Updates on naringinase: structural and biotechnological aspects ...

Appl Microbiol Biotechnol (2012) 93:49–60 DOI 10.1007/s00253-011-3679-3

MINI-REVIEW

Updates on naringinase: structural and biotechnological aspects Munish Puri

Received: 7 June 2011 / Revised: 11 October 2011 / Accepted: 27 October 2011 / Published online: 13 November 2011 # Springer-Verlag 2011

Abstract Naringinases has attracted a great deal of attention in recent years due to its hydrolytic activities which include the production of rhamnose, and prunin and debittering of citrus fruit juices. While this enzyme is widely distributed in fungi, its production from bacterial sources is less commonly known. Fungal naringinase are very important as they are used industrially in large amounts and have been extensively studied during the past decade. In this article, production of bacterial naringinase and potential biotechnological applications are discussed. Bacterial rhamnosidases are exotype enzymes that hydrolyse terminal non-reducing α-L-rhamnosyl groups from α-L-rhamnose containing polysaccharides and glycosides. Structurally, they are classified into family 78 of glycoside hydrolases and characterized by the presence of Asp567 and Glu841 in their active site. Optimization of fermentation conditions and enzyme engineering will allow the development of improved rhamnosidases for advancing suggested industrial applications. Keywords Family 78 glycosides hydrolases . Rhamnosidase . Rhamnose . Pruning . Debittering . Pro-drug

Introduction Naringinase (EC 3.2.1.40), a hydrolytic enzyme detected in different species of microorganisms, hydrolyzes naringin (4, M. Puri (*) Centre for Biotechnology, Chemistry and System Biology (Biodeakin), Institute of Technology Research and Innovation (ITRI), Deakin University, Warrnambool, Victoria 3217, Australia e-mail: [email protected] M. Puri Fermentation and Protein Biotechnology Laboratory, Department of Biotechnology, Punjabi University, Punjab, India

5, 7-trihydroxy flavonone 7-rhamnoglucoside) to release Lrhamnose and naringenin (4, 5, 7-trihydroxy flavonone) (Puri et al. 2011a, b). Naringinase is a combination of two glycosidase activities (α-rhamnosidase and β-glucosidase) which are located on two separate polypeptides. Many rhamnosidases are members of the glycosyl hydrolase family GH78, which are widely distributed in Archaea, Bacteria, and Eukarya. Glycoside hydrolases (GH) from both fungal and bacterial sources are widely used in industrial applications. The terms “naringinase” and “α-rhamnosidase” are frequently used synonymously in the literature but are distinguished here in further detail. Naringin is rutinosylated flavanone and 1, 2-runtinose contains two sugar residues, a glucosidic, and rhamnosidic residue. De-glycosylation must be performed by two different glycosides, an α-rhamnosidase and a β-glucosidase (Fig. 1). Naringinase is of particular interest to the structure determination of polysaccharides, glycosides, and glycolipids (Kamiya et al. 1985). Naringin and its hydrolyzed product naringenin are useful for inhibiting acyl-coA-cholesterol-oacyltransferase activity, which helps in preventing accumulation of macrophage–lipid complex that causes hepatic diseases (Bok et al. 2000). It has several important industrial applications which have already been reviewed in detail (Puri and Banerjee 2000; Ribeiro 2011). This enzyme has attracted growing interest from biotech groups for its role in rhamnose production (Kaur et al. 2010) and the biotransformation of antibiotics and steroids (Thirkettle 2000). Moreover, its hydrolyzed products can be used as components or process materials in the synthesis of pharmaceutical, cosmetic, and food products (Busto et al. 2007). It also has biotechnological applications for the removal of the bitter agent “naringin” in citrus juices through conversion into less bitter substances (Busto et al. 2007; Kaur et al. 2009; Puri et al. 2010a, b). Therefore, naringinase (an enzyme containing both α-Lrhamnosidase and β-glucosidase activities) is an important enzyme not only for elucidation of the structures of bioactive

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Appl Microbiol Biotechnol (2012) 93:49–60

Fig. 1 Stepwise degradation of naringin by the action of naringinase expressing α-L-rhamnosidase and β-glucosidase activities. The structures of hydrolyzed products of naringin (prunin, L-rhamnose, naringenin, and D-glucose) are shown above [reproduced from Puri et al. 2011b]

compounds containing L-rhamnose but also for potential application in the food and beverage industries (Puri et al. 2010a, b). The α-L-rhamnosidase cleaves the L-rhamnose from the glycosides (such as naringin), thus enhancing its use in determining structures of compounds. Recent reviews on the status of naringinases are somewhat scarce (Yadav et al. 2010; Ribeiro 2011). This report aims to fill the gaps by focusing on bacterial sources, structural, and molecular biology aspects of rhamnosidase and their potential applications in biotechnology.

Naringinase assay Spectrophotometric determination of flavonones was used to evaluate naringinase activity. In this method, naringin reacted with diethylene glycol in alkaline solution to produce a yellow chalcone, which is measured at a wavelength of 420 nm (Davis 1947). There are difficulties in assaying naringinase activity. Habelt and Pittner (1983) elegantly distinguished between the content of naringin, prunin, and naringenin present in the assay mixture. The amount of these compounds can be estimated by combining two spectrophotometric procedures, namely treatment with strong alkali (determination of naringenin, naringin, and prunin) and treatment of the liberated aldohexoses with o-aminodiphenyl, as detailed in the published article. This method can be used for future research as it solves much of the previous inherent confusion. Romero et al. (1985) used p-nitrophenyl-L-rhamnopyranoside for the measurement of L-rhamnosidase activity of naringinase by colorimetrically following the appearance of p-nitrophenol. The use of a synthetic substrate did not affect the pH, temperature, or ionic strength optima of the enzyme. Using naringin as substrate, naringinase activity was deter-

mined by HPLC on a C18 μBondapak reverse-phase column and detection of absorbance at 280 nm (Yusof et al. 1990). A fast, effective HPLC-PAD method for the simultaneous determination of naringin, prunin, and naringenin was developed. The method was linear, precise, and selective for naringin and naringenin identification and quantification (Ribeiro and Ribeiro 2008). The developed gradient analytical method was suitable for process control of the enzymatic naringin bioconversion into prunin and naringenin. Naringinase activity can also be followed through rhamnose and glucose determination. To that end, the 2, 4-dinitrosalicylic acid (DNS) method (Miller 1959) was applied. The DNS macro-assay was modified into a micro-assay using a 96microwell plate, allowing higher speed, and large sample analysis number, sample volume reduction, and better repeatability (Nunes et al. 2010; Vila-Real et al. 2010).

Production of naringinases Microorganism are the main sources of naringinase, although these enzymes have also been found in liver tissues of the marine gastropod Turbo cornutus (Kurosawa et al. 1973) and common pig (Qian et al. 2005). There are also reports where naringinase has been isolated from plant sources such as celery seeds (Hall 1938), grapefruit leaves (Thomas et al. 1958), Rhamnus daurica (Suzuki 1962), and Fagopyrum esculentum (Bourbouze et al. 1976). It should however be noted that only processes based on microbial naringinases are feasible for industrially practicable. Naringinase is produced by many microorganisms (Table 1), mainly filamentous fungi (Aspergillus, Circinella, Eurotium, Fusarium, Penicillium, Rhizopus, and Trichoderma) (Scaroni et al. 2002). Most of the work related to


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Table 1 Sources for naringinase production

submerged fermentation of naringinase is reported elsewhere (Puri and Banerjee 2000), although a few gaps in the literature on production (Ribeiro 2011) are filled in the following paragraphs. Biochemical characterization of fungal naringinase has been carried out mainly on enzymes purified from culture filtrates and on commercial enzyme preparations from Aspergillus species such as Aspergillus terreus (Gallego et al. 2001), Aspergillus niger (Manzanares et al. 1997; Puri et al. 2005), and Aspergillus nidulans (Manzanares et al. 2007). A large number of 348 fungal isolates from 11 different sources in Thailand and China were collected to evaluate naringinase activity (Thammawat et al. 2008). Secondary screening was performed measuring α-L-rhamnosidase and β-glucosidase activities at 40°C, pH 4.0. Optimization of the medium for enzyme production in submerged fermentation allowed Czapek-Dox as the suitable medium, containing 0.1% naringin. The maximum naringinase production (117.77 U mg−1) was obtained in a medium supplemented with 3.75 g L−1 rhamnose as carbon source and 2.5 g L−1 sodium nitrate as nitrogen source. Naringinase produced by several fungi, especially A. niger, is used for elimination of bitter flavor from naringin in citrus fruit juices (Bram and Solomons 1965). A postharvest pathogenic fungus (Penicillium ulaiense) of citrus described for the first time in Taiwan has been found to produce α-L-rhamnosidase activity. A process for the production of α-L-rhamnosidase employing P. ulaiense in a stirred-batch reactor using rhamnose as the main carbon source was run for 12 days without pH control. The kinetic parameters for the growth of the fungi and enzyme production were calculated from experimental values. The purified enzyme showed good thermostability up to 60°C and good operational stability in white wine (Rajal et al. 2009). Flavonoids like naringin, naringenin, rutin, quercetin, and hesperidin (widely distributed in plants) are functional chemicals with sought-after properties in the fields of health care and food processing (Perez-Vizcaino and Duarte 2010). Different flavonoids and sugars tested for the induction of naringinase showed an increase in enzyme activity. Simultaneous production of the three enzymes was optimized following the examination of rutin as an inducer in the fermentation medium with Penicillium decumbens. Maximum enzyme activities were observed when the fungus was grown at 30°C with an initial pH of 7.0, using 8.0 g L−1 rutin and 9.0 g L−1 di-ammonium hydrogen phosphate as carbon and nitrogen sources, respectively (Mamma et al. 2004). The use of naringin to induce naringinase production from A. niger MTCC 1344 has also been documented. The highest enzyme titer of 9.68 U of enzyme per milliliter was achieved using a step-wise addition of small amounts of naringin up to 7 days of fermentation in a complex medium containing molasses, peptone, and salts (Kumar 2010). This

Source

Microorganism

References

Plant

Celery seeds (Apium graveolens) Rhamnus daurica Buckwheat (Fagopyrum esculentum) Grapefruit leaves Turbo cornutus Pig liver Aspergillus niger Penicillium sp. Aspergillus niger A. aculeatus A. kawachii A. nidulans Aspergillus terrus Rhizopus nigricans

Hall 1938

Gastropod Mammal Fungi

Yeast

Bacteria

Aspergillus niger (MTCC1344) Penicillium decumbens Aspergillus niger (BCC 25166) Aspergillus kawachii Penicillium ulaiense Aspergillus sojae Hanshula anomala, Debaryomyces polymorphus Pichia angusta X349 Cryptococcus laurentii Bacteriodes distasonis, JY-1 Thermomicrobium roseum Pseudomonas paucimobilis Clostridium stercorarium Bacillus sp. GL1 Geothermus vaporicell Sphingomonas paucimobilis Burkholderia cenocepacia Ralstonia pickettii Pseudoalteromonas sp. Lactobacillus ulaiense Lactobacillus acidophillus Lactobacillus plantarum Staphylococcus xylosus Pediococcus acidilactici

Suzuki 1962 Bourbouze et al. 1976

Ting 1958 Kurosawa et al. 1973 Qian et al. 2005 Bram and Solomons 1965 Young et al. 1989 Manzanares et al. 1997 Manzanares et al. 2001 Koseki et al. 2008 Orejas et al. 1999 Gallego et al. 2001 Shanmugam and Yadav 1995 Puri et al. 2005 Mamma et al. 2004 Thammawat et al. 2008 Koseki et al. 2008 Rajal et al. 2009 Chang et al. 2011 McMahon et al. 1999

Yanai and Sato 2000 Lei et al. 2011 Jang and Kim 1996 Jang and Kim 1996 Miake et al. 2000 Zverlov et al. 2000 Hashimoto et al. 2003 Birgisson et al. 2004 Hashimoto and Murata 1998; Miyata et al. 2005 Cardona et al. 2006 Gaston Orillo et al. 2007 Mazzaferro et al. 2008 Rajal et al. 2009 Beekwilder et al. 2009 Avila et al. 2009 Puri et al. 2010b; Puri et al. 2011a Michlmayr et al. 2011

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study was based on earlier work where naringenin (an end product of naringin hydrolysis) was used as an inducer for naringinase production from A. niger MTCC 1344. An increase in the enzyme production rate was observed when metal ions (calcium) were added to the medium (Puri et al. 2005). Reported inducers for naringinase production are rhamnose (Thammawat et al. 2008), hesperidin (Fukumoto and Okada 1973), naringin (Bram and Solomons 1965; Puri et al. 2008), and citrus peel powder (Puri et al. 2011a). Naringinase activity in a culture of Aspergillus sojae isolated from fermented Korean soybean was recently documented (Chang et al. 2011). This fungus has been safely used as a koji mould in the manufacture of soy sauce. The culture was grown in a medium containing naringin (0.5 g L−1). The purified enzyme presented a molecular weight of 70 kDa. The α-L-rhamnosidase activity of this enzyme was optimal at pH 6.0 and stable in the pH range of 5.5–8.0. The enzymatic bioconversion of naringin to prunin was efficiently performed with a 91% yield and negligible amount of naringenin (Chang et al. 2011). A low molecular weight α-L-rhamnosidase has been isolated from decaying lemon fruit fungi. The fungal strain was identified as Aspergillus flavus and assigned accession number MTCC 9606. The enzyme was purified from the culture filtrate using ultrafiltration and cation exchange chromatography on carboxy methyl (CM) cellulose. The molecular mass of the purified enzyme determined by SDSPAGE was 41 kDa. The pH and temperature optima for the enzyme were 11°C and 50°C, respectively. The purified enzyme was inhibited by alcohols and ethyl acetate, suggesting that it would not to be a promising candidate for the aroma enhancement of grape wine (Yadav et al. 2011). Low levels of rhamnosidase activity have been found in screenings performed on enological yeast strains (Rodriguez et al. 2004). Some yeast like Sacchromyces cerevisiae, Hanshula anomala, and Debaryomyces polymorphus show low level of α-L-rhamnosidase activity (McMahon et al. 1999). However, Pichia angusta X 349 is a remarkable producer of α-L-rhamnosidase (Yanai and Sato 2000). Recently, a yeast strain named Jmedeb008 has been reported to produce naringinase. Phylogenetic information generated from the 26S rDNA and 5.8S-ITS confirmed the identity of the isolate as Cryptococcus laurentii which has naringinase producing abilities. It was grown in a medium with naringin as the sole carbon source; however, it was found that the synthesis of naringinase was suppressed when glucose was available (Li et al. 2010).

Bacterial naringinase The characteristics of naringinase from fungi and plants have been thoroughly documented, although the production


of this enzyme in bacteria has to date received little attention. Few naringinases have been reported from bacterial origin, partially because their range of profitable applications has resulted in industry leading a somewhat clandestine approach to the research. Despite this, some reports on α-L-rhamnosidase activity from bacteria have been published recently. These reports have documented purification of α-L-rhamnosidases from Bacillus sp. (Hashimoto et al. 2003), Burkholderia cenocepacia (Cardona et al. 2006), Clostridium stercorarium (Zverlov et al. 2000), Lactobacillus acidophilus (Beekwilder et al. 2009), S. paucimobilis (Hashimoto and Murata 1998), and Thermomicrobium roseum (Jang and Kim 1996). Some Pseudoalteromonas species and Ralstonia pickettii, which are found in the ocean waters of subantartic environments, exhibits α-L-rhamnosidases activities in low temperature ranges of −1–8°C (Gaston Orillo et al. 2007). S. paucimobilis and Bacillus sp. GL1 show substantial α-Lrhamnosidase activities in a medium containing gellan as a carbon source (Hashimoto et al. 2003). Two new thermostable α-L-rhamnosidases from the thermophillic bacterium PRI-1686 have been reported by Birgisson et al. (2004). The α-L-rhamnosidase from Lactobacillus species has been reported (Beekwilder et al. 2009; Avila et al. 2009). In the genome of Lactobacillus plantarum, two putative rhamnosidase genes (ram1LP, and ram2LP) were identified, while in L. acidophilus, one rhmanosidase gene was found (ramALa). Gene products from all three genes were produced after introduction into Escherichia coli and were tested for enzyme activity. The RamALa enzyme was able to cleave naringin (1—— > 2 linked rhamnose) from tomato pulp; however, efficient conversion required adjustment of the pulp to pH 6. This system was used for the fermentation of tomato pulp, with the aim of improving the bioavailability of flavonoids in processed tomato products (Beekwilder et al. 2009). A bacterial strain was isolated from soil for the production of α-L-rhamnosidase activity. The phylogenetic analysis based on the 16S rDNA sequence identified the isolate as Staphylococcus xylosus, a non-pathogenic member of CNS (coagulase-negative staphylococci) family (Puri and Kaur 2010). A response surface optimization study was later conducted to optimize the fermentation medium for the improvement of naringinase production by S. xylosus. In the first step, the sources of carbon (sucrose), nitrogen (sodium nitrate), inducer (naringin), and pH levels significantly affected naringinase production. In the second step, the 22a full factorial central composite design was applied to determine the optimal levels of each of the significant variables. Using this methodology, the optimum values for the critical components were obtained as follows: sucrose, 10.0%; sodium nitrate, 10.0%; pH 5.6; biomass concentration, 1.58%; and naringin, 0.50% (w/v), respectively. Under


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optimal conditions, the experimental naringinase production was 8.45 U mL−1 (Puri et al. 2010b). In continuing the work on enhancing naringinase yield, the author used citrus peel powder to enhance enzyme activity. Puri et al. (2011a) produced naringinase with S. xylosus MAK 2 in a stirred tank reactor. The effect of different physico-chemical parameters, such as pH, temperature, agitation, and inducer concentration was investigated. The addition of Ca2+ stimulated the naringinase activity, at optimal pH of 5.5 and 30°C. A twofold increase in naringinase production was achieved by the addition of citrus peel powder to the medium. A large number of variables were optimized, in a 5-L bioreactor, leading to significantly improved enzyme production, namely an increase in sugar concentration (15 g L−1) in the fermentation medium, further increased naringinase production (8.9 IU mL−1).

Purification and characterization of bacterial naringinases Purification and characterization of bacterial naringinase are covered in this section (Table 2), while discussions on fungal naringinase are left to the existing literature (Ribeiro 2011; Yadav et al. 2010; Puri and Banerjee 2000). A soil bacterium with α-L-rhamnosidase gene was isolated from a cumulative mixed culture containing a polysaccharide of gellan as a carbon source and identified to be of Bacillus sp. known as a potent producer of gellan. α-L-rhamnosidase purified from Bacillus sp. R1 grown in the presence of naringin was a monomer with a molecular mass of 110 kDa and most active at pH 8 and 50°C. The enzyme required divalent metal ions for the activity and released α-L-

rhamnosidase from various rhamnosyl glycosides (Hashimoto and Murata 1998). Two α-L-rhamnosidases of the Bacillus sp. strain GL1 were purified and characterized. Both RhaA and RhaB were highly specific for rhamnosyl saccharides, including gellan disaccharide (rhamnosyl glucose) and naringin, and released rhamnose from substrates most efficiently at pH 6.5–7.0 and 40°C. Bacillus sp. strain GL1 cells grown in a gellan medium produced only RhaB, indicating that RhaB plays a crucial role in the complete metabolism of gellan (Hashimoto et al. 2003). Attempts were made to isolate a bacterium (Pseudomonas paucemobilis FP2001) with wider substrate specificity. α-Lrhamnosidase was extracted and purified from the cells of the bacterium with 19.5% yield. The enzyme activity was accelerated by Ca2+. The optimum pH was 7.8 and the optimum temperature was 45°C. The enzyme was purified by ammonium sulphate precipitation, dialysis, and affinity chromatography and was checked for purity on an SDSPAGE and with isoelectric focusing. α-L- rhamnosidase extracted from this strain remained stable for several months when stored at −20°C. The Km, Vmax, and Kcat for pnpR were 1.18 mM, 92.4 μmol/min, and 117,000/min, respectively (Miake et al. 2000). Two new thermostable α-L-rhamnosidases from a novel thermophilic bacterium exhibited temperature optima 70°C. Rhm A had an optimal pH 7.9 and Rhm B had a broad pH optimum of 5.0–6.9. Over 50% of activity was demonstrated by Rhm A in the pH interval 5.0–8.7 by Rhm B in the pH interval 4.0–7.9. Both enzymes exhibited over 20% residual activity after 24 h incubation was carried out at 60°C. Rhm A and Rhm B had Km values of 0.46 and 0.66 mM and Vmax values of 134 and 352 U/mg, respectively, on pnpR substrate (Birgisson et al. 2004).

Table 2 Molecular properties of bacterial rhamnosidases Microorganism

pH

Temperature (°C)

Mol wt. (kDa)

pI

Reference

Bacteroides sp. Bacillus sp. Clostridium stercorarium Pseudomonas paucimobilis Bacillus sp.

7 6.5–7 7.5 7.8 6.5–7

40 60 45 40

120 98 95 112 106

4.2 n.d. n.d. 4.6 n.d.

Jang and Kim 1996 Hashimoto and Murata 1998 Zverlov et al. 2000 Miake et al. 2000 Hashimoto et al. 2003

Thermomicrobium sp. Sphingomonas paucimobilis Ralstonia pickettii Lactobacillus sp. Staphylococcus xylosus C. stercorarium Pediococcus acidilactici

7.9 5–6.9 6.0 6.0 5.6 5.5 5.5

70 70 40 45 37 60 60

104 107 n.d. n.d. n.d. 98 74

4.6 4.5 n.d. n.d. n.d. n.d. n.d.

Birgisson et al. 2004 Miyata et al. 2005 Gaston Orillo et al. 2007 Beekwilder et al. 2009 Puri et al. 2010b Puri et al. 2011b Michlmayr et al. 2011

n.d. Not detected

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Miyata et al. (2005) reported the cloning of rhamnosidase gene (rhaM) of S. paucimobilis FP2001. The rhamnosidase clone was sequenced and expressed in E. coli. Rham has a sugar-binding domain of glycoside hydrolase family 2, which has been well conserved in β-glucuronidase, bmannosidase, and β-galactosidase in its C-terminal region. Rham belonged to a new bacterial subfamily in the glycoside hydrolase family 78 (α-L-rhamnosidase). Gaston Orillo et al. (2007) reported the production of coldactive α-L-rhamnosidase from psychrotolerant bacteria isolated from a subantarctic ecosystem in Argentina. Molecular identification revealed that four out of five selected isolates were closely related to the Pseudoalteromonas genes, while the fifth was identified as R. pickettii. The α-L-rhamnosidase activity was predominantly intracellular. The reaction demonstrated optimum pH of 6 and temperature of 40°C, maintaining 6% activity at 4°C. The enzyme was found to be thermosensitive, presenting a half-life of 4 min at 50°C. Cold-active α-L-rhamnosidase could have application in food processing technologies. The purified rhamnosidase (Ram1LP and RamALa) from Lactobacillus exhibited maximum activity at a temperatures range of 37–45°C. For both enzymes, the optimal pH was close to 6. Kinetic constants (Km) for both enzymes were determined (0.7 and 0.3 mM for rhamnose-pNP and rutin). Vmax values indicated that the activity of RamALa with rhamnose-pNP was significantly higher than that with rutin (Beekwilder et al. 2009).

Molecular and structural biology of naringinases Compared with the naringinases that have been purified and characterized, the number of genes that encode rhamnosidase activity is significantly lower. Cloning has been performed mainly by two methods: (1) construction of a library followed by selection of clones by screening for α-rhamnosidase activity using pNPR assay and (2) the construction of a library followed by selection of clones by screening for α-rhamnosidase production with polyclonal antibodies (Manzanares et al. 2007). The first α-rhamnosidase encoding gene (ramA), isolated from thermophillic anaerobic bacterium C. stercorarium (Zverlov et al. 2000) consists of 874 codons and codes for protein with a predicted molecular mass of 100 kDa and secondary structure, as detected by CD spectroscopy, consisting of 27% α-helices and 50% β-sheets. To date, only few α-rhamnosidase genes have been cloned and heterologously expressed: those from C. stercorarium, Aspergillus aculeatus, Aspergillus kawachii, and L. plantarum (Zverlov et al. 2000; Manzanares et al. 2001; Koseki et al. 2008; Beekwilder et al. 2009). The rhamnosidase (ramA) gene cloned from C. stercorarium has been expressed in E. coli (pWS1261) cells, and its


protein (Ram78A) was purified using three liquid chromatography steps (Zverlov et al. 2000). Two thermostable α-Lrhamnosidase genes from the thermophilic bacterium T. roseum were then characterized, and the protein was purified by a three-column strategy. Another microorganism (Sphingomonas paucimobilis) has been reported for the production of alpha-L-rhamnosidase (Miyata et al. 2005).The gene (rhaM) encoding α-Lrhamnosidase (Rham) from S. paucimobilis consisted of 3,354 nucleotides and further encodes a protein sequence which consisted of 1,117 amino acids. Interestingly, this Rham has no similarity to other known rhamnosidases. Analysis of rhaM mRNA indicated that the induction of the enzyme was facilitated by the addition of L-rhamnose on the transcriptional level. Some microbial sources like A. kawachii have been exploited recently for the production of thermo-stable α-Lrhamnosidase. PCR product from the amplified genomic DNA was ligated into pGEM-T easy vector (Promega, US) and transformed into E. coli strain DH5α cells. Based on the amino acid sequence deduced from cDNA, the protein possessed 13 potential N-glycosylation recognition sites and exhibited up to 75% sequence identity with the αrhamnosidase belonging to the glycoside hydrolase family 78. The purified enzyme from this fungus had a molecular mass of 90 kDa and retained more than 80% of its activity following incubation at 60°C for 1 h. Its T50 was determined to be 72°C. The enzyme was able to hydrolyze 1, 2- and a-1, 6-glycosidic bonds (Koseki et al. 2008). All studies pertaining to the purification of recombinant α-L-rhamnosidase to date utilized multiple purification steps that ultimately lead to a low recovery rate of the enzyme. A histidine tag (Hexa “His” tag) has been engineered on the recombinant α-L-rhamonisdase. In this study, a His tag was genetically attached to the rhamnosidase gene ramA from C. stercorarium to facilitate its purification from E. coli BL21 (DE3) cells containing a pET-21 d/ ramA plasmid. This work documents the first attempt at implementing a single-step method for the purification of a recombinant rhamnosidase used for the hydrolysis of naringin (Puri et al. 2010b). The purified recombinant enzyme was immobilized in Ca2+ alginate beads for investigating its hydrolysis in citrus fruit juice (Kaur et al. 2009). The histidine tag facilitated the availability of purified bacterial rhamnosidase making it attractive for an array of biotechnological applications. The properties of intracellular rhamnosidase from food lactic acid bacteria (Pediococcus acidilactici) have only recently been reported. The putative rhamnosidase genes (ram and ram 2) were heterologously expressed in E. coli with deduced protein sequences containing 653 and 525 amino acids, respectively. Surprisingly, both enzymes were


unable to hydrolyze the natural flavonone glycoside naringin (Michlmayr et al. 2011).

Three dimensional (3D) structure of naringinase The 3D structure of one of the important activities of this enzyme (α-rhamnosidase) has been determined. The enzyme from Bacillus sp. was crystallized at 20°C using the hangingdrop vapor diffusion technique with polyethylene glycol 8000 as a precipitant (Cui et al. 2006). Furthermore, same workers solved the X-ray crystal structure of rhamnosidase from Bacillus sp. by single-wavelength anomalous diffraction at 1.9 Å resolutions. The rhamnosidase forms a homodimer in the a crystal structure containing 1,908 amino acids, 43 glycerol molecules, four calcium ions, and 1,755 water molecules. The overall structure consists of five domains, four of which are β-sandwich structures designated as domains N, D1, D2, and C, and an (α/α) 6-barrel structure designated as domain A (Fig. 2). The enzyme obeys catalytic mechanism specific to glycoside hydrolase (GH) family 78 enzymes. The key residues involved in the Fig. 2 Three dimensional structure of Rhamnosidase (αRha) in complex with rhamnose. a Overall structure of monomer I in αRha–Rha (ribbon stereodiagram). Red domain N, blue domain D1, green domain D2, yellow domain A, cyan domain C. The omit map of N2.5σ corresponding to a rhamnose molecule in domain A is shown as thin blue lines. b Amino acid residues surrounding the rhamnose molecule (stereo diagram). The omit map contoured at 3 sigma corresponding to the rhamnose molecule in domain A is shown as thin blue lines (this figure has been reproduced from Cui et al. 2007)

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enzyme catalysis and or substrate recognition have been investigated by site-directed mutagenesis. Acidic residues generally function as general acid and base catalysts in glycoside hydrolases. Several negatively charged residues, such as Asp567, Glu572, Asp579, and Glu841, conserved in GH family 78 enzymes, interact with rhamnose, and mutant of these residues have drastically reduced enzyme activity, indicating that the residues are crucial for enzyme catalysis (Cui et al. 2007). During the refinement of the 3D structure, two strong electron densities regions were identified, one for the calcium located near Val423 in the loop domain D2, and the other for the calcium located near Lys953 in domain C, indicating that activity of the rhamnosidase may be dependent on divalent cations (Cui et al. 2007). Active site in GH Glycoside hydrolases are classified into EC 3.2.1 as enzymes catalyzing the hydrolysis of O- or S-glycosides. Glycoside hydrolases can also be classified according to the stereochemical outcome of the hydrolysis reaction. Hydrolysis of

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the glycosidic bond occurs exclusively by one of two mechanisms, either with retention or inversion of the anomeric configuration. In both mechanisms, a pair of carboxylic acids is utilized in the active site. Inverting enzymes use one residue as a general acid and the other as a general base catalyst, and are suitably placed, about 10.5 Å apart, to allow both a substrate and a water molecule between them (Fig. 3a). To the contrary, in retaining enzymes, one residue acts as a nucleophile and the other as a general acid/base catalyst and are separated by only 5.5 Å (Fig. 3b) (Sinnott 1990; McCarter and Withers 1994). Glu572 in combination with Asp567 or Glu841 are key residues in the catalytic mechanism, thus indicating putative general acid–base catalysis obeyed by bacterial rhamnosidase.

Biotechnological applications of naringinases With recent advances, enzyme technology presents an alternative to chemical processes, reducing both energy and material consumption and thus minimizing the generation of waste. In this context, naringinases have been demonstrated to Fig. 3 Putative mechanism of glycoside hydrolysis by (a) inverting and (b) retaining glycoside hydrolases (this figure has been cited from McCarter and Withers 1994; http://afmb.cnrs-mrs.fr/CAZY/ fam/acc_GH.html)


have applications in biotechnology, with a possible foray into industrial uses. These applications are mainly based on the hydrolytic activity of naringinase and are discussed with respect to the food and pharmaceutical industry. (1) Deglycosylation of flavonoids in Cleome arabica leaf extracts (CALE) by naringinase improves the beneficial effect of this plant extract on polymorphonuclear leukocytes (PMNs). The ability of C. arabica leaf extract treated with naringinase to reduce chemotaxis in human neutrophils may be an important therapeutic factor in the treatment of chronic diseases (Bouriche and Arnhold 2010). (2) Gellan depolymerisation: With respect to the manufacture of food additives, rhamnosidase could be used in the preparation of food additives from biopolymers as well as in the preparation of sweeteners (Giavasis et al. 2000). Rhamnosidases play an important natural role in the modification of the viscous property of gellan gum. The α-L-rhamnosidase from Bacillus sp. GL1 was used for complete depolymerization of gellan (a heteropolysaccharide to a tetrasaccharide (unsaturated


(3)

(4)

(5)

(6)

glucuronyl-glucosyl-rhamnosyl-glucose). The enzyme acted on the gellan-degrading product (rhamnosylglucose) formed after successive reactions catalyzed by gellan lyase (Hashimoto and Murata 1998). Enzymatic production of rhamnose: Large amounts of citrus peel (rich in glycosylated poly-phenolic compounds) are generated as a byproduct of the juice processing industry. This study facilitated the hydrolysis of naringin extracted from citrus peel waste. The potential of recombinant alpha-L-rhamnosidase in the manufacture of rhamnose from citrus peel was investigated. The result indicated that recombinant L-rhamnosidase has industrial applicability as well as being an interesting candidate for the production of rhamnose and prunin from citrus peel waste (Kaur et al. 2010). An efficient method was developed to produce rhamnose by inactivating b-glucosidase expressed by naringinase (Vila-Real et al. 2010). Naringin extraction from kinnow peel waste: A carbohydrate containing substrate “naringin” extracted recently from kinnow peel was investigated by the author. Kinnow (a hybrid between Citrus deliciosa and Citrus nobilis) peel, a waste rich in glycosylated phenolic substances, is the principal by-product of the citrus fruit processing industry. Recombinant α-L-rhamnosidase purified from E. coli cells using immobilized metalchelate affinity chromatography (IMAC) was used for naringin hydrolysis. The purified enzyme was inhibited by Hg 2+ (1 mM), 4-hydroxymercuribenzoate (0.1 mM) and cyanamide (0.1 mM). The purified enzyme established hydrolysis of naringin extracted from kinnow peel, thus endorses its industrial applicability for producing rhamnose (Puri et al. 2011b). Birgisson et al. (2004) produced α-L-rhamnosidase from E. coli and developed a bioreactor for narigin hydrolysis. Tomato pulp digestion: The rhamnosidases from L. plantarum have been shown to convert flavonoid rutinosides (such as rutin from tomato) into wellabsorbed glucosides. Such activity implies that probiotic lactobacilli when present in gut microflora may enhance flavonoid bioavailability (Beekwilder et al. 2009). Debittering citrus fruit juices: Bitterness in citrus fruit juices is due to the presence of naringin, which may be removed by treating the juice with naringinase (Puri et al. 2008). In this direction, naringinase has been immobilized on various supports for achieving hydrolysis of naringin and eventually debittering citrus fruit juice (Puri and Banerjee 2000; Prakash et al. 2002; Ferreira et al. 2008; Ribeiro 2011). The immobilized naringinase on polyvinyl alcohol has been used for the debittering of juices (Busto et al. 2007). Additionally, the use of

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purified recombinant enzyme for treating citrus juice, a maiden report where recombinant alpha-L-rhamnosidase has been immobilized in Ca2+ alginate beads for extrapolating its hydrolysis in kinnow fruit juice. This study reports activity and stability of the free and immobilized enzyme (Puri et al. 2010a). Recently, immobilization of naringinase using mesoporous silica MCM-41 via adsorption with glutaraldehyde has been documented. These supports possess tunable pore size, large surface area, high adsorption capacity, and an ordered porous network for free diffusion of the substrates and reaction products. The immobilized catalyst showed excellent thermal stability and storage stability and could be recycled six times for the treatment of white grapefruit juice (Lei et al. 2011). (7) Preparation of prunin: Prunin possesses anti-inflammatory and antiviral activity against DNA/RNA viruses (Kaul et al. 1985). Pure prunin in high yield was obtained from naringin when immobilized naringinase pretreated with alkaline buffer was used for the preparation. The obtained flavonoid prunin has variable antiviral activity against DNA/RNA viruses. The flavonoid prunin possesses anti-inflammatory activity and may be used as sweetening agent in diabetes therapy. The natural flavonone glycoside of naringenin has also been reported to prevent gastric mucosal ulceration in animal models. The studies later observe gastroprotective effect of the glycoside and naringin on ethanol-induced gastric injury. Other rhamnosides acts as cytotoxic rhamnosylated terpenoids, as signal substances in plants or play a role in antigenicity of pathogenic bacteria (Roitner et al. 1984). The separation of bittering components (by enzyme rhamnosidase) from the citrus juices was investigated to identify compounds (limonin, naringin, and naringenin) that possess anticarcinogenic benefits. Plant flavonoids may be useful for the treatment of cardiovascular disease as well as associated conditions such as obesity, hepatic steatosis, and type 2 diabetes. Flavonoid naringenin-7-O-glucoside is a potential therapeutic agent for treating or preventing cardiomyopathy associated with doxorubicin (Wood and Bhat 1988; Han et al. 2008). (8) Aroma enhancement: Rhamnosidase activity of naringinase in combination with β-glucosidase and arabinosidase was considered suitable for aroma enhancement in wine making. The enzyme was immobilized to a solid carrier with the aim of developing a continuous process for wine aroma enhancement (Gallego et al. 2001). (9) Prodrug therapy: A drug is released from its prodrug by enzyme action. A system was designed in which drugs of interest are capped with rhamnose and are released by rhamnosidase enzyme. The carbohydrate

58

structure of rhamnosidase was specifically engineered through enzymatic deglycosylation and chemical reglycosylation to activate α-L-rhamnopyranoside to determine its use in prodrug therapy (LEAPT-lectindirected enzyme activated prodrug therapy). Highly localized prodrug activation was achieved, and glycosylation enhanced the rate of uptake of the protein component from the serum, while a reduction in time for potential immunogenic exposure was reported (Robinson et al. 2004). Prodrugs of doxorubicin and 5-fluorouracil capped by the nonmammalian L-rhamnosyl were released by rhamnosidase to its liver target (Garnier et al. 2010). (10) Naringinase has been used to hydrolyze glucolipids, leading to the synthesis of unique special fatty acids. A di-rhamnolipid was cleaved by naringinase from P. decumbens leading to a mono-rhamnolipid and Lrhamnose (Magario et al. 2009).

Further research There is a great deal of enthusiasm towards the application of this enzyme in biotech industry, but realizing this will not be possible unless more studies on “fermentation processes” on an industrial scale are carried out to secure cost-effective availability of α-L-rhamnosidases. Although many rhamnosidase coding genes are identified from various bacteria, none have been used for commercial production of the enzyme. There is also a scientific need to crystallize α-L-rhamonisdase from other sources to obtain structural variation and complete hydrolysis of its natural substrate “naringin”, which is responsible for bitterness in fruit juices. Random mutagenesis and rational protein design are typical tools for improving enzyme activity in the field of protein engineering (Bornscheuer and Pohl 2001). Of this tools, error prone PCR (epPCR) and random method of mutagenesis can easily be applied to investigate selectivity of naringinase with respect to hydrolyzing flavonoids. Directed evolution may result in mutants with remarkable/unexpected changes that eventually lead to increased enzyme activity. Thus, optimization of fermentation conditions and enzyme engineering will allow the development of improved rhamnosidase. Acknowledgement The author would like to thank Director, Centre for Biotechnology and Interdisciplinary Sciences (CBIS), ITRI for providing the necessary facility to carry out this work at Deakin University, Australia. Some of the data presented in this write-up has emanated from the research work carried at Department of Biotechnology, Punjabi University, India. Conflicts of interest The author declare that he has no conflict of interest.


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