ToxSci Advance Access originally published online on November 12, 2007
Toxicological Sciences 2008 102(1):179-186; doi:10.1093/toxsci/kfm278
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Mitochondrial Modulation of Phosphine Toxicity and Resistance in Caenorhabditis elegans
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* School of Molecular and Microbial Sciences, University of Queensland, St Lucia, Queensland 4072, Australia
School of Integrative Biology, University of Queensland, St Lucia 4072, Brisbane, Australia
1 To whom correspondence should be addressed at Goddard Building, School of Integrative Biology, University of Queensland, St Lucia, Queensland 4072, Australia. Fax: +61-7336-51655. E-mail: p.ebert{at}uq.edu.au.
Received September 7, 2007; accepted November 6, 2007
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONFUNDINGREFERENCES |
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Phosphine is a fumigant used to protect stored commodities from infestation by pest insects, though high-level phosphine resistance in many insect species threatens the continued use of the fumigant. The mechanisms of toxicity and resistance are not clearly understood. In this study, the model organism, Caenorhabditis elegans, was employed to investigate the effects of phosphine on its proposed in vivo target, the mitochondrion. We found that phosphine rapidly perturbs mitochondrial morphology, inhibits oxidative respiration by 70%, and causes a severe drop in mitochondrial membrane potential (
m) within 5 h of exposure. We then examined the phosphine-resistant strain of nematode, pre-33, to determine whether resistance was associated with any changes to mitochondrial physiology. Oxygen consumption was reduced by 70% in these mutant animals, which also had more mitochondrial genome copies than wild-type animals, a common response to reduced metabolic capacity. The mutant also had an unexpected increase in the basal m, which protected individuals from collapse of the membrane potential following phosphine treatment. We tested whether directly manipulating mitochondrial function could influence sensitivity toward phosphine and found that suppression of mitochondrial respiratory chain genes caused up to 10-fold increase in phosphine resistance. The current study confirms that phosphine targets the mitochondria and also indicates that direct alteration of mitochondrial function may be related to phosphine resistance. Key Words: phosphine; mitochondria; Caenorhabditis elegans; pesticide; fumigant.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONFUNDINGREFERENCES |
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Phosphine (hydrogen phosphide, PH3) is a fumigant that is used worldwide to protect stored commodities against pest insects and rodents (reviewed in Chaudhry, 1997
). Its prominence as a fumigant is due to its low cost, ease of application, lack of residues, and potency. Phosphine also decomposes into environmentally safe compounds and does not deplete the ozone layer, as does the highly restricted fumigant, methyl bromide (MBTOC, 2006; UN, 1991). Unfortunately, insect resistance to phosphine has developed to a level, which threatens its continued use (Athie et al., 1998; Collins et al., 2002).
Although its mode of toxicity is not well understood, phosphine has been found to inhibit aerobic respiration in a number of species and tissues (Chefurka et al., 1976; Dua and Gill, 2004; Jian et al., 2000; Singh et al., 2006). In vitro studies on isolated mitochondria and submitochondrial particles have revealed that cytochrome c oxidase (complex IV) of the electron transport chain is inhibited by phosphine (Chefurka et al., 1976; Nakakita, 1976). However, it is still unclear as to whether this interaction is the primary cause of toxicity as phosphine inhibits complex IV activity less dramatically in vivo than in vitro (Jian et al., 2000; Price, 1980). In insects (Nakakita, 1987), mites (Jian et al., 2000), rats (Dua and Gill, 2004), and humans (Singh et al., 2006), phosphine only partially inhibits complex IV activity in vivo. Thus, phosphine poisoning is different from other complex IV inhibitors such as cyanide, which strongly inhibits complex IV activity in vivo (Price and Walter, 1987). As a result, alternative hypotheses that do not rely on inhibition of aerobic respiration as the mechanism of phosphine toxicity have been developed.
One such proposal is that phosphine is directly involved in the chemical reactions that result in oxidative damage as phosphine and hydrogen peroxide can interact to form the highly reactive hydroxyl radical (Quistad et al., 2000). This is consistent with an observed increase in hydroxyl radical associated damage, such as lipid peroxidation, in vitro (Hsu et al., 1998; Quistad et al., 2000) and in vivo (Hsu et al., 2002a,b; Quistad et al., 2000) in phosphine-treated mammalian cell lines. Additionally, phosphine has been shown to inhibit the antioxidants catalase and peroxidase in a variety of insects (Chaudhry, 1997), cause hypersensitivity to a glutathione depleting compound in Caenorhabditis elegans (Valmas and Ebert, 2006) and mammals (Hsu et al., 2002b), as well as hypersensitivity toward free iron (Cha'on et al., 2007). These observations are consistent with an oxidative stress model of phosphine toxicity, and because mitochondrial activity is intimately linked with oxyradical production, the mitochondrial mode of action of phosphine may also be supported by these observations.
The mechanism of resistance to phosphine is even less well understood than the mechanism of its toxicity. Resistance is not due to alterations in the respiratory chain that prevent phosphine from inhibiting complex IV in vitro and presumably in vivo (Chaudhry, 1997; Price, 1980). A recent study has also shown that carbon dioxide production and phosphine resistance levels are correlated in field collected insect strains, indicating a direct relationship between metabolic rate and phosphine sensitivity (Pimental et al., 2007).
Phosphine is toxic toward all actively aerobically respiring organisms. In this report, we chose to examine the toxic effects of phosphine in vivo in the model organism C. elegans. This nematode is ideal for studying fumigant toxicology, particularly because of its tremendous reproductive capacity and rapid life cycle, which facilitates toxin screening. Genetic resources (http://www.wormbase.org/) are readily available and the small size and transparency of these nematodes allows direct in situ visualization and analysis of mitochondria, which in the case of phosphine toxicity is of utmost importance. Phosphine was found to alter mitochondrial morphology, strongly inhibit aerobic respiration, and severely decrease the mitochondrial membrane potential (m) of living nematodes, indicating a direct inhibition of mitochondrial function. Interestingly, the pre-33 mutant, a phosphine-resistant line of C. elegans, had more copies of the mitochondrial genome than normal and a constitutively higher m but also had a lower respiratory rate. Furthermore, experimental suppression of mitochondrial respiration resulted in phosphine resistance. Therefore, our results indicate that phosphine directly targets mitochondria in vivo and that resistance can be achieved by modulating respiratory activity.
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MATERIALS AND METHODS |
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Nematode strains.
The wild-type C. elegans, N2 (var. Bristol), used for these studies was kindly provided by Dr Warwick Grant, AgResearch, New Zealand. GFP (green fluorescence protein) reporter strain CB5600, which contains a myo-3::Ngfp-lacZ;myo-3::Mtgfp transgene (Labrousse et al., 1999
), was obtained from the Caenorhabditis Genetics Center. The phosphine-resistant strain, pre-33, was previously created using EMS mutagenesis in an N2 background (Cheng et al., 2003). The nematodes were cultured and exposed to phosphine on nematode growth medium (NGM) at 20°C unless otherwise stated (Brenner, 1974) and were fed the attenuated Escherichia coli OP50 strain in all but the RNAi experiments.
Phosphine treatment.
Phosphine generation and exposure was performed as previously described (Valmas and Ebert, 2006). Briefly, nematodes on NGM agar plates were placed into airtight desiccators and sealed. A measured amount of phosphine gas was then injected into these desiccators to obtain the desired concentration. Nematode development was synchronized (Cheng et al., 2003) prior to phosphine exposure by allowing eggs to hatch in the absence of food. For mortality assays, young adult nematodes were used that had been allowed to develop for 48 h after reinitiation of feeding. After exposure, the nematodes were removed from the desiccators and allowed to fully recover in fresh air for up to 48 h before mortality was assessed. Individuals were counted as dead if they did not initiate movement in response to submersion in M9 buffer (Cheng et al., 2003).
Confocal microscopy of mitochondria.
To obtain large, postreproductive nematodes for microscopy, CB5600 nematodes were allowed to develop for 135-h postsynchronization prior to phosphine exposure. Older and larger nematodes were used for this experiment as young adult nematodes proved to be too small to obtain detailed images of mitochondria in situ. These older nematodes have been found to have equal tolerance to phosphine as young adult nematodes (unpublished observations). A phosphine concentration of 215 ppm at 25°C was selected as it represents the LC20 at 24 h. After exposures, which lasted for 1, 5, 10, and 24 h, the nematodes were immediately harvested and paralyzed with M9 buffer containing 0.1% tricaine (Sigma, St Louis, MO) and 0.01% tetramisole (Sigma). The live but paralyzed nematodes were then mounted under a cover slip on a 2% agarose pad (Wallenfang and Seydoux, 2000). Microscopic analysis of the GFP nematodes was performed with a Nikon Eclipse E600 and photographs were taken using a BIO-RAD Radiance 2000 laser scanning system camera. BIO-RAD LaserSharp 2000 software (version 4.1): Hercules, CA was used to control image acquisition. All photographs were captured using the same gain and exposure settings.
Oxygen consumption rate.
Synchronized populations of young adult nematodes were exposed to phosphine (70 ppm, standard sublethal concentration) for 1 or 5 h before being immediately washed in S-basal buffer and collected. We monitored oxygen consumption rates using a mitocell ST200 chamber (Strathkelvin Instruments, Motherwell, UK). Measurements were performed at 20°C and continued for as long as necessary to accurately obtain a linear rate of oxygen consumption (usually 5 min). Oxygen consumption rates were normalized to either protein content of matched sample aliquots (Bradford protein assay, Sigma, St Louis, MO), or to the approximate number of animals used in the assay.
Quantification of mitochondrial DNA copy number using real-time PCR.
Quantitative real-time PCR was used to determine the relative copy number of mitochondrial DNA (mtDNA) to nuclear DNA. Total cellular DNA was extracted from nematodes and amplified with primers specific to either mtDNA (locus: 6.254 kb; F: TTATCTACGGGATTTCACGGAATT, R: TCCAACCCCAGATGATGATTATAAT) or nuclear DNA (locus: F53G12; F: CGGTGAGCTCCTCCAAATGT, R: CCTCTGCATCAACAAGCTCG). Quantitative PCR was carried out on the DNA template using an ABI Model 7000 Sequence Detector (Applied Biosystems, Foster City, CA). PCR reactions (25 µl) contained 0.1µM of each primer; 1 ng of template DNA, and 12.5 µl of SYBR Green PCR Master Mix (Applied Biosystems). The standard SYBR Green PCR program of 10 min at 95°C, and 40 cycles of 15 s at 95°C, 20 s at 60°C, and 20 s at 72°C was used. DNA was extracted from N2 and pre-33 nematodes that were 1, 5, 10, and 24 h past the young adulthood stage of their development (48 h old). These times were used to match the age of the nematodes in the other mitochondrial physiology experiments.
Analysis of mitochondrial membrane potential (m).
Mitochondrial membrane potential was measured as relative uptake of the fluorescent dye tetramethylrhodamine ethyl ester (TMRE; Sigma) into the mitochondrial matrix of whole live nematodes. This lipophilic cationic dye accumulates in mitochondria in proportion to the strength of the m (Ehrenberg et al., 1988) and has been used previously with C. elegans (Yoneda et al., 2004). Synchronized animals were grown to young adulthood and exposed to 350 ppm phosphine for 1, 5, 10, or 24 h. A phosphine concentration of 350 ppm at 20°C was selected as it represents the LC20 at 24 h. However, the nematodes were first transferred to NGM plates containing 0.1µM of TMRE so that in all cases nematodes were exposed to TMRE for a total of 24 h (including the phosphine exposure time). After phosphine treatment, the animals were prepared for fluorescence microscopy as described for confocal microscopy. Live animals were then photographed using a Nikon fluorescence microscope at 10x magnification. All fluorescent images were taken using the same gain and exposure settings and the amount of dye accumulated in the mitochondrial matrix was quantified using Adobe Photoshop CS2 (Kirkeby and Thomsen, 2005) to reveal the mean red channel intensity of individual nematodes from three independent experiments (n = 9 for each treatment).
RNA interference of mitochondrial genes.
RNA interference (RNAi) clones in the RNase III mutant E. coli host strain, HT115, were obtained from MRC GeneService (Cambridge, UK) and sequenced to confirm correct genetic inserts. RNAi was performed using the feeding protocol based on the recommendations of MRC GeneService. Briefly, each bacterial strain was cultured for 7 h in Luria Broth containing 100 µg/ml ampicillin at 37°C. The bacteria were then seeded onto NGM agar plates containing 1mM IPTG and 25 µg/ml carbenicillin. double-stranded RNA production was induced by Isopropyl β-D-1-thiogalactopyranoside (IPTG) overnight at 25°C. The following day, synchronized L1 stage nematodes were added to the RNAi plates and allowed to develop to young adulthood for use in subsequent resistance analysis.
Statistical analysis.
Results from Figures 2 to 6 are presented as means with error bars representing the standard errors of the means (SEM). The Student t-test was used to determine the level of significance of differences in respiratory rate, mtDNA content, and phosphine sensitivity in RNAi nematodes. Genstat 7.2 (VSN International, Ltd., Hemel Hempstead, UK) was used for statistical analysis of dose-dependent phosphine mortality curves (Fig. 3). The phosphine-induced mortality of each biological replicate was adjusted using Abbott's formula and then pooled for analysis using linear regression, whereas dose–response data were subject to probit regression. Transformation was performed on the data (complementary log–log on the response variate; and logarithm on the explanatory variate) to approximate the does–response relationship.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONFUNDINGREFERENCES |
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Phosphine Alters Mitochondrial Morphology In Vivo
To observe the effect of phosphine on mitochondria in vivo, we used nematodes expressing a GFP fusion gene (myo-3::GFP) that labels the mitochondrial and nuclear compartments of muscle cells of the body-wall (Figs. 1A and 1B). Mitochondria of nematodes exposed to air alone for up to 24 h appeared as elongated networks of tubules interspersed with large round nuclei (Figs. 1C–F). However, treatment with sublethal doses of phosphine rapidly disrupted these mitochondrial networks and induced the formation of smaller, more circular vesicles from 5 h of treatment onwards (Figs. 1G-J). This difference was especially evident after 24 h of treatment (compare Figs. 1F and 1J). The presence of unaffected nuclei in phosphine treated animals suggests that the effect was specific to mitochondria and not the result of widespread damage to cellular membranes.
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Phosphine Inhibits Respiration in Wild-Type Nematodes
Exposure of wild-type (N2) nematodes to 70 ppm phosphine gas, a dose that is sublethal even for an exposure period of 24 h, drastically inhibited respiration after just 1 h exposure onwards (Figs. 2A and 2B). Interestingly, the phosphine-resistant mutant pre-33 was found to have a constitutively reduced basal rate of respiration under normal conditions. pre-33 nematodes are much more resistant to phosphine than are wild-type nematodes, with LC50 values of 2150 and 600 ppm, respectively, at 20°C (Fig. 3). Phosphine treatment had no effect on the respiration rate of the mutant animals with the air treated and phosphine-treated pre-33 nematodes exhibiting almost identical rates of oxygen consumption (Fig. 2A). A similar effect was seen after 5 h of phosphine exposure (Fig. 2B). Under normal conditions, pre-33 nematodes also had a significantly greater mtDNA content relative to wild-type nematodes (Fig. 4). It is possible that mtDNA replication is upregulated in these mutants in response to their low rates of respiration, as mtDNA copy number is closely linked with energy status in C. elegans (Tsang and Lemire, 2002
).
Phosphine Exposure Lowers Mitochondrial Membrane Potential (m)
Using the fluorescent dye TMRE, which accumulates in intact respiring mitochondria in proportion to the strength of the m (Ehrenberg et al., 1988), we observed greater fluorescence in the phosphine-resistant pre-33 mutant than in wild-type animals (Fig. 5). The m was unaffected by 1-h exposure to phosphine, but thereafter, exposure resulted in an equivalent rate of reduction in m in both pre-33 and N2 nematodes. In N2 animals, it reached a minimum by 5 h phosphine exposure. The lack of a further reduction may have been caused by the exclusion of dead nematodes from the analysis as these individuals exhibited no fluorescence. Despite a similar rate of reduction in the m of both pre-33 and N2 animals under phosphine treatment, the unusually strong m of pre-33 individuals at the start of the experiment ensured that the m was conserved much better in these mutants than in N2 animals (Figs. 5I and 5J). The difference between wild-type and resistant nematodes was most apparent after 5 h of phosphine exposure as live N2 animals exhibited very weak TMRE fluorescence, whereas pre-33 animals experienced a comparatively slight decrease in m.
Suppression of Mitochondrial Gene Expression Causes Phosphine Resistance
The reduced oxygen consumption shown previously for the phosphine-resistant mutant (Fig. 2) presented the intriguing possibility that phosphine resistance results directly from reduced mitochondrial respiration. We tested this by experimentally inhibiting mitochondrial respiratory function by individually silencing 21 mitochondrial respiratory chain genes in nematodes using RNAi, each of which potentially inhibited oxidative respiration (Lee et al., 2003). Following gene suppression, we screened the nematodes for phosphine sensitivity by exposing the animals to 350 and 700 ppm of phosphine (Table 1). The silenced genes encode subunits of the electron transport chain complex I (NADH [nicotinamide adenine dinucleotide phosphate, reduced]/ubiquinone oxidoreductase), complex II (succinate/ubiquinone oxidoreductase), complex III (cytochrome c reductase), and complex IV (cytochrome c oxidase). This screening experiment indicated that silencing of 15 of the 21 genes resulted in a > 20% decrease in phosphine-induced mortality relative to control nematodes, whereas the remainder gave weak or ambiguous results (Table 1). We selected the most resistant lines and performed a more detailed analysis of phosphine sensitivity over a greater range of concentrations (Fig. 6). RNAi of complex III genes conferred the strongest resistance toward phosphine, which was particularly notable when T02H6.11 and ucr-1 were silenced (Figs. 6B and 6C).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONFUNDINGREFERENCES |
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Mitochondria are critical to the survival of all obligatory aerobic eukaryotes as indicated by the extreme toxicity of compounds that disrupt mitochondrial function. Here we have shown that the fumigant phosphine is able to disrupt mitochondrial function in the model organism C. elegans at toxicologically significant concentrations. Phosphine caused rapid inhibition of aerobic respiration (Fig. 2), which is consistent with a conserved mode of action in insects, nematodes, and other eukaryotes (Chefurka et al., 1976
; Dua and Gill, 2004; Jian et al., 2000; Nakakita, 1987; Singh et al., 2006). We also observed specific alterations in mitochondrial morphology (Fig. 1) as well as decreases in m after treatment with phosphine (Fig. 5). Owing to the advantageously small size and transparency of C. elegans, these observations were made in situ in living animals and collectively indicate that phosphine directly and rapidly targets mitochondria in vivo. Because phosphine disrupted the capacity to maintain m and respiration, our results are consistent with the prevailing model of phosphine toxicity that invokes the inhibition of complex IV and the disruption of electron transport (Chefurka et al., 1976; Dua and Gill, 2004; Jian et al., 2000; Nakakita, 1987; Singh et al., 2006).
It is possible that phosphine toxicity is exerted through its ability to reduce the strength of the m. In phosphine-resistant pre-33 mutants, we observed that phosphine treatment did not lower m to the same degree as in sensitive N2 nematodes (Fig. 5). This was because the m of pre-33 animals was initially greater than that of N2 animals (Fig. 5). Limiting the collapse in m may contribute to the phosphine resistance phenotype in pre-33 mutants. Indeed, a prediction resulting from this work is that exacerbation of the decrease in m caused by phosphine would enhance its toxicity.
Other unique features of pre-33 physiology may also have protected the mutants from phosphine toxicity. Metabolic rate appeared to be constitutively reduced in pre-33 mutants as indicated by a markedly lower rate of oxygen consumption than in N2 animals (Fig. 2). Other forms of evidence also confirm a reduction in metabolism in resistant pre-33 nematodes. For example, pre-33 animals have a greater ratio of mtDNA genomes to nuclear genomes (Fig. 4). mtDNA copy number is usually increased in response to heightened cellular energy demand as it allows for the production of more oxidative phosphorylation components (Lopez-Lluch et al., 2006; McCabe et al., 2000; Moraes, 2001). This may reflect an increase in energy demand induced by the lowered respiratory output in these mutants and is consistent with a general decrease in metabolic rate. Furthermore, pre-33 mutants have reduced developmental rates, lowered fecundity, and extended longevity, all of which are consistent with reduced metabolism (Cheng et al., 2003).
Thus, the results reported in this paper strongly support a link between metabolic rate and the degree of sensitivity toward phosphine, which has been previously proposed (Schlipalius et al., 2006). Indeed, factors that alter aerobic metabolic rate are directly correlated with the toxicity of phosphine. For example, near anaerobic conditions render phosphine nontoxic (Kashi, 1981). Similarly, reduced temperatures, which lowers metabolism, reduce sensitivity toward phosphine (Chaudhry, 1997). Furthermore, a number of phosphine resistant strains of beetle were recently found to have a correlated reduction in metabolism as measured by reduced CO2 production (Pimental et al., 2007).
Silencing genes encoding components of the mitochondrial respiratory chain also increased resistance to phosphine (Table 1 and Fig. 6). Suppressing these genes by RNAi has previously been shown to reduce growth rate, adenosine triphosphate (ATP) content, and respiration in C. elegans (Dillin et al., 2002; Lee et al., 2003). Thus, genetically reducing respiration and metabolism by an alternative method also enhances phosphine resistance in C. elegans. The ongoing mapping of the pre-33 mutation suggests that there are no genes encoding respiratory chain subunits within the narrowed chromosomal region of the mutation (C. Cheng, personal communication). So, although similarities may be drawn between the changes in life traits and the reductions in respiratory rates, nematodes treated with respiratory chain RNAi and pre-33 mutants may achieve these metabolic alterations by fundamentally different mechanisms. Nevertheless, it is noteworthy that suppression of mitochondrial respiratory genes leads not only to phosphine resistance, but longevity as well (Dillin et al., 2002; Lee et al., 2003). This parallels the situation with the pre-33 mutant in which a reduction in respiration and metabolism is associated with not only phosphine resistance, but also with greater longevity (Cheng et al., 2003).
It is now clear that multiple factors that influence metabolism, and specifically mitochondrial function, have a direct influence on phosphine toxicity as well as resistance against its toxic effects. Mitochondrial membrane potential, rate of electron flow through the mitochondrial respiratory chain, ATP levels, metabolic supply versus demand, and mitochondrial generated oxidative stress are all metabolic factors that may contribute to phosphine sensitivity or resistance. Biochemical and genetic evidence indicates that multiple resistance mechanisms exist, so it is possible that multiple metabolic factors contribute to resistance in unique ways. This work identifies m and electron flow through the mitochondrial respiratory chain as potential resistance mechanisms that have not previously been described.
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FUNDING |
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Australian Research Council Discovery Project Grant (ARC-DP0558507) to P.E.; and Australian Postgraduate Awards to S.Z. and J.K.
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ACKNOWLEDGMENTS |
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We thank Nick Valmas, David Schlipalius, Andrew Tuck, and Yosep Mau for useful discussion and comments. We also thank Robert Simpson for assistance with quantitative real-time-polymerase chain reaction and Prof. Craig Franklin for assistance with measurement of oxygen consumption rates.
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