Article | Open Access | Published:

Systemic regulation of L-carnitine in nutritional metabolism in zebrafish, Danio rerio

Scientific Reports volume 7, Article number: 40815 (2017) | Download Citation

  • A Corrigendum to this article was published on 23 March 2017

This article has been updated


Excess fat accumulation has been observed widely in farmed fish; therefore, efficient lipid-lowering factors have obtained high attention in the current fish nutrition studies. Dietary L-carnitine can increase fatty acid β-oxidation in mammals, but has produced contradictory results in different fish species. To date, the mechanisms of metabolic regulation of L-carnitine in fish have not been fully determined. The present study used zebrafish to investigate the systemic regulation of nutrient metabolism by dietary L-carnitine supplementation. L-carnitine significantly decreased the lipid content in liver and muscle, accompanied by increased concentrations of total and free carnitine in tissues. Meanwhile, L-carnitine enhanced mitochondrial β-oxidation activities and the expression of carnitine palmitoyltransferase 1 mRNA significantly, whereas it depressed the mRNA expression of adipogenesis-related genes. In addition, L-carnitine caused higher glycogen deposition in the fasting state, and increased and decreased the mRNA expressions of gluconeogenesis-related and glycolysis-related genes, respectively. L-carnitine also increased the hepatic expression of mTOR in the feeding state. Taken together, dietary L-carnitine supplementation decreased lipid deposition by increasing mitochondrial fatty acid β-oxidation, and is likely to promote protein synthesis. However, the L-carnitine-enhanced lipid catabolism would cause a decrease in glucose utilization. Therefore, L-carnitine has comprehensive effects on nutrient metabolism in fish.


L-carnitine is an important factor that regulates fatty acid (FA) and glucose metabolism to achieve balanced cellular energy metabolism1. L-carnitine can be synthesized naturally from lysine and methionine in microorganisms, plants and animals2. L-carnitine plays a vital role in the β-oxidation of long chain fatty acids (LCFAs) to produce energy by transporting LCFAs from the cytosol into the mitochondrial matrix, and can modulate the ratio of acetyl-CoA/CoA3,4. In humans and other mammals, L-carnitine deficiency causes severe hyperlipidaemia and systemic metabolic syndrome5,6. Accordingly, dietary supplementation of L-carnitine had been reported widely to improve lipid metabolism and/or alleviate the high fat diet (HFD)-associated excess lipid accumulation in humans and other land animals7,8,9.

Compared with other farmed animals, fish generally have a lower ability to use carbohydrates as energy sources; thus requiring higher levels of dietary proteins10. With the increasing cost and limited supplies of fish feed protein, HFDs are used currently in aquaculture for their “protein sparing effect”, which means that the energy that is normally sourced from protein consumption is replaced by lipid-sourced energy11,12. However, HFDs often lead to severe lipid accumulation in the liver and abdominal adipose tissues in farmed fish, and induce metabolic disturbances, such as fatty liver and excess mesenteric fat deposition12,13. Therefore, regulatory factors that could reduce lipid deposition in fish, especially during HFD feeding, have attracted increased research attention. The effective lipid-lowering effects of L-carnitine have led to the hypothesis that L-carnitine in fish would have similar effects to those in mammals and would improve lipid utilization.

During recent decades, the potential nutritional functions of carnitine have been studied in many fish species; however, the results and conclusions differed. In European sea bass14, red sea bream15 and hybrid striped bass16, carnitine promoted growth. However, this effect was not observed in channel catfish17, rainbow trout18 and hybrid tilapia19. In European sea bass14, Mozambique tilapia20, Indian major carp rohu21, dietary carnitine supplementation could lower the lipid content in the body and liver, and it improved the utilization of certain FAs, such as eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA) in red sea bream15, and African catfish22. However, carnitine supplementation had no effects on lipid metabolism in African catfish23, hybrid tilapia19 and hybrid striped bass16. Indeed, carnitine even reduced lipid metabolism in red sea bream15, yellow catfish24 and rainbow trout25.

The contradictory and complicated results of carnitine supplementation in fish have made the further utilization of carnitine in fish feed controversial. However, compared with the intensive mechanistic studies of carnitine in mammals, the majority of the studies of carnitine in fish have only focused on the effects of carnitine on the fish’s growth and body composition, whereas studies on the systemic regulation of nutritional metabolism by carnitine in fish, and the underlying mechanisms, are lacking. This has largely restricted our understanding of the function of carnitine in fish, and an explanation of the conflicting results of the application of carnitine in fish remains elusive. Therefore, the mechanisms of the effects of carnitine in fish should be studied.

As a widespread model animal in research areas such as development, molecular genetics, toxicology and biomedicine26,27, zebrafish (Danio rerio) has been used in mechanistic studies of fish nutrition28,29. In the present study, zebrafish was used to evaluate the systemic regulation of lipid, protein and carbohydrate metabolism by L-carnitine in fish in feeding and fasting states, respectively. Moreover, the metabolic pathways associated with the nutritional regulation by dietary L-carnitine supplementation in zebrafish were determined. To the best of our knowledge, this is the first study to demonstrate the metabolic mechanisms of nutritional regulation by dietary L-carnitine supplementation in fish.


The effect of dietary L-carnitine supplementation on the carnitine concentration, carnitine synthesis gene expression and triglyceride levels in tissues

During the experiment, the fish were in good health and showed no growth differences compared with control fish (data not shown). To test the effects of exogenous carnitine supplementation on the endogenous carnitine concentrations, the carnitine concentrations in various fish tissues were measured. In the liver and muscle, both in the feeding and fasting states, the free carnitine and total carnitine concentrations in the carnitine supplementation group were significantly higher than in the control group (Fig. 1A,B). The mRNA levels of BBOX1, the key enzyme for carnitine synthesis, were not significantly different in the liver and in fasting state muscle; however, in feeding state muscle, the BBOX1 mRNA level in the carnitine supplementation group was significantly higher than that in the control group (Fig. 1C). No significant difference was found in the whole fish lipid level in all groups (Fig. 1D). Compared with the control group, the triglyceride (TG) content was significantly decreased in the carnitine supplementation group in the liver and muscle in both the feeding and fasting states; however, the TG content in the viscera was comparable in all groups (Fig. 1E). There were no interaction effects of the nutritional states and L-carnitine found in the whole fish lipid level and TG content in the different organs, except in the BBOX1 mRNA level (Supplemental Table 1), showing BBOX1 is simultaneously regulated by nutritional state and L-carnitine supplementation. The above data indicated that dietary carnitine supplementation could increase the in vivo carnitine concentration and reduce the TG content in the liver and muscle.

Figure 1: Effect of the dietary L-carnitine on the carnitine concentration, gene of carnitine synthesis and triglyceride in tissues of zebrafish.
Figure 1

(A) Free carnitine; (B) Total canitine; (C) The relative mRNA abundance of carnitine synthesis (gamma-butyrobetaine hydroxylase 1, BBOX1); (D) The crude lipid content of whole fish; (E) TG content of liver, muscle, viscera. All values are means ± SEM (n = 6). Values with *,**statistically differ at P < 0.05, P < 0.01.

The effect of dietary L-carnitine supplementation on mitochondrial and peroxisomal β-oxidation

As shown in Fig. 2, in the liver, the mitochondrial and total β-oxidation capability in carnitine supplementation group in the feeding and fasting state were significantly increased (Fig. 2A,C); however, peroxisomal β-oxidation capability was not affected. In the muscle, the mitochondrial and total β-oxidation capability in the feeding state were higher in the carnitine supplementation group than in the control, but showed no difference in the fasting state (Fig. 2A,C). There were no significant differences in the peroxisomal β-oxidation capability between the control and carnitine supplementation groups (Fig. 2B). The significant interaction between the nutritional state and L-carnitine was only seen in the mitochondrial and total β-oxidation capability in muscle (Supplemental Table 2). The results showed that dietary L-carnitine supplementation targets the liver and muscle, and has a lipid-lowing effect that functions mainly through mitochondrial FA β-oxidation, but not via peroxisomal FA β-oxidation.

Figure 2: Effect of the dietary L-carnitine on mitochondrial and peroxisomal β-oxidation capability of zebrafish.
Figure 2

(A) The mitochondrial β-oxidation capability; (B) The peroxisomal β-oxidation capability; (C) The total β-oxidation capability. All values are means ± SEM (n = 4). Values with *,**statistically differ at P < 0.05, P < 0.01.

The effect of dietary L-carnitine on the mRNA expression of lipid metabolism genes

Figure 3 shows the effects of dietary L-carnitine on the mRNA expressions of genes related to lipid metabolism. In the L-carnitine supplementation group, the mRNA level of the mitochondrial β-oxidation-related gene CPT1 was significantly increased (Fig. 3A), whereas the mRNA levels of the adipogenesis-related genes, such as ACC, FAS, and DGAT2, were significantly decreased (Fig. 3D–F), either in the liver or muscle. The mRNA level of LPL increased in the carnitine supplementation group in the liver in the feeding state but decreased in the muscle in the fasting state (Fig. 3G). The other lipid metabolism-related genes, including HAD, ACOX3, CD36, ATGL and HSL, remained unchanged in the livers and muscles of the two groups in the two states. Interaction effects between the nutritional state and carnitine levels were observed in the mRNA levels of ACC, FAS, and DGAT2 in liver and muscle, and ATGL and LPL in liver (Supplemental Table 3), showing nutritional state and L-carnitine both regulated the lipid synthesis and lipolysis. The results suggested that dietary L-carnitine could upregulate the mRNA expression of mitochondrial FA β-oxidation genes, and downregulate the mRNA expression of FA and TG synthesis genes in the liver and muscle.

Figure 3: The effect of the dietary L-carnitine on mRNA expression of genes of lipid metabolism.
Figure 3

All values are means ± SEM (n = 6). Values with *,**statistically differ at P < 0.05, P < 0.01.

The effect of the dietary L-carnitine on glycogen and glucose metabolism

The whole fish glycogen levels remained invariant in feeding states, but increased in fasting state (Fig. 4A), and the mRNA expression of glycogen synthase (Gys) was comparable among all groups (Fig. 4I). However, the genes related to glucose metabolism were affected by dietary L-carnitine supplementation. The mRNA level of PFK, a key glycolysis-related gene, was significantly decreased (fasting state) and increased (feeding state) in the muscle, but was comparable in the liver (Fig. 4B). The mRNA level of PK, another key glycolysis-related gene, was decreased in the muscle, and the decrease was significant in the feeding state (Fig. 4C). The mRNA expressions of PECK1 and G6Pa, which are key gluconeogenesis-related genes, were significantly upregulated by dietary L-carnitine in the liver, but did not change in the muscle (Fig. 4D,E). The mRNA expressions of insulin sensitivity-related genes, including insulin, Ira and Irb, were mostly unaffected by dietary L-carnitine supplementation (Fig. 4F–H), except for Irb in the carnitine supplementation group, which was expressed at a much higher level in the liver in the feeding state (Fig. 4H). The interaction effects between the nutritional state and L-carnitine were only seen in the mRNA levels of PFK in muscle and PECK1 and Irb mRNA levels in liver (Supplemental Table 4). In general, the above data suggested that dietary L-carnitine supplementation tended to increase gluconeogenesis, but decreased glycolysis; therefore, L-carnitine supplementation lowered the utilization of glucose.

Figure 4: The effect of the dietary L-carnitine on glycogen and mRNA expression of the genes related to glucose metabolism.
Figure 4

All values are means ± SEM (n = 6). Values with *,**statistically differ at P < 0.05, P < 0.01.

The effect of the dietary L-carnitine on proteins and genes of protein metabolism

To further understand the effect of dietary L-carnitine on the possible “protein-sparing effect” in zebrafish, the whole fish protein content and the mRNA expressions of genes related to protein metabolism were assayed. As shown in Fig. 5A, the crude protein level in the carnitine supplementation group was significantly increased compared with the control group in the feeding state, but was significantly decreased in the fasting state (Fig. 5A). The mRNA expressions of aminopeptidase N (APN) and oligopeptide transporter (PEPT1), which function in protein and amino acid digestion and absorption, and glutamate dehydrogenase 1a and 1b (GDH1a, GDH1b), which are essential in amino acid catabolism, were not affected significantly by dietary L-carnitine supplementation in the liver and muscle (Fig. 5B–E). The mRNA level of asparagine synthetase (ASNS), which plays roles in protein synthesis, was significantly decreased in the L-carnitine group in the fasting state, both the liver and muscle (Fig. 5F). Notably, in the L-carnitine supplementation group, the mRNA expression of mTOR, the regulatory factor for protein synthesis, was significantly increased in the liver in the feeding state (Fig. 5G). The interaction effects between the nutritional state and L-carnitine were observed in the whole fish protein content and the mRNA levels of PEPT1 in the muscle (Supplemental Table 5). These data suggested that dietary L-carnitine supplementation tended to increase protein degradation by inhibiting protein synthesis in the fasting state. However, the increased crude protein levels in whole fish and the increased mTOR expression in the liver of carnitine-fed zebrafish also showed that L-carnitine has positive effects on protein synthesis in the normal feeding period.

Figure 5: The effect of the dietary L-carnitine on protein and mRNA expression of the genes related to protein metabolism.
Figure 5

All values are means ± SEM (n = 6). Values with *,**statistically differ at P < 0.05, P < 0.01.

The effect of dietary L-carnitine on inflammatory gene expression

The expressions of three inflammation-related genes were measured to investigate the potential effects of dietary L-carnitine on inflammation. Among all groups, the mRNA expressions of IL-1β were comparable (Fig. 6A). The mRNA level of TNF-α, a strong inflammatory factor, was significantly decreased in the muscle of the L-carnitine supplementation group, in both the feeding and fasting states; however, no significant differences in the liver expression were found among the groups (Fig. 6B). The mRNA expression of TGF-β1, a known anti-inflammatory factor, was significantly increased by dietary L-carnitine supplementation in the muscle in the feeding and fasting states; however, its expression in the liver was not changed by L-carnitine supplementation (Fig. 6C). The interaction effects between the nutritional state and L-carnitine were observed in the mRNA levels of IL-1β and TGF-β1 in liver and TNF-α in muscle (Supplemental Table 6). These data indicated that dietary L-carnitine is likely to play roles in inflammation processes.

Figure 6: The effect of the dietary L-carnitine on mRNA expression of the genes related to inflammation.
Figure 6

All values are means ± SEM (n = 6). Values with *,**statistically differ at P < 0.05, P < 0.01.


As an essential factor in mitochondrial FA β-oxidation, L-carnitine has been used to alleviate fat accumulation-related metabolic syndromes in humans and other mammals for decades8,9,30. By contrast, in those fish that generally prefer to utilize protein, but not carbohydrate, to produce energy, carnitine has not only been used to decrease excess fat deposition in fish tissues, but is also thought to “save” the protein, which is used for energy through increasing the degradation of lipid for energy production. However, the effects of dietary carnitine supplementation in different fish species are contradictory31, but few previous studies performed comprehensive assays of lipid metabolism. In the present study, although growth promotion was not observed in zebrafish, dietary L-carnitine supplementation decreased the TG content in liver and muscle significantly, but did not affect the level in the viscera. Considering that liver and muscle, but not adipose tissue, are the main tissues for FA β-oxidation, the liver and muscle should be the main target tissues of dietary L-carnitine supplementation in fish. Indeed, the carnitine concentration in the liver and muscle of the experimental zebrafish increased with dietary L-carnitine supplementation.

By measuring the β-oxidation of [1-14C] palmitate in the mitochondria or peroxisome of the liver or muscle, the present study first indicated that dietary L-carnitine supplementation mainly increased mitochondrial FA β-oxidation activity, but not peroxisomal activity, at least in zebrafish. This was also confirmed by the high expression of CPT1 mRNA in the liver and muscle of the L-carnitine-fed zebrafish, and the comparable mRNA expressions of ACO in both tissues between the groups. The present study also indicated that dietary L-carnitine supplementation downregulated significantly the expression of adipogenesis-related genes, such as ACC, FAS and DGAT. Therefore, the lipid-lowing effects of dietary L-carnitine supplementation in the present study are likely to be caused by increased mitochondrial FA β-oxidation activity and decreased lipid synthesis, which was in consistent with some mammalian studies32,33.

Mammalian and human studies showed that dyslipidaemia is accompanied frequently by inflammation34; however, the L-carnitine-induced upregulation of CPT1 has been shown to prevent inflammation by decreasing inflammatory cytokines, such as TNF-α, in serum and the liver30,35. Similarly, in the present study, L-carnitine supplementation significantly increased the expression of CPT1, accompanied by decreasing and increasing the mRNA expression of TNF-α and the anti-inflammatory factor TGF-β1, respectively, in zebrafish muscle. This is the first report of this anti-inflammatory effect of L-carnitine in fish. Lipid accumulation-related metabolic dysfunctions have been observed widely in aquatic animals; therefore, the anti-inflammatory effects of L-carnitine in fish require further study.

Compared with mammals, fish have lower abilities to use carbohydrates as energy sources, thus fish cannot use glucose to produce energy efficiently10. A number of studies have indicated that, as compared with mammals, the responses of the activities of many glucose metabolism-related key enzymes, such as GK, PFK and PK, are not sensitive to the increased dietary carbohydrate content in fish36,37. In many fish species, high dietary carbohydrate induced lower growth, excess lipid deposition, and decreased stress resistance38,39,40. Therefore, fish nutritionists have tried different routes to promote the carbohydrate/glucose utilization in fish, including exploring the optimal sources and contents of dietary carbohydrate41,42, adding supplementary elements (e.g. chromium)43,44, or even developing gene-modified fish species45,46. However, in many fish species, these efforts are not ideal, and the regulatory mechanisms of carbohydrate metabolism are still poorly understood in fish. Recently, the interaction between lipid metabolism and carbohydrate metabolism has been observed in fish47,48. The phenomena that high-fat diet impairs glucose homeostasis has been reported in rainbow trout49,50. Our recent work further illustrated that glycolysis-related genes are upregulated or downregulated during low-fat diet or high-fat diet feeding, respectively51. In the present study, dietary L-carnitine supplementation improved lipid catabolism significantly, but increased the whole body glycogen deposition in the fasting state, indicating that glucose degradation was inhibited. This was also supported by the downregulated expressions of glycolysis-related genes (PFK and PK) in muscle, and the upregulated expressions in gluconeogenesis-related genes (PECK1 and G6Pa) in the liver. Similarly, dietary L-carnitine supplementation also increased the pyruvate carboxylase (PC) activity and glucose production in the liver in Atlantic salmon52. This evidence indicated that in fish, increasing the catabolism of lipids decreases the energy portion sourced from glucose degradation, resulting in increased gluconeogenesis and glycogen synthesis. This could be explained if, during energy homeostasis in fish, the endogenous acetyl-CoA concentration is relatively stable; therefore, if the increased lipid catabolism produces more acetyl-CoA for energy production, the portion of the acetyl-CoA sourced from other nutrients, such as glucose, would decrease correspondingly. Thus, the glycolysis pathway would be downregulated and gluconeogenesis would be upregulated. The relationships of the metabolic pathways are illustrated in Fig. 7.

Figure 7: Systemic regulation of L-carnitine in nutritional metabolism of zebrafish.
Figure 7

(A) The regulation of L-carnitine in feeding state. (B) The regulation of L-carnitine in fasting state. ACS, acyl-CoA synthetase. CACT, carnitine/acylcarnitine translocase.

One of the concerns associated with the use of L-carnitine in fish feed is to increase energy supply from lipid catabolism and “save” the protein from energy production to increase protein deposition. A number of studies in different fish species have indicated that dietary L-carnitine reduced the lipid content and increased the protein content, or decreased the protein oxidation in tissues and/or the whole body, showing a significant protein-sparing effect14,17,22,52. However, the underlying molecular basis remained unknown. In the present study, dietary L-carnitine supplementation increased the protein content of the whole body in the feeding trial, and did not affect the mRNA expressions of genes associated with protein and amino acid catabolism. However, notably, dietary L-carnitine significantly increased the mRNA expression of mTOR in the liver in the feeding state. The mTOR protein is one of the most important regulatory elements of protein synthesis53; therefore, dietary supplementation with L-carnitine might have positive effects on protein synthesis in zebrafish. In Atlantic salmon, dietary L-carnitine supplementation increased the amino acid concentration in plasma and the liver, especially the three branched-chain amino acids, including leucine, isoleucine, and valine, and increased the protein synthesis capacity, accompanied by the accumulation of protein in organs52. The studies in mammals and rainbow trout reported that leucine can effectively stimulate mTOR signalling54,55. Thus, we deduced that dietary L-carnitine supplementation can stimulate the expression of mTOR in fish by increasing the leucine concentration in the feeding state. The specific mechanism of the protein sparing effect of L-carnitine is detailed in Fig. 8. However, the present study also indicated that dietary L-carnitine supplementation significantly decreased the whole body protein content in the fasting state, and the gene expressions of ASNS and mTOR in the L-carnitine groups were also downregulated in the muscle in the fasting state. This might be explained if L-carnitine accelerated the degradation of lipids and proteins in the fasting state, and inhibited the synthesis of lipids and proteins to larger degree compared with the control. Compared with the whole body glycogen content, which was increased in the L-carnitine supplementation group in the fasting state, the decreased lipid and protein content in L-carnitine supplementation group in the fasting state confirmed that fish prefer to utilize proteins and lipids rather than carbohydrates.

Figure 8: The possible mechanism of the L-carnitine on protein synthesis.
Figure 8

To illustrate the systemic regulation of nutritional metabolism by L-carnitine in zebrafish, the interaction of different metabolic pathways in the L-carnitine-fed fish is shown in Fig. 7. Briefly, in the feeding state, dietary L-carnitine supplementation can increase lipid catabolism to produce more energy by stimulating mitochondrial FA β-oxidation and reducing FA and TG synthesis, utilising the “protein sparing effect” to elevate protein synthesis. Meanwhile, dietary L-carnitine supplementation also inhibited glycolysis and enhanced gluconeogenesis to decrease glucose-derived energy production (Fig. 7A). In the fasting state, dietary L-carnitine supplementation had similar effects on lipid and glucose metabolism to the feeding state, but also increased protein degradation and reduced protein synthesis (Fig. 7B). Nevertheless, our results also indicated that the interaction effects between nutritional state and L-carnitine supplementation existed in many parameters, including nutrient compositions, biochemical activities and metabolism-related gene expressions. This indicates that the regulatory mechanisms of L-carnitine in feeding or fasting states as shown in Fig. 7A,B could not be understood separately. Furthermore, it is of note that the changes of gene mRNA expression do not directly reflect the enzyme activities or protein functions, therefore, the regulatory mechanisms of L-carnitine at the transcriptional level still need further functional validation.


The present study indicated that dietary L-carnitine supplementation could increase the concentrations of carnitine in the liver and muscle, and decrease the lipid content in both tissues, possibly through enhanced mitochondrial β-oxidation and reduced adipogenesis. Dietary L-carnitine supplementation also increased the expression of mTOR in the liver, suggesting that it has positive roles in protein synthesis. However, dietary L-carnitine supplementation decreased glycolysis, because the increase in lipid-sourced ATP from L-carnitine supplementation might change the balance of energy homeostasis between lipids and carbohydrates. This is the first study to explore the mechanism of the positive effect of L-carnitine on the nutritional metabolism at transcriptional and biochemical levels in fish.

Materials and Methods

Animal ethics

All experiments were conducted strictly under the Guidance of the Care and Use of Laboratory Animals in China. This study was approved by the Committee on the Ethics of Animal Experiments of East China Normal University.

Fish, Diets and Sampling

In order to avoid the metabolic disturbance of estrogen during the sexual maturation of female fish, only male zebrafish (0.25 ± 0.03 g) were bought from Chinese National Zebrafish Resource Center (Wuhan, China). Before experiments, fish were acclimated in 100 cm × 45 cm × 45 cm tanks for 1 week and fed with commercial zebrafish diet (protein ≥ 50, lipid ≥ 8%) (Shengsuo Co., Shandong, China). After acclimation, six hundred zebrafish were randomly divided into 2 groups (3 tanks per group, 100 fish per tank): control group and carnitine group (Fig. 9). In the feeding trial, the carnitine was given by feeding fish with small wheat flour-dough particles containing L-carnitine (carnitine group) or not (control group), before basic diet feeding. In the preparation of the L-carnitine-contained wheat flour-dough particles, L-carnitine was first dissolved in pure water, and the carnitine solution was mixed with given amount of wheat flour to make wet dough. The wet dough was rubbed to small particles which size was proper for zebrafish, and then dried under 60 °C. When the dough particles were fed with 1% body weight, the L-carnitine composition in the dry wheat flour-dough particles could promise the final carnitine intake was 0.05% body weight, which is in the accordance with the dose used in other animals56,57. In the every morning, the dough particles containing L-carnitine or not were first fed to the carnitine group and control group, respectively, with 1% body weight. Afterwards, the same basic diet was fed to both groups with 3% body weight. The formulations of the basal diet and wheat flour-dough particle are listed in Table 1. The basal diet and dough particles were pelleted to a proper size (about 0.2 mg per particle) for zebrafish. After 6 weeks, the control and carnitine groups were divided into four groups: control-feeding, control-fasting, carnitine-feeding and carnitine-fasting (3 tanks/group and 50 fish/tank) (Fig. 9). In the two feeding groups, the feeding strategy was the same as that in the previous 6 weeks. In the two fasting groups, only dough particles were fed in the morning and the amount was decreased to 0.45% body weight, while the L-carnitine composition in the dough was increased to 10% to promise the same daily intake of L-carnitine with 0.05% body weight. Because the physiological effects of dietary L-carnitine supplementation were normally observed after 6–8 weeks in other animals24,57, the duration of the present experiment was 7 weeks. The experimental design was shown in Fig. 9. During the experiment, the photoperiod was 12 h:12 h, and the temperature was kept at 26 ± 2 °C. The weight of the fish in each tank was recorded every one week, and the feeding amount was adjusted correspondingly. At the end of the experiment, the whole fish in each tank were anthesthetized on ice, and sampled to collect liver, muscle and visceral for molecular and biochemical indexes. Hepatic, muscle and visceral triglyceride (TG) and whole fish glycogen were assessed by the commercial kit (Jiancheng Biotech Co., China). The crude lipid of the whole fish body was tested by using methanol and chloroform (1:2) as previously described58. Whole fish protein was measured by KjeltecTM 8200 (FOSS, Sweden). Because of the little mass of zebrafish organs (e.g. liver is 3–5 mg), samples were normally pooled from 2–4 fish. In the measurements and statistical calculations, 4–6 pooled samples were used (n = 4–6, see Figure legends).

Figure 9: The design and the protocol of the present experiment.
Figure 9
Table 1: Formulation of the basic diet and wheat flour-dough particles.

Carnitine concentration determination

After the feeding trial, the whole liver and parts of muscle of 4 fishes collected from each group were weighted and homogenized by pure-water (1:10, w/v) by using a drill-driven Teflon glass homogenizer (Scientz Biotec, China) with 3–4 strokes. Then the tissue homogenate centrifuged at 2000 rpm/min for 15 min (Centrifuge 5804R, Eppendorf, Germany). One hundred microliter aliquot of tissue homogenate supernatant was first incubated with 50 μL KOH (1 mol/L) at 37 °C for 30 min to fully hydrolyze the combined carnitine, and 10 μL HCl (1 mol/L) was then added to neutralize the solution (sample A). Sixty microliter distilled water was added directly into another 100 μL aliquot of tissue homogenate (sample B). Protein in sample A and B was removed by precipitating with 200 μL of cold ACN (containing 1 μg/mL carbachol, the internal standard, IS) and centrifuging at 16,900 × g for 20 min. One microliter of the supernatant was injected for LC-MS/MS analysis. Total carnitine was defined as the concentration of carnitine in sample A, free carnitine was defined as the concentration of carnitine in sample B.

An 1290 HPLC system (Agilent Technologies, Palo Alto, USA) and a 6460 triple-quadrupole mass spectrometer (Agilent Technologies, Palo Alto, USA) were used. The data was acquired and analyzed using MassHunter software (version; Agilent Technologies, Palo Alto, USA). Chromatographic separation was performed on a Phenomenex Luna 3u HILIC 200A Column (50*2 mm; Phenomenex, USA) due to the hydrophilicity and amphotericity of the analytes. The mobile phase system consisted of two phases, phase A (water) and phase B (water/acetonitrile, 1:9, v/v), and 10 mM of ammonium acetate was added to both of the phases. The analytes were separated on chromatographic column by isocratic elution (98% phase B) at the flow rate of 0.3 mL/min. The temperature of column oven was set at 40 °C, and the injection volume was 1 μL. The total run time for sample analysis was 4.5 min, and the flow was switched to MS during 0.8 to 3.1 min.

The mass spectrometer was operated in the positive electrospray ionization (ESI) mode, and the MS/MS analysis was performed in the multiple reaction monitoring (MRM) mode, monitoring the transitions of m/z 162.0 → 103.0 for carnitine, m/z 147.1 → 88.1 for IS, respectively. Fragmentation voltages/collision energy were optimized at 120 V/16 eV for carnitine, 90 V/14 eV for IS, respectively, for the best response of the transitions. Ion source conditions were set as follows: drying gas, 8 L/min at 300 °C; nebulizer gas, 20 psi; sheath gas, 9 L/min at 350 °C; collision gas (N2), 1.6 MPa; capillary voltage, 4000 V and nozzle voltage, 500 V.

Mitochondrial and peroxisomal [1-14C] palmitate oxidation in liver and muscle

After the feeding trial, the whole liver and parts of muscle of 3 fish collected from each group were weighted and homogenized in the ice-cold 0.25M-sucrose medium containing 2 mM-EGTA and 10 mM-Tris-Cl, pH 7.4,(1:40 and 1:10, w/v) by using a drill-driven Teflon glass homogenizer with 3–4 strokes. The samples of homogenate were used for the immediate measurement of mitochondrial and peroxisomal [1-14C] palmitate β-oxidation. Palmitate oxidation rates were detected at 28 °C using two media as already described59, the first media allowing both mitochondrial and peroxisomal β-oxidation to occur, and the second one allowing peroxisomal β-oxidation only. After 0.5 h, the radio activity initially held by [1-14C] palmitate was recovered on labelled short molecules released from the β-oxidative cycle and soluble in perchloric acid (acid-soluble products, ASP). The pure radioactive ASP medium was collected using 0.45 μm membrane filters and measured after mixing with the scintillation cocktail described by Du et al.60 in a liquid scintillation spectrometer MicroBeta2 Plate Counter (Perkin, USA).

Quantitative real-time PCR

Total RNA was isolated by using a Tri Pure Reagent (Aidlab, Cnina). The quality and quantity of total RNA were tested by NANODROP 2000 Spectrophpto (Thermo, USA). cDNAs of tissues total RNA were synthesized using a PrimerScriptTM RT reagent Kit with a gDNA Eraser (Perfect Real Time)(Takara, Japan) by S1000TM Thermal Cycler (Bio-Rad, USA). Elongation factor 1 alpha (EF1α) and β-actin were used as the reference genes61. The primers of EF1α and target genes (Table 2) for Quentitative PCR (qPCR) were designed to overlap intron. qPCR (20 μL) was carried out by using 2 × Ultra SYBR Mixture (CWbio, China) in 12 × 8 well plates in a CFX Connect Real-Time System (Bio-Rad), containing 10 μL of SYBR Mixture, 2 μL cDNA, 1.6 μL of qPCR primers (4 μM), and 7.4 μL nuclease-free water. The program of qPCR reaction included 95 °C for 10 min, 40 cycles of 95 °C for 5 s and 60 °C for 15 s. The melting curves of amplified products were generated to ensure the specificity of assays at the end of each PCR. qPCR efficiency was between 98% and 102% and the correlation coefficient was over 0.97 for each gene. Each qPCR run performed in triplicate and negative controls (no cDNA) were conducted. The method of 2−ΔΔCt was used for estimating the relative cDNA abundance (control feeding group as control).

Table 2: The primers used in the experiment.

Statistical analyses

All results are presented as mean ± SEM. Independent-samples t-test was performed to evaluate the significant difference (P < 0.05) of variables between control and carnitine group in feeding or fasting states. Two-way ANOVA analysis was used to explore the possible interactions existing between nutritional states and L-carnitine supplementation in all parameters. All analyses were conducted using the SPSS Statistics 19.0 software (IBM, USA).

Additional Information

How to cite this article: Li, J.-M. et al. Systemic regulation of L-carnitine in nutritional metabolism in zebrafish, Danio rerio. Sci. Rep. 7, 40815; doi: 10.1038/srep40815 (2017).

Publisher's note: Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Change history

  • 23 March 2017

    A correction has been published and is appended to both the HTML and PDF versions of this paper. The error has been fixed in the paper.


  1. 1.

    , & The regulation of energy metabolism pathways through L-carnitine homeostasis, Role of the Adipocyte in Development of Type 2 Diabetes (ed. ). 107–128 (INTECH Open Access Publisher, 2011).

  2. 2.

    Carnitine. Annu. Rev. Biochem. 57, 261–283 (1988).

  3. 3.

    . Metabolism and functions. Physiol. Rev. 63, 1420–1480 (1983).

  4. 4.

    , , & Carnitine, mitochondrial function and therapy. Adv. Drug Del. Rev. 61, 1353–1362 (2009).

  5. 5.

    et al. The syndrome of systemic carnitine deficiency. Clinical, morphologic, biochemical, and pathophysiologic features. Neurology 25, 16–24 (1975).

  6. 6.

    et al. Dietary eicosapentaenoic acid supplementation accentuates hepatic triglyceride accumulation in mice with impaired fatty acid oxidation capacity. Biochim. Biophys. Acta 1831, 291–299 (2013).

  7. 7.

    , , , & Effect of L-carnitine and soybean oil on growth performance and body composition of early-weaned pigs. J. Anim. Sci. 74, 1612–1619 (1996).

  8. 8.

    , , , & The effects of oral L-carnitine treatment on blood lipid metabolism and the body fat content in the diabetic patient. Asia Pac. J. Clin. Nutr. 7, 192–195 (1998).

  9. 9.

    , , & L-carnitine treatment in the hyperlipidemic rabbit. Metabolism 36, 1192–1196 (1987).

  10. 10.

    Utilization of dietary carbohydrate by fish. Aquaculture 124, 67–80 (1994).

  11. 11.

    et al. Regulation of feed intake, growth, nutrient and energy utilisation in European sea bass (Dicentrarchus labrax) fed high fat diets. Aquaculture 231, 529–545 (2004).

  12. 12.

    , , , & Protein/lipid ratios in extruded diets for Atlantic cod (Gadus morhua L.): effects on growth, feed utilisation, muscle composition and liver histology. Aquaculture 203, 101–119 (2001).

  13. 13.

    et al. Effect of dietary lipid level on growth performance, lipid deposition, hepatic lipogenesis in juvenile cobia (Rachycentron canadum). Aquaculture 249, 439–447 (2005).

  14. 14.

    & Effects of supplemental dietary carnitine on growth and lipid metabolism of hatchery-reared sea bass (Dicentrarchus labrax L.). Aquaculture 59, 177–186 (1986).

  15. 15.

    , & The Effect of Dietary L-Carnitine on Growth Performance and Lipid Composition in Red Sea Bream Fingerlings. Fish. Sci. 61, 1004–1008 (1995).

  16. 16.

    & Effect of dietary carnitine on growth rates and body composition of hybrid striped bass (Morone saxatilis male ×M. chrysops female). Aquaculture 187, 153–161 (2000).

  17. 17.

    & Dietary Carnitine and Lysine Affect Channel Catfish Lipid and Protein Composition. J. World. Aquac. Soc. 25, 169–174 (1994).

  18. 18.

    , , & The Effect of Dietary Carnitine Supplementation on Growth of Rainbow Trout Fingerlings. Fish. Sci. 63, 321–322 (1997).

  19. 19.

    , , & Influence of dietary l -carnitine on growth, biological traits and meat quality in Tilapia. Aquac. Res. 40, 1374–1382 (2009).

  20. 20.

    , & Effect of Dietary L-carnitine on Growth and Reproductive Performance of Male Oreochromis mossambicus (Peters). Fishery Technology 33, 84–90 (1996).

  21. 21.

    & Effect of dietary L-carnitine supplements on growth and body composition of fingerling rohu, Labeo rohita (Hamilton). Aquacult. Nutr. 4, 83–88 (1998).

  22. 22.

    et al. Dietary carnitine supplements increased lipid metabolism and decreased protein oxidation in African catfish (Clarias gariepinus) juveniles fed high fat levels. J. Aquacult. Trop. 18, 225–238 (2003).

  23. 23.

    , & The effect of dietary L-carnitine on the growth performance in fingerlings of the African catfish (Clarias gariepinus) in relation to dietary lipid. Br. J. Nutr. 69, 289–299 (1993).

  24. 24.

    et al. Dietary L-carnitine supplementation increases lipid deposition in the liver and muscle of yellow catfish (Pelteobagrus fulvidraco) through changes in lipid metabolism. Br. J. Nutr. 112, 698–708 (2014).

  25. 25.

    et al. Effects of dietary L-carnitine and chromium picolinate supplementations on performance and some serum parameters in rainbow trout (Oncorhynchus mykiss). Aquac. Int. 18, 213–221 (2010).

  26. 26.

    Zebrafish embryology builds better model vertebrate. Science 272, 1103–1103 (1996).

  27. 27.

    et al. Expression profiling identifies novel Hh/Gli-regulated genes in developing zebrafish embryos. Genomics 91, 165–177 (2008).

  28. 28.

    et al. Programming effects of high-carbohydrate feeding of larvae on adult glucose metabolism in zebrafish, Danio rerio. Br. J. Nutr. 111, 808–818 (2014).

  29. 29.

    , , & Zebrafish as a model organism for nutrition and growth: towards comparative studies of nutritional genomics applied to aquacultured fishes. Rev. Fish Biol. Fish. 21, 649–666 (2011).

  30. 30.

    et al. L-carnitine prevents progression of non-alcoholic steatohepatitis in a mouse model with upregulation of mitochondrial pathway. PLoS One 9, e100627 (2014).

  31. 31.

    L-carnitine and its attributed functions in fish culture and nutrition—a review. Aquaculture 249, 3–21 (2005).

  32. 32.

    , , & L-carnitine stimulates lipolysis via induction of the lipolytic gene expression and suppression of the adipogenic gene expression in 3T3-L1 adipocytes. J. Med. Food 9, 468–473 (2006).

  33. 33.

    , , , & Carnitine in metabolic disease: potential for pharmacological intervention. Pharmacol. Ther. 120, 149–156 (2008).

  34. 34.

    , , & The Relationship between Dyslipidemia and Inflammation, Nutritional and Metabolic Bases of Cardiovascular Disease (ed. ). 202–210 (Wiley‐Blackwell, 2011).

  35. 35.

    et al. L-carnitine ameliorates cancer cachexia in mice by regulating the expression and activity of carnitine palmityl transferase. Cancer Biol. Ther. 12, 125–130 (2011).

  36. 36.

    & Chapter 4 Tissue carbohydrate metabolism, gluconeogenesis and hormonal and environmental influences. Biochemistry & Molecular Biology of Fishes 4, 65–100 (1995).

  37. 37.

    , , & Glucose dehydrogenase, glucose-6-phosphate dehydrogenase and hexokinase in liver of rainbow trout (Salmo gairdneri). Effects of starvation and temperature variations. Comparative Biochemistry & Physiology B Comparative Biochemistry 99, 687–691 (1991).

  38. 38.

    , & Effects of dietary carbohydrate kind and level on growth, body composition and glycemic response of juvenile sunshine bass (Morone chrysops ♀× M. saxatilis ♂). Aquaculture 161, 187–199 (1998).

  39. 39.

    et al. Effect of dietary carbohydrate on haematology, respiratory burst activity and histological changes in L. rohita juveniles. Fish Shellfish Immunol. 19, 331–344 (2005).

  40. 40.

    , , & Effect of dietary carbohydrate on non-specific immune response, hepatic antioxidative abilities and disease resistance of juvenile golden pompano (Trachinotus ovatus). Fish Shellfish Immunol. 41, 183–190 (2014).

  41. 41.

    et al. Effect of dietary carbohydrate levels on growth performance, body composition, intestinal and hepatic enzyme activities, and growth hormone gene expression of juvenile golden pompano, Trachinotus ovatus. Aquaculture 437, 390–397 (2015).

  42. 42.

    et al. Effects of dietary carbohydrate source on growth performance, diet digestibility and liver glucose enzyme activity in blunt snout bream, Megalobrama amblycephala. Aquaculture 438, 75–81 (2015).

  43. 43.

    , , & Growth performance and starch utilization in common carp (Cyprinus carpio L.) in response to dietary chromium chloride supplementation. J. Trace Elem. Med. Biol. 27, 45–51 (2013).

  44. 44.

    & Carbohydrate utilization and digestibility by tilapia, Oreochromis niloticus × O. aureus, are affected by chromic oxide inclusion in the diet. J. Nutr. 125, 976–982 (1995).

  45. 45.

    et al. Growth hormone transgenesis influences carbohydrate, lipid and protein metabolism capacity for energy production in coho salmon (Oncorhynchus kisutch). Comparative Biochemistry & Physiology Part B Biochemistry & Molecular Biology 154, 121–133 (2009).

  46. 46.

    et al. Glucose metabolic gene expression in growth hormone transgenic coho salmon. Comparative Biochemistry & Physiology Part A Molecular & Integrative Physiology 170, 38–45 (2014).

  47. 47.

    et al. Regulation of de novo hepatic lipogenesis by insulin infusion in rainbow trout fed a high-carbohydrate diet. J. Anim. Sci. 89, 3079–3088 (2011).

  48. 48.

    et al. High dietary lipid level is associated with persistent hyperglycaemia and downregulation of muscle Akt-mTOR pathway in Senegalese sole (Solea senegalensis). PLoS One 9, e102196 (2014).

  49. 49.

    , & High dietary lipids induce liver glucose-6-phosphatase expression in rainbow trout (Oncorhynchus mykiss). J. Nutr. 132, 137–141 (2002).

  50. 50.

    , , , & High levels of dietary fat impair glucose homeostasis in rainbow trout. J. Exp. Biol. 215, 169–178 (2012).

  51. 51.

    et al. Systemic adaptation of lipid metabolism in response to low‐and high‐fat diet in Nile tilapia (Oreochromis niloticus). Physiological Reports 3, e12485 (2015).

  52. 52.

    , & Atlantic salmon (Salmo salar) fed L-carnitine exhibit altered intermediary metabolism and reduced tissue lipid, but no change in growth rate. J. Nutr. 126, 1937–1950 (1996).

  53. 53.

    & The mTOR pathway in the control of protein synthesis. Physiology 21, 362–369 (2006).

  54. 54.

    & Signaling pathways and molecular mechanisms through which branched-chain amino acids mediate translational control of protein synthesis. J. Nutr. 136, 227S–231S (2006).

  55. 55.

    et al. L-leucine, L-methionine, and L-lysine are involved in the regulation of intermediary metabolism-related gene expression in rainbow trout hepatocytes. J. Nutr. 141, 75–80 (2011).

  56. 56.

    , , , & Dietary L-carnitine increases plasma insulin-like growth factor-I concentration in chicks fed a diet with adequate dietary protein level. Br. Poult. Sci. 43, 117–121 (2002).

  57. 57.

    , , , & Investigation of Protective Effect of L-Carnitine on L-Asparaginase-Induced Acute Pancreatic Injury in Male Balb/c Mice. Dig. Dis. Sci. 60, 1290–1296 (2015).

  58. 58.

    & Seasonal variations in biochemical composition during the reproductive cycle of the intertidal gastropod Thais lamellosa Gmelin (Gastropoda, Prosobranchia). Can. J. Zool. 52, 305–318 (1974).

  59. 59.

    , , & Incomplete palmitate oxidation in cell-free systems of rat and human muscles. Biochim. Biophys. Acta 753, 399–410 (1983).

  60. 60.

    et al. Biochemical hepatic alterations and body lipid composition in the herbivorous grass carp (Ctenopharyngodon idella) fed high-fat diets. Br. J. Nutr. 95, 905–915 (2006).

  61. 61.

    , , , & Validation of Zebrafish (Danio rerio) Reference Genes for Quantitative Real‐time RT‐PCR Normalization. Acta Biochimica et Biophysica Sinica 39, 384–390 (2007).

Download references


This research was funded by National Basic Research Program of China (973 program 2014CB138600), National Natural Science Fund (31272676 and 31472290), and Program for New Century Excellent Talents in University.

Author information


  1. Laboratory of Aquaculture Nutrition and Environmental Health (LANEH), School of Life Sciences, East China Normal University, Shanghai, China

    • Jia-Min Li
    • , Ling-Yu Li
    • , Li-Jun Ning
    • , Dong-Liang Lu
    • , Dong-Liang Li
    • , Mei-Ling Zhang
    •  & Zhen-Yu Du
  2. Shanghai Key Laboratory of Regulatory Biology, Institute of Biomedical Sciences and School of Life Sciences, East China Normal University, Shanghai, China

    • Xuan Qin
    •  & Xin Wang


  1. Search for Jia-Min Li in:

  2. Search for Ling-Yu Li in:

  3. Search for Xuan Qin in:

  4. Search for Li-Jun Ning in:

  5. Search for Dong-Liang Lu in:

  6. Search for Dong-Liang Li in:

  7. Search for Mei-Ling Zhang in:

  8. Search for Xin Wang in:

  9. Search for Zhen-Yu Du in:


Z.Y.D., X.W. and J.M.L. designed the research. J.M.L., L.Y.L., X.Q. and L.J.N. conducted the research. J.M.L., D.L.L. and M.L.Z. analyzed data. D.L.L. and X.W. contributed to the final writing of the paper. J.M.L. and Z.Y.D. wrote the manuscript. All authors have read and approved the final manuscript.

Competing interests

The authors declare no competing financial interests.

Corresponding authors

Correspondence to Xin Wang or Zhen-Yu Du.

Supplementary information

Word documents

  1. 1.

    Supplementary Dataset

About this article

Publication history





Further reading


By submitting a comment you agree to abide by our Terms and Community Guidelines. If you find something abusive or that does not comply with our terms or guidelines please flag it as inappropriate.