Abstract
Termites depend on their gut microbes for digestion of complex polysaccharides of wood into simpler molecules. Cellulose is a major polymeric carbohydrate present in the wood which is broken down to simpler byproducts through metabolic steps by the hindgut microbes. Termite gut microbes also produce gases during the cellulose degradation process, of which methane is the major product. Gut microbes belong to three major groups, namely, bacteria, archaea and protozoa. They show a mutualistic relationship and typically convert 95% of cellulose into simple sugars within 24 h. More than 200 species of microbes form this community, producing different types of wood-busting enzymes, mainly cellulases, cellubiases, hemicellulases, glucosidases and gluconases, during wood degradation. Studies suggest that lower termites utilize both endogenous and protozoal enzymes for cellulose digestion, while higher termites acquire enzymes from their diet instead of protozoal enzymes. Some termite species change their feeding habits with seasonal variations. These affect gut microbes population and therefore are responsible for enhancing their survival under changed environmental conditions.
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4.1 Introduction
Termites are among the most efficacious groups of insects on Earth that colonize most landmasses except Antarctica, with colonies assorting from a couple of hundred to several million individuals (Cranshaw 2013). Lower termites predominantly harbour species of oxymonads, trichomonads and hypermastigote flagellates as symbionts in the paunch of hindgut, that aid in wood ingestion. Higher termites (Termitidae) consist of about 75% of all termite species and show instead low protozoan populations (Sanderson 1996). The symbiotic associations in termites are crucial due to the presence of three groups of microorganisms. These are mainly bacteria, protozoa (inhabiting the hindgut) and fungi cultivated as ‘fungus gardens’ or ‘fungus combs’ in some species (Darlington 1994). Most termite species reside in the tropical, subtropical and warmer temperate zones of the world and subsist on a diet rich in cellulose either in the form of living or dead wood, woody tissue of plants, humus or dung (Higashi and Abe 1997). They, therefore, play an important role as terrestrial decomposers (Brune and Friedrich 2000).
4.2 Termite Gut
The digestive system of termites consists of three main parts: the foregut, midgut and hindgut (Ptacek et al. 2013; Noirot and Noirot-Timothee 1969). Foregut includes the crop and muscular gizzard and contributes in the secretion of digestive enzymes as well as absorption of soluble nutrients. Hindgut also aids in digestion and absorption of nutrients (Breznak 1994). It may be divided into five successive segments, namely: proctodeal segment, enteric valve, paunch (abundant in symbiotic microorganisms), colon and rectum. Enteric valve prevents return of paunch contents back into the midgut or foregut. Malpighian tubules enter the gut at the junction of the midgut and first proctodeal segment, just in front of the proctodeal valve. Some higher termites have an elongated midgut known as the ‘mixed segment’ or mesenteron, due to prolongation of one of the intestinal tube faces (Fig. 4.1).
Gut segments of a typical termite (worker) (Ptacek et al. 2013)
The hindgut, being the largest part of the intestinal tract, is anaerobic. Here the symbiotic gut microbes depolymerize cellulose and hemicellulose and ferment resulting carbohydrates into short-chain fatty acids, which are later absorbed and oxidized by the host (Brune and Friedrich 2000) (Fig. 4.2). Guts of termites maintain anoxic conditions due to their steep oxygen gradients at the oxic-anoxic interface, which drives a continuous influx of O2 into the gut. According to microsensor studies, O2 can travel at a rate of 50–200 μm in the gut, therefore creating a microoxic periphery around an anoxic centre. In the lower termite Reticulitermes flavipes, radiotracer studies show an influx of O2 through gut epithelium which gets reduced at the periphery of the hindgut (Boga and Brune 2003). The microbe-filled paunch region of lower and higher termites has a microoxic periphery where pH and redox potential undergo significant transitions along the guts anterior to posterior axis (Bignell 1994). However, the paunch region measurements are circumneutral and anoxic with a comparatively low redox potential of −150 to -250 mV. The pH values of hindguts also show neutrality from 6.2 to 7.6, but it can undergo alterations with environment changes (Abe et al. 2000).
Collaboration of the host and symbionts in lignocellulose digestion in R. flavipes (a) and different components of the digestive tract (b) showing the oesophagus (E), salivary glands (SG), foregut (FG), midgut (MG), Malpighian tubules, (MT) hindgut (HG) and rectum (R) (Scharf et al. 2011)
Soil-feeding termites exhibit extreme pH changes along the gut axis with pH values as high as 11. This is an evolutionary adaptation to their diets, rich in tannins or other polyphenolic constituents as it prevents precipitation of digestive enzymes and enhances solubility of dietary proteins (Breznak 1994). The high gut alkalinity separates organic nutrients from the organo-mineral aggregates, as soil-feeding termite ingests organic matter from soil (Kappler 1999). The guts of the wood-feeding and soil-feeding termites are characterized by noticeable axial dynamics of O2 and H2 partial pressure followed by intestinal pH. These differences bring uneven distribution of H2 production, methanogenesis and reductive acetogenesis by gut microbes (Brune and Friedrich 2000).
Numerous prokaryotes have been isolated in pure culture, and many of them are unique to the termite gut habitat (Graber et al. 2004), including oxygen reduction by anaerobic bacteria, such as lactic acid bacteria (Bauer et al. 2000), homoacetogenic bacteria (Boga and Brune 2003) and sulphate-reducing bacteria (Frohlich et al. 1999). Lower and higher termites differ in their gut microflora. Bacteria and archaea inhabit guts of both higher and lower termites. However, cellulolytic flagellates participating in wood digestion are present only in lower termites (Dietrich et al. 2014) that without them starve (Breznak and Brune 1994). Some higher termites exhibit symbiosis with the fungus Termitomyces, or Bacteroidetes and Firmicutes, while Spirochaetes and Fibrobacteres outnumber the guts of other higher termites with a cellulose-rich diet (He et al. 2013; Breznak 1994).
The ‘mixed segment’ present only in higher termites is composed of a mesenteric epithelium which occupies half of the gut wall, while the remaining area is covered by the proctodeal epithelium. The mixed segment has an elevated pH (Brune and Kuhl 1996) along with an alkaline fluid rich in potassium ions. Symbiotic bacteria occupy the mesenteric side of the mixed segment (Tokuda et al. 1997). A peritrophic membrane separates bacteria from wood particles present in the lumen as shown by in situ hybridization and electron microscopy studies. Due to their non-uniform distribution around the peritrophic membrane, the lumen is not a nutritional source for these bacteria. Electron microscopy indicates a close relationship between bacteria and mesenteric epithelium suggesting that symbiotic bacteria utilize some substances secreted by the mesenteric tissue of the mixed segment. Midgut cells are very quickly replaced in cockroaches (Dow 1986) and termites via endocytosis of the neighbouring epithelial cells, which are later digested by phagolysosomes (Yamaoka and Nagatani 1980). Similar phagosomes have been observed throughout the midgut columnar cells, including the mixed segment of Nasutitermes takasagoensis where the symbionts trapped among the microvilli of old cells can be endocytosed by young columnar cells and digested along with old cells. Since bacteria cannot pass through the peritrophic membrane, it is believed that they infect termites only when their peritrophic membrane is intact (Tellam et al. 1999). Phylogeny and distribution studies of the symbiotic bacteria dwelling in the mixed segment suggest that these bacteria have a significant role in the termites gut physiology. Microflora of the mixed segments of soil-feeding and wood-feeding termites are different; therefore, it is possible that these symbionts bear a close relationship with the feeding habits, gut physiology and phylogeny of their hosts (Tokuda et al. 2000).
The neutral pH value of the midgut of the wood-eating higher termite N. nigriceps increases in the mixed segment and reaches to 10.23 ± 0.46 in the first proctodeal segment, while a decrease in oxygen concentration in the mixed segment occurs, which ultimately becomes zero in the first proctodeal segment (Brune et al. 1995). The space between the gut wall and peritrophic membrane of the mixed segment is termed the ‘ectoperitrophic space’ and is the shelter home for the bacterial species. Cocci and rods have been identified in the wood-eating termite N. exitiosus (Czolij et al. 1985), while spirochaetes and actinomycete-like bacteria populate the guts of the soil-feeding termites Cubitermes severus and Procubitermes aburiensis (Bignell and Eggleton 2000). The symbiotic bacteria present in the mixed segment resemble the Clostridium group but differ from the bacteria of wood-eating lower termite Reticulitermes speratus (Ohkuma and Kudo 1996) as well as from the clostridia isolated from higher termites (Hethener et al. 1992). However, no low-GC-content Gram-positive bacteria have been yet discovered from the gut of the drywood-eating lower termite Cryptotermes domesticus (Kudo et al. 1998). Clostridium species are usually obligate anaerobes, while some are facultative anaerobes appearing as rods, short rods or cocci (Rieu-Lesme et al. 1996). Some of them are able to degrade polysaccharides by producing acetone, alcohol, acetate, lactate, CO2 and hydrogen (Rainey et al. 1996), while others can ferment nitrogenous or lipidic compounds playing, therefore, an important role in the nutritional physiology of termites (Hethener et al. 1992).
4.3 Termite Gut Microbiome
Termite gut unveils one of the most complex microbial communities entailing diverse microbes from the three domains of life (Bacteria, Archaea and Eukarya) (Ohkuma and Kudo 1996). Detritivorous subsocial cockroaches believed as termite ancestors (Inward et al. 2007) can digest wood with the help of cellulolytic flagellates (Engel et al. 2009; Ohkuma et al. 2009). The evolution of termites coincides with three major events associated with dietary diversification and microbial symbiosis: (i) cellulolytic, flagellate protozoa inhabit the hindguts of all five lower termite families except Termitidae, (ii) an ectosymbiotic relationship with basidiomycete fungi in the litter-feeding Macrotermitinae and (iii) independent evolution of soil-feeding behaviour among the three higher termite subfamilies Nasutitermitinae, Apicotermitinae and Termitinae (Ohkuma and Brune 2011). The role of the cellulolytic flagellates in lower termites is well known, but the exact functions of uncultivated bacterial symbionts in digestion, especially in flagellate-lacking higher termites, need further research (Hongoh 2011). Subterranean termites primarily feed on woody tissue containing lignocellulose as their diets are deficient in vitamins and essential components for protein and fat synthesis. These termites produce their own cellulases (Inoue et al. 1997), but in order to meet their nutritional requisites, the association with symbionts becomes crucial and is achieved by coprophagy and trophallaxis (Grimaldi 2001). These symbionts display a mutualistic beneficial relationship by augmenting nutrients and energy to the termites and by gaining a stable food supply as well as protection under the constant gut environment (Nalepa et al. 2001). Each worker termite acquires an initial inoculum of symbionts from the parents or nest mates, right after hatching as well as after each moult (Thorne 1997).
The gut microbiota varies markedly among termite species comprising many unique phylogenetic clusters (Hongoh et al. 2006). The presence of microbes in the guts of cockroaches (Schauer et al. 2012) and termites (Noda et al. 2009) suggests that the bacterial microbiota of both originated from a common dictyopteran ancestor (Dietrich et al. 2014) (Table 4.1). Termites rely predominantly upon the symbiotic protozoa (metamonads) and flagellate protists for cellulose digestion as well as absorption of the end products (Ikeda-Ohtsubo and Brune 2009). Gut protozoa, such as Trichonympha, further rely on the symbiotic bacteria rooted on their body surfaces for the production of the necessary digestive enzymes. Termites of the family Termitidae, although able to produce their own cellulases, depend on bacteria as well (Li et al. 2013).
4.3.1 Bacteria
Microscopic cell counts approximate presence of 106 to 107 bacteria in comparison with the protozoa (4 · 104) in the gut of R. flavipes where the prokaryote groups principally consist of methanogens (Archaea), Proteobacteria, Actinobacteria, Firmicutes, Bacteroidetes and Spirochaeta (Eubacteria) (O’Brien and Slaytor 1982). Three species of methanogenic Archaea placed in the genus Methanobrevibacter have already been cultured from R. flavipes gut (Leadbetter et al. 1998). The culture-independent sequencing of archaeal 16S rRNA genes also indicates the presence of methanogens in the gut of R. speratus (Shinzato et al. 2001). Some methanogen species are both ecto- and endosymbionts of protozoa (Tokura et al. 2000). Methanogenesis is followed by acetogenesis in the wood-feeding termites and acetogenic bacteria compete with the methanogens for hydrogen (Brauman et al. 1992).
The hindgut of xylophagous lower termite harbours both protozoa and bacteria. However, due to their mutualistic relationship with termites and cellulose-degrading activity, more research has been done on protozoa, although prokaryotes are also important for the termites’ survival (Mauldin 1977). The presence of bacteria is necessary for the persistence of protozoa as well as for normal termite nutrition (Breznak 1975). Nitrogen fixation by some bacteria shows a role in nitrogen utilization by termites, as many species exist in intimate physical association with the gut epithelium, thereby reflecting biochemical interactions (Thayer 1976). An anaerobic, cellulolytic actinomycete (Micromonospora propionici) from the guts of Amitermes minimus was isolated, but it turned out to be of limited importance in situ (Hungate 1946). Anacanthotermes ahngerianus and A. turkestanicus harbour 1.3 · 106–4.3 · 109 viable bacterial cells per hindgut, depending on the developmental stage or caste of termite (Husseneder 2010; Krasil’nikov and Satdykov 1970).
The intestinal bacterial microflora of R. speratus contains taxa related to the enteric bacteria, such as Enterobacter, Citrobacter, Desulfovibrio, Treponema, Bacteroides and Clostridium which are either strict or facultative anaerobes and have been frequently isolated from animal intestines (Brauman et al. 1992). Apart from Citrobacter freundii, Enterobacter agglomerans, Clostridium and Desulfovibrio spp. also partake in nitrogen fixation in several termite species (Fuhrman et al. 1993). Species of Sporomusa, Acetonema and Clostridium act as CO2-reducing acetogens within the termite guts (Kane and Breznak 1991). Bacteroides species (Bacteroides termitidis) participates in the uric acid metabolism (Potrikus and Breznak 1980). The termite Desulfovibrio cluster points out the presence of significant quantity of the sulphate-reducing bacteria in situ within gut, functioning in interspecies hydrogen (H2) transfer as H2 donors by utilizing small organic compounds. These include, for example, pyruvate, lactate and sugar monomers as oxidizable electron donors in comparison to the H2 acceptors oxidizing sulphur compounds.
Streptococcus sp. occurs as the major bacterial species in several termites (Breznak and Brune 1994). The gut microbes enrich the termites with the required amount of carbon, nitrogen and energy to such an extent that survival of termites becomes almost impossible without them (Brune and Friedrich 2000). A predominance of five bacterial groups, viz., Cytophaga-Flexibacter-Bacteroides (CFB) group, low-G + C Gram-positive bacteria, Proteobacteria, Spirochaeta and the newly discovered ‘termite group I’ (TG I), has been found in many species (Hongoh et al. 2003). However, spirochaetes occur more in the gut of R. speratus as compared to the gut contents of Nasutitermes lujae, followed by the Bacteroides-related, clostridial and TG I clones which form the second most dominant group (Husseneder 2010; Paster et al. 1996).
Spirochaetes comprise a monophyletic group of motile bacteria with a characteristic spiral or wavy shape (Paster et al. 1991) and a coiled or undulate protoplasmic cylinder confined by a cell wall-cytoplasmic membrane complex, containing genomic DNA, ribosomes and other cytoplasmic constituents. An outer membranous sheath surrounds the protoplasmic cylinder. Between the outer sheath and the protoplasmic cylinder one or more periplasmic flagella are present (Canale-Parola 1984), functioning as organelles of motility (Charon et al. 1992). Spirochaetes are widely distributed in nature for example, as free-living forms in freshwater, marine and hypersaline waters or associated with invertebrate and vertebrate hosts, through commensalism, mutualism and parasitism. Spirochaetes were first documented by Joseph Leidy (1874–1881, 1877) in the hindgut of the eastern subterranean termite, Termes (now Reticulitermes) flavipes. Spirochaetes account for up to 50% of all prokaryotes in the hindgut of some termites and about a dozen of different morphological types can be identified on the basis of cell length, width, wavelength and amplitude or pitch (Paster et al. 1996; Leadbetter et al. 1999).
The nucleotide sequence of MDS1clone of the Australian termite, Mastotermes darwiniensis (family Mastotermitidae) (Berchtold et al. 1994), R. speratus (family Rhinotermitidae) (Ohkuma and Kudo 1996) and Cryptotermes domesticus (family Kalotermitidae) (Ohkuma 1998) indicates that the spirochaetes residing within the guts of these species are either anaerobes or microaerophiles (Breznak 1994). Termite gut spirochaetes are placed into two phylogenetic clusters namely Treponema clusters I and II. Cluster I contains diverse phylotypes of the gut spirochaetes containing the strains isolated from termite gut, while cluster II is smaller, belonging to the Treponema bryantii subgroup.
Spirochaetes can exist freely in the gut fluid or are attached as ectosymbionts on the cell surface of gut protists (Iida et al. 2000). They have specialized attachment sites on the protists as shown by ultrastructural observations (Bloodgood and Fitzharris 1976). Treponema strains ZAS-1 and ZAS-2 have been isolated from the dampwood termite Zootermopsis angusticollis (family Termopsidae) (Leadbetter et al. 1999; Breznak 2002). Recently, ectosymbiotic spirochaetes (Treponema sp. strains ZAS-1 and ZAS-2) associated with Zootermopsis angusticollis have been clustered with the major cluster I sequences, with function as H2 and CO2 consumers by environment absorbance from inside the protist. The other major ectosymbiotic spirochaetes (cluster II) placed in the T. bryantii subgroup can enhance the cellulolytic activity of other microbes (Ohkuma and Kudo 1996). Members of cluster II are ectosymbiotic spirochaetes of oxymonad protists as identified in the termite species R. speratus, Hodotermopsis sjoestedti and Neotermes. However, various species of devescovinid (Devescovina sp.), calonymphid and hypermastigote protists (Holomastigotoides mirabile in Coptotermes formosanus), in addition to the oxymonad protists, also harbour dense populations of ectosymbiotic spirochaetes (Iida et al. 2000).
The unique ultrastructure of spirochaete-protist attachment sites is known for several protists of termites and wood-feeding cockroaches, and two types of attachment structures have been identified in the symbiotic protists of R. flavipes, R. tibialis and Cryptocercus punctulatus. One structure is a narrow nose-like appendage making direct contact with the plasma membrane of the host cell, while the other is the flattened end of the spirochaete in contact with the protistan membrane, with a thick layer of electron dense material. The ectosymbiotic spirochaetes are involved in the maintenance of ‘motility symbiosis’ (Kitade et al. 1997). The termite gut bacteria are deliberated to be vertically transmitted from generation to generation via proctodeal trophallaxis known for the gut symbiotic protists in lower termites (Inoue et al. 2000). However, horizontal transfer among congeneric termites is also known for the bacterial community profiles of Microcerotermes species M1 and M2. Ambient temperature, food quality and humidity also affect variation in bacterial gut microbiota within congeneric termites (Donovan et al. 2004).
Gut epithelium is also an important habitat for the dense colonization of the symbionts. Methanogenic archaea utterly occupy the gut wall of R. flavipes, while in other termite species, they are located within some protist cells (Leadbetter and Breznak 1996). The methanogenic species present on the gut wall, as well as within the protists cells in R. speratus and Hodotermopsis sjoestedti, are phylogenetically different (Tokura et al. 2000). This is due to the radical changes in the physico-chemical conditions, especially fluctuations of oxygen and hydrogen partial pressures, found inside the gut (Brune and Friedrich 2000).
Bacterial communities also diversify at the phylum or phylotype levels within the gut wall and gut luminal fractions, where bacteria residing on the gut wall are believed to be more dynamic than those populating the gut lumen (Noda et al. 2003). Divergence index, rarefaction curve and Chao1 richness estimators of biodiversity also confirm that a wide variety of bacteria are able to colonize the gut wall, directly or indirectly. However, the populations on the gut wall are five times lower in comparison to those on the gut luminal fraction (Iida et al. 2000).
Bacterial groups densely colonizing the gut wall fraction include species of Actinobacteria, Bacteroidales, Clostridiales and Lactococcus. Many of them are nonmotile species. However, gut lumen shows abundance of highly motile Spirochaetes and Desulfovibrio species (Wenzel et al. 2003). In order to resist fluid flow within the gut, bacterial symbionts also subordinate with the protists, for example, spirochaetes and TG1 bacteria that form associations with the gut protists. The central part of the gut is anoxic as oxygen penetrates the gut via its wall, thereby maintaining a steep oxygen gradient near the gut wall (Brune and Friedrich 2000). This affects the gut metabolism as already displayed by the isolated lactic acid (Tholen and Brune 1997) or sulphate-reducing bacterial strains (Kuhnigk et al. 1996) highlighting the presence of strict aerobic communities inside the gut (Moriya et al. 2003).
Bacteroidales endosymbionts dominate the gut bacterial community of C. formosanus and Pseudotrichonympha sp. by living intracellularly within eukaryotic cells, while few Bacteroidetes or the Cytophaga-Flavobacterium-Bacteroides (CFB) phylum bacteria occur as intracellular endosymbionts of eukaryotic cells (Bandi et al. 1995). Intracellular Blattabacterium sp. dwell within the bacteriocytes of fat bodies of cockroaches and Mastotermes darwiniensis. Intracellular acanthamoebae bacteria affiliated as Flavobacterium or ‘Candidatus Amoebophilus asiaticus’ resemble with the endosymbionts of a tick and a whitefly through a novel phylogenetic lineage (Horn et al. 2001).
The ectosymbiotic Bacteroidales carry a typical Gram-negative cell wall along with inner and outer membranes. However, endosymbionts have lost the cell wall as they localize within the cytoplasm of the host protist. Ectosymbionts colonizing the protist cell surface are incorporated into vacuoles of the host cytoplasm, as structures required for attachment with the host cell membrane occur in the vacuoles (Stingl et al. 2004).
Endomicrobia (formerly termite group 1) represent a deep branching clade of the uncultivated bacteria of the phylum Elusimicrobia with single isolate, Elusimicrobium minutum which occurs as intracellular symbiont of termite gut flagellates and transferred vertically. Many genes of E. proavitum are highly similar to the closely related ‘Candidatus Endomicrobium trichonymphae’ strain Rs-D17 (Brune 2014).
Bacteria of the Cytophaga/Flexibacter/Bacteroides (CFB) phylum predominate the guts of Macrotermes michaelseni, fungus-cultivating (Macrotermes gilvus) (Hongoh et al. 2006), wood-feeding (Ohkuma et al. 2004) and soil-feeding termites (Schmitt-Wagner et al. 2003). Many CFB phylum bacteria are capable of degrading plant fibres and proteins which form the ingredients of the termite diet (Shah 1992). Proteobacteria, Desulfovibrio sp., Escherichia hermannii and E. senegalensis also colonize the gut of M. gilvus instead of CFB phylum members (Hongoh et al. 2006). Desulfovibrio spp. are strict anaerobes partaking in sulphate reduction and nitrogen fixing. Clostridia are also present in the mixed segment of Nasutitermes takasagoensis (Shiraki) (Tokuda et al. 2000) and M. gilvus (Hongoh et al. 2006) where they degrade polysaccharides and produce acetone, alcohol, acetate, lactate, CO2 and hydrogen (Chen 1995). However, Clostridium mayombei present in the mixed segment of Cubitermes speciosus brings about acetogenesis (Kane et al. 1991). The intestinal bacteria in termites aid in the production of short-chain fatty acids from carbohydrates or amino acid synthesis, just like they do in humans (Cummings and Macfarlane 1997).
The Anaerobaculum-Thermoanarovibrio, also denoted as the ‘Synergistes group’, shows coevolution with termites although the exact role of these members in the termite guts is unknown. It is believed that they are anaerobic amino acid degraders participating in the amino acid turnover in natural anaerobic ecosystems (Godon et al. 2005). The ‘Synergistes’ strains Aminomonas paucivorans (Baena et al. 1999a), Thermoanaerovibrio acidaminovorans (Baena et al. 1999b), Dethiosulfovibrio sp. (Surkov et al. 2001), Aminobacterium mobile (Baena et al. 2000) and Aminobacterium colombiense (Baena et al. 1998), for example, degrade amino acids, but some strains such as Thermanaerovibrio velox (Zavarzina et al. 2000) and Anaerobaculum sp. can utilize carbohydrates as well (Rees et al. 1997).
The population of spirochaete-like cells in the gut contents of fungus-cultivating termites is generally low, ranging 2–3% in Pseudacanthotermes, 6–10% in Odontotermes sp. (Liu et al. 2013), 11–19% in Ancistrotermes and 22–29% in Microtermes sp. (Makonde et al. 2013).
Firmicutes and Bacteroidetes, placed in the Treponema I lineage, form the major fraction of the bacterial community in the Macrotermitinae (Dietrich et al. 2014) although their actual is not clear, but several of them resemble isolates from the lower termite hindguts. These species either partake in fermentation of mono- and oligosaccharides by producing acetate and other products or are homoacetogenic (Droge et al. 2008). Uncultured Treponema lineages in higher termites can carry out reductive acetogenesis from H2 + CO2 (Warnecke et al. 2007). Methanogenesis dominates the reductive acetogenesis as hydrogen sink in Macrotermitinae. Therefore, Treponema shows a faint representation which is consistent with the reduced acetogenesis observed in the guts of Macrotermes mulleri, Postelectrotermes militaris and Pseudacanthotermes spiniger (Brauman et al. 1992). The reason behind this is, however, unknown (Hongoh 2011). Variation in the microbiotas of Macrotermitinae reflects their ecological differences which is governed by the plant diet and Termitomyces association.
Diet shapes the gut communities of wood-feeding termites (Huang et al. 2013), but such variation in the Macrotermitinae is generally not very clearly understood (Hongoh 2010). Macrotermes spp., with the exception of Macrotermes malaccensis, are primarily leaf litter feeders (Hyodo et al. 2000). Odontotermes and Ancistrotermes sp. predominantly feed on wood, and P. militaris feeds on both leaf litter and wood (Hyodo et al. 2003). The role of Termitomyces also varies among different termite hosts (Nobre et al. 2011). Although this needs more attention, some termite species mainly gain access to cellulose via the lignolytic activity of Termitomyces (Hyodo et al. 2000). Some species exploit Termitomyces as a protein-rich food source (Hyodo et al. 2003), while other species extract cellulases and xylanases from Termitomyces for the decomposition of plant substrate (Rouland et al. 1991). Therefore, differences in gut communities among the termite hosts may be the result of specific diets or symbiont functions.
The role of the bacterial community in lignocellulose breakdown in the Macrotermitinae is not fully elucidated. Metagenomic studies suggest their possible contributions in the cellulose digestion in the wood-feeding termites (He et al. 2013). Glycosyl hydrolases showing cellulase activity in the P3 lumen metagenome of the wood-feeding Nasutitermes sp. are now taxonomically confined to the phyla Fibrobacteres and Spirochaetes (Warnecke et al. 2007). These phyla, along with the TG3 phylum, show high abundance in wood-feeding termites of the subfamilies Nasutitermitinae (Mikaelyan et al. 2014) and Termitinae (Dietrich et al. 2014).
4.3.2 Protozoa
The symbiotic associations between termites and their gut microorganisms are continuously under study since the beginning of the century, when earlier work was centred on the intestinal protozoa of lower termites and their role in digestion. Currently more attention is given indeed to the bacterial and archaeal populations, their metabolic activities, structure and function.
Intestinal protozoa provide shelter to the prokaryotes in the hindgut of lower termites by forming intimate associations with them. Previously, it was difficult to obtain protozoan cultures in vitro and maintain pure cultures of prokaryotic symbionts, so only a morphological description of the different associations was known. With advanced molecular biology tools, informations regarding the symbiotic associations between prokaryotes and termite gut flagellates have now become clear.
Symbiotic flagellates exclusively inhabit the lower termites and closely related cockroaches (Cryptocercus), whereas higher termites largely harbour prokaryotic microbiota. Molecular phylogeny of termites and Cryptocercus clarified that gut protists share a common ancestor (Lo 2003). The beneficial nature of these peculiar symbionts came into light for the first time by the studies of Lespes in 1856 (Leidy 1874–1881), who classified them as parasites, while Cleveland (1926) pointed out that termites cannot live without the gut flagellates. The symbionts heavily populate the hindgut paunch, with a fresh weight that may account for more than 50% of the fresh termite weight (Katzin and Kirby 1939). Phylogenetically, gut flagellates are extremely diverse, and almost 450 distinct species, from about 200 termite species, are into account till date (Yamin 1979).
Termite gut flagellates were earlier considered as primitive, primarily amitochondriate eukaryotes. Recent molecular data indicated that they share two separate eukaryotic lineages and can be placed under three distinct taxa, namely, trichomonads, hypermastigids and oxymonads (Yamin 1979). Phylogenetic studies conducted by 18S rRNA gene sequence analysis confirmed that majority of termite gut flagellates fall within two classes of the phylum Parabasalia, i.e. Trichomonadea and Hypermastigea (Gerbod et al. 2004). On the other hand, the phylogeny of Oxymonadea was under dark for a long time, but comparative sequence analysis indicated that they are a sister taxon of unidentified protists which are now classified in the phylum Loukozoa (Stingl et al. 2005).
Hypermastigea are exclusive termite gut symbionts, while Oxymonadea and Trichomonadea inhabit other habitats as well, such as the intestinal tract or body cavities of other animals, including humans (Cavalier-Smith 2002). Out of the 440 species of amitochondriate protists belonging to Trichomonadida, Hypermastigida and Oxymonadida are exclusively gut symbionts of wood-eating termites (Mastotermitidae, Kalotermitidae, Hodotermitidae, Termopsidae, Rhinotermitidae and Serritermitidae) and the wood-eating roach, Cryptocercus.
Bacterial symbionts associate either on the cell surface or in specific cytoplasm regions of the protozoan, as confirmed by morphology-based descriptions. Trichomonads and other amitochondriate protist symbionts of wood-eating termites and Cryptocercus can be easily identified by the presence of their motility structures (mastigont) while in Parabasalia (trichomonads, hypermastigids) by the parabasal body (Golgi complex). The key characters include number and arrangement of flagella (undulipodia), presence and shape of accessory structures (e.g. costa and cresta) and arrangement of connected microtubular structures, axostyle and pelta. The parabasal body bears a distinct shape as well as size and can branch or spirally coiled around the axostyle. It is connected to the mastigont in the trichomonads and arranged in multiple copies in the hypermastigids. Every termite species inhabits a characteristic community of gut protists. Therefore, in general the protist species are not restricted to one termite species only (Simpson 2006).
The phylum Parabasalia has a monophyletic but complex assemblage of diverse species of flagellated protists characterized by a unique parabasal apparatus (Golgi complex associated with striated fibres), closed mitosis with an external spindle (cryptopleuromitosis) and anaerobic energy-generating organelles (hydrogenosomes) (Brugerolle 2005). On the basis of their morphological characters, more than 80 genera and 400 parabasalid species are now known (Yamin 1979). Most parabasalids occur in the digestive tract of animal hosts either as commensals, parasites or symbionts, whereas the symbiotic parabasalids present in the gut of termites and wood-eating cockroaches have a key role in cellulose digestion (Brune and Ohkuma 2011). This symbiotic relationship between termites and parabasalids also provides an evidence for the evolution of social behaviour in the hosts and also carries an ecological significance for plant litter decomposition in terrestrial ecosystems (Ohkuma 2003).
Mixotricha paradoxa, a protozoan species with multiple bacterial symbionts living in the gut of the Australian termite species, Mastotermes darwiniensis, was first described in 1933 by Australian biologist J.L. Sutherland (Cleveland and Grimstone 1964). Mixotricha bears four anterior flagella which are used for steering rather than for locomotion. Locomotion is performed by cilia-like movements of approximately 250,000 hairlike Treponema spirochaetes attached to the cell surface. Mixotricha also shows the presence of rod-shaped bacterial Bacteroides-like species, arranged in an ordered pattern on the cell surface. Both these bacterial types (Spirochaetes and Bacteroides) are ectosymbionts. Apart from the ectosymbionts, Mixotricha has spherical bacteria inside the cell (endosymbionts) functioning as mitochondria, which are absent in this protozoan. Just like in its relatives, including Trichonympha, these bacteria help in cellulose digestion, as without Mixotricha the host termites fail to survive (Radek and Nitsch 2007).
Pseudotrichonympha (class Hypermastigea and order Trichonymphida) carries Hoplonympha, Barburanympha, Urinympha and Staurojoenina as ectosymbionts that form a monophyletic lineage. The endosymbionts of Pseudotrichonympha are distinct (Ohkuma et al. 2005) as they closely resemble the ectosymbionts of Devescovina protists (order Cristamonadida; class Trichomonadea). Two other protist species, Holomastigotoides mirabile and Spirotrichonympha leidyi also dwell in the gut of C. formosanus. Endosymbionts rarely occur within the cells of H. mirabile as this protist carries attached spirochaetes (ectosymbionts) of the cluster I of termite treponemas (Noda et al. 2003). The concentration of ectosymbiotic spirochaetes is less than 200 cells per H. mirabile cell. However, cells of Spirotrichonympha leidyi have an endosymbiotic methanogen (Methanobrevibacter) and only 300 cells of the methanogen occur within a single cell (Shinzato et al. 2005).
4.3.3 Fungi
The nests of the fungus-growing macrotermitine termites can occupy high volumes and can persist for decades, with million sterile helper individuals produced by a single queen (Shellman-Reeve 1997). This agricultural symbiosis with fungi has enabled this termite species to occupy niches loaded with abundant resources that were previously inaccessible (Waller 1988).
The fungal symbionts of Macrotermitinae produce sexual fruiting bodies (Katoh et al. 2002). Symbiotic relationship has a significant role in termite evolution, as it involves a wide range of intestinal microorganisms (Bignell 2000). Macrotermitinae is the only example of a single Termitidae subfamily displaying a mutualistic ectosymbiosis with a white-rot fungi, Termitomyces [tribe Termitomyceteae (Julich) Singer, family Tricholomataceae Roze, Basidiomycotina]. The fungus is capable of digesting lignin and provides food, either directly, when termites consume fungal nodules containing asexual spores, or indirectly, when they ingest partially degraded fungal biomass (Bignell 2000). The fungus is cultivated on a specialized structure within the nest, known as the fungus comb, which is maintained via continuous addition of predigested plant substrate. However, the older comb material is also consumed (Rouland-Lefevre et al. 2002). Approximately 40 species of Termitomyces symbiont are known till date, in Macrotermitinae (Kirk et al. 2001).
Just like most Basidiomycetes, Termitomyces shows homokaryotic and heterokaryotic stages in the life cycle, with one and two genetically different types of haploid nuclei (De Fine Licht et al. 2005). The fungus mycelium is heterokaryotic in all natural colonies, which emphasizes that the incipient termite colonies must contain at least two compatible and genetically different sexual spores, with a relatively short-lived homokaryotic stage (Aanen et al. 2007). The cultivation is beneficial to both the termites and the fungus in a number of ways, for example, the fungus is able to chemically degrade complex substances (e.g. lignin) in an easily accessible form for the termites and also the fungal component increases the termite diet N/C ratio, enabling the use of more diverse cellulose sources. On the other hand, the fungus gains advantages by accessing plant material for easier penetration, along with an increased surface area, in a suitable microclimate optimal for establishment, and, last but not the least, termite secretions prevent spread of microbial infections (Darlington 1994).
The Macrotermitinae nests exhibit an expanded thermoregulation by maintaining constantly high temperatures along with high relative humidity (Korb and Linsenmair 2000a, b). This provides an optimal microclimate for fungal cultivation (Wood and Thomas 1989) as well as a platform for the synergistic interaction of the complementary enzyme systems (enzymes derived from the termite and fungus) for cellulose digestion (Veivers et al. 1991).
The morphospecies of Termitomyces and its fruiting bodies form a monophyletic group intimately associated with the termite nests (Rouland-Lefevre et al. 2002), suggesting a single evolutionary origin of mutualistic symbiosis with termites. Cospeciation and specificity patterns are, however, consistent with the fungal symbiont transmission from host to host, outside the vertical host lineage [horizontal transmission] (Frank 1996).
Horizontal fungal transmission occurs in most of the Macrotermitinae-Termitomyces associations where the fungus produces fruiting bodies (basidiocarps) bearing sexual spores which are carried towards the newly formed nests by the first workers of the new colony through their first foraging trips (Darlington 1994). Laboratory trials confirm that alates (winged sexuals) fail to establish a colony unless provided with external fungal spores (Sieber 1983) and also the fruiting bodies of the fungus are in synchronization with the emergence of first fully developed foraging workers (Johnson 1981). Few Macrotermitinae termites also show vertical, uniparental symbiont transmission, in which reproductive units of termite and fungus are aligned together. Microtermes sp. and Macrotermes bellicosus alates (either male or female) carry in their foregut a bolus of conidia (asexual spores) from the fungus combs of the parent colony for inoculation of first fungus combs in their newly formed colonies (Wood and Thomas 1989). In M. bellicosus males transmit the fungus, while females transmit the fungus in Microtermes sp. However, fungal fruiting bodies have never been identified in these termite species (Darlington 1994). Phylogenetic and molecular investigations have supported this difference in sex-specificity in fungus transmission, pointing out an independent origin of uniparental and vertical transmission in these termite species. Termites with vertical transmission do not form a monophyletic group but fall in two unrelated clades, suggesting horizontal transmission as ancestral mode of transmission of which uniparental transmission is a derived trait, having two independent origins (Aanen et al. 2002).
Like most basidiomycetes, Termitomyces also have a heterothallic (i.e. outcrossing) life cycle where germinating spores form a monokaryon, and all cells carry a single nucleus. Later, two monokaryons of same species with different mating types fuse to form a stable dikaryon, and all cells have two nuclei, one derived from each monokaryon. This dikaryon can form fruiting bodies by meiosis, finally forming a spore. Macrotermitinae fruiting symbionts showing horizontal transmission follow this pattern, while some fruiting Termitomyces fungi also have a homothallic (i.e. non-outcrossing) mating system, in which only a single fungal spore completes the life cycle.
Two main models have been proposed for the successive evolution of the fungus in social insects. The first model is the traditional ‘consumption first’ model where consumption is followed by cultivation and the transmission fungi become essential component of the insect’s diet (vertical fungus transmission). The second model is the alternative ‘transmission first’ model, in which transmission is preceded by consumption, resulting in the cultivation of a specialized fungus which is dispersed by the insect. Insects cultivate the fungus by the addition of substrate (Mueller et al. 2001).
Xylaria (Ascomycotina, Xylariales) occupies a wide variety of habitats including dead or live plant material as endophytic in living plants (Davis et al. 2003) and also a vast number of fungus-growing termite nests, where the termites actively control species composition by continuous excretion of antimicrobial peptides (Fuller 2007). This active suppression of spore germination or mycelial growth by termites results in a patchy distribution of Xylaria across fungus combs. Xylaria species can degrade lignin. Therefore, they cause white rot in wood and plant debris (Osono and Takeda 1999). Termites bring the inocula of Xylaria into their nests by their foraging activities. Certain Xylaria species (X. escharoidea, X. furcate and X. nigripes) show coevolution with termites as they carry smaller spores (Rogers et al. 2005) that can be easily ingested or carried by insects, enhancing the chances of dispersal (Rogers 2000). Termite-associated Xylaria act like ‘sit-and-wait saprotrophs’, foliar endophytes latently present on the leaf and start degrading processes only when the leaf falls from the tree (Herre et al. 2007).
4.4 Physiological Roles of the Microorganisms
The termite gut symbionts carry out the following physiological roles:
4.4.1 Nitrogen Fixation
Nitrogen fixation is a fundamental aspect of symbiosis in termites and is controlled by the gut microbes due to low nitrogen content of the termite diet. The rate of nitrogen fixation greatly varies among and within the same termite species (Braun et al. 1999). A nitrogenase complex catalyses the biological nitrogen fixation (Dean and Jacobson 1992) with the help of a molybdenum (Mo)-containing nitrogenase enzyme, encoded by the nif HDK operon. The cofactors of Mo-independent nitrogenases coordinate either with vanadium or lack Mo/vanadium (alternative nitrogenase) which in turn is encoded by the vnfH-vnf DGK and anfH DGK operons. The nifH, vnfH and anfH genes bear a high degree of sequence conservation (Widmer et al. 1999). Although there is a simultaneous regulation of genes within the single operon, the three operons (nif, vnf and anf) are differentially regulated. Availability of fixed nitrogen strictly regulates transcription of all the three nitrogenase operons, while Mo availability differentially affects expression of nitrogenase genes during transcription (Ohkuma and Kudo 1996).
The nif operon is repressed in the absence of Mo, but vnf and anf operons are repressed in the presence of Mo. A high nitrogen fixation activity, where more than half of the fixed nitrogen comes from the atmospheric N2, is known in the drywood termite, Neotermes koshunensis (Tayasu et al. 1994). A wide diversity of nifH genes has been documented in the gut of N. koshunensis and other termite species as well (Ohkuma et al. 1999). In the symbiotic microbial community of N. koshunensis, the anf gene, connected with termite anf-methano cluster I, is the most critical gene responsible for fixation. It is believed that inadequate amount of Mo in the diet of this termite species is responsible for the expression of anf genes as the ordinary Mo-dependent nitrogenases require Mo as a cofactor for nitrogen-fixing activity. Termite species lacking anf genes in the gut community are possibly able to obtain sufficient amounts of Mo from their food (Ohkuma et al. 1996).
Some methanogenic archaea have nifH genes associated with the anf-methano group of nifH genes, but their nifD genes are phylogenetically grouped with Mo-dependent nitrogenases (Kirshtein et al. 1991). Mo-independent regulation of anf gene expression suggests encoding of a Mo-dependent ordinary enzyme, just like in methanogens. The anfH, anfD and anfG in the anf gene cluster encode both Mo- and V-independent alternative nitrogenases.
Gene organization and sequence features of the termite anf gene distantly resemble with those of well-characterized organisms. For example, in the Archaea domain, a Mo-independent alternative nitrogenase and anfD and anfG orthologous genes have not been reported yet, but the anfH gene resembles bacterial anfH gene. The Archaea domain including all diazotrophic methanogenic archaea contains two ORFs between nifH and nifD genes. In Clostridium cellobioparum, a sequence homologous to ORF105 has been recently identified, but the nucleotide sequence of the DNA region corresponding to ORF122, or even existence of ORF122-like genes, has not been documented yet. Presence of ORF105 in the genome of the bacteria domain indicates that it is not a characteristic of the Archaea domain only. The two ORFs of the termite anf gene cluster closely resemble the methanogen, Methanosarcina barkeri (Chien and Zinder 1996).
N2fixation by the gut microbes contributes to about 60% of the total nitrogen in some termite colonies (Breznak 2000). Several strains of spirochaetes residing in termite gut also fix N2 (Tayasu et al. 1994), and recently ZAS strains, along with their two homologs of nifH, also exhibit nitrogenase activity, in which ZAS-9 shows greatest specific activity (100-fold greater) than ZAS-1 and ZAS-2 (Lilburn et al. 2001). ZAS-9 also fixes 15N2. The NifH amino acid sequences of several spirochaetes along with the ZAS strains are identical or nearly identical to various NifHs observed in termite guts (Noda et al. 1999), indicating their spirochaete origin (Lilburn et al. 2001).
4.4.2 Acetogenesis
The symbiotic protozoa and bacteria present in the hindgut of Reticulitermes flavipes carry out homoacetic fermentation of cellulose. Cellulolytic protozoa first hydrolyse cellulose by fermenting every C (CO2) and H2 (Breznak 1984):
CO2-reducing acetogenic bacteria then convert H2 and CO2 to an additional acetate molecule (Breznak and Switzer 1986):
The three acetates formed per glucose monomer are absorbed and oxidized by the termite to supply 100% of the insect’s respiratory requirement (Odelson and Breznak 1983):
H2 and CH4 (methane is formed by reduction of CO2 by methanogenic bacteria) are also emitted by termites. The rate of CO2 reduction to acetate in the wood- and grass-feeding termites is greater in comparison to the fungus-growing or soil-feeding termites, while the rate of CH4 emission by soil-feeding and fungus-growing termites (lesser extent) is greater than wood-feeding termites. Three strains of CO2-reducing acetogenic bacteria from the guts of a higher and a lower wood-feeding and a higher soil-feeding termite have been described (Kane and Breznak 1991). Each one was a novel and different bacterial species capable of fermenting a variety of organic substrates for energy, including methoxylated aromatics (components of lignin). One of these isolates, Sporomusa termitida, is mixotrophic deriving energy by simultaneous usage of organic and inorganic (H2 + CO2) substrate mixtures (Breznak and Blum 1991). Mixotrophy enhances the ability of acetogens to outcompete methanogens for CO2 reduction in wood- and grass-feeding termites.
Termite emissions can be considered as a significant source of total annual global CH4 production, ranging from <5% to >40%. The hydrogenotrophic activity of acetogenic hindgut bacteria of the wood- and grass-feeding termites produces <10% CH4, while fungus-growing and soil-feeding termites do not have significant levels of bacterial acetogenesis from H2 + CO2 so are more potent sources of CH4 emission (Khalil et al. 1990).
H2/CO2 acetogenesis by termite gut spirochaetes is already known (Breznak 1975). The spirochaetes in higher termites and protozoa in lower termites display the highest H2 concentration (up to 50,000 ppmv) in the luminal region, as confirmed by microelectrode-determined radial profiles (Ebert and Brune 1997). H2-consuming methanogens (nonspirochetal prokaryotes) in R. flavipes lie on or near the hindgut epithelium (Leadbetter and Breznak 1996) and maintain low H2 concentrations (Tholen and Brune 2000). The attachment of spirochaetes on the surface of hindgut protozoa as well as their ability to grow by H2/CO2 acetogenesis is responsible for ‘motility symbiosis’ (Cleveland and Grimstone 1964) allowing access to major sites of H2 production. Recently, fluorescent rRNA-targeted oligonucleotide probes showed that only few distinct phylogenetic types of spirochaetes attach to protozoa (Iida et al. 2000).
Treponema strains ZAS-1 and ZAS-2, although capable of H2/CO2 acetogenesis but not restricted to the substrate like many so-called homoacetogens (anaerobic microbes producing acetate as major fermentation product), also ferment various mono- or disaccharides either alone or by H2 consumption. ZAS-2 also carries out homoacetogenesis by using methyl groups of methoxylated aromatic compounds (Graber and Breznak 2000). Therefore, spirochaetes contribute to demethoxylation of lignin (Esenther and Kirk 1974) as well as other methoxylated aromatic components of termite food, but all termite gut spirochaetes are not homoacetogens. The Treponema strain ZAS-9 ferments sugars and produces acetate and other products, including H2, but cannot conduct H2/CO2 acetogenesis (Graber and Breznak 2000).
4.4.3 Lignin Degradation
Lignin degradation in the gut of insects is carried out by the diversified microbial community. Cellulose degradation in insect guts is very well known (Breznak and Brune 1994), but the fate of lignin needs further research (Brune 2007) as it is widely accepted that insect gut systems lack the capability to degrade lignin (Ohkuma 2003). Apart from this fact, many previous studies have indicated that several wood-feeding insects can overcome the lignin barrier either by feeding on pre-degraded wood (Kukor et al. 1988) or via exosymbiotic relationships with wood-degrading fungi (Johjima et al. 2006) and that a few insect species can feed on the inner wood of alive healthy trees (Taprab et al. 2005). The major products of undegraded wood are 3,4-dimethoxybenzaldehyde (G4) from guaiacyl (G) lignin and 3,4,5-trimethoxybenzaldehyde (S4) from syringyl (S) lignin. The fungal lignin degradation has three main reactions: (i) propyl side-chain oxidation/cleavage, (ii) ring hydroxylation and (iii) demethylation (Filley 2003). Side-chain oxidation is responsible for Cα–Cβ cleavage/depolymerization of lignin in the white-rot fungi. The oxidative alteration of lignin propyl side chain produces higher amounts of 3,4-dimethoxybenzoic acid (G6) and 3,4,5-trimethoxybenzoic acid (S6), increasing the G6/G4 and S6/S4 ratios (Filley et al. 2000). Ring hydroxylation of guaiacyl units brings hydroxylation of either intact or side-chain-oxidized lignin. The diastereomeric pair of the enantiomers 1-(3,4-dimethoxy)-1,2,3-trimethoxypropane represents integral lignin. Syringyl lignin can be more easily degraded and depolymerized (Chiang and Funaoka 1990).
The recent metagenome sequencing of the hindgut microbe community of the higher termite species Nasutitermes corniger strengthens the view that termites can degrade lignin although no genes encoding lignin-degrading enzymes are yet known (Warnecke et al. 2007). It is presumed that hindgut microbes are also anaerobic and can degrade lignin. The lignin modification by the lower termite Cryptotermes brevis showed only minor changes in the molecule, without modifying the side chains (Katsumata et al. 2007). Lignin biodegradation in Zootermopsis angusticollis, a lower termite feeding on coniferous trees, shows side-chain oxidation along with demethylation. All three reactions (side-chain oxidation, ring hydroxylation and demethylation) in this termite gut are possibly related to the brown-rot fungal decay, as brown and white fungi are already known to be associated with termite guts. Excluding the fungal components, aromatic degrading bacteria (actinomycetes) in termite guts are also known. However, they participate in evading lignin barrier although their actual biochemical abilities are not well defined (Delalibera et al. 2005). Actinomycetes plays similar aromatic degrading role in the gut of Anoplophora glabripennis (Schloss et al. 2006).
The well-coordinated cooperation between termites and fungi is responsible for the efficient utilization of lignocellulose. Old workers collect plant litter, while young workers masticate and ingest the collected plant litter which passes down the termite gut without digestion in the form of faecal pellets (primary faeces) used in the formation of fungus comb for the growth of the symbiotic fungi. The fungi form mycelia as well as fungus nodules, and lignin content progressively decreases, with the maturation of the fungus comb (Shary et al. 2007). The in vitro digestibility of cellulose in a matured fungus comb is approximately three times higher as compared to that in a newly formed one. Fungus nodules are consumed by young workers while old workers consume old combs for producing final faeces, but it is almost impossible to observe final faeces which suggest a highly efficient decomposition as well as complete biorecycling of plant litter. It is now very well known that symbiotic fungi degrade lignin bringing easy degradation of cellulose in comparison to the cellulase produced by the termites (Filley et al. 2006).
Lignocellulose digestion needs efficient cellulases along with glycoside hydrolases for the degradation of the cellulose and hemicellulose present in the plant cell wall, in addition to a mechanism for handling lignin barrier which is actually a combined effort of termite and symbionts (Brune 2014). Both higher and lower termites produce enzymes for the first stage of the tricarboxylic acid (TCA) cycle but lack an enzyme capable for the conversion of pyruvate to acetyl CoA or acetate (Breznak and Brune 1994).
Cellulose hydrolysis starts with endoglucanases released by the termites (salivary glands secrete endoglucanases in lower termites while they are secreted within the midgut epithelium in higher termites). Numerous exoglucanases, exoglucanases, β-glucosidases and numerous other glycoside hydrolases are produced by the gut flagellates in lower termites. Higher termites also produce many cellulases, xylanases and other glycoside hydrolases. Metaproteomic studies of the hindgut of N. corniger have shown the presence of almost a quarter of the 886 proteins identified as enzymes of which 36 are glycoside hydrolases (Burnum et al. 2011). These findings suggest that these enzymes are important for the symbiotic relationship of the hindgut microbes and termites. The metaproteomic and metatranscriptomic analysis carried out by He et al. (2013) on N. corniger and Amitermes wheeleri (both higher termites with different diets and habitats) displayed differences in the abundance of certain bacteria in the guts due to their varied diets, but both species contained many glycoside hydrolases for cellulose degradation. No lignin degradation genes were identified in hindguts of the two species (Fig. 4.3). (The details of the cellulose degradation will be discussed in another chapter.)
Major metabolic differences between two higher termites from the metatranscriptomic study of He et al. (2013) (Green lines indicate the genes more abundant in A. wheeleri while red to N. corniger)
4.5 Conclusion
Termites play a vital role in recycling waste material important for the maintenance of the ecological balance. They produce about 11% of the atmospheric methane via cellulose breakdown. Symbiotic protozoa, as well as other flagellate protists residing in their guts, carry out the cellulose degradation. Gut protozoa (Trichonympha) are dependent on symbiotic bacteria for the production of necessary digestive enzymes. The higher termites (family Termitidae), however, produce their own cellulase enzymes. Some termite species practice fungiculture by maintaining ‘fungal gardens’ of specialized fungi Termitomyces, which are being nourished by their excrements. Researchers are trying to understand the details of the relationship between the termite digestive tract and the microbial endosymbionts.
References
Aanen, D. K., Eggleton, P., Rouland-Lefevre, C., Guldberg-Frøslev, T., Rosendahl, S., & Boomsma, J. J. (2002). The evolution of fungus growing termites and their mutualistic fungal symbionts. Proceedings of the National Academy of Sciences of the United States of America, 99, 14887–14892.
Aanen, D. K., Ros, V. I. D., Licht, H. H. D., Mitchell, J., de Beer, Z. W., Slippers, B., Rouland-Lefevre, C., & Boomsma, J. J. (2007). Patterns of interaction specificity of fungus-growing termites and Termitomyces symbionts in South Africa. BMC Evolutionary Biology, 7, 115.
Abe, Y., Bignell, D. E., & Higashi, M. (2000). Termites: Evolution, sociality, symbioses, ecology (p. 466). Dordrecht: Kluwer Academic Publishers.
Baena, S., Fardeau, M. L., Labat, M., Ollivier, B., Thomas, P., Garcia, J. L., & Patel, B. K. (1998). Aminobacterium colombiense sp. nov., an amino-acid-degrading anaerobe isolated from anaerobic sludge. Anaerobe, 4, 241–250.
Baena, S., Fardeau, M. L., Ollivier, B., Labat, M., Thomas, P., Garcia, J. L., & Patel, B. K. (1999a). Aminomonas paucivorans gen. nov., sp. nov., a mesophilic, anaerobic, amino-acid-utilizing bacterium. International Journal of Systematic Bacteriology, 49, 975–982.
Baena, S., Fardeau, M. L., Woo, T. H., Ollivier, B., Labat, M., & Patel, B. K. (1999b). Phylogenetic relationships of three amino-acid-utilizing anaerobes, Selenomonas acidaminovorans, ‘Selenomonas acidaminophila’ and Eubacterium acidaminophilum, as inferred from partial 16S rDNA nucleotide sequences and proposal of Thermanaerovibrio acidaminovorans gen. nov., comb. nov. and Anaeromusa acidaminophila gen. nov., comb. nov. International Journal of Systematic Bacteriology, 49, 969–974.
Baena, S., Fardeau, M. L., Labat, M., Ollivier, B., Garcia, J. L., & Patel, B. K. (2000). Aminobacterium mobile sp. nov., a new anaerobic amino-acid-degrading bacterium. International Journal of Systematic and Evolutionary Microbiology, 50, 259–264.
Bandi, C., Sironi, M., Damiani, G., Magrassi, L., Nalepa, C. A., Laudani, U., & Sacchi, L. (1995). The establishment of intracellular symbiosis in an ancestor of cockroaches and termite. Proceedings of the Royal Society of London – Series B: Biological Sciences, 259, 293–299.
Bauer, S., Tholen, A., Overmann, J., & Brune, A. (2000). Characterization of abundance and diversity of lactic acid bacteria in the hindgut of wood- and soil-feeding termites by molecular and culture dependent techniques. Archives of Microbiology, 173, 126–173.
Berchtold, M., Ludwig, W., & Konig, H. (1994). 16S rDNA sequence and phylogenetic position of an uncultivated spirochete from the hindgut of the termite Mastotermes darwiniensis Froggatt. FEMS Microbiology Letters, 123, 269–273.
Bignell, D. E. (1994). Soil-feeding and gut morphology in higher termites. In J. H. Hunt & C. A. Nalepa (Eds.), Nourishment and evolution in insect societies (pp. 131–158). Boulder: Westview Press.
Bignell, D. E. (2000). Introduction to symbiosis. In T. Abe, D. E. Bignell, & M. Higashi (Eds.), Termites: Evolution, sociality, symbioses, ecology (pp. 189–208). Dordrecht: Kluwer Academic Publishers.
Bignell, D. E., & Eggleton, P. (2000). Termites in ecosystems. In T. Abe, D. E. Bignell, & M. Higashi (Eds.), Termites: Evolution, sociality, symbiosis, ecology (pp. 363–387). Dordrecht: Kluwer Academic Publishers.
Bloodgood, R. A., & Fitzharris, T. P. (1976). Specific associations of prokaryotes with symbiotic flagellate protozoa from the hindgut of the termite Reticulitermes and the wood-eating roach Cryptocercus. Cytobios, 17, 103–122.
Boga, H., & Brune, A. (2003). Hydrogen-dependent oxygen reduction by homoacetogenic bacteria isolated from termite guts. Applied and Environmental Microbiology, 69, 779–786.
Brauman, A., Kane, M. D., Labat, M., & Breznak, J. A. (1992). Genesis of acetate and methane by gut bacteria of nutritionally diverse termites. Science, 257, 1384–1387.
Braun, S. T., Proctor, L. M., Zani, S., Mellon, M. T., & Zehr, J. P. (1999). Molecular evidence for zooplankton-associated nitrogen-fixing anaerobes based on amplification of the nifH gene. FEMS Microbiology Ecology, 28, 273–279.
Breznak, J. A. (1975). Symbiotic relationships between termites and their intestinal microbiota. In D. H. Jennings & D. L. Lee (Eds.), Symbiosis (Society for experimental biology symposium ser., no. 29) (pp. 559–580). Cambridge: Cambridge University Press.
Breznak, J. A. (1984). Biochemical aspects of symbiosis between termites and their intestinal microbiota. In J. M. Anderson, A. D. M. Rayner, & D. W. H. Walton (Eds.), Invertebrate microbial interactions (pp. 173–203). London: Cambridge University Press.
Breznak, J. A. (1994). Acetogenesis from carbon dioxide in termite guts. In H. L. Drake (Ed.), Acetogenesis (pp. 303–330). New York: Chapman and Hall.
Breznak, J. A. (2000). Ecology of prokaryotic microbes in the guts of wood- and litter-feeding termites. In T. Abe, D. E. Bignell, & M. Higashi (Eds.), Termites: Evolution, sociality, symbiosis, ecology (pp. 209–231). Dordrecht: Kluwer Academic Publishers.
Breznak, J. A. (2002). Phylogenetic diversity and physiology of termite gut spirochetes. Integrative and Comparative Biology, 42, 313–318.
Breznak, J. A., & Blum, J. S. (1991). Mixotrophy in the termite gut acetogen, Sporomusa termitida. Archives of Microbiology, 156, 105–110.
Breznak, J. A., & Brune, A. (1994). Role of microorganisms in the digestion of lignocellulose by termites. Annual Review of Entomology, 39, 453–487.
Breznak, J. A., & Switzer, J. M. (1986). Acetate synthesis from H2 plus CO2 by termite gut microbes. Applied and Environmental Microbiology, 52, 623–630.
Brugerolle, G. (2005). The amoeboid parabasalid flagellate Gigantomonas herculea of the African termite Hodotermes mossambicus reinvestigated using immunological and ultrastructural techniques. Acta Protozoologica, 44, 189–199.
Brune, A. (2007). Microbiology: Woodworker’s digest. Nature, 450, 487–488.
Brune, A. (2014). The family Elusimicrobiaceae. In E. Rosenberg, E. F. DeLong, S. Lory, E. Stackebrandt, & F. Thompson (Eds.), The prokaryotes (Vol. 11., 4th ed, pp. 637–640). Berlin: Springer Verlag.
Brune, A., & Friedrich, M. (2000). Microecology of the termite gut: Structure and function on a microscale. Current Opinion in Microbiology, 3, 263–269.
Brune, A., & Kuhl, M. (1996). pH profiles of the extremely alkaline hindguts of soil-feeding termites (Isoptera: Termitidae) determined with microelectrodes. Journal of Insect Physiology, 42, 1121–1127.
Brune, A., & Ohkuma, M. (2011). Role of the termite gut microbiota in symbiotic digestion. In D. E. Bignell, Y. Roisin, & N. Lo (Eds.), Biology of termites: A modern synthesis (pp. 439–475). Dordrecht: Springer.
Brune, A., Emerson, D., & Breznak, J. A. (1995). The termite gut microflora as an oxygen sink: Microelectrode determination of oxygen and pH gradients in guts of lower and higher termites. Applied and Environmental Microbiology, 61, 2681–2687.
Burnum, K. E., Callister, S. J., Nicora, C. D., Purvine, S. O., Hugenholtz, P., Warnecke, F., Scheffrahn, R. H., Smith, R. D., & Lipton, M. S. (2011). Proteome insights into the symbiotic relationship between a captive colony of Nasutitermes corniger and its hindgut microbiome. The ISME Journal, 5, 161–164.
Canale-Parola, E. (1984). Order I. Spirochaetales Buchanan 1917, 163AL. In N. R. Krieg & J. G. Holt (Eds.), Bergey’s manual of systematic bacteriology (pp. 38–39). Baltimore: Williams & Wilkins.
Cavalier-Smith, T. (2002). The phagotrophic origin of eukaryotes and phylogenetic classification of Protozoa. International Journal of Systematic and Evolutionary Microbiology, 52, 297–354.
Charon, N. W., Greenberg, E. P., Koopman, M. B., & Limberger, R. J. (1992). Spirochaete chemotaxis, motility, and the structure of the spirochetal periplasmic flagella. Research in Microbiology, 143, 597–603.
Chen, J. S. (1995). Alcohol dehydrogenase: Multiplicity and relatedness in the solvent-producing clostridia. FEMS Microbiology Reviews, 17, 263–273.
Chiang, V. L., & Funaoka, M. (1990). The difference between guaiacyl and guaiacyl-syringyl lignins in their responses to kraft delignification. Holzforschung, 44, 309–313.
Chien, Y. T., & Zinder, S. H. (1996). Cloning, functional organization, transcript studies, and phylogenetic analysis of the complete nitrogenase structural genes (nifHDK2) and associated genes in the archaeon Methanosarcina barkeri. Journal of Bacteriology, 178, 143–148.
Cleveland, L. R. (1926). Symbiosis among animals with special reference to termites and their intestinal flagellates. The Quarterly Review of Biology, 1, 51–64.
Cleveland, L. R., & Grimstone, A. V. (1964). The fine structure of the flagellate Mixotricha paradoxa and its associated micro-organisms. Proceedings of the Royal Society of London. Series B, Biological Sciences, 159, 668–686.
Cranshaw, W. (2013). Bugs rule: An introduction to the world of insects (p. 188). Princeton: Princeton University Press.
Cummings, J. H., & Macfarlane, G. T. (1997). Role of intestinal bacteria in nutrient metabolism. JPEN, 21, 357–365.
Czolij, R., Slaytor, M., & O’Brien, R. W. (1985). Bacterial flora of the mixed segment and the hindgut of the higher termite Nasutitermes exitiosus Hill (Termitidae, Nasutitermitinae). Applied and Environmental Microbiology, 49, 1226–1236.
Darlington, J. E. C. P. (1994). Nutrition and evolution in fungus-growing ants. In J. H. Hunt & C. A. Nalepa (Eds.), Nourishment and evolution in insect societies (pp. 105–130). Boulder: Westview Press.
Davis, E. C., Franklin, J. B., Shaw, A. J., & Vilgalys, R. (2003). Endophytic Xylaria (Xylariaceae) among liverworts and angiosperms: Phylogenetics, distribution, and symbiosis. American Journal of Botany, 90, 1661–1667.
De Fine Licht, H. H., Andersen, A., & Aanen, D. K. (2005). Termitomyces sp. associated with the termite Macrotermes natalensis has a heterothallic mating system and multinucleate cells. Mycological Research, 109, 314–318.
Dean, D. R., & Jacobson, M. R. (1992). Biochemical genetics of nitrogenase. In G. Stacy, R. H. Burris, & H. J. Evans (Eds.), Biological nitrogen fixation (pp. 763–834). New York: Chapman and Hall.
Delalibera, I., Handelsman, J., & Raffa, K. F. (2005). Contrasts in cellulolytic activities of gut microorganisms between the wood borer, Saperda vestita (Coleoptera: Cerambycidae) and the bark beetles, Ips pini and Dendroctonus frontalis (Coleoptera: Curculionidae). Environmental Entomology, 34, 541–547.
Dietrich, C., Kohler, T., & Brune, A. (2014). The cockroach origin of the termite gut microbiota: Patterns in bacterial community structure reflect major evolutionary events. Applied and Environmental Microbiology, 80, 2261–2269.
Donovan, S. E., Purdy, K. J., Kane, M. D., & Eggleton, P. (2004). Comparison of Euryarchaea strains in the guts and food-soil of the soil-feeding termite Cubitermes fungifaber across different soil types. Applied and Environmental Microbiology, 70, 3884–3892.
Dow, J. A. T. (1986). Insect midgut function. Advances in Insect Physiology, 19, 188–328.
Droge, S., Rachel, R., Radek, R., & Konig, H. (2008). Treponema isoptericolens sp. nov., a novel spirochaete from the hindgut of the termite Incisitermes tabogae. International Journal of Systematic and Evolutionary Microbiology, 58, 1079–1083.
Ebert, A., & Brune, A. (1997). Hydrogen concentration profiles at the oxic–anoxic interface: A microsensor study of the hindgut of the wood-feeding lower termite Reticulitermes flavipes (Kollar). Applied and Environmental Microbiology, 63, 4039–4046.
Engel, M. S., Grimaldi, D. A., & Krishna, K. (2009). Termites (Isoptera): Their phylogeny, classification, and rise to ecological dominance. American Museum Novitates, 3650, 1–27.
Esenther, G. R., & Kirk, T. K. (1974). Catabolism of aspen sapwood in Reticulitermes flavipes (Isoptera: Rhinotermitidae). Annals of the Entomological Society of America, 67, 989–991.
Filley, T. R. (2003). In B. Goodell, D. D. Nicholas, & T. P. Schultz (Eds.), ACS symposium series in wood deterioration and preservation: Advances in our changing world (pp. 119–139). Washington, DC: American Chemical Society.
Filley, T. R., Hatcher, P. G., Shortle, W. C., & Praseuth, R. T. (2000). The application of 13C-labeled tetramethylammonium hydroxide (13C-TMAH) thermochemolysis to the study of fungal degradation of wood. Organic Geochemistry, 31, 181–198.
Filley, T. R., Nierop, K. G. J., & Wang, Y. (2006). The contribution of polyhydroxyl aromatic compounds to tetramethylammonium hydroxide lignin-based proxies. Organic Geochemistry, 37, 711–727.
Frank, S. A. (1996). Host-symbiont conflict over the mixing of symbiotic lineages. Proceedings of the Royal Society of London B: Biological Sciences, 263, 339–344.
Frohlich, J., Sass, H., Babenzien, H.-D., Kuhnigk, T., Varma, A., Saxena, S., Nalepa, C., Pfeiffer, P., & Konig, H. (1999). Isolation of Desulfovibrio intestinalis sp. nov. from the hindgut of the lower termite Mastotermes darwiniensis. Canadian Journal of Microbiology, 45, 145–152.
Fuhrman, J. A., McCallum, K., & Davis, A. A. (1993). Phylogenetic diversity of subsurface marine microbial communities from the Atlantic and Pacific Oceans. Applied and Environmental Microbiology, 59, 1294–1302.
Fuller, C. A. (2007). Fungistatic activity of freshly killed termite, Nasutitermes acajutlae, soldiers in the Caribbean. Journal of Insect Science, 7, 14.
Gerbod, D., Sanders, E., Moriya, S., Noel, C., Takasu, H., Fast, N. M., Delgado-Viscogliosi, P., Ohkuma, M., Kudo, T., Capron, M., Palmer, J. D., Keeling, P. J., & Viscogliosi, E. (2004). Molecular phylogenies of Parabasalia inferred from four protein genes and comparison with rRNA trees. Molecular Phylogenetics and Evolution, 31, 572–580.
Godon, J.-J., Moriniere, J., Moletta, M., Gaillac, M., Bru, V., & Delgenes, J.-P. (2005). Rarity associated with specific ecological niches in the bacterial world: The ‘Synergistes’ example. Environmental Microbiology, 7, 213–224.
Graber, J. R., & Breznak, J. A. (2000). Nutrition and physiological properties of termite gut spirochaetes. Abstracts of the American Society for Microbiology no. N-104.
Graber, J. R., Leadbetter, J. R., & Breznak, J. A. (2004). Description of Treponema azonutricum sp. nov and Treponema primitia sp. nov., the first spirochetes isolated from termite guts. Applied and Environmental Microbiology, 70, 1307–1314.
Grimaldi, D. (2001). Insect evolutionary history from Handlirsch to Hennig, and beyond. Journal of Paleontology, 75, 1152–1160.
He, S., Ivanova, N., Kirton, E., Allgaier, M., Bergin, C., Scheffrahn, R. H., Kyrpides, N. C., Warnecke, F., Tringe, S. G., & Hugenholtz, P. (2013). Comparative metagenomic and metatranscriptomic analysis of hindgut paunch microbiota in wood- and dung-feeding higher termites. PLoS One, 8, e61126.
Herre, E. A., Mejia, L. C., Kyllo, D. A., Rojas, E., Maynard, Z., Butler, A., & Van Bael, S. A. (2007). Ecological implications of anti-pathogen effects of tropical fungal endophytes and mycorrhizae. Ecology, 88, 550–558.
Hethener, P., Braumann, A., & Garcia, J.-L. (1992). Clostridium termitidis sp. nov., a cellulolytic bacterium from the gut of the wood feeding termite, Nasutitermes lujae. Systematic and Applied Microbiology, 15, 52–58.
Higashi, M., & Abe, T. (1997). Global diversification of termites driven by the evolution of symbiosis and sociality. In T. Abe, S. A. Levin, & M. Higashi (Eds.), Biodiversity: An ecological perspective (pp. 83–112). New York: Springer-Verlag.
Hongoh, Y. (2010). Diversity and genomes of uncultured microbial symbionts in the termite gut. Bioscience, Biotechnology, and Biochemistry, 74, 1145–1151.
Hongoh, Y. (2011). Toward the functional analysis of uncultivable, symbiotic microorganisms in the termite gut. Cellular and Molecular Life Sciences, 68, 1311–1325.
Hongoh, Y., Ohkuma, M., & Kudo, T. (2003). Molecular analysis of bacterial microbiota in the gut of the termite Reticulitermes speratus (Isoptera; Rhinotermitidae). FEMS Microbiology Ecology, 44, 231–242.
Hongoh, Y., Deevong, P., Hattori, S., Inoue, T., Noda, S., Noparatnaraporn, N., Kudo, T., & Ohkuma, M. (2006). Phylogenetic diversity, localization and cell morphologies of the candidate phylum TG3 and a subphylum in the phylum Fibrobacteres, recently found bacterial groups dominant in termite guts. Applied and Environmental Microbiology, 72, 6780–6788.
Horn, M., Harzenetter, M. D., Linner, T., Schmid, E. N., Muller, K.-D., Michel, R., & Wagner, M. (2001). Members of the Cytophaga-Flavobacterium-Bacteroides phylum as intracellular bacteria of acanthamoebae: proposal of ‘Candidatus Amoebophilus asiaticus.’. Environmental Microbiology, 3, 440–449.
Huang, X. F., Bakker, M. G., Judd, T. M., Reardon, K. F., & Vivanco, J. M. (2013). Variations in diversity and richness of gut bacterial communities of termites (Reticulitermes flavipes) fed with grassy and woody plant substrates. Microbial Ecology, 65, 531–536.
Hungate, R. E. (1946). Studies on cellulose fermentation. II. An anaerobic cellulose-decomposing actinomycete, Micromonospora propionici, n. sp. Journal of Bacteriology, 51, 51–56.
Husseneder, C. (2010). Symbiosis in subterranean termites: A review of insights from molecular studies. Environmental Entomology, 39, 378–388.
Hyodo, F., Inoue, T., Azuma, J. I., Tayasu, I., & Abe, T. (2000). Role of the mutualistic fungus in lignin degradation in the fungus-growing termite Macrotermes gilvus (Isoptera; Macrotermitinae). Soil Biology and Biochemistry, 32, 653–658.
Hyodo, F., Tayasu, I., Inoue, T., Azuma, J.-I., Kudo, T., & Abe, T. (2003). Differential role of symbiotic fungi in lignin degradation and food provision for fungus-growing termites (Macrotermitinae: Isoptera). Functional Ecology, 17, 186–193.
Iida, T., Ohkuma, M., Ohtoko, K., & Kudo, T. (2000). Symbiotic spirochetes in the termite hindgut: Phylogenetic identification of ectosymbiotic spirochetes of oxymonad protists. FEMS Microbiology Ecology, 34, 17–26.
Ikeda-Ohtsubo, W., & Brune, A. (2009). Cospeciation of termite gut flagellates and their bacterial endosymbionts: Trichonympha species and ‘Candidatus Endomicrobium trichonymphae’. Molecular Ecology, 18, 332–342.
Inoue, T., Murashima, K., Azuma, J.-I., Sugimoto, A., & Slaytor, M. (1997). Cellulose and xylan utilisation in the lower termite Reticulitermes speratus. Journal of Insect Physiology, 43, 235–242.
Inoue, T., Kitade, O., Yoshimura, T., & Yamaoka, I. (2000). Symbiotic associations with protists. In T. Abe, D. E. Bignell, & M. Higashi (Eds.), Termites: Evolution, sociality, symbioses, ecology (pp. 275–288). Dordrecht: Kluwer Academic Publishers.
Inward, D., Beccaloni, G., & Eggleton, P. (2007). Death of an order: A comprehensive molecular phylogenetic study confirms that termites are eusocial cockroaches. Biology Letters, 3, 331–335.
Johjima, T., Taprab, Y., Noparatnaraporn, N., Kudo, T., & Ohkuma, M. (2006). Large-scale identification of transcripts expressed in a symbiotic fungus (Termitomyces) during plant biomass degradation. Applied Microbiology and Biotechnology, 73, 195–203.
Johnson, R. A. (1981). Colony development and establishment of the fungus comb in Microtermes sp. nr. Usambaricus (Sjöst) (Isoptera, Macrotermitinae) from Nigeria. Insectes Sociaux, 28, 3–12.
Kane, M. D., & Breznak, J. A. (1991). Acetonema longum gen. nov. sp. nov., an H2/CO2 acetogenic bacterium from the termite, Pterotermes occidentis. Archives of Microbiology, 156, 91–98.
Kane, M. D., Brauman, A., & Breznak, J. A. (1991). Clostridium mayombei sp. nov., an H2/CO2 acetogenic bacterium from the gut of the African soil-feeding termite, Cubitermes speciosus. Archives of Microbiology, 156, 99–104.
Kappler, A. (1999). Influence of gut alkalinity and oxygen status on mobilization and size-class distribution of humic acids in the hindgut of soil-feeding termites. Applied Soil Ecology, 13, 3.
Katoh, H., Miura, T., Maekawi, K., Shinzato, N., & Matsumoto, T. (2002). Genetic variation of symbiotic fungi cultivated by the macrotermitine termite Odontotermes formosanus (Isoptera: Termitidae) in the Ryukyu Archipelago. Molecular Ecology, 11, 1565–1572.
Katsumata, K. S., Jin, Z. F., Hori, K., & Iiyama, K. (2007). Structural changes in lignin of tropical woods during digestion by termite, Cryptotermes brevis. Journal of Wood Science, 53, 419–426.
Katzin, L. I., & Kirby, H. (1939). The relative weight of termites and their protozoa. The Journal of Parasitology, 25, 444–445.
Khalil, M. A. K., Rasmussen, R. A., French, J. R., & Holt, J. (1990). The influence of termites on atmospheric trace gases: CH4, CO2, CHCl3, N2O, CO, H2, and light hydrocarbons. Journal of Geophysical Research, 95, 3619–3634.
Kirk, P. M., Canno, P. F., David, J. C., & Stalpers, J. A. (2001). Ainsworth & Bigby’s dictionary of the fungi. Wallingford: CAB International.
Kirshtein, J. D., Paerl, H. W., & Zehr, J. (1991). Amplification, cloning, and sequencing of a nifH segment from aquatic microorganisms and natural communities. Applied and Environmental Microbiology, 57, 2645–2650.
Kitade, O., Maeyama, T., & Matsumoto, T. (1997). Establishment of symbiotic flagellate fauna of Hodotermopsis japonica (Isoptera: Termopsidae). Sociobiology, 30, 161–167.
Korb, J., & Linsenmair, K. E. (2000a). Thermoregulation of termite mounds: What role does ambient temperature and metabolism of the colony play? Insectes Sociaux, 47, 357–363.
Korb, J., & Linsenmair, K. E. (2000b). Ventilation of termite mounds: New results require a new model. Behavioral Ecology, 11, 486–494.
Krasil’nikov, N. A., & Satdykov, S. I. (1970). Bacteria of termites’ intestines. Microbiology, 39, 562–564.
Kudo, T., Ohkuma, M., Moriya, S., Noda, S., & Ohtoko, K. (1998). Molecular phylogenetic identification of the intestinal anaerobic microbial community in the hindgut of the termite, Reticulitermes speratus, without cultivation. Extremophiles, 2, 155–161.
Kuhnigk, T., Branke, J., Krekeler, D., Cypionka, H., & Konig, H. (1996). A feasible role of sulfate-reducing bacteria in the termite gut. Systematic and Applied Microbiology, 19, 139–149.
Kukor, J., Cowan, D., & Martin, M. (1988). The role of ingested fungal enzymes in cellulose digestion in the larvae of cerambycid beetles. Physiological Zoology, 61, 364–371.
Leadbetter, J. R., & Breznak, J. A. (1996). Physiological ecology of Methanobrevibacter cuticularis sp. nov. and Methanobrevibacter curvatus sp. nov., isolated from the hindgut of the termite Reticulitermes flavipes. Applied and Environmental Microbiology, 62, 3620–3631.
Leadbetter, J. R., Crosby, L. D., & Breznak, J. A. (1998). Methanobrevibacter filiformis sp. nov., a filamentous methanogen from termite hindguts. Archives of Microbiology, 169, 287–292.
Leadbetter, J. R., Schmidt, T. M., Graber, J. R., & Breznak, J. A. (1999). Acetogenesis from H2 plus CO2 by spirochetes from termite guts. Science, 283, 686–689.
Leidy, J. (1877). On the intestinal parasites of Termes flavipes. Proceedings of the Academy of Natural Sciences (Philadelphia), 29, 146–149.
Leidy J (1874–1881) The parasites of the termites. Journal of the Academy of Natural Sciences (Philadelphia) 8, 425–447.
Li, Z. Q., Liu, B. R., Zeng, W. H., Xiao, W. L., Li, Q. J., & Zhong, J. H. (2013). Character of cellulase activity in the guts of flagellate-free termites with different feeding habits. Journal of Insect Science, 13, 1–8.
Lilburn, T. G., Kim, K. S., Ostrom, N. E., Byzek, K. R., Leadbetter, J. R., & Breznak, J. A. (2001). Nitrogen fixation by symbiotic and free-living spirochaetes. Science, 292, 2495–2498.
Liu, N., Zhang, L., Zhou, H., Zhang, M., Yan, X., Wang, Q., Long, Y., Xie, L., Wang, S., Huang, Y., & Zhou, Z. (2013). Metagenomic insights into metabolic capacities of the gut microbiota in a fungus-cultivating termite (Odontotermes yunnanensis). PLoS One, 8, e69184.
Lo, N. (2003). Evidence for cocladogenesis between diverse dictyopteran lineages and their intracellular endosymbionts. Molecular Biology and Evolution, 20, 907–913.
Makonde, H. M., Boga, H. I., Osiemo, Z., Mwirichia, R., Mackenzie, L. M., Goker, M., & Klenk, H. P. (2013). 16S-rRNA-based analysis of bacterial diversity in the gut of fungus-cultivating termites (Microtermes and Odontotermes species). Antonie van Leeuwenhoek, 104, 869–883.
Mauldin, J. K. (1977). Cellulose catabolism and lipid synthesis by normally and abnormally faunated termites, Reticulitermes flavipes. Insect Biochemistry, 7, 27–31.
Mikaelyan, A., Strassert, J. F. H., Tokuda, G., & Brune, A. (2014). The fiber-associated cellulolytic bacterial community in the hindgut of wood-feeding higher termites (Nasutitermes spp.) Environmental Microbiology, 16, 2711–2722.
Moriya, S., Dacks, J. B., Takagi, A., Noda, S., Ohkuma, M., Doolittle, W. F., & Kudo, T. (2003). Molecular phylogeny of three oxymonad genera: Pyrsonympha, Dinenympha and Oxymonas. The Journal of Eukaryotic Microbiology, 50, 190–197.
Mueller, U. G., Schultz, T. R., Currie, C. R., Adams, R. M. M., & Malloch, D. (2001). The origin of the attine ant-fungus mutualism. The Quarterly Review of Biology, 76, 169–197.
Nalepa, C. A., Bignell, D. E., & Bandi, C. (2001). Detritivory, coprophagy, and the evolution of digestive mutualisms in Dictyoptera. Insectes Sociaux, 48, 194–201.
Nobre, T., Rouland-Lefevre, C., & Aanen, D. K. (2011). Comparative biology of fungus cultivation in termites and ants. In D. Bignell, Y. Roisin, & N. Lo (Eds.), Biology of termites: A modern synthesis (pp. 193–210). Dordrecht: Springer.
Noda, S., Ohkuma, M., Usami, R., Horikoshi, K., & Kudo, T. (1999). Culture-independent characterization of a gene responsible for nitrogen fixation in the symbiotic microbial community in the gut of the termite Neotermes koshunensis. Applied and Environmental Microbiology, 65, 4935–4942.
Noda, S., Ohkuma, M., Yamada, A., Hongoh, Y., & Kudo, T. (2003). Phylogenetic position and in situ identification of ectosymbiotic spirochetes on protists in the termite gut. Applied and Environmental Microbiology, 69, 625–633.
Noda, S., Hongoh, Y., Sato, T., & Ohkuma, M. (2009). Complex coevolutionary history of symbiotic Bacteroidales bacteria of various protists in the gut of termites. BMC Evolutionary Biology, 9, 158.
Noirot, C., & Noirot-Timothee, C. (1969). The digestive system. In K. Krishna & F. M. Weesner (Eds.), Biology of termites (pp. 49–88). New York: Academic Press.
O’Brien, R. W., & Slaytor, M. (1982). Role of microorganisms in the metabolism of termites. Australian Journal of Biological Sciences, 35, 239–262.
Odelson, D. A., & Breznak, J. A. (1983). Volatile fatty acid production by the hindgut microbiota of xylophagous termites. Applied and Environmental Microbiology, 45, 1602–1613.
Ohkuma, M. (1998). Phylogenetic analysis of the symbiotic intestinal microflora of the termite Cryptotermes domesticus. FEMS Microbiology Letters, 164, 389–395.
Ohkuma, M. (2003). Termite symbiotic systems: Efficient bio-recycling of lignocellulose. Applied Microbiology and Biotechnology, 61, 1–9.
Ohkuma, M., & Brune, A. (2011). Diversity, structure, and evolution of the termite gut microbial community. In D. E. Bignell, Y. Roisin, & N. Lo (Eds.), Biology of termites: A modern synthesis (pp. 413–438). Dordrecht: Springer.
Ohkuma, M., & Kudo, T. (1996). Phylogenetic diversity of the intestinal bacterial community in the termite Reticulitermes speratus. Applied and Environmental Microbiology, 62, 461–468.
Ohkuma, M., Noda, S., Usami, R., Horikoshi, K., & Kudo, T. (1996). Diversity of nitrogen fixation genes in the symbiotic intestinal microflora of the termite Reticulitermes speratus. Applied and Environmental Microbiology, 62, 2747–2752.
Ohkuma, M., Noda, S., & Kudo, T. (1999). Phylogenetic diversity of nitrogen fixation genes in the symbiotic microbial community in the gut of diverse termites. Applied and Environmental Microbiology, 65, 4926–4934.
Ohkuma, M., Yuzawa, H., Amornsak, W., Sornnuwat, Y., Takematsu, Y., Yamada, A., Vongkaluang, C., Sarnthoy, O., Kirtibutr, N., Noparatnaraporn, N., Kudo, T., & Inoue, T. (2004). Molecular phylogeny of Asian termites (Isoptera) of the families Termitidae and Rhinotermitidae based on mitochondrial COII sequences. Molecular Phylogenetics and Evolution, 31, 701–710.
Ohkuma, M., Iida, T., Ohtoko, K., Yuzawa, H., Noda, S., Viscogliosi, E., & Kudo, T. (2005). Molecular phylogeny of parabasalids inferred from small subunit rRNA sequences, with emphasis on the Hypermastigea. Molecular Phylogenetics and Evolution, 35, 646–655.
Ohkuma, M., Noda, S., Hongoh, Y., Nalepa, C. A., & Inoue, T. (2009). Inheritance and diversification of symbiotic trichonymphid flagellates from a common ancestor of termites and the cockroach Cryptocercus. Proceedings of the Royal Society of London – Series B: Biological Sciences, 276, 239–245.
Osono, T., & Takeda, H. (1999). Decomposing ability of interior and surface fungal colonizers of beech leaves with reference to lignin decomposition. European Journal of Soil Biology, 35, 51–56.
Paster, B. J., Dewhirst, F. E., Weisburg, W. G., Tordoff, L. A., Fraser, G. J., Hespell, R. B., Stanton, T. B., Zablen, L., Mandelco, L., & Woese, C. R. (1991). Phylogenetic analysis of the spirochetes. Journal of Bacteriology, 173, 6101–6109.
Paster, B. J., Dewhirst, F. E., Cooke, S. M., Fussing, V., Poulsen, L. K., & Breznak, J. A. (1996). Phylogeny of not-yet-cultured spirochetes from termite guts. Applied and Environmental Microbiology, 62, 347–352.
Potrikus, C. J., & Breznak, J. A. (1980). Uric acid-degrading bacteria in guts of termites [Reticulitermes flavipes (Kollar)]. Applied and Environmental Microbiology, 40, 117–124.
Ptacek, P., Brandstetr, J., Soukal, F., & Opravil, T. (2013). Investigation of subterranean termites nest material composition, structure and properties. In Y. Mastai (Ed.), Materials science-advanced topics. London: InTech. https://doi.org/10.5772/55145.
Radek, R., & Nitsch, G. (2007). Ectobiotic spirochetes of flagellates from the termite Mastotermes darwiniensis: Attachment and cyst formation. European Journal of Protistology, 43, 281–294.
Rainey, F. A., Ward-Rainey, N. L., Janssen, P. H., Hippe, H., & Stackebrandt, E. (1996). Clostridium paradoxum DSM 7308T contains multiple 16S rRNA genes with heterogeneous intervening sequences. Microbiology, 142, 2087–2095.
Rees, G. N., Patel, B. K., Grassia, G. S., & Sheehy, A. J. (1997). Anaerobaculum thermoterrenum gen. nov., sp. nov., a novel, thermophilic bacterium which ferments citrate. International Journal of Systematic Bacteriology, 47, 150–154.
Rieu-Lesme, F., Dauga, C., Morvan, O., Bouvet, P., Grimont, P. A., & Dore, J. (1996). Acetogenic coccoid spore-forming bacteria isolated from the rumen. Research in Microbiology, 147, 753–764.
Rogers, J. D. (2000). Thoughts and musings on tropical Xylariaceae. Mycological Research, 104, 1412–1420.
Rogers, J. D., YM, J., & Lehmann, J. (2005). Some Xylaria species on termite nests. Mycologia, 97, 914–923.
Rouland, C., Lenoir, F., & Lepage, M. (1991). The role of the symbiotic fungus in the digestive metabolism of several species of fungus-growing termites. Comparative Biochemistry and Physiology. Part A, Physiology, 99, 657–663.
Rouland-Lefevre, C., Diouf, M. N., Brauman, A., & Neyra, M. (2002). Phylogenetic relationships in Termitomyces (Family Agaricaceae) based on the nucleoticle sequence of ITS: A first approach to elucidate the evolutionary history of the symbiosis between fungus-growing termites and their fungi. Molecular Phylogenetics and Evolution, 22, 423–429.
Sanderson, M. G. (1996). Biomass of termites and their emissions of methane and carbon dioxide: A global database. Global Biogeochem Cycles, 10, 543–557.
Scharf, M. E., Karl, Z. J., Sethi, A., & Boucias, D. G. (2011). Multiple levels of synergistic collaboration in termite lignocellulose digestion. PLoS One, 6, e21709.
Schauer, C., Thompson, C. L., & Brune, A. (2012). The bacterial community in the gut of the cockroach Shelfordella lateralis reflects the close evolutionary relatedness of cockroaches and termites. Applied and Environmental Microbiology, 78, 2758–2767.
Schloss, P. D., Delalibera, I., Handelsman, J., & Raffa, K. F. (2006). Bacteria associated with the guts of two wood-boring beetles: Anoplophora glabripennis and Saperda vestita (Cerambycidae). Environmental Entomology, 35, 625–629.
Schmitt-Wagner, D., Friedrich, M. W., Wagner, B., & Brune, A. (2003). Phylogenetic diversity, abundance, and axial distribution of bacteria in the intestinal tract of two soil-feeding termites (Cubitermes spp). Applied and Environmental Microbiology, 69, 6007–6017.
Shah, H. N. (1992). The genus Bacteroides and related taxa. In A. Balows, H. G. Truper, M. Dworkin, W. Harder, & K. H. Schleifer (Eds.), The Prokaryotes (pp. 3593–3607). New York: Springer.
Shary, S., Ralph, S. A., & Hammel, K. E. (2007). New insights into the ligninolytic capability of a wood decay ascomycete. Applied and Environmental Microbiology, 73, 6691–6694.
Shellman-Reeve, J. S. (1997). In J. C. Choe & B. J. Crespi (Eds.), Social competition and cooperation in insects and arachnids (pp. 52–93). Cambridge: Cambridge University Press.
Shinzato, N., Matsumoto, T., Yamaoka, I., Oshima, T., & Yamagishi, A. (2001). Methanogenic symbionts and the locality of other host lower termites. Microbes and Environments, 16, 43–47.
Shinzato, N., Muramatsu, M., Matsui, T., & Watanabe, Y. (2005). Molecular phylogenetic diversity of the bacterial community in the gut of the termite Coptotermes formosanus. Bioscience, Biotechnology, and Biochemistry, 69, 1145–1155.
Sieber, R. (1983). Establishment of fungus comb in laboratory colonies of Macrotermes michaelseni and Odontotermes montanus (Isoptera, Macrotermitinae). Insectes Sociaux, 30, 204–209.
Simpson, A. G. B. (2006). Comprehensive multigene phylogenies of excavate protists reveal the evolutionary positions of “primitive” eukaryotes. Molecular Biology and Evolution, 23, 615–625.
Stingl, U., Maass, A., Radek, R., & Brune, A. (2004). Symbionts of the gut flagellate Staurojoenina sp. from Neotermes cubanus represent a novel, termite- associated lineage of Bacteroidales: Description of ‘Candidatus Vestibaculum illigatum’. Microbiology, 150, 2229–2235.
Stingl, U., Radek, R., Yang, H., & Brune, A. (2005). “Endomicrobia”: Cytoplasmic symbionts of termite gut protozoa form a separate phylum of prokaryotes. Applied and Environmental Microbiology, 71, 1473–1479.
Surkov, A. V., Dubinina, G. A., Lysenko, A. M., Glockner, F. O., & Kuever, J. (2001). Dethiosulfovibrio russensis sp. nov., Dethosulfovibrio marinus sp. nov. and Dethosulfovibrio acidaminovorans sp. nov., novel anaerobic, thiosulfate and sulfur-reducing bacteria from ‘Thiodendron’ sulfur mats in different saline environments. International Journal of Systematic and Evolutionary Microbiology, 51, 327–337.
Taprab, Y., Johjima, T., Maeda, Y., Moriya, S., Trakulnaleamsai, S., Noparatnaraporn, N., Ohkuma, M., & Kudo, T. (2005). Symbiotic fungi produce laccases potentially involved in phenol degradation in fungus combs of fungus-growing termites in Thailand. Applied and Environmental Microbiology, 71, 7696–7704.
Tayasu, I., Sugimoto, A., Wada, E., & Abe, T. (1994). Xylophagous termites depending on atmospheric nitrogen. Naturwissenschaften, 81, 229–231.
Tellam, R. L., Wijffels, G., & Willadsen, P. (1999). Peritrophic matrix proteins. Insect Biochemistry and Molecular Biology, 29, 87–101.
Thayer, D. W. (1976). Facultative wood-digesting bacteria from the hind-gut of the termite Reticulitermes hesperus. Journal of General Microbiology, 95, 287–296.
Tholen, A., & Brune, A. (1997). Location and in situ activities of homoacetogenic bacteria in the highly compartmentalized hindgut of soil feeding higher termites (Cubitermes spp). Applied and Environmental Microbiology, 65, 44975–44405.
Tholen, A., & Brune, A. (2000). Impact of oxygen on metabolic fluxes and in situ rates of reductive acetogenesis in the hindgut of the wood-feeding termite Reticulitermes flavipes. Environmental Microbiology, 2, 436–449.
Thorne, B. L. (1997). Evolution of eusociality in termites. Annual Review of Ecology and Systematics, 28, 27–54.
Tokuda, G., Watanabe, H., Matsumoto, T., & Noda, H. (1997). Cellulose digestion in the wood-eating higher termite, Nasutitermes takasagoensis (Shiraki): Distribution of cellulases and properties of endo-β -1,4-glucanase. Zoological Science, 14, 83–93.
Tokuda, G., Yamaoka, I., & Noda, H. (2000). Localization of symbiotic clostridia in the mixed segment of the termite Nasutitermes takasagoensis (Shiraki). Applied and Environmental Microbiology, 66, 2199–2207.
Tokura, M., Ohkuma, M., & Kudo, T. (2000). Molecular phylogeny of methanogens associated with flagellated protists in the gut and with the gut epithelium of termites. FEMS Microbiology Ecology, 33, 233–240.
Veivers, P. C., Muhlemann, R., Slaytor, M., Leuthold, R. H., & Bignell, D. E. (1991). Digestion, diet and polyethism in two fungus-growing termites: Macrotermes subhyalinus Rambur and M. michaelseni Sjostedt. Journal of Insect Physiology, 37, 675–682.
Waller, D. A. (1988). Ecological similarities of fungus growing ants (Attini) and termites (Macrotermitinae). In J. C. Trager & E. J. Brill (Eds.), Advances in myrmecology (pp. 337–345). New York: E. J. Brill.
Warnecke, F., Luginbuhl, P., Ivanova, N., et al. (2007). Metagenomic and functional analysis of hindgut microbiota of a wood-feeding higher termite. Nature, 450, 560–565.
Wenzel, M., Radek, R., Brugerollec, G., & Konig, H. (2003). Identification of the ectosymbiotic bacteria of Mixotricha paradoxa involved in movement symbiosis. European Journal of Protistology, 39, 11–23.
Widmer, F., Shaffer, B. T., Porteous, L. A., & Seidler, R. J. (1999). Analysis of nifH gene pool complexity in soil and litter at a Douglas fir forest site in the Oregon Cascade mountain range. Applied and Environmental Microbiology, 65, 374–380.
Wood, T. G., & Thomas, R. J. (1989). The mutualistic association between Macrotermitinae and Termitomyces. In N. Wilding, N. M. Collins, P. M. Hammond, & J. F. Webber (Eds.), Insect-fungus interactions (pp. 69–92). New York: Academic Press.
Yamaoka, I., & Nagatani, Y. (1980). Phagocytic cells in the midgut epithelium of the termite, Reticulitermes speratus (Kolbe). Zoological Magazine, 89, 308–311.
Yamin, M. A. (1979). Termite flagellates. Sociobiology, 4, 1–119.
Zavarzina, D. G., Zhilina, T. N., Tourova, T. P., Kuznetsov, B. B., Kostrikina, N. A., & Bonch-Osmolovskaya, E. A. (2000). Thermanaerovibrio velox sp. nov., a new anaerobic, thermophilic, organotrophic bacterium that reduces elemental sulfur, and emended description of the genus Thermanaerovibrio. International Journal of Systematic and Evolutionary Microbiology, 50, 1287–1295.
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Maurice, N., Erdei, L. (2018). Termite Gut Microbiome. In: Khan, M., Ahmad, W. (eds) Termites and Sustainable Management. Sustainability in Plant and Crop Protection. Springer, Cham. https://doi.org/10.1007/978-3-319-72110-1_4
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