Chitin Degradation and Modification
Insects must periodically replace their old cuticle with a new one because it is too rigid to allow for growth. Key to this process is the elaboration of the molting fluid with an assortment of chtitinases and proteases. Chitinases are among a group of proteins that insects use to digest the structural polysaccharide in their exoskeletons and gut linings during the molting process (Kramer et al., 1985; Kramer and Koga, 1986; Kramer and Muthukrish-nan, 1997; Fukamizo, 2000). Precise regulation of chi-tin metabolism is a complex and intricate process that is critical for insect growth, metamorphosis, organogenesis, and survival (Arakane and Muthukrishnan, 2010). Chitin content, which fluctuates throughout the life cycle of the insect, is directly influenced not only by chitin synthases (CHSs), but also by chitinases (CHTs, EC 3.2.1.14) and P-A-acetylglucosaminidases (NAGs, EC 3.2.1.52).
Figure 5 Phenotypes of T. castaneum larvae after RNAi for genes of chitin metabolism. dsRNAs for the indicated genes (200 ng per insect, n = 20) were injected into penultimate instar larvae (young larvae), last instar larvae, pharate pupae as indicated above each panel. All animals injected with dsRNA for CHS-A, TcCHT10, TcNAG1, and TcCDA1 died at the ensuing molt. Unlike RNAi of TcCHT10, injection of dsRNA (200 ng per insect) for TcCHT5 into penultimate instar and last instar larvae as well as pharate pupae prevented only adult molt. When dsRNA for TcCHT7 (200 ng per insect) was injected into pharate pupae, normal phenotypes were observed in the pupal stage. However, unlike buffer-injected controls, TcCHT7 dsRNA-treated insects failed to expand their adult elytra and their wings did not fold properly (modified from Zhu et al., 2008c). Animals injected with control dsRNA for EGFP developed in a normal fashion, and had no mortality or abnormal phenotype.
Chitin is digested in the cuticle and PM to GlcNAc by a binary enzyme system composed of CHT and NAG (Fukamizo and Kramer, 1985a, 1985b; Filho et al., 2002). The former enzyme from molting fluid hydrolyzes chitin into oligosaccharides, whereas the latter, which is also found in the molting fluid, further degrades the oligomers to the monomer from the non-reducing end. In some cases, additional unrelated proteins that possess one or more chitin-binding domains (CBD), but are devoid of chi-tinolytic activity, enhance degradation of chitin (Vaaje-Kolstad et al., 2005). This system also probably operates in the gut during degradation of PM, and increases the porosity of the PM. It may also help in the digestion of chitin-containing prey (Bolognesi et al., 2005; Khajuria et al., 2010).
The precise control of chitin content is critical not only for the survival of the insect, but also for optimal function of individual anatomical structures such as wings and other appendages. In addition, modulation of the physical properties of chitin-containing structures of insects is accomplished, in part, by the deacetylation of the polysaccharide by chitin deacetylases (CDAs, EC 3.5.1.41). Partially deacetylated chitin may have different protein-binding and physical properties than those of chitin. The process of partially deacetylating chitin and the importance of this modification for insect growth and development have emerged as new areas of research in insect molecular science (Luschnig et al., 2006; Wang et al., 2006; Arakane et al, 2009).
Insect Chitinases
Cloning of genes encoding insect chitinases and chitinase-like proteins Since the first report of an insect chitinase, its cDNA and its corresponding gene from M. sexta (MsCHT5) (Koga et al, 1987; Kramer et al., 1993; Choi et al., 1997; Kramer and Muthukrishnan, 1997), numerous insect CHT genes and cDNAs have been cloned and characterized from several insect species belonging to different orders, including dipterans, lepidopterans, coleopterans, hemipterans, and hymenopterans (Kramer and Muthukrishnan, 2005). The organization of most of these genes is very similar to that of MsCHT5, and most of the proteins display a domain architecture consisting of catalytic, linker, and/or chitin-binding domains (CBD) similar to MsCHT5. These genes/ enzymes include epidermal chitinases from the silkworm Bombyx mori (Kim et al., 1998; Abdel-Banat and Koga, 2001), the fall webworm Hyphantria cunea (Kim et al., 1998), wasp venom from Chelonus sp. (Krishnan et al., 1994), the common cutworm Spodoptera litura (Shinoda et al. , 2001), the fall armyworm Spodoptera frugiperda (Bolognesi et al., 2005), a molt-associated chitinase from the spruce budworm Choristoneura fumiferana (Zheng et al., 2002), and midgut-associated chitinases from the malaria mosquito A. gambiae (Shen and Jacobs-Lorena, 1997), yellow fever mosquito Ae. aegypti (de la Vega et al., 1998; Khajuria et al., 2010), the beetle Phaedon cochleariae (Girard and Jouanin, 1999), and the sand fly Lutzomyia longipalpis (Ramalho-Ortigao and Traub-Cseko, 2003), as well as several deduced from Drosophila genome data. A smaller linkerless fat body-specific chitinase from the tsetse fly Glossina morsitans (Yan et al., 2002), and a very large epidermal chitinase with five copies of the catalytic domain and multiple chitin-binding domain from the yellow mealworm Tenebrio molitor (Royer et al., 2002), have also been described.
Daimon et al. (2003) described a gene encoding another type of chitinase from the silkworm, BmCHT-h. The encoded chitinase shared extensive similarities with microbial and baculoviral chitinases (73% amino acid sequence identity to Serratia marcescens chitinase, and 63% identity to Autographa californica nuclear polyhe-drosis virus chitinase). Even though this enzyme had the signature sequence characteristic of a family 18 chitinase, it had a rather low percentage of sequence identity with the family of insect chitinases. It was suggested that an ancestral species of B. mori acquired this chitinase gene via horizontal gene transfer from Serratia or a baculovirus. A gene encoding a CHT-like protein that is highly related to BmCHT-h was also found in the pea aphid, Acyrthosiphon pisum (Nakabachi et al., 2010).
Only after the complete genome sequences became available was it recognized that insect genomes contain a large number of genes encoding CHT-like proteins widely divergent not only in their DNA and amino acid sequences, but also in the organization of their domains (Zhu et al, 2004, 2008a; Arakane and Muthukrishnan, 2010). The number of CHT genes per insect genome is in the range of 7 to 24 for D. melanogaster, A. gambiae, Ae. aegypti, B. mori, A. pisum, and T. castaneum. This range excludes genes encoding CHT-like proteins whose consensus sequences are poorly conserved (see section 7.1.2.2; Khajuria et al, 2010; Nakabachi et al, 2010; Zhu et al., 2004, 2008a). ‘The 22 genes that encode CHTs or chitinase-like proteins (CHLPs) in T. castaneum have been divided into eight subgroups, based on sequence similarity and domain organization (Figure 6) (Arakane and Muthukrishnan, 2010).
Figure 6 Domain organization of T. castaneum chitinase gene family. The program SMART was used to analyze the identified domains. TcCHT7 and TcCHT11 have a single transmembrane span at the N-terminal region. Blue boxes, signal peptide; pink boxes, catalytic domain; green boxes, chitin binding domain; red boxes, transmembrane span; lines, linker regions.
The chitinases in all insect species can be similarly classified into multiple groups (Figure 7). There is only one copy of the gene encoding a group I chitinase (CHT5) in all species except for A. gambiae, Ae. aegypti, and the human body louse Pedicu-lus humanus corporis, in which obvious gene duplications have occurred, resulting in one to four additional copies (Khajuria et al., 2010). To date, only one gene representing each of the groups II, III, VI, VII, and VIII (CHT10, 7, 6, and 11, respectively) has been found in various insect species. Interestingly, in addition to the group III CHT genes (CHT7s with two catalytic domains) identified in fully sequenced insect genomes such as T. castaneum, D. melanogaster, A. gambiae, Ae. aegypti, C. pipiens, A. mel-lifera, N. vitripennis, A. pisum, and P. corporis, orthologs have also been found in non-insect arthropod genomes, including those of the crustacean water flea Daphnia pulex, and the arachnid deer tick Ixodes scapularis, indicating an ancient origin of CHT7 that predates separation of the class Chelicerata more than five million years ago. Group IV appears to be the largest group in all insect species studied, containing 5, 8, 10, and 14 genes in D. melanogaster, A. gambiae, Ae. aegypti, and T. castaneum, respectively. The sole exception so far is A. pisum. No chitinase gene encoding a protein that belongs to this group was identified in A pisum (Nakabachi et al., 2010). In T. castaneum, most group IV CHT genes form a large cluster within a small region of the genome, suggesting the occurrence of a recent gene duplication event. Group V is composed of the genes encoding CHLPs such as imaginal disc growth factors (IDGFs). The number of genes for this group ranges from one in B. mori and A. pisum to as many as six in D. melanogaster (Arakane and Muthukrishnan, 2010; Nakabachi et al, 2010).
Domain organization of insect chitinases Insect CHTs belong to family-18 glycosylhydro-lases (the GH-18 super family) and function in hydrolysis of chitin in the exoskeleton and PM-associated chitin in the midgut, utilizing an endo-type cleavage mechanism during the molting process (Kramer and Muthukrishnan, 1997, 2005). Members of the CHT family contain a multidomain structural organization that includes a leader peptide and/or a transmembrane span, one to five catalytic domains (GH-18), multiple Ser/Thr-rich linker regions that are usually heavily glycosylated, and zero to seven six-cysteine-containing chitin-binding domains (CBDs) related to the peritrophin A domain (Figure 6; Royer et al., 2002; Arakane et al., 2003; Zhu et al., 2008b). The catalytic domains of all insect CHTs, which are comprised of about 370 amino acids, assume a fi8a8-barrel structure and possess signature motifs of family 18 glycosylhydrolases (Kramer et al., 1993; Perrakis et al., 1994, Terwisscha van Scheltinga et al., 1994; de la Vega et al, 1998; Fusetti et al., 2002; Varela et al, 2002; Tsai et al., 2004; Arakane and Muthukrishnan, 2010). The consensus sequence for conserved motif I is KXX(V/L/I)A(V/L)GGW in the ^3-strand, where X is a non-conserved amino acid. The conserved motif II is FDG(L/F)DLDWE(Y/F)P, which is known to be located in or near the catalytic site (^4-strand) of the enzyme, with a glutamate residue (E) being the most critical residue in this motif as the putative proton donor in the catalytic mechanism (Watanabe et al., 1993; Lu et al., 2002; Zhang et al., 2002). Conserved motifs III and IV are MXYDL(R/H)G in the ^6-strand and GAM(T/V) WA(I/L)DMDD in the ^8-strand.
CBDs found in insect CHTs all belong to carbohydrate-binding module 14 (CBM-14, pfam 01607; ChtBD2 family = SMART family 00494, Boraston et al., 2004). Insect CBDs are only about 60 amino acids long and have less conserved amino acid sequences, with the exception of the six cysteines and several aromatic residues whose relative locations are highly conserved (Jasrapuria et al., 2010). The proposed function(s) of the CBD is to help anchor the enzyme onto the insoluble chitin to enhance chitin degradation efficiency (Linder et al., 1996; Arakane et al., 2003). As described in section 7.4.1.1, based on the amino acid sequence similarity and domain architecture, insect CHTs can be classified into eight groups (Figures 6 and 7). Group I CHTs (CHT5s) represent the prototypical and enzymatically characterized CHTs purified from molting fluid and/or integument of M. sexta and B. mori (Koga et al., 1983, 1997). All of these group members contain a signal peptide, one catalytic domain, a Ser/Thr-rich linker region, and one CBD. Group II CHTs (CHT10s) are rather diverse in their domain architecture, and have four or five catalytic domains, together with four to seven CBDs. Dipterans and A. pisum (hemiptera) appear to be unique in having only four catalytic domains and four CBDs. The domain corresponding to the most N-terminal catalytic domain and one CBD found in group II chitinases from other species appear to be missing in the dipteran CHT10s (Zhu et al., 2008b; Arakane and Muthukrishnan, 2010; Nakabachi et al, 2010). The second catalytic unit of all CHT10s (the first catalytic unit in the case of the dipteran and A. pisum proteins) is predicted to lack chitinolytic activity due to a substitution of the most critical amino acid residue glutamate (E) with asparagine (N) in conserved motif II. Group III CHTs (CHT7s) possess two catalytic domains and one C-termi-nal CBD. The first catalytic domains of the group III proteins from all insect species studied share greater sequence similarity with each other than they do to the second catalytic domain, suggesting a unique function and/or evolutionary origin for each of the catalytic domains. Unlike most insect CHTs, CHT7s are predicted to have an N-terminal transmembrane segment, and are likely to be membrane-bound proteins. Indeed, recombinant T. casta-neum CHT7 (TcCHT7) that was expressed in Hi-5 insect cells using the baculovirus protein expression system was found to be in the cell pellet rather than in the medium, as expected for secreted proteins.
Figure 7 Phylogenetic analysis of putative chitinases and chitinase-like proteins (IDGFs) in insects. ClustalW software was used to perform multiple sequence alignments prior to phylogenetic analysis. The phylogenetic tree was constructed by MEGA 4.0 software using UPGMA (Tamura et al., 2007). Protein sequences obtained from GenBank as follows: Tribolium castaneum, TcCHT2 (AY873913); TcCHT4 (EF125543); TcCHT5 (AY675073); TcCHT6 (AY873916); TcCHT7 (DQ659247); TcCHT8 (DQ659248); TcCHT9 (DQ659249); TcCHT10 (DQ659250); TcCHT11 (DQ659251);TcCHT12 (XM_967709); TcCHT13 (DQ659252); TcCHT14 (XM_967912); TcCHT15 (XM_967984); TcCHT16 (AY873915); TcCHT17 (XP_972719); TcCHT18 (XP_973161); TcCHT19 (XP_973119); TcCHT20 (NP_001034516); TcCHT21 (NP_001034517); TcCHT22 (NP_001038095); TcIDGF2 (DQ659253); TcIDGF4 (DQ659254); Aedes aegypti, AaCHT1 (XP_001656232); AaCHT2 (XP_001662520); AaCHT3 (XP_001663568); AaCHT4 (XP_001663099); AaCHT5 (XP_001656234); AaCHT6 (XP_001662588); AaCHT7 (XP_001650020); AaCHT8 (XP_001663098); AaCHT9 (XP_001663099); AaCHT10 (XP_001655973); AaCHT11 (XP_001654045); AaCHT12 (XP_001658836); AaCHT13 (XP_001656231); AaCHT14 (XP_001656233); AaBR1 (XP_001660745); AaBR2 (XP_001660748); Apis mellifera, AmCHT2 (XP_623744); AmCHT5 (XP_623995); AmCHT6 (XP_393252); AmCHT7 (XP_396925); AmCHT10 (XP_395734); AmCHT11 (XP_395707); AmIDGF (XP_396769); Drosophila melanogaster, DmCHT2 (NP_477298); DmCHT5 (NP_650314); DmCHT6 (NP_572598); DmCHT7 (NP_647768); DmCHT10 (NP_001036422); DmCHT11 (NP_572361); DmIDGF1 (NP_477258); DmIDGF2 (NP_477257); DmIDGF3 (NP_723967); DmIDGF4 (NP_727374); DmIDGF5 (NP_611321); DmDS47 (NM_057733); Bombyxmori, BmCHT2 (BGIBMGA009695); BmCHT5 (BGIBMGA010240); BmCHT6 (BGIBMGA009890); BmCHT7 (BGIBMGA005539); BmCHT10 (BGIBMGA006874); BmCHT11 (BGIBMGA005859); BmIDGF (BGIBMGA000648); Anopheles gambiae, AgCHT2 (XP_315650); AgCHT5 (XP_001237469); AgCHT6 (AGAP000198); AgCHT7 (XP_308858); AgCHT10 (XP_001238192); AgCHT11 (XP_310662); AgBR1 (AAS80137); AgBR2 (AY496421); Nasonia vitripennis, NvCHT2 (XP_001601416); NvCHT5 (NP_001155084); NvCHT6 (); NvCHT7 (XP_001604515); NvCHT10 (XR_036825); NvCHT11 (XP_001604954); NvIDGF (XP_001599305.
The washed cell pellet containing recombinant TcCHT7 could hydrolyze chitin added to the culture medium, suggesting that the catalytic domains of this putative membrane-bound protein face the extracellular space (Arakane, unpublished data). Group IV CHTs comprise the largest and most divergent group of proteins. CHTs in this group have a signal pep-tide and one catalytic domain. Most (but not all) of the members lack a CBD (Figure 6). Group V chitinase-like proteins (CHLPs) include the imaginal disc growth factors (IDGFs) and the hemocyte aggregation inhibitor protein (HAIP, Kanost et al., 1994; Pan et al, 2010). CHLPs have a signal peptide, one catalytic domain, and no CBDs. Like other family-18 proteins, the crystal structure of D. mela-nogaster IDGF2 and homology modeling of all proteins in this group revealed the j8a8-TIM barrel structure (Varela et al., 2002). However, members of this group have an additional loop sequence located between the j4-strand and the a4-helix immediately after conserved region II. Although these proteins possess all four of the family-18 conserved motifs, the glutamate residue in conserved motif II is substituted by a glutamine in all members of the group, with the exception of two T. castaneum IDGFs (TcIDGF2 and TcIDGF4; Zhu et al, 2008b). TcIDGF2 and TcIDGF4 retain the glutamate residue in conserved region II but lack chitinase activity, either due to a D to A substitution in the conserved motif II, or to an extra loop stretching between the j4-strand and the a4-helix that possibly interferes with a productive substrate-enzyme interaction (Zhu et al., 2008a), or both. Group VI CHTs (CHT6s) exhibit a domain architecture similar to that of group I (a signal peptide, one catalytic domain, and one CBD), but they have a very long C-terminal stretch (e.g., 1819 amino acids in length after the CBD in TcCHT6) that has no predicted conserved domain (Figure 6) except for the A. pisum enzyme, which possesses an additional CBD at the C-terminal region (Nakabachi et al., 2010). Group VII CHTs (CHT2s) possess a domain architecture similar to that of group IV CHTs, which have a signal peptide, one catalytic domain, and no CBDs. They are classified as a separate group because phylogenetic analysis clearly indicates that these CHTs form a different clade near group II CHT10s. Group VIII CHTs (CHT11s) have one catalytic domain and no CBD. Interestingly, they have a predicted transmembrane segment instead of a signal peptide at the N-terminus, and they fall into a branch next to group III (CHT7s), all of which are predicted to be membrane-bound proteins.
Gene expression and functions of insect chitinases The redundancy of genes for CHTs raises important questions about their functions. Several insect CHT cDNAs have been obtained from epidermis, gut, and fat body, and extensively characterized (Kramer and Muthukrishnan, 2005). The epidermal endochitinases presumably function in turnover of the old cuticle, as these enzymes are found in the molting fluid along with A-acetylglucosaminidases, whereas the gut CHTs are thought to participate in the breakdown of chitin in the PM. In T. castaneum, tissue specificity and developmental patterns of expression of the 22 TcCHT and TcCHLP genes were analyzed by RT-PCR using cDNAs prepared from RNAs isolated at different developmental stages, such as embryo, larva, pharate pupa, pupa, and adult (Zhu et al., 2008c; Arakane and Muthukrishnan, 2010). The group I gene TcCHT5, group II gene TcCHT10, group III gene TcCHT7, group V genes TcIDGF2 and TcIDGF4, group VI gene TcCHT6, group VII gene TcCHT2, and group VIII gene TcCHTll are expressed at all stages analyzed, with some variation, whereas all group IV genes (TcCHTs 2, 4, 8, 9, and 12 to 22) were predominantly expressed in the feeding stages (larva and adult). In addition, all chitinase genes belonging to group IV were expressed in larval gut tissue but not in the carcass (whole body minus gut), suggesting a possible function of these TcCHTs in PM-associated chitin turnover or digestion of dietary chitin (Zhu et al., 2008c). Khajuria et al. (2010) recently reported that orally feeding dsRNA for a midgut-specific chitinase gene (encoding a group IV CHT) from larvae of O. nubilalis (OnCHT) significantly reduced the transcript levels of this gene and led to a significant increase of chitin content in the PM. The body weight of dsRNA OnCHT-fed larvae was decreased by 54% as compared with that of control dsRNA GFP-fed larvae, suggesting that some group IV CHTs are critical for regulating PM-chitin content, insect growth, and development. Interestingly, A. pisum appears to have no group IV CHT genes (Nakabachi et al., 2010). A. pisum (hemipteran) possesses a perimicrovillar membrane (PMM) that is devoid of chitin, suggesting that group IV CHTs may not play a role in the PM turnover. Instead, one CHT gene, ApCHT6 (encoding a group VIII CHT), was highly expressed in the midgut of A. pisum. Similarly, TcCHTll (encoding a group VIII CHT) was expressed in larval midgut, but not in the carcass (Arakane and Muthukrishnan, 2010). Group VIII CHTs, as well as group VI CHTs, may play critical roles in PM/PMM chitin degradation and turnover.
RNAi for group IV chitinases in T. castaneum for individual chitinases (and some combinations of chitinases) failed to produce any visible phenotypes, perhaps reflecting the redundant functions of this large group of chitinolytic enzymes. In contrast, injection of dsRNA for all chitinases belonging to groups I, II, III, and V resulted in unique lethal phenotypes. The most severe molting defect was observed after injection of dsRNA for TcCHTlO (encoding a group II CHT). Injections of dsRNA for TcCHTlO prevented the embryo from hatching and also averted all types of molts, including larval-larval, larval-pupal, and pupal-adult, depending on the timing of administration of the dsRNA (Figure 5; Zhu et al, 2008c). These results suggest a critical role for group II CHTs at every molt and developmental stage. Other CHTs (e.g., CHT5, also expressed in the epidermis) could not compensate for the loss of function of a group II CHT.
Unlike RNAi for TcCHT10, injection of dsTcCHT5 (encoding a group I CHT) prevented only the pupal— adult molt (Figure 5). Although the gene encoding this prototypical CHT was expressed throughout all developmental stages, and the corresponding enzymes from several other insect species have been found in larval molting fluid, the failure to obtain a larval-larval or larval-pupal molting arrest probably indicates that one or more of the other CHTs (e.g., group II CHT, TcCHT10) could compensate for TcCHT5 at all molts except during adult eclosion. Group III CHTs, which appear to encode membrane-bound enzymes with two catalytic domains and one CBD at the C-terminus, appear to be critical for tissue differentiation, rather than chitin degradation associated with molting. Indeed, in D. melanogaster, expression of the DmCHT7 (CG1869) gene increased more than 40-fold in the wing during the 32- to 40-h pupal wing differentiation period (Ren et al., 2005). In T. castaneum, injection of dsRNA for CHT7 resulted in a defective ely-tral and hindwing expansion without affecting molting (Figure 5; Zhu et al., 2008c). Group V is composed of IDGFs that are known to be involved in cell proliferation and differentiation (Kawamura et al., 1999; Zhang et al., 2006). It is worthy of note that although group V CHTs have no chitinolytic activity (Zhu et al., 2008b), they appear to be important for the adult molt. Injection of dsRNA for one of these CHLPs in T. castaneum, TcIDGF4, prevented adult eclosion (Zhu et al., 2008c). It is possible that TcIDGF4 may be required for tracheal proliferation during adult metamorphosis. Two A. gam-biae proteins, AgBR1 and AgBR2, which belong to this group, were induced specifically in the hemolymph by bacterial challenge (Shi and Paskewitz, 2004), suggesting that some members of the CHLP group (and/or members of other CHT groups) may have a role in the immune response.
