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Keywords:

  • aging;
  • C. elegans;
  • longevity;
  • metabolic rate;
  • metabolism

Summary

  1. Top of page
  2. Summary
  3. Introduction
  4. Life history characteristics of C. elegans
  5. Growth of C. elegans in the laboratory
  6. C. elegans metabolic measurements
  7. Indirect measures of C. elegans metabolism
  8. Long-lived C. elegans mutants and aging
  9. Comparisons of metabolic rate between wild-type and long-lived mutants
  10. Factors potentially complicating metabolic measurements in C. elegans
  11. Conclusion
  12. Acknowledgments
  13. References

Much of the recent interest in aging research is due to the discovery of genes in a variety of model organisms that appear to modulate aging. A large amount of research has focused on the use of such long-lived mutants to examine the fundamental causes of aging. While model organisms do offer many advantages for studying aging, it also critical to consider the limitations of these systems. In particular, ectothermic (poikilothermic) organisms can tolerate a much larger metabolic depression than humans. Thus, considering only chronological longevity when assaying for long-lived mutants provides a limited perspective on the mechanisms by which longevity is increased. In order to provide true insight into the aging process additional physiological processes, such as metabolic rate, must also be assayed. This is especially true in the nematode Caenorhabditis elegans, which can naturally enter into a metabolically reduced state in which it survives many times longer than its usual lifetime. Currently it is seen as controversial if long-lived C. elegans mutants retain normal metabolic function. Resolving this issue requires accurately measuring the metabolic rate of C. elegans under conditions that minimize environmental stress. Additionally, the relatively small size of C. elegans requires the use of sensitive methodologies when determining metabolic rates. Several studies indicating that long-lived C. elegans mutants have normal metabolic rates may be flawed due to the use of inappropriate measurement conditions and techniques. Comparisons of metabolic rate between long-lived and wild-type C. elegans under more optimized conditions indicate that the extended longevity of at least some long-lived C. elegans mutants may be due to a reduction in metabolic rate, rather than an alteration of a metabolically independent genetic mechanism specific to aging.


Introduction

  1. Top of page
  2. Summary
  3. Introduction
  4. Life history characteristics of C. elegans
  5. Growth of C. elegans in the laboratory
  6. C. elegans metabolic measurements
  7. Indirect measures of C. elegans metabolism
  8. Long-lived C. elegans mutants and aging
  9. Comparisons of metabolic rate between wild-type and long-lived mutants
  10. Factors potentially complicating metabolic measurements in C. elegans
  11. Conclusion
  12. Acknowledgments
  13. References

There is a great deal of interest in the use of the nematode C. elegans as a model organism in aging studies. Much of this interest is due to the identification of numerous genes in C. elegans that can significantly increase worm longevity when mutated (Johnson et al., 2000). These mutations potentially identify genes specifically involved in aging, and an extensive research effort is currently underway to elucidate the mechanisms responsible for the extended longevity of these mutants (Guarente & Kenyon, 2000). Care must be taken though in interpreting the significance of the means by which longevity is extended in C. elegans mutants. Taken alone, measures of the chronological longevity of an organism may not be meaningful without additional studies on other physiological processes in the organism (Sohal et al., 2000). Measures of physiological processes are especially important when considering factors that increase longevity in an ectothermic organism such as C. elegans.

One example of the importance of physiological processes to longevity is the influence of growth temperature on ectotherm life span. Chronological longevity can be increased in most ectotherms simply by reducing the temperature at which the organism is reared, and C. elegans grown at low temperatures live around four times longer than high-temperature counterparts (Klass, 1977; Van Voorhies & Ward, 1999). However, despite this increase in chronological longevity, the lifetime metabolic output of C. elegans reared at lower temperatures is the same, or even reduced, compared to animals reared at higher temperatures (Van Voorhies & Ward, 1999). The large effect of temperature on longevity is thought to be mediated by the influence of temperature on metabolic rate, with metabolic rate linked to aging via the production of free-radicals and other oxidants formed during aerobic respiration (Ragland & Sohal, 1975; Miquel et al., 1976; McArthur & Sohal, 1982; Beckman & Ames, 1998; Sohal et al., 2000).

A similar reduction in metabolic rate could be responsible for the increased longevity of long-lived C. elegans mutants. This explanation predicts that the increased longevity of these mutants is a consequence of a reduction in their metabolic rate, rather than an alteration of a metabolically independent genetic mechanism specific for aging. Thus, these long-lived mutants could live their lives at high temperature as if they are at a low temperature, producing so-called ‘refrigerator mutants’ (Martin, 2002). The reduced metabolic rate of long-lived mutants could also explain their increased stress resistance (Johnson et al., 2000), since many of the factors that reduce metabolic rate also generally increase the ability of the organism to withstand stress (Sohal et al., 2000). As such it would not be necessary to invoke any other novel or unknown pathways of stress resistance to explain the increased longevity of these mutants.

Evaluating this explanation requires a careful assessment of the metabolic rate of wild-type and long-lived C. elegans mutants. While several studies measuring metabolic rate in C. elegans have been conducted, contradictory conclusions have been drawn regarding the metabolic rate of long-lived mutants relative to wild-type C. elegans. These different conclusions appear to be largely due to differences in the measurement techniques and the conditions used when assaying C. elegans metabolic rates. Accurately determining the standard metabolic rate of any organism requires the maintenance of conditions that minimize environmental stresses, and takes into account the evolutionary history and natural environment of the organism. For such measurements, the organism should be maintained within its thermal limits, not be nutritionally challenged, and subjected to a minimum of physiological stress. In other words, studies correlating the metabolic rate of an organism to its longevity should measure metabolic rates under conditions that closely match the conditions used to assay longevity. These conditions are essential for establishing a reference point from which the metabolic effects of other factors can be compared. Careful control of environmental conditions is especially important when measuring metabolism in C. elegans. Endothermic animals, such as mammals, typically maintain high, relatively constant metabolic rates. In contrast, ectotherms can greatly reduce their metabolic rate and continue to maintain required physiological functions, although at lower levels of activity. This allows C. elegans to potentially survive environmental conditions or metabolic mutations that induce a state of metabolic depression that would be lethal to endothermic organisms.

Life history characteristics of C. elegans

  1. Top of page
  2. Summary
  3. Introduction
  4. Life history characteristics of C. elegans
  5. Growth of C. elegans in the laboratory
  6. C. elegans metabolic measurements
  7. Indirect measures of C. elegans metabolism
  8. Long-lived C. elegans mutants and aging
  9. Comparisons of metabolic rate between wild-type and long-lived mutants
  10. Factors potentially complicating metabolic measurements in C. elegans
  11. Conclusion
  12. Acknowledgments
  13. References

The starting point when conducting metabolic rate studies on an organism is to carefully consider the natural history of the organism. Several aspects of C. elegans natural history that are important in metabolic studies include the nutrition state of the worms, the growth media used and the measurement temperature. C. elegans is a terrestrial, free-living nematode that feeds on soil-dwelling bacteria. C. elegans survives growth temperatures ranging from 6 to 26 °C, but reproduces and grows poorly at either temperature extreme (Klass, 1977). Fecundity in C. elegans is particularly sensitive to high temperatures and decreases rapidly above 24 °C (Byerly et al., 1976; Hirsh et al., 1976). Under abundant food conditions, C. elegans ingests up to several times its body mass per day in bacteria (Seymour et al., 1983; Ferris et al., 1997) by contracting its pharynx over 200 000 times per day to engulf bacteria into the lumen where they are concentrated, ground-up and passed into the intestine for digestion (Anderson, 1978; Avery & Horvitz, 1990; Avery, 1993; Raizen et al., 1995). C. elegans can convert these ingested bacteria into body mass, either in the form of an increase in body size or as egg production, at rates approaching 10% of its body mass per hour (Byerly et al., 1976). Consistent with its rapid growth and food ingestion rate, the mass-specific metabolic rate (metabolic rate per unit body mass) of C. elegans is approximately that of highly trained human athletes at maximal performance (Mortola et al., 1999). Equally impressive, while humans can only sustain these high metabolic rates for a relatively short period, C. elegans maintains this high metabolic rate for days at a time.

The maintenance of this high metabolic rate requires appropriate environmental conditions and an abundant food supply. Under stressful conditions, C. elegans can cease feeding and survive for many weeks entirely on stored energy reserves. C. elegans’ most starvation-resistant state, the dauer larva, enters a period of dormancy characterized by a reduction in activity and metabolic rate and is hyper-resistant to a variety of environmental stresses including high temperatures, oxidative stress and hypoxia (Anderson, 1978; Larsen, 1993; Riddle et al., 1997). Dauer larvae survive up to several months, many times longer than the 2- to 3-week-long life span typical for C. elegans under optimized growth conditions (Klass & Hirsh, 1976). The dauer formation pathway in C. elegans is complex, and several thousand genes appear to be expressed only in dauer larvae (Jones et al., 2001). The ability of C. elegans to rapidly switch from ingesting and metabolizing several times its body mass in bacteria per day to a total cessation of food intake is consistent with its ecology. Food is ephemeral in a soil environment, and soil nematodes must often survive periods of reduced food availability (Ferris et al., 1997). C. elegans can also quickly recover normal function even after greatly reducing its metabolism (Van Voorhies & Ward, 2000). The ability of C. elegans to quickly reduce its metabolism in response to stressful conditions has important implications when measuring both its longevity and metabolic rate.

Growth of C. elegans in the laboratory

  1. Top of page
  2. Summary
  3. Introduction
  4. Life history characteristics of C. elegans
  5. Growth of C. elegans in the laboratory
  6. C. elegans metabolic measurements
  7. Indirect measures of C. elegans metabolism
  8. Long-lived C. elegans mutants and aging
  9. Comparisons of metabolic rate between wild-type and long-lived mutants
  10. Factors potentially complicating metabolic measurements in C. elegans
  11. Conclusion
  12. Acknowledgments
  13. References

Laboratory populations of C. elegans are typically grown either on solid agar or in liquid culture medium (Wood, 1988). The growth rate, fecundity, metabolic rate and longevity of C. elegans differ widely between these two media. When reared in solid medium, C. elegans are grown on Petri plates filled with a nematode growth medium agar. A ‘lawn’ of Escherichia coli is grown on the surface of this agar as a food source for the worms (Wood, 1988). C. elegans can rapidly move across the agar surface using a sinusoidal motion characteristic of free-living nematodes and use chemotaxis to locate bacteria on which they feed continuously.

When reared in liquid medium, worms are placed in buffer medium containing cholesterol and fed on either E. coli or a food source comprised of a small amount of haemoglobin, yeast extract and soy protein (Gandhi et al., 1980; Jansson et al., 1986; Ebert et al., 1996). In liquid as compared to solid medium, C. elegans grows more slowly, has an altered morphology, cannot mate, has reduced fertility and adult worms are subject to up to 90% mortality rates from internal egg hatching (Braeckman et al., 2000; Mitchell et al., 1979; Gandhi et al., 1980). Some mutants cannot be grown in liquid culture (Hekimi, 2000). Worms which avoid internal egg hatching when grown in liquid culture also live about 50–100% longer than on solid medium (Mitchell et al., 1979; Braeckman et al., 2000). This increased longevity may appear paradoxical, but probably results from the reduced levels of bacteria or nutrients present in liquid medium inducing a state of caloric restriction on the worms. In liquid culture, the density of E. coli is typically around 109 per mL of medium, and higher bacterial concentrations have detrimental effects on C. elegans longevity (Klass, 1977). This bacterial density is more than two orders of magnitude lower than the bacterial lawn on which C. elegans feeds in solid medium which, assuming that half of a bacterial lawn is pore space, has an estimated density of ∼5 × 1011E. coli mL−1 (Ingraham et al., 1983). C. elegans in liquid culture could theoretically adjust to lower bacterial concentrations by increasing their pharyngeal pumping rate or the amount of bacteria ingested per contraction. Neither of these, however, would be sufficient to compensate for the reduced concentration of bacteria in liquid medium (Seymour et al., 1983; Avery & Horvitz, 1990) and C. elegans grown in liquid appear semi-starved (Jansson et al., 1986; Liu et al., 1997). Another difficulty of growing C. elegans in liquid culture is that oxygen diffusion rates are approximately four orders of magnitude slower in liquid than in air (Denny, 1993). Probably because of this, the growth and reproduction of C. elegans in liquid culture are very sensitive to aeration levels (Gandhi et al., 1980).

C. elegans metabolic measurements

  1. Top of page
  2. Summary
  3. Introduction
  4. Life history characteristics of C. elegans
  5. Growth of C. elegans in the laboratory
  6. C. elegans metabolic measurements
  7. Indirect measures of C. elegans metabolism
  8. Long-lived C. elegans mutants and aging
  9. Comparisons of metabolic rate between wild-type and long-lived mutants
  10. Factors potentially complicating metabolic measurements in C. elegans
  11. Conclusion
  12. Acknowledgments
  13. References

Several studies have used a variety of techniques to measure the metabolic rate of C. elegans or to make inferences about its metabolism. Such measurements typically either measure the metabolic rate of intact, living C. elegans, or measure a biochemical process or concentration in killed worms as a proxy for metabolism during life. Techniques used to measure metabolic rates in live C. elegans include heat flux measurements (microcalorimetry) and determinations of oxygen consumption or CO2 production. These measurements are based on the biochemistry of aerobic respiration, during which metabolic substrates are oxidized to carbon dioxide and water with free energy from this reaction used to drive the phosphorylation of ADP to ATP (Hochachka & Somero, 1984). Aerobic metabolism in C. elegans appears very similar to mammalian respiratory metabolism (Murfitt et al., 1976; Anderson, 1978; Blum & Fridovich, 1983; O’Riordan & Burnell, 1990). C. elegans requires the use of aerobic respiration to move, develop and survive (Anderson, 1978; Van Voorhies & Ward, 2000), and mutations in mitochondrial respiratory complex genes can be lethal (Tsang et al., 2001).

Microcalorimetry directly assays heat produced by an organism and, assuming that the organism is not carrying out physical work or growing, measures the organism's total metabolic rate including aerobic and anaerobic metabolism. Heat production and oxygen consumption are directly related to each other by a calorimetric–respirometric ratio (Gnaiger & Kemp, 1990), which varies from −470 kJ mol−1 O2 during fully aerobic metabolism to negative infinity during completely anaerobic metabolism. While microcalorimetry can be highly sensitive and does measure the total metabolism of an organism, in practice it is rather cumbersome to use. Consequently, the metabolic rate of aerobic organisms is most commonly determined by measures of oxygen consumption. Measures of metabolic rate by oxygen consumption are potentially the most relevant to aging since reactive oxygen species formation during oxidative phosphorylation is the mechanism most commonly proposed to link metabolic rate and aging.

CO2 production by an organism is also frequently used to measure the rate of metabolism, especially for long-term metabolic measurements, or for metabolic measurements that require high sensitivity (Walsberg & Wolf, 1995; Nagy et al., 1999). While the use of CO2 flux to measure metabolic rate is a widely used and accepted method, several factors must be considered when using CO2 production to calculate energy expenditure (Elia, 1991; Walsberg & Wolf, 1995). CO2 is produced as a direct by-product of oxidative metabolism, but the ratio of CO2 produced to oxygen consumed varies with the metabolic substrate utilized. This ratio, often called the respiratory quotient (RQ), typically ranges from 1.0 for starch and other carbohydrate substrates to 0.71 for ingested lipid (Hochachka & Somero, 1984). Accurately calculating energy expenditures from CO2 production requires a knowledge of the animal species used, the end products of metabolism (e.g. nitrogen end products), the respiratory quotient and the energy balance of the organism (Gessaman & Nagy, 1988; Elia, 1991; Walsberg & Wolf, 1995). However, by carefully considering these parameters, CO2 production can be used to measure metabolic rate with accuracy similar to measurements based on heat flux or oxygen consumption (Elia, 1991; Gessaman & Nagy, 1988). With the techniques and instrumentation currently available the most sensitive and accurate method for assaying metabolic rate in small organisms such as C. elegans is through the detection of CO2 output. This is particularly true when CO2 measurements are combined with measurements on larger numbers of worms that simultaneously determine both CO2 production and oxygen consumption. The combination of these parameters allows measuresments of CO2 production to be accurately converted to standard energy units. Such a combination of measurements also accounts for the use of alternative metabolic pathways that could potentially bias metabolic rate measurements based solely on CO2 production.

Estimates of the metabolic rate of C. elegans as determined by several different investigators are summarized in Table 1. To minimize the potentially confounding effects of differences in body size, strain differences and worm age, only the metabolic rates of young adult, wild-type C. elegans are listed. There is approximately a five-fold range of reported metabolic rates. One factor explaining some of this variation is that most investigators measured metabolic rate in C. elegans by placing unfed worms in liquid and monitoring changes in dissolved oxygen levels. A major complication of this protocol is that the metabolic rate of unfed worms is rapidly reduced. As shown in Fig. 1, under such conditions the metabolic rate of C. elegans decreases several-fold relative to worms maintained on an abundant food supply.

Table 1.  Metabolic rates in wild-type C. elegans. The term solid or liquid refers to the worms either being on solid agar medium on in liquid medium. The listed food source is either E. coli or an axenic medium containing yeast extract, and soy protein instead of bacteria. Metabolic measurements on fed C. elegans were done on animals fed heat-killed bacteria. C. elegans grow normally and have normal longevity on this bacteria and heat-killed bacteria do not significantly contribute to the measured CO2 signal. The number of worms used in the metabolic measurements should be considered rough estimates except for (De Cuyper & Vanfleteren, 1982; Van Voorhies & Ward, 1999). ND indicates no data. Metabolic measurements reported in values per mg protein (Vanfleteren & De Vreese, 1996; Braeckman et al., 1999, 2002) were converted to wet weight values by dividing by 5 (Vanfleteren, personal communication). The two values listed for Braeckman et al. (2002) were obtained using either microcalorimetry (7.5 µW – this group also appears to have been measured with food) or with oxygen consumption methods (19 µW). Different measurement units were converted to the International System of Units (SI) unit of Watts using standard oxycaloric values (Gnaiger & Kemp, 1990). Measurements made at 25 °C were extrapolated to 20 °C assuming a Q10 value of 2.0 (Anderson, 1978; Dusenbery et al., 1978; Van Voorhies & Ward, 1999)
StudyGrowth conditionsMeasurement conditionsMetabolic rate (µW/mg wet wt)
De Cuyper & Vanfleteren (1982)Liquid, axenic, 21 °CLiquid, no food, 20 °C, 7 worms 7
Dusenbery et al. (1978)Solid, E. coli, 20 °CLiquid, no food, 20 °C, 1000s worms13
Anderson (1978)Solid, E. coli, 20 °CLiquid, no food, 20 °C, ND13
De Cuyper (1982)Liquid, axenic, 21 °CLiquid, no food, 20 °C, 7 worms 7
Ferris et al. (1997)Solid, E. coli, 20 °CLiquid, no food, 20 °C, 1000s worms14
Vanfleteren & De Vreese (1996)Liquid, E. coli, NDLiquid, no food, 25 °C, 1000s worms22
Vanfleteren (1996)Liquid, E. coli, NDLiquid, no food, 25 °C, 1000s worms22
Van Voorhies & Ward (1999)Solid, E. coli, 20 °CSolid, E. coli, 20 °C, 50 worms28
Braeckman et al. (1999)Liquid, E. coli, 24 °CLiquid, no food, 25 °C, 1000s worms38
Braeckman et al. (2002)Liquid, axenic, 24 °CLiquid, no food, 25 °C, 1000s worms 7.5/19
image

Figure 1. Effect of food deprivation on metabolic rate in C. elegans. Data are from groups of hundreds of young adult C. elegans placed in metabolic chambers without food. To avoid the effect of progeny production and growth on metabolic rate a sterile mutant (fer-1) was used. Longevity in this strain is the same as the C. elegans wild-type strain N2. Data are recorded in a flow-through respirometry chamber with CO2 output recorded with a 5-s sample resolution. For clarity data are plotted from 30-min intervals. The relative metabolic rate of an age-matched cohort of fed worms remains constant or increases slightly over this time period. The graph plots the means and standard errors from four groups of worms.

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It is less clear what factors are responsible for the large range of metabolic rates reported by the same investigators under seemingly similar measurement conditions. Also puzzling are data from Braeckman et al. (2002) that measured metabolic rates using both microcalorimetric and oxygen consumption techniques. These data indicate that the energy value of the oxygen consumed is more than two-fold higher that the measured heat flux. Such values result in a calorimetric–respirometric ratio well under −470 kJ mol−1 O2, a ratio difficult to explain with conventional biochemical reactions, particularly since the caloric equivalent of oxygen is very constant across both taxonomic level and reaction conditions (Gnaiger & Kemp, 1990).

Indirect measures of C. elegans metabolism

  1. Top of page
  2. Summary
  3. Introduction
  4. Life history characteristics of C. elegans
  5. Growth of C. elegans in the laboratory
  6. C. elegans metabolic measurements
  7. Indirect measures of C. elegans metabolism
  8. Long-lived C. elegans mutants and aging
  9. Comparisons of metabolic rate between wild-type and long-lived mutants
  10. Factors potentially complicating metabolic measurements in C. elegans
  11. Conclusion
  12. Acknowledgments
  13. References

Less direct estimates of metabolism in C. elegans include measures of ATP concentration, superoxide production and reducing equivalents in freeze-thawed worms. Such indirect measures of metabolism suffer from the disadvantage of extrapolating from values measured on dead animals at a single time point, to those expected in the living organism. It has been proposed that ATP levels provide an estimate of the amount of energy available to C. elegans (Braeckman et al., 1999, 2002). For these assays large numbers of C. elegans are grown in liquid culture, collected and usually frozen. ATP levels are determined on the thawed samples with a luciferin/luciferase assay. Studies using this method concluded that long-lived clk-1 mutants have at least the same energy available to them as wild-type worms (Braeckman et al., 1999, 2002). There are two basic problems with this assay, the first conceptual and the second methodological. Conceptually, the use of ATP levels as a measure of metabolic activity assumes that the ATP content of an organism accurately reflects its metabolism. It is a well known biological principle that ATP is not stored by an organism but is rapidly turned-over. Because of this, ATP is linked to metabolic rate through the rate of ATP/ADP turnover, rather than ATP concentration per se. ATP levels and metabolic rate are not generally correlated and ATP levels remain relatively constant across taxa ranging from bacteria to humans, despite enormous differences in metabolic rate (Atkinson, 1977). The lack of correlation between ATP levels and metabolic rate is also seen within species. For example, ATP levels in the killifish (Austrofundulus limnaeus) can remain relatively stable even over 10-fold variations in metabolic rate (Podrabsky & Hand, 1999). The lack of a correlation between ATP pool and metabolic rate limits the usefulness of ATP concentration to estimate energy use in C. elegans.

Methodologically, the rapid turnover rate of ATP requires careful analytical techniques for its accurate measurement. Any protocol used to measure ATP levels must quickly kill the organism, and simultaneously inactivate enzymes that catalyse reactions in which ATP is consumed, produced or interconverted. Determinations of ATP levels which do not carefully control for these factors have been called ‘worse than useless because they are actively misleading’ (Atkinson, 1977). Accepted methods for measuring ATP concentrations follow protocols that flash freeze the sample in liquid nitrogen and then extract ATP in the presence of ATPase-inactivating compounds (Podrabsky & Hand, 1999). In contrast, protocols relying only on sample boiling to inactivate ATPase (Braeckman et al., 1999, 2002) are likely to be inaccurate because significant amounts of ATP can hydrolyse before heat inactivation of ATPases occurs (Vinogradov, 2000). Such ATP degradation may account for reports showing approximately 30-fold decreases in ATP levels between young and old C. elegans, while oxygen consumption rates decreased only 2- to 4-fold over the same period. This large decline in ATP levels also contrasts with studies showing that organisms usually maintain relatively constant ATP levels (Atkinson, 1977), and observations that relatively small reductions (25%) in ATP levels induce apoptotic cell death (Lieberthal et al., 1998). While ATP/ADP ratios have been used as an indicator of the balance between energy-forming and energy-utilizing processes in a cell (Atkinson, 1980), their value as a measure of the energy availability in C. elegans is highly questionable.

Another indirect assay of metabolism measures lucigenin-mediated light production in freeze-thawed C. elegans. This luminescence, assumed to correlate with superoxide formation in the mitochondria during oxidative phosphorylation, is proposed to be a reliable estimate of the potential for metabolic activity (Braeckman et al., 2002). A limitation of this method is that superoxide production by mitochondria in respiration is dependent on the integrity of the inner mitochondrial membrane. This membrane integrity is lost in freeze-thawed mitochondrial samples. Compromises in membrane integrity may be responsible for the high levels of variation reported for this method. As seen in Table 2, light production values from the same laboratory measured on wild-type C. elegans of similar age varied up to 275-fold. While some of these differences are due to modifications in the assay conditions (Vanfleteren et al., 1998), there are also large differences in rate of chemiluminescence decline in aging worms. Some studies found a 2- to 3-fold decrease in light production levels between young adult and middle-aged adult C. elegans (Braeckman, 2002) while other studies of worms in the same age groups showed up to a 40-fold decline in light production levels (Braeckman et al., 1999). The light production levels also appear to typically decline far more quickly with worm age than measures of oxygen consumption. For example, oxygen consumption rates decreased 2- to 4-fold between 4 and 10 days of age in C. elegans, while light production levels decreased up to 40-fold over the same period (Table 2). These data imply that the production rate of superoxides from oxidative phosphorylation is greatly reduced as C. elegans age. Such results are difficult to reconcile with known mechanisms of superoxide formation mediated by oxidative phosphorylation.

Table 2.  Comparison of chemiluminescence values and oxygen consumption values reported for wild-type C. elegans measured at approximately the same two ages. Chemiluminescence units have been standardized to 104 counts (min.mg protein), while oxygen consumption units have been standardized to µg O2 (min.mg protein). Oxygen consumption data from Braeckman et al. (1999) have been corrected from mg O2 to µg O2 (Vanfleteren personal communication)
SourceChemiluminescenceOxygen consumption
Worm age (days)Relative changeWorm age (days)Relative change
410410
Vanfleteren (1994)   50  2.520-foldNDND 
Vanfleteren & De Vreese (1995)   50  2.520-foldNDND 
Vanfleteren & De Vreese (1996)   47  1.434-fold37191.9-fold
Vanfleteren et al. (1998)  5301803-foldNDND 
Braeckman et al. (1999)1300032540-fold50143.6-fold
Braeckman et al. (2000)  200 258-fold26132.0-fold
Braeckman (2002)  2801003-fold29132.2-fold

Another concern with the use of the lucigenin-mediated light production assay is that lucigenin can directly react with oxygen to produce light even in the absence of superoxide (Liochev & Fridovich, 1997; Liochev & I., 1998). Because of this auto-oxidization reaction it was recommended that lucigenin-mediated luminescence should not be used to quantify, or even detect, superoxide levels (Liochev & Fridovich, 1997; Liochev & I., 1998). While Braeckman et al. (2002) reported that auto-oxidization of lucigenin appears minimal under certain conditions, it is not clear what levels of auto-oxidized light production would occur under the conditions used when measuring luminescence in C. elegans samples. In summary, the lucigenin-mediated light production assay used by Braeckman et al. lacks both a theoretical basis and a means of validation, and should not be used for metabolic comparisons in C. elegans.

Long-lived C. elegans mutants and aging

  1. Top of page
  2. Summary
  3. Introduction
  4. Life history characteristics of C. elegans
  5. Growth of C. elegans in the laboratory
  6. C. elegans metabolic measurements
  7. Indirect measures of C. elegans metabolism
  8. Long-lived C. elegans mutants and aging
  9. Comparisons of metabolic rate between wild-type and long-lived mutants
  10. Factors potentially complicating metabolic measurements in C. elegans
  11. Conclusion
  12. Acknowledgments
  13. References

While accurately determining the metabolic rate of C. elegans is essential, an issue of broader interest regards the metabolic rate of long-lived mutants vs. wild-type worms. Long-lived mutants with normal metabolic rates are far more likely to reveal basic mechanisms specifically modulating aging than mutants with reduced metabolic rates. Studies of the factors responsible for extending longevity in C. elegans must consider both chronological life span and physiological life span, the number of physiological events carried out over a lifetime (Lindstedt & Calder, 1981; Strauss, 1999) when evaluating the mechanism(s) by which longevity is extended. An example of the importance of this is seen in mutations that disrupt mitochondrial respiratory chain function in C. elegans. While the chronological longevity of these mutants is extended, the mutants never develop into adults and are smaller than wild-type worms of the same age and developmental state (Tsang et al., 2001). As such, the extended longevity of these mutants is unlikely to reveal much about aging. Consistent with this view, after completing the first genetic screen for long-lived C. elegans mutants, Klass et al. (1983) concluded that the extended longevity of the mutants was due to the mutants either spontaneously forming dauer larva or having a reduced feeding ability. Thus, these mutants were unlikely to provide much insight into novel mechanisms of reducing aging.

Two main classes of long-lived mutants have been used for C. elegans metabolic and aging studies; Clk mutants, and mutants that affect dauer formation. Clk mutants were originally identified in genetic screens for mutations affecting worm development and behaviour. Compared to wild-type, Clk mutants have a slower development, reduced rate of egg production, reduced progeny production and reduced feeding rate (Wong et al., 1995; Lakowski & Hekimi, 1996; Branicky et al., 2000). By themselves, Clk mutations have a relatively modest and variable effect on longevity. On solid medium, average adult longevity of clk-1 mutants is the same as (Larsen & Clarke, 2002; Thaden & Shmookler Reis, 2000) or extended up to 40% relative to wild-type C. elegans (Branicky et al., 2000). These phenotypes would be expected from a mutation that reduces metabolic rate and is consistent with the molecular characterization of clk-1. clk-1 encodes a polypeptide required for the synthesis of ubiquinone or coenzyme Q (CoQ), an essential component of the mitochondrial electron transfer chain (Vajo et al., 1999; Miyadera et al., 2001). Because CoQ is required for typical aerobic respiration, it was originally proposed that either the clk-1 gene was not critically involved in CoQ biosynthesis (Branicky et al., 2000) or clk-1 mutants used alternative electron carriers (Miyadera et al., 2001). Jonassen et al. (2001) provided a simpler explanation by demonstrating that clk-1 mutants obtain CoQ from ingested E. coli. Therefore, it appears that clk-1 mutants essentially use the same method of electron transport as used in wild-type C. elegans. Levels of CoQ also appear to be important in wild-type C. elegans longevity because C. elegans fed on E. coli lacking CoQ live longer than when feeding on CoQ-producing E. coli (Larsen & Clarke, 2002). The increased longevity of C. elegans fed on CoQ-lacking E. coli could potentially be mediated by a reduced metabolic rate in this group compared to worms fed on CoQ-producing E. coli.

A second class of mutations extending longevity in C. elegans affect dauer formation (Daf mutants). Most long-lived Daf mutants were originally identified in genetic screens for mutations affecting dauer formation, rather than for mutations that increase longevity. In addition to their increased longevity, long-lived Daf mutants form dauer larvae under conditions in which wild-type C. elegans will develop to adulthood. The extended longevity of Daf mutants may be mediated through the effects of these mutations on insulin-like signalling pathway(s) (Ogg & Ruvkun, 1998; Guarente & Kenyon, 2000). Many aspects of the dauer larvae appear to be present in long-lived Daf mutants, including patterns of gene expression, physiological and metabolic activity levels, and increased resistance to stress (Guarente & Kenyon, 2000; Vanfleteren & De Vreese, 1995; Jones et al., 2001). In addition to their extended longevity, long-lived Daf mutants have other altered phenotypes that can include slower developmental and growth rates, reduced fertility and a high rate of death from internal egg hatching (Gems et al., 1998).

Comparisons of metabolic rate between wild-type and long-lived mutants

  1. Top of page
  2. Summary
  3. Introduction
  4. Life history characteristics of C. elegans
  5. Growth of C. elegans in the laboratory
  6. C. elegans metabolic measurements
  7. Indirect measures of C. elegans metabolism
  8. Long-lived C. elegans mutants and aging
  9. Comparisons of metabolic rate between wild-type and long-lived mutants
  10. Factors potentially complicating metabolic measurements in C. elegans
  11. Conclusion
  12. Acknowledgments
  13. References

Considering the importance of metabolic rate to C. elegans longevity, surprisingly few studies have measured the metabolic rates in long-lived mutants. Studies from Vanfleteren's laboratory using measures of oxygen consumption, ATP levels and lucigenin-mediated light production concluded that the metabolic rates of long-lived Clk and Daf mutants, and wild-type C. elegans are essentially the same (Vanfleteren & De Vreese, 1996; Braeckman et al., 1999, 2002). This contrasts with studies by Van Voorhies & Ward (1999) measuring C. elegans CO2 production that concluded that the metabolic rates of long-lived mutants were typically reduced compared to wild-type. It has also been reported that ammonium-nitrogen excretion is reduced by a factor of 2–3 in long-lived mutants relative to wild-type (Cherkasova et al., 2000; Thaden & Shmookler Reis, 2000), although the use of ammonium–nitrogen excretion as a measure of metabolic rate must be viewed with caution. The most likely explanation for these different conclusions relates to the conditions used when measuring metabolic rates.

Studies of the importance of metabolic rate to C. elegans longevity require conducting metabolic and longevity assays under optimized environmental conditions to both accurately measure standard metabolic rate and to allow the animal to develop a normal aging phenotype. The optimal environment to grow and maintain C. elegans can be debated, but a practical definition is the maintenance of growth conditions under which fecundity is maximized. In C. elegans this entails growing worms between 15 and 24 °C, on solid medium, with an abundance of E. coli as a food source. The stresses that C. elegans is subjected to in liquid medium make it difficult to interpret metabolic or longevity measurements since the measurements are done under suboptimal conditions. Because C. elegans rapidly reduces its metabolic rate under stressful conditions, measurements done in such environments are not likely to accurately represent the metabolic rate of worms in optimized conditions. An example of the sensitivity of C. elegans metabolic rate to environmental conditions is seen in Fig. 1, which shows the effect of food availability on metabolic rate. The metabolic rate of actively feeding worms placed on media devoid of food is halved within 5 h, and drops approximately 4-fold relative to fed worms after 24 h of starvation. This decline in metabolic rate is not due to starvation-induced mortality as adult C. elegans survive up to 4 days of starvation with > 96% survivorship.

Wild-type worms with potentially high metabolic rates are most likely sensitive to environmental factors which reduce metabolic function, while mutants whose metabolic rate is already depressed will be less affected. This may be responsible for the similar metabolic rates reported by some investigators for wild-type and long-lived mutants. In these cases the strains were typically maintained in liquid medium, and metabolic rates were measured on unfed worms, both conditions expected to depress metabolic rates, especially in wild-type C. elegans.

Factors potentially complicating metabolic measurements in C. elegans

  1. Top of page
  2. Summary
  3. Introduction
  4. Life history characteristics of C. elegans
  5. Growth of C. elegans in the laboratory
  6. C. elegans metabolic measurements
  7. Indirect measures of C. elegans metabolism
  8. Long-lived C. elegans mutants and aging
  9. Comparisons of metabolic rate between wild-type and long-lived mutants
  10. Factors potentially complicating metabolic measurements in C. elegans
  11. Conclusion
  12. Acknowledgments
  13. References

Several important factors proposed potentially to affect metabolic comparisons between long-lived and wild-type C. elegans include embryonic metabolism, body size and alternative metabolic pathways (Braeckman et al., 2002). None of these factors, however, appears to be responsible for the relatively higher metabolic rates reported for wild-type C. elegans (Van Voorhies & Ward, 1999).

Embryonic metabolism Metabolic measurements of C. elegans may include egg-laying adults. These metabolic measurements could be biased if the embryo developing in the egg had a high metabolic rate. Two factors minimizing the influence of this are the reproductive period of C. elegans, and the metabolic rate of a developing embryo. At 20 °C a C. elegans hermaphrodite lays eggs for only 3 days, with the rate of laying decreasing sharply as the worm ages. A 5-day-old worm lays < 1 egg per hour, and egg laying essentially ceases in a 6-day-old worm (Byerly et al., 1976; Hirsh et al., 1976), so developing embryos will not significantly contribute to the metabolic measurements of older worm populations. The contribution of embryonic metabolism to metabolic measurements is also minimized because developing embryos have both low metabolic rates and a small size compared to adult worms (Hirsh et al., 1976; De Cuyper & Vanfleteren, 1982). Thus, eggs laid by worms over a metabolic measurement period will have a relatively minor effect on the metabolic measurement.

The minor metabolic contribution of eggs to adult metabolic measurements indicates that pharmacologically blocking reproduction in C. elegans by inhibiting DNA synthesis with fluorodeoxyuridine (FUdR) is not required to measure adult metabolic rates accurately. Some studies of C. elegans metabolic rate include FUdR in the medium to maintain age-synchronized populations and avoid embryonic metabolism (Braeckman et al., 1999, 2002). FUdR can induce a variety of side-effects that include reducing C. elegans egg production, movement and pharyngeal pumping rates, and cause abnormalities in movement patterns, body size and shape, and internal morphology, and increase superoxide dismutase levels and worm longevity (Mitchell et al., 1979; Gandhi et al., 1980; Bolanowski et al., 1981; Vanfleteren & De Vreese, 1995). The many potential side-effects of FUdR in C. elegans makes it difficult to make informative metabolic measurements on FUdR treated worms.

Body size differences A difficulty that is encountered when biological traits are compared between organisms is differences in body mass. In general, the effects of body size on metabolic rate and aging require more careful consideration than is typically given. For example, within a species, smaller individuals tend to live longer, and long-lived mutants are often smaller than wild-type animals (Bartke, 2000) although the reasons for this are not known. Consistent with this trend many long-lived C. elegans mutants are smaller than wild-type worms. The supposition, however, that the reduced metabolic rate reported for some long-lived C. elegans mutants is due only to the reduced size of those mutants is incorrect. Van Voorhies & Ward (1999) controlled for this effect by comparing metabolic rates of worms between these groups at two points when they were of equivalent size and developmental stage. When compared in this manner, wild-type worms had metabolic rates averaging 2.5 times higher than long-lived mutants.

Investigators often attempt to control for body size effects by expressing the result in a mass-specific unit, such as metabolic rate per unit body mass or unit protein content (McNab, 1999). There are several potential problems with expressing data in such a manner. Expressing data in mass-specific units assumes that the relationship between the dependent variable being adjusted and the mass unit is a simple linear function with an intercept of 0 (Carpenter et al., 1995; Weathers & Siegel, 1995). Metabolic rate data expressed in mass-specific units typically assume that body mass and metabolic rate have a scaling exponent (b) of 1.0, when the relationship between metabolic rate (Y) and body mass (M) is expressed in an allometric relationship of Y = aMb (a is a mass coefficient) (Heusner, 1982; Calder, 1984; Schmidt-Nielsen, 1984).

Such assumptions on the relationship between body mass and metabolic rate are often invalid (Chappell & Bachman, 1995; Hayssen & Lacy, 1985; Hack, 1997; Degen et al., 1998), and expressing data in mass-specific units can result in erroneous conclusions. For example, in nematodes the relationship between body mass and metabolic rate can be bi- or triphasic and the scaling exponent for metabolic rate can range from 0.33 to 0.75 (Atkinson, 1980; De Cuyper & Vanfleteren, 1982). Extrapolating from these exponents predicts that doubling the size of a nematode would increase its metabolic rate by a factor of 1.26–1.68 rather than 2.0 as assumed if C. elegans metabolic rates are only plotted in mass-specific units. Controlling for the effects of body mass on a trait requires the careful determination of the appropriate method with respect to the data set being analysed and the use of appropriate statistical methods (Packard & Boardman, 1999). Expressing metabolic rate only as body mass ratio seldom removes the effects of body size on the variable of interest and statistical analysis of such data is commonly biased and unreliable; thus, expressing data in such terms can hinder rather than help with understanding the complex factors affecting an organism's metabolic rate (Packard & Boardman, 1988, 1999; Hayes, 2001). While there is no universal method for correcting for differences in body mass, at a minimum the whole animal body size and the number of animals used to calculate the metabolic rates must be reported.

Alternative metabolic pathways In addition to oxidative phosphorylation, C. elegans can potentially produce CO2 from metabolic pathways such as the glyoxylate cycle (Liu et al., 1997) or pentose phosphate shunt. The glyoxylate cycle converts lipids to gluconeogenic precursors that can be used to form carbohydrates or for anaplerotic reactions to form intermediates used in replenishing metabolic pathways. C. elegans most likely uses this pathway to synthesize carbohydrates from stored lipids for biosynthetic purposes rather than the conversion of lipids to carbohydrates for subsequent use in the Krebs cycle (Bolla, 1980). A much greater use of these metabolic pathways by long-lived mutants compared to wild-type worms could potentially complicate the interpretation of CO2-based metabolic measurements. The impact, however, of the glyoxylate cycle or pentose phosphate shunt to C. elegans CO2 production appears to be minor in well-fed C. elegans. A diet of E. coli already supplies large amounts of carbohydrates (Ingraham et al., 1983), minimizing the need for feeding C. elegans to use the glyoxylate cycle to obtain carbohydrates. Also, the utilization of the glyoxylate cycle in C. elegans, even in dauer larvae, appears limited (Wadsworth & Riddle, 1989; O’Riordan & Burnell, 1990; but see Vanfleteren & De Vreese, 1995) and is reduced in adult C. elegans compared to larval worms (Liu et al., 1997). Additionally the respiratory quotient of C. elegans embryos, a developmental stage at which the use of the glyoxylate cycle should be maximized, is near 0.70 (Fig. 2), indicating that only lipid substrates are being metabolically utilized and the pentose phosphate shunt and the glyoxylate cycle are not a significant source of CO2 production. Figure 2 also shows that while the respiratory quotient of C. elegans varies with larval development, the variation is relatively modest. This range of respiratory quotients is consistent with C. elegans predominately using oxidative phosphorylation-based metabolism. This indicates that measures of CO2 output can be used accurately to infer oxygen consumption rates in C. elegans. Finally, this figure shows that the respiratory quotient of a long-lived C. elegans mutant is essentially the same as that of normal lived worms, demonstrating that the ratio of CO2 production to oxygen consumption is very similar between wild-type and at least some long-lived C. elegans mutants. As such, CO2 production provides an accurate and robust method of comparing metabolic rates between these groups.

image

Figure 2. The respiratory quotient in C. elegans during development and compared between a normal-lived and long-lived mutant. RQ was measured in a long-lived daf-2 mutant (e1368) which lives approximately twice as long as wild-type. Worms were grown at 23 °C for all experiments. At this temperature eggs develop into adults in around 2.5 days. (•) fer-1 worms; (▵) daf-2 worms. The graph plots the means and standard errors derived from four groups of worms for RQ values measured during development. Data from three groups of long-lived mutant worms and two groups of wild-type worms are plotted for 4–5-day-old worms. Each group of worms comprised hundreds of individual worms.

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Conclusion

  1. Top of page
  2. Summary
  3. Introduction
  4. Life history characteristics of C. elegans
  5. Growth of C. elegans in the laboratory
  6. C. elegans metabolic measurements
  7. Indirect measures of C. elegans metabolism
  8. Long-lived C. elegans mutants and aging
  9. Comparisons of metabolic rate between wild-type and long-lived mutants
  10. Factors potentially complicating metabolic measurements in C. elegans
  11. Conclusion
  12. Acknowledgments
  13. References

Interpreting and understanding the mechanism(s) by which mutations can increase C. elegans longevity requires the careful assessment of metabolic function in these mutants. The extended longevity of at least some long-lived C. elegans mutants appears to be due to reductions in metabolic rate. Studies correlating the metabolic rate of an organism to its longevity should measure metabolic rates under conditions that closely match the conditions used to assay longevity. Accurate metabolic measurements require careful attention to both conditions under which the organism's metabolism is measured and the use of appropriate techniques to measure metabolic rate. Most of the commonly studied long-lived C. elegans mutants typically live around twice as long as wild-type worms (Gems et al., 1998; Johnson et al., 2000). If there is an approximately linear relationship between a biochemical parameter, such as free radical generation and aging, the difference in this parameter between long-lived mutants and wild-type would also be around two-fold. This indicates that measurement methods used to assay for physiological or biochemical differences between these groups should have the accuracy and sensitivity to reproducibly distinguish relatively small differences in a trait. As currently used, indirect measures of metabolic rate or capacity, such as ATP levels or superoxide production, are inadequate for assaying metabolism in C. elegans.

It is also important to realize that many factors besides metabolic rate affect longevity. Therefore, mutations that reduce metabolic rate will not always increase longevity, suggesting that a fine line exists between compromising metabolic function and extending longevity. Many features of C. elegans make it a valuable model organism to study aging. However, studies of C. elegans and aging must assay both chronological life span and physiological life span to provide true insight into the aging process. This same criterion applies to other studies of genetic mutations or methods that increase longevity. Methods that increase longevity in any organism need also to demonstrate that other aspects of the organism's physiology, such as growth and metabolic rates, remain normal, when ascertaining the significance of the means by which longevity is increased.

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  1. Top of page
  2. Summary
  3. Introduction
  4. Life history characteristics of C. elegans
  5. Growth of C. elegans in the laboratory
  6. C. elegans metabolic measurements
  7. Indirect measures of C. elegans metabolism
  8. Long-lived C. elegans mutants and aging
  9. Comparisons of metabolic rate between wild-type and long-lived mutants
  10. Factors potentially complicating metabolic measurements in C. elegans
  11. Conclusion
  12. Acknowledgments
  13. References
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