The liver plays a crucial role in preventing hypoglycemia during exercise (27), and it is generally believed that strategies that enhance liver glycogen after exercise will increase exercise capacity in a subsequent exercise bout (6). Most studies have investigated the role of muscle glycogen after exercise (reviewed in Beelen et al. (1) and Jentjens and Jeukendrup (13)), but very few studies have focused on the potentially very important role of the substrates in the liver. Evidence from early rodent studies suggests that when CHO are available after exercise, liver glycogen resynthesis is the first priority and muscle glycogen synthesis is secondary (7). Only one study has investigated liver glycogen synthesis after exercise in healthy humans. In this study, it was found that a single feeding of glucose or fructose (1 g·kg−1 body mass) was sufficient to initiate liver glycogen synthesis after exercise, without affecting muscle glycogen synthesis (6).
Fructose has been shown to have different effects on liver glycogen metabolism than glucose (22). The bulk of an intravenous or oral fructose load is taken up by the liver and converted to glucose (27). However, although gluconeogenesis is markedly increased with fructose ingestion, this does not result in an increased hepatic glucose output, suggesting that fructose could be stimulating hepatic glycogen synthesis, reducing hepatic glycogenolysis, or both. In line with this, small amounts of intraduodenal or intraportally infused fructose given concurrently with a glucose load stimulated the synthesis and storage of liver glycogen and reduced postprandial hyperglycemia and hyperinsulinemia in dogs (32). There is also evidence that small amounts of fructose stimulate glucokinase and glycogen synthase in the liver, the two rate-limiting enzymes of liver glycogen synthesis (22). Combinations of multiple transportable CHO (i.e., glucose and fructose) have also been shown to result in higher exogenous CHO oxidation rates during exercise (14,15,37); for review, see Jeukendrup (16), suggesting better CHO absorption than from a similar amount of glucose only. Very high exogenous CHO oxidation rates were reported for a mixture of maltodextrins and fructose (37). Wallis et al. (36) suggested that this more rapid CHO delivery could also help postexercise muscle glycogen synthesis, but this did not seem to be the case. Muscle glycogen synthesis rates were equally high with maltodextrin-fructose compared with isoenergetic maltodextrin. One explanation could be that most of the fructose was directed toward liver glycogen synthesis and not muscle glycogen.
Another CHO that could be beneficial for liver glycogen synthesis is galactose. The liver is the major site of galactose uptake and metabolism in humans and galactose clearance from the blood after a galactose load has been used to assess the general functional capacity of this organ. In the liver, galactose can be converted to glucose and subsequently stored as glycogen or immediately released into the circulation. In rats, formation of glycogen accounts for the great majority of galactose taken up by the liver (26). Accordingly, a study with isolated perfused rat liver (34) showed that galactose stimulated liver glycogen synthesis in the presence of glucose with a concomitant increase in the activity of glycogen synthase and a decrease in the activity of glycogen phosphorylase. However, animal and in vitro studies (25,38) have shown that the infusion or ingestion of galactose results in lower liver glycogen synthesis rates compared with after the administration of glucose.
Therefore, combinations of glucose with either fructose or galactose could prove to be attractive possibilities when the aim is to amplify liver glycogen resynthesis in humans after a glycogen-depleting exercise. To our knowledge, this has not been done before. However, energy-dense glucose solutions are hypertonic and may interfere with gastrointestinal (GI) fluid delivery. Maltodextrin (MD), a glucose polymer, tastes less sweet than glucose and has a lower osmolality. Because gastric emptying (33) and metabolic availability (30,37) of glucose polymer drinks are not less than for glucose drinks, MD was used for delivering much energy per unit time. In the present work, we used magnetic resonance (MR) techniques, which have made it possible to noninvasively quantify hepatic glycogen metabolism in humans, to investigate the hypothesis that the postexercise ingestion of MD + fructose or MD + galactose results in an enhanced liver glycogen synthesis compared with the ingestion of a matched iso-osmolar and isocaloric MD + glucose control.
The study was approved by the local ethics committee and conformed to the Declaration of Helsinki. Ten healthy male endurance-trained volunteers gave their written informed consent to participate. They were characterized as follows: height = 184 ± 2 cm, weight = 74 ± 2 kg, age = 29 ± 1 yr, body mass index = 22 ± 1 kg·m−2, aerobic capacity (V˙O2max) = 64 ± 1 mL·kg−1·min−1, and maximum power (Wmax) = 373 ± 14 W (mean ± SEM).
This was a double-blind, triple crossover, randomized clinical trial (recruitment started December 1, 2005). After preliminary testing, each subject completed three experimental trials on separate days at least 1 wk apart. After cycling exercise to exhaustion to deplete body glycogen stores, liver glycogen content was evaluated using 13C MR spectroscopy (MRS). This was followed by a 6.5-h recovery period in which subjects consumed one of three CHO drinks (Latin square design) containing a combination of two-thirds of maltodextrin (MD) and one-third of fructose (FRU drink), galactose (GAL drink), or glucose (GLU drink). During recovery, liver glycogen was measured approximately every 2 h; blood was collected for analysis of insulin, glucose, and free fatty acids (FFA); and a questionnaire on GI comfort and sensations was administered (details in the "Experimental trials" section and in Fig. 1).
Before study commencement, subjects underwent a medical examination where anthropometric measurements were obtained. Subsequently, an aerobic capacity test was performed using a graded exercise protocol to volitional exhaustion on an electromagnetically braked bicycle ergometer (Excalibur Sport; Lode, Groningen, The Netherlands) to evaluate V˙O2max and Wmax, as described elsewhere (37) with the following modifications. The exercise test started at a work intensity of 100 W for 5 min, after which intensity was increased by 30 W every 2 min until the volunteer could no longer maintain the required pedaling frequency (>60 rpm). HR was recorded continuously using radiotelemetry (Polar Vantage NV; Polar Electro Ltd., Kempele, Finland), whereas oxygen consumption and carbon dioxide production were monitored continuously throughout the test using an online automated gas analysis system (Oxycon Alpha; Jaeger-Toennies, Höchberg, Germany).
Test drinks were supplied to the experimenter as powder in blinded sachets (one sachet = a 45-g CHO dose). The MD-to-monosaccharide ratio was analyzed in the sample sachets (Nestlé Quality Assurance Laboratory, Croydon, UK) to be 2:1 for each of the FRU, GAL, and GLU mixes. Maltodextrin Glucidex with dextrose equivalent DE12 was obtained from Roquette Frères (Lestrem, France), crystalline fructose Fruitose S from Galam Ltd. (via Sugro AG, Basel, Switzerland), d-(+)-galactose from Aldrich (Fluka Chemie GmbH, Buchs, Switzerland), and d-(+)-glucose (Roferose ST) from Roquette. On the day of each experimental trial, 10 sachets (450 g) were dissolved in 3 L of water (15% solution) plus some drops of lemon juice to maximize the palatability for each volunteer.
Subjects kept a dietary and training log 2 d before the first experimental trial. For the subsequent two trials, they were asked to repeat the recorded activities and dietary patterns. In particular, they refrained from strenuous training and had no alcohol consumption the day before a trial. Subjects attended the laboratory in the morning after an overnight fast to undergo a standardized cycling exercise protocol to deplete glycogen stores (19). The exercise started with a 10 min warm-up at 50% Wmax. Thereafter, the fasted participants cycled for 2-min periods alternating between 90% and 50% Wmax, until they were no longer able to complete 2 min at 90%. At this point, the high-exercise block was reduced to 80% Wmax, and the alternating process continued until this intensity could no longer be maintained, after which the high-intensity block was decreased to 70% Wmax. When the volunteers were no longer able to switch between 70% and 50% at 2-min intervals, they were motivated to maintain the 50% level until complete exhaustion. Water was provided ad libitum during the exercise protocol. After exhaustion, participants took a brief shower and returned to the laboratory within 30 min. While supine, a catheter was inserted into an antecubital vein of the forearm to allow for repeated blood sampling. At this time, subjects replied to the questionnaire to evaluate baseline GI sensations. Subsequently, they underwent a basal MR measurement for approximately 45 min (Fig. 1), followed by the initial resting blood sample (7 mL), and the ingestion of the first drink at T0 (T0 = zero time). Nine further blood samples and GI questionnaires were collected or administered, respectively, at 45, 85, 140, 185, 220, 275, 320, 355, and 405 min (Fig. 1). The irregular time sequence of blood draws was related to time constraints of the MR measurements because blood sampling could not be done concurrently. T0 was the time of the end of the baseline MR measurement. For the three subsequent measurements, T1 (124 min), T2 (258 min), and T3 (392 min), the times were taken half-way during the acquisition of MR spectra (SD = 4 min for the precision of T times).
Test drinks were consumed at the same times as the first eight blood draws (0-320 min). A single dose (45 g, 1 × 300 mL) was given, except at 0 and 135 min, where two doses (2 × 300 mL) were given (Fig. 1). This 2-1-1 schedule was chosen in an attempt to catch up for the time spent in the magnet, where drinking was not possible. Therefore, the effective rate of CHO and fluid intake was 1.5 g·min−1 (90 g·h−1) and 600 mL·h−1 during the first 4 h, but it was 1.15 g·min−1 and 462 mL·h−1 on average during the 6.5-h recovery until the last MR measurement at T3.
Magnetic resonance measurements.
MR images and 1H-decoupled natural abundance 13C-MR spectra were recorded on a standard 1.5-T GE SIGNA system (General Electric, Milwaukee, WI) equipped with a home-built decoupler console for decoupling and nuclear Overhauser effect build up, as described previously (4,9). The body coil was used to obtain MR images for positioning and volume determination, whereas a double-tuned flexible surface coil (Medical Advance, Milwaukee, WI) was used for 13C signal excitation and reception (square surface coil of 11.3 cm × 11.3 cm) and for a localizer 1H imaging and 1H decoupling (Helmholtz-type coil, flexible, approximately 16 cm). Volunteers were positioned reproducibly within a few millimeters on an examination bed that had been specifically built for a foregoing study (4) with an analogous acquisition of 13C MR spectra in the liver. In addition, the position of the right lower corner of the 12th thoracic vertebra was determined and the relative position of the 13C-coil was calculated. In the following sessions, the position of the body was adjusted relative to the right lower edge of the 12th thoracic vertebra. After fixation of the surface coil, one series in axial orientation was repeated to document any change in position during fixation of the surface coil. A typical sequence of spectra is shown in Figure 2.
Quantitation of glycogen.
Quantitation of the glycogen signal was obtained in two steps. First, careful positioning of the surface coil helped to obtain the same sensitivity distribution in all sessions for a given subject. Spectra were analyzed using jMRUI (java-based Magnetic Resonance User Interface; http://www.mrui.uab.es/mrui/mrui_Overview.shtml) with the following prior knowledge: fixed position of glycogen, line width relative to the lipid resonances, and two components with different line width (28). Variations of the signal amplification and coil loading were monitored by the simultaneous acquisition from two acetone vials fixed close to the coil center. The acetone resonance showed no systematic variations between volunteers or sessions, such that no correction for coil loading was necessary. The obtained institutional units (IU) are comparable within one volunteer, but they need an individual correction factor depending on the distance between coil and liver. Therefore, in a second step, IU were translated into absolute units (mmol·L−1; 180 g·mol−1) after measuring a 95-mmol·L−1 glycogen solution in a liver-shaped phantom at the same distance from the coil center as the liver in the volunteers. These distances had been determined by MRI in volunteers and phantom. Finally, rates of glycogen replenishment during intermediate periods (T0 to T1, T1 to T2, and T2 to T3) were calculated after correcting for the individual time intervals between measurements. The glycogen result for one GLU subject at T2 had to be discarded because of technical problems of the MR scanner.
Quantitation of liver volume.
After the acquisition of localizer images, two series of images were taken in axial (13 images, field of view = 40 cm) and coronal (16 images, field of view = 48 cm) directions with a fast-spoiled-gradient sequence (body coil, slice = 8 mm, flip angle = 60°, TE = 1.5 ms, TR = 100 ms, matrix = 512 × 512). To check for systematic effects of the slice orientation in volume determination, both series were analyzed using a newly developed tool for the evaluation of body composition (5). The determination of liver volumes in the two slice directions showed excellent agreement (r = 0.957, slope = 0.88-0.98, 95% confidence interval); therefore, the average of the two measurements was used.
The GI questionnaire consisted of 16 questions. Each question started with "Have you suffered from…?" and was answered by ticking a box on a 10-point scale ranging from 1 (not at all) to 10 (very, very much). Six questions related to upper GI symptoms (general stomach problems, nausea, urge to vomit, belching, heartburn, cramps in the stomach), four questions related to lower GI symptoms (flatulence, an urge to defecate, cramps in the intestine, diarrhea), and six questions related to central or other symptoms (dizziness, headache, an urge to urinate, a bloated feeling, side aches left, side aches right).
Blood was drawn into prechilled tubes and centrifuged. Aliquots of plasma were stored at −20°C until analysis for glucose (enzymatic; Roche Diagnostics, Berthoud, Switzerland) and FFA (enzymatic colorimetric; Wako Bioproducts, Richmond, VA) using an autoanalyzer (XPAND; Dade Behring, Inc., Eschborn, Germany). Plasma insulin was determined by ELISA (IBL; Immuno-Biological Laboratories, Hamburg, Germany). Plasma residues have been used for a metabolomics analysis (published elsewhere) that was not related to the objectives of this study.
Statistical analysis of glycogen data was performed with SPSS 18 for Windows (SPSS, Inc., Chicago, IL) using general linear models (GLM), one-sample tests to compare means against zero, and a nonparametric Friedman test to evaluate the responses to the questionnaires on GI sensations. GLM were with fixed factors for treatments, random factors for volunteers, and main interactions. Pairwise comparisons were Bonferroni corrected for multiple tests. Linear regression and the calculation of slopes for determining replenishment rates were performed in Microsoft Excel 2007 (Microsoft Corp., Redmond, WA). One single glycogen result of 120 was missing because of low spectral quality (second postdepletion value for one volunteer with one drink) and was, therefore, not used for the calculation of the slope. A GLM for repeated measures over time was applied to blood concentrations of glucose, insulin, and FFA data, which were log-transformed to achieve approximate normal distribution of the residues. GLM was followed by tests of the within-subject contrasts for treatment × time. Statistical significance was accepted at P < 0.05. The Friedman tests for the 16 questionnaires were Bonferroni corrected such that a significance level of 0.003 was required. Data are presented as means ± SEM if not stated otherwise.
The glycogen depletion ride to exhaustion lasted approximately 2 h and was not different between treatments (FRU = 119 ± 2 min, GAL = 116 ± 2 min, and GLU = 118 ± 3 min, NS). The total workload of 1564 kJ (CV = 18%) corresponded to 59% of Wmax (CV = 6%) on average (weighted mean of the alternating intensities).
Values at depletion.
At exhaustion, the liver glycogen concentration was 99 mmol·L−1 (CV = 49%), without significant differences between treatments (FRU = 97 ± 16 mmol·L−1, GAL = 82 ± 16 mmol·L−1, and GLU = 117 ± 13 mmol·L−1, NS). Liver volume was 1598 mL (CV = 13%), with a trend (P = 0.05) to slight differences between treatments (FRU = 1648 ± 70 mL, GAL = 1540 ± 61 mL, and GLU = 1606 ± 64 mL). Total liver glycogen content before replenishment was 29.2 g (CV = 58%), without significant differences between treatments (FRU = 29.6 ± 5.9 g, GAL = 23.6 ± 5.5 g, and GLU = 34.3 ± 4.5 g, NS).
Liver glycogen and volume rates.
During recovery, liver glycogen concentration increased consistently in all volunteers (Fig. 3). An analysis of replenishment rates for glycogen concentrations found an overall effect of the treatments (P < 0.001) and of the subjects (P = 0.002). Furthermore, rates were significantly higher with FRU (24 ± 2 mmol·L−1·h−1) and GAL (28 ± 3 mmol·L−1·h−1) compared with GLU (13 ± 2 mmol·L−1·h−1). Pairwise tests resulted in the following: FRU versus GAL, P = NS; FRU versus GLU, P < 0.001; and GAL versus GLU, P < 0.001.
Similarly, during replenishment, liver volume significantly increased after FRU (24 ± 4 mL·h−1, P < 0.001) and GAL (22 ± 3 mL·h−1, P < 0.001), but liver volume remained unchanged after GLU (3 ± 3 mL·h−1, NS; Fig. 4).
Total glycogen content (i.e., glycogen concentrations multiplied by the respective liver volumes) increased significantly after all three drinks (time effect, P < 0.001) with FRU (8.1 ± 0.6 g·h−1), GAL (8.6 ± 0.9 g·h−1), and GLU (3.7 ± 0.5 g·h−1) (Fig. 5); however, FRU and GAL led to significantly higher replenishment than GLU (pairwise tests resulted in FRU vs GAL, P = NS; FRU vs GLU, P < 0.001; and GAL vs GLU, P < 0.001). The increase in liver glycogen content seemed linear for all three treatments for the group averages (FRU r2 = 0.964, GAL r2 = 0.976, GLU r2 = 0.886).
Absolute changes in liver glycogen and volume.
The increase in glycogen concentrations within the duration of the experiment (approximately 6.5 h) was significantly different between treatments (P < 0.001) and between subjects (P = 0.005). The increase was larger after FRU (154 ± 14 mmol·L−1) and GAL (179 ± 22 mmol·L−1) than after GLU (77 ± 10 mmol·L−1). Pairwise tests resulted in the following: FRU versus GAL, P = NS; FRU versus GLU, P < 0.001; and GAL versus GLU, P = 0.001.
At the end of the recovery period, changes in the liver volumes were significantly different between treatments (P < 0.001) and between subjects (P < 0.001). An increase occurred after FRU (153 ± 27 mL, P < 0.001) and GAL (152 ± 22 mL, P < 0.001) but not after GLU (35 ± 21 mL, NS).
The increase in the glycogen content, i.e., differences in glycogen concentrations multiplied by the respective liver volumes, was significantly different between treatments (P < 0.001) and between subjects (P = 0.004). It reached 52 ± 4 g after FRU and 56 ± 6 g after GAL, as against 23 ± 3 g after GLU.
Plasma concentrations of glucose and insulin during recovery are shown in Figure 6. In addition to the obvious changes of plasma glucose and plasma insulin with time (P < 0.001), the time trends of plasma glucose and insulin were also significantly different for the three treatments (treatment × time interaction terms, P < 0.001, adjusted for sphericity by the Huynh-Feldt correction). A noticeable effect of the treatment was detected by the tests for within-subject contrasts at time 10 (P < 0.001) where FRU changed the plasma parameters in 3 of the 10 volunteers significantly. Plasma FFA concentrations decreased in response to CHO ingestion from the resting values (FRU = 0.74 ± 0.15 mmol·L−1, GAL = 0.87 ± 0.05 mmol·L−1, GLU = 0.83 ± 0.05 mmol·L−1) to <0.05 mmol·L−1 at 2 h and onward (time main effect, P < 0.001; time × drink interaction, P < 0.001). Tests for within-subject contrasts showed a significant difference at time 10 (P < 0.001), with FRU higher than GLU (0.12 ± 0.06 vs 0.01 ± 0.01 mmol·L−1).
A main effect of the drinks was observed in association with the upper GI tract. Scores during the consumption of GAL were significantly (P < 0.003 required to correct for multiple tests) higher than with the other drinks for general stomach problems (P = 0.002), heartburn (P = 0.001), and stomach cramps (P < 0.001); however, these symptoms remained very modest, with average group maxima not exceeding 1.3 on the 1 to 10 scale at any time point after ingestion of the drinks.
On examination of the time course of the complaints, minor dizziness was reported between exhaustion and consumption of the first drink. Some headaches were observed after exhaustion, which remained stable during the replenishment period, independent of the drinks. The obvious symptom was the occurrence of an urge to urinate, with two peaks coincident with the periods of confinement in the magnet after the consumption of four boluses of a 300-mL drink within 1.5 h (1.2 L).
Liver glycogen resynthesis was measured in male cyclists who were fed different CHO mixes during recovery after exhaustive exercise. Our major finding was a doubling of liver glycogen deposition with combinations of MD + fructose or MD + galactose compared with an isoenergetic, iso-osmotic combination of MD + glucose.
Using the biopsy technique, early investigators observed that fructose infusion resulted in greater increases in liver glycogen than glucose (2). More recently using 13C MR, Casey et al. (6) fed subjects glucose, sucrose (76 g, one bolus), or a placebo and found only a small net liver glycogen resynthesis during a 4-h period and no difference between glucose and sucrose. The most significant differences between our study and the previous ones (6) are the use of CHO mixes and the larger doses, maximizing the intestinal CHO transport, which resulted in considerable differences in net liver glycogen synthesis rates.
The enhancing effect of FRU on hepatic glycogen synthesis may be partially explained at the level of the intestinal absorption, subsequently to the liver. Postprandially, glucose absorption is increased through the recruitment of the facilitative glucose transporter GLUT2 in addition to the active transporter SGLT1 (18). With the high level of CHO ingestion used in this study (90 g·h−1 in the first 4 h), glucose absorption capacity would have reached saturation (14,17), resulting in submaximal delivery to the liver. In contrast, the intestinal absorption of fructose is GLUT5 dependent and it is readily saturated when fed in isolation, but it is facilitated by the simultaneous ingestion of glucose (35). High consumption (>60 g·h−1) of glucose and fructose blends, which recruit multiple transporters (SGLT1, GLUT2, and GLUT5), have previously been shown to enhance gastric emptying and fluid delivery compared with glucose-only drinks (8,17,31). Accordingly, it could be hypothesized that postexercise CHO delivery to the liver was enhanced with the FRU drink compared with GLU, resulting in increased hepatic glycogen storage. This notion seems to be supported by our observation of unique changes in plasma values with FRU at the final blood draw (85 min after the last drink). Falls in glucose and insulin concentrations (Fig. 6) and the rise in FFA concentration may be the first signals that a postabsorptive situation is reestablishing faster with FRU than with the other blends; however, because this has been observed only in 3 of the 10 volunteers, studies with more subjects would be necessary to confirm this observation.
A factor of potentially larger magnitude in enhancing liver glycogen synthesis is the differential postabsorptive fates of fructose and glucose. Glucose is a relatively poor direct substrate for liver glycogen synthesis (24,27). Much of it is released from the liver into the systemic circulation to be stored as muscle glycogen (3,7). In contrast, fructose is primarily taken up by the liver where it is either phosphorylated and converted to glycogen or metabolized to lactate and pyruvate, which can then be released to the circulation (21). In addition, fructose is a potent acute regulator of liver glucose uptake and glycogen synthesis (29,32) and suppresses net hepatic glycogenolysis (40). Using 13C MR and an euglycemic hyperinsulin clamp, Petersen et al. (29) measured hepatic glycogen synthesis with or without a low-dose fructose infusion (∼40 mg·min−1) in fasted healthy subjects. These authors observed a threefold increase in the stimulated net rate of hepatic glycogen synthesis with a 13:1 glucose-to-fructose ratio compared with glucose alone (29). This suggests that the amount of fructose that will maximize liver glycogen storage in postexercise feedings lies somewhere between the substantial amount that maximizes intestinal CHO delivery (>60 g·h−1) and a catalytic one operating at the hepatic level.
We are not aware of any previous work addressing the role of galactose on quantitative liver glycogen synthesis in humans. This seems to be the first demonstration that replenishment of liver glycogen proceeds twice as fast when one-third of glucose was replaced by galactose in a postexercise CHO drink. Although galactose and glucose are transported equally well by SGLT1 (39), we noticed a slightly higher occurrence of GI problems upstream of the pylorus with GAL (general stomach problems, burns, and cramps) compared with the other drinks. If galactose absorption were competitively inhibited when coingested with glucose (10), a duodenal feedback inhibition (23) could cause upper GI symptoms. Notwithstanding hypothetical absorptive obstacles, hepatic glycogen replenishment was higher with GAL than with GLU, indicating the dominance of hepatic factors over any intestinal limitation. The relatively poor net glycogenic activity reported for galactose when supplied alone has been associated with its weaker insulin stimulation and by urinary galactose loss (25,38). In contrast, our results are consistent with observations of an increased activation of glycogen synthase-in proportion to the rate of infused glucose and independent of insulin-and a decreased glycogen phosphorylase activity when galactose is co-perfused with larger amounts of glucose in vitro (34). This suggests that, with GAL, glucose is a major contributor to the neoformed liver glycogen.
Liver size increased during those trials leading to faster glycogen replenishment (FRU and GAL). This enlargement accounted for a minor fraction (∼9%) of glycogen storage, the bulk of the gain being due to liver glycogen concentration changes. Liver volume may change from day to day in relation to exercise, feeding, and hydration, and within a day, a mean diurnal change of 17% in liver size has been reported (20).
In practical exercise and dietary terms, muscle and liver glycogen concentrations may both determine the point of fatigue (11,12). Some information on the relative response of both liver and muscle glycogen replenishment to loading with fructose-containing drinks can be drawn by considering the present study, together with a recent study of muscle glycogen replenishment (36) using comparable study design and subject population during 4 h of recovery. In the study by Wallis et al. (36), recovery drinks were identical with FRU and GLU in the current study, except that free glucose replaced glucose polymers. No significant difference was found in postexercise muscle glycogen accumulation between the CHO treatments (36). Combining the 4-h data of both studies, quantitative estimates of glycogen storage can be derived assuming 10 kg of active muscle mass with a wet-to-dry mass ratio of 4.28 (13). Estimates of whole-body (muscle + liver) glycogen storage amount to 101 g (65 + 36 g) with the fructose-containing drink and 92 g (74 + 18 g) with the glucose-only drink. In other words, whole-body glycogen storage seems to be slightly better when fructose is present.
The effects of these findings on performance are unclear. Of course, hypoglycemia has been described as a possible cause of fatigue during prolonged endurance exercise (12), and hepatic glycogenolysis contributes significantly in the prevention of hypoglycemia. Therefore, having higher liver glycogen stores could be beneficial. CHO drinks containing fructose and galactose could help in situations where athletes have to exercise twice in 1 d with relatively little recovery. Clearly, more research needs to be done to study the possible performance effects as well as to elucidate the underlying mechanisms. Nevertheless, this is the first study in humans to clearly show that CHO blends including fructose and galactose can significantly improve postexercise liver glycogen resynthesis.
In summary, this study demonstrates that ingestion of ∼70 g·h−1 of maltodextrin + fructose (2:1) or maltodextrin + galactose (2:1) drinks consumed during the short-term postexercise recovery results in a twofold increase in the rates of liver glycogen replenishment compared with an isoenergetic, iso-osmotic maltodextrin + glucose control.
The authors thank D. Grathwohl, A. Fracheboud, C. Ammon-Zufferey, A. Blondel-Lubrano, J. Farrar, J.P. Schmid, R. Koenig, E. Schaller, and T. Stellingwerff.
This study was supported by Nestec S.A., Lausanne, Switzerland. The jMRUI software package was kindly provided by the participants of the EU Network programs: Human Capital and Mobility (CHRX-CT94-0432) and Training and Mobility of Researchers (ERB-FMRX-CT970160). Part of the salaries were funded by an SNF grant (310000-118219).
The study was instigated and financed by Nestec. J.D. and R.J. were employees of Nestec.
None of the authors had a conflict of interest to declare.
The results of the present study do not constitute endorsement by the American College of Sports Medicine.
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