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Lipid, glycogen, proteins, uric acid, and body water in termites and other insects.


Like other insects, termites store fat and carbohydrate in certain types of cells. Uric acid, the main waste product of protein digestion for terrestrial insects, also accumulates in specialized fat body cells of termites as well as the closely related cockroach. Uric acid can be utilized as a nutrient by termites and cockroaches following digestion by uricolytic bacteria harbored in the hindgut or fat body. These bacterial symbionts convert uric acid, which contains four nitrogen atoms, to ammonia that is used for protein synthesis by the insect host. Body water must be maintained within a narrow limit for an insect's survival. Subterranean termites must remain in humid, enclosed areas, or will quickly lose body water and die. This literature review covers food molecules, uric acid, and body water involved in insect nutrition, metabolism, and digestion, with termites given emphasis.

Uric Acid

Uric acid is probably synthesized in insect fat body (Candy 1985) and is the primary metabolic waste product of most insects (Chapman, 1998; Nation 2002). The uricolytic pathway is the main route for uric acid production in insects, starting with the digestion of proteins (Nation 2002). The pathway also occurs in mammals and birds (Downer 1982). Less common is the nucleolytic pathway, with uric acid precursors of nucleic acids, purines, or pyrimidines rather than proteins (Downer 1982).

Some insects store uric acid in specialized fat body cells called urocytes (Chapman 1998). Stored uric acid has been described as precipitated spherules of potassium or sodium urates (Mullins 1979). Insects that store uric acid include tobacco hornworm larvae, larval Chrysopa lacewings, and adult mosquitoes (Chapman 1998).

Termites and cockroaches store uric acid at all life stages. In laboratory culture these insects have been reported to store the molecule in large quantites (Mullins and Cochran 1975; Potrikus and Breznak 1980a; Arquette et al. 2006). American cockroaches fed a high protein diet accumulated up to 31% uric acid (dry weight) (Mullins and Cochran 1975). Workers of eastern subterranean termites (Reticulitermes flavipes) measured 45% uric acid (dry weight) after 18 months in a laboratory, compared to 1% at the time of field collection (Potrikus and Breznak 1980a). However, the degree of uric acid accumulation in captivity varies widely by colony (Arquette et al. 2006).

In contrast to numerous studies demonstrating uric acid mobilization from fat body of cockroaches (Cochran 1975), only limited study has been performed in this area with termites (Breznak 2000). There is controversy as to whether uric acid accumulation in termite fat body is permanent or temporary. One theory holds that uric acid in fat body is transported via the Malpighian tubules to the hindgut, where it undergoes anaerobic digestion by uricolytic bacteria (Breznak 2000). Three species of uricolytic bacteria have been identified from R. flavipes hindgut, each converting uric acid to carbon dioxide, ammonia, and acetate (Breznak 1982 and 2000). Ammonia is subsequently used for manufacturing proteins in termite fat body (Potrikus and Breznak 1981; Breznak 2000). Uricolytic bacteria are present in sufficient numbers to provide the termite with significant dietary nitrogen (Potrikus and Breznak 1981). Most uric acid reaching the hindgut is digested by uricolytic bacteria, with little excreted in feces (Potrikus and Breznak 1980b). Feces from two additional subterranean termite species, Coptotermes formosanus and Reticulitermes virginicus, also contain little or no uric acid (Arquette and Rodriguez 2012).

A second theory submits termites permanently excrete uric acid to fat body, with nitrogen for amino acid synthesis predominantly provided by nitrogen-fixing spirochetes in the hindgut rather than uricolytic bacteria (Slaytor and Chappell 1994). From this view, uricolytic bacteria obtain uric acid from cannibalized termites (Slaytor and Chappell 1994; Breznak 2000). High uric acid accumulation in termite fat body in captivity may result from loss of nitrogen fixing ability, which has consistently been reported to occur soon after field collection (Slaytor 2000), as well as from digestion of cellular proteins during starvation (Slaytor and Chappell 1994).

Limited investigations have considered whether uric acid is involved in termite metabolism (Breznak 2000). However, for the closely related cockroach, the storage, mobilization, and metabolism of uric acid has been well documented (Cochran 1975). One of these studies (Mullins and Cochran, 1975) reported the body weight of female American cockroaches (Periplaneta americana) increased to 30% uric acid (dry weight) when the insects were fed a high protein diet, but levels dropped when starved (Mullins and Cochran 1975). Another study found that removal of blattabacteria, which are mutualistic uricolytic bacteria harbored in cockroach fat body, resulted in smaller and slower-maturing insects with diminished fecundity (Brooks and Richards 1956). Fat body of cockroaches with blattabacteria removed contained twentyfold higher levels of uric acid than control insects. Transplanting normal cockroach fat body into nymphs lacking blattabacteria caused the growth rate to increase. Blattabacteria-free regions of fat body appeared white due to accumulated uric acid, while transplanted fat body containing blattabacteria was translucent from lack of uric acid (Brooks and Richards 1956).

Blattabacteria synthesize uricase and xanthine dehydrogenase (Wren and Cochran 1987; Breznak 2000) and are capable of breaking down uric acid to pyruvate in vitro (Donnellan and Kilby 1967). The pathway by which blattabacteria metabolize uric acid begins with the conversion of uric acid to allantoin, followed by allantoic acid, glyoxylate, glycerate, and finally pyruvate (Donnellan and Kilby 1967). Mycetocytes, which are specialized fat body cells harboring uricolytic bacteria, have been identified from only one termite species, Mastotermes darwiniensis (Bandi and Sacchi 2000). As all other termites only harbor uricolytic bacteria in the hindgut, stored uric acid would not be digested unless mobilized to the hindgut. Mobilization of uric acid from termite fat body remains controversial because this has not been directly demonstrated by experiment (Slaytor and Chappell 1994).


The various classes of insect proteins include transport proteins, such as lipophorins in the hemolymph; regulatory proteins, including hormones of insects; defense proteins; and structural proteins, a variety of which are found in the cuticle (Neville 1975; Chapman 1998; Lenhninger et al. 2000). Over 100 kinds of proteins are known to occur in insect cuticle, functions of which include pigmentation and cuticular hardness (Hopkins and Kramer 1992; Chapman 1998).

Many insect proteins are mobile. For example, lipophorin is a conjugated protein that shuttles diacylglycerols via hemolymph (Chapman 1998). Urates may also be transported through hemolymph by proteins (Cochran 1975). Some proteins are transported to specific sites as required, such as cuticular proteins secreted from the epidermis during molting that originate in hemolymph (Reviewed in Neville 1975; Hopkins and Kramer 1992).

Although the core food of termites is lignocellulose, some protein is also obtained from the diet. Fungal tissue is a primary food source of subterranean termites (Waller and Curtis 2003) and is 20%-40% protein (dry weight). Other dietary sources of protein include dead protozoans obtained from proctodeal feeding, cannibalism and necrophagy, and coprophagy (Noirot and Noirot-Timothee 1969; Collins 1983; Hunt and Nalepa 1994; Nation 2002; Arquette et al. 2012). All termites produce proteolytic digestive enzymes (Collins 1983).

Insects predominantly synthesize amino acids in fat body (Chapman 1998). Synthesis of amino acids occurs at intermediate steps of glycolysis and the Krebs cycle (Lehninger et al., 2000). Lower termites such as the Formosan subterranean termite (Coptotermes formosanus) obtain essential amino acids synthesized by bacterial and protozoan symbionts (Mauldin et al. 1978). Intracellular bacterial symbionts of the German cockroach synthesize amino acids from urate precursors (McFarlane 1985). Amino acids are stored in hemolymph at 100-300 times the levels found in human blood, in excess of what is needed for protein synthesis (Chen 1985). In addition to comprising primary protein structure, amino acids also serve as precursor molecules. For example, glucogenic amino acids are converted to glucose during starvation (Lenhninger et al. 2000).

In aging adult insects, the rate of protein synthesis may decline due to an inability to repair or replace mitochondria (Sohal 1985). Therefore protein synthesis and subsequently enzyme activity is diminished (Sohal 1985; Brunk and Terman 2002).

The diet of termites is too low in nitrogen, amino acids, and protein for growth and egg development (Chen 1985). Nitrogen for adequate protein synthesis has been hypothesized to come from various sources. Ammonia produced from nitrogen fixation by bacterial symbionts is thought to provide termites with high enough levels of dietary nitrogen to compensate for the low nitrogen content of wood (Slaytor and Chappell 1994). Uricolytic bacteria harbored in the paunch of subterranean termites may provide the insect with significant amounts of dietary nitrogen for protein production (Breznak 2000). Dead protozoan symbionts provide termites with dietary nitrogen (Breznak 1982; Collins 1983). Chitin contains a significant amount of nitrogen, about 7 percent by weight (Waller and LaFage, 1987). It is always associated with proteins in nature, and may be a source of nitrogen that termites can metabolize (Breznak 1982; Chapman 1998). Chitinase activity has been measured from R. virginicus workers (Arquette, 2011). This chitinase likely was from the gut, as the termites assayed had full gut contents, indicating they were not molting. When an insect molts its cuticle, digested chitin, and protein is contained in molting fluid, which is either re-absorbed through epidermal cells, or taken in through the mouth or anus (Neville 1975; Breznak 1982).


Glycogen is synthesized in insects by the addition of glucose molecules to a glucose primer, a process catalyzed by glycogen synthetase (Nation 2002). Insects store glycogen in fat body and epithelial gut cells (Nation 2002). Glucose cleaved from glycogen does not need to be transported, so can be used as an immediate fuel source (Candy 1985). Glycogen of insects is converted to glucose during periods of vigorous activity such as flight (Chapman, 1998) as well as during starvation (Wigglesworth 1942; Satake et al. 2000). Glycogen is also a source of trehalose for insects (Candy 1985). Glucose is cleaved from glycogen branches by the activated form of glycogen phosphorylase. Glycogen phosphorylase b is a stored, inactive form of the enzyme, and is activated to glycogen phosphorylase a to release glucose or phosphorylated glucose units from glycogen (Friedman 1985).

Gluconeogenesis, or glucose synthesis from non-carbohydrate precursors, occurs in starved insects (Candy 1985). Gluconeogenesis is essentially the reverse of glycolysis, except for three irreverable steps of glycolysis which are bypassed by alternate pathways (Candy 1985; Lenhninger et al. 2000). Amino acids are the main precursor molecules for gluconeogenesis. Other precursors are pyruvate, lactate, and glycerol (Candy 1985). The conversion of amino acids to carbohydrates by insects was first demonstrated in the Aedes aegypti mosquito, with glycogen levels shown to increase when mosquitoes are fed alanine or glutamine (Wigglesworth, 1942).

Glycogen levels of termites remain relatively steady for laboratory reared R. flavipes workers (Arquette et al. 2006). However, glycogen content varies widely for termites collected from different environments. Glycogen levels measured from crude extracts of R. flavipes workers collected from a home lawn on an Atlantic barrier island were 25-fold or more higher compared with colonies from forested locations (Arquette and Forscher, 2006). The reason for this difference is unknown.


Lipids, mainly in the form of glycerides, provide insects with a major source of stored energy and metabolic water. At least 78% of insect lipids are triglycerides, with the proportion in fat body about 90% (Fast 1964; Beenakkers et al. 1985). Diglycerides in hemolymph are an important energy reserve for insects, comprising about 3% of the total lipid content (Bailey 1975; Beenakkers et al. 1985). Other lipids are integral components of waxes and other cuticular layers (Fast 1964; Nation 2002).

Insects obtain lipids from food and microbial symbionts (Chapman 1998) or synthesize it in fat body from carbohydrate or other precursor molecules (Bailey 1975). Acetyl CoA formed at the start of cellular respiration is used in long-chain fatty acid synthesis (Bailey 1975). Long-chain fatty acids and glycerols are esterified, forming triglycerides for storage in fat body (Bailey 1975). Lower termites also synthesize glycerides using acetate produced from fermentation by protozoan symbionts (Potrikus and Breznak 1981). Carter et al. (1972) reported that oleic acid makes up about 60% of the fatty acids of R. flavipes termites, while lineolic acid (18 carbon atoms) and palmitic acid (16 carbon atoms) each constitute about 10% of the total.

Insects cannot synthesize sterols (McFarlane 1985), and obtain it either from food or microbial symbionts of the gut (Chapman 1998). Clayton (1960) reported the production of a sterol, 22-dehydrocholesterol, from radiolabeled acetate fed to German cockroaches. He concluded this sterol was probably done by bacterial symbionts from an initial conversion of acetate to ergosterol, then to 22-dehydrocholesterol.

Most glycerides stored in fat body must be transported to other sites for oxidation (Candy 1985). Triglycerides are converted to diglycerides in fat body cells (trophocytes), which in turn are attached to lipophorins prior to entering hemolymph for transport (Bailey 1975). Lipophorins are reusable shuttles for transporting diglycerides in hemolymph to muscle where oxidation occurs (Beenakkers et al. 1985). When fatty acid chains of diglycerides are hydrolyzed, most of the remaining glycerol is transferred to hemolymph (Candy 1985).

Lipid content has been reported for termites maintained in laboratory culture and just after field collection (Arquette and Forschler, 2006; French et al., 1984; Carter et al., 1972). For R. flavipes workers, lipid content was measured between 2.3% and 6.7% from field populations (Carter et al. 1972; Arquette and Forschler, 2006). Lipid levels fluctuate for some termite species in laboratory culture. Fat content of Coptotermes acinaciformis doubled after 2 weeks in a laboratory, then dropped to low levels by 6 weeks (French et al., 1984). Arquette et al. (2006) reported 2-3 fold higher fat levels for R. flavipes workers from separate colonies after 2 weeks in laboratory culture. Lipid content of workers from two colonies was maintained or increased under laboratory conditions, but those of a third colony eventually dropped to low levels. For Mastotermes darwiniensis, French et al. (1984) reported lipid content remained relatively steady over 4 weeks in a laboratory.

Decreasing levels of lipid for termites in laboratory culture appears to result from starvation, when stored lipids would be the main energy source of insects. For starved American cockroaches, lipids provide 66% of the insect's metabolic energy, compared to 22% from glycogen and 11% from protein (Fast 1964). Mauldin et al. (1977) reported R. flavipes measured 5.3% lipid (wet weight) shortly after field collection, then decreased to 1.3-1.4% after partial or total elimination of microbial symbionts, which in turn caused the termites to starve.

Body Water

Water has various physiological roles in insects. These include regulation of body temperature, providing a hydrostatic skeleton for body support, and transport of food molecules, hormones, respiratory gases, and excretory products (Hadley 1994). Insects obtain water from drinking, food, water vapor absorption, and metabolism (Hadley 1994) and lose water through the cuticle, spiracles, mouth, and anus (Edney 1977; Hadley 1994). Factors influencing water loss in insects include warm temperature, low humidity, air currents, molting, egg production, nutritional state, abrasion of the waterproofing wax layer of the cuticle, and unknown effects of aging (Becker 1969; Edney 1977; Hadley 1994).

Water homeostasis is the maintenance of proper cellular osmolarity (Hadley 1994; Nation 2002). Without physiological regulation of proper body water levels, too much water would result in an insect's cells swelling and rupturing, while excessive water lost results in precipitation of soluble metabolites, causing irreversible harm (Hadley 1994). To maintain optimal body water levels, an insect must take in enough water to offset the amount lost through respiration, transpiration, or excretion (Hadley 1994).

Insects absorb water vapor through the mouth and rectum (Edney 1977; Hadley 1994). The critical equilibrium humidity is the relative humidity at which absorption of water vapor and dessication are balanced (Hadley 1994). Critical equilibrium humidity differs between insect species, ranging from about 43% to near saturated humidity (Edney 1977 and Hadley 1994). While body mass usually decreases from water loss, some terrestrial arthropods still lose water at saturated humidity levels (Hadley 1994).

Metabolic water is an important source of water for many insects including termites (Lee and Wood 1971; Nation 2002). Metabolic water is produced by insects from oxidation of carbohydrates, lipids, and proteins from food or stored forms (Hadley 1994; Nation 2002).

Fats are a major source of metabolic water (Nation 2002). For example one molecule of palmatic acid (16 carbons) produces 108 water molecules when oxidized, while the breakdown of one glucose molecule nets four water molecules (Nation 2002). About 1.07 ml of water is produced per gram of metabolized fat (Hadley 1994). This is about twice the amount of metabolic water produced by weight from the metabolism of carbohydrates, and about 2.5 times the rate produced from the oxidation of protein to urea (Hadley 1994). However, fat requires about 2.5 times more oxygen for metabolism than glucose (Hadley 1994). Also, water is required for the synthesis of fats (Hadley 1994).

Cuticular lipids of the epicuticle provide insects with a barrier against body water loss (Chapman 1998). However, some water still diffuses through the cuticle even if heavily sclerotized, and exact sites of cuticular water loss is unknown (Hadley 1994). Cuticular transpiration increases when lipids in the epicuticle are disrupted from temperature, chemicals, adsorption by dusts, and mechanical disruption (Hadley 1994).

Spiracles are a major site of body water loss in terrestrial arthropods (Hadley 1994). Most insects reduce transpiration through spiracles with valves that open and close (Hadley 1994). The valves open in the presence of elevated C[O.sub.2] in the trachea, as well as by the action of motor neurons of the central nervous system (Hadley 1994). Metabolism increases with higher temperatures and activity levels, thus requiring spiraclular valves to stay open longer to exchange respiratory gases, and allowing more water to be lost (Chapman 1998). Specific amounts of water lost through spiracles are unknown for most insect species due to the difficulty in obtaining measurement (Hadley 1994).

Another main source of insect body water loss is urine. Some water is from urine is absorbed through the ileum and rectum prior to excretion (Collins 1969; Nation 2002).

Land arthropods are vulnerable to dessication, and usually stay in moist environments (Cloudsley-Thompson 1988). By burrowing into the soil arthropods avoid the dessicating environment above the surface (Hadley 1994). Humidity in burrows comes from the surrounding soil or the tunnel's inhabitants (Hadley 1994). High humidity required for the survival of subterranean termites is maintained in the shelter tubes they construct (Lee and Wood 1971; Traniello and Leuthold 2000).

Hadley (1994) described a simple method of measuring insect body water content by obtaining the live weight followed by re-weighing after drying at around 60[degrees]C. The difference between live and dry weight is divided by the live weight to give the percent water content. Vacuum drying is another method for determining water content of arthropods (Hadley 1994). Most studies of the dessication tolerance of live insects measure water loss at 5% or less relative humidity (Hadley 1994). Sponsler and Appel (1990) reported lower survivable limits of dehydration for different C. formosanus and R. flavipes castes. R. flavipes workers placed in 0% relative humidity died when they lost 50% of their body water, which occurred after about 5 hours. R. flavipes lose water faster than other Reticulitermes species (Collins 1969).

While moist conditions are required for survival of subterranean termites, high humidity has been described as fatal to a drywood termite, Cryptotermes brevis (Walker) (Collins, 1969). When placed in 86% or higher relative humidity, this species swells up and dies due to an inability to transpire metabolically produced water to humid air (Collins 1969). There are three North American termite species that inhabit arid environments and cannot tolerate high humidity (Collins 1969).

Body water levels of termites have been considered as possible indicators of the vitality of termite populations (Arquette et al., 2006; French et al., 1984). French et al. (1984) did not regard body water levels as a reliable method for describing termite vitality. However Arquette et al. (2006) concluded R. flavipes workers are less healthy below a threshold of approximately 75% body water.


I greatly appreciate the assistance of Dr. Richard R. Mills, Virginia Commonwealth University (retired) for reviewing this manuscript and providing helpful suggestions.


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Tim J. Arquette (1,2)

(1) Mississippi State University Formosan Termite Research Laboratory, Poplarville, MS

(2) Mississippi State University Department of Biochemistry, Molecular Biology, Entomology, and Plant Pathology, Mississippi State, MS

(3) Mississippi State University Electron Microscope Center, Mississippi State, MS

(4) USDA Agricultural Research Service, Poplarville, MS

(5) State Chemical Laboratory, Mississippi State, MS

Corresponding author: Tim Arquette
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Date:Oct 1, 2013
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