the action of endo-PG can act a defence factor against pathogenic fungi (Ridley et al. 2001). These findings suggest that, during infection, fungal PGs not only cause the disease symptoms in host tissue but also provide the innate immunity to host plants by synthesis of defence factors and up-regulation of plant defence proteins such as PGIP. Conclusions and future perspectives Although, PGs are versatile proteins some of their biochemical properties are very common, for example,

Helpful for penetration and colonization of fungal pathogen in the host tissue and modulate host plant physiology (Idnurm & Howlett 2001).

majority PGs show optimum catalytic activity at acidic pH (3 to 6) (Table 1), they are monomeric proteins, some of them being natural glycoproteins. The activesite residues (carboxylate, histidine, arginine and tyrosine) are highly conserved and the X-ray structures confirm similarities also in secondary structure and the active-site cleft, despite the amino acid sequence similarity may be low. Most fungal organisms produce multiple forms of the enzyme, suggesting either that several genes are involved, or that post-translational modification results in different physical properties. During degradation of pectin or during pathogenesis, fungal pathogens express several set of PG genes. However, it is not clear why the fungal organisms express several PG genes and multiple PG proteins in host tissue. Several researchers have been investigated the regulation of PG genes and suggested that these genes are tightly controlled by glucose, the pH of environment or local host cell-sap pH. If the local pH of the host tissue is not optimal with respect to PG activity, then it is likely that – though the protein may be produced by the pathogen – it may be in an inactive state. It is essential to take into attention the biochemical data, such as pH optimum, pH stability, temperature optimum, specific activity, affinity of enzyme towards substrate (Km ), turnover number (kcat ) and knowledge about the selective inhibitor of each PG before using these enzymes in the various biotechnological applications. It should be noted that most of the fungal organisms produced several toxins in culture media, therefore application of these PGs are big task in commercial sector. Making the recombinant PG proteins may thus represent the alternative approach. The novel discovery of PGIP protein and its role in controlling PG regulation is a dynamic process. Since PGIP recognizes various PGs their specificities and in-

Interact with PGIP, stimulate plant defense factors and provide innate immunity to plant (Federici et al. 2006). Over-expression in transgenic plants induced growth and development, and change phenotype (Atkinson et al. 2002).

PGs

Used in food technology, fruit juice clarification, extraction of oil, processing of alcoholic beverages, etc. (Lang et al. 2000).

Important in host-pathogen interaction from the ecological point of view (Niture 2003). Biodegradation of pectic substances, recycling of carbon in the environment and pollution control (Kashyap et al. 2001).

Fig. 4. Possible roles of fungal PGs in nature.



15

hibition kinetics are highly variable. Several reports suggest that PGIP interacts with PG and this complex stimulates several defence factors in host tissue and controls the pathogenicity of fungal pathogen. These all different aspects of PG suggest that this protein is not only involved in plant pathogenesis but is also performing several other functions in the nature. The possible roles of PGs in nature are briefly summarized in Figure 4. Fungal PG research has made fast progress on molecular level. The future research should: (i) focus on discovery of novel PGs especially from fungi originated from extreme environment; (ii) determine biological functions of PG especially in biotechnological applications (using biochemical and structural data); and (iii) design selective inhibitors by molecular modelling approaches. Signalling of fungal PG expression, regulation mechanism between different PGs, gene evolution and generating new PGs with protein engineering may be future challenges for PG researchers. In addition to PG, the knowledge about other cell wall degrading enzymes will also be essential for controlling the fungal pathogenesis in plants. Acknowledgements SKN had a research fellowship from CSIR New Delhi, India.

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Received April 16, 2007 Accepted October 25, 2007