Introduction
Understanding the dynamics of insect–plant interactions provides insights into diverse aspects of relationships between plant-feeding insects and their host plants (Walling, Reference Walling2000; Mitchell, Reference Mitchell2004; Smith, Reference Smith2005; Ali & Agrawal, Reference Ali and Agrawal2012; Raman, Reference Raman2012). Recognition of an appropriate plant by an insect and the consequent response – either resistant or susceptible – of the plant can be examined in different ways. Because such relationships are usually triggered by feeding actions, an analysis of the saliva offers scope to understand insect–plant relationships better. For example, aphid-secreted salivary proteins are considered similar to plant-pathogenic effectors and therefore considered to function by perturbing host-cell processes (Arimura et al., Reference Arimura, Ozawa and Maffei2011). Adequate evidence exists that the feeding behaviours of phytophagous biting–chewing insects on the one hand and sap-sucking insects on the other are strikingly different: the former triggers a jasmonic-acid signalling pathway, whereas the latter triggers the jasmonic acid–ethylene and salicylic-acid signalling pathways (Zhao et al., Reference Zhao, Davis and Verpoorte2005). Moreover, understanding the structure and roles of salivary proteins provides scope to determine evolutionary trends in phytophagous insects and their host plants (Howe & Jander, Reference Howe and Jander2008).
Stress in plants is a state in which increasing demands are made on a plant, which leads to an immediate perturbation of its functions, followed by gradual reversal to normality of cell function and usually resulting in a better capacity of resistance (Larcher, Reference Larcher1987). During feeding, insects stress plants in multiple ways by affecting plant tissue (Backus et al., Reference Backus, Serrano and Ranger2005b ) generating stress-mitigating molecular responses (Moran & Thompson, Reference Moran and Thompson2001; Thompson & Goggin, Reference Thompson and Goggin2006), and redirecting the plant's energy in the production of novel secondary compounds (Mutikainen et al., Reference Mutikainen, Walls, Ovaska, Keinänen, Julkunen-Tiitto and Vapaavuori2002). Physical symptoms of plant stress could manifest as wilting, lesions and either localized or extended necrosis. Insect induced galls — specialized representations in insect–plant interactions — also manifest plant-stress responses (Raman, Reference Raman2011). Cells of the stressed tissue present anomalous wall structures, irregular and abnormal-wall thickenings, slack-cell membranes, extended and hyaline cytoplasm including large vacuoles, and callose formation (Raman et al., Reference Raman, Beiderbeck and Herth2009; Atkinson & Urwin, Reference Atkinson and Urwin2012), which are, by and large, similar to the stress inflicted by abiotic factors on plant cells (Harper & Horne, Reference Harper and Horne2012). When plants are unable to resist the stress effect, they experience production of reactive-oxygen species (Atkinson & Urwin, Reference Atkinson and Urwin2012), resulting ultimately either in cell and tissue death or in cell proliferation resulting in gall.
Feeding strategies
Most hemipteroids belong to the sap-sucking guild. Feeding by sap-sucking insects involves the secretion of saliva and ingestion of plant-cell components. Hemipteran mouth parts consist of mandibles and maxillae that are modified into two pairs of stylets, whereas those of the Thysanoptera have assymetrical mouthparts with either reduced or absent right mandible, which enable them to both salivate and ingest simultaneously and efficiently. They utilize one of the following strategies during feeding: (i) stylet-sheath forming, (ii) osmotic-pump feeding and (iii) cell-rupture feeding. The last strategy consists of four sub-strategies, which include lacerate-and-flush, lacerate-and-sip, lance-and-ingest and macerate-and-flush (Hori, Reference Hori, Schaefer and Panizzi2000; Backus et al., Reference Backus, Serrano and Ranger2005b ). Sheath-forming hemipteroids feed mostly on vascular tissues whereas the cell-rupture feeders feed mostly intracellularly on either mesophyll or stem parenchyma cells (Backus, Reference Backus1988). The cell-rupture feeding strategy occurs in the Typhlocybinae in the Auchenorrhyncha, Cimicomorpha and some families of the Pentatomomorpha in the Heteroptera. This type of feeding behaviour includes variation in movement of the outer – the mandibular – stylets and the inner – maxillary – stylets (Backus, Reference Backus1988). Cell-rupture tactics include (i) active and rapid laceration without complete salivation in lacerate-and-sip feeding behaviour; (ii) alternate salivation and ingestion with slow movement of stylets in lacerate-and-flush feeding; (iii) long probing time and ingestion from phloem by forming salivary pseudosheaths in lance-and-ingest type of feeding behaviour (Backus et al., Reference Backus, Serrano and Ranger2005b ) and (iv) enzyme-dependent cell degradation in macerate-and-flush feeding (Hori, Reference Hori, Schaefer and Panizzi2000). Similar to cell-rupturing technique, sheath former also forms sheaths by either movement of entire stylet into plant tissue (species of Pentatomomorpha and Sternorrhyncha) or by penetrating the maxillary stylets deeper into plant tissue compared with branched mandibular stylets (species of Auchenorrhyncha) (Backus et al., Reference Backus, Serrano and Ranger2005b ).
In a few hemipterans such as Myzus persicae (Aphididae), Bemisia tabaci (Aleyrodidae) and Nilaparvata lugens (Delphacidae), which are salivary-sheath formers, the insects insert their stylets at the feeding sites damaging a few cells. Some insects with stylet-sheath forming behaviour also employ cell-degrading enzymes, such as cellulase, amylase and pectinase, which facilitate minimal mechanical injury to the plant tissue (Miles, Reference Miles1999; Hori, Reference Hori, Schaefer and Panizzi2000; Backus et al., Reference Backus, Andrews, Shugart, Greve, Labavitch and Alhaddad2012). The stylet-sheath pathway can be either intercellular (e.g., Brevicoryne brassicae, M. persicae, Rhopalosiphum padi, all Aphididae) (Prado & Tjallingii, Reference Prado and Tjallingii2007) or intracellular (e.g., Nephotettix cincticeps, Cicadellidae) (Hattori et al., Reference Hattori, Nakamura, Komatsu, Tsuchihara, Tamura and Hasegawa2012). The Coreidae employ an osmotic-pump feeding strategy by enhancing the osmotic potential of the intercellular fluid, viz., the apoplast (Mitchell, Reference Mitchell2004). Whereas in Mictis profana (Coreidae) that employ the osmotic-pump feeding mechanism, salivary sucrase increases osmotic concentration of intercellular fluids enabling the insect to suck plant sap (Miles & Taylor, Reference Miles and Taylor1994; Taylor & Miles, Reference Taylor and Miles1994). The Lygaeidae feed by lacerating and flushing, cut the plant tissue and push-and-pull their stylets during feeding. Other cell-rupturing families, such as the Miridae, on the other hand, feed by macerating and flushing; their feeding action degenerates cell walls using specific cell-wall digesting enzymes, such as pectinases and cellulases. For example, in those hemipterans, which feed by macerating and flushing, salivary pectinase macerates the tissue (e.g., Deraeocoris nebulosus, Miridae; Boyd et al., Reference Boyd, Cohen and Alverson2002). Sap sucking (e.g., Frankliniella occidentalis, Terebrantia: Thripidae; Kindt et al., Reference Kindt, Joosten, Peters and Tjallingii2003) and sap-sucking–sheath-forming (e.g., M. profana, Coreidae; Miles & Taylor, Reference Miles and Taylor1994; Taylor & Miles, Reference Taylor and Miles1994) hemipteroids are known to feed on host-plant parenchyma and either xylem (e.g., Philaenus spumarius, Aphrophoridae; N. cincticeps, Cicadellidae; Crews et al., Reference Crews, Mccully, Canny, Huang and Ling1998; Hattori et al., Reference Hattori, Nakamura, Komatsu, Tsuchihara, Tamura and Hasegawa2012) or phloem (Acyrthosiphon pisum, Carolan et al., Reference Carolan, Fitzroy, Ashton, Douglas and Wilkinson2009). Selection of the feeding site varies according to developmental stage and stylet length. For example, short-styleted Thysanoptera feed on the upper layers of leaf tissue (Kindt et al., Reference Kindt, Joosten, Peters and Tjallingii2003), whereas the long-styleted Hemiptera (e.g., Kerria lacca, Kerridae) feed on deeper-lying stem tissues (Ahmad et al., Reference Ahmad, Kaushik, Ramamurthy, Lakhanpaul, Ramani, Sharma and Vidyarthi2012). Stylet passage can be either inter- or intra-cellular and the extent of tissue damage depends on probing strategies (Ahmad et al., Reference Ahmad, Kaushik, Ramamurthy, Lakhanpaul, Ramani, Sharma and Vidyarthi2012).
Feeding behaviour and salivary composition
Salivary glands of the Hemiptera and Thysanoptera vary in the structure and number of lobes. In the Hemiptera, one pair of glands occur, each usually comprising a principal gland functioning as a reservoir, and an accessory gland in supplying the fluid to watery saliva in the form of haemolymph ultrafiltrate (Miles, Reference Miles1999). In the Thysanoptera, two pairs of glands consist of well-differentiated structures: one pair comprises long, tubular glands that run parallel to the intestine, and a second pair that are short, ovoid and usually confined to the thorax (Del Bene et al., Reference Del Bene, Cavallo, Lupetti and Dallai1999). Feeding behaviour is critical in regulating the salivary chemistry of these insects. Most of the Heteroptera and Sternorrhyncha secrete saliva in two ways: as gelly and watery saliva. The gel saliva is composed of lipoproteins, phospholipids and conjugated carbohydrates whereas watery saliva is mainly composed of different enzymes (Miles, Reference Miles1999; Backus et al., Reference Backus, Serrano and Ranger2005b ). Recent developments, however, have enabled us to understand the enzymatic composition of the two different salivary secretions (Miles, Reference Miles1999). For example, immunolocalization techniques indicate that salivary proteins in the principal and accessory glands of Schizaphis graminum (Hemiptera: Aphididae) are different. Proteins of molecular weight of 66–69 kDa proteins were found in the watery and gel saliva, whereas 154 kDa protein in watery saliva (Cherqui & Tjallingii, Reference Cherqui and Tjallingii2000). However, in the Miridae saliva is secreted as a single type, possibly a combination of gelling and watery components (Miles, Reference Miles1999). Within the Heteroptera salivary enzymes have been characterized in the Pentatomidae, Coreidae, Lygaeidae, Dinidoridae, Pyrrhocoridae, Miridae, Acanthosomatidae, Aradidae, Cydnidae, Largidae, Scutelleridae, Berytidae and Tingidae. They have been characterized in the Cicadellidae and Delphacidae in the Auchenorrhyncha. In the Sternorrhyncha they have been characterized in the Psyllidae, Aphididae and Aleyrodidae. In Thysanoptera salivary enzymes have been characterized in Terebrantia and Tubulifera (table 1).
Host-plant stress inducing salivary proteins in the hemipteroids.
EC – Enzyme Commission number [Nomenclature Committee of the International Union of Biochemistry and Molecular Biology (NC-IUBMB)].
1 Activity of enzyme is reported.
2 Adapted molecular weight.
Damage caused by hemipteroid feeding
While feeding, the members of the Heteroptera damage plant tissues resulting in tissue thinning, malformation and necrosis (Baxendale et al., Reference Baxendale, Heng-Moss and Riordan1999; Schaefer & Panizzi, Reference Schaefer, Panizzi, Schaefer and Panizzi2000), those of the Auchenorrhyncha inflict tip wilting, plant stunting and chlorosis (Backus et al., Reference Backus, Serrano and Ranger2005b ), whereas those of the Sternorrhyncha induce necrosis and galls (Miles, Reference Miles1999). Feeding action of the Terebrantia induces necrosis in plant tissue (Hunter & Ullman, Reference Hunter and Ullman1989), whereas some of the Tubulifera (e.g., Liothrips, Gynaikothrips) result in galls (Raman, Reference Raman2003). Direct damage is due to mechanical injury caused by movement of stylets as well as chemical injury caused by salivary enzymes (Backus et al., Reference Backus, Serrano and Ranger2005b ). However sap-sucking hemipteroids do not inflict as much mechanical damage as the biting and chewing coleopteroids would; but hemipteroids may, nevertheless, inflict intense physiological changes in the host. For example, feeding actions of Diuraphis noxia and R. padi (Aphididae) on Triticum aestivum and Avena sativa (both Poaceae) alter total-protein contents, activities of peroxidase, catalase and polyphenol oxidase (Ni et al., Reference Ni, Quisenberry, Heng-Moss, Markwell and Sarath2001). In the Pentatomorpha and Miridae, salivary enzymes such as pectinase, protease, amylase and cellulase play specific roles in degrading parts of host cells in facilitating stylet insertion. Sap-sucking insects inflict more intense transcriptomic changes in plants compared with the chewing insects. Feeding by M. persicae (Aphididae) triggers changes in the expression of 2181 genes in the host-plant Arabidopsis thaliana (Brassicaceae), whereas during feeding of Pieris rapae (Lepidoptera: Pieridae) only 186 genes are activated (De Vos et al., Reference De Vos, Van Oosten, Van Poecke, Van Pelt, Pozo, Mueller, Buchala, Métraux, Van Loon, Dicke and Pieterse2005).
Apart from affecting host plants through direct interaction resulting either in necrosis or in cell damage or in gall induction, in other subtle physiological changes (Morkunas et al., Reference Morkunas, van Chung and Gabrys2011), sap-sucking hemipteroids also transmit pathogenic microbes (Purcell & Almeida, Reference Purcell and Almeida2005). Species of the Aphidoidea, Psylloidea, Aleyrodoidea, Cicadoidea, Fulgoroidea (Hemiptera) and the Terebrantia (Thysanoptera) are established vectors of many plant pathogens (Hogenhout et al., Reference Hogenhout, Oshima, Ammar, Kakizawa, Kingdom and Namba2008a ). There are two major types of insect-transmitted pathogens: circulative and non-circulative. Circulative pathogens penetrate the gut epithelial cells of their insect vectors, migrate into the haemolymph, and then to the salivary glands before vectors re-introduce them into plants via their saliva (Hogenhout et al., Reference Hogenhout, Ammar, Whitfield and Redinbaugh2008b ). Immediate salivation of the non-circulative pathogens is necessary (Powell, Reference Powell2005), since the insect gut does not have an appropriate retention capacity for non-circulative viruses in particular, as shown in the transmission of the cauliflower-mosaic virus, which binds specifically to the lining cells of the salivary canal of B. brassicae and has to be quickly transmitted to Brassica oleracea var. botryris (Brassicaeae) (Uzest et al., Reference Uzest, Gargani, Drucker, Hebrard, Garzo, Candresse, Fereres and Blanc2007). Non-circulative pathogens adhere to the stylets of vectors, subsequently are re-introduced into the plant during feeding, without circulating in the hemolymph. For example, the acrostyle, a recently discovered structure on the maxillary stylets of A. pisum is houses the stylet-borne pathogens (Uzest et al., Reference Uzest, Gargani, Dombrovsky, Cazevieille, Cot and Blanc2010). The acrostyle seems to enable either stiffening of the stylet tip or stimulation of the protein–protein interaction in Vicia faba (Fabaceae)–A. pisum interactions. It seems that the acrostyle may also be helpful in acquiring the virus from one plant and inoculating it into another, as well as launching appropriate salivary contents at the appropriate time, such as release of Ca ++ -binding proteins with watery saliva, i.e., after stylet-sheath formation preventing sieve-tube occlusion (Uzest et al., Reference Uzest, Gargani, Dombrovsky, Cazevieille, Cot and Blanc2010). Foregut-borne, non-circulative pathogens also rely on the saliva for transmission. Salivary β-glucosidase of Homalodisca vitripennis (Cicadellidae) enables the dispersal of Xylella fastidiosa bacterial cells (Xanthomonadales: Xanthomonadaceae), which occur as a dense biofilm in Vitis vinifera (Vitaceae) and the vector's foregut. β-glucosidase is considered the carrier of bacterial cells egested into the plant, thus initiating the infective process of X. fastidiosa leading to Pierce's disease in Vitis (Backus et al., Reference Backus, Andrews, Shugart, Greve, Labavitch and Alhaddad2012).
Among the stylet-sheath forming Hemiptera, the Aphididae are the most investigated. The gelly and watery saliva in the Aphididae include a variety of proteins: phenol oxidases, peroxidases, pectinases, amylases in the watery saliva; polyphenol oxidase, peroxidases and 1,4-glucosidases in the gel saliva, which perform a variety of functions. While tracking phloem cell in the host plant, taxa of the Aphididae injure many mesophyll-parenchyma cells activating wound-signalling pathways (Martinez de Ilarduya et al., Reference Martinez de Ilarduya, Xie and Kaloshian2003), whereas those of the Aleyrodoidea ‘tactically’ avoid apoplastic plant-defence compounds in host-cell vacuoles (Walling, Reference Walling2008). The saliva of these insects, while interacting with plants, either transmits microbial pathogens or induces galls. Reactive sites are vital for gall induction; salivary enzymes play a role in stimulating appropriate host physiology resulting in gall induction (Raman, Reference Raman, Seckbach and Dubinsky2010). Among gall-inducing Phlaeothripidae, salivary proteins play a critical role in gall induction. The saliva of the gravid females of Arrhenothrips ramakrishnae includes greater quantities of proteases, amylases and lipases than that of the adult males. This is significant because only adult gravid females induce galls on the leaves of Mimusops elengi (Sapotaceae); the saliva of the first and second larvae includes maximal levels of amylases, which contribute to gall growth in M. elengi (Raman et al., Reference Raman, Rajadurai, Mani and Balakrishna1999).
To the best of our knowledge, no consolidated discussion on the functions of salivary proteins of phytophagous sap-sucking hemipteroids on the levels of physiological changes in plants exists. In such a context, this article discusses salivary proteins of the hemipteroids (oxidoreductases, hydrolases, transferases, lyases, Ca++-binding proteins, the effector proteins and the newly found salivary proteins such as lamin 1, ficolins) and their role in phytophagy.
Salivary proteins and functions
Amylases, proteases, phenol oxidases, α-glucosidases, catechol-oxidases and pectinases are the most studied enzymes in hemipteroid saliva (Miles, Reference Miles1999; Hori, Reference Hori, Schaefer and Panizzi2000). Trehalases, esterases, lipases, acid and alkaline phosphotases, α-galactosidases and peptidases are equally well known (Miles, Reference Miles1999; Hori, Reference Hori, Schaefer and Panizzi2000). Whereas the salivary proteins of a majority of the Sternorrhyncha have been implicated in stressing host-plant tissues, a minority has been shown to play a role in modifying host-plant defences (Kaloshian & Walling, Reference Kaloshian and Walling2005). Detoxification of plant-defence compounds is a critical function of both the gel and watery saliva. Polyphenol oxidase and peroxidases in the gel saliva polymerize phenolics of the plant-cell apoplast as an induced-defence mechanism (Miles, Reference Miles1999). The level of volatiles emitted is low in plants attacked by phloem-feeding Hemiptera and this could be due to salivary enzymes and proteins that can inhibit synthesis of volatiles (Walling, Reference Walling2008). Activity of esterases, glutathione transferases and cytochrome P-450-dependent mono-oxygenases is known in the saliva of the polyphagous F. occidentalisis responsible for the detoxification of different plant allelochemicals, such as organic cyanides, terpenoids and alkaloids (Feyereisen, Reference Feyereisen1999; Jensen, Reference Jensen2000; Li et al., Reference Li, Williams, Loh, Lee and Howe2002).
Oxidoreductases
Catalases, catechol oxidases, superoxide dismutases, ascorbate oxidases, peroxidases, cytochromes and glucose oxidases are present in the hemipteran saliva (Miles, Reference Miles1999; Hori, Reference Hori, Schaefer and Panizzi2000; Ni et al., Reference Ni, Quisenberry, Pornkulwat, Figarola, Skoda and Foster2000; DeLay et al., Reference DeLay, Mamidala, Wijeratne, Wijeratne, Mittapalli, Wang and Lamp2012). By altering the redox balance, these proteins detoxify phenolic compounds in plant-defence reactions (Miles & Oertli, Reference Miles and Oertli1993). Alterations to the redox balance are responsible for tissue damage (Miles, Reference Miles1999; Ni et al., Reference Ni, Quisenberry, Pornkulwat, Figarola, Skoda and Foster2000; Sarker & Mukhopadhyay, Reference Sarker and Mukhopadhyay2006). For example, glutathione peroxidase in the saliva of A. pisum degrades reactive-oxygen species (ROS) generated in V. faba to achieve redox balance; in addition, glutathione peroxidase also reduces lipidhydroperoxides to their corresponding alcohols and also reduces the free H2O2 to H2O (Carolan et al., Reference Carolan, Caragea, Reardon, Mutti, Dittmer, Pappan, Cui, Castaneto, Poulain, Dossat, Tagu, Reese, Reeck, Wilkinson and Edwards2011). Oxidases in the hemipteran saliva act on the aglycones, produced through hydrolase action on glycosides, converting them into non-toxic compounds (Miles, Reference Miles1999).
Other flavin-adenine dinucleotide-dependent oxidoreductases such as glucose–methanol choline (GMC) as shown in the saliva of A. pisum mediate the oxidative detoxification of allelochemicals such as lactic, benzoic, p-hydroxybenzoic, vanillic, adipic, succinic, malic, glycolic and p-hydroxyphenylacetic acids in V. faba (Asaduzzaman & Toshiki, Reference Asaduzzaman and Toshiki2012) thus suppressing plant-defence mechanisms (Carolan et al., Reference Carolan, Fitzroy, Ashton, Douglas and Wilkinson2009). This action is similar to that of the salivary glucose oxidase (which is also a GMC oxidoreductase) in Helicoverpa zea (Lepidoptera: Noctuidae) feeding on Nicotiana tabacum (Solanaceae) reducing the nicotine-defence pathways (Eichenseer et al., Reference Eichenseer, Mathews, Bi, Murphy and Felton1999). Glucose oxidase in the saliva of M. persicae induces weak wound responses in V. faba by suppressing defence mechanisms (Harmel et al., Reference Harmel, Létocart, Cherqui, Giordanengo, Mazzucchelli, Guillonneau, De Pauw, Haubruge and Francis2008). Synthesis of glucose oxidases could be a dominant strategy in plant-feeding hemipteroids, because it affects jasmonic-acid-biosynthesis-regulating genes in plants, when attacked by different species of Aphididae (Harmel et al., Reference Harmel, Létocart, Cherqui, Giordanengo, Mazzucchelli, Guillonneau, De Pauw, Haubruge and Francis2008).
Dehydrogenases elicit plant-signalling responses to feeding by different Aphididae (Couldridge et al., Reference Couldridge, Newbury, FordLloyd, Bale and Pritchard2007). For example, M. persicae while feeding on the leaves of Solanum tuberosum (Solanaceae) rapidly enhances activities of glutamine synthase and glutamate dehydrogenase at the feeding site and an elevated activity of glutamine synthase at distant leaves (Giordanengo et al., Reference Giordanengo, Brunissen, Rusterucci, Vincent, van Bel, Dinant, Girousse, Faucher and Bonnemain2010), reinforcing the involvement of multiple genes in NO3 and sugar remobilization (Divol et al., Reference Divol, Vilaine, Thibivilliers, Amselem, Palauqui, Kusiak and Dinant2005). Glucose dehydrogenase functions similarly to glucose oxidase in suppressing plant defences (Cox-Foster & Stehr, Reference Cox-Foster and Stehr1994). However, it is unclear whether it alters the redox balance in plants. Glucose dehydrogenase occurs in the saliva of A. pisum, D. noxia (Aphididae) and M. persicae (Harmel et al., Reference Harmel, Létocart, Cherqui, Giordanengo, Mazzucchelli, Guillonneau, De Pauw, Haubruge and Francis2008; Carolan et al., Reference Carolan, Fitzroy, Ashton, Douglas and Wilkinson2009; Cooper et al., Reference Cooper, Dillwith and Puterka2010). Zn-binding dehydrogenases from the saliva of M. persicae (Cooper et al., Reference Cooper, Dillwith and Puterka2010) detoxify plant allelochemicals, especially the alcohols. Alcohol NADP+ oxidoreductases reduced aldehydes to alcohol and triggered salicylic-acid, methyl-jasmonate and ethylene biosynthesis pathways in tested plants (Somssich et al., Reference Somssich, Wernert, Kiedrowski and Hahlbrock1996; Montesano et al., Reference Montesano, Hyytiainen, Wettstein and Palva2003).
Phenol oxidases cause browning of individual plant cells by accumulating o-quinones and by triggering hydroxylation of monophenols to o-diphenols and oxidation of o-diphenols to quinines (Urbanska et al., Reference Urbanska, Leszczynski, Laskowska, Matok, Nieto and Dixon1998). Phenol oxidases are vital for the detoxification of phenolic compounds; salivary phenol oxidases from Aphis gossypii, Macrosiphum euphorbiae, Macrosiphum rosae, Sitobion avenae, M. persicae (Aphididae) and some taxa of the Miridae (Hori, Reference Hori, Schaefer and Panizzi2000; Sarker & Mukhopadhyay, Reference Sarker and Mukhopadhyay2006). Phenol-oxidase activity is also known from the salivary sheaths of different aphids and in halos around the sheaths on artificial diets (Urbanska et al., Reference Urbanska, Tjallingii and Leszczynski1994). Phenol oxidases, in high likelihood, oxidize the plant polyphenols to o-quinones. During penetration by stylets, the host plant produces phenolics as a defence reaction and phenol oxidases detoxify them by hydrolysation (Miles, Reference Miles1999).
Ascorbate oxidase – a phenol oxidase – occurs in D. noxia and R. padi and detoxifies plant phenolics (Ni et al., Reference Ni, Quisenberry, Pornkulwat, Figarola, Skoda and Foster2000). Laccase and catechol oxidase have been shown in N. cincticeps, while feeding on Oryza sativa (Poaceae) (Hattori et al., Reference Hattori, Konishi, Tamura, Konno and Sogawa2005), where catechol oxidase is indicated to play a role in overcoming plant defences (Miles, Reference Miles1999). Catechol oxidase in the saliva of N. cincticeps oxidizes and polymerizes phenolic compounds that accumulate around the salivary sheath. Monolignols in O. sativa produce quinine methides, when plant cells are ruptured by stylet action of N. cincticeps, and these quinine methides are protein-alkylating agents, which harm N. cincticeps (quinines acts on proteins and diminishes dietary protein value for insects by alkylation; Duffey & Stout, Reference Duffey and Stout1996) (Hattori et al., Reference Hattori, Konishi, Tamura, Konno and Sogawa2005). Laccase oxidizes monolignols to lignin in O. sativa and is responsible for making the host-plant consumable.
Peroxidases dehydrogenate phenolic substances (e.g., chlorophenols) and produce phenoxy radicals in the presence of H2O2 to form phenolic polymers. Different peroxidases occur in A. gossypii, Therioaphis trifolii maculata, M. euphorbiae, M. rosae, R. padi, S. avenae and M. persicae, which appear to mimic phenol oxidases by acting on host-plant phenolics, particularly on alkaloids (e.g., DIMBOA [2,4-dihydroxy-7-methoxy-1,4-benzoxazin-3-one], gramine) on which phenol oxidase cannot act. H2O2 is critical for peroxidase activity; phenol oxidase releases H2O2 acting on plant phenolics. Hence, the activity of phenol oxidase and peroxidase is possibly synergistic (Miles, Reference Miles1999; Cherqui & Tjallingii, Reference Cherqui and Tjallingii2000; Ni et al., Reference Ni, Quisenberry, Pornkulwat, Figarola, Skoda and Foster2000). While feeding on Camellia sinensis (Theaceae), these enzymes in the saliva of Helopeltis theivora (Miridae) induce necrosis in growing shoots (Sarker & Mukhopadhyay, Reference Sarker and Mukhopadhyay2006). Oxidation of phenols is necessary for insect survival on plants, but this process also produces H2O2, which can damage plant cells. The concurrent presence of peroxidase in insect saliva may act on the synthesized H2O2 and reduce it, thus preventing a hypersensitive response of the plant and enabling the insect to consume the plant without interruption. Catalases and superoxide dismutase act on H2O2 and convert it into H2O and O2. Catalase is reported from the salivary secretions of D. noxia and is responsible for leaf chlorosis in susceptible wheat plants either by reducing chlorophyll synthesis or by degrading chlorophyll and indirectly affecting the redox balance of the plant (Ni et al., Reference Ni, Quisenberry, Pornkulwat, Figarola, Skoda and Foster2000).
Cytochrome P-450 (Cyt P-450) is a major enzyme class in the saliva of almost all insects. They act as mixed-function oxidases and mono-oxygenases. Cyt P-450 is critical for defending insects against plant chemicals. Adaptation of insects to a particular host plant also depends on plant-chemical break down using Cyt P-450 (Feyereisen, Reference Feyereisen1999). Salivary Cyt P-450 in D. noxia, while feeding on T. aestivum, detoxifies allelochemicals, such as p-hydroxybenzoic, trans-p-coumaric, cis-p-coumaric, syringic, vanillic, trans-ferulic and cis-ferulic acids, DIMBOA (Wu et al., Reference Wu, Haig, Pratley, Lemerle and An2000; Nicholson et al., Reference Nicholson, Hartso and Puterka2012). Cyt P-450 sequences are vital in mediating isoprenoid biosynthesis; isoprenoids function in plant–arthropod signalling as herbivore repellents and as attractants of arthropod parasitoids (Boyko et al., Reference Boyko, Smith, Thara, Bruno, Deng, Starkey and Klaahsen2006), although this aspect has not yet been demonstrated in plant-feeding hemipteroids.
Hydrolases
Hydrolases induce phytotoxicosis, such as wilting and necrosis of plant tissues, as shown in Eucalyptus regnans and Eucalyptus nitans (Myrtaceae) consequent to feeding by Amorbus obscuricornis (Coreidae) (Steinbauer et al., Reference Steinbauer, Taylor and Madden1997) and browning of cells in S. tuberosum following feeding by S. avenae (Urbanska et al., Reference Urbanska, Leszczynski, Laskowska, Matok, Nieto and Dixon1998). Pectinase, cellulase and amylase soften the tissue facilitating stylet entry and its movement in host tissue. Simultaneously, pectinases provide gustatory clues and render the plant amenable to feeding. To elicit the gustatory clue, the aphids initially taste cell contents. For example, M. persicae feeds on artificial diets that include 2, 3-diacetyl pectin, a component in its preferred host Beta vulgaris (Amaranthaceae). Monophagous (e.g., B. brassicae) and oligophagous aphids (e.g., Melanocallis caryaefoliae) show greater gustatory sensitivity for plant polysaccharides compared with polyphagous aphids (e.g., M. persicae). Aphids with a lesser capability to discriminate between different polysaccharides usually remain polyphagous (Campbell et al., Reference Campbell, Jones and Dreyer1986). While degrading plant tissue, damage due to hydrolases varies according to feeding sites: when species of Lygus feed on mesophyll tissue they induce simple lesions, but when they feed on meristematic tissue, they inflict severe damage resulting in tissue malformation (Hori, Reference Hori, Schaefer and Panizzi2000).
Pectinase degrade pectin, one of the principal components of the middle lamella of plant cells. As part of the watery saliva, pectinases induce degenerative changes in host cells, as shown in the saliva of Lygus hesperus (Miridae) in disintegrating parenchyma cells (Strong & Kruitwagen, Reference Strong and Kruitwagen1968) and in pre-empting a wound response by producing pectin fragments (Miles, Reference Miles1999; Hori, Reference Hori, Schaefer and Panizzi2000). Activity of pectinase is reported from different aphids and the Heteroptera (table 1).
Cellulase activity is known from S. graminum enabling the depolymerization of xylans and arabinogalactans of cell walls, which on secretion into plant tissue render cellulose ingestible. Hydrolysis of cell-wall polysaccharides by salivary cellulase facilitates stylet penetration, although the mechanical properties of the cell wall regulate its movement within, as shown in Acrosternum hilare (Pentatomidae) in enabling stylet penetration in Sorghum vulgare (Poaceae), but due to degradation of cellulose, occurrence of wound responses due to cellulose action is a possiblity (Miles, Reference Miles1999). β -1,4-glucanase (which is related to cellulase) in the saliva of H. vitripennis (Cicadellidae) hydrolyses hemicelluloses in cell walls, facilitating the sealing of the sheath-encased stylet tips into xylem elements. Enzymes in the saliva are also hypothesized to loosen populations of Pierce's-disease bacterium in the vector's foregut, allowing subsequent egestion to inoculate the bacteria into the xylem of V. vinifera (Backus et al., Reference Backus, Andrews, Shugart, Greve, Labavitch and Alhaddad2012).
Polygalacturonase is a pectin-hydrolysing enzyme that enables intercellular-stylet movement (Campbell & Dreyer, Reference Campbell, Dreyer, Campbell and Eikenbary1990). Endo- and exo-polygalacturonases are known from the hemipteran saliva (Laurema & Nuorteva, Reference Laurema and Nuorteva1961; Miles, Reference Miles1999; Celorio-Mancera et al., Reference Celorio–Mancera, Greve, Teuber and Labavitch2009). Activity of these enzymes is recorded for Lygus rugulipennis, Lygus pratensis, Orthops kalmii (Miridae), Adelphocoris lineolatus (Miridae) Closterotomus norwegicus (Miridae) (Frati et al., Reference Frati, Galletti, Lorenzo, Salerno and Conti2006), S. graminum, A. pisum and M. persicae (Miles, Reference Miles1999; Cherqui & Tjallingii, Reference Cherqui and Tjallingii2000). Exo-polygalacturonase produces lesser quantities of galacturonides by acting on the oligouronides compared with other pectinases. Endo-polygalacturonases degrade pectin and generate oligosaccharides (Celorio-Mancera et al., Reference Celorio–Mancera, Greve, Teuber and Labavitch2009). Polygalacturonase activity is shown in the saliva as well as in other body parts of the Hemiptera, in halos of watery saliva restored in artificial diets and in salivary proteins (Miles, Reference Miles1999; Frati et al., Reference Frati, Galletti, Lorenzo, Salerno and Conti2006).
Amylases are the most prevalent enzymes in the saliva of most of the phytophagous hemipteroids (Urbanska & Leszczynski, Reference Urbanska and Leszczynski1997; Hori, Reference Hori, Schaefer and Panizzi2000; Harmel et al., Reference Harmel, Létocart, Cherqui, Giordanengo, Mazzucchelli, Guillonneau, De Pauw, Haubruge and Francis2008). α- and β-amylase and amyloglucosidase occur in the saliva of Anoplocnemis curvipes, Clavigralla tomentosicollis, Clavigralla shadabi (all Coreidae), Riptortus dentipes and Mirperus jaculus (both Alydidae) that feed on Vigna unguiculata (Fabaceae). Whereas the α-amylase and amyloglucosidase hydrolyse starch to release energy, β-amylase hydrolyses 1,4-α-d-glycosidic linkages of starch and releases β-maltose. Amyloglucosidase removes glucose units from amylopectin and hydrolyses a greater proportion of starch from V. unguiculata (Soyelu et al., Reference Soyelu, Akingbohungbe and Okonji2007). α-amylase is also known to occur in Eurygaster maura (Scutelleridae), which feeds on wheat seeds (Mehrabadi & Bandani, Reference Mehrabadi and Bandani2009). Lower-molecular weight of salivary α-amylase suggests that salivary α-amylase is an isozyme (Zibaee et al., Reference Zibaee, Hoda and Fazeli-Dinan2012). In the Heteroptera, amylase activity is known from different species of the Pentatomidae, Coreidae, Lygaeidae, Dinidoridae, Pyrrhocoridae, Miridae, Aradidae, Cydnidae, Largidae, Scutelleridae, Berytidae and Tingidae (Hori, Reference Hori, Schaefer and Panizzi2000) (table 1). In the Sternorrhyncha, amylases have been recorded in R. padi and S. avenae (Miles, Reference Miles1999). While feeding on V. faba, S. avenae releases α−1,4-glucan-4-glucanohydrolase, which is an amylase, that metabolizes the carbohydrates in V. faba, but simultaneously stressV. faba. Salivary α-amylase in M. persicae catalyses the hydrolysis of oligosaccharides and polysaccharides and renders glucose to M. persicae (Harmel et al., Reference Harmel, Létocart, Cherqui, Giordanengo, Mazzucchelli, Guillonneau, De Pauw, Haubruge and Francis2008). Amylase activity has also been reported from A. ramakrishnae (Thysanoptera: Phlaeothripidae) and B. tabaci (Cohen & Hendrix, Reference Cohen and Hendrix1994; Raman et al., Reference Raman, Rajadurai, Mani and Balakrishna1999).
Trehalase degrades trehalose to two molecules of glucose. Trehalose (1,1-α-d-glucopyranosyl α-d-glucopyranoside) has both protective and adverse effects on plants in that it is critical for the infectivity of pathogens and in eliciting defence responses to abiotic and biotic stresses, although its exact role and mechanism in biotic stress is being debated (Fernandez et al., Reference Fernandez, Béthencourt, Quero, Sangwan and Clément2010). Trehalose has been implicated for defence response in Arabidopsis thaliana against M. persicae. Trehalose-PO4-synthase 11 (TPS11) is critical for antixenosis and antibiosis against M. persicae by promoting relocation of C into starch; TPS11 thus enhances accumulation of starch in plant tissue in lieu of sucrose. Trehalase occurs not only in the saliva of A. pisum but also in its midgut (Cristofoletti et al., Reference Cristofoletti, Riberio, Deraison, Rahbé and Terra2003; Carolan et al., Reference Carolan, Caragea, Reardon, Mutti, Dittmer, Pappan, Cui, Castaneto, Poulain, Dossat, Tagu, Reese, Reeck, Wilkinson and Edwards2011; Nicholson et al., Reference Nicholson, Hartso and Puterka2012). Degradative action of trehalase may affect trehalose-based plant defensive responses (trehalose delays programmed-cell death, elicits plant defence, promotes stress responses to proteins such as ϕ -glutathione S-transferase 2 [AtGSTF2], flavin mononucleotide-binding flavodoxin-like quinone reductase 1 [FQR1], cytosolic dehydroascorbate reductase 1 [DHAR1] and S-adenosylmethionine synthetase 2 [SAMS2]) (Baea et al., Reference Baea, Hermanb, Baileya, Baec and Sicher2005), besides enabling insect survival on V. faba (Nicholson et al., Reference Nicholson, Hartso and Puterka2012).
α-glucosidase, α−1,4-glucosideglucohydrolase, amyloglucosidase, β-glucosidases (β−1,4-glucoside glucohydrolase), 1,3-glucosidase and 1,4-glucosidase are known in the Hemiptera. These enzymes mediate hydrolysis of glucose molecules and the ‘toxic’ phenolic glycosides in plants, which are released during stylet insertion in mesophyll tissues. Glucosidases convert phenolic glycosides into aglycones. Glucosidases have been isolated from the salivary-gland homogenates of S. avenae and R. padi and from gut extract and watery saliva of M. profana (Miles, Reference Miles1999; Hori, Reference Hori, Schaefer and Panizzi2000). Amyloglucosidase in the saliva of A. curvipes, C. tomentosicollis, C. shadabi, R. dentipes and M. jaculus hydrolyse greater quantities of starch as shown in V. unguiculata compared with other salivary amylases in providing energy to these Coreidae (Soyelu et al., Reference Soyelu, Akingbohungbe and Okonji2007). 1,3-glucosidases break pectin contents of plant tissue and may also act on the callose on sieve plates (Miles, Reference Miles1999). α- and β-galactosidase, are also carbohydrate digesting enzymes, whose weak activity is reported from saliva of Palomena angulosa (Pentatomidae), Coreus marginatus (Coreidae) and Orthocephalus funestus (Miridae).
β-d-fructofuranosidase and α-d-glucohydrolase are sucrose, maltose and trehalose hydrolysers. Urbanska & Leszczynski (Reference Urbanska and Leszczynski1997) confirmed that probing by S. avenae and R. padi into substrates containing sucrose releases glucose by the activity of β-d-fructofuranosidase, which is available as a carbon source for the feeding insects. In the saliva of M. profana (Coreideae), β-d-fructofuranosidase enables degradation of sucrose to glucose and fructose and enhancing apoplast osmotic pressure and helps M. profana to feed from apoplast (Miles & Taylor, Reference Miles and Taylor1994; Taylor & Miles, Reference Taylor and Miles1994). α-dihydroxy-glucohydrolase is known in the saliva of R. padi and S. avenae; however, its role is confusing because the usual function of this enzyme is to unload the phloem content; yet this function is not required by these aphids. However, gall-inducing aphids may use this enzyme for feeding indirectly from sieve-tube elements (Miles, Reference Miles1999). Sucrase in the saliva of A. obscuricornis and Gelonus tasmanicus (Coreidae) possibly enables the insects to feed on phloem and in consequence, induces wilting and necrosis in the leaves of Eucalyptus regans and Eucalyptus obliqua (Steinbauer et al., Reference Steinbauer, Taylor and Madden1997).
Another less frequently occurring hydrolase is chito-oligosaccharidolytic β-N-acetylglucosaminidase (NAGase) from the saliva of D. noxia feeding on T. aestivum (Poaceae). NAGase is a chitinase protein and is involved principally in the regeneration of the exoskeleton of D. noxia. It is also known to enhance fecundity in M. persicae, when overexpressed in S. tuberosum (Saguez et al., Reference Saguez, Hainez, Cherqui, Van Wuytswinkel, Jeanpierre, Lebon, Noiraud, Beaujean, Jouanin, Laberche, Vincent and Giordanengo2005) and functions as an antifungal compound in other plants by hydrolysing N-glycans of polysaccharides and glycoproteins (Altmann et al., Reference Altmann, Staudacher, Wilson and März1999). Because of such a role, in the saliva of D. noxia, it possibly inhibits fungal infection in the probed-plant tissues; it appears that NAGase-s act in concert with chitinase and chitin synthase providing opportunity for controlled lysis and synthesis of chitin is known (Horsch et al., Reference Horsch, Mayer, Sennhauser and Rast1997). In plants, NAGase functions as a defence protein against fungal pathogens therefore NAGase in the aphid saliva possibly protects the stylet from the host plant's NAGase activity (Nicholson et al., Reference Nicholson, Hartso and Puterka2012).
Peptidase (protease, proteinase) are the protein-hydrolysing enzymes, reported from a range of hemipteroids. Proteases are important for initiating gall induction in M. elengi by A. ramakrishnae. In pod-sucking Coreidae, protease occur abundantly in saliva and are responsible for characteristic symptoms such as shrivelling of young pods, partially filled older pods and dimpled seeds in mature pods (Soyelu et al., Reference Soyelu, Akingbohungbe and Okonji2007).
CLIP-domain serine protease (clip-SP, paper-clip-like domain) occurs in the saliva of A. pisum, where it inhibits phenol oxidase-based innate defences of V. faba (Carolan et al., Reference Carolan, Caragea, Reardon, Mutti, Dittmer, Pappan, Cui, Castaneto, Poulain, Dossat, Tagu, Reese, Reeck, Wilkinson and Edwards2011). M2 metalloprotease (angiotensin-converting enzyme) and M1 zinc metalloprotease are also known from the saliva of A. pisum, where these enzymes destroy plant-defence proteins and improve food quality for phloem-feeding insects by increasing the level of free-amino acids in phloem. Although the exact mechanism is unclear, M2 metalloprotease degenerates signalling peptides, such as hormones and neuropeptides (Carolan et al., Reference Carolan, Caragea, Reardon, Mutti, Dittmer, Pappan, Cui, Castaneto, Poulain, Dossat, Tagu, Reese, Reeck, Wilkinson and Edwards2011).
Phosphatases are responsible for dephosphorylation of proteins. Alkaline phosphatase (ALP) is recorded from the saliva of B. tabaci and B. argentifolii (Aleyrodidae). ALP in other tissues plays a secondary role in the production of sheath and glue from colleterial glands for the attachment of eggs to foliage (Funk, Reference Funk2001; Cooper et al., Reference Cooper, Dillwith and Puterka2010; Yan et al., Reference Yan, Peng, Liu, Wan and Harris2011). It is also reported from salivary glands of other Hemiptera such as one species of Lygaeus, Dysdercus koenigii and Coridius janus (Dinidoridae; Hori, Reference Hori, Schaefer and Panizzi2000). Acid phosphatase is reported from Oncopeltus fasciatus, C. janus, D. koenigii, L. rugulipennis, and Cletus signatus, but not in any taxon of the Aphididae (Hori, Reference Hori, Schaefer and Panizzi2000).
Lipase, esterase and pectin-methylesterase (pectinesterase) form another group of hemipteran salivary enzymes which act on lipids. Lipase activity is shown in the salivary glands of A. ramakrishnae (Raman et al., Reference Raman, Rajadurai, Mani and Balakrishna1999), and Trioza jambolanae (Triozidae) (Rajadurai et al., Reference Rajadurai, Mani, Balakrishna and Raman1990) and a few taxa of the Lygaeidae and Miridae (Hori, Reference Hori, Schaefer and Panizzi2000; Sarker & Mukhopadhyay, Reference Sarker and Mukhopadhyay2006). Pectinesterase is important in intercellular-stylet penetration by dissolving the middle lamellae (Campbell & Dreyer, Reference Campbell, Dreyer, Campbell and Eikenbary1990) and reported in A. pisum, D. noxia, M. persicae, R. padi and S. graminum (Ma et al., Reference Ma, Reese, Black and Bramel-cox1990; Cherqui & Tjallingii, Reference Cherqui and Tjallingii2000; Ni et al., Reference Ni, Quisenberry, Pornkulwat, Figarola, Skoda and Foster2000) and carboxylesterhydrolases in O. fasciatus (Hori, Reference Hori, Schaefer and Panizzi2000). Presence of pectinesterase in Phenacoccus manihoti (Pseudococcidae) is shown to be responsible for degrading middle lamellae of Manihot esculenta (Malpighiales: Euphorbiaceae) (Calatayud et al., Reference Calatayud, Boher, Nicole and Geiger1996).
Transferases and lyases
Information on transferases in insect saliva is limited, although the role of glutathione S-transferase in insects is established for inducing resistance against insecticides (Kostaropoulos et al., Reference Kostaropoulos, Papadopoulos, Metaxakis, Boukouvala and Papadopoulou-Mourkidou2001). Activity of cell-degrading phosphorylase is known in Pyrrhocoris apterus (Pyrrhocoridae) (Hori, Reference Hori, Schaefer and Panizzi2000). The function of phosphorylase, in general is to confer or add a phosphate group to a protein or a compound. They hydrolyse starch into glucose through a cascade of events (Rathore et al., Reference Rathore, Garg, Garg and Kumar2009) and also phosphorylate proteins (Giordanengo et al., Reference Giordanengo, Brunissen, Rusterucci, Vincent, van Bel, Dinant, Girousse, Faucher and Bonnemain2010) or possibly facilitate the generation of ROS (Manda et al., Reference Manda, Nechifor and Neagu2009) thereby stressing the host plant. Lyases degrade the substrate without hydrolysis and oxidation. In hemipteroid saliva, lyase activity has not been much explored but the role of hydroperoxide lyase is demonstrated in hemipteroids, such as in Sogatella furcifera. Hydroperoxide lyase plays a role in inducing resistance in O. sativa to bacterial blight, Xanthomonas oryzae (Gomi et al., Reference Gomi, Satoh, Ozawa, Shinonaga, Sanada, Sasaki, Matsumura, Ohashi, Kanno, Akimitsu and Takabayashi2010). Hydroxymethylglutaryl-CoA lyase has been detected in salivary constituents of D. noxia. This is an enzyme of mitochondrial origin and primarily helps in ketone-body production. It seems to degrade plant proteins and interfere in lipid signalling and hence may alter host-plant physiology (Nicholson et al., Reference Nicholson, Hartso and Puterka2012).
Ca++-binding proteins
Ca++-binding protein (Calmoludin) are recorded in the saliva of many aphids. After puncturing sieve-tube elements, aphids feed passively with phloem flowing into their stylet canals enabled by a turgor-pressure gradient. Innate plant-defence mechanisms enable occlusion of sieve-tube elements by callose, a high-molecular weight β-(1,3)-glucan polymer, in response to the wound inflicted by insertion of the stylets. Callose formation is preceded by the formation of proteinaceous materials (Furch et al., Reference Furch, van Bel, Fricker, Felle, Fuchs and Hafke2009), such as the forisomes (the contractile protein bodies that can modify their structures from crystalloid to sphaeroid formations) shown in Fabaceae, the parietal-phloem proteins (e.g., GFP–SEO proteins) in Cucurbitaceae, and the phloem-protein network in Brassicaceae (Sjölund, Reference Sjölund1997). However, what is common among the three studied plant families is that the occlusion of sieve elements is Ca++ion dependent (Knoblauch & van Bel, Reference Knoblauch and van Bel1998). To prevent sieve-tube occlusion, the Hemiptera use salivary Ca++-binding proteins such as calmodulin, calreticulin, C002 protein, angiotensin and PR1-like protein SMP-30 (regucalcin), NcSP84 and Calreticulin-like isoform 1, which bind with Ca++ influx in the phloem restricting sieve-tube occlusion. The presence of these proteins was detected in saliva of A. pisum, Megoura viciae, D. noxia, Aphis fabae, M. euphorbiae, S. graminum and N. cincticeps (Carolan et al., Reference Carolan, Fitzroy, Ashton, Douglas and Wilkinson2009, Reference Carolan, Caragea, Reardon, Mutti, Dittmer, Pappan, Cui, Castaneto, Poulain, Dossat, Tagu, Reese, Reeck, Wilkinson and Edwards2011; Will et al., Reference Will, Kornemann, Furch, Tjallingii and van Bel2009; Hattori et al., Reference Hattori, Nakamura, Komatsu, Tsuchihara, Tamura and Hasegawa2012; Nicholson et al., Reference Nicholson, Hartso and Puterka2012). This is also coincide with study of feeding behaviour by electrical penetration graph (EPG) (Prado & Tjallingii, Reference Prado and Tjallingii1994; Tjallingii, Reference Tjallingii2006). EPG study of A. pisum, M. viciae, M. euphorbiae, B. brassicae, A. gossypii and M. persicae shows that while continuously ingesting phloem sap, aphids switch from ingestion to secretion of watery saliva. Wounding triggers the plant for Ca++ influx, enabling sieve-tube occlusion, which, in turn, enables a drop in sieve-tube pressure stimulating the aphids to secrete Ca++ including watery-saliva (Will et al., Reference Will, Kornemann, Furch, Tjallingii and van Bel2009). The acrostyle on the maxillary stylets of A. pisum is believed to be responsible for releasing Ca++-binding proteins with watery saliva (Uzest et al., Reference Uzest, Gargani, Dombrovsky, Cazevieille, Cot and Blanc2010).
Calreticulin interferes with Ca++ influx in probed plant cells through chelation and may circumvent Ca++-mediated wound responses of the host plant (Carolan et al., Reference Carolan, Caragea, Reardon, Mutti, Dittmer, Pappan, Cui, Castaneto, Poulain, Dossat, Tagu, Reese, Reeck, Wilkinson and Edwards2011). Calreticulin-like isoform-1 enables A. pisum to ingest phloem sap. It alters the mechanism of blocking the sieve-tube element, which usually eventuates as a plant response to insect attack (Nicholson et al., Reference Nicholson, Hartso and Puterka2012). C002 protein helps the insect to feed on phloem sap of the plant by altering the blocking mechanism, which is induced due to plant response to insect attack. In addition, it is also implicated in converting forisomes of the sieve element to contract, preventing blockage of sieve elements (Mutti et al., Reference Mutti, Louis, Pappan, Pappan, Begum, Chen, Park, Dittmer, Marshall, Reese and Reeck2008). PR1-like proteins of insect saliva, which are a homologue of the PR1 protein of plants, can interfere with the function of PR1 in plants. PR1 are considered lipid-transfer proteins that mediate the signalling of systemic defence responses in plants, thereby altering defence mechanisms (van Loon et al., Reference van Loon, Rep and Pieterse2006; Carolan et al., Reference Carolan, Fitzroy, Ashton, Douglas and Wilkinson2009). SMP-30 (Regucalcin) is a Ca++-binding protein reported from A. pisum feeding on V. faba. This protein is also known to have a suppressive effect on intracellular calcium ion homoeostasis and thus regulate intracellular signalling (Carolan et al., Reference Carolan, Fitzroy, Ashton, Douglas and Wilkinson2009). Other than aphids, recently Ca++-binding proteins (NcSP84) are reported from N. cincticeps, which feeds on phloem and xylem of O. sativa (Youn, Reference Youn1998; Hattori et al., Reference Hattori, Nakamura, Komatsu, Tsuchihara, Tamura and Hasegawa2012).
Effector proteins
In insect–plant interactions, an induced plant-defence mechanism is critical. This mechanism is a part of the plant's innate immune system and works in two phases, as shown in microbial pathogenesis (Hogenhout & Bos, Reference Hogenhout and Bos2011). The first phase depends on the recognition of microbial-associated molecular patterns of pathogens, such as ‘flagellin’ (flg), a protein, which is recognized by the host plant's pattern-recognition receptors. This recognition induces a ‘microbial-associated molecular pattern-triggered immunity’ (Jones & Dangl, Reference Jones and Dangl2006). The microbial pathogens counter the host-plant's triggered immunity by generating effector molecules, which could be proteins suppressing such a first-phase immunity. These effectors induce an effector-triggered immunity, which is associated with plant disease resistance genes (e.g., R genes) as the second phase. Similar effectors, e.g., Mp10, Mp42 and MpC002 have been reported in the saliva of M. persicae (Mp), feeding on Nicotiana benthamiana (Bos et al., Reference Bos, Prince, Pitino, Maffei, Win and Hogenhout2010). These effector proteins inflict chlorosis in N. benthamiana, when overexpressed. Mp-10 effector causes chlorosis and local cell death; however, it does not show response in other plants on which it was tested, viz., N. tabacum and Solanum lycopersicum. Mp10 suppresses flg22-induced oxidative burst responses of the plant-defence mechanism, but does not suppress any chitin-induced oxidative burst response, which could be caused by the stylet action. Moreover, overexpression of Mp10 also causes reduced fecundity of aphids. Similarly, Mp 42 also reduces fecundity of aphids but MpC002 increases fecundity of aphids (Bos et al., Reference Bos, Prince, Pitino, Maffei, Win and Hogenhout2010).
Non-enzymatic proteins
A putative ficolin-3-like protein is known in D. noxia. This protein contributes to the innate immunity of animals. Ficolins, powerful molecules in host defence (Endo et al., Reference Endo, Matsushita and Fujita2011), can recognize N-acetyl compounds such as lipopolysaccharides of bacterial and fungal cell walls. Ficolins can activate the associated complementary compounds such as lectins enabling phagocytosis and the breakdown of pathogenic microbes. In sap-sucking insects ficolins prevent secondary infection of the host plant during stylet insertion. Moreover, cellular Ca++ influxes during aphid feeding may provide suitable conditions for ficolin activity as Ca++ is required as a cofactor for ficolin activity (Nicholson et al., Reference Nicholson, Hartso and Puterka2012). Nuclear lamin-like protein (L1 alpha) is also known from D. noxia. It functions as intermediate filaments (IF) proteins. IF-proteins are, however, capable of modifying their configuration and adapt to performing new functions. Elasticity is another property of this protein. IF-proteins are capable of cushioning cellular mechanical stress. Presence of exoskeleton in insects is explained as a reason behind the absence of cytoplasmic IF in insects, and the IF-proteins can be compensated by other proteins in insects (Herrmann & Strelkov, Reference Herrmann and Strelkov2011). However, their exact role is not known yet but, L1 alpha may function in reducing mechanical stress during stylet insertion by aphids into plant tissue (Nicholson et al., Reference Nicholson, Hartso and Puterka2012).
Vesicular-fusion protein, N-ethylmaleimide-sensitive factor 1 (Nsf1) is an ATPase. ATPases occur in all eukaryotic cells. Nsf-s are concerned with membrane fusion and can regulate neurotransmission. Expression of one of the negative mutants of Nsf inflicts cell death (Zhao et al., Reference Zhao, Xu, Qian, Lv, Ji, Chen, Zhao, Zheng, Gu, Xie and Mao2008). In plants after wounding, sieve plates occlusion occurs, preventing phloem-sap flow. In the saliva of D. noxia, the presence of Nsf1 may hinder vesicle formation, which is necessary for sieve-tube element fusion (Nicholson et al., Reference Nicholson, Hartso and Puterka2012).
Actin, a multifunctional protein, occurs in all eukaryotes, and is an integral component of cytoskeleton. In D. noxia saliva, three putative actin-binding and depolymerizing proteins are known. Actin-depolymerizing factors are essential for Meloidogyne incognita (Nematoda: Heteroderidae) infestation (Clément et al., Reference Clément, Ketelaar, Rodiuc, Banora, Smertenko, Engler, Abad, Hussey and de Almeida Engler2009). Stretchin–myosin light chain kinase protein that enables the assembly of actin filaments in its host-plant tissue is known in D. noxia. Putative cofilin–actin depolymerizing factor-like protein facilitates actin depolymerization. Hence, these proteins may prevent activation of defence responses of plants to insect feeding, which depends on actin polymerization (Nicholson et al., Reference Nicholson, Hartso and Puterka2012).
Microtubule-associated protein futsch (MAP1) known in the nerve signalling and microtubulular organization in Drosophila melanogaster (Diptera: Drosophilidae) (Bettencourt da Cruz et al., Reference Bettencourt da Cruz, Schwärzel, Schulze, Niyyati, Heisenberg and Kretzschmar2005), is also known from plants, although their role in plants is not yet established in microtubular organization (Gardiner & Marc, Reference Gardiner and Marc2011). MAP1 in the saliva of D. noxia is therefore inferred to act similarly to plant MAP proteins, obstructing cell signals and thus facilitating the feeding action of insects (Nicholson et al., Reference Nicholson, Hartso and Puterka2012). Lava lamp (Lva), a golgin protein, in D. noxia is implicated in cellularization, which is the separation of a multi-nucleate cell into several uninucleate cells. The Lva domains have been demonstrated to bind the microtubule-dependent motility factors and inhibit Golgi movement leading to cellularization in Drosophila melanogaster. It is proposed that Lva may interfere with the Golgi particle-related process of protein synthesis in companion cells of the phloem and surrounding plant tissues and thus can be toxic in companion cells (Nicholson et al., Reference Nicholson, Hartso and Puterka2012).
Guanosine triphosphate (GTP)-binding Di-Ras2-like protein is a GTPase and can hydrolyse GTP. Whereas the ‘Ras’ proteins activate nerve-tissue formation (Hall & Lalli, Reference Hall and Lalli2010), overexpression of Di-Ras2-like protein inhibits cell growth and cell survival as shown in human tissue (Gasper et al., Reference Gasper, Sot and Wittinghofer2010). Di-Ras2-like protein, shown in D. noxia saliva, may inflict damage through vacuolization in host-plant cells (Nicholson et al., Reference Nicholson, Hartso and Puterka2012).
Lipases catalyse either formation or cleavage of fats. In insect saliva, the only lipase known is apolipophorin from A. pisum (Carolan et al., Reference Carolan, Fitzroy, Ashton, Douglas and Wilkinson2009) and D. noxia (Nicholson et al., Reference Nicholson, Hartso and Puterka2012). In many other insects including Manduca sexta (Lepidoptera: Sphingidae) and Schistocerca gregaria (Orthoptera: Acrididae) apolipophorin is implicated in lipid transportation (Wang et al., Reference Wang, Sykes and Ryan2002; van der Horst & Rodenburg, Reference van der Horst and Rodenburg2010). Apolipophorin is abundant in insect haemolymph and participates in the insect's immune system. It has the capacity to interact and alter the plant-defensive sterols, fatty acids and carotenoids (Ma et al., Reference Ma, Hay, Li, Asgari and Schmidt2006; Zdybicka-Barabas & Cytryńska, Reference Zdybicka-Barabas and Cytryńska2011). Secreted apolipophorins could interfere with signalling of a plant's cellular immune response, apolipophorins after binding to lipid elicitor molecule undergoes a confirmational change and induces plant-immune response.
Conclusion
Feeding behaviours among the hemipteroids vary widely as indicated by EPG studies on R. padi (Prado & Tjallingii, Reference Prado and Tjallingii1994), Diaphorina citri (Psyllidae; Bonani et al., Reference Bonani, Fereres, Garzo, Miranda, Appezzato-da-Gloria and Lopes2010), Phenacoccus solenopsis (Pseudococcidae) (Huang et al., Reference Huang, Tjallingii, Zhang, Zhang, Lu and Lin2012), H. vitripennis (Backus et al., Reference Backus, Habibi, Yan and Ellersieck2005a ), Orosius orientalis (Cicadellidae) (Trebicki et al., Reference Trebicki, Tjajjingii, Harding, Rodoni and Powell2012) and F. occidentalis (Kindt et al., Reference Kindt, Joosten, Peters and Tjallingii2003). Feeding guilds, too, in the hemipteroids vary equally in magnitude.
Thysanoptera, with their relatively short, characteristically asymmetrical mouth parts damage multiple host cells (e.g., epidermal and upper mesophyll cells) during feeding (Moritz, Reference Moritz, Parker, Skinner and Lewis1995; Kirk, Reference Kirk and Lewis1997). Among those belonging to the gall-inducing guild, e.g., many phlaeothripids, one gravid female triggers gall development, although the final gall shape is realized only by the collective feeding impact of all of her offspring (Raman et al., Reference Raman, Ananthakrishnan and Swaminathan1978; Raman, Reference Raman2003). Unfortunately, not much is known on the salivary composition of the Thysanoptera other than amylases, proteases and lipases are implicated in damaging epidermal cells (Raman et al., Reference Raman, Rajadurai, Mani and Balakrishna1999). Pectinases, known in the saliva of Heteroptera, e.g., L. hesperus (Hori, Reference Hori, Schaefer and Panizzi2000) have not yet been demonstrated in the Thysanoptera leaving the question open whether the amylases, proteases and lipases can by themselves perform the function of host-cell degradation. Evolution of thrips from a plesiotypic life style (Mound & Morris, Reference Mound, Morris, Raman, Schaefer and Withers2005) to feeding on leaves (e.g., A. ramakrishnae on M. elengi) and fruits (e.g., Scirtothrips citri; Thripidae, on fruits of different species of Citrus (Rutaceae) on the one hand, and on fungal mycelia and spores (e.g., Allothrips bournieri; Phlaeothripidae), pollen (e.g., Thrips fuscipennis; Thripidae on Rosa sp.; Rosaceae) on the other, and their capability to induce complex galls by modifying the vegetative terminal meristems to develop into large enclosing pouches (e.g., Austrothrips cochinchinensis; Phlaeothripidae on Calycopteris floribundus; Combretaceae) indicate that the enzyme machinery in thrips saliva – details still to be determined – is highly varied. What can be determined is that the feeding action of the Thysanoptera is unwieldy compared with that of the Sternorrhyncha, but is vaguely similar to the majority of the Auchenorrhyncha.
Both Auchenorrhyncha and Sternnorrhyncha are phytophagous. The Sternorhyncha feeds on plant sap, from leaves (e.g., D. citri), stems (e.g., K. lacca) and roots (e.g., Pemphigus betae; Aphididae). The adults of a majority of the Aphidoidea, which are obligate-phloem feeders, seldom rely on surface signals (Powell & Hardie, Reference Powell and Hardie2000; Powell et al., Reference Powell, Tosh and Hardie2006). Their stylet pathway is intercellular, but they puncture mesophyll cells randomly along the path during probing and tasting. Adults of the Aleyrodoidea, which essentially feed on phloem, occasionally ingest xylem sap and, contrary to that found in the Aphidoidea, make fewer attempts probing and tasting. They, thus, inflict fewer intracellular punctures in the mesophyll cells (Walling, Reference Walling2008; Stafford et al., Reference Stafford, Gregory and Ullman2012). A majority of enzymes characterized in the Sternorrhyncha is in different Aphidoidea and have been shown to perform varied functions such as degradation of cells by amylases and cellulases, detoxifying plant-defence compounds by peroxidases and phosphatases and the proteins responsible in plate formation during cell divisions (e.g., actin), and binding mannans during host-plant metabolism (e.g., ficolin), whereas in the Aleyrodoidea, the key enzymes are phosphatases (Funk, Reference Funk2001). Other Sternorrhyncha insert their stylets intercellularly but, similar to the Aphidoidea probe and taste host parenchyma before accessing phloem (Raman & Takagi, Reference Raman and Takagi1992; Gullan et al., Reference Gullan, Miller, Cook, Raman, Schaefer and Withers2005). The Psylloidea feed either on phloem (D. citri; Bonani et al., Reference Bonani, Fereres, Garzo, Miranda, Appezzato-da-Gloria and Lopes2010; D. truncata; Balakrishna & Raman, Reference Balakrishna and Raman1992) or on xylem (Bactericera cockerelli; Triozidae; Butler et al., Reference Butler, Walker and Trumble2012). Compared with the probing—tasting and feeding behaviour of the Auchenorrhyncha and the Heteroptera, the Sternorrhyncha inflict less mechanical damage but, due to their innate salivary chemistry, they alter the physiology of the host plant, such as aggravated transcriptomic changes (De Vos et al., Reference De Vos, Van Oosten, Van Poecke, Van Pelt, Pozo, Mueller, Buchala, Métraux, Van Loon, Dicke and Pieterse2005) and gall induction (Raman, Reference Raman2011). Presence of salivary sheath helps the sternorrhynchan insects to avoid apoplastic plant-defence compounds and enzymes such as peroxidases and phosphatases simultaneously enabling in detoxifying the encountered plant-defence compounds. Gall induction capability is an extreme capability in modifying the host-organ morphology among the Sternorrhyncha. This may be due to the occurrence of specific proteins such as α-dihydroxy-glucohydrolase (Miles, Reference Miles1999) during feeding indirectly on phloem sap. Similarly the presence of Ca++-binding proteins in the saliva of the Sternorrhyncha, could be a facility for their phloem feeding in reducing the occlusion of sieve-tube elements, which would occur as an induced defence response in plants.
The Auchenorrhyncha, on the other hand, feed on plant sap, but a few species belonging to the Cercopoidea, Membracoidea, Cicadoidea feed on fungal mycelia and moss thalli (Nickel, Reference Nickel2003). Nevertheless, the feeding process among the Auchenorrhyncha is not as subtly developed as it has in the Sternorrhyncha, because many of the adult Auchenorrhyncha damage phloem tissue by their stylet bundles of larger dimensions than those of the Sternorrhyncha. This habit – obviously – costs them immensely in terms of the energy spent, since the damaged phloem cannot respond to their feeding action with adequate subcellular pressure (Backus, Reference Backus, Nault and Rodriguez1985); consequently these Auchenorrhyncha have to spend more energy to extract the preferred quantities of phloem sap. While feeding on xylem against negative pressure, the cibarial muscles play a key role (Dugravot et al., Reference Dugravot, Backus, Reardon and Miller2008). Membracoidea and Fulgoroidea insert stylets intracellularly and feed on xylem (e.g., P. spumarius, Cercopidae) (Crews et al., Reference Crews, Mccully, Canny, Huang and Ling1998), mesophyll (e.g., Empoasca fabae; Cicadellidae) (Hunter & Backus, Reference Hunter and Backus1989), and phloem (e.g., N. lugens) (He et al., Reference He, Chen, Chen, Zhang, Chen, Shenc and Zhud2011). Activity of β-1, 4-endoglucanase in saliva of H. vitripennis, which is a cellulose-degrading enzyme, demonstrates the relevance of such proteins in pathogen transfer (Backus et al., Reference Backus, Andrews, Shugart, Greve, Labavitch and Alhaddad2012). However, the ‘unique’ gall-induction capability by the instar I nymphs of Scenergates viridis (Cicadellidae) on the leaves of Alhagi maurorum (Fabaceae) (Rakitov & Appel, Reference Rakitov and Appel2012) illustrates the probability of specific salivary enzymes, not yet characterized. Not much is known about the salivary enzymes of the Auchenorrhyncha. However, the presence of most of cell-degrading enzymes, such as amylases, lipases and trypsin, with detoxifying enzymes such as superoxide dismutase is known from E. fabae (DeLay et al., Reference DeLay, Mamidala, Wijeratne, Wijeratne, Mittapalli, Wang and Lamp2012). With our current knowledge, we can only infer that similar to that in the Heteroptera, cell-degrading enzymes play a critical role in the feeding process of the Auchenorrhyncha.
Cell-rupturing feeding behaviour reinforces the plentiful occurrence of cell-degrading enzymes (amylases, proteases and pectinases) in the Heteroptera. Among these insects, the labium plays a key role in dabbing before site selection and stylet insertion. By secreting pectinases, the heteropterans (e.g., Leptocorisa chinensis Alydidae; Ishizaki et al., Reference Ishizaki, Yasuda and Watanabe2007) macerate the plant tissue. Heteropteran feeding substrates are staggeringly diverse: detritus (e.g., Corixidae), other arthropods (e.g., Anthocoridae) and blood of mammals, birds and reptiles (e.g., Reduviidae), and this renders a generalization difficult. Facultatively carnivorous heteropterans (e.g., Pentatomidae, Miridae) feed on nitrogen-rich plants parts, such as fruits and seeds, as well as on animals (Schaefer, Reference Schaefer and Raman1997; Schaefer & Panizzi, Reference Schaefer, Panizzi, Schaefer and Panizzi2000). In comparison with the saliva of the Culicidae (Diptera) that includes trypsin active at an alkaline pH, the blood-feeding Heteroptera include cathepsin-like proteinases active at acidic pH. This argument illustrates that the salivary physiology varies with insect groups although their feeding guilds are similar, which possibly has been driven by different evolutionary pathways (Lehane, Reference Lehane and Lehane2005). Extending on this, the incidence of hydrolases in the Heteroptera stands as a useful example: in a majority of the plant-feeding heteropterans, hydrolases occur as the predominant enzyme machinery, whereas in the in the blood-feeding heteropterans, hydrolases occur negligibly (see table 1). While feeding on leaves (e.g., Anasa tristis; Coreidae, on Citrullus lanatus; Cucurbitaceae), fruits (e.g., Campylomma verbasci; Miridae, on Malus domestica; Rosaceae) and seeds (e.g., O. fasciatus; Lygaeidae on Asclepias sp. Asclepiadaceae), these Heteroptera employ cell-rupturing mechanism, thus acquiring greater energy levels quickly (Hori, Reference Hori, Shorthouse and Rohfritch1992). Due to a higher degree of mechanical damage and different salivary elicitors, L. hesperus and Nezara viridula (Pentatomidae) influence plant-volatile production through damage to Gossypium hirsutum (Malvaceae) and Zea mays (Poaceae), respectively (Williams et al., Reference Williams, Rodriguez-Saona, Paré and Crafts-Brandner2005).
The ability of the hemipteroids to transmit pathogens is closely linked to the feeding strategy and the nature of the target tissue. Sternorrhyncha and Auchenorrhyncha are the more efficient vectors of microbial pathogens than the Heteroptera (Mitchell, Reference Mitchell2004). Enzyme ‘weaponry’ decides the transmission modes of microbial pathogens (Hogenhout et al., Reference Hogenhout, Ammar, Whitfield and Redinbaugh2008b ); they also act as elicitors for induced-plant responses. For instance, glucosidase known in Pieris brassicae (Tumlinson & Lait, Reference Tumlinson and Lait2005) and glucose oxidase known in Heliothis zea (Lepidoptera: Noctuidae) (Morkunas et al., Reference Morkunas, van Chung and Gabrys2011) trigger emission of volatiles (e.g., volicitin: N-(17-hydroxylinolenoyl)-l-glutamine) in their respective host plants. Given that glucosidases and glucose oxidase are known in the saliva of different hemipteroids, their role in eliciting plant volatiles cannot be overlooked. Induction of plant volatiles due to the feeding effect of L. hesperus and N. viridula could be the starting point of our need to understand the importance of elicitors in hemipteroid saliva (Williams et al., Reference Williams, Rodriguez-Saona, Paré and Crafts-Brandner2005). Feeding behaviour of the hemipteroids immensely influences the nature and chemistry of enzymes of the salivary glands and gut. A majority of hydrolyses are known from the Heteroptera and they are generally known for their function in cell degradation; but their incidence appears to trigger gall induction (e.g., amylases and lipases; Raman et al., 1999; Miles, Reference Miles1999), in manipulating the plants osmotic pressure to elicit mobilization of compounds (e.g., sucrases; Miles & Taylor; Taylor & Miles, Reference Taylor and Miles1994), and in detoxifying plant-defence compounds (e.g., catalases; Miles, Reference Miles1999; Ni et al., Reference Ni, Quisenberry, Pornkulwat, Figarola, Skoda and Foster2000).
This review has attempted to consolidate the available information on the salivary enzymology of the hemipteroids. Hemipteroids, among Insecta, are a complex group showing a range of adaptations to different habitats and with an equally complex feeding guilds and behaviours. A better understanding of enzymatic proteins and effectors should shed better light on the interrelationship between different hemipteroid groups, their feeding behaviour and various other aspects of insect–plant interaction.
Acknolwedgements
We thank Elaine Backus (USDA—ARS, Pacific West, Parlier, California, USA), Scott J. Nicholson (USDA—ARS, Plant Science Research Laboratory, Stillwater, Oklahoma, USA), Donald Miller (Department of Biological Sciences, California State University, Chico, California, USA) and Soundararajan Madhavan (The Beadle Center, University of Nebraska, Lincoln, USA) for their helpful reviews and insightful remarks.
