Background
Oils from different plant sources differ in fatty acid composition. Coconut oil is rich in lauric acid (C12:0), comprising about 45–53% of the total fatty acid composition, while having very low content of fatty acids above C14 [
1‐
3]. Soybean oil, on the other hand, is rich in polyunsaturated and monounsaturated fatty acids particularly linolenic acid (C18:3), linoleic acid (C18:2) and oleic acid (C18:1) occupying 7.8, 53.2 and 23.4%, respectively of its total fatty acid composition while almost absent of C14 fatty acids and below [
3,
4]. Differences in fatty acid content and composition not only lead to diverging physico-chemical properties but also metabolic fates. Medium chain fatty acids (MCFAs; C8-C14) can enter cells without requiring fatty acid transporters unlike long chain fatty acids (LCFAs; C16-C22) [
5]. However, both types require carnitine activation in the muscle mitochondria [
5]. Moreover, most MCFAs are absorbed and metabolized in the liver for conversion to ketone bodies or incorporation to liver triglycerides (TG) especially with prolonged feeding [
6,
7]. These observations imply that MCFAs function as direct or precursor energy substrates for non-hepatic organs.
Training leads to several adaptations in whole body and organs. In the muscle, prolonged training increases glycogen storage, fatty acid uptake and utilization, and mitochondrial biogenesis among others [
8]. Trained muscle is then able to produce energy to sustain endurance. These adaptations are brought about by contraction- and high AMP/ATP ratio-induced AMP-activated protein kinase (AMPK) activation accompanying prolonged training [
8]. The peroxisome proliferator-activated receptors (PPARs) family and the estrogen-related receptors (ERRs) family are involved in these transcriptional adaptations with the coactivator PPARγ-coactivator 1α (PGC1A) synchronizing signals from AMPK and therefore of exercise [
9].
Fats with varying fatty acid composition differentially affect PPAR isotypes. PPARα and PPARβ/δ are highly expressed in peripheral tissues such as the muscle and control genes for oxidative metabolism while PPARγ is present mainly in adipose tissues orchestrating adipogenesis [
10]. LCFAs activate PPARs while the intensity of activation is influenced by the type of fatty acid and the PPAR isotype [
11,
12]. MCFAs, particularly C10 and C12 strongly activate PPARγ while C10 and C14 activate PPARα and PPARβ/δ to a certain extent [
11‐
14]. Also, MCFAs upregulate mitochondrial biogenesis better than LCFAs [
15]. Therefore, MCFAs and LCFAs function as direct or indirect signaling molecules for mitochondrial oxidative capacity possibly through PPARα and/or PPARβ/δ [
9,
15‐
17]. Unlike PPARs, no endogenous ligands for ERRs have been identified. However, ERR isotypes are constitutively active and the binding of PGC1A potentiates their activity [
18,
19]. Therefore, downstream effects of training on transcriptional adaptations are also influenced not only by the expression of these transcription factors but also by available fatty acids for activation.
The effects of short- and long-term feeding of MCFAs and LCFAs with or without training have been investigated. In our laboratory, MCFAs (coconut oil) improved swimming capacity in trained mice and increased mitochondrial enzyme activities relative to LCFAs (soybean oil) [
20]. With training, MCFAs (C8 and C10) increased energy expenditure in rats relative to LCFAs (rapeseed oil) [
21]. In untrained mice, increased mitochondrial markers with MCFAs (coconut oil) relative to LCFAs (lard) was associated with whole body and localized muscle oxygen consumption as assessed with in vitro, ex vivo and in vivo experiments [
15]. In rats, while MCFAs (C8 to C12) did not affect endurance performance, LCFAs (primarily C16) decreased endurance associated with increased cardiac mitochondrial uncoupling [
22]. In contrast, LCFAs (soybean oil) with training did not impair but rather improved endurance on a treadmill in wild-type C57BL/6J mice [
23]. Despite these previous investigations, literature on the comparative effects of MCFAs and LCFAs on training adaptations remains scarce. Furthermore, the absence of comparison with high-carbohydrate/low-fat diet may limit the adaptability of these diets in athletic dietary management.
The objective of the study was to update the current knowledge of physiological adaptations occurring during exercise training with low-fat diet, and medium-fat diets containing coconut oil or soybean oil specifically on whole-body metabolism at rest and during exercise, substrate metabolism, mitochondrial functions, and genetic adaptive responses in the muscle and liver under the treadmill exercise modality.
Methods
Animals
Seven (7)-week old male C57BL/6N mice were purchased from Shimizu Laboratory Supplies (Hamamatsu, Shizuoka, Japan). Mice were housed in an animal room at 22 ± 0.5 °C and 50% humidity with a 12 h light-dark cycle (lights on and off at 6:00 and 18:00, respectively). Mice were acclimatized to this environment with ad libitum access to a standard chow diet (Oriental Yeast Co., Tokyo, Japan) and water for 7 to 10d before changing to assigned diets. Mice were randomly assigned to the following purified diets: low-fat diet (L; 20% kcal from casein, 70% kcal from cornstarch and 10% kcal from soybean oil); coconut oil diet (C; 20% kcal from casein, 50% kcal from cornstarch, 10% kcal from soybean oil and 20% kcal from coconut oil); and soybean oil diet (S; 20% kcal from casein, 50% kcal from cornstarch and 30% kcal from soybean oil). All diets contained vitamin and mineral mixes and were prepared by Research Diets (NJ, USA) based on the D12450K formulation. Training of mice was commenced following diet assignment. Dietary groups were divided into untrained (U) and trained (T) groups. Animal experiments were performed according to the Kyoto University Guidelines for the Ethical Treatment of Laboratory Animals as approved by the Kyoto University Animal Experimentation Committee with the number (29–39).
Treadmill training
Training was conducted daily for 30d from 6:00 using a treadmill for rodents (MK-680; Muromachi, Tokyo, Japan). On the first 15d, mice ran for 1 h at 15 m/min at 3° incline. From the 16th day, intensity was increased to 18 m/min. Mice were forced to run by poking. All mice completed the training program.
Basal indirect calorimetry
Basal indirect calorimetry using the ARCO-2000 system (Tokyo, Japan) under ad libitum feeding and resting conditions as described in [
24] was performed on a subset of mice. Mice were assigned to calorimetry chambers on the 28th day of training to facilitate acclimatization. Actual measurement was performed 24 h to 48 h post-training which represents a full light-dark cycle devoid of acute exercise effects. Mice were sacrificed after measurements. Sample collection was based on Manio, et al. [
23].
Exercise indirect calorimetry and endurance test
Indirect calorimetry during exercise with treadmill endurance test was performed on a subset of mice. About 1.5-2 h before the run (6:00), mice were placed in sealed treadmill chambers (Mousebelt-200; Arco System, Tokyo, Japan) at 10° incline to acclimatize. After warming-up for 2 min, intensity was increased to 15 m/min. After 30 min, the intensity was increased to 18 m/min and maintained for 30 min. Then, the intensity was increased to 21 m/min and kept herein until exhaustion. Mice were stimulated with electrical stimulus (0.2 mA) and occasional noise and poking. Exhaustion was ruled if mice remained on electrodes or could not sustain running for 20s despite additional stimulation.
Fixed time run
Fixed time run was performed on a subset of mice. Food was removed 2 h before running. Mice were individually placed on a moving treadmill set at an intensity of 10 m/min. After 2 min of warm-up, mice were transferred to a moving treadmill set at 15 m/min, 10° incline. After exactly 30 min, mice were removed from the treadmill and immediately sacrificed. Experiments were performed in the same conditions as exercise indirect calorimetry.
Blood chemistry and tissue metabolites
Serum was measured for glucose, triglycerides (TG), non-esterified fatty acids (NEFA) and beta-hydroxybutyrate (β-HB). Glycogen and TG were measured in organs [
25‐
27]. Measurements are detailed in [
23].
Protein extraction and enzyme activities
Muscle and liver were lysed in a 1% NP-40 buffer as detailed in [
23]. Beta-hydroxyacyl-CoA dehydrogenase (β-HAD) activity was measured according to a procedure by Holloway, et al. [
28]. Succinyl-CoA:3-oxoacid CoA-transferase (SCOT) activity was measured based on Williamson, et al. [
29]. Citrate synthase (CS) activity was measured according to Srere [
30]. Cytochrome c oxidase of the mitochondrial electron transport chain, also known and hereby referred to as Complex IV, was measured based on Mac Arthur, et al. and Spinazzi, et al. [
31,
32]. Acetoacetyl-CoA thiolase (AACT) and deacylase (AACD) activities were measured based on Williamson, et al. [
33]. All enzyme activity measurements were modified to adapt to a 96-well plate system as detailed in [
23].
Reverse transcriptase quantitative polymerase chain reaction (RT-qPCR)
Total RNA was extracted with Tripure Isolation Reagent (Roche, Mannheim, Germany) and GenElute Mammalian Total RNA Miniprep Kit (Sigma-Aldrich, MO, USA) as detailed in [
23]. Total RNA (1.8 μg and 1.5 μg for muscle and liver, respectively) was reverse transcribed with Transcriptor First Strand cDNA Synthesis Kit (Roche, Mannheim, Germany). Messenger RNA (mRNA) expression levels were quantified using intron-spanning primers and corresponding Universal Library Probes (Roche, Mannheim, Germany) listed in Table
1. Values were rationalized to
Hprt expression [
34].
Table 1
Primers and probes in RT-qPCR
PPARγ Coactivator 1α, (Pgc1a) | F: tgtggaactctctggaactgc | 63 | NM_008904.2 |
R: agggttatcttggttggcttta |
PPARα, (Ppara) | F: ccgagggctctgtcatca | 11 | NM_011144.6 |
R: gggcagctgactgaggaa |
PPARβ/δ, (Pparb/d) | F: atgggggaccagaacacac | 11 | NM_011145.3 |
R: ggaggaattctgggagaggt |
ERRα, (Erra) | F: gtgggcatgctcaaggag | 29 | NM_007953.2 |
R: ggaaaggcaaagggtcca |
ERRβ, (Errb) | F: ggcgttcttcaagagaacca | 49 | NM_011934.4 |
R: tccgtttggtgatctcacatt |
ERRγ, (Errg) | F: aagtgggcatgctgaaagaa | 29 | NM_011935.3 |
R: cagcatctattctgcgcttg |
Lipoprotein lipase, (Lpl) | F: tggataagcgactcctacttcag | 22 | NM_008509.2 |
R: tccctagcacagaagatgacc |
Carnitine palmitoyltransferase 1B, (Cpt1b) | F: ccatcattgggcacctct | 104 | NM_009948.2 |
R: gtctccgtgtagcccaggt |
Glucose transporter, 4 (Glut4) | F: tcgtcattggcattctggt | 104 | NM_009204.2 |
R: agcagtggccacagggta |
Fatty acid transport protein, 1 (Fatp1) | F: cttcctaaggctgccattgt | 49 | NM_011977.3 |
R: ggcagtcatagagcacatcg |
Myosin heavy chain, 2a (Myh2) | F: tcttctctggggcacaaact | 22 | NM_001039545.2 |
R: cccttcttcttggcaccttt |
Carnitine palmitoyltransferase, 1A (Cpt1a) | F: aaagcaccagcacctgtacc | 34 | NM_013495.2 |
R: aacctccatggctcagacag |
3-Hydroxy-3-methylglutaryl-CoA synthase, 2 (Hmgcs2) | F: ctgtggcaatgctgatcg | 93 | NM_008256.4 |
R: tccatgtgagttcccctca |
Hypoxanthine-guanine phosphoribosyl transferase, (Hprt) | F: cctcctcagaccgcttttt | 95 | NM_013556.2 |
R: aacctggttcatcatcgctaa |
Immunoblotting
Protein concentration of lysates was adjusted with lysis buffer and 4× Laemilli buffer containing 20% mercaptoethanol. Samples were loaded on 8% polyacrylamide gels. Semi-dry transfer to PVDF membranes was performed in a transfer buffer containing 20% methanol. After transfer, Ponceau S staining was performed. Excess stain was removed in distilled water. Membranes were visualized and digitized (LAS-3000; Fujifilm, Tokyo, Japan). Membranes were blocked in 5% skim milk powder in Tris-buffered saline with 0.1% Tween-20 buffer (TBST) containing 0.05% ProClin 300 (Sigma Aldrich, MO, USA). Membranes were cut at sections corresponding to regions of molecular weight previously identified to contain the protein of interest. Membranes were incubated in goat anti-fatty acid translocase/cluster of differentiation 36 (CD36) antibody (1:2000; AF2519, R&D Systems, MN, USA), goat anti-pyruvate dehydrogenase kinase 4 (PDK4) antibody (1:1000; C-16, Santa Cruz Biotechnology, CA, USA), rabbit anti-PGC1A antibody (1:1000; H-300, Santa Cruz Biotechnology, CA, USA) or rabbit anti-glucose transporter 4 (GLUT4) antibody (1:2000, AB1346, Chemicon International, CA, USA) at 4 °C for 20 h. Membranes were washed 3× in TBST to remove excess primary antibody. Horseradish peroxidase-labelled secondary antibody incubation was performed in anti-goat IgG (1:1000; P0449; Dako, Tokyo, Japan) or anti-rabbit IgG (1:1000; P0399; Dako, Tokyo, Japan) for 3 h at 4 °C. After washing 3× in TBST to remove excess secondary antibody, membranes were visualized by chemiluminescent detection (Western Lightning Plus ECL; Perkin Elmer, MA, USA). Signals corresponding to PGC1A (91 kDa), CD36 (88 kDa), GLUT4 (58 kDa), PDK4 (47 kDa) and Ponceau S signals were quantified using the software MultiGauge V3.2 (Fujifilm, Tokyo, Japan) with automatic background detection.
Statistical analyses
Statistical analyses were performed using the Prism 5.0 software (Graphpad Software, CA, USA). Points in exercise indirect calorimetry time-course data are presented as means ± SEM. Each time point was analyzed by one-way analysis of variance (ANOVA) to compare groups of different diets and by Student’s unpaired t-test to compare groups of different training states receiving the same diet. Other data are presented as means ± SEM and were analyzed accordingly using one-way ANOVA followed by Newman-Keuls post-hoc test and Student’s t-test. Significance level (α) was set at 0.05.
Discussion
Fat as a bioactive compound influences metabolism by inducing muscle and liver phenotypic remodeling through transcriptional activation of PPARs [
15,
17,
38‐
40]. We hypothesized that diets varying in fat source and proportion together with training would lead to different adaptations in the muscle and liver consequently affecting whole-body metabolism and endurance. Comparative studies on the effects of different diets with training have been conducted [
20,
41,
42]. However, data on energy expenditure at rest and during exercise, substrate utilization, and gene transcription are scarce. Coconut oil is a good source of MCFAs which are rapidly metabolized relative to LCFAs [
43]. MCFAs with training improve endurance in swimming [
20] but other aspects of adaptation required further investigation. We aimed to update the current knowledge on the effects of MCFAs and fat types. We show that fat source and content in the diet exert variably influence different aspects of basal and treadmill training adaptations particularly on endurance, exercise whole-body metabolism, energy substrate storage and utilization, and genetic and biochemical characteristics of the muscle and liver.
Medium-fat diets increased FAT regardless of fat type and training promoted CHO at rest without affecting energy expenditure. Utilizing a lower intensity training protocol with the same soybean oil diet increased CHO, but it accompanied increased energy expenditure without affecting FAT [
23] suggesting that different training intensities differentially affect resting metabolism [
44,
45]. Our data in relation to [
15] suggest that higher absolute MCFAs content may increase VO
2 even without training relative to LCFAs.
Whole-body metabolism during exercise under different diets is associated with changes in VO
2max [
42]. Unfortunately, we could not measure VO
2max because of technical limitations. During exercise under slight food deprivation, training but not diet influenced whole-body metabolism suggesting that at rest with ad libitum feeding, diet composition determined differences in resting energy metabolism while general effects were due to training. Moreover, training lowered VO
2 (and energy expenditure) implying that exercise economy increased in trained groups during exercise [
46]. In our previous study using a lower training intensity, decreased RER without changes in VO
2 was observed suggesting higher fat utilization [
23]. These observations further indicate that changes in energy metabolism at rest or during exercise is influenced by training intensity. It is important to note that inaccurate calculations in CHO and FAT, especially in C with ketosis during exercise, may exist because the complete oxidation of acetoacetate and β-HB give respiratory quotient values of 1.0 and 0.89, respectively [
47]. Unfortunately, corrections for ketone body oxidation and its relative contribution to substrate metabolism could not be performed because a time-course ketone body profile in the blood was not available.
Our group and others show that diets high in fat induce muscle mitochondrial biogenesis and also imply that MCFAs promote mitochondrial biogenesis better than LCFAs at the same absolute concentration [
15,
17]. β-HAD activity increased in C without training. Also, in C2C12 muscle cells, C10 and C12 fatty acids increased succinate dehydrogenase activity [
15]. However, changes in the activities of other enzymes may not be entirely dependent on fatty acid species but on their amount as seen with SCOT and CS. We also showed that training universally increased mitochondrial function suggesting mitochondrial biogenesis. Therefore, elevated oxidative capacity in C, albeit small, likely improved endurance in the untrained state as measured by work while robust increase in mitochondria improved endurance in trained mice regardless of diet [
48,
49]. In contrast with the swimming modality, MCFAs increased swimming time attributed to increased CS and SCOT relative to LCFAs [
20] underscoring the notion that different training modalities variably influence adaptation, and potentially, endurance.
Exercise increases PGC1A and this co-activates or potentiates the PPARs and ERRs in the control of mitochondrial biogenesis [
9]. We did not observe changes in PGC1A mRNA and protein despite increased mitochondrial function suggesting differential effects of training intensity on their half-lives [
23,
50,
51]. However, we show that diets influenced basal and training-induced changes in mRNA expression of PPARs and ERRs. ERRs had decreased expression in medium-fat diets. This did not negatively affect mitochondrial enzyme activities,
Pgc1a or
Erra in the untrained state suggesting that at the basal level, homeostatic control and/or other ERR isotypes likely compensated for decreased mRNA expression of ERRγ [
52,
53]. Training increased
Erra and
Errb, which could explain the increased mitochondrial biogenesis in the muscle [
49].
High-fat diets increases
Ppara but not the other isotypes in rats [
54]. We did not observe significant elevations in
Ppara in the untrained state possibly due to a relatively lower fat content of our diets. On the other hand, only
Pparb/d increased with training in contrast to increased
Ppara and unchanged
Pparb/d when trained at a lower intensity even with the same soybean oil diet [
23] suggesting that PPARs respond differently with training intensity. Interestingly, coconut oil impaired the training-induced upregulation of
Pparb/d. Consistent with increased
Pparb/d with training, target genes related to glucose and fat utilization, and fiber type remodelling (
Glut4,
Cpt1b,
Fatp1 and
Myh2) responded similarly especially in S [
55‐
58]. Because exercise increases fatty acid uptake by translocation of CD36 to the sarcolemma, not only increased expression of PPARβ/δ but also increased fatty acid-induced activation and availability could upregulate these targets [
23,
59] which may explain some of the differences between S and L with training. On the other hand, these changes could not be attributed to increased CD36 as its mRNA and protein did not respond as expected [
23] indicating that training intensity affects specific genetic adaptations.
The expression of
Cpt1b and
Lpl with medium-fat diets in the untrained state is probably related to PPARα as this isotype also controls their transcription in the muscle [
60]. Because functions of PPAR isotypes overlap in some of these genes, we could not discount the contribution of PPARγ. Although PPARγ is abundant in adipose tissues, it is also present in skeletal muscle and MCFAs and LCFAs strongly activate PPARγ [
11‐
14,
61].
Total caloric intake was similar among diet groups. This means that the minimum amount of consumed soybean oil was similar among diet groups suggesting that coconut oil inhibited training-induced upregulation of PPARβ/δ and some downstream targets. Whether MCFAs inhibited LCFAs by competitive binding in PPARβ/δ activation requires further research. While competitive binding assays between fatty acids and synthetic agonists have been performed [
11,
12,
14], competitive binding assay among fatty acids to PPARs has yet to be undertaken. Overall, PPAR-related gene transcription as a training adaptation was influenced by the type of fat in the diet. Also, these adaptations reflect the route of catabolism of energy substrates within these diets during exercise.
In the untrained state, diets did not affect pre-exercise muscle glycogen possibly due to similar circulating lipids or serum β-HB as these influence glycogen storage [
62‐
65]. Training increases muscle glycogen but ketone bodies, particularly acetoacetate, inhibit insulin-stimulated glucose uptake that occurs during feeding after training [
66]. This may explain the inhibited glycogen accumulation in C. Furthermore, while GLUT4 is not essential for glycogen repletion per se it could influence glycogen accumulation with insulin post-exercise by increasing the rate of glucose transport [
67‐
69]. Liver glycogen accumulation was also impaired in C and to a lesser extent in S which was emphasized by training. This could be partly explained by exhaustion of hepatic glycogen reserves with MCFAs and the glycogen replenishing effect carbohydrates [
70,
71].
Increased muscle glycogen and glycogen sparing improves endurance by slowing the utilization of circulating glucose and liver glycogen [
72,
73]. Muscle glycogen is spared by improved utilization of fatty acids and ketone bodies [
74‐
77] linking glycogen sparing with high serum β-HB, decreased serum NEFA and intramuscular TG especially in C with training post-exercise. These suggest the existence of compensatory mechanisms to preserve endurance despite low pre-exercise muscle glycogen in this group. Nevertheless, muscle glycogen availability and utilization together with increased oxidative and mitochondrial functions likely promoted robust endurance improvement in all trained groups.
In the liver, unlike LCFAs, MCFAs can bypass fatty acid transport proteins to enter the cell and mitochondria for oxidation and these undergo β-oxidation for complete oxidation or ketogenesis [
36,
59,
78‐
81]. Improved oxidative capacity with increased upstream β-HAD activity suggested increased capacity to produce ketones particularly in C [
82] despite higher downstream ketogenic enzyme activities in S than C. Furthermore, because MCFAs are undetectable in the serum at rest, the overwhelming increase in β-HB during exercise in C suggests that liver and adipose tissues stored MCFAs, released them to circulation during exercise and were immediately catabolized [
6,
20,
79,
83‐
85].
Fatty acid oxidation and ketogenesis in the liver is controlled by PPARα [
10]. Diets did not influence
Ppara but training increased its expression in C. Fibroblast growth factor 21 (FGF21) is induced by acetoacetate via an upstream regulator and this upregulates PPARα [
38,
86] thus connecting the link between training, coconut oil, increased ketone bodies during training, and increased
Ppara. While non-significant, increased
Cpt1a and β-HAD activity in this group suggest PPARα activation in the liver [
82].
Hmgcs2, the gene that encodes the first enzyme of ketogenesis [
36], was unaffected by diet nor training suggesting that high β-HB observed post-exercise in C was primarily caused by increased supply of ketogenic precursors for β-oxidation in C rather than changes in ketogenic activity. On the other hand, relatively lower β-HB with training at post-exercise is likely because of increased muscular utilization accompanying increased SCOT activity. Whether increased oxidative capacity prevented increase of liver weight in C with training was not investigated. Overall, coconut oil with training promoted liver remodeling to an oxidative phenotype without influencing mitochondrial biogenesis.