Chitinase from Thermomyces lanuginosus SSBP and ...

Chitinase from Thermomyces lanuginosus SSBP and its biotechnological applications

Faez Iqbal Khan, Krishna Bisetty, Suren Singh, Kugen Permaul & Md. Imtaiyaz Hassan Extremophiles Microbial Life Under Extreme Conditions ISSN 1431-0651 Extremophiles DOI 10.1007/s00792-015-0792-8

1 23

Your article is protected by copyright and all rights are held exclusively by Springer Japan. This e-offprint is for personal use only and shall not be self-archived in electronic repositories. If you wish to self-archive your article, please use the accepted manuscript version for posting on your own website. You may further deposit the accepted manuscript version in any repository, provided it is only made publicly available 12 months after official publication or later and provided acknowledgement is given to the original source of publication and a link is inserted to the published article on Springer's website. The link must be accompanied by the following text: "The final publication is available at link.springer.com”.

1 23

Author's personal copy Extremophiles DOI 10.1007/s00792-015-0792-8

REVIEW

Chitinase from Thermomyces lanuginosus SSBP and its biotechnological applications Faez Iqbal Khan1,2 · Krishna Bisetty1 · Suren Singh2 · Kugen Permaul2 · Md. Imtaiyaz Hassan3

Received: 7 September 2015 / Accepted: 3 October 2015 © Springer Japan 2015

Abstract Chitinases are ubiquitous class of extracellular enzymes, which have gained attention in the past few years due to their wide biotechnological applications. The effectiveness of conventional insecticides is increasingly compromised by the occurrence of resistance; thus, chitinase offers a potential alternative to the use of chemical fungicides. The thermostable enzymes from thermophilic microorganisms have numerous industrial, medical, environmental and biotechnological applications due to their high stability for temperature and pH. Thermomyces lanuginosus produced a large number of chitinases, of which chitinase I and II are successfully cloned and purified recently. Molecular dynamic simulations revealed that the stability of these enzymes are maintained even at higher temperature. In this review article we have focused on chitinases from different sources, mainly fungal chitinase of T. lanuginosus and its industrial application. Keywords Chitinases · TIM-barrel fold · Thermomyces lanuginosus · Biotechnological applications · Thermophilic microorganisms · Thermostable enzymes

Communicated by S. Albers. * Faez Iqbal Khan [email protected] 1

Department of Chemistry, Durban University of Technology, SB Campus, POBOX 1334, Durban 4000, South Africa

2

Department of Biotechnology and Food Technology, Durban University of Technology, Durban 4000, South Africa

3

Center for Interdisciplinary Research in Basic Sciences, Jamia Millia Islamia, New Delhi 110025, India

Introduction Chitinase enzymes hydrolyze the naturally occurring polysaccharide chitins, a linear polymer of β (1,4) N-acetyl β-dglucosamine (Henrissat 1999). These are ubiquitous and present in all forms of life including bacteria, fungi, plants, animals and also in viruses (Hamid et al. 2013). They form a class of highly conserved enzymes and exhibit diverse function. Chitinases possess an extraordinary ability to hydrolyze highly insoluble chitin polymer directly to the low molecular weight chito-oligomers, which are widely used in agricultural, biotechnological, industrial and medical fields (Di Giambattista et al. 2001). In recent years, chitinase has been a key focus of research owing to its important biophysiological functions and applications (Di Giambattista et al. 2001). It plays a crucial role in the defense of plants against chitin-containing pathogens and also serve as an effective biocontrol agent against phytopathogens (Khoushab and Yamabhai 2010). Fungal chitinase primarily belongs to the family of 18 glycosyl hydrolases (Gilkes et al. 1991) and displays a high sequence homology with class III plant chitinases (Hayes et al. 1994). The family of 18 fungal chitinases comprises of 5 domains, viz., catalytic domain, N-terminal signal peptide region, chitin-binding domain, serine/threonine-rich region and C-terminal extension region (Hartl et al. 2012). The signal peptide at the N-terminus is an indicator of the protein secretion that targets the protein outside the cells through secretory pathways (Neuhaus et al. 1991). The catalytic domain is the most important domain of this enzyme which is responsible for the chitin hydrolysis. A comparison of the amino acid sequences of chitinases reveals two highly conserved motifs in the family of GH18 chitinases: DXXDXDXE and SXGG, corresponding to the catalytic

13

Author's personal copy Extremophiles

domains and substrate-binding sites, respectively (Henrissat and Bairoch 1996). Glu (E) and Asp (D) are highly conserved within the catalytic domains of chitinases, indicating their direct involvement in the hydrolysis of glycosidic bond (Li and Greene 2010; Liu et al. 2012). In fungi, chitinases have diverse role in nutrition, morphogenesis, defense and development processes (Hartl et al. 2012). Other significant application of fungal chitinases include the possibility for improvement of plant resistance using genetic manipulation techniques (Punja and Zhang 1993). Fungal chitinases are also employed in insect control. Chitinases have several industrial and agricultural applications: the ability to degrade varieties of pest and convert them into valuable products for biocontrol application (Scholte et al. 2004; Hamid et al. 2013). In addition, chitinases play important roles in the fungal protoplast generation, single-cell protein production, preparation of bioactive chitooligosaccharides and in the degradation of shellfish waste (Chang et al. 2007). In the previous study, chitinases from Thermomyces lanuginosus was isolated, cloned, expressed and purified (Meng Zhang et al. 2015; Khan et al. 2015). Structure predictions of the thermostable chitinases using “Homology modeling” and “ab initio” methods revealed the presence of TIM-barrel fold in its framework (Khan et al. 2015). The molecular dynamic (MD) simulations implemented in GROMACS (Van Der Spoel et al. 2005) suggested that the chitinases were able to maintain their stability at temperatures ranging from 300 to 350 K, comparable to experimental results (Meng Zhang et al. 2015; Khan et al. 2015). The present review focuses mainly on the fungal chitinase obtained from T. lanuginosus, a thermophilic deuteromycete that grows between 20 and 60 °C, with an optimum growth temperature of 50 °C, and optimum growth pH of 6.5. Some of them are thermostable and very suitable for industrial applications (Singh et al. 2003).

Chitinase Chitinases are hydrolytic enzymes that cleave the β (1,4) linkage of chitin polymer into N-acetyl β-d-glucosamine monomers (Muzzarelli 1999). This accounts for an extraordinary performance since the substrate chitin is highly insoluble and crystalline solid which occurs in innumerable forms depending on the role in the living tissue. They form a class of highly conserved enzymes and exhibit diverse functions in almost all organisms. Presently, chitinase has been a key focus to researchers owing to its diverse function and application (Hartl et al. 2012).

13

Chitinase sources Chitinases are present in plant seeds, stem, tuber and flower. It is induced by the attack of phytopathogens as pathogenesis-related proteins in plant self-defense or by contact with elicitors such as chitooligosaccharides or growth regulators such as ethylene (Koga et al. 1997). Not only defense, but also they have additional role in growth and development supported by the fact that plant chitinases have been detected in the early stages of seed development (Gijzen et al. 2001; Santos et al. 2007). In plants, chitinases generally belong to endochitinase category and have smaller molecular mass than the insect chitinases. They are found to inhibit fungal growth along with other enzymes like the β-1,3-glucanases that are present in citrus fruits, corn, potatoes, tobacco, beans, tomatoes, yam and peas (Flach et al. 1992; Mayer et al. 1996). Chitinases of insects are degrading enzymes that function during ecdysis (Zhang et al. 2011). These are primarily endochitinases that randomly cleave chitin present in the cuticle to chitooligosaccharides that are further hydrolyzed by exoenzymes to N-acetyl-glucosamine monomer, which is reused to synthesize a new cuticle (Chaudhari et al. 2013). Chitinases from Bombyx mori, Manduca sexta have been extensively studied. In addition, insect chitinases have a defensive role against their own parasites (Koga et al. 1997). Chitinases are largely found in crustaceans such as prawns, shrimps and krills, where they are induced before molting process in the integument (Aronson et al. 2003). Bacteria that produces chitinases include Serratia, Chromobacterium, Klebsiella, Bacillus and Streptomyce, while fungal chitinase producers are Trichoderma, Oenicillium, Lecanicillium, Neurospora, Mucor, Metarhizium, Beauveria, Lycoperdon and Aspergillus. These microorganisms produce large amount of chitinase than animals and plants (Ubhayasekera and Karlsson 2012; Hamid et al. 2013). Most of the bacterial chitinases seem to have different catalytic centers than the plant chitinases due to fewer residue deletions between the putative catalytic residues (Hoell et al. 2006). Chitin is not degraded intracellularly because of its size, molecular complexity, insolubility and heterogeneous composition (Itoh et al. 2013). The microbes extracellularly secrete enzymes with different specificity to transform or hydrolyze chitin (Cottrell et al. 1999). Yeast and fungal chitinases participate in morphogenesis where hydrolytic cleavage of the chitin polymer is crucial for hyphal growth, branching, septum formation and spore germination (Selvaggini et al. 2004). Until 1994, it was believed that humans do not produce chitinases due of the lack of endogenous chitin in human tissues suggesting no requirement for enzymatic turnover


and restructuring of chitin molecules. However, recent studies have reported 9 mammalian chitinases or chitinaselike genes that exist in human and all belonging to the family 18 glycosyl hydrolase (Boot et al. 2001; Fusetti et al. 2002; Bierbaum et al. 2005; Lee et al. 2007). The first chitinase in human was described by Hollak et al. (1994). They observed high chitinolytic activity in serum of Gaucher disease-affected patients. It was named chitotriosidase because of its ability to hydrolyze chitotriose. In the progressive year, Bleau et al. (1999) have identified and characterized four other mammalian chitinase-like enzymes in search of other chitinases that could possibly compensate for chitotriosidase deficiency (Ober and Chupp 2009; Lee et al. 2011). Boot et al. (2001) discovered another acidic mammalian chitinase (AMCase) which is expressed in the lung macrophages, gastric epithelia, and in the pulmonary epithelia during asthmatic inflammation and possesses high catalytic activity at pH 2.0 (Zhu et al. 2004).

Classification of chitinases On the basis of sequence similarity, chitinases are grouped into glycosyl hydrolase families 18 and 19 (Henrissat and Bairoch 1993). These two families differ in their sequence similarity and display different three-dimensional structure suggesting their evolution from different ancestors. Chitinases form a unique group of enzyme that can efficiently hydrolyze polymeric chitin by degrading into chitooligosaccharides with a minimum chain length of n = 2. The family 18 chitinases possess a characteristic (α/β)8 TIM-barrel domain, comprising of eight α-helices and eight β-strands (Khan et al. 2015). They are widely distributed in bacteria, fungi, viruses, insects, plants and animals. The family 19 chitinases are similar to lysozyme and

N-acetyl-β-glucosaminidase

Chitobiosidase

chitosanase and are found in plants and some Streptomyces strain. Their catalytic domains possess high α-helical content. Substrate-assisted catalysis is preferred catalytic mechanism of family 18 chitinases. While, a general acid and base mechanism has been reported in case of family 19 chitinases (Sasaki et al. 2006). The family 18 chitinases are sensitive to a potent chitinase inhibitor allosamidin, while family 19 chitinases are not (Koga et al. 1997). Family 18 chitinases are capable of cleaving GlcNAc–GlcNAc and GlcNAc–GlcN linkages, whereas members of family 19 chitinases cleave GlcNAc–GlcNAc and GlcN–GlcNAc (Watanabe et al. 1999). The chitinases are also classified according to their N-terminal sequence, isoelectric pH, localization, inducers and presence or absence of signal peptide. Class I chitinases are reported in plants, whereas class II enzymes are found in bacteria, fungi and plants. Class III includes endochitinases that have a shallow and open active site architecture (Terwisscha van Scheltinga et al. 1996) and do not display any sequence similarity to class I or II enzymes. Class IV chitinases exhibit a similar characteristics to that of class I chitinases, but they are significantly smaller than class I chitinases (Patil et al. 2000). Class V chitinases are exochitinases comprising a deep and tunnel-shaped active site (van Aalten et al. 2000; Bortone et al. 2002) and have been detected in studies on symbiotic plant–microbe interaction (Salzer et al. 2004). The enzymatic hydrolysis of chitin is carried out by a chitinolytic system classified into: endochitinases (EC 3.2.1.1.4), exochitinases or chitobiosidases (EC 3.2.1.29), and N-acetylβ-glucosaminidases (EC 3.2.1.52) (Fig. 1). Endochitinases cleave along the internal chain of chitin randomly, generating low molecular mass oligomers of N-acetyl glucosamine and eventually giving diacetylchitobiose as a predominant product. On the contrary, exochitinases hydrolyze terminal

Endochitase and Lysoyme

Fig. 1 Schematic representation of action of chitinases on chitin

13


non-reducing sugars releasing diacetylchitobiose without production of N-acetyl glucosamine or oligomers. N-acetylβ-glucosaminidases remove sugar residues from the nonreducing end of the chitin chain and catalyze the breakdown of diacetylchitobiose as well as chitotriose and chitotetraose into N-acetyl glucosamine monomers (Jung and Park 2014).

Fungal chitinases Fungal chitinases primarily belong to the family 18 of the glycosyl hydrolase superfamily (Henrissat 1999) and display a high sequence homology with class III plant chitinase (Hayes et al. 1994). The basic structure of family 18 fungal chitinases comprised of 5 domains: (1) catalytic domain, (2) N-terminal signal peptide region, (3) chitinbinding domain, (4) serine/threonine-rich region and (5) C-terminal extension region (Merzendorfer and Zimoch 2003). The signal peptide is an indicator of protein secretion that targets the protein outside cell via secretory pathway (Khan et al. 2015). The signal peptide is cleaved by signal peptidase releasing the mature protein outside of the cell during protein secretion across the cell membrane. Chitinases lacking this domain are intracellular and may function during morphogenesis (Takaya et al. 1998b; Seidl et al. 2005). The catalytic domain is most important domain because it is responsible for chitin substrate hydrolysis. Comparison of amino acid sequence of chitinases reveals two highly conserved motifs in family GH18 chitinase: DXXDXDXE and SXGG correspond to the catalytic domain and substrate-binding site, respectively (Henrissat and Bairoch 1996). Glu and Asp residues are highly conserved within the catalytic domain of chitinases indicating their direct involvement in hydrolysis of the glycosidic bond (Arimori et al. 2013). Degradation of chitin begins when insoluble chitin binds to the chitin-binding domain. Lack of this domain has no effect on enzyme activity to soluble chitin, but activity is lost to insoluble substrate (Tjoelker et al. 2000). Insoluble chitin binds to chitin-binding domain by highly conserved aromatic residues like tryptophan, phenylalanine and/or tyrosine (Zhang et al. 2002; Ferrandon et al. 2003). Serine/threonine-rich region acts as a linker between the catalytic domain and the chitin-binding domain and is necessary for maintenance of fungal chitinase stability. These regions of fungal chitinase are generally glycosylated with sugar chains which help to resist hydrolysis by proteases (Arakane et al. 2003). However, most of the fungal chitinases lack the serine/threonine-rich region, chitin-binding domain, and C-terminal extension region suggesting that these domains are unnecessary for chitinase

13

activity because naturally occurring chitinases that lack these regions are still enzymatically active (Takaya et al. 1998a). In general, the molecular masses of fungal chitinase range between 30 and 200 kDa and pH optima range from 4.0 to 7.0. The optimal temperature for fungal chitinase activity varies between 20 and 40 °C. The enzyme activity of fungal chitinases is significantly affected by metal ions. Generally Mg2+, Ca2+, Na+ and K+ act as an activator of fungal chitinase, while heavy metal ions like Hg2+, Ag+, Fe2+ and Cu2+ have significant inhibitory effect (DuoChuan 2006; Hartl et al. 2012). In fungi, chitinases have diverse roles in nutrition, morphogenesis, defense and development processes. For example, mutation of the chitinase gene (CTS1) in Saccharomyces cerevisiae prevents the separation of cells after division and results in cell clumping. While, exogenous expression of chitosanase and chitinase influences morphogenesis in the Schizosaccharomyces pombe (Shimono et al. 2002). Reports suggest a role of fungal cell wall chitinases in filamentous fungal sporulation. Since, the chitinase inhibitors demethylallosamidin or allosamidin resulted in inhibition of fragmentation of hyphae into arthroconidia (Sandor et al. 1998). Trichoderma spp. are considered as powerful biocontrol agents against soil-borne fungal pathogens amongst other chitinolytic fungi and bacteria (Lorito et al. 1993). Other reports have emphasized on the mycoparasitic effect of Trichoderma on fungi which results in induced resistance and increased development in plants (Bolar et al. 2000). Other significant applications of fungal chitinases include the possibility for improvement of plant resistance using genetic manipulation techniques. For example, the chi42 gene of T. harzianum encodes an endochitinase possessing a much stronger anti-fungal activity against a number of phytopathogenic fungi. It is constitutively expressed in apple, tobacco and potato (Lorito et al. 1998; Bolar et al. 2000). Fungal chitinases are also employed in insect control. A thorough phylogenetic study of chitinases from sequenced fungal genomes classified them into three different subgroups A, B and C as shown in Fig. 2 (Seidl et al. 2005). Subgroups A and B included all previously recognized fungal chitinases, while subgroup C comprised a novel group of high molecular weight chitinases. Subgroup A chitinase has an average molecular mass of 40–50 kDa and possesses a catalytic domain, but lack chitin-binding domain (CBM). Also they include highly conserved and profusely expressed chitinases of the fungal kingdom. Several subgroup A chitinases are extracellular proteins expressing an N-terminal signal peptide which guides them to the secretory pathway while others either posses an


Subgroup A

SP

GH 18

Subgroup B

SP

GH 18

distinguishes it from subgroups A and B chitinases indicating towards a specialized and distinctive role of these enzymes in chitin hydrolysis (Gruber et al. 2011).

(Cellulose)

Subgroup C LysM–BD Chitin-BD

SP

GH 18

Fig. 2 Domain organization of fungal chitinases. SP signal peptide, GH 18 glycoside hydrolase family 18, BD binding domain

ER-targeting sequence or are predicted to be intracellular proteins (Gruber et al. 2011). Subgroup B chitinases are highly variable in their domain structure as well as size. Their molecular masses range between 30 and 90 kDa. A feature that distinguishes subgroup B chitinases from other members is the presence of CBMs frequently (Seidl et al. 2005; Gruber et al. 2011). The number of subgroup B members highly varies among different fungal species. Most ascomycetes (Mg. grisea., N. crassa, A. niger, A. nidulans, Mycosphaerella fijiensis, Myc. graminicola and Coccidoides immitis) and basidiomycetes (Postia placenta, Phanerochaete chrysosporium, Laccaria bicolor) are characterized by the presence of only 2–3 ORFs encoding subgroup B chitinases (Seidl et al. 2005; Yang et al. 2013). Subgroup C comprises of a novel category of fungal chitinases. Their molecular masses range between 140 and 170 kDa. They possess a characteristic N-terminal signal peptide that targets them to the secretory pathway. Presence of a CBM18 and several LysM-motifs clearly distinguishes them from other fungal chitinases. Interestingly, the unique domain architecture of subgroup C chitinase members Table 1 Enzymes from Thermomyces lanuginosus with their optimum pH and temperature

Thermostable chitinase Thermomyces lanuginosus was first isolated in 1899 (Sutherland et al. 1989). It is a thermophilic Deuteromycete (unicellular or septate and reproduces asexually by forming aleurioconidia) that grows between 20 and 60 °C with an optimum growth temperature of 50 °C (Singh et al. 2003). The optimum growth pH of most strains of T. lanuginosus is 6.5. It is reported to be non-cellulolytic and probably grows commensally with cellulolytic fungi by utilizing the sugar generated by these fungi and possibly using their mycelial breakdown products (Singh et al. 2000b; Gramany et al. 2015). Some of them are thermostable and very suitable for the industrial applications. T. lanuginosus secretes many enzymes (Fischer et al. 1995; Chaudhuri and Maheshwari 1996; Li et al. 1997; Bharadwaj and Maheshwari 1999; Singh et al. 2000a; Odibo and Ulbrich-Hofmann 2001; Gulati et al. 2007; Rezessy-Szabo et al. 2007; Khucharoenphaisan and Sinma 2010; Fang et al. 2014) that are listed in Table 1. Table 1 suggests that thermophilic microorganisms like T. lanuginosus produces thermostable enzymes that are able to survive at high temperatures as well as extremes of pH conditions (Haki and Rakshit 2003). These thermostable enzymes include xylanases, lipases, amylases and chitinases that exhibit very high specificity and efficiency at extreme conditions and, therefore, have substantial potential in many industrial purposes (Haki and Rakshit 2003). The utility of these enzymes in the paper, pharmacy and food industries are extensively studied (Haki and Rakshit 2003). Xylanase was produced by cloning of rapid-amplifying mesophiles using recombinant DNA technology (Stephens

Enzyme

Optimal pH

Optimal temperature (°C)

References

Lipase β-Galactosidase Invertase Protease Trehalase Hemicellulases Glucoamylase Α-Amylase Phytase

8.0 6.7 to 7.2 6.5 5.0 and 9.0 5.0 6.0 6.0 5.0 5.0

60 – 60 70 50 50 70 60 70

Arima et al. (1972) Fischer et al. (1995) Chaudhuri and Maheshwari (1996) Li et al. (1997) Bharadwaj and Maheshwari (1999) Singh et al. (2000a) Odibo and Ulbrich-Hofmann (2001) Odibo and Ulbrich-Hofmann (2001) Gulati et al. (2007)

α-Galactosidase

5–5.5

65

Rezessy-Szabo et al. (2007)

Xylanases

6.0–7.0

60–75

Khucharoenphaisan and Sinma (2010)

13


et al. 2014). In industry, xylanase has been used in the process of pulp-bleaching for reducing the chlorine chemicals (Gruber et al. 1998). Despite their superior stability at high temperatures, various efforts such as site-directed mutagenesis, introduction of extra disulfide bonds have been performed to increase the thermostability (Gruber et al. 1998). Various alkali-stable and thermostable mutants of the xylanases were created using random mutagenesis and DNA-shuffling techniques (Stephens et al. 2014). Chitinase is another enzyme produced from T. lanuginosus that possesses numerous application in diverse fields like food industry, medicine, biotechnology, agriculture, etc. The present day research work focuses on the isolation, characterization, cloning, expression, modeling, simulations and stable mutant predictions of the thermostable chitinase are going on in many labs. Most of chitinolytic fungi have been found to produce more than one kind of chitinase. The carbon source in medium can significantly affect the chitinase production of fungi. The chitinolytic activity of T. lanuginosus chitinase was found when colloidal chitin as carbon source was present in the medium. Chitinase production of T. lanuginosus SSBP is strictly controlled by repressor/inducer system, in which by-products function as inducer whereas carbon source as repressors. Chitinolytic enzyme activity of T. lanuginosus slowed down after 96 h of incubation due to accumulation of by-products, which repress chitinase secretion by negative feedback mechanism. The recombinant chitinase I and II enzymes exhibited optimum activity at pH 5.0 and 4.0, respectively. The chitinase I has an optimal activity at 50 °C and retained 56 % of its activity at 60 °C after 30 min, while chitinase II is optimally active at 40 °C and retained 71 % of its activity at 50 °C after 60 min. Both enzymes produce chitobiose as the major product using different substrates. The chitinase II displays antifungal activity against Penicillium verrucosum and Aspergillus niger. These activities could be useful in the environmental degradation of chitinous wastes as well as for biotechnological applications. Sequence analysis suggested that chitinase I and II introns are 58.5 and 56.7 % A + T-rich, respectively, as compared to the 44.3 %, 15.8 % A + T-rich coding regions. The chitinases of T. lanuginosus have G + C-rich coding regions (54–56 %) that may offer stability to protein molecules (Meng Zhang et al. 2015; Khan et al. 2015).

Structure analysis Chitinase II (GenBank: AIW06012) was similar to the fungal class III chitinase with the highest sequence identity of 65 % belonging to Byssochlamys spectabilis chitinase III having an e value of 3e−157 in the NCBI non-redundant database. In the protein data bank (PDB) search we

13

identified Saccharomyces cerevisiae chitinase as a suitable template for the structure prediction (Khan et al. 2015). Similarly, chitinase I (GenBank: AIW06013) was very much similar to the chitinase of T. lanuginosus which showed 98 % identity. Chitinase I is similar to chain A of the chitinase present in Aspergillus fumigatus (PDB ID: 1WNO) with an identity of 62 % selected to as a suitable template for chitinase I. InterProScan (Quevillon et al. 2005) analysis revealed that both sequences contain a domain that is similar to the glycoside hydrolase family 18, a group of enzymes that hydrolyzes the glycosidic linkages between the carbohydrate units. The given proteins were also found to be a member of 4164978 and 136700 clusters of ProtoNet (Sasson et al. 2003) and SYSTERS (Krause et al. 2000), respectively, which are groups of chitinase-like enzymes. Structure analysis of a protein provides detailed information about its function and mechanism of action (Hassan et al. 2008, 2010a, b, 2012, 2013; Hassan and Ahmad 2011). Hence, we modeled the structure of both chitinase I and II to relate the structure with their functions. Furthermore, structure analysis will offer a new insight into ligand/ substrate binding and recognition (Hassan et al. 2007a, b; Hassan 2015). The topology for the given structures of chitinases was generated using PDBsum (de Beer et al. 2014) to understand a detailed structural features. The structurally conserved (β/α)8 TIM-barrel was observed in the framework of chitinase I and II (Fig. 3). The structure of chitinase I showed (trans) glycosidases like α/β TIM-barrel as a characteristic domain of its structure with 75 (18.8 %), 109 (27.2 %) and 30 (7.5 %) amino acids are β-strand, α-helix and 3–10 helix, respectively (Fig. 3a). The structure of chitinase I revealed two β-sheets which are composed of nine mixed β-strands and five anti parallel β-strands. There are 22 α-helices with 16 helix–helix interactions as well as 4 β–α–β motifs present in the structure of chitinase I. Using the information present in the literature and the comparison of sequence with template, the active pocket of the chitinase II possesses Tyr36, Phe70, Asp174, Glu176, Gln208, Gln232, Tyr234 and Trp307. The Glu176 was predicted to be essential for the activity of the chitinase (Khan et al. 2015). The active pocket of the chitinase I contains Tyr10, Phe38, Asp135, Asp137, Glu139, Met202, Tyr204, Asp205, Tyr259 and Trp344 residues. The chitinase II contains 18.7 % β-strands (56 aa), 27.3 % α-helix (82 aa) and 0.9 % 3–10 helix (3 aa). The structure shows the presence of 2 β-sheets which comprises 8 parallel strands, 2 anti-parallel strands, 1 β-hairpin, 10 α-helices with 9 helix–helix interactions and 6 characteristic β–α–β motifs that may be responsible for the activity of enzyme (Fig. 3b). Moreover, there are two disulfide cysteine bridges between 58Cys–113Cys and

Author's personal copy Extremophiles Fig. 3 Three-dimensional structure of a chitinase I, b chitinase II, and c superimposed structure of both the enzymes. The predicted active site residues present in the pocket are shown in stick model

209Cys–238Cys. There is an extra possible chance of disulfide bond between 92Cys and 103Cys in the structure (Khan et al. 2015). The superimposed structures of both chitinases showed a marginal difference in the overall structure and fold (Fig. 3c). However, many remarkable differences are observed such as size and orientation of loops and secondary structures may be responsible for the differences in their stability.

Molecular dynamic simulations How the sequence and a protein folds into a well-defined three-dimensional native structure is a big question

(Rahaman et al. 2008; Zaidi et al. 2014). Apart from the sequence, environmental conditions play a great role in the folding and stability of a protein (Alam Khan et al. 2009, 2010; Rahaman et al. 2015). Molecular dynamic (MD) simulations of chitinase provided a detailed information on the fluctuations and conformational changes (Khan et al. 2015). MD simulations can track system behavior and understanding of protein folding (Ubaid-Ullah et al. 2014; Haque et al. 2015; Naiyer et al. 2015). These methods are used to investigate the structure, dynamics and thermodynamics of biological molecules (Anwer et al. 2015; Hoda et al. 2015; Naz et al. 2015). In explicit conditions, all atoms of a protein and surrounding water molecules move in a series of time steps. At each time step, the forces on

13


each atom, the atomic positions and the velocities are computed (Borhani and Shaw 2012). MD simulation of chitinase displayed an acceptable stability profile at a temperature of 300, 325 and 350 K while instability was observed at 375 K. A relatively stable nature of the protein was observed for all temperatures by average total energy values. Similarly, root mean square deviation (RMSD), root mean square fluctuations (RMSF), and radius of gyration (Rg) values were found to be low at temperatures ranging from 300 to 350 K (Khan et al. 2015). The RMSD is a fundamental property to establish whether the protein is stable and close to the experimental structure (Kuzmanic and Zagrovic 2010). Vibrations around the equilibrium are not random but depend on the local structure flexibility. The RMSF calculate the average fluctuations of all residues during the simulations. Moreover, the Rg is a parameter linked to the tertiary structural volume of a protein and has been applied to obtain insight into the stability of the protein in a biological system (Lobanov et al. 2008).

applied in degradation of chitin and production of SCPs (Dahiya et al. 2006).

Industrial usage of chitinases

Bio‑control

Chitinases have several industrial and agricultural applications. The ability of chitinases to degrade chitin makes them valuable for pollution treatment. In addition, chitinases play significant role in the fungal protoplast generation, single-cell protein production, preparation of bioactive chitooligosaccharides and in the degradation of shellfish waste. Besides, chitinases can be used in human health care, in fungal disease therapy and as additives in antifungal creams (Dahiya et al. 2006). In this section, we discuss a detail of industrial application of chitinases.

Biocontrol is an alternative method in modern agriculture for reducing the dependence on chemical fungicides that may cause environmental pollution and produce resistant strains. Since, chitin is the major cell wall component of plant pathogens like Botrytis cinerea, Rhizoctonia solani, Sclerotinia sclerotiorum or Fusarium sp.; therefore, chitinases can be used as biocontrol agents and they are considered to be crucial enzymes in mycoparasitism (HerreraEstrella and Chet 1999; Zhang et al. 2001).

Fungal protoplast generation

Conclusions and perspectives

Fungal protoplasts are used to study cell wall synthesis, enzyme synthesis and secretion or strain improvement (Dahiya et al. 2006). Since fungi contain chitin in their cell walls, chitinases can be used in fungal protoplast formation. Dahiya et al. (2006) produced chitinase in submerged and solid-state fermentation with Enterobacter sp. NRG4 which was effective in release of protoplasts from Pleurotus florida, Trichoderma reesei, Agaricus bisporus and A. niger (Dahiya et al. 2006).

Chitinases can be exploited for their use as food preservatives to increase the shelf life of the foods, like xylanase (Stephens et al. 2014) has been used in the process of pulpbleaching for reducing the chlorine chemicals (Gruber et al. 1998) due to their superior stability at high temperatures. Thermostable chitinase also exhibits very high specificity and efficiency even at extreme conditions and hence it may have significant industrial values. Various efforts such as site-directed mutagenesis and introduction of extra disulfide bonds have been made to increase the thermostability of chitinases (Gruber et al. 1998). The thermostable property of chitinases can also be improved by random mutagenesis and DNA-shuffling techniques. There is a possibility of using chitinases as anti-tumor drugs since chitohexaose and chitoheptaose have shown anti-tumor activity (Hamid et al. 2013).

Single‑cell protein production Single-cell proteins (SCPs) are proteins produced from the culture of a single-celled microorganism and can be used as food or feed supplement or substitute. Using chitinous wastes as substrates the chitinolytic enzymes can be

13

Production of chitooligosaccharides Chitooligosaccharides, N-acetyl glucosamines and glucosamines have a tremendous pharmaceutical potential and can be used in medicine. Chitohexose and chitoheptose, for example, have antitumor activity (Dahiya et al. 2006). Glucosamine is commonly used for osteoarthritis treatment, while N-acetyl glucosamine can be used as a nutritional supplement, where it is important for intestinal function. Chitooligosaccharides and N-acetyl glucosamine are either produced as a result of acid hydrolysis of chitin or chemical acetylation of glucosamine and its oligomers. Recently, production of N-acetyl glucosamine has been reported by enzymes derived from Serratia marcescens, Bacillus huringiensis subsp. pakistani, Aeromonas hydrophila H2330, Trichoderma viride, Acremonium cellulolyticus and Aeromonas sp. (Jung and Park 2014).


The present review offers insights into the structural and dynamical features of family 18 chitinases isolated from T. lanuginosus. In recent years, thermostable chitinases from thermophilic microorganisms have gained wide attention in industrial, medical, environmental and biotechnological applications due to their inherent stability at high temperatures and a wide range of pH optima. The review focuses on chitinases I and II isolated from T. lanuginosus due to their stability at high temperature. The predicted 3D model of chitinase I and II showed the characteristic (α/β)8 TIM-barrel conformations. MD simulation analyses suggested that these chitinases maintain their structure in extremes of environmental condition and hence may offer wide industrial uses. Acknowledgments MIH is thankful to the Indian Council of Medical Research, Government of India for financial assistance. We sincerely thank Jamia Millia Islamia and Durban University of Technology for providing high-speed computing facilities. Compliance with ethical standards Conflict of interest The authors have no substantial financial or commercial conflicts of interest with the current work or its publication.

References Alam Khan MK, Das U, Rahaman MH, Hassan MI, Srinivasan A, Singh TP, Ahmad F (2009) A single mutation induces molten globule formation and a drastic destabilization of wild-type cytochrome c at pH 6.0. J Biol Inorg Chem 14:751–760 Alam Khan MK, Rahaman MH, Hassan MI, Singh TP, MoosaviMovahedi AA, Ahmad F (2010) Conformational and thermodynamic characterization of the premolten globule state occurring during unfolding of the molten globule state of cytochrome c. J Biol Inorg Chem 15:1319–1329 Anwer K, Sonani R, Madamwar D, Singh P, Khan F, Bisetty K, Ahmad F, Hassan MI (2015) Role of N-terminal residues on folding and stability of C-phycoerythrin: simulation and ureainduced denaturation studies. J Biomol Struct Dyn 33:121–133 Arakane Y, Zhu Q, Matsumiya M, Muthukrishnan S, Kramer KJ (2003) Properties of catalytic, linker and chitin-binding domains of insect chitinase. Insect Biochem Mol Biol 33:631–648 Arima K, Liu W-H, Beppu T (1972) Studies on the lipase of thermophilic fungus Humicola lanuginosa. Agric Biol Chem 36:893–895 Arimori T, Kawamoto N, Shinya S, Okazaki N, Nakazawa M, Miyatake K, Fukamizo T, Ueda M, Tamada T (2013) Crystal structures of the catalytic domain of a novel glycohydrolase family 23 chitinase from Ralstonia sp. A-471 reveals a unique arrangement of the catalytic residues for inverting chitin hydrolysis. J Biol Chem 288:18696–18706 Aronson NN Jr, Halloran BA, Alexyev MF, Amable L, Madura JD, Pasupulati L, Worth C, Van Roey P (2003) Family 18 chitinaseoligosaccharide substrate interaction: subsite preference and anomer selectivity of Serratia marcescens chitinase A. Biochem J 376:87–95 Bharadwaj G, Maheshwari R (1999) A comparison of thermal characteristics and kinetic parameters of trehalases from a thermophilic and a mesophilic fungus. FEMS Microbiol Lett 181:187–193

Bierbaum S, Nickel R, Koch A, Lau S, Deichmann KA, Wahn U, Superti-Furga A, Heinzmann A (2005) Polymorphisms and haplotypes of acid mammalian chitinase are associated with bronchial asthma. Am J Respir Crit Care Med 172:1505–1509 Bleau G, Massicotte F, Merlen Y, Boisvert C (1999) Mammalian chitinase-like proteins. EXS 87:211–221 Bolar JP, Norelli JL, Wong KW, Hayes CK, Harman GE, Aldwinckle HS (2000) Expression of endochitinase from Trichoderma harzianum in transgenic apple increases resistance to apple scab and reduces vigor. Phytopathology 90:72–77 Boot RG, Blommaart EF, Swart E, Ghauharali-van der Vlugt K, Bijl N, Moe C, Place A, Aerts JM (2001) Identification of a novel acidic mammalian chitinase distinct from chitotriosidase. J Biol Chem 276:6770–6778 Borhani DW, Shaw DE (2012) The future of molecular dynamics simulations in drug discovery. J Comput Aided Mol Des 26:15–26 Bortone K, Monzingo AF, Ernst S, Robertus JD (2002) The structure of an allosamidin complex with the Coccidioides immitis chitinase defines a role for a second acid residue in substrateassisted mechanism. J Mol Biol 320:293–302 Chang WT, Chen YC, Jao CL (2007) Antifungal activity and enhancement of plant growth by Bacillus cereus grown on shellfish chitin wastes. Bioresour Technol 98:1224–1230 Chaudhari SS, Arakane Y, Specht CA, Moussian B, Kramer KJ, Muthukrishnan S, Beeman RW (2013) Retroactive maintains cuticle integrity by promoting the trafficking of Knickkopf into the procuticle of Tribolium castaneum. PLoS Genet 9:e1003268 Chaudhuri A, Maheshwari R (1996) A novel invertase from a thermophilic fungus Thermomyces lanuginosus: its requirement of thiol and protein for activation. Arch Biochem Biophys 327:98–106 Cottrell MT, Moore JA, Kirchman DL (1999) Chitinases from uncultured marine microorganisms. Appl Environ Microbiol 65:2553–2557 Dahiya N, Tewari R, Hoondal GS (2006) Biotechnological aspects of chitinolytic enzymes: a review. Appl Microbiol Biotechnol 71:773–782 de Beer TA, Berka K, Thornton JM, Laskowski RA (2014) PDBsum additions. Nucleic Acids Res 42:D292–D296 Di Giambattista R, Federici F, Petruccioli M, Fenice M (2001) The chitinolytic activity of Penicillium janthinellum P9: purification, partial characterization and potential applications. J Appl Microbiol 91:498–505 Duo-Chuan L (2006) Review of fungal chitinases. Mycopathologia 161:345–360 Fang Z, Xu L, Pan D, Jiao L, Liu Z, Yan Y (2014) Enhanced production of Thermomyces lanuginosus lipase in Pichia pastoris via genetic and fermentation strategies. J Ind Microbiol Biotechnol 41:1541–1551 Ferrandon S, Sterzenbach T, Mersha FB, Xu MQ (2003) A single surface tryptophan in the chitin-binding domain from Bacillus circulans chitinase A1 plays a pivotal role in binding chitin and can be modified to create an elutable affinity tag. Biochim Biophys Acta 1621:31–40 Fischer L, Scheckermann C, Wagner F (1995) Purification and characterization of a thermotolerant beta-galactosidase from Thermomyces lanuginosus. Appl Environ Microbiol 61:1497–1501 Flach J, Pilet PE, Jolles P (1992) What’s new in chitinase research? Experientia 48:701–716 Fusetti F, von Moeller H, Houston D, Rozeboom HJ, Dijkstra BW, Boot RG, Aerts JM, van Aalten DM (2002) Structure of human chitotriosidase. Implications for specific inhibitor design and function of mammalian chitinase-like lectins. J Biol Chem 277:25537–25544 Gijzen M, Kuflu K, Qutob D, Chernys JT (2001) A class I chitinase from soybean seed coat. J Exp Bot 52:2283–2289

13

Author's personal copy Extremophiles Gilkes NR, Henrissat B, Kilburn DG, Miller RC Jr, Warren RA (1991) Domains in microbial β-1, 4-glycanases: sequence conservation, function, and enzyme families. Microbiol Rev 55:303–315 Gramany V, Iqbal Khan F, Govender A, Bisetty K, Singh S, Permaul K (2015) Cloning, expression and molecular dynamics simulations of a xylosidase obtained from Thermomyces lanuginosus. J Biomol Struct Dyn. doi:10.1080/07391102.2015.1089186 Gruber K, Klintschar G, Hayn M, Schlacher A, Steiner W, Kratky C (1998) Thermophilic xylanase from Thermomyces lanuginosus: high-resolution X-ray structure and modeling studies. Biochemistry 37:13475–13485 Gruber S, Vaaje-Kolstad G, Matarese F, Lopez-Mondejar R, Kubicek CP, Seidl-Seiboth V (2011) Analysis of subgroup C of fungal chitinases containing chitin-binding and LysM modules in the mycoparasite Trichoderma atroviride. Glycobiology 21:122–133 Gulati H, Chadha B, Saini H (2007) Production, purification and characterization of thermostable phytase from thermophilic fungus Thermomyces lanuginosus TL-7. Acta Microbiol Immunol Hung 54:121–138 Haki GD, Rakshit SK (2003) Developments in industrially important thermostable enzymes: a review. Bioresour Technol 89:17–34 Hamid R, Khan MA, Ahmad M, Ahmad MM, Abdin MZ, Musarrat J, Javed S (2013) Chitinases: an update. J Pharm Bioallied Sci 5:21–29 Haque MA, Zaidi S, Ubaid-Ullah S, Prakash A, Hassan MI, Islam A, Batra JK, Ahmad F (2015) In vitro and in silico studies of urea-induced denaturation of yeast iso-1-cytochrome c and its deletants at pH 6.0 and 25 °C. J Biomol Struct Dyn 33:1493–1502 Hartl L, Zach S, Seidl-Seiboth V (2012) Fungal chitinases: diversity, mechanistic properties and biotechnological potential. Appl Microbiol Biotechnol 93:533–543 Hassan I (2015) Recent advances in the structure-based drug design (SBDD) and discovery. Curr Top Med Chem 16(9):1–2 Hassan I, Ahmad F (2011) Structural diversity of class I MHC-like molecules and its implications in binding specificities. Adv Protein Chem Struct Biol 83:223–270 Hassan MI, Kumar V, Singh TP, Yadav S (2007a) Structural model of human PSA: a target for prostate cancer therapy. Chem Biol Drug Des 70:261–267 Hassan MI, Kumar V, Somvanshi RK, Dey S, Singh TP, Yadav S (2007b) Structure-guided design of peptidic ligand for human prostate specific antigen. J Pept Sci 13:849–855 Hassan MI, Bilgrami S, Kumar V, Singh N, Yadav S, Kaur P, Singh TP (2008) Crystal structure of the novel complex formed between zinc α2-glycoprotein (ZAG) and prolactin-inducible protein (PIP) from human seminal plasma. J Mol Biol 384:663–672 Hassan MI, Aijaz A, Ahmad F (2010a) Structural and functional analysis of human prostatic acid phosphatase. Expert Rev Anticancer Ther 10:1055–1068 Hassan MI, Toor A, Ahmad F (2010b) Progastriscin: structure, function, and its role in tumor progression. J Mol Cell Biol 2:118–127 Hassan MI, Saxena A, Ahmad F (2012) Structure and function of von Willebrand factor. Blood Coagul Fibrinolysis 23:11–22 Hassan MI, Waheed A, Grubb JH, Klei HE, Korolev S, Sly WS (2013) High resolution crystal structure of human beta-glucuronidase reveals structural basis of lysosome targeting. PLoS ONE 8:e79687 Hayes CK, Klemsdal S, Lorito M, Di Pietro A, Peterbauer C, Nakas JP, Tronsmo A, Harman GE (1994) Isolation and sequence of an endochitinase-encoding gene from a cDNA library of Trichoderma harzianum. Gene 138:143–148 Henrissat B (1999) Classification of chitinases modules. EXS 87:137–156

13

Henrissat B, Bairoch A (1993) New families in the classification of glycosyl hydrolases based on amino acid sequence similarities. Biochem J 293(Pt 3):781–788 Henrissat B, Bairoch A (1996) Updating the sequence-based classification of glycosyl hydrolases. Biochem J 316(Pt 2):695–696 Herrera-Estrella A, Chet I (1999) Chitinases in biological control. Exs 87:171–184 Hoda N, Naz H, Jameel E, Shandilya A, Dey S, Hassan MI, Ahmad F, Jayaram B (2015) Curcumin specifically binds to the human calcium-calmodulin-dependent protein kinase IV: fluorescence and molecular dynamics simulation studies. J Biomol Struct Dyn. doi:10.1080/07391102.2015.1046934 Hoell IA, Dalhus B, Heggset EB, Aspmo SI, Eijsink VG (2006) Crystal structure and enzymatic properties of a bacterial family 19 chitinase reveal differences from plant enzymes. FEBS J 273:4889–4900 Hollak CE, van Weely S, van Oers MH, Aerts JM (1994) Marked elevation of plasma chitotriosidase activity. A novel hallmark of Gaucher disease. J Clin Invest 93:1288–1292 Itoh T, Hibi T, Fujii Y, Sugimoto I, Fujiwara A, Suzuki F, Iwasaki Y, Kim JK, Taketo A, Kimoto H (2013) Cooperative degradation of chitin by extracellular and cell surface-expressed chitinases from Paenibacillus sp. strain FPU-7. Appl Environ Microbiol 79:7482–7490 Jung WJ, Park RD (2014) Bioproduction of chitooligosaccharides: present and perspectives. Marine drugs 12:5328–5356 Khan FI, Govender A, Permaul K, Singh S, Bisetty K (2015) Thermostable chitinase II from Thermomyces lanuginosus SSBP: cloning, structure prediction and molecular dynamics simulations. J Theor Biol 374:107–114 Khoushab F, Yamabhai M (2010) Chitin research revisited. Marine drugs 8:1988–2012 Khucharoenphaisan K, Sinma K (2010) Beta-xylanase from Thermomyces lanuginosus and its biobleaching application. Pak J Biol Sci 13:513–526 Koga D, Sasaki Y, Uchiumi Y, Hirai N, Arakane Y, Nagamatsu Y (1997) Purification and characterization of Bombyx mori chitinases. Insect Biochem Mol Biol 27:757–767 Krause A, Stoye J, Vingron M (2000) The SYSTERS protein sequence cluster set. Nucleic Acids Res 28:270–272 Kuzmanic A, Zagrovic B (2010) Determination of ensemble-average pairwise root mean-square deviation from experimental B-factors. Biophys J 98:861–871 Lee P, Waalen J, Crain K, Smargon A, Beutler E (2007) Human chitotriosidase polymorphisms G354R and A442V associated with reduced enzyme activity. Blood Cells Mol Dis 39:353–360 Lee CG, Da Silva CA, Dela Cruz CS, Ahangari F, Ma B, Kang MJ, He CH, Takyar S, Elias JA (2011) Role of chitin and chitinase/ chitinase-like proteins in inflammation, tissue remodeling, and injury. Annu Rev Physiol 73:479–501 Li H, Greene LH (2010) Sequence and structural analysis of the chitinase insertion domain reveals two conserved motifs involved in chitin-binding. PLoS ONE 5:e8654 Li D-C, Yang Y-J, Shen C-Y (1997) Protease production by the thermophilic fungus Thermomyces lanuginosus. Mycol Res 101:18–22 Liu T, Zhou Y, Chen L, Chen W, Liu L, Shen X, Zhang W, Zhang J, Yang Q (2012) Structural insights into cellulolytic and chitinolytic enzymes revealing crucial residues of insect β-N-acetyl-dhexosaminidase. PLoS ONE 7:e52225 Lobanov M, Bogatyreva NS, Galzitskaia OV (2008) Radius of gyration is indicator of compactness of protein structure. Mol Biol 42:701–706 Lorito M, Hayes CK, Di Pietro A, Harman GE (1993) Biolistic transformation of Trichoderma harzianum and Gliocladium virens using plasmid and genomic DNA. Curr Genet 24:349–356

Author's personal copy Extremophiles Lorito M, Woo SL, Garcia I, Colucci G, Harman GE, Pintor-Toro JA, Filippone E, Muccifora S, Lawrence CB, Zoina A, Tuzun S, Scala F (1998) Genes from mycoparasitic fungi as a source for improving plant resistance to fungal pathogens. Proc Natl Acad Sci USA 95:7860–7865 Mayer RT, McCollum TG, Niedz RP, Hearn CJ, McDonald RE, Berdis E, Doostdar H (1996) Characterization of seven basic endochitinases isolated from cell cultures of Citrus sinensis (L.). Planta 200:289–295 Meng Zhang AKP, Algasan G, Zhengxiang W, Suren S, Kugenthiren P (2015) The multi-chitinolytic enzyme system of the compostdwelling thermophilic fungus Thermomyces lanuginosus. Process Biochem 50(2):237–244 Merzendorfer H, Zimoch L (2003) Chitin metabolism in insects: structure, function and regulation of chitin synthases and chitinases. J Exp Biol 206:4393–4412 Muzzarelli RA (1999) Native, industrial and fossil chitins. EXS 87:1–6 Naiyer A, Hassan MI, Islam A, Sundd M, Ahmad F (2015) Structural characterization of MG and pre-MG states of proteins by MD simulations, NMR, and other techniques. J Biomol Struct Dyn 33:2267–2284 Naz F, Singh P, Islam A, Ahmad F, Imtaiyaz Hassan M (2015) Human microtubule affinity-regulating kinase 4 is stable at extremes of pH. J Biomol Struct Dyn. doi:10.1080/07391102.2015.1074942 Neuhaus JM, Sticher L, Meins F Jr, Boller T (1991) A short C-terminal sequence is necessary and sufficient for the targeting of chitinases to the plant vacuole. Proc Natl Acad Sci USA 88:10362–10366 Ober C, Chupp GL (2009) The chitinase and chitinase-like proteins: a review of genetic and functional studies in asthma and immunemediated diseases. Curr Opin Allergy Clin Immunol 9:401–408 Odibo FJC, Ulbrich-Hofmann R (2001) Thermostable α-amylase and glucoamylase from Thermomyces lanuginosus F1. Acta Biotechnol 21:141–153 Patil RS, Ghormade VV, Deshpande MV (2000) Chitinolytic enzymes: an exploration. Enzyme Microb Technol 26:473–483 Punja ZK, Zhang YY (1993) Plant chitinases and their roles in resistance to fungal diseases. J Nematol 25:526–540 Quevillon E, Silventoinen V, Pillai S, Harte N, Mulder N, Apweiler R, Lopez R (2005) InterProScan: protein domains identifier. Nucleic Acids Res 33:W116–W120 Rahaman H, Khan KA, Hassan I, Wahid M, Singh SB, Singh TP, Moosavi-Movahedi AA, Ahmad F (2008) Sequence and stability of the goat cytochrome c. Biophys Chem 138:23–28 Rahaman H, Alam Khan MK, Hassan MI, Islam A, Moosavi-Movahedi AA, Ahmad F (2015) Heterogeneity of equilibrium molten globule state of cytochrome c induced by weak salt denaturants under physiological condition. PLoS ONE 10:e0120465 Rezessy-Szabo JM, Nguyen QD, Hoschke A, Braet C, Hajos G, Claeyssens M (2007) A novel thermostable α-galactosidase from the thermophilic fungus Thermomyces lanuginosus CBS 395.62/b: purification and characterization. Biochim Biophys Acta 1770:55–62 Salzer P, Feddermann N, Wiemken A, Boller T, Staehelin C (2004) Sinorhizobium meliloti-induced chitinase gene expression in Medicago truncatula ecotype R108-1: a comparison between symbiosis-specific class V and defence-related class IV chitinases. Planta 219:626–638 Sandor E, Pusztahelyi T, Karaffa L, Karanyi Z, Pocsi I, Biro S, Szentirmai A, Pocsi I (1998) Allosamidin inhibits the fragmentation of Acremonium chrysogenum but does not influence the cephalosporin-C production of the fungus. FEMS Microbiol Lett 164:231–236 Santos IS, Oliveira AE, Da Cunha M, Machado OL, Neves-Ferreira AG, Fernandes KV, Carvalho AO, Perales J, Gomes VM (2007)

Expression of chitinase in Adenanthera pavonina seedlings. Physiol Plant 131:80–88 Sasaki C, Varum KM, Itoh Y, Tamoi M, Fukamizo T (2006) Rice chitinases: sugar recognition specificities of the individual subsites. Glycobiology 16:1242–1250 Sasson O, Vaaknin A, Fleischer H, Portugaly E, Bilu Y, Linial N, Linial M (2003) ProtoNet: hierarchical classification of the protein space. Nucleic Acids Res 31:348–352 Scholte EJ, Knols BG, Samson RA, Takken W (2004) Entomopathogenic fungi for mosquito control: a review. J Insect Sci 4:19 Seidl V, Huemer B, Seiboth B, Kubicek CP (2005) A complete survey of Trichoderma chitinases reveals three distinct subgroups of family 18 chitinases. FEBS J 272:5923–5939 Selvaggini S, Munro CA, Paschoud S, Sanglard D, Gow NA (2004) Independent regulation of chitin synthase and chitinase activity in Candida albicans and Saccharomyces cerevisiae. Microbiology 150:921–928 Shimono K, Matsuda H, Kawamukai M (2002) Functional expression of chitinase and chitosanase, and their effects on morphologies in the yeast Schizosaccharomyces pombe. Biosci Biotechnol Biochem 66:1143–1147 Singh S, Pillay B, Dilsook V, Prior BA (2000a) Production and properties of hemicellulases by a Thermomyces lanuginosus strain. J Appl Microbiol 88:975–982 Singh S, Reddy P, Haarhoff J, Biely P, Janse B, Pillay B, Pillay D, Prior BA (2000b) Relatedness of Thermomyces lanuginosus strains producing a thermostable xylanase. J Biotechnol 81:119–128 Singh S, Madlala AM, Prior BA (2003) Thermomyces lanuginosus: properties of strains and their hemicellulases. FEMS Microbiol Rev 27:3–16 Stephens DE, Khan FI, Singh P, Bisetty K, Singh S, Permaul K (2014) Creation of thermostable and alkaline stable xylanase variants by DNA shuffling. J Biotechnol 187:139–146 Sutherland LA, Wong WM, Nazar RN (1989) Sequence and putative regulatory elements in a 5S rRNA gene from a eukaryotic thermophile, Thermomyces lanuginosus. Nucleic Acids Res 17:10504 Takaya N, Yamazaki D, Horiuchi H, Ohta A, Takagi M (1998a) Cloning and characterization of a chitinase-encoding gene (chiA) from Aspergillus nidulans, disruption of which decreases germination frequency and hyphal growth. Biosci Biotechnol Biochem 62:60–65 Takaya N, Yamazaki D, Horiuchi H, Ohta A, Takagi M (1998b) Intracellular chitinase gene from Rhizopus oligosporus: molecular cloning and characterization. Microbiology 144(Pt 9):2647–2654 Terwisscha van Scheltinga AC, Hennig M, Dijkstra BW (1996) The 1.8 A resolution structure of hevamine, a plant chitinase/ lysozyme, and analysis of the conserved sequence and structure motifs of glycosyl hydrolase family 18. J Mol Biol 262:243–257 Tjoelker LW, Gosting L, Frey S, Hunter CL, Trong HL, Steiner B, Brammer H, Gray PW (2000) Structural and functional definition of the human chitinase chitin-binding domain. J Biol Chem 275:514–520 Ubaid-Ullah S, Haque MA, Zaidi S, Hassan MI, Islam A, Batra JK, Singh TP, Ahmad F (2014) Effect of sequential deletion of extra N-terminal residues on the structure and stability of yeast iso1-cytochrome-c. J Biomol Struct Dyn 32:2005–2016 Ubhayasekera W, Karlsson M (2012) Bacterial and fungal chitinase chiJ orthologs evolve under different selective constraints following horizontal gene transfer. BMC Res Notes 5:581 van Aalten DM, Synstad B, Brurberg MB, Hough E, Riise BW, Eijsink VG, Wierenga RK (2000) Structure of a two-domain chitotriosidase from Serratia marcescens at 1.9-A resolution. Proc Natl Acad Sci USA 97:5842–5847

13

Author's personal copy Extremophiles Van Der Spoel D, Lindahl E, Hess B, Groenhof G, Mark AE, Berendsen HJ (2005) GROMACS: fast, flexible, and free. J Comput Chem 26:1701–1718 Watanabe T, Kanai R, Kawase T, Tanabe T, Mitsutomi M, Sakuda S, Miyashita K (1999) Family 19 chitinases of Streptomyces species: characterization and distribution. Microbiology 145(Pt 12):3353–3363 Yang J, Yu Y, Li J, Zhu W, Geng Z, Jiang D, Wang Y, Zhang KQ (2013) Characterization and functional analyses of the chitinase-encoding genes in the nematode-trapping fungus Arthrobotrys oligospora. Arch Microbiol 195:453–462 Zaidi S, Hassan MI, Islam A, Ahmad F (2014) The role of key residues in structure, function, and stability of cytochrome-c. Cell Mol Life Sci 71:229–255

13

Zhang Z, Yuen GY, Sarath G, Penheiter AR (2001) Chitinases from the plant disease biocontrol agent, Stenotrophomonas maltophilia C3. Phytopathology 91:204–211 Zhang H, Huang X, Fukamizo T, Muthukrishnan S, Kramer KJ (2002) Site-directed mutagenesis and functional analysis of an active site tryptophan of insect chitinase. Insect Biochem Mol Biol 32:1477–1488 Zhang H, Liu M, Tian Y, Hu X (2011) Comparative characterization of chitinases from silkworm (Bombyx mori) and bollworm (Helicoverpa armigera). Cell Biochem Biophys 61:267–275 Zhu Z, Zheng T, Homer RJ, Kim YK, Chen NY, Cohn L, Hamid Q, Elias JA (2004) Acidic mammalian chitinase in asthmatic Th2 inflammation and IL-13 pathway activation. Science 304:1678–1682