Lipid and Carbohydrate Metabolism in Caenorhabditis elegans

Abstract

Lipid and carbohydrate metabolism are highly conserved processes that affect nearly all aspects of organismal biology. Caenorhabditis elegans eat bacteria, which consist of lipids, carbohydrates, and proteins that are broken down during digestion into fatty acids, simple sugars, and amino acid precursors. With these nutrients, C. elegans synthesizes a wide range of metabolites that are required for development and behavior. In this review, we outline lipid and carbohydrate structures as well as biosynthesis and breakdown pathways that have been characterized in C. elegans. We bring attention to functional studies using mutant strains that reveal physiological roles for specific lipids and carbohydrates during development, aging, and adaptation to changing environmental conditions.

INTEREST in lipid and carbohydrate metabolism has skyrocketed in recent years along with the human obesity and diabetes epidemics. All animals must eat to survive and reproduce, and dietary macronutrients are vital precursors required to build cellular material for growth and reproduction, even though overnutrition leads to a variety of modern diseases in humans. In recent years, there has been a growing appreciation of the dynamic roles of fat depots and the regulation of energy homeostasis, especially the interplay between the nervous system and various metabolic pathways. Caenorhabditis elegans is best known for powerful genetic and genomic analysis dissecting mechanisms of development, nervous system function, and aging. These studies have contributed to an appreciation of the roles of macronutrients and conserved metabolic pathways in these vital biological processes. The sections on Fatty Acids, Storage Lipids, Membrane Lipids, and Sterol and Prenol Lipids of this review outline lipid structures and metabolic pathways leading to the synthesis and metabolism of a wide range of lipids and describe specific functions for lipids in development and aging. Introduction to Carbohydrates and Functions and Metabolism of Monosaccharides introduce carbohydrate structures, metabolic pathways, and functions for simple and complex carbohydrates in development, aging, and protection from environmental stressors.

Lipids are small organic molecules that are insoluble in water, but are soluble in organic solvents. Biochemically, they originate entirely or in part from carbanion-based condensations of thioesters, forming fatty acids, which are components of triacylglycerols (TAGs), phospholipids, and sphingolipids; or by carbocation-based condensation of isoprene units, forming isoprenol derivatives including sterols (Fahy et al. 2009) (Figure 1). C. elegans has the potential to synthesize thousands of distinct lipid species, many of which have yet to be characterized.

Figure 1.

Structures of C. elegans lipid classes. (A) Fatty acids. (B) Sterol and prenol lipids. (C) Glycerophospholipids. Fatty acyl groups are outlined in red, the glycerol backbone in pink, and the phosphate group is outlined in blue, head group structures abbreviated as X are shown below. (D) A PE structure shows an ether linkage at C1 and an ester linkage at C2. (E) TAG molecule showing the fatty acyl chains outlined in red. (F) Sphingolipid structure showing the branched long chain base, head group structures abbreviated as X are shown below.

Lipids perform many essential functions in the cell. Their highly reduced state renders them efficient energy storage molecules. They are the hydrophobic units of bilayers that form cellular and organellar membranes, and they are potent signaling molecules. In membranes, the diversity of lipid species contributes to cellular and organellar functions. For example, in most organisms, the outside leaflet of the plasma membrane is composed of entirely different lipid species than the inside leaflet (Shevchenko and Simons 2010). Plasma membrane lipid composition differs among various tissues, as does lipid composition of endoplasmic reticulum (ER), mitochondria, and other cellular organelles (van Meer et al. 2008). Because the distinct lipid composition is actively maintained by ATP-consuming enzymes, it is believed that the lipid asymmetry is important for distinct cellular functions. Additionally, specific lipid composition influences membrane protein folding and topology (Vitrac et al. 2015). Questions regarding functions of distinct lipids in membranes or lipid requirements for protein functions are only beginning to be addressed in C. elegans. To date, research in the C. elegans lipid field has been ongoing to characterize lipid biosynthetic genes, regulatory genes affecting lipid synthesis, storage, and breakdown, as well as the neurological and developmental consequences that occur when specific lipids are not synthesized or correctly or when lipid homeostasis is incorrectly regulated.

Fatty Acids, Storage Lipids, Membrane Lipids, and Sterol and Prenol Lipids will follow the LIPID MAPS classification scheme (Fahy et al. 2009) to review the current state of knowledge concerning the biosynthesis, breakdown, and functions of specific lipid classes in C. elegans. Although 471 putative lipid metabolism genes have been identified based on the KEGG database and homology to mammalian lipid metabolism (Zhang et al. 2013), this review will focus on genes that have been experimentally verified based on biochemical assays, changes in nematode lipid composition in loss of function strains, or functional phenotypes in C. elegans. Roles for specific lipids have often been identified in forward or reverse genetic screens, which offer unbiased determinations of functions of specific lipid classes (Han 2015; Zhang et al. 2015). Methods for analysis of specific lipids will be mentioned briefly in the appropriate sections, and more information regarding lipidomics analysis methods can be found in a recent comprehensive review (Witting and Schmitt-Kopplin 2016).

Fatty Acids

Characteristics of C. elegans fatty acids

Fatty acids are carboxylic acids with long aliphatic chains, which in C. elegans lipids contain 14–20 carbons (Tanaka et al. 1996; Watts and Browse 2002) (Figure 1A). Fatty acids are the building blocks and precursors for storage lipids (TAGs), membrane lipids (phospholipids and sphingolipids), and signaling lipids (fatty acyl amides, eicosanoids, and others). Fatty acids can be saturated, containing no double bonds, monounsaturated, containing one C–C double bond, or polyunsaturated, containing two or more C–C double bonds. C. elegans double bonds are in the “cis” configuration, and are methylene interrupted, meaning double bonds are spaced along the carbon chain in intervals of three carbons. Fatty acids are named by common (trivial) names, systematic chemical names, and chemical abbreviations. For example, the systematic name of linoleic acid is 9,12 octadecanoic acid, which is abbreviated 18:2 (18 carbons with two double bonds). Further information regarding double bond location is given by the designation n-6 or ω6, which indicates the position of the terminal double bond. Thus, using the abbreviations 18:3n-3 and 18:3n-6, α-linolenic (9,12,15-octadecatrienoic) can be distinguished from γ-linolenic (6,9,12-octadecatrienoic). C. elegans differs from most animals in that it has the capacity to synthesize a range of monomethyl and polyunsaturated fatty acids starting with acetyl-CoA or isobutyryl-CoA as precursors. In contrast, most animals must obtain polyunsaturated fatty acids from their diet, because they do not possess the enzymes to convert monounsaturated fatty acids to polyunsaturated fatty acids (Wallis et al. 2002).

Methods to analyze fatty acid composition

Fatty acids from worm populations are commonly analyzed using acidic methylation to produce fatty acid methyl esters, which are separated from each other by gas chromatography (GC), and detected with either mass spectrometry (MS) or flame ionization detection (FID) (Watts and Browse 2002). This is a sensitive method, because use of a polar column such as SP-2340 or Omegawax aids in the separation of fatty acyl isomers with identical chemical composition but different double bond positions. Because C. elegans synthesizes and accumulates several isomers of 18:1 and 20:4, it is important to use authentic standards to distinguish among isomers during analysis.

When using single quad GC/MS analysis to characterize the fatty acid composition of distinct lipid classes, such as triglycerides or phospholipids, the lipids must first be separated before GC/MS analysis of fatty acids. One method to separate neutral, glycerophospholipids, and sphingolipids is using a solid state column (Perez and Van Gilst 2008). Another method, thin layer chromatography (TLC), is useful because distinct phospholipid species as well as neutral lipids can be separated using a two-solvent system (Shi et al. 2013). After solid state or TLC separation, the fatty acyls can then be derivatized using acidic methylation for GC/MS analysis. The disadvantage of the solid state or TLC method for lipid separation is that it requires large amounts of material, typically 10,000 young adult C. elegans per separation.

Fatty acid composition of intact lipids can be analyzed by liquid chromatography/mass spectrometry (LC/MS) (Entchev et al. 2008; Castro et al. 2012). The use of triple quad mass spectrometers and shotgun lipidomics allows lipid analysis without separating lipid classes (Witting and Schmitt-Kopplin 2016). In C. elegans, due to the large number of fatty acyl classes, this has the potential to generate over 1000 distinct lipid species (Entchev et al. 2008; Shi et al. 2016; Witting and Schmitt-Kopplin 2016). Methods to analyze intact lipids are required to identify sphingolipids and ether-linked lipids, because the acidic methylation technique does not liberate the fatty acyl of the sphingosine base or the alkyl group on the ether-linked lipid. A disadvantage of intact lipid analysis methods for C. elegans fatty acids is the inability to identify distinct isomers. For example, the cyclopropane fatty acids derived from Escherichia coli are simply identified by their mass as 17:1 or 19:1, which has propagated the myth that C. elegans synthesizes a considerable amount of odd chain fatty acids, when in fact these represent mmBCFA C17iso or dietary cyclopropane fatty acids (C17∆ and C19∆). Only very small amounts of straight, odd-chain fatty acids accumulate in C. elegans lipids. Another disadvantage of intact lipid analysis methods is that all species are identified based on their total mass, making the identification of specific fatty acyl groups ambiguous; for example, PC-40:8 could contain one 20:3 and one 20:5, or two 20:4 (which could be either the 20:4n-3or the 20:4n-6 isomers).

C. elegans fatty acids are synthesized de novo, or are derived from dietary fatty acids

Stable isotope labeling studies using dietary ¹³C have been used to study synthesis and turnover of fatty acids (Perez and Van Gilst 2008; Dancy et al. 2015); ∼7–20% of C. elegans fatty acids are synthesized de novo from acetyl-CoA, with the exception of the monomethyl fatty acids, which cannot be derived from E. coli precursors and are 100% synthesized when C. elegans feeds on E. coli OP50 (Perez and Van Gilst 2008). The remaining fatty acids are incorporated or modified from E. coli fatty acids. Fatty acyl turnover is rapid, with ∼4.5% of membrane fatty acid acyl components of phospholipids being turned over every hour, and 2.7% of neutral (storage) lipids being turned over every hour (Dancy et al. 2015). This means that the majority of fatty acyl components of membrane lipids are replaced daily. When grown on nematode growth media (NGM) plates to stationary phase, E. coli OP50 and HT115 membranes are composed of ∼37% saturated fatty acid (31% 16:0, 6% 14:0, and trace amounts of 18:0), 11% monounsaturated fatty acids, (trace amounts of 16:1n-7 and 11% 18:1n-7), and 49% cyclopropane fatty acids (Brooks et al. 2009). The E. coli HB101 strain contains considerably more monounsaturated fatty acids, including 16:1n-7 and 18:1n-7, and reduced amounts of cyclopropane fatty acids (Brooks et al. 2009). In worms, the relative composition of cyclopropane fatty acids and 18:1n-7reflects the dietary E. coli source (Brooks et al. 2009). Neither strain of E. coli produces monomethyl branch chain fatty acids or polyunsaturated fatty acids.

The de novo synthesis of fatty acyl chains using the two-carbon subunit acetyl-CoA is achieved by the activity of fatty acid synthase (FAS). This is a multi-enzyme protein encoded by one large gene (fasn-1) and contains six catalytic activities, including beta-ketoacyl synthase, acetyl/malonyl transacylase, beta-hydroxyacyl dehydratase, enoyl reductase, β-ketoacyl reductase, acyl carrier protein, and thioesterase (Chirala and Wakil 2004). This reaction requires NADPH, and the condensation of malonyl-CoA and the following reaction series are repeated seven times to generate palmitate (16:0) (Figure 2).

Figure 2.

Pathway of de novo fatty acid synthesis in C. elegans. Enzyme names and activities are enclosed in ovals. ELO, elongase; C13iso, 11-methyldodecanoic acid; C15iso, 13-methyltetradecanoic acid; C17iso, 15-methylhexanoic acid; SA, stearic acid; OA, oleic acid; LA, linoleic acid; ALA, alpha linoleic acid; GLA, gamma linoleic acid; STA, stearidonic acid; DGLA, dihommo gamma linoleic acid; ETA, eicosatetraenoic acid; AA, arachidonic acid; EPA, eicosapentaenoic acid.

The rate-limiting step of de novo fatty acid synthesis is the generation of malonyl-CoA by acetyl-CoA carboxylase (ACC), encoded by the pod-2 gene in C. elegans (Rappleye et al. 2003). The ACC enzyme requires biotin, and uses energy generated by the hydrolysis of ATP to catalyze the carboxylation of acetyl-CoA to malonyl-CoA (Waite and Wakil 1962). Malonyl CoA is the substrate for each round of fatty acid synthesis, with CO₂ being liberated during each condensation reaction. After de novo synthesis of 16:0, malonyl-CoA is used as the substrate for the elongation of fatty acids, which uses separate condensation, reductase, dehydratase, and thiosterase enzymes, including the ELO-1,ELO-2, and ELO-3 synthases (Watts and Browse 2002; Kniazeva et al. 2004; H. Zhang et al. 2011), the LET-767 β-ketoacyl dehydratase (Entchev et al. 2008), and likely the HPO-8 β-hydroxyacyl dehydratase, and the ART-1 enoyl reductase.

Monomethyl branch chain fatty acids serve as precursors of sphingolipids

An understudied family of fatty acids containing a methyl branch is synthesized by bacteria and is a significant component of the human diet, especially of consumers of dairy and ruminant meat (Ran-Ressler et al. 2014). C. elegans synthesize monomethyl branch chain fatty acids and these play important functions in the organism (reviewed in Han 2015). The pathway to synthesize monomethyl branched chain fatty acids (mmBCFAs) requires ACC and FAS, similar to the synthesis of straight chain fatty acids, except that the starting material is isovaleryl-CoA, which is derived from the branched chain amino acid leucine (Kniazeva et al. 2004) (Figure 2). The biosynthesis pathway involves the branched-chain ketoacid dehydrogenase complex (BCKDC), fatty acid synthase (FASN-1), acetyl-coA carboxylase (POD-2), fatty acyl elongases (ELO-5 and ELO-6), the LET-767 β-ketoacyl dehydratase, and an acyl CoA synthetase (ACS-1), After five rounds of condensation of malonyl-CoA by FAS, the ¹³C-iso fatty acids are elongated by activities of elo-5, elo-6, and let-767 to form 13-methyl myristic acid (C15iso) and 15-methyl hexadecanoic acid (C17iso), which accumulate in C. elegans storage lipids, glycerophospholipids, and sphingolipids (Kniazeva et al. 2004; Entchev et al. 2008; Zhu et al. 2013; Jia et al. 2016).

The elo-5 mutants, which lack mmBCFA, arrest as L1 larvae with a phenotype similar to starvation arrest (Kniazeva et al. 2004, 2008). Involvement with the insulin signaling pathway and the correlation with 17iso in TAG and fat stores indicate that accumulation or synthesis of these fatty acids could indicate nutrient status (Kniazeva et al. 2008; Brooks et al. 2009). The larval arrest phenotype is suppressed for one generation by mutations in tat-2, which encodes a P-type ATPase/flippase that likely allows for the translocation of lipids within the bilayer or between organelles (Seamen et al. 2009). In addition, a mutation in a peroxisomal regulatory gene partially restored development as well as mmBCFA levels (Wang et al. 2013).

While C15iso and C17iso are found in several lipid species, including TAGs, PC, and PE, an important role for mmBCFA are as structural components of sphingolipids. All C. elegans sphingolipids contain C17iso as the long chain acyl component of the sphingoid base (Chitwood et al. 1995; Zhu et al. 2013). During sphingolipid biosynthesis, the mmBCFA C15iso condenses with the amino acid serine to form an unusual 17-carbon branched chain sphingoid base. This is different from mammals, which typically synthesize the sphingoid base from palmitic acid (16:0) and serine. The specialized sphingoid base is important for function, because the developmental phenotypes of elo-5 mutants depend on C17iso base of sphingolipids (Zhu et al. 2013). More information on sphingolipid synthesis and function is found below in Sphingolipid biosynthesis and function.

Dietary and endogenous synthesized monounsaturated fatty acids regulate fat accumulation and longevity

Monounsaturated fatty acids are abundant in C. elegans lipids, especially cis-vaccenic acid (18:1n-7), which can be directly obtained from the E. coli diet. Alternatively, monounsaturated fatty acids can be produced from saturated fatty acid precursors by desaturation. Desaturase enzymes are diiron proteins, which use conserved histidine residues to align the iron and cytochrome b₅ to extract protons from acyl chains (Shanklin et al. 1994). Desaturase enzymes are characterized by specificity for the position along the acyl chain. Stearoyl-CoA desaturase, or ∆ 9 desaturase, insert the first double bond at carbon 9 into a saturated fatty acyl chain, and are among the most extensively studied desaturases in mammals. C. elegans encodes three ∆9 desaturases. The FAT-5 desaturase is specific for 16:0, producing 16:1n-7, which can then be elongated to 18:1n-7 (cis-vaccenic acid) (Watts and Browse 2000). The FAT-6 and FAT-7 desaturases mainly act on 18:0, producing oleic acid, 18:1n-9 (Watts and Browse 2000). Unlike in mammals, in C. elegans 18:1n-9 can be further desaturated and elongated to form polyunsaturated fatty acids, thus 18:1-9 only accumulates to a small degree in C. elegans membranes and neutral lipids, even though it is a much more significant component of mammalian membranes (Wallis et al. 2002). In spite of dietary MUFA provided by the E. coli diet, endogenous production of monounsaturated fatty acids is essential, as the fat-5;fat-6;fat-7 triple mutant strain is lethal (Brock et al. 2006).

In mammals and in C. elegans, reduction of ∆9 desaturase activity leads to reduced fat accumulation, and thus ∆9 desaturases are considered to be lipogenic enzymes (Ntambi et al. 2002; Brock et al. 2007; Sampath and Ntambi 2011). In mammals, monounsaturated fatty acids, such as oleic acid, are the preferred substrate for acyl transferase enzymes that synthesize triglycerides (Cases et al. 2001). However, in C. elegans, triglyceride fatty acids consist mainly of dietary fatty acids obtained from E. coli (Perez and Van Gilst 2008; Brooks et al. 2009). In both mammals and C. elegans, stearoyl-CoA desaturase knockdown leads to increased fatty acid oxidation (Flowers and Ntambi 2008; Shi et al. 2013), which can mitigate lipotoxicity caused by excess saturated fatty acids (Listenberger et al. 2003).

Because the ∆12 desaturase activity allows polyunsaturated fatty acids (PUFAs) to be synthesized from monounsaturated fatty acids, the ∆9 desaturases are vital for providing precursors for further desaturation and elongation. In addition to this role, the fat-6;fat-7 double mutants show reduced turnover of fatty acyl groups, which may be necessary for optimal healthspan and longevity (Dancy et al. 2015). In a more specific context, the ∆9 desaturases have been implicated in a signaling event that regulates long lifespan in nematodes lacking a germ line. The long lifespan depends on two nuclear hormone receptors, NHR-80 and NHR-49, both of which regulate ∆9 desaturase activity (Van Gilst et al. 2005a; Brock et al. 2006; Goudeau et al. 2011; Ratnappan et al. 2014). In addition to ∆9 desaturation, the longevity of germline-less nematodes depends on a functional autophagy pathway, which is described in Neutral lipid catabolism. Finally, chromatin modification in the germ line also influences ∆9 desaturation, and mutants defective in H3K4me3 transferase activity have a long lifespan and increased monounsaturated fatty acid levels (Han et al. 2017).

Saturated fatty acid levels and temperature adaptation

C. elegans lipids contain relatively low levels of saturated fatty acids compared to mammalian cells, with ∼5% of fatty acids in total lipid extracts consisting of 16:0, 6% 18:0, and small amounts (<2%) of 14:0 and 20:0 (Tanaka et al. 1996; Watts and Browse 2002). The addition of just one double bond to a saturated fatty acyl chain changes the melting temperature of the fatty acid drastically. The homeoviscous theory of membranes adaptation states that membrane saturation levels are regulated in order to adapt to environmental temperature changes (Sinensky 1974). It appears that low levels of saturated fat found in C. elegans lipids likely allow for growth in cool environments, because the ∆9 desaturase gene fat-7 is induced upon transfer to cold temperature (Murray et al. 2007), and C. elegans fat-6;fat-7 double mutants, which accumulate high levels of saturated fatty acids, are cold sensitive (Brock et al. 2007), as are fat-5;fat-7(RNAi) (Murray et al. 2007). Other cold-sensitive and cold-tolerant mutant strains also influence the activity of ∆9 desaturation, including the phosphoinositide-3 kinase mutant age-1 (Savory et al. 2011), the adiponectin receptor homolog paqr-2 (Svensson et al. 2011; Svensk et al. 2013), and iglr-2 (Svensk et al. 2016). Mutations in the acyl-CoA dehydratase gene acdh-11 were found to suppress paqr-2 by upregulating fat-7 (Ma et al. 2015). Structural studies of ACDH-11 suggest that short chain fatty acids become sequestered by binding to ACDH-11, and that the sequestration of these fatty acids inhibits expression of fat-7, reducing fatty acid desaturation such that membranes become more saturated in response to increased temperature (Ma et al. 2015).

Cyclopropane fatty acids are obtained from the diet and are not essential

C. elegans grown on NGM plates with E. coli (OP50) contain large amounts of cyclopropane fatty acids in their lipids. These fatty acids are formed in bacteria by addition of a methylene group across the double bond of an unsaturated fatty acid by the enzyme CFA synthase, and they accumulate during stationary phase of E. coli growth (Grogan and Cronan 1997). E. coli strains with fewer cyclopropane fatty acids accumulate more unsaturated fatty acids, and vice versa (Brooks et al. 2009). The two cycloproane fatty acids found in E. coli and C. elegans are cis-9,10-methylenehexadecanoic acid, abbreviated C17∆, and cis-11,12-methyleneoctadecanoic acid, abbreviated C19∆ (Figure 1A). In C. elegans lipids, the majority of cyclopropane fatty acids are components of triglyceride storage lipids, and are incorporated in membrane phospholipids to a lesser degree (Table 1). C. elegans grown on CFA synthase mutants of E. coli are viable and do not contain cyclopropane fatty acids in lipid extracts (Kaul et al. 2014). This demonstrated that C. elegans do not synthesize cyclopropane FAs, and that cyclopropane FA are not essential for C. elegans. The same study also showed that C. elegans recover more efficiently from dauer arrest in the absence of cyclopropane fatty acids than in their presence (Kaul et al. 2014), suggesting that, for some processes, dietary cyclopropane fatty acids may be detrimental.

Table 1. Fatty acid composition of isolated lipid classes in wild type C. elegans.

	15iso	16:0 (%)	17iso	18:0DMA	17∆ (%)	18:0 (%)	18:1n-9 (%)	18:1n-7 (%)	18:2 (%)	19∆	20:3 (%)	20:4	20:4n-3	20:5
PC	2.0%	2.3	1.5%	0.1%	8.4	2.2	2.3	8.4	6.4	15.0%	4.7	3.0%	7.2%	32.8%
PE	1.8%	7.7	6.3%	7.5%	9.2	14.3	2.1	12.9	6.3	12.4%	3.4	1.3%	3.5%	9.3%
PI	0.7%	4.3	1.3%	0.1%	1.6	26.8	1.9	8.6	1.9	13.5%	1.2	1.8%	1.6%	35.1%
PS	0.5%	5.6	3.8%	0.4%	3.1	26.4	3.0	7.7	3.3	5.0%	6.6	3.2%	5.1%	26.1%
CL	ND	6.5	ND	ND	2.4	8.9	1.3	4.9	2.8	ND	21.5	ND	44.9%	4.4%
TAG	5.2%	4.5	3.9%	0.2%	32.2	2.9	6.0	8.5	2.9	23.5%	1.7	0.5%	0.9%	2.6%
Total lipids	3.3%	4.1	3.5%	2.0%	20.3	5.2	3.9	9.9	4.6	17.9%	3.4	1.6%	4.0%	12.5%

Sources for fatty acid composition: Synchronized, young adult stage N2 hermaphrodites grown at 20° were used for these measurements. For all lipids except CL, values are average from Shi et al. (2013), (2016); for CL, values are from Hou et al. (2014). ND, Not detected; 15iso, 13-methyltetradecanoic acid; 17:iso, 15-methylhexanoic acid; 18:0DMA, 18:0 dimethylacetal, formed from 18:0 with vinyl ether linkage; 17∆, cis-9,10-methylenehexadecanoic acid; 19∆, cis-11,12-methylene octadecanoic acid, PC, phosphatidylcholine; PE, phosphatidylethanolamine; PI, phosphatidylinositol; PS, phosphatidylserine; CL, cardiolipin; TAG, triacylglycerols.

PUFAs are required for growth, reproduction, and neurotransmission

A unique aspect of C. elegans fatty acid metabolism is the ability to synthesize a wide range of PUFAs de novo. There are four fatty acid desaturases that convert 18:1n-9 into a range of C18 and C20 PUFAs. The FAT-2 (∆12) desaturase catalyzes the conversion of oleic acid (18:1n-9) into the PUFA linoleic acid (18:2n-6), and the FAT-1 desaturase catalyzes the conversion of 18-carbon and 20-carbon omega-6 fatty acids into omega-3 fatty acids (Figure 2) (Spychalla et al. 1997; Peyou-Ndi et al. 2000; Watts and Browse 2002). The FAT-1 and FAT-2 desaturases are similar to the plant desaturases ∆12 and omega-3 desaturases encoded by FAD2, FAD3, FAD6, FAD7, and FAD8, and homologs are not normally present in animals, although they have been identified in other nematode species. Two C. elegans desaturases, the ∆5 desaturase FAT-4 and the ∆6 desaturase FAT-3 are homologs of human FADS1 and FADS2, respectively (Napier et al. 1998; Watts and Browse 1999; Marquardt et al. 2000). The fatty acid desaturases function as part of an electron transport pathway requiring cytochrome b₅ and a cytochrome b₅ reductase to extract two protons from an acyl chain. Fatty acid composition analysis indicates that hpo-19 and T05H4.4 encode cytochrome b₅ reductases that act in this pathway (Y. R. Zhang et al. 2016).

The sum of the C18 and C20 PUFAs comprise ∼28% of fatty acids in total worm lipids (Shi et al. 2013). PUFAs are not distributed evenly in worm lipids, for example, triacylglycerol storage lipids contain relatively low levels of PUFAs (9% of total fatty acids), while phosphatidylcholine (PC) consists of 56% PUFAs (Shi et al. 2013) (Table 1). The most abundant PUFA in all C. elegans lipid classes is eicosapentaenoic acid (EPA, 20:5)—the final fatty acid in the desaturation pathway (Figure 1A and Figure 2). Overall, PUFA levels, especially 20:5, increase during fasting (Van Gilst et al. 2005b).

C. elegans severely deficient in PUFAs are viable, although they exhibit many growth, reproduction, and neurological defects (Watts and Browse 2002). The ∆12 desaturase fat-2 mutants contain large amounts of oleic acid and only 1% PUFA. These mutants display many defects, including slow growth, very low brood size, and uncoordinated movement. The ∆6 desaturase fat-3 mutants contain C18 PUFAs, but not C20 PUFAs. They grow better and display higher brood size than fat-2 mutants, but show many defects compared to wild type (Lesa et al. 2003; Watts et al. 2003). The fat-3 mutants have uncoordinated movement, which may be due to depleted synaptic vesicles and reduction of neurotransmitter release (Lesa et al. 2003). The fat-3 mutants are defective in chemosensation, olfactory adaptation, touch sensitivity, and adaptation to alcohol (Kahn-Kirby et al. 2004; O’Halloran et al. 2009; Raabe et al. 2014; Vasquez et al. 2014). The fat-4 and fat-1 mutants contain different species of PUFAs and greatly different ratios of omega-6 and omega-3 species, although growth, movement, and reproduction are essentially normal (Watts and Browse 2002). Therefore, many reproductive and neurological functions of PUFAs can be fulfilled by either omega-6 or omega-3 fatty acids. However, dietary omega-6 fatty acids, such as dihommogamma linolenic acid (DGLA, 20:3) adversely affect germ cell maintenance, leading to sterility above certain concentrations of dietary DGLA (Watts and Browse 2006; Webster et al. 2013). More details regarding the functions of PUFAs in C. elegans can be found in a recent review (Watts 2016).

Polyunsaturated fatty acids as precursors for signaling molecules

In response to signaling events, PUFAs are liberated from membranes by phospholipases and further metabolized to form powerful short-range signaling molecules, collectively termed eicosanoids. Eicosanoids are formed by several pathways that introduce molecular oxygen in highly stereo specific reactions. In mammals, eicosanoids are formed by cyclooxygenase, lipoxygenase, and cytochrome P450 (CYP) enzymes (Funk 2001; Spector and Kim 2015). While early papers noted that C. elegans do not contain homologs of cyclooxygenase enzymes that are responsible for synthesizing prostaglandins, several F-series prostaglandins have been identified in worm lipid extracts using mass spectrometry (Hoang et al. 2013). These F-series prostaglandins are synthesized from several C20PUFA precursors in the female germline and they function to attract sperm to oocytes (Kubagawa et al. 2006; Hoang et al. 2013). While the precise mechanism of prostaglandin formation has not been elucidated, it is likely that in both C. elegans and mammals, prostaglandins are synthesized using a cyclooxygenase-independent pathway (McKnight et al. 2014).

CYP enzymes produce eicosanoids from PUFAs by way of two pathways. One pathway uses ω-hydroxylation to convert PUFAs such as arachidonic acid to hydroxyeicosatetraenoic acids (HETEs), and another uses epoxygenase activity to generate epoxyeicosatrienoic acids (EETs); both pathways are active in C. elegans and the eicosanoid products of PUFAs have potent biological effects (Kulas et al. 2008). CYP-33E2 is expressed in the pharynx, and it produces EETs that are required for efficient pharyngeal pumping (Kosel et al. 2011). CYP-33E2 also acts on DGLA (20:3) and mediates the sterility that occurs after administration of dietary DGLA (Deline et al. 2015). Finally, the epoxygenase activity of CYP-13A12 is required for a behavior response to reoxygenation after oxygen deprivation (Ma et al. 2013; Keller et al. 2014).

Storage Lipids

Synthesis of neutral glycolipids

In animals, plants, and fungi, fatty acids are stored in the form of TAGs, which are efficient energy storage molecules due to their reduced state (Figure 1E). De novo synthesized or dietary fatty acids are esterified onto glycerol molecules to produce TAGs for storage. In addition, other nutrients, such as carbohydrates and amino acids, can be broken down into acetyl CoA for de novo fatty acid synthesis, and other metabolites can enter the glycerol-3-phosphate pathway to form diacylglycerols (DAG) and TAGs (Figure 3). Importantly, phospholipid and neutral lipid synthesis are linked, because both biosynthetic pathways share a common intermediate: DAG. C. elegans contains homologs of the glycerol-3 phosphate acyl transferase (GPAT) and the lysophosphatidic acid acyl transferase (LPAAT) families, which are involved in the transfer of acyl groups onto glycerol-3 phosphate and lysophosphatidic acid to synthesize phosphatidic acid (PA). Biochemical activity or substrate specificities of the many C. elegans GPAT and LPAAT homologs have not been determined. PA is then dephosphorylated by the lipin homolog LPIN-1 to generate DAG (Golden et al. 2009). TAGs are synthesized from DAGs by the activity of diacylglycerol acyl transferases (DGATs), one isoform of which is encoded by dgat-2 (Xu et al. 2012). Fatty acyl-CoAs for TAG synthesis are either synthesized de novo, or obtained from the diet. The E. coli (OP50) strain does not synthesize or store TAGs; however, bacterial membranes are composed of phospholipids composed of fatty acids. These fatty acids are liberated from phospholipids during the digestion process, after which they are absorbed and are either directly incorporated into TAG or phospholipids, or further modified before incorporation into C. elegans lipids.

Figure 3.

Biosynthesis of neutral lipids and glycerophospholipids. Substrates are depicted in yellow ovals, minor lipids and intermediates are depicted in blue ovals, major lipid products are depicted in red ovals, and enzymes are shown in purple ovals. G3P, glucose-3 phosphate; FA-coA, fatty acyl coenzymeA; Ino, inositol; Ser, serine; Etn, ethanolamine; SAM, s-adenosyl methionine; Cho, choline; CTP, cytidine triphosphate; LPA, lysophosphatidic acid; PA, phosphatidic acid; DAG, diacylglycerol; CDP-DAG, cytidine diphosphate diacylglycerol; PG, phosphatidylglycerol; CDP-Etn, cytidine diphosphate ethanolamine; CDP-Cho, cytidine diphosphate choline; P-Etn, phosphoethanolamine; P-Cho, phosphocholine; TAG, triacylglycerol; PI, phosphatidylinositol; PS, phosphatidylserine; CL, cardiolipin; PE, phosphatidylethanolamine; PC, phosphatidylcholine; GPAT, glycerol-3 phosphate acyl transferase; LPAAT, lysophosphatidic acid acyl transferase; DGAT, diacylglycerol acyl transferase; EK, ethanolamine kinase; CK, choline kinase.

While digestion has not been actively studied in C. elegans, the intestine is acidic, and many genes encoding secreted lipases have been identified as intestinally expressed genes (McGhee 2007). Digestion begins with the disruption of bacteria in the pharynx, and further disruption of cells by secreted lysozymes. Because intact phospholipids cannot be imported across membranes, lipids are absorbed after dietary bacterial membrane lipids are acted on by secreted phospholipase enzymes that hydrolyze phospholipids into lysophospholipids and free fatty acids, which can then be imported across membranes into intestinal cells. Studies of the pept-1 mutants, which are defective in the transport of dietary peptides into the intestine, reveal a relationship between the peptide transporter PEPT-1, the sodium proton exchanger NHX-2, and fatty acid intake, which, in PEPT-1 deficiency, leads to greatly increased fat accumulation (Spanier et al. 2009). Digestion and lipid assimilation are very efficient processes, given that, at the peak of egg laying, each C. elegans hermaphrodite consumes several million bacteria, and converts her entire body mass into embryos every day (McGhee 2007).

Lipid droplet and yolk lipoprotein structure

TAGs are the major component of lipid droplets, which accumulate in the intestine, epidermis, and germ line of C. elegans. Electron microscopy and lipidomic analysis of lipid droplets reveal that lipid droplets consist of a core of TAGs, surrounded by a monolayer of phospholipid and protein (Zhang et al. 2010b; Vrablik et al. 2015). TAGs are also a component of yolk, similar to mammalian lipoproteins, which is synthesized in the intestine and transferred to developing oocytes during reproduction (Kimble and Sharrock 1983; Hall et al. 1999). The major protein components of yolk are the vitellogenin proteins YP170A, YP170B, YP115, and YP88, which are encoded by the genes vit-1, -2, -3, -4, -5, and -6 and exist as two distinct lipoproteins complexes (Spieth and Blumenthal 1985; Heine and Blumenthal 1986; Sharrock et al. 1990). Yolk granules are estimated to be ∼85% protein and 15% lipid by mass (Sharrock et al. 1990) and are ∼0.5–1 μm in size. Yolk is synthesized in intestinal cells, exocytosed, and taken up by RME-2 receptor in oocytes, and also by coelomocytes (Grant and Hirsh 1999). Lipidomic analysis revealed that yolk contains relatively greater amounts of phospholipids than TAGs (Sharrock et al. 1990; Kubagawa et al. 2006). In contrast, lipid droplets contain a much higher relative amount of TAGs compared to phospholipids, and also are composed of much less protein compared to yolk granules (Vrablik et al. 2015). Lipid droplet are typically 0.5–2 μm in size, but can reach sizes up to 10 μm (Li et al. 2016). Neither yolk nor lipid droplets contain measurable amounts of cholesterol or cholesterol esters (Kubagawa et al. 2006; Vrablik et al. 2015). The relationship between the biosynthesis of yolk and lipid droplets is still unclear, and it is unknown how lipid droplets are transported to, or generated in, the germ line. Interestingly, vitellogenin proteins have been identified in C. elegans lipid droplet proteomes (Zhang et al. 2012; Vrablik et al. 2015), and a recent study reported that VIT-2 is involved in the transfer of dietary lipids to lipid droplets (Wang et al. 2016).

The major lipid droplet protein is PLIN-1, previously called MDT-28 (Zhang et al. 2012; Chughtai et al. 2015; Na et al. 2015; Vrablik et al. 2015). PLIN-1 is the closest C. elegans homolog of perlipins, which are a family of major structural lipid droplet proteins found in a wide variety of animals and fungi (Brasaemle 2007). When PLIN-1 is absent, F22F7.1, the homolog of the mammalian lipid droplet protein CGI-49, increases in abundance on the surface of lipid droplets (Na et al. 2015). PLIN-1 is especially abundant in embryos, and embryos produced by plin-1 RNAi contain lipid droplets that clump together, indicating the importance of PLIN-1 in stabilizing the structure of lipid droplets in embryos (Na et al. 2015; Vrablik et al. 2015). Other C. elegans lipid droplet-associated proteins include the ACS-22(FAT1P) and DGAT-2 complex, which is required for the generation of lipid droplets (Mak 2012; Xu et al. 2012), DHS-3 (Zhang et al. 2012; Na et al. 2015), ACS-4 (Vrablik et al. 2015), ATGL-1 (Zhang et al. 2010a), and LID-1 (Lee et al. 2014).

Several activities are important for the formation of lipid droplets and for the regulation of lipid droplet size. The FITM-2 protein, a member of the FIT family of conserved ER proteins required in yeast and mammals for the generation of lipid droplets at the ER, is also required for lipid droplet formation in C. elegans (Choudhary et al. 2015). The fitm-2 mutants are lethal, implying that lipid droplet formation is essential. Lipid biosynthesis genes, including FAT-6 and FAT-7 are required for lipid droplet expansion, even in genetic backgrounds that normally promote increased fat stores (Shi et al. 2013). The atlastin GTPase is required for proper ER structure and lipid droplet size expansion (Klemm et al. 2013). The peroxisomal β-oxidation proteins MAOC-1, DHS-28, and DAF-22 are also required for the regulation of lipid droplet size (Zhang et al. 2010a). Mutants in these genes all accumulate large lipid droplets, indicating a requirement for the peroxisome in the utilization of fat from lipid droplets. Several recent genetic screens have identified a range of lipid droplet size regulators in embryos (Schmokel et al. 2016) and in adults, including several new proteins involved in lipid droplet fusion (Li et al. 2016).

Neutral lipid catabolism

To utilize fatty acids for energy, they must first be liberated from TAG molecules, and then broken down by β-oxidation, the sequential cleaving of two-carbon units to produce acetyl-CoA, which then enters the citric acid cycle to produce CO₂, NADH, and ATP. A reverse genetic screen of lipases identified ATGL-1, the homolog of mammalian adipocyte triglyceride lipase as the major lipase involved in liberating fatty acids from TAGs from lipid droplets during fasting (Lee et al. 2014). ATGL-1 is responsible for the rapid burning of fat in dauer larvae carrying mutations in AMP kinase, and thus participates in the rapid fat burning that contribute to the demise of AMP kinase-deficient dauer larvae (Narbonne and Roy 2009). The phosphorylation of ATGL-1 by AMP kinase generates 14-3-3 binding sites on ATGL-1, which leads to the dissociation of ATGL-1 from lipid droplets and eventual protesosomal degradation of ATGL-1 (Xie and Roy 2015a). A C. elegans homolog of the mammalian lipid droplet-associated protein CGI-58, called ABHD-5.2 in C. elegans, was shown to coactivate ATGL-1 by tethering it to lipid droplets (Xie and Roy 2015b). In addition, CGI-58/ABHD-5.2 acts independently of ATGL-1 to prevent fusion of lipid droplets, which might stem from changes in PUFA levels of the phospholipids surrounding lipid droplets (Xie and Roy 2015b).

Autophagy is the process of recycling of cellular material, including damaged proteins, mitochondria, peroxisomes, and lipid droplets to the lysosome for degradation. During fasting, the lysosome plays an important role in the early catabolic steps of lipid degradation (Settembre and Ballabio 2014). C. elegans contains at least three lysosomal lipases, LIPL-1, LIPL-3, and LIPL-4 (Wang et al. 2008; Lapierre et al. 2011; O’Rourke and Ruvkun 2013; Folick et al. 2015). LIPL-4 has been studied the most because of the link between autophagy, lysosomal lipase activity, and the extension of lifespan in germline-less nematodes (Wang et al. 2008; Lapierre et al. 2011). As described in Dietary and endogenous synthesized monounsaturated fatty acids regulate fat accumulation and longevity, ∆9 desaturases and their regulators NHR-80 and NHR-49 are required for the generation of lipid signals that promote longevity (Goudeau et al. 2011; Ratnappan et al. 2014). Thus, lipid synthesis and lipid catabolism are both required for increased longevity in animals lacking a germline. This may be explained by the recent findings that increased yolk lipoproteins are detrimental to longevity (Seah et al. 2016), and that, in constantly feeding worms, the lack of germline leads to energy imbalance in which greatly increased yolk accumulation occurs in intestinal cells and in pseudocoelomic spaces (Steinbaugh et al. 2015). The excess yolk has been proposed to trigger autophagy pathways regulated by HLH-30 to induce lipases required for the breakdown of excess lipids, while at the same time contributing to the production of lipid signals, such as oleyl ethanolamide (OEA), which is transported to the nucleus by way of the lysosomal lipid chaperone LBP-8, and serves as a ligand to activate NHR-80 and NHR-49 (Folick et al. 2015). Similar lipid signals derived from oleic acid could activate SKN-1, which, together with NHR-49, activate lipid catabolic pathways and other protective pathways leading to increased longevity (Ratnappan et al. 2014; Steinbaugh et al. 2015). In addition, the transcription factors DAF-16/FOXO and TCER-1 regulate lipid metabolism pathways in animals lacking a germline (Amrit et al. 2016). Interestingly, even though autophagy and lysosomal lipases are important components of lipid catabolism, well fed, fertile C. elegans mutants defective in autophagy processes actually have reduced, not increased, lipid accumulation compared to wild type (Lapierre et al. 2012). This indicates that autophagy is required for overall metabolic homeostasis in C. elegans.

β-Oxidation of fatty acids

Homologs of mitochondrial and peroxisomal genes encoding proteins involved in β-oxidation of fatty acids are present in C. elegans (Van Gilst et al. 2005b). The mechanism of the core β-oxidation cycle involves a dehydrogenation, hydration, dehydrogenation again, and a thiolytic cleavage, in which each cycle produces acetyl-CoA, NADH, and FADH₂, and an acyl group that is two carbons shorter than before. The four enzymes required include acyl-CoA dehydrogenase in the mitochondria or acyl-CoA oxidase in peroxisomes, enoyl-CoA hydratase, 3-hydroxylacyl-CoA dehydrogenase, and 3-ketoacyl-CoA thiolase (Figure 4). Peroxisomal β-oxidation has been studied more in C. elegans than mitochondrial beta oxidation, because the peroxisomal enzymes are required for synthesis of ascarosides, signaling glycolipids based on the sugar ascarylose linked to short chain fatty acids [Dideoxysugars (ascarosides); Ludewig and Schroeder 2013]. For peroxisomal oxidation and ascaroside formation, the genes encoding the major activities of the pathway include various acox genes, maoc-1, dhs-28, and daf-22 (Figure 4) (Jorgensen and Mango 2002; Butcher et al. 2009; von Reuss et al. 2012).

Figure 4.

Lipid catabolism. (A) Location of phospholipase activity on a hypothetical phospholipid. (B) Location of TAG lipase cleavage on triacylglycerol. (C) Pathway of β-oxidation in peroxisomes and mitochondria. FA-coA, fatty acyl coenzymeA; CPT, carnitine palmitoyl transferase; ACOX, acyl coenzyme A oxidase; ECH, enoyl CoA hydratase; 3-HACD, 3-hydroxyacyl CoA dehydrogenase; FA-2C, a fatty acyl group two carbons shorter than when it started the cycle.

A β-oxidation-like pathway is used in C. elegans for the breakdown of propionate under conditions of limiting vitamin B12 (Watson et al. 2016). Propionate is a three-carbon short-chain fatty acid breakdown product of odd-chain fatty acids and branched chain amino acids that is normally catabolized by a carboxylation pathway that depends on vitamin B12. In the alternative pathway, the short-chain acyl-CoA dehydrogenase ACDH-1 converts propionyl-CoA to acrylyl-CoA, which is oxidized by the β-oxidation enzyme ECH-6, and further broken down into acetyl-CoA by the activities of HACH-1, HPHD-1, and ALH-8 (Watson et al. 2016).

In mammals, the peroxisome is involved in breaking down very long chain fatty acids, while mitochondrial oxidation breaks down long, medium, and short chain fatty acids. However, in plants, and in most fungi, fatty acid oxidation takes place solely in peroxisomes (Wanders and Waterham 2006). While the relative proportions of fat oxidation occurring in mitochondria or peroxisomes are not known in C. elegans, genetic screens point to the importance of peroxisomal oxidation for the breakdown of fats in large lipid droplets (Zhang et al. 2010a; Li et al. 2016). A biochemical assay for C. elegans β-oxidation has been developed (Elle et al. 2012). In addition to fat oxidation defects in AMP kinase mutants and atgl-1 mutants, this assay has been used to demonstrate altered β-oxidation in fat-6;fat-7 double mutants (Shi et al. 2013), acs-3 mutants (Mullaney et al. 2010) and daf-2 mutants (Elle et al. 2012). A functional role for β-oxidation has been shown using RNAi knockdown of β-oxidation genes to implicate increased fat oxidation as the mechanism of altered fat accumulation in various studies, including that of cebp-2 mutants (Xu et al. 2015), klf-3 mutants (J. Zhang et al. 2011), and in genetic backgrounds in which serotonin signaling is increased (Srinivasan et al. 2008).

Visualization of neutral lipids

Due to the human obesity epidemic and the serious diseases associated with obesity, there has been great interest in using C. elegans genetics to identify novel fat regulatory genes. Initial genetic screens used the lipophilic dye Nile Red in nematodes to visualize fat accumulation in live animals (Ashrafi et al. 2003). Feeding of Nile Red was later determined to be problematic because Nile Red dye accumulates in lysosomal related organelles (LROs), also called gut granules, and this accumulation does not necessarily correspond to TAG levels in the worms (Schroeder et al. 2007; O’Rourke et al. 2009). Fixation of nematodes with isopropanol or paraformaldehyde and staining with Sudan Black, Nile Red, or Oil Red O produces visual patterns that are representative of biochemical determinations of fat levels (Brooks et al. 2009; O’Rourke et al. 2009; Pino et al. 2013; Wahlby et al. 2014). Lipid droplets in the intestine, germ line, and in epidermal tissues can be visualized with this technique. Another family of vital stains, BODIPY-labeled fatty acids, stains both lipid droplets and LROs in live nematodes, and has been successfully used to screen for mutants with supersized lipid droplets (Zhang et al. 2010a,b; Li et al. 2016). Lipids and fat depots have been visualized in label-free live animals using Coherent Anti-Raman Stokes (CARS) microscopy (Hellerer et al. 2007; Yen et al. 2010; Folick et al. 2011), whole-body imaging mass spectrometry (Hameed et al. 2015), and nonlinear microscopy imaging including second and third harmonic generation phenomena, which can detect ectopic fat deposits in muscle and neuronal tissues in aging worms (Mari et al. 2015; Palikaras et al. 2016).

Several strains expressing lipid droplet markers fused to GFP have been used to characterize lipid droplet size and abundance, although not every lipid droplet expresses the same lipid droplet proteins. Lipid droplets expressing DGAT-2 can be visualized in the intestine with the DGAT-2::GFP strain (Mak 2013), a subset of lipid droplets in the germline can be visualized with a ATGL-1::GFP and Drosophila PLIN-1::GFP, which localizes to C. elegans lipid droplets (Z. Liu et al. 2014).

Regulation of neutral lipid synthesis and breakdown

Lipid metabolism in C. elegans is regulated in response to environmental inputs, such as temperature and food availability, as well as by physiological state, including development, reproduction, and aging. Fat accumulation is altered depending on the bacterial strain used in culture (Brooks et al. 2009), food availability (Van Gilst et al. 2005b), temperature (Murray et al. 2007; Ma et al. 2015), iron (Wang et al. 2016), and various dietary treatments, such as supplementation of plant-derived extracts (Bhattacharya et al. 2013; El-Houri et al. 2014; Zheng et al. 2014; Ding et al. 2015; Gao et al. 2015; Peng et al. 2016; Sun et al. 2016).

The regulation of fat synthesis, desaturation, and lipid turnover is likely mediated at many levels, including transcription, translation, and protein stability. Of these processes, transcriptional regulation has been studied the most in C. elegans. The homolog of mammalian SREBP-1, SBP-1, regulates transcription of genes encoding proteins for fatty acid biosynthesis, especially fat-5, fat-6, fat-7, elo-5, elo-6, and genes encoding proteins required for phospholipid synthesis (see Biosynthesis of phospholipids below) (Kniazeva et al. 2004; Yang et al. 2006; Walker et al. 2011). SBP-1 null mutants arrest during early larval stages (McKay et al. 2003), while a hypomorphic allele, sbp-1(ep79), develops to adulthood, but displays slow growth, reduced reproduction, and low fat accumulation (Liang et al. 2010). Mutants defective in fat synthesis, including sbp-1, fat-6;fat-7, and pod-2 fail to display normal satiety quiescence, in which worms stop feeding and reduce their activity—an activity analogous to sleep in mammals (You et al. 2008; Hyun et al. 2016). Thus, altered fat synthesis is detrimental to many aspects of C. elegans physiology.

Another major transcription factor regulating lipid metabolism in C. elegans is NHR-49 (Van Gilst et al. 2005a). This nuclear receptor is one of the large family of C. elegans HNF4-type NHRs, and it shows functional homology to mammalian PPARα—a mammalian regulator of lipid metabolism (Antebi 2015). NHR-49 regulates ∆9 desaturases, fat oxidation genes, sphingolipid synthesis genes, as well as nonlipid metabolism genes during normal development and also in response to fasting (Van Gilst et al. 2005b; Pathare et al. 2012). Both SBP-1 and NHR-49 associate with the mediator subunit MDT-15 to regulate transcription of lipid metabolism genes (Taubert et al. 2006; Yang et al. 2006). Gain-of-function alleles of nhr-49 and mdt-15, together with genes required for PC synthesis, were isolated in a screen for suppressors of the cold-sensitive paqr-2 strain, suggesting that the adiponectin homolog PAQR-2 regulates fatty acid desaturation during cold adaptation, in part by modulating PC abundance and SBP-1 activation (Svensk et al. 2013). In mutant strains carrying gain of function nhr-49 alleles, many NHR-49 targets are upregulated, even those not involved in lipid metabolism (Lee et al. 2016). Another HNF4-type NHR, NHR-64, also regulates lipid metabolism, and RNAi of nhr-64 partially suppresses the slow growth and low fat phenotypes of sbp-1 and fat-6;fat-7 mutants (Liang et al. 2010). In forward and reverse genetic screens, additional NHRs were identified as playing roles in regulating metabolic responses to various dietary inputs, as well as in regulating feeding behavior (Watson et al. 2013; Hyun et al. 2016). Hence, various nuclear receptors mediate a complex gene regulatory network regulating metabolic responses to the environment.

Fasting induces transcription of lipases, including fasting-induced lipases FIL-1 and FIL-2 (Jo et al. 2009), and the lysosomal lipase LIPL-1(O’Rourke and Ruvkun 2013). LIPL-1 is regulated by the bHLH transcription factor MLX-3, which represses lipl-1 expression in the fed state, while, after food removal, another bHLH transcription factor HLH-30 activates expression of lipl-1 and other lipases (O’Rourke and Ruvkun 2013). HLH-30 is also required in well-fed worms to limit ectopic lipid deposition in neuronal and muscle tissue in aging worms (Palikaras et al. 2016). Transcription of the adipocyte triglyceride lipase homolog atgl-1 is repressed by the bHLH homolog HLH-11 in the presence of food, and then activated by NHR-76 in response to serotonin signaling (Noble et al. 2013; Srinivasan 2015).

Serotonin, also known as 5-HT, controls many food-related behaviors, including food intake, energy expenditure, locomotion, and reproduction (Horvitz et al. 1982; Loer and Kenyon 1993; Waggoner et al. 1998; Sawin et al. 2000; Sze et al. 2000). Increasing serotonin signaling leads to rapid decrease of body fat as well as increased locomotion and feeding. However, increased feeding behavior and increased fat expenditure are regulated by independent neuronal circuits, in which the serotonin-gated chloride channel MOD-1 is involved in as the receptor required for loss of body fat, yet has no role in the behavioral responses to increased serotonin (Srinivasan et al. 2008). Three G-protein coupled serotonin receptors, SER-1, SER-5, and SER-7 are required for changes in feeding rates, but do not affect body fat composition (Srinivasan et al. 2008; Cunningham et al. 2012; Song and Avery 2012). For the regulation of body fat levels, serotonin and octopamine act together in a neural circuit to sustain body fat expenditure (Noble et al. 2013). A more detailed review of the neuronal control of fat accumulation is found in several recent publications (Lemieux and Ashrafi 2015; Srinivasan 2015).

A post-transcriptional regulator of fat accumulation was recently identified in a screen for genes required for survival during cold adaptation. The RNAse REGE-1 promotes the accumulation of fat by degrading ETS-4, a transcription factor that promotes the expression of fat catabolism genes (Habacher et al. 2016). Interestingly, ETS-4 also promotes expression of REGE-1, establishing an auto-regulatory loop for fine control of fat breakdown.

Two well-studied endocrine pathways required to promote reproductive growth under favorable nutrient conditions are the TGFβ/DAF-7 pathway and the insulin-like growth factor pathway, including the insulin-like growth factor receptor DAF-2. These developmental fate regulators are important in adult stage nematodes for body fat homeostasis, as reduction of function of daf-2 or daf-7 during adulthood leads to greatly increased fat accumulation (Kimura et al. 1997; Greer et al. 2008). DAF-7 is produced by a pair of ASI neurons and acts as a food sensor as well as a cell-nonautonomous regulator of fat synthesis and utilization (Greer et al. 2008). The DAF-2 receptor is expressed widely in the nervous system, and regulates fat via DAF-16/FOXO transcription factor activity in the intestine. The daf-2 mutants accumulate large lipid droplets (Shi et al. 2013), and exhibit a higher rate of de novo fat synthesis than wild type (Perez and Van Gilst 2008).

Mechanistic target of rapamycin (mTOR) is a conserved serine/threonine kinase that acts as a nutrient sensor and promoter of growth. The mTOR kinase acts in two distinct complexes, TORC1, which consists of DAF-15/Raptor and TORC2, which consists of RICT-1/Rictor. Null mutants of let-363/mTOR and daf-15/Raptor arrest as L3 stage larvae (Long et al. 2002; Jia et al. 2004), while mutants in rict-1/Rictor display delayed growth rates, increased fat stores, and long lifespan (Jones et al. 2009; Soukas et al. 2009). The effects of TORC2 are mediated by SGK-1, a homolog of serum- and glucocorticoid-induced kinases. Recently it was reported that the lin-4 and let-7 microRNA program, which is best known for acting in the epidermis to control developmental timing, also signals to the intestine through SGK-1 and TORC2 to coordinate yolk protein synthesis upon the initiation of reproduction (Dowen et al. 2016).

The TORC1 complex is regulated by nutrient intake, when nutrients are scarce, AAK-1/AMP kinase acts on the TSC1 complex to inhibit the conversion of GTPase activating protein RHEB to RHEB-GTP, leading to repression of TORC1 complex activity. While the components of this signaling pathway are not well defined in C. elegans, a recent study revealed that ATX-2, a homolog of human Ataxin-2, acts as a TORC1 repressor, and atx-2 mutants exhibit increased body size, cell size, and fat accumulation (Bar et al. 2016). A downstream target of the TORC1 complex is the S6 kinase, called RSKS-1 in C. elegans. The S6 kinase regulates translation efficiency by phosphorylating ribosomal protein S6 and other proteins. C. elegans rsks-1 mutants grow slowly, have a reduced brood size and a smaller pool of germline progenitor cells (Korta et al. 2012). The rsks-1 mutant also have increased fat accumulation compared to wild type (Shi et al. 2013). This reduced progenitor pool is also observed in nutrient sensing-defective, high fat accumulating daf-2 and daf-7 mutants (Michaelson et al. 2010; Dalfo et al. 2012). Thus, in these three strains, as well as others, such as glp-1 mutants, there is often a correlation between increased fat accumulation and reduced reproductive output, and often these strains exhibit lifespan extension as well (Lemieux and Ashrafi 2016).

Membrane Lipids

In C. elegans, membranes are composed of glycerophospholipids, sphingolipids, and very small amounts of cholesterol. Glycerophospholipids, hereafter called phospholipids, make up the bulk of membrane lipids, with PC and phosphatidylethanolamine (PE) being the most abundant (Satouchi et al. 1993) (Figure 1C).

Biosynthesis of phospholipids

Phospholipids are synthesized from several pathways including the CDP-DAG pathway and the Kennedy pathway (Figure 3). Both pathways begin with the addition of two acyl groups by acyl-transferase enzymes (GPAT and LPAAT homologs) to glycerol-3 phosphate (G3P), generating PA—a central metabolite in both neutral and phospholipid synthesis. Before transfer, the acyl groups must first be activated to acyl-CoAs, a reaction that is catalyzed by ACS enzymes. C. elegans has 23 ACS homologs, which potentially activate fatty acids for use in a variety of processes, including lipid synthesis, β-oxidation, protein acylation, and fatty acyl transport across membranes (Watkins et al. 2007). Several C. elegans ACS proteins that have been biochemically assayed, including ACS-1, which is required for mmBCFA synthesis (Kniazeva et al. 2008), and ACS-20 and ACS-22, which are FAT1p homologs required for the incorporation of C26:0 in sphingomyelin (Kage-Nakadai et al. 2010). Recently, ACS-4 was shown to have protein myristolation activity, and to regulate fertility by translating fatty acid levels into regulatory cues affecting the germline sex-determination pathway (Tang and Han 2017). Other enzymes involved in phospholipid synthesis include the two acyl-transferase activities that contribute to PA synthesis from G3P, which are likely encoded by one or more of the 14 acyl-CoA ligases (ACL) enzymes that are homologs of GPAT and LPAAT. While the GPAT and LPAAT activities involved in PA biosynthesis have not been characterized in C. elegans, four ACL enzymes required for other activities have been characterized. These include ACL-7, the mammalian GNPAT homolog that acts in the peroxisome for the first steps of the synthesis of ether-linked lipids (see Synthesis of ether-linked PE species) (Shi et al. 2016) and ACL-8, ACL-9, and ACL-10/BUS-18, which, together with the phospholipase ipla-1, are responsible for the remodeling of PI such that stearic acid is transferred to the sn1 position (Imae et al. 2010).

In the CDP-DAG pathway, PA and CTP are converted to CDP-DAG by CDP-DAG synthase, encoded by the cdgs-1 gene. The CDP-DAG pathway permits synthesis of phosphatidyl inositol (PI) and phosphatidyl glycerol (PG) by the addition of the inositol (pisy-1) and glycerol-3 phosphate (pgs-1 to CDP-DAG (Figure 3). The mitochondrial lipid cardiolipin (CL) is synthesized from two PG molecules by cardiolipin synthase, CRLS-1 (Sakamoto et al. 2012).

The Kennedy pathway also operates in C. elegans, in which choline and ethanolamine are phosphorylated by the activities of choline- and ethanolamine-kinases (encoded by cka-1, cka-2, ckb-1, ckb-2, ckb-3, ckb-4), then activated by CTP-phosphocholine cytidylyl transferase (pcyt-1) (Walker et al. 2011) and CTP-phosophoethanolamine cytidylyl transferase. The activated CDP-choline and CDP-ethanolamine then react with DAG to form PE and PC, with the enzyme CEPT-1. In many organisms, PC can by synthesized from PE by three sequential methylation reactions, using s-adenysyl methionine (SAM) as a substrate and methyl transferase enzymes (Brendza et al. 2007; Walker et al. 2011). Interestingly, while most animals can convert PC from PE using PE methyl transferases, C. elegans perform this methylation in a manner similar to plants, using phosphoethanolamine as a substrate and the phosphoethanolamine methyl transferase enzymes PMT-1 and PMT-2 (Brendza et al. 2007) (Figure 3).

Phosphatidylserine (PS) is formed from the exchange of serine with choline or ethanolamine moieties of PC or PE by the PS synthase enzymes 1 and 2 (PSSY). In mammals, PTDSS1 (homolog of PSSY-1) can use either PE or PC as a substrate, while PTDSS2 (homolog of PSSY-2) only uses PE. Substrate specificities of PSSY-1 and PSSY-2 have not been determined in C. elegans. PS can be converted back to PE with the activity of PS decarboxylase (PSD-1) (Figure 3).

The importance of the phosphocholine methylation pathway for PC synthesis is apparent when observing sams-1 mutants. These mutants exhibit low levels of PC, which can be rescued by dietary choline (Walker et al. 2011). In the absence of dietary choline, the sams-1 worms exhibit large lipid droplets, and high PE and TAG levels, which may be due to lipid homeostasis disruption that occurs when DAG precursors are not used for PC synthesis, and instead are diverted into the neutral lipid synthesis pathway (Li et al. 2011; Walker et al. 2011; Ehmke et al. 2014). During a moderate, post embryonic dietary restriction regime, lipid stores increase, and reproduction slows, even though the restricted worms produce normal numbers of progeny over their lifetime (Palgunow et al. 2012; Miersch and Doring 2013). Interestingly, dietary choline abrogates this increased lipid droplet size and TAG levels, indicating that methyl donors and one carbon metabolites are key factors for normal lipid homeostasis (Ehmke et al. 2014).

Membrane asymmetry and remodeling

In animals, membrane lipid composition varies depending on cell and organelle type, but, in C. elegans, phospholipid analysis has only been performed using whole worms because of the difficulty in isolating enough pure tissues for lipid analysis. It is likely that the distribution of various phospholipids is not uniform throughout the membranes of intercellular organelles or among distinct tissues. For example, based on lipid analysis of other animals, cardiolipin is mainly a component of mitochondrial membranes, while sphingomyelin is mainly localized to the outer leaflet of plasma membranes.

Class IV P-type ATPases translocate PS or PE between two leaflets of membranes, maintaining a higher concentration of PE and PS on the inner leaflet of the plasma membrane (Folmer et al. 2009). C. elegans encodes six genes predicted to encode class IV P-type ATPases: tat-1–tat-6. Mutants lacking tat-1 show increased PS staining on the outer surface of germ cells, indicating that TAT-1 functions to maintain PS in the inner membrane leaflet (Darland-Ransom et al. 2008). The externalization of PS serves as an engulfment signal during the conserved process of apoptosis, and tat-1 mutants were found to exhibit a random loss of neurons and muscle cells that was dependent on the PS-dependent phagocyte recognizing receptor PSR-1 (Darland-Ransom et al. 2008). Furthermore, TAT-1 and its chaperone CHAT-1 are required for proper membrane tubule formation, and, in their absence, endocytic sorting and recycling are defective (Chen et al. 2010). In the embryo, tat-5 mutants release excessive extracellular vesicles, leading to disruptions in cell adhesion and morphogenesis during embryogenesis (Wehman et al. 2011). Staining embryos with duramycin, which binds to PE, revealed excessive PE on the outer membrane surface of tat-5 mutant embryos, demonstrating a link between membrane lipid asymmetry and extracellular vesicle formation (Wehman et al. 2011).

The requirement for regulated phospholipid composition within the nuclear envelope was demonstrated in studies of C. elegans mutants defective in the PA phosphatase LIPIN (lpin-1 mutants) as well as mutant strains of the nuclear envelope-localized LIPIN regulator CTDNEP1 (cnep-1 mutants). In both mutant strains, the nuclear envelope fails to disassemble during mitosis (Golden et al. 2009; Gorjanacz and Mattaj 2009; Bahmanyar et al. 2014). Studies of the cnep-1 mutants showed that ectopic ER sheets formed around the nuclear envelope, preventing breakdown of the nuclear membrane during mitosis (Bahmanyar et al. 2014). The formation of the ectopic ER sheets correlated with higher levels of PI in the cnep-1 mutants, and both PI levels and the ectopic sheet formation were rescued by RNAi inhibition of pisy-1 or cdgs-1, which reduced PI synthesis (Bahmanyar et al. 2014). These studies demonstrate that spatial regulation of membrane phospholipid composition is critical for key cellular and developmental events.

The various phospholipids have distinct fatty acyl compositions (Table 1). It is likely significant fatty acyl remodeling is occurring to maintain the specific composition. For example, cardiolipin PUFA composition is unique, and, unlike other phospholipids, contains almost no ∆5 desaturated fatty acids (Hou et al. 2014) (Table 1). Adding fatty acyl groups to lysophospholipids—an activity important for membrane remodeling—uses membrane bound o-acyltransferases, abbreviated MBOA. C. elegans encodes seven mboa genes, and the biochemical activities of two MBOAs have been characterized. MBOA-6 has been shown to incorporate PUFAs into PC, PS, and PE (Matsuda et al. 2008). MBOA-7 incorporates long chain PUFA specifically into PI (Lee et al. 2008), and, as such, plays an important role in phosphoinositide signaling (Lee et al. 2012).

Synthesis of ether-linked PE species

Ether lipids are glycerolipids in which one of the fatty acyl groups, typically a saturated fatty acid, is linked to the sn1 position of glycerol backbone by an ether bond, rather than by the usual ester bond. Ether lipids can be in the alkylacyl form, or the alkenylacyl, which contain a double bond adjacent to the oxygen, and are also known as plasmalogens (Figure 1D). In C. elegans, ether-linked lipids are found mainly in PE, with very small amounts in PC (Satouchi et al. 1993; Drechsler et al. 2016; Shi et al. 2016); ∼20% of PE species contain an alkylacyl bond, and 14% contain an alkenyl bond (Satouchi et al. 1993).

Ether-linked lipids are unusual in that their synthesis begins in the peroxisome (Brites et al. 2004). In the peroxisome, the acyl transferase ACL-7 transfers an acyl group to dihydroxyacetone phosphate, creating 1-acyl-DHAP. The ether-linkage is formed by the ADS-1 enzyme, which catalyzes the synthesis of 1-alkylacyl DHAP by using a reduced fatty alcohol, rather than a fatty acyl, as a substrate. The reduced fatty alcohol is produced by the fatty alcohol reductase FARD-1. Mutants in acl-7, ads-1, and fard-1 lack the ability to synthesize ether-linked lipids (Shi et al. 2016). Analysis of the ether-lipid deficient mutants demonstrate that ether lipids are required for optimal reproduction, lifespan, survival at cold temperatures, and resistance to oxidative stress (Drechsler et al. 2016; Shi et al. 2016). All of the C. elegans ether-lipid deficient strains accumulate high levels of 18:0 in all lipids, presumably to compensate for the structural properties of ether-linked lipids, which are predicted to contribute to membrane rigidity.

The activity of AGMO-1, alkyglycerol monooxygenase, is required to break down ether-linked lipids (Watschinger et al. 2010). The C. elegans agmo-1 gene is expressed in the epidermis, and agmo-1 mutants, along with mutant strains lacking the ability to synthesize tetrahydrobiopterin, an AGMO-1 cofactor, have a fragile cuticle and are sensitive to exogenous chemicals (Loer et al. 2015). This indicates that oxidation of ether lipids is an important process involved in the formation of the nematode cuticle.

Phospholipid catabolism and signaling

Both synthesis and degradation of phospholipids occurs constantly, as membrane are consumed for cellular processes such as intracellular trafficking, organelle biogenesis, β-oxidation, and cellular repair. Phospholipid turnover has been measured in C. elegans using a ¹⁵N incorporation assay analyzed by HPLC-MS/MS (Dancy et al. 2015). For this study, 210 distinct PC and PE species were followed, and the authors found that ∼2% of phospholipid species are turned over every hour, with PE species turning over at rates higher than PC species (Dancy et al. 2015). The rates of fatty acyl alterations in phospholipids are even higher; thus, the majority of C. elegans lipids turnover daily (Dancy et al. 2015).

Fatty acyl groups are liberated from phospholipids by the activities of phospholipase enzymes (Figure 4A). There are five types of phospholipases that are defined by the position in a phospholipid in which they cleave. Phospholipases A₁ and A₂ cleave off a fatty acyl group at the sn-1 and sn-2 positions of phospholipids, respectively. Phospholipiase B enzymes possess both A₁ and A₂ activity. Phospholipase C enzymes cleave at the at the phosphate group, creating DAG and a phosphorylated headgroup, while Phospholipase D enzymes cleave on the other side of the phosphate group, creating PA and an alcohol. Phospholipases that are secreted into the lumen of the intestine aid in digestion of bacterial membranes, allowing for the uptake of bacterial fatty acids and lysophospholipids. Intracellular phospholipases play important roles in signaling, creating powerful short-range signaling molecules such as DAG, phosphoinositides, fatty acyl amides, and eicosanoids. Each family of phospholipases is represented by multiple gene products in C. elegans, and most phospholipase activities and substrate specificities have not been experimentally determined. In addition to their role in signaling, phospholipases also contribute to membrane maintenance by removing fatty acyl groups to allow acyl transferase enzymes to insert specific fatty acyl species for membrane remodeling.

C. elegans contains multiple gene encoding phospholipase A₂ homologs, and the group VIA, calcium-independent phospholipase A₂ homologs have been identified by sequence similarity to the human gene (Morrison et al. 2012). The C. elegans IPLA-1 was expressed and purified for biochemical studies examining its conformation, complex formation, and substrate specificity, and it was found to bind to many species of acidic phospholipids, including PA, PG, PS, CL, and phosphorylated derivatives of PI (Morrison et al. 2012). Knockdown of the six homologs showed that IPLA-2 is necessary for optimal progeny production (Morrison et al. 2012). A study of the roles of lysosomes in stress responses, including short-term fasting, identified IPLA-2 as a target of NHR-49 in attenuating the overaccumulation of lysosomal lipids during fasting (Huang et al. 2014).

Among the best studied phospholipases are the phospholipase C family, which produces three distinct signaling molecules from PI: DAG, inositol 1,4,5-trisphosphate (IP₃), and phosphatidylinositol 4,5 bisphosphate (PIP₂) (Putney and Tomita 2012). Phospholipase C activity is regulated by calcium, and all three of the lipid signaling molecules regulate ion channels. C. elegans encodes at least five active PLC isoforms belonging to four PLC families: plc-2 and egl-8 (PLC-β), plc-3 (PLC-γ), plc-4 (PLC-δ), and plc-1 (PLC-ε) (Gower et al. 2005). These PLC activities are important for a range of reproductive, neurological, and host defense processes (Gower et al. 2005; Vazquez-Manrique et al. 2008; Ziegler et al. 2009; Iwasa et al. 2010; Kawli et al. 2010; Xing and Strange 2010; Xu and Chisholm 2011; Kovacevic et al. 2013; Yu et al. 2013; Chun et al. 2015; Najibi et al. 2016; C. Zhang et al. 2016).

Phospholipase D activity produces N-acyl amides, lipid signaling molecules that include endogenous ligands of mammalian cannabinoid receptors (De Petrocellis et al. 2000). In C. elegans, at least six N-acetylethanolamines (NEAs) are generated from PE molecules by the phospholipase D isoforms NAPE-1 and NAPE-2 (Lucanic et al. 2011; Harrison et al. 2014). NEAs promote larval development, and are present at very low levels in starved L1 or dauer larvae. Dietary EPEA, derived from PE containing C20:5, rescues constitutive dauer formation in chemosensory mutants, TGFβ mutants and insulin signaling mutants required for food sensation and promotion of reproductive growth (Lucanic et al. 2011). Low levels of NEAs present in dietary restricted nematodes promote long lifespan, and dietary NEAs obliterate the lifespan extension (Lucanic et al. 2011). NAEs are degraded by fatty acid amide hydrolase FAAH-1, and overexpression of faah-1 causes a developmental delay (Lucanic et al. 2011). OEA levels are affected by the lysosomal lipase LIPL-4, a lipase linked to longevity of germline deficient nematodes (Wang et al. 2008; Lapierre et al. 2012; Folick et al. 2015). OEA binds to LBP-8, a lipid chaperone required for lysosomal signaling to the nucleus (Folick et al. 2015). The nuclear OEA then binds to NHR-80, which acts with NHR-49 to regulate fatty acid desaturation and promote longevity (Folick et al. 2015).

Sphingolipid biosynthesis and function

Sphingolipids differ from glycerophospholipids in that they contain a sphingoid base, which is a long-chain aliphatic amine (Figure 1F). In most animals, the sphingoid base is typically the saturated straight chain of 18 carbons containing a double bond at C4, hydroxyl groups at C1 and C3, and an amino group at C2. In C. elegans, the long-chain sphingoid base is derived from the branched chain fatty acid C15:iso, which condenses with serine to form a d17:iso-sphinganine (Chitwood et al. 1995; H. Zhang et al. 2011). The condensation reaction uses the enzyme serine palmitoyl transferase, forming 3-keto sphinganine, which is reduced by the sphinganine reducatase and NADPH to form sphinganine (Figure 5). The serine palmitoyl transferase homologs are encoded by three genes in C. elegans, sptl-1, sptl-2, and sptl-3. RNAi of sptl-1 results in L1 arrest with intestinal lumen polarity defects that resulted in L1 lethality (H. Zhang et al. 2011). This L1 lethality can be rescued by dietary supplementation of d17iso-sphinganine (Zhu et al. 2013).

Figure 5.

Sphingolipid synthesis and breakdown. Yellow ovals depict substrates and breakdown metabolites, blue ovals depict minor lipids and intermediates, green ovals depict signaling lipids, red ovals depict major lipid products, and purple ovals depict enzymes. C15iso, 13-methyltetradecanoic acid; FA-coA, fatty acyl coenzymeA; Glu, glucose; Mann, mannose; FFA, free fatty acid; PC, phosphatidylcholine; DAG, diacylglycerol.

The simplest sphingolipid, ceramide, consists of a fatty acid linked to the amine group of the sphinganine base (Figure 1F). In C. elegans, the amino linked fatty acids are primarily straight-chained saturated or 2-hydroxylated fatty acids of length C18–C26 (Chitwood et al. 1995; H. Zhang et al. 2011; Cutler et al. 2014). The long chain saturated or hydroxylated fatty acyl-CoA is added to the 17iso-sphinganine by ceramide synthase enzymes encoded by the hyl-1, hyl-2, and lagr-1 genes. This forms d17iso-dihydroceramide, which is then desaturated by dihydroceramide desaturases, encoded by ttm-5 and F33D4.4, to form ceramide. In C. elegans, in addition to its role as a structural membrane lipid, de novo ceramide synthesis is required to regulate apoptosis of germ cells after radiation damage (Deng et al. 2008), as well as for surveillance of mitochondrial function after genetic or drug-induced treatments that disable mitochondria (Y. Liu et al. 2014).

The ceramide synthases HYL-1 and HYL-2 transfer distinct acyl chains to the d17iso sphinganine. HYL-1 is specific for C24–C26 acyl chains, while HYL-2 preferentially catalyzes the reaction with C20–C22 acyl chains (Menuz et al. 2009). Mutants in hyl-1 and hyl-2 have opposing phenotypes, hyl-1 mutants are resistant to anoxia, while hyl-2 mutants are more sensitive than wild type, indicating that specific ceramides produced by HYL-2 confer resistance to anoxia (Menuz et al. 2009). The hypoxia sensitivity of hyl-2 mutants can be suppressed by reduction of CYP-25A1 or CYP-33C8 (Ladage et al. 2016). Similarly, hyl-2 mutants have short lifespans, while a double mutant of hyl-1;lagr-1 produced worms with extended lifespan (Mosbech et al. 2013). Interestingly, the long lifespan of hyl-1;lagr-1 double mutants can be suppressed by sphk-1 RNAi, which reduces the activity of sphingosine kinase, which acts in the sphingolipid degradation pathway (Mosbech et al. 2013) (Figure 5). This provides evidence that specific sphingolipid species have opposing roles in lifespan determination.

Ceramides are modified to form more complex sphingolipids by the addition of phosphate or carbohydrate groups to form sphingomyelins, cerebrosides, or gangliosides (Figure 1F). Sphingomyelin is synthesized by the addition of a phosphatidylcholine group to C1 of ceramide, using sphingomyelin synthases encoded by sms-1, sms-2, sms-3 (Huitema et al. 2004) (Figure 5). Sphingomyelins are the most abundant group of sphingolipids. In mammals, these lipids are typically localized to the outer leaflet of plasma membranes, and often are components of lipid microdomains that form a distinct structure for membrane-bound signal transduction proteins. The ACS-20 and ACS-22 activities are required for the addition of long chain fatty acyl groups to sphingomyelin, and acs-20 and acs-20;acs-22 double mutants exhibit surface barrier defects, suggesting that sphingomyelins are components of the nematode cuticle (Kage-Nakadai et al. 2010).

Carbohydrate groups are added to ceramide by the action of three ceramide glucosyl transferases, CGT-1, CGT-2, and CGT-3 (Marza et al. 2009). Single mutants of cgt-1, cgt-2 show no discernable phenotypes; however, cgt-3 mutants have greatly reduced brood size and early embryo defects, and cgt-3;cgt-1 double mutant has growth arrest as L1 (Nomura et al. 2011). Indeed, RNAi of cgt-1 and cgt-3 leads to aberrant lumen formation and L1 arrest, as does RNAi of pod-2, let-767, acs-1, elo-3, and hpo-8, all of which are involved in the synthesis and elongation of saturated fatty acids (H. Zhang et al. 2011). This systematic analysis of L1 intestinal lumen defects led to the identification of d17iso glucosylceramide (d17iso-GlcCer) as the sphingolipid required for proper polarity of intestinal cells and lumen formation in L1 larvae (H. Zhang et al. 2011; Zhu et al. 2013).

An interesting relationship between d17iso-GlcCer and the TORC1 signaling pathway was identified by an unbiased genetic screen in which the requirements for d17iso-GlcCer in larval development were ameliorated by a reduction of function mutation in nplr-3, which encodes a homolog of a NPR3—a nitrogen permease regulator that acts as a negative regulator of the TORC1 pathway (Zhu et al. 2013). Furthermore, hyperactivation of TORC1 also bypasses the need for mmBCFA and d17iso-GlcCer synthesis, leading to the idea that d17iso-GlcCer is required for TORC1 activation, which leads to proper L1 development (Zhu et al. 2013). Furthermore, the NPR-3/TORC1 signaling pathway acts downstream of the d17iso-GlcCer pathway to regulate foraging behavior, which depends on the proper differentiation of sensory neurons in the L1 larval stage, and on proper food-seeking locomotion behavior in the adult (Kniazeva et al. 2015).

Finally, d17iso GlcCer can be further glycosylated by the β-mannosyl transferase BRE-3 and the glycosyltrasferases BRE-2, BRE-4, and BRE-5 (Griffitts et al. 2001, 2003, 2005). An array of complex glycosphingolipids can be formed through these activities. These glycosylated sphingolipids are produced by nematodes and arthropods, but not by mammals, and are targets of Bacillus thuringiensis toxins (Griffitts et al. 2001, 2003, 2005).

Breakdown of sphingolipids and ceramide signaling

The breakdown of plasma membrane sphingomyelins generates ceramides that act as signaling molecules in response to environmental stresses, such as heat shock, infection, radiation, or chemotherapy (Schuchman 2016). C. elegans contains acidic, but not neutral, sphingomyelinase activity (Lin et al. 1998). The acid sphingomyelinases are encoded by the asm-1, asm-2, and asm-3 genes. Ceramides can diffuse through the membrane and activate various stress pathways, such as the JNK and ERK pathways that promote apoptosis and inflammation.

Ceramide is degraded through an acid ceramidase, encoded by asah-1, which deacylates to 17iso-sphingosine, which is subsequently phosphorylated by SPHK-1 sphingosine kinase to 17iso-sphingosine-1-phosphate (Zhang et al. 2015). This can be further degraded by sphingosine-1-phosphate lyase (spl-1), yielding ethanolamine phosphate and a C17iso aldehyde, which can be utilized for production of various other lipids. The spl-1 gene is expressed in intestinal cells after hatching throughout larval development and adulthood (Mendel et al. 2003). Knockdown of spl-1 by RNAi leads to severe defects in intestinal and reproductive function, demonstrating that the breakdown of sphongosine-1 phosphate is essential in C. elegans (Mendel et al. 2003).

Sterol and Prenol Lipids

The mevalonate pathway

Sterol and prenol lipids differ from the acyl-based lipids described above, in that the basic units are the five-carbon branched isoprene groups, isopentyl pyrophosphate (IPP) and dimethylallyl pyrophosphate (DMAPP) (Figure 1B). In plants and animals, these precursors are used to synthesize a large, diverse class of lipids including cholesterol, vitamin D, dolichol, coenzyme Q, and steroid hormones. In C. elegans, RNAi of genes encoding the enzymes of the mevalonate pathway leading to IPP and DMAPP result in embryo lethality, indicating that the synthesis of isoprenenoid precursors is essential (Kamath et al. 2003; Rauthan and Pilon 2011). These precursors are the substrates for various biosynthetic pathways that lead to the synthesis of coenzyme Q, dolichol, isopenteneyl pyrophosphate tRNAs, and prenylated proteins (Figure 6). Unlike mammals, C. elegans does not synthesize cholesterol and other steroids using the mevalonate pathway, because it lacks squalene synthase activity (Vinci et al. 2008). Details of the genes, enzymes, and biochemical reactions of the mevalonate pathway in C. elegans have been described in an extensive review (Rauthan and Pilon 2011).

Figure 6.

Abbreviated pathway of prenol and sterol lipid synthesis in C. elegans. Substrates are shown in yellow ovals, lipid intermediates are depicted in blue ovals, products are depicted in red ovals, and enzymes are shown in purple ovals. -PP, pyrophosphate.

The rate-limiting step in the mevalonate pathway is HMG-CoA reductase, the enzyme that is targeted by statin drugs, which are widely used to treat high cholesterol in humans. Statins induce embryo lethality and induction of ER unfolded protein response (UPR^ER) in C. elegans (Morck et al. 2009). A mutant in the HMG-CoA reductase, hmgr-1, shows phenotypes corresponding to those of the statin-treated C. elegans. The embryonic lethality and UPR^ER phenotypes of the hmgr-1 mutant can be rescued with dietary mevalonate, which enabled the establishment of mevalonate levels required for various biological processes (Ranji et al. 2014). Reproductive processes require the highest levels of mevalonate, while rescuing the growth from L1 to adult stage requires 10-fold less mevalonate. GFP reporter strains revealed that protein prenylation and the distribution of muscle mitochondria are abnormal in both hmgr-1 mutants as well as in nematodes treated with statins (Ranji et al. 2014).

Aside from HMG-CoA reductase, work in C. elegans has revealed that another regulated enzyme in the mevalonate pathway is HMG-CoA synthase. A recent study revealed that HMG-CoA synthase is regulated by post-translational ubiquination and age-related sumoylation (Sapir et al. 2014). The sumoylation levels increase with age, and sumoylation is balanced by the age-regulated activity of ULP-4, a SUMO protease (Sapir et al. 2014).

An interesting indirect role for the mevalonate pathway is its contribution to the silencing of genes by microRNAs (miRNAs). Reduction of activity of several genes, including hmgr-1, and hmgs-1 in the mevalonate pathway by RNAi leads to inappropriate expression of several genes that are normally repressed by miRNAs (Shi and Ruvkun 2012). Specifically, the dolicol branch of the mevalonate pathway is responsible for proper silencing function of miRNAs. Dolicol is a lipid carrier of the oligosaccharide used for N-glycosylation of proteins, a process that is required for proper function of the miRISC complex (Shi and Ruvkun 2012).

The C. elegans gro-1 gene encodes an isopentenylpyrophosphate:tRNA transferase (IPT), which modifies tRNAs (Lemieux et al. 2001). This modification is thought to increase the efficiency of protein translation (Persson et al. 1994). In C. elegans, GRO-1 is required maternally for normal rates of embryogenesis, postembryonic development, and normal brood size. Intriguingly, mitochondrially localized GRO-1 expression, but not cytoplasmic expression, rescues the developmental phenotypes of gro-1 mutants (Lemieux et al. 2001).

Roles of coenzyme Q in C. elegans

Coenzyme Q is an electron carrier of the mitochondrial electron transport chain. Its reduced form, ubiquinol, acts as an antioxidant, inhibiting lipid peroxidation in most subcellular membranes, and recycling other antioxidants such as vitamin E and vitamin C (Ernster and Dallner 1995). The structure of coenzyme Q consists of a benzyoquinone ring that can be reversibly reduced and oxidized, with a prenylated tail of varying number of isoprene units. Bacteria predominately synthesize CoQ₈, while C. elegans and rodents synthesize CoQ₉, and humans predominately contain CoQ₁₀. Eight gene products are required for the synthesis of coenzyme Q from farnesyl PP. These are encoded by the coq-1-coq-8 genes in Saccharomyces cerevisiae and C. elegans, although in C. elegans coq-7 is known as clk-1, because mutants were isolated based on their slow development and long lifespan phenotypes (Wong et al. 1995; Ewbank et al. 1997; Larsen and Clarke 2002). While most mutant strains lacking an enzyme required for coenzyme Q biosynthesis die as larvae, the clk-1 strain is viable. Even though clk-1 mutants fail to synthesize CoQ₉, and instead accumulate dimethoxy CoQ, the CoQ₈ they obtain from their bacterial diet permits development and fertility (Jonassen et al. 2002, 2003). A coq-8 deletion strain synthesizes ∼7% of wild type levels of CoQ9, and mutants show developmental defects, including a detachment of the cuticle from muscle cells and reduced fertility (Asencio et al. 2009). A reduction of CoQ9 and related metabolites occurs during dietary restriction in C. elegans, and this reduction may be due to reduced expression of the coenzymeQ biosynthesis genes, including coq-1 and coq-5 (Fischer et al. 2015). Thus, endogenous coenzyme Q synthesis modulates both development and aging in C. elegans.

Roles of cholesterol in C. elegans

In mammals, cholesterol is an important membrane component required to regulate membrane fluidity, and an important structural component of membrane microdomains. Compared to other animals, C. elegans accumulates very low levels of cholesterol in cellular membranes (Matyash et al. 2001; Merris et al. 2003). Even though C. elegans lacks the ability to synthesize cholesterol (Vinci et al. 2008), they can readily absorb sterols from the diet, and C. elegans are routinely grown in the laboratory in the presence of cholesterol. Cholesterol depletion experiments indicate that very small amounts of cholesterol are required for molting and larval growth, and achieving depletion is difficult due to small amounts of sterols contaminating agar and other media ingredients (Kurzchalia and Ward 2003). Using organic solvent-extracted agar, C. elegans grow normally and are fertile until the F1 generation, which become arrested during larval growth, indicating that sterols are essential for C. elegans development.

Consistent with the requirement for small amounts of steroid precursors, bile-acid like steroid hormones and receptors are required for reproduction and dauer formation in C. elegans. Sterol-derived ligands were shown to bind the NHR DAF-12 (Motola et al. 2006). Synthesis of the ligands, called ∆7 dafachronic acid and ∆4 dafachronic acid, whose structures resemble mammalian bile acids, require the activity of the DAF-9 CYP450, which hydroxylates cholesterol twice at the C26 position to form a carboxyl group (Gerisch et al. 2001; Jia et al. 2002). Other modifications require the DAF-36 Rieske-like monooxygenase, which acts as a 7-desaturase, to convert cholesterol to 7-dehydrocholesterol (Rottiers et al. 2006), as well as the HSD-1 hydroxysteroid dehydrogenase for synthesis (Patel et al. 2008). These and other steroid-derived ligands (Mahanti et al. 2014), together with the DAF-12 receptor, play a major role in regulating developmental and reproductive responses to the environment (Antebi 2015). Synthesis of bile acid ligands are regulated by NHR-8, a homolog of sterol-sensing receptors which acts upstream of DAF-12. NHR-8 regulates the transcription of genes involved in synthesizing dafachronic acids, fatty acid desaturation, and apolipoprotein production (Magner et al. 2013). During dietary restriction, which leads to longer lifespan in C. elegans and many other animals, levels of ∆7 dafachronic acid are increased, and signaling through NHR-8 and LET-363/mTOR mediates the lifespan extension (Thondamal et al. 2014). Thus, steroid hormone signaling is essential for development as well as lifespan regulation in C. elegans.

Introduction to Carbohydrates

Carbohydrates are hydrated organic molecules consisting of carbon (C), hydrogen (H), and oxygen (O), characterized by the formula C_x(H₂O)_y. Chemically, most carbohydrates are polyhydroxy aldehydes, ketones, alcohols, and acids. They are typically classified based on their structural complexity; the simplest moiety is a monosaccharide, a single molecule following the formula C_x(H₂O)_y that cannot be hydrolyzed into smaller carbohydrates. Monosaccharides form polymers referred to as disaccharides, oligosaccharides, and polysaccharides, which consist of repeats monosaccharide moieties with acetal type linkages. These polymers follow the formula [C_x(H₂O)_y]_n, where n = 2 for disaccharides, n = 2–11 for oligosaccharides, and n > 10 for polysaccharides, which can reach up to several thousands of monosaccharide units. Many mono and disaccharides are known as sugars, due to their sweet taste.

Carbohydrates exist in all living kingdoms and have a variety of essential functions. Carbohydrates are a major energy source for all nonphotosynthetic organisms. Animals store glycogen, a polymer of thousands of monosaccharide units, while plants store long polymers of amylose and amylopectin. Other polysaccharides serve vital structural functions, such as chitin in animals, including C. elegans, and cellulose and hemicellulose in plants. Five-carbon carbohydrates such as ribose and deoxyribose are essential components of RNA and DNA, respectively. Ribose is essential for the biosynthesis of biochemical cofactors such as nicotinamide adenine dinucleotide (NAD), flavin adenine dinucleotide (FAD), adenosine mono/di/triphosphate (AMP/ADP/ATP), and others.

Functions and Metabolism of Monosaccharides

Monosaccharides are simple carbohydrates that are classified by the intramolecular position of the carbonyl (carbon–oxygen double bond) group, as well as the number and chirality of the carbons. Stereochemical chirality is defined by the position of the various hydroxyl-groups. Carbonyl groups can either be ketones, defining the corresponding sugar as a ketose, or aldehydes giving rise to aldoses. Thus, monosaccharides carry defining names such as aldohexoses, which include glucose.

Glucose transport, metabolism and function

d-Glucose (Figure 7A) is the best biochemically characterized monosaccharide, while its rare enantiomer l-glucose (Figure 7B) is inaccessible to enzymatic breakdown. The process of glycolysis breaks down d-glucose and converts it into lactate or pyruvate, to generate readily accessible energy equivalents including ATP (Figure 8). Besides β-oxidation and subsequent oxidative phosphorylation (OXPHOS), glycolysis is the most relevant energy-providing process in the animal kingdom, and it is highly conserved and well-characterized in C. elegans (Gebauer et al. 2016; Yilmaz and Walhout 2016). Unlike β-oxidation and OXPHOS, glycolysis does not necessarily require oxygen, and promotes nematodal survival in hypoxic states (LaMacchia et al. 2015).

Figure 7.

Structures of glucose and other monosaccharides. (A) Various projections and isomers of d-glucose. Fisher projections of the respective open-chain forms of (B) l-glucose, (C) d-fructose, and (D) d-galactose, respectively. Structures of (E) the dideoxy sugar ascarolose and (F) the ascaroside daumone-1, also known as ascr#1.

Figure 8.

Glucose uptake, glycolysis and interacting pathways. In C. elegans, glucose is taken up by the FGT-1 glucose transporter. Glucose is phosphorylated by hexokinase and enters various metabolic pathways, including glycolysis, glycogen synthesis, and interacting pathways. –P, phosphate; PEPCK, phosphoenolpyruvate carboxykinase.

Cellular glucose uptake in mammals is mediated by specific glucose transporters (GLUTs) (Herman and Kahn 2006). Recently, eight putative GLUT orthologs have been identified in C. elegans. These were characterized by heterologous overexpression in Xenopus oocytes (Feng et al. 2013; Kitaoka et al. 2013). Only one candidate gene, namely fgt-1, was found to mediate glucose uptake in both the oocyte expression system as well as in nematodes (Feng et al. 2013; Kitaoka et al. 2013). The C. elegans transporter appears to be less substrate-specific than the corresponding, and glucose-specific, GLUTs in mammals, since FGT-1 is capable of transporting other hexoses like fructose, galactose, and mannose (Kitaoka et al. 2013).

Following cellular uptake, glucose is either stored as glycogen, or metabolized by the glycolytic pathway (Figure 8). The early RNAi-based screening approaches have found that inhibition of the glycolytic enzyme glucose phosphate isomerase 1 (GPI-1) extends lifespan (Lee et al. 2003; Hamilton et al. 2005; Schulz et al. 2007), suggesting that reduced glucose metabolism promotes longevity. Consistently, blockade of glycolysis with chemical agents like 2-deoxyglucose, glucosamine, or RNAi-mediated knock-down of fgt-1 also extends lifespan of nematodes (Schulz et al. 2007; Feng et al. 2013; Weimer et al. 2014) and mice (Weimer et al. 2014), respectively.

In mammals, a significant portion of glucose uptake depends on insulin signaling-mediated and translocation-dependent activation of GLUT4 (Huang and Czech 2007). In C. elegans, impaired insulin/IGF-1/INS-18 signaling extends lifespan (Friedman and Johnson 1988; Kenyon et al. 1993), and, consistent with mammalian insulin signaling, reduces glucose uptake (Zarse et al. 2012). Conversely, impairing nematodal glucose transport by inhibition of FGT-1 epistatically interacts with AGE-1 and DAF-2 signaling regarding extension of lifespan, indicating that impaired glucose uptake by knock-down of the transporter (fgt-1 RNAi) and impaired insulin/IGF-1 (AGE-1, DAF-2) likewise reducing glucose share a functional denominator (Feng et al. 2013).

Early approaches to establish axenic media conditions demonstrated than C. elegans can be maintained on glucose as the only energy source, as well as on trehalose (a glucose dimer, see below) and glycogen (a glucose polymer, see below) (Lu and Goetsch 1993). By contrast, other saccharides like fructose or sucrose exerted toxicity. Also, high concentrations of trehalose and glucose inhibited growth (Lu and Goetsch 1993). Researchers have repeatedly demonstrated that high, i.e., millimolar, concentrations of glucose (in addition to the normal E. coli OP50 food source) impair the lifespan of nematodes (Schulz et al. 2007; Lee et al. 2009, 2015; Schlotterer et al. 2009; Choi 2011; Fitzenberger et al. 2013; Deusing et al. 2015a,b; Dobson et al. 2017). Notably, this effect is not observed in males (Miersch and Doring 2012; Liggett et al. 2015), possibly due to sex-specific differences in expression of genes involved into carbohydrate catabolism (Miersch and Doring 2012).

Experimental evidence points to several mechanisms mediating the lifespan reduction by glucose. Reducing glycolysis has been shown to induce mitochondrial OXPHOS to generate a lifespan-extending reactive oxygen species (ROS) signal (Schulz et al. 2007) due to a process called mitohormesis (Ristow 2014) that has been confirmed in multiple organisms, including C. elegans (Zarse et al. 2012; K. Schmeisser et al. 2013; S. Schmeisser et al. 2013; De Haes et al. 2014; Tauffenberger et al. 2016). Conversely, increased glucose levels reduce both OXPHOS and ROS, impairing lifespan (Schulz et al. 2007; Zarse et al. 2012). Independently, it has been shown that increased glucose concentrations direct excess glucose-6-phosphate into the hexosamine pathway causing formation of methylglyoxal and advanced glycated end products to limit longevity (Morcos et al. 2008; Schlotterer et al. 2009; Chaudhuri et al. 2016). Third, because glucose affects formation of the metabolic intermediate glycerol, the aquaporin AQP-1 has been linked to osmotic stress tolerance and lifespan regulation following exposure to glucose (Lee et al. 2009). Fourthly, and since blocking GPI-1 by RNAi or chemical inhibitors forces glucose-6-phosphate into the pentose phosphate shunt (Figure 8), subsequent alterations in nematodal redox state have been proposed to mediate the phenotype (Ralser et al. 2008; Kruger et al. 2011; Penkov et al. 2015), given that the pentose phosphate shunt is responsible for generation of NADPH, which, in turn, affects levels of reduced glutathione (GSH), besides others. Along this line, it was subsequently shown that depletion of NADPH also initiates dauer formation due to impaired biosynthesis of dafachronic acid from cholesterol—a NADPH-dependent process (Penkov et al. 2015). Consistently, it was shown that carnitine protects from glucose toxicity in a DAF-12-dependent manner, notably the receptor for dafachronic acid (Deusing et al. 2015a). Other mechanism that ameliorate glucose toxicity are (i) modulation of membrane lipid composition via PAQR-2 and IGLR-2 (Svensk et al. 2016), (ii) reduction of glucose-mediated triglyceride accumulation via SREBP and MDT-15 (Lee et al. 2015), and lastly O-linked N-acetylglucosamine cycling (Love et al. 2010; Mondoux et al. 2011). Lastly, exposure to glucose reduces fecundity in a transgenerational manner (Tauffenberger and Parker 2014).

Despite this extensive body of evidence regarding glucose toxicity and its underlying mechanisms, excess glucose has unexpectedly been demonstrated to reduce proteotoxicity in several C. elegans models of protein aggregation (Tauffenberger et al. 2012). It remains to be shown how this interesting observation translates into higher organisms, particularly since diabetes mellitus, and hence elevated glucose levels, have been linked to increased incidence of Alzheimer’s disease in multiple human cohorts (Zhang et al. 2017).

Fructose and galactose

As stated above, in comparison to glucose and glucose-derived di- and polymers, the monosaccharide fructose (Figure 7C) has been shown to be significantly less effective in maintaining viability of nematodes in axenic culture (Lu and Goetsch 1993). Very recently, fructose has been shown to extend lifespan at lower doses while higher doses shorten lifespan, both in liquid culture and due to unknown mechanisms (Zheng et al. 2017). Moreover, fructose at lower doses appears to reduce intestinal lipid deposition, while higher doses induce the opposite, potentially reflecting the effects of this sugar on hepatic lipid accumulation in mammals (Zheng et al. 2017).

Galactose (Figure 7D) is a monosaccharide that serves as an important component in the formation of—widely undefined—glycolconjugates, i.e., the plethora of lipid or protein molecules with at least one sugar moiety attached to it. While beyond the primary/metabolic scope of this text, such changes will be discussed below.

Independently, galactose metabolism has been analyzed in C. elegans, and essentially reflects the so-called Leloir pathway (Holden et al. 2003), where galactose is converted into glucose derivatives to make it accessible for glycolytic catabolism (Figure 9). Enzymatic defects of this pathway have been linked to an inherited human disease named galactosemia. The Leloir pathway requires four enzymes: the first step is the conversion of β-galactose into to α-galactose by an enzyme called galactose mutarotase, which has been crystallized for both humans and C. elegans (Thoden et al. 2004). The second step is catalyzed by galactokinase to yield galactose-1-phosphate. The third enzyme is galactose-1-phosphate uridyl-transferase, which transfers an UMP group from UDP-glucose to galactose-1-phosphate, thereby generating glucose 1-phosphate and UDP-galactose. The last step is the conversion of UDP-galactose into UDP-glucose by UDP-galactose 4-epimerase, named GALE-1 in nematodes. Complete loss-of-function mutations of gale-1 cause lethality, while partial loss-of-function mutants are viable (Brokate-Llanos et al. 2014). Moreover, the authors observe hypersensitivity to galactose supplementation, as well as to infections in partial loss-of-function mutants of gale-1, as well as interactions with the unfolded protein response (Brokate-Llanos et al. 2014).

Figure 9.

Galactose metabolism. The conserved Leloir-pathway converts galactose to glucose-6 phosphate using the activity of four key enzymes.

Sugar alcohols

Sorbitol (Figure 10A) is an alcohol produced from glucose by the enzyme aldose reductase, which, in mammals, occurs as a consequence of supraphysiologically increased glucose levels. This reaction requires NADPH, and may hence impair redox defense of the cell by depleting GSH (see above). Sorbitol can further be metabolized to fructose by sorbitol dehydrogenase, which requires NAD+ as a cofactor. Collectively, these two enzymatic steps are known as the polyol pathway (Figure 10B). Sorbitol was found to be most strikingly elevated in a global screen for aging-associated metabolites in C. elegans, and, consistently, a change of redox state was observed in the same experimental setting (Copes et al. 2015).

Figure 10.

Sorbitol and the polyol pathway. (A) The structure of the sugar alcohol sorbitol. (B) The interconversion of glucose, sorbitol, and fructose by the polyol pathway.

By contrast, sorbitol supplementation has been shown to extend nematodal lifespan, likely due to induction of osmotic stress and subsequent induction of stress response pathways (Chandler-Brown et al. 2015; Hipkiss 2015). The same mechanism may also apply to fructose-mediated lifespan extension (see above) (Zheng et al. 2017), as well as to the concomitant effects of the disaccharide trehalose (see below).

Dideoxysugars (ascarosides)

Ascarosides are glycolipids that, by definition, always contain the dideoxysugar ascarylose (3,6-dideoxy-l-arabino-hexose) (Figure 7E) linked to a fatty acid of varying chain length. One well-characterized example is the first ascaroside isolated from C. elegans, daunome-1, also known as ascr#1, which carries a saturated C7 fatty acid, i.e., enanthic or heptanoic acid (Figure 7F). Fatty acids linked to ascarylose can be saturated (as in daunome-1), or unsaturated. Ascarosides were initially identified in the roundworm family Ascaridia spp., and have been subsequently identified in C. elegans to function as extracellular signaling molecules. In the latter, ascarosides control the dauer state as well as male attraction, hermaphrodite repulsion, olfactory plasticity, and aggregation. They act mainly through G protein-coupled receptors (GPCRs) (Pungaliya et al. 2009; Ludewig et al. 2013), and have been excellently described elsewhere in this publication (Ludewig and Schroeder 2013).

Functions and Metabolism of Disaccharides, Oligosaccharides, and Polysaccharides

Trehalose

Disaccharides, by definition, contain two monosaccharides formed by glycosidic linkage, i.e., by a condensation reaction at two hydroxyl groups. While sucrose, lactose, and maltose are the most relevant disaccharides in mammalian biology, trehalose (Figure 11A) is the only disaccharide that has been characterized in nematodes to a significant extent.

Figure 11.

Structure and metabolism of trehalose and maradolipids. (A) Structure of the disaccharide trehalose. (B) Biosynthesis and catabolism of trehalose. Trehalose-P synthase is encoded by tps-1 and tps-2 genes, Trehalose-6-phosphate phosphatase is encoded by gob-1, and Trehalase is encoded by tre-1-5 genes (C) General structure of maradolipids. R, fatty acyl chains, which frequently are branched (see text for details).

Trehalose is found in many lower organisms, including plants, fungi, and metazoae, and serves different purposes importantly including energy storage and stress resistance. Chemically, trehalose is formed from two molecules of d-glucose by an α,α-1,1-glucoside bond (Figure 11B). Its precursor, trehalose-6-phosphate, is synthesized from glucose-6-phosphate and UDP-glucose by an enzyme called trehalose-6-phosphate synthase, encoded by tps-1 and tps-2 in C. elegans. Impairing expression of both tps genes reduces trehalose content in C. elegans by >90% (Pellerone et al. 2003). Trehalose-6-phosphate is next dephosphorylated by an enzyme named trehalose-6-phosphate phosphatase (TPP), which, in C. elegans, is encoded by gob-1. While loss-of-function mutants of gob-1 are lethal, impairment of tps-1 and tps-2 reverts gob-1 lethality (Kormish and McGhee 2005). This indicates that accumulation of trehalose-6-phosphate, rather than lack of trehalose, causes lethality in nematodes.

Like glycogen, nematodes store trehalose to conserve energy (Roberts and Fairbairn 1965), and the disaccharide is made reavailable to glycolysis by cleavage to glucose mediated by an enzyme called trehalose (Figure 11B). This enzyme has been proposed to be encoded by the tre gene family in C. elegans (Pellerone et al. 2003). Consistent with its role in energy storage, trehalose has been involved in the hatching process in nematodes (Perry and Clarke 1981). Nevertheless, the trehalose-to-glucose ratio is 5.4 times higher in males than in hermaphrodites (Miersch and Doring 2012).

Trehalose has a long-standing role as a stress protectant in various species (Van Laere 1989). Consistently, trehalose feeding extends C. elegans lifespan, and daf-2-mediated extension of lifespan is reduced by impairing tps-1 or tps-2 (Honda et al. 2010), while daf-2 mutants show increased levels of tps-1 and -2 mRNA expression (Lamitina and Strange 2005; McElwee et al. 2006; Honda et al. 2010; Zarse et al. 2012). Moreover, C. elegans increases endogenous trehalose production in states of extreme desiccation to ensure dauer survival in states of anhydrobiosis (Erkut et al. 2011). Preconditioning by mild desiccation for 4 days potentiated survival rates in states of extreme desiccation. Interestingly, mutants unable to synthesize trehalose lost the ability to survive desiccation in the dauer state, indicating that trehalose is not only associated with, but also the cause of, survival in states of anhydrobiosis (Erkut et al. 2011). Subsequently, the same authors showed that trehalose biosynthesis in states of desiccation mainly depends on the glyoxylate shunt, where sugars are synthesized from fatty acids, particularly acetate (Erkut et al. 2016), a mechanism also observed in S. cerevisiae, with possible translation into mammals in states of ketosis (Hu et al. 2014). The NADPH-dependent biosynthesis of trehalose during adverse conditions may also initiate dauer formation, since it was shown that depletion of NADPH promotes dauer formation due to impaired biosynthesis of dafachronic acid from cholesterol, a NADPH-dependent process (Penkov et al. 2015).

While trehalose is absent in mammals, trehalose supplementation nevertheless induces autophagy and prevents hepatic steatosis in murine cells, possibly by inhibiting the SLC2A protein (DeBosch et al. 2016). Moreover, trehalose has been successfully used for cryopreservation of mammalian and particularly human cells (Eroglu et al. 2000; Guo et al. 2000).

Maradolipids

Fatty acids can form condensation products with sugars—a process contributing to the glycosylation of membranes and other moieties. The disaccharides trehalose can form specific condensation products with two—typically different—types of fatty acid molecules. These condensation products have been named maradolipids (Penkov et al. 2010) and chemically reflect 6,6′-di-O-acyl-trehaloses (Figure 11C). Typically, one of the acyl moieties reflects oleic acid (18:1n-9), while the other moiety appears to be a branched-chain fatty acid of variable molecular structure (Penkov et al. 2010). Maradolipid-like components are widely found in plants and prokaryotes, and contribute to desiccation resistance in these species (Harland et al. 2008). Consistently, maradolipids are preferentially synthesized during dauer formation in C. elegans (Penkov et al. 2010). Consistent with the fact that maradolipid synthesis requires branched-chain fatty acids, RNA interference against elo-5, which is required for formation of such fatty acids (Kniazeva et al. 2004, 2008), prevents maradolipid biosynthesis in nematodes (Penkov et al. 2010). Likewise, impairing enzymes like tps-1 or tps-2 that are required for trehalose biosynthesis (Honda et al. 2010) impairs formation of maradolipids (Penkov et al. 2010).

Glycogen

In addition to trehalose, glycogen is the main carbohydrate storage in C. elegans. Glycogen consists of up to several thousands of glucose molecules, which can be released from glycogen by an enzymatic process called glycogenolysis. Inversely, excess glucose is stored as glycogen by activation of glycogen synthase (Figure 8). There is longstanding evidence for glycogen storage as well as glucose mobilization from glycogen in C. elegans (O’Riordan and Burnell 1989). In the dauer state, nematodes not only accumulate significant amounts of lipids (see above) but also glycogen (Popham and Webster 1979). Consistent with its role as energy buffer, glycogen is preferentially mobilized in the dauer state, while glycogen synthesis is downregulated (O’Riordan and Burnell 1989; Holt and Riddle 2003; Depuydt et al. 2014). Since glucose can be used to generate ATP in the absence of oxygen by means of anaerobic glycolysis (O’Riordan and Burnell 1989; Braeckman et al. 2009) (Figure 8), it has been consistently demonstrated that increased glycogenolysis mediates survival in states of anoxia (Frazier and Roth 2009; Larue and Padilla 2011; Depuydt et al. 2014; LaMacchia et al. 2015) and hypo-osmotic stress (LaMacchia et al. 2015).

Chitin

Chitin is a long-chain polymer of N-acetyl-glucosamine. The latter chemically qualifies as an N-glycosylated alanine (see below). N-Acetyl-glucosamine is derived from glucose through the hexosamine pathway, and has been shown to extend lifespan in C. elegans by inducing protein quality control (Denzel et al. 2014).

Chitin is an important structural component of C. elegans. Chitin is polymerized from N-acetyl-glucosamine monomers by two isoforms of chitin synthase, chs-1 and chs-2 (Veronico et al. 2001): chs-1 mediates chitin deposition into the eggshell, and is expressed during later larval stages and adulthood, while chs-2 provides chitin for the nematodal pharynx, and is expressed before the molt (Veronico et al. 2001; Zhang et al. 2005).

Glycosylation

Glycosylation is defined as the covalent addition of a carbohydrate molecule to lipid, nucleic acids, or protein moieties, leading to characteristic and functionally relevant modifications. Often oligosaccharides, consisting of >2 but <11 monosaccharide units, contribute to the glycation of proteins and lipids. Glycosylation is a specific and enzyme-directed process, and genetic defects in glycosylation cause numerous inherited diseases in humans. Particularly, the addition of sugars to the N- or the C-terminus of proteins to form N- and O-glycans has been intensely studied. Glycosylation of lipids typically reflects O-linked glycosylation. The numerous specific aspects of glycosylation in C. elegans have been excellently reviewed elsewhere (Berninsone 2006).

Summary and Future Directions

Metabolic pathways of lipid and carbohydrate metabolism are well conserved across the animal kingdom, including C. elegans. Interestingly, C. elegans also possess pathways that are rare in animals, but common in plants, such as the ability to synthesize polyunsaturated fatty acids in the absence of essential dietary fatty acids, and the synthesis of maradolipids for protection from desiccation. Specific lipids and carbohydrates play vital roles in development, and C. elegans research has been especially useful for unraveling the roles of lipids and carbohydrates during aging and stress. As in other animals, lipid and carbohydrate metabolism pathways converge, because animals have a limited capacity to store dietary carbohydrates, and thus carbons from dietary carbohydrates are converted to acetyl-CoA, where they can be diverted into fatty acid synthesis. The consequences of disrupting this linkage are devastating, as glucose toxicity is greatly elevated in both worms and humans when fatty acid synthesis regulators and pathways are disrupted (Lee et al. 2015; Listenberger et al. 2003).

Future research in lipid and carbohydrate metabolism in C. elegans will benefit from the rapid improvement of metabolomic analysis. In addition to lipids and carbohydrates, it will be important to analyze the metabolism of amino acids and intermediates of other metabolic pathways with respect to various developmental and environmental manipulations. Because the C. elegans diet consists of bacteria, and the major macronutrient by weight in bacteria is protein, it will be useful for future research to focus on catabolic and anabolic pathways of amino acid metabolism, including the interplay between amino acid breakdown products and lipid and carbohydrate metabolism. Combined with the powerful genetic approaches common in C. elegans research, this will lead to more insights into mechanisms of energy homeostasis and roles for macronutrients in development, aging, and stress responses.