From a quantitative point of view, muscle glycogen represents the most important fuel source during moderate- to high-intensity endurance-type exercise (37,45). Because a direct relationship exists between fatigue and muscle glycogen depletion (6,7), postexercise recovery time is mainly determined by the rate of muscle glycogen repletion. Ingestion of 1.2 g·kg−1·h−1 CHO, when provided at frequent intervals (i.e., every 15–30 min), allows optimal postexercise muscle glycogen synthesis rates (8,12,19,21,23,41,44,47). Further increasing CHO intake, up to 1.6 g·kg−1·h−1, does not augment postexercise muscle glycogen storage (19). However, two nutritional intervention strategies have recently been suggested to accelerate muscle glycogen synthesis during postexercise recovery.
Postexercise amino acid and/or protein coingestion with CHO has been established as an effective dietary strategy to strongly augment postprandial insulin release (35,46,48). As insulin stimulates both glucose uptake and glycogen synthase activity in skeletal muscle tissue (10,22,50), it has been suggested that coingestion of an insulinotropic amino acid and/or protein mixture can further accelerate postexercise muscle glycogen synthesis. In accordance, we (47) as well as others (4,20,52) showed that amino acid and/or protein coingestion with CHO (0.5–0.8 g·kg−1·h−1) accelerates postexercise glycogen synthesis. However, several other studies reported no additional benefits of protein coingestion on postexercise muscle glycogen synthesis when more than 1.0 g·kg−1·h−1 CHO was administered (19,23,44). It has been suggested that the latter might be attributed to the fact that muscle glycogen repletion was assessed during a restricted timeframe of only 3 (23) and 4 h (19,44) of postexercise recovery. Such a short timeline might be insufficient to assess the full potential of insulin on postexercise muscle glycogen synthesis rates because glucose uptake and glycogen synthesis become more insulin dependent throughout the latter stages of the recovery period (22,34). So far, no study has assessed the effect of coingesting an insulinotropic amino acid/protein mixture on postexercise muscle glycogen synthesis during 6 h of recovery when an optimal amount of CHO (1.2 g·kg−1·h−1) is provided.
Recently, Pedersen et al. (31) reported a substantial increase in postexercise muscle glycogen synthesis rate when caffeine (2 mg·kg−1·h−1) was coingested with CHO (1.0 g·kg−1·h−1). The latter was remarkable as previous work generally reports a negative effect of caffeine administration on glucose disposal (1,17,29,33,43). Although the mechanism(s) that might explain the proposed stimulating effect of caffeine coingestion on muscle glycogen repletion remained unresolved, the authors (31) speculated that caffeine coingestion might stimulate intestinal glucose absorption (49,51). Studies are warranted to assess whether caffeine coingestion accelerates postexercise muscle glycogen repletion when an optimal amount of CHO (1.2 g·kg−1·h−1) is provided.
In the present study, we tested both hypotheses that coingestion of an insulinotropic amino acid/protein mixture or caffeine with 1.2 g CHO·kg−1·h−1 accelerates postexercise muscle glycogen synthesis when compared with ingestion of CHO only. Therefore, we subjected 14 male cyclists to glycogen depletion exercise on 3 occasions, after which they ingested CHO (1.2 g·kg−1·h−1), CHO (1.2 g·kg−1·h−1) with an amino acid/protein mixture (0.3 g·kg−1·h−1), or CHO (1.2 g·kg−1·h−1) with added caffeine (1.7 mg·kg−1·h−1) during a 6-h recovery period. Muscle biopsies were collected immediately after cessation of exercise and after 6 h of postexercise recovery to assess the increase in muscle glycogen content. Furthermore, [U-13C6]-labeled glucose was administered orally to assess potential differences in the appearance rate of the ingested glucose in the circulation.
Fourteen well-trained male cyclists participated in this study (age = 24 ± 1 yr, bodyweight = 71.6 ± 2.5 kg, body mass index = 21.7 ± 0.4 kg·m−2, maximal workload capacity (Wmax) = 387 ± 11 W, maximal oxygen uptake capacity (V˙O2max) = 61.5 ± 1.2 mL·kg−1·min−1). Subjects cycled at least 100 km·wk−1 and had a training history of >3 yr. Subjects were fully informed on the nature and possible risks of the experimental procedures, before their written informed consent was obtained. The study was approved by the Medical Ethical Committee of the Maastricht University Medical Centre, Maastricht, The Netherlands.
All subjects participated in a screening session, which was performed 1 wk before the first experiment. Subjects performed an incremental exhaustive exercise test (27) on an electronically braked cycle ergometer (Lode Excalibur, Groningen, The Netherlands) to assess V˙O2max and Wmax.
Diet and Activity before the Experiments
All subjects received the same standardized dinner (81 ± 3 kJ·kg−1 body weight, consisting of 58 energy% (En%) CHO, 31 En% fat, and 11 En% protein) the evening before each test day. All volunteers refrained from any sort of exhaustive physical labor and/or exercise and kept their diet as constant as possible 2 d before each experimental day. In addition, subjects filled in food intake and physical activity questionnaires for 2 d before the start of the first experiment, which were used to standardize food intake and physical activity before the second and third experimental day. Furthermore, subjects were instructed to refrain from caffeine containing food products like coffee, tea, cola, and chocolate for 2 d before each experimental day. Subjects’ regular caffeine use was 3.5 ± 0.6 U·d−1, in which a unit represents one cup of coffee with a caffeine content of ∼70 mg.
Subjects performed 3 randomized tests, each separated by at least 1 wk. During each test, they were first subjected to a glycogen depletion protocol. Thereafter, subjects were studied for 6 h while ingesting only CHO in the control trial (CHO), CHO plus a protein plus leucine mixture in the CHO + PRO trial, or CHO plus caffeine in the CHO + CAF trial. During the 6 h postexercise recovery period, subjects remained in supine rest. Beverages were provided every 30 min, and tests were performed in a randomized double-blind order. Muscle biopsies were taken immediately after exercise and at the end of the 6-h recovery period to assess changes in muscle glycogen content.
Subjects reported to the laboratory at 8:00 a.m. after an overnight fast. Muscle glycogen depletion was established by performing an intense exercise protocol on a cycle ergometer (26). This muscle glycogen depletion protocol started with a 10-min warm-up period at a 50% Wmax workload. Thereafter, subjects were instructed to cycle 2-min block periods at alternating workloads of 90% and 50% Wmax, respectively. This was continued until subjects were no longer able to complete the 2 min at 90% Wmax. That moment was defined as the inability to maintain cycling speed at 60 rpm. At that moment, the high-intensity block was reduced to 80% Wmax. Again, subjects had to cycle until they were unable to complete a 2-min block at 80% Wmax, after which the high-intensity block was reduced to 70% Wmax. Subjects were allowed to stop when pedaling speed could not be maintained at 70% Wmax. Water was provided ad libitum during the exercise protocol of the first test day, and the same amount of water was provided during the second and third test day. A fan was placed 1 m from the subjects to provide cooling and air circulation during the exercise protocol. After cessation of exercise, a muscle biopsy was taken from the vastus lateralis muscle. Thereafter, a Teflon catheter (Baxter BV, Utrecht, The Netherlands) was inserted in an antecubital vein, a resting blood sample was taken, and subjects received the first bolus of the test drink (t = 0 min). Subjects were observed for the following 6 h during which they received a beverage with a volume of 3 mL·kg−1 every 30 min until t = 330 min. Blood samples were taken at 15-min intervals for the first 90 min of recovery and every 30 min thereafter until t = 360 min. Immediately after acquiring the final blood sample, a second biopsy was taken from the vastus lateralis muscle of the other leg.
Subjects were asked to fill out a questionnaire using a 10-point scale (1 = not at all, 10 = very, very much) at t = −5, 175, and 355 min. This questionnaire contained questions regarding the presence of gastrointestinal (GI) distress and addressed the following complaints: nausea, bloated feeling, belching, stomach problems and GI cramping, vomiting, diarrhea, the urge to urinate and/or defecate, headache, and dizziness. One question regarding the taste of the test drink was also conducted (1 = horrible, 10 = very tasty).
Subjects received a beverage volume of 3 mL·kg−1 every 30 min during recovery to ensure a given dose of 1.2 g·kg−1·h−1 CHO (CHO), 1.2 g·kg−1·h−1 CHO with 0.2 g·kg−1·h−1 protein hydrolysate and 0.1 g·kg−1·h−1 leucine (CHO + PRO), or 1.2 g·kg−1·h−1 CHO with 1.7 mg·kg−1·h−1 caffeine (CHO + CAF). To all test drinks, 0.32 g·L−1 [U-13C6]-labeled glucose was added. The CHO source consisted of 50% glucose and 50% maltodextrin (AVEBE, Veendam, The Netherlands). The casein protein hydrolysate (PeptoPro, 85.3% protein) was prepared by DSM Food Specialties (Delft, The Netherlands). Leucine was obtained from Ajinomoto (Tokyo, Japan), caffeine was from Fagron (Nieuwekerk a/d IJssel, The Netherlands), and [U-13C6] glucose was from Cambridge Isotope Laboratories, Inc. (Andover, MA). To make the taste comparable, all solutions were flavored by adding 0.05 g·L−1 sodium saccharinate, 0.9 g·L−1 citric acid, and 5.0 g·L−1 cream vanilla flavor (Givaudan Nederland B.V., Barneveld, The Netherlands). Treatments were performed in a randomized order, with test drinks provided in a double-blind fashion.
Muscle biopsies were obtained from the middle region of the vastus lateralis muscle (∼15 cm above the patella) and approximately 2 cm below the entry through the fascia using the percutaneous needle biopsy technique described by Bergstrom et al. (5). All samples were carefully freed from any visible adipose tissue and blood, immediately frozen in liquid nitrogen (biochemistry) or in liquid nitrogen–cooled isopentane (histochemistry), and stored at −80°C until subsequent analysis.
Blood samples (8 mL) were collected in EDTA-containing tubes and centrifuged at 1000g and 4°C for 10 min. Aliquots of plasma were frozen in liquid nitrogen and stored at −80°C until analysis. Plasma glucose (Uni Kit III, 07367204; Roche, Basel, Switzerland), lactate (18), and free fatty acids (FFA, NEFA-C; Wako Chemicals, Neuss, Germany) concentrations were analyzed with a COBAS-FARA semiautomatic analyzer (Roche). Insulin was analyzed by radioimmunoassay (Linco, Human Insulin RIA kit; LINCO Research, Inc., St. Charles, MO). Concentrations of plasma catecholamines (39), sampled with heparin and put into glutathione-containing tubes on ice, and plasma caffeine (Clin Rep Komplettkit für Theophyllin, Theobromin und Coffein; Recipe Chemical + Instruments Labortechnik, Munich, Germany) were determined by using high-performance liquid chromatography (HPLC). Plasma [U-13C6] glucose enrichment was determined using the method of Pickert et al. (32), modified for use with gas chromatography–combustion–IRMS (GC-C-IRMS). Briefly, plasma samples were extracted with methanol–chloroform (2.3:1, v/v) and then chloroform–water (pH 2.0) (2:1, v/v). After drying under nitrogen gas, samples were derivatized to butylboronic acid–acetate, using butylboronic acid as a derivatizing agent. The glucose derivative (1 μL) was injected into the GC (split ratio 1:15) and analyzed by GC-C-IRMS (GC, Trace GC Ultra; C, GC Combustion III; IRMS, Delta Plus XP; all Thermo Finnigan, Herts, UK). The measured 13C enrichment was corrected by a factor of 16/6 to account for isotopic carbon dilution from the butlyboronic acid–acetyl derivative. Plasma [U-13C6] glucose enrichments are expressed as tracer-to-tracee ratios.
Muscle tissue, ∼25 mg w.w., was freeze-dried after which collagen, blood, and other nonmuscle fiber material were removed from the muscle fibers under a microscope. The isolated muscle fiber mass (∼5 mg) was weighed, and 1000 μL of 1 M HCl was added. After heating for 3 h at 100°C to hydrolyze the glycogen to glycosyl units and cooling down to room temperature, 400 μL of the solution was neutralized by adding 250 μL of Tris–KOH (precise amount of Tris–KOH is determined by titration to pH 7.0). Thereafter, 150 μL of this solution was analyzed for glucose concentration (Uni Kit III, 07367204; Roche) with a COBAS FARA semiautomatic analyzer (Roche).
Multiple serial sections (5 μm) from biopsy samples collected immediately after (t = 0 min) and 6 h after exercise (t = 360 min) were thaw mounted on uncoated, precleaned glass slides for each subject. The antibody against laminin was used to select individual muscle fibers and the antibody against human myosin heavy chain I to differentiate between the Type I and II muscle fibers. To assess intramyocellular glycogen content, we used the modified PAS stain as described previously (38). After 24 h, glass slides were examined using a Nikon E800 fluorescence microscope (Uvikon, Bunnik, The Netherlands) coupled to a Basler A113 C progressive scan color CCD camera, with a Bayer color filter. Epifluorescence signal was recorded using an (fluorescein isothiocyanate) excitation filter (465–495 nm) for laminin, a 4′,6-diamidino-2-phenylindole UV excitation filter (340–380 nm) for nuclear staining, and Texas red excitation filter (540–580 nm) for muscle fiber type. Digitally captured images were processed and analyzed using Lucia 4.8 software (Nikon, Düsseldorf, Germany). PAS-stained sections were captured in full color using bright-field light microscopy. The PAS signal was recorded for each muscle fiber, resulting in a total of 123 ± 4 muscle fibers analyzed for each muscle cross section (85 ± 3 Type I and 38 ± 2 Type II muscle fibers). The bright-field images of the PAS stains were converted post hoc to 8-bit grayscale values. The mean optical density of the PAS-raised signal per individual fiber was determined by averaging the optical density measured in every pixel in the cell, corrected for the mean optical density of the background stain measured in a field-of-view containing no muscle fibers. Mixed muscle glycogen content, as determined by PAS staining, has previously been shown to correlate well with muscle glycogen content measured using the biochemical assay (38).
All data are expressed as means ± SEM. The plasma insulin and glucose responses were calculated as area under the curve. A two-factor repeated-measures ANOVA with time and treatment as factors was used to compare differences between treatments over time. In case of significant F-ratios, Bonferroni post hoc tests were applied to locate the differences. For non–time-dependent variables, a paired Student’s t-test was used to compare differences between treatment and control. The results from the questionnaires were analyzed by the Friedman nonparametric test. Statistical significance was set at P < 0.05. All calculations were performed using SPSS Statistics 15.0 (SPSS, Inc., Chicago, IL).
Glycogen depletion protocol.
Maximal workload capacity measured during the pretesting averaged 387 ± 11 W (5.4 ± 0.1 W·kg−1). Consequently, average workload settings in the depletion protocol were 194 ± 5, 271 ± 8, 310 ± 9, and 349 ± 10 W at 50%, 70%, 80%, and 90% Wmax, respectively. On average, subjects cycled a total of 23 ± 3, 23 ± 2, and 23 ± 2 high-intensity blocks, which resulted in a total cycling time of 106 ± 10, 104 ± 10, and 106 ± 10 min in the CHO, CHO + PRO, and CHO + CAF experiments, respectively. Total cycling time did not differ between experiments.
Drink ingestion and gastrointestinal complaints.
All drinks were well tolerated by the subjects, although some subjects had difficulty ingesting the last two test drinks. Nonetheless, 95% ± 2% of the total volume of test drink (2.6 ± 0.9 L) was ingested, with no differences between treatments. The main complaints according to the questionnaires were bloated feeling, belching, and the urge to urinate. There were no differences between experiments, although the taste of the CHO + PRO drink was rated significantly lower than the CHO and CHO + CAF drinks (2.5 ± 0.2, 5.1 ± 0.3, and 4.5 ± 0.4, respectively; P < 0.01).
In all experiments, plasma insulin concentrations increased during the first 90 min of postexercise recovery, after which concentrations plateaued (Fig. 1A). Plasma insulin concentrations were higher in the CHO + PRO compared with CHO treatment (P < 0.05; Fig. 1B). Plasma glucose concentrations increased during the first 60 min of postexercise recovery, after which concentrations declined to baseline levels (Fig. 2A). Plasma glucose responses did not differ between experiments (Fig. 2B). Plasma [U-13C6] glucose enrichments are shown in Figure 3. No differences in plasma [U-13C6] glucose enrichments were observed between treatments.
Plasma lactate, FFA, adrenalin, and noradrenalin concentrations are presented in Table 1. Plasma lactate concentrations over time were lower in CHO + PRO compared with CHO (P = 0.012). There were no differences between CHO and CHO + CAF. Plasma FFA concentrations over time were higher in CHO + CAF compared with CHO (P < 0.01). There were no differences between CHO and CHO + PRO. Plasma adrenalin concentrations over time did not differ between experiments. Plasma noradrenalin concentrations over time were lower in CHO + PRO compared with CHO. There were no differences in plasma noradrenalin concentrations over time between the CHO and CHO + CAF experiment.
Plasma caffeine concentrations increased to 4.7 ± 0.2 and 9.3 ± 0.5 mg·L−1 or 24.3 ± 1.1 and 47.9 ± 2.4 μmol·L−1 at t = 180 and t = 360 min, respectively (P < 0.01), in the CHO + CAF experiment. Caffeine concentrations were below the detection limit (<0.05 mg·L−1) in the CHO and CHO + PRO experiments.
Postexercise muscle glycogen concentrations did not differ between experiments and averaged 172 ± 40, 184 ± 32, and 138 ± 22 mmol·kg−1 dry weight (d.w.) in the CHO, CHO + PRO, and CHO + CAF treatments, respectively. After 6 h of postexercise recovery, muscle glycogen concentrations had increased to 360 ± 31, 384 ± 25, and 326 ± 24 mmol·kg−1 d.w., respectively (NS). Glycogen resynthesis rates are shown in Figure 4. Histochemical analyses of the muscle biopsies revealed no differences in muscle glycogen content between experiments or between Type I and Type II fibers (Fig. 5). There was a strong significant correlation between mixed muscle glycogen content, as determined by PAS staining, and muscle glycogen content measured using the biochemical assay, with a Pearson correlation coefficient of 0.84 (P < 0.01).
The present study shows that coingesting 0.3 g·kg−1·h−1 of an insulinotropic protein plus leucine mixture or 1.7 mg·kg−1·h−1 caffeine with 1.2 g·kg−1·h−1 CHO does not further accelerate postexercise muscle glycogen synthesis when compared with the ingestion of 1.2 g·kg−1·h−1 CHO only.
The rate of postexercise muscle glycogen synthesis depends on numerous factors, which include the magnitude of muscle glycogen depletion, the amount of CHO ingestion, the rate of gastric emptying and intestinal glucose uptake, and the insulin-stimulated uptake of glucose and subsequent conversion to muscle glycogen by glycogen synthase (3,22).
The exercise protocol in our study resulted in postexercise muscle glycogen concentrations that did not differ between treatments (i.e., 172 ± 40, 184 ± 32, and 138 ± 22 mmol·kg−1 d.w. in the CHO, CHO + PRO, and CHO + CAF treatments, respectively). The postexercise muscle glycogen levels are comparable to values published previously applying similar exercise protocols to strongly reduce muscle glycogen levels (23,47). Muscle glycogen synthesis has been shown to occur in two distinct phases (15,28,34,36), with the first insulin-independent phase being characterized by higher muscle glycogen synthesis rates. Several studies have shown that this fast phase of muscle glycogen synthesis occurs when concentrations are reduced to less than 150 mmol·kg−1 d.w. (22,28,30,34). In our study, postexercise muscle glycogen concentrations were below this level in the CHO + CAF experiment only, but this did not result in higher muscle glycogen synthesis rates during the 6-h recovery phase.
Coingestion of protein, protein hydrolysates, and/or free amino acids with CHO has been shown to strongly augment postprandial insulin release (24,35,46,48). In the present study, we confirm the insulinotropic potential of protein and free leucine coingestion, with a 76% ± 11% greater insulin response in the CHO + PRO versus CHO experiment. However, the greater insulin response did not augment postexercise muscle glycogen synthesis. The latter seems to be in agreement with three other studies (19,23,44) that reported no additional benefits of protein coingestion on postexercise muscle glycogen synthesis when much CHO were administered during relatively short recovery periods of 3 and 4 h. Because glucose uptake and glycogen synthesis become more insulin dependent throughout the latter stages of postexercise recovery (22,34), we hypothesized that the glycogen-stimulating effect of protein coingestion with an optimal amount of CHO would become apparent after a more extended recovery period of 6 h. In contrast to our hypothesis, data from the present study indicate that circulating insulin levels are not the rate-limiting factor for muscle glycogen synthesis when postexercise CHO intake is optimal (1.2 g·kg−1·h−1).
From the present findings, we conclude that further elevation of circulation insulin levels does not accelerate muscle glycogen synthesis during postexercise recovery. Nonetheless, it could be speculated that stimulating insulin release could still be beneficial for other aspects of recovery. Although recent studies have shown that increasing insulin does not further stimulate muscle protein synthesis when ample protein is ingested (16,25,40), it may inhibit exercise-induced muscle protein breakdown. The latter might improve net protein balance and, as such, improve postexercise muscle reconditioning. Furthermore, elevating postexercise insulin levels might accelerate the replenishment of liver glycogen stores, which is an aspect of postexercise recovery that has received much less attention (11,14).
It might be speculated that the increased osmolality and energy density of the CHO + PRO beverage delayed gastric emptying, thereby decreasing the delivery of exogenous CHO for muscle glycogen synthesis. However, in the present study, the coingestion of protein or caffeine did not result in differences in exogenous [U-13C6] glucose appearance, suggesting that gastric emptying rates did not differ between treatments.
Recently, Pedersen et al. (31) showed a 66% greater increase in muscle glycogen synthesis rate during postexercise recovery when caffeine was coingested with CHO. The latter seems at odds with previous work in which no stimulating effect of caffeine supplementation on muscle glycogen synthesis has been observed (2). However, in the latter study, subjects were administered caffeine before and during exercise, whereas the CHO-containing beverages were provided during postexercise recovery. Consequently, it could be suggested that caffeine can only exert its effect on muscle glycogen synthesis when coingested with the CHO supplements. The mechanism by which caffeine coingestion might accelerate postexercise muscle glycogen synthesis remains unclear. In resting conditions, caffeine ingestion has been shown to actually impair insulin-mediated glucose disposal (1,17,29,33,43), likely attributed to β-adrenergic stimulation and/or adenosine receptor antagonism (1,43). However, exercise seems to attenuate these detrimental effects of caffeine on insulin action in skeletal muscle tissue (42). Furthermore, studies that were performed during exercise suggest that caffeine coingestion with CHO can stimulate intestinal glucose absorption by stimulating the jejunal sodium–glucose transporter protein SGLT1 via cAMP (49,51).
To assess the potential stimulating properties of caffeine coingestion on the intestinal uptake of ingested glucose during postexercise recovery, we added a [U-13C6]-labeled glucose tracer to the beverages. The increase in plasma [U-13C6] glucose enrichments did not differ between treatments, implying that there were no differences in exogenous glucose appearance rates between treatments. Consequently, our findings do not support the suggestion that caffeine coingestion stimulates intestinal glucose absorption during postexercise recovery. In accordance, caffeine coingestion did not accelerate muscle glycogen synthesis rates. There is no clear explanation for the differences between our findings and the recent work by Pedersen et al. (31). The apparent discrepancy may be attributed to differences in study design. In the present study, we applied the exhaustive exercise session in the morning after an overnight fast, whereas Pedersen et al. included an additional exercise session the evening before the experimental day followed by the ingestion of a low-CHO diet. Consequently, postexercise muscle glycogen levels were generally lower in the study by Pedersen et al. when compared with the present findings. However, these differences would unlikely modulate the proposed efficacy of caffeine to accelerate intestinal glucose absorption or subsequent muscle glycogen synthesis. Furthermore, there were distinct differences in CHO supplementation regimen between studies. Pedersen et al. provided their subjects with 1.0 g·kg−1·h−1 CHO with/without 2.0 mg·kg−1·h−1 caffeine at 2-h intervals during a 4-h recovery period. In contrast, we provided subjects with 1.2 g·kg−1·h−1 CHO with or without 1.7 mg·kg−1·h−1 caffeine via boluses provided every 30 min during a more extensive 6-h recovery period. A more frequent provision of CHO has generally shown to maximize the rate of postexercise glycogen repletion. Although the absolute amount of caffeine that was provided during the recovery phase was similar between studies, Pedersen et al. provided their supplements at 0 and 2 h of postexercise recovery. The latter might explain the slightly higher plasma caffeine concentrations in their study. However, this is unlikely responsible for the ∼70% greater glycogen resynthesis rate in the CHO plus caffeine treatment. So far, there is little evidence and/or mechanistic background to suggest that caffeine coingestion after exercise will accelerate postexercise muscle glycogen synthesis when ample amounts of CHO are provided at (more) frequent intervals.
After 6 h of postexercise recovery, muscle glycogen concentrations had increased to 360 ± 31, 384 ± 25, and 326 ± 24 mmol·kg−1 d.w. in the CHO, CHO + PRO, and CHO + CAF experiments, respectively. We did not measure basal muscle glycogen concentrations, but previous studies have reported preexercise values in athletes to range between 500 and 600 mmol·kg−1 d.w. (9,13). The latter implies that muscle glycogen stores cannot entirely be replenished within 6 h of postexercise recovery even when ingesting much CHO with or without additional protein or caffeine. Moreover, for athletes training or competing twice daily or recovering overnight, it is not always feasible to ingest such substantial amounts of CHO. It has previously been shown that the coingestion of protein with a smaller amount of CHO stimulates muscle glycogen synthesis to the same extent when compared to the ingestion of a greater amount of CHO (47). Based on those findings, it was speculated that the greater postprandial insulin response after protein coingestion facilitates the uptake of the ingested CHO in the more insulin-sensitive tissues, like skeletal muscle. Therefore, under circumstances where the total amount of CHO that can be ingested is limited, it might be preferred to coingest some protein to facilitate glucose uptake in muscle. However, more work is warranted to determine the effect of a greater postprandial insulin response on glucose uptake and glycogen deposition in various tissues. Furthermore, the coingestion of caffeine during short-term recovery might enhance subsequent exercise performance via its stimulating effect on neuromuscular function.
In summary, we show that coingestion of 0.3 g·kg−1·h−1 protein with 1.2 g·kg−1·h−1 CHO during postexercise recovery strongly increases insulin release but does not further accelerate muscle glycogen synthesis. Furthermore, coingestion of 1.7 mg·kg−1·h−1 caffeine with 1.2 g·kg−1·h−1 CHO has no effect on the uptake of ingested glucose from the gut and does not increase postexercise muscle glycogen synthesis. We conclude that coingestion of an insulinotropic amino acid/protein mixture or caffeine does not further accelerate postexercise muscle glycogen synthesis when an optimal amount of CHO (1.2 g·kg−1·h−1) is already ingested.
This study did not receive any funding from governmental or nongovernmental organizations.
The authors are grateful for the assistance of Ernst Fluitman, Antoine Zorenc, and Jos Stegen and the enthusiastic support of the subjects who volunteered to participate in these experiments.
M.B. and L.J.C.v.L. designed the study. M.B. organized and carried out the clinical experiments. J.v.K. and J.M.S. performed the stable isotope tracer analyses. M.B. performed the statistical analysis and wrote the manuscript together with H.K. and L.J.C.v.L.
None of the authors has any conflict of interest with companies or manufacturers that will benefit from the results of the present study.
The results of the present study do not constitute endorsement by the American College of Sports Medicine.
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Keywords:©2012The American College of Sports Medicine
INSULIN; LEUCINE; GLYCOGEN SYNTHASE; AMINO ACIDS; 13C-GLUCOSE