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First published online February 15, 2006
Journal of Experimental Biology 209, 871-880 (2006)
Published by The Company of Biologists 2006
doi: 10.1242/jeb.02071
Fuel use during glycogenesis in rainbow trout (Oncorhynchus mykiss Walbaum) white muscle studied in vitro
Department of Biology, The University of Western Ontario, London, Ontario, Canada N6A 5B7
* Author for correspondence (e-mail: milligan{at}uwo.ca)
Accepted 4 January 2006
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Summary |
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Key words: lactate metabolism, white muscle, glycogenesis, rainbow trout, Oncorhynchus mykiss
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Introduction |
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TOPSummaryIntroductionMaterials and methodsResultsDiscussionReferences |
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; Richards et al.,
2002a; Richards et al.,
2002b). At exhaustion, glycogen, adenosine 5'-triphosphate
(ATP) and phosphocreatine (PCr) stores in working muscle are reduced and
lactate and H+ levels are elevated
(Richards et al., 2002a).
Clearance of anaerobic metabolites and the restoration of energy reserves by
muscle are necessary to prepare the fish for subsequent bouts of exercise.
The current model for the restoration of muscle energy reserves in fish
after exhaustive exercise suggests that in adult fish glycogen is
resynthesized in situ with lactate as the main substrate
(Milligan and Girard, 1993;
Wang et al., 1997;
Richards et al., 2002a).
Glucose, the main glycogenic substrate in mammalian muscle, is probably not an
important glycogenic or oxidative substrate for trout white muscle since
muscle has low hexokinase activity (Knox
et al., 1980; Storey,
1991) and the expression of glucose transporters in the muscle
membrane is extremely low and their physiological role is unclear
(Legate et al., 2001;
Teerijoki et al., 2001;
Capilla et al., 2002). Clearly,
if lactate is the main glycogenic substrate and glucose is a relatively
unimportant fuel for muscle, other extracellular substrates must be fuelling
glycogenesis and muscle metabolism as a whole, during recovery from exercise.
The concentration of alanine, a major form of inter-tissue transport of amino
acid-derived carbon (Mommsen et al.,
1980), increases in both plasma and white muscle following
exercise (Milligan, 1997),
though its role in fueling muscle recovery from exercise is not known. Overall
it is believed that protein does not make a significant contribution to
fuelling aerobic metabolism, despite it being plentiful in the white muscle
(Lauff and Wood, 1996).
Trout white muscle has considerable intramuscular lipid stores and Richards
et al. (Richards et al.,
2002a) have suggested a prominent role for lipid oxidation in
fueling exercise recovery, thus sparing intracellular lactate carbon for
glycogenesis.
Thus, whereas we have some insights into what fuels are probably important
for supporting muscle glycogenesis, we have very little understanding of what
fuels are preferred and how these fuels are used by muscle (i.e. carbon for
glycogen synthesis or for oxidation). Therefore, the purpose of this study was
to test the hypothesis that intracellular lactate is the major substrate for
trout muscle glycogenesis and the muscle relies upon extracellular substrates
for oxidation to provide the ATP necessary to drive glycogen synthesis. We
used the in vitro isolated white muscle slice preparation, which has
been shown to be metabolically viable, capable of glycogenesis and affords the
opportunity to control substrate availability
(Frolow and Milligan, 2004).
Our objective was to determine which substrates support glycogenesis and,
using appropriate radiotracers, determine how these extracellular substrates
are used by muscle (glycogenic versus oxidative fate).
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Materials and methods |
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). All holding and handling of fish was in accordance with the
Canada Council on Animal Care Guidelines and approved by the University of
Western Ontario Senate Animal Use Subcommittee.
Experimental protocol
Fish were exhaustively exercised by chasing them in a 300 l circular tank
for 5 min at which point they were unresponsive to further manual stimulation.
Previous studies have shown that this form of exercise leads to exhaustion and
a significant reduction in muscle glycogen (e.g.
Milligan, 1996). Fish were
transferred in a water-filled acrylic, 4 l box to a tank containing 0.3 g
MS-222 (tricaine methane sulfonate; Syndel, Vancouver, BC, Canada) buffered to
a pH of 7.8 with NaHCO3 in 3 l of dechlorinated tapwater. Fish died
within 1–2 min without struggling. Resting fish were killed in the same
manner but were not exercised.
Following anesthetization, a 1.5 cm3 block of muscle tissue of
approximately 1.5 g was rapidly excised from epaxial muscle close to the
mid-dorsal line of the fish, as described by Frolow and Milligan
(Frolow and Milligan, 2004).
The block of muscle tissue was then placed in ice-cold modified Cortland's
saline containing 140 mmol l–1 NaCl, 3.5 mmol
l–1 KCl, 1.0 mmol l–1 MgSO4
7H2O, 3.0 mmol l–1 NaHPO4
H2O, 4.5 mmol l–1 NaHCO3, 1.0 mmol
l–1 CaCl2, 10.0 mmol l–1 Hepes
and 0.30% defatted bovine serum albumin oxygenated with humidified 99.5%
O2/0.5% CO2 and adjusted to pH 7.8. Tissue slices of
approximately 1.3–1.4 mm in thickness and weighing 243.5±1.8 mg
(N=136) were obtained from tissue blocks maintained in iced saline
using a Stadie-Riggs microtome and tissue slicer blade (Thomas Scientific,
Swedesboro, New Jersey, USA). One slice was immediately blotted dry and frozen
by freeze clamping between aluminum blocks cooled with liquid nitrogen to
determine the metabolic status of the tissue at the time of sampling (referred
to as `time 0' slice). The remaining slices were incubated in 3.0 ml of
modified Cortland's saline (as above) continuously oxygenated with humidified
99.5% O2/0.5% CO2 for 1 h at 15°C in a 25 ml
Erlenmeyer flask in a shaking water bath. Following incubation, tissue slices
were removed, blotted dry, and freeze-clamped. Frozen tissue slices were
ground to a fine powder using an insulated mortar and pestle cooled with
liquid nitrogen and stored at –80°C until time of analysis.
Experimental series
To quantify the oxidation of extracellular substrates,
[14C]CO2 production was measured using a modified
CO2 trapping system (French et
al., 1981) adapted by Walsh et al.
(Walsh et al., 1988). A closed
CO2 trapping apparatus consisted of a glass filter paper, wetted
with 75 µl of 10 mol l–1 benzylthonium hydroxide, placed
in a well suspended in a 25 ml Erlenmeyer flask, containing 3.0 ml of aerated
incubation medium. Muscle slices were added to incubation flasks, the flasks
were sealed and either [U-14C]lactate (specific activity: 50 mCi
mmol–1; 1 Ci=3.7x1010 Bq),
[U-14C]glycerol (specific activity: 140 mCi
mmol–1) or [U-14C]palmitate (specific activity:
400 mCi mmol–1) (all from ICN Radiochemicals, Montreal, PQ,
Canada) was added to yield the final specific activities in the incubation
saline given in Table 1. The
substrate concentrations for each experiment are given in
Table 1 and are typical levels
seen post-exercise in trout plasma (e.g.
Richards et al., 2002a).
Palmitate was in the form of sodium palmitate, solubilized according the
methods described by Berry et al. (Berry et
al. 1991). The flasks were incubated in a shaking water bath at
15°C for 1 h. In every experiment, a flask containing everything except
tissue was incubated with the experimental flasks to account for spontaneous
[14C]O2 production from the universally labeled
14C substrates and the subsequent [14C]O2
production rates were corrected accordingly.
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After 1 h, 50 µl of 70% HClO4 was added to each flask to halt metabolism and liberate 14CO2. The sealed flasks were vigorously shaken at room temperature for 60 min on an orbital shaker at 100 revs min–1 to ensure complete collection of [14C]O2. At the end of this 60 min period, the muscle slices were freeze-clamped and analyzed for muscle glycogen and lactate and the filter papers removed and added to 5.0 ml of Ready Safe (Beckman, Mississauga, Canada) scintillation fluid in a scintillation vial and counted for total 14C radioactivity. Initial experiments using [14C]NaHCO3 (specific activity: 50 mCi mmol l–1; ICN Radiochemicals) indicated that the [14C]O2 trapping efficiency of the system is 84±2% (N=8) and [14C]O2 production rates were corrected accordingly.
To determine incorporation of extracellular substrate carbon into glycogen carbon, a 500 µl sample of the tissue glycogen suspension (see below), was added to 5.0 ml of Ready Safe fluor and counted for total 14C radioactivity. For each experiment, each treatment was performed in duplicate on tissue slices obtained from the same fish, and N in the figures refers to the number of fish used per experiment.
Analytical techniques and calculations
Frozen tissue slices were individually ground to a fine powder using an
insulated mortar and pestle cooled with liquid nitrogen. Muscle glycogen
content was assayed by isolating the glycogen (Hassid and Abraham, 1959) and
measuring the free glucose after digestion of the glycogen with
amyloglucosidase (Bergmeyer,
1965). In our laboratory, we typically recovery 95–100% of
the glycogen with this method. Muscle concentrations of lactate, adenosine
triphosphate (ATP) and phosphocreatine (PCr) were measured on approximately 50
mg of muscle ground to a fine powder in a liquid N2-cooled mortar
and vigorously resuspended in 1 ml 8% HClO4. Lactate, ATP and PCr
were then measured in the supernatant neutralized with 3 mol
l–1 KOH (Bergmeyer,
1965).
Tissue [14C]glycogen was determined by adding 500 µl of the
resuspended glycogen (see above) (Hassid and Abraham, 1959) to 5 ml of Beckman
Ready Safe scintillation cocktail. The incorporation of extracellular
substrate into the muscle glycogen pool was calculated using the specific
activities of the substrate in the saline, and the specific activity of muscle
glycogen and CO2 (Table
1) after Pagnotta and Milligan
(Pagnotta and Milligan, 1991)
and according to the equation:
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The contribution of extracellular substrate to glycogen synthesis was calculated from the substrate incorporation and the amount of glycogen synthesized over the 1 h period.
All samples were counted on a Packard 1900TR Liquid Scintillation Counter using automatic quench correction. All biochemicals were purchased from Sigma Chemical Co. (Mississauga, ON, Canada), Boehringer-Mannheim Chemical Co. (Laval, PQ, Canada), and all enzymes were purchased from Worthington Biochemical Corp. (Lakewood, NJ, USA).
Statistical analyses
Values presented are means ± 1 s.e.m. (N). Statistical
analyses were performed using a one-way analysis of variance (ANOVA) followed
by Kruskal–Wallis comparison tests or a Student's t-test and
significant differences were accepted when P<0.05.
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After 1 h incubation in 5 mmol l–1 lactate there was net glycogenesis (Fig. 1A) and significant incorporation of extracellular lactate, as indicated from 14C activity, into both the glycogen and CO2 pools. The amount of extracellular lactate incorporated into CO2 was about 100-fold greater than that incorporated into glycogen (Fig. 1B,C). Further, the contribution of extracellular lactate to total glycogen synthesis was trivial, accounting for less than 0.1% of the total glycogen synthesized (compare Fig. 1A and B).
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The combination of 0.3 mmol l–1 palmitate and 5 mmol l–1 lactate did not affect net glycogen synthesis, or lactate incorporation into glycogen or CO2 compared to either substrate alone (Fig. 1). However, incubation in palmitate plus glycerol plus lactate eliminated the stimulatory effect of glycerol on glycogenesis and lactate oxidation (Fig. 1A,B).
Incubation of muscle slices in glycerol plus lactate consistently stimulated glycogen synthesis compared to that in tissues incubated in glycerol alone (Fig. 2A). However, the incorporation of extracellular glycerol carbon into glycogen synthesized was very small, with only 0.34 nmol g–1 entering the glycogen pool, accounting for less than 0.01% of the total glycogen synthesized (Fig. 2B), which is one tenth of the contribution of lactate carbon to glycogen (compare Fig. 1B and Fig. 2B). Extracellular glycerol carbon made a greater contribution to oxidative metabolism than glycogenesis, as there was a tenfold greater incorporation into CO2 than into glycogen (Fig. 2B,C). However, the oxidation of extracellular glycerol was only 1% that of extracellular lactate oxidation (compare Fig. 1C and Fig. 2C) and was unaffected by the presence of lactate.
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When all substrates were present at a concentration of 1 mmol l–1, muscle synthesized glycogen (Fig. 4A) and the stimulatory effects of glycerol on glycogenesis in slices incubated in glycerol plus lactate or glycerol plus palmitate were still evident (Fig. 4A). The net amount of glycogen synthesized was also unaffected by the altered substrate concentrations (compare Fig. 4A with Figs 1A, 2A and 3A).
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Extracellular substrate use during muscle glycogenesis
In the absence of any extracellular substrates, there was no net glycogen
synthesis, despite clearance of a significant amount of muscle lactate,
suggesting that under these conditions, the major fate of endogenous lactate
is oxidation (Fig. 5 #1). It is
unlikely that the disappearance of lactate from the muscle is a result of
lactate efflux since the white muscle membrane is relatively impermeant to
lactate efflux and as a consequence, the bulk of the lactate generated is
retained within the muscle (Sharpe and
Milligan, 2003). The presence of extracellular lactate, glycerol
or palmitate supported glycogenesis, presumably by serving as oxidative fuels
(Fig. 5#2,#3,#4). Their
presence probably reduced the reliance upon oxidation of intracellular
lactate, sparing some of it for glycogenesis
(Fig. 5#5). Although there was
net glycogenesis when lactate, palmitate or glycerol was available, the
reduction in intracellular lactate was in excess of that accounted for in the
amount of glycogen synthesized, suggesting either some intracellular lactate
was still being oxidized or the `missing' lactate (5–6 µmol
g–1; Table 2)
was trapped in glycogenic intermediates. The observation that when tissues
were incubated in glycerol plus lactate or glycerol plus palmitate, all the
lactate cleared could be accounted for by the glycogen synthesized could mean
that the availability of oxidizable substrates is limiting to glycogenesis and
determines the fate of intracellular lactate.
Lactate was the preferred extracellular substrate for oxidation by muscle
during glycogen resynthesis, contributing as much as 16–30% of the ATP
required for the observed glycogenesis. This suggest that at least 70% of the
ATP needed for glycogenesis came from oxidation of intracellular substrates,
probably fatty acids (Richards et al.,
2002a). These estimates assume that substrates were completely
oxidized (allows calculation of ATP yield) and that glycogen was synthesized
from intracellular lactate (allows calculation of ATP requirement). This
represents a true muscle preference for extracellular lactate as an oxidizable
substrate and not a consequence of differences in substrate concentrations,
because when all substrates were present at the same concentration, lactate
was still preferred over the other fuels. Lactate is clearly taken up by the
muscle, despite the fact that the intracellular lactate concentration was four
times that of the extracellular concentration (20 vs 5 µmol
g–1 wet tissue); a gradient that should favor net efflux.
Lactate uptake by trout muscle is facilitated by a monocarboxylate (MCT)-like
transporter located in the sarcolemmal membrane
(Fig. 5#6) (Laberee and
Milligan, 1999) and the major fate of this extracellular lactate is clearly
oxidative (Figs 1C,
5#7). The amount of
extracellular lactate-derived carbon entering the CO2 pool is about
100 times that entering the glycogen pool (compare
Fig. 1B,C) and its oxidation
appears to `spare' lactate of intracellular origin for glycogenesis
(Fig. 5#5).
This model suggests that the fate of extracellular lactate is separate from
that of intracellular, glycolytically derived lactate; in other words, lactate
metabolism is compartmentalized in trout skeletal muscle. Compartmentation of
carbohydrate metabolism has been described for vascular smooth muscle
(Lynch and Paul, 1983) in
which extracellular glucose was the sole precursor for aerobic glycolysis,
whereas glycogenolysis was the precursor for oxidative phosphorylation.
Similarly, in insect flight muscle, glycolytic metabolites generated from
glucose do not mix with those from glycogen
(Srere and Knull, 1998). More
recently compartmentation of lactate metabolism has been suggested to explain
the simultaneous influx and efflux of lactate in the isolated rat heart
(Chatham and Forder, 1996;
Chatham et al., 2001). The data
from Chatham et al. (Chatham et al.,
2001), in which they perfused the rat heart with
[3-13C]lactate, indicated that lactate of extracellular origin was
preferentially oxidized whereas glycolytically derived lactate (endogenous)
was preferentially transported out of the cardiac muscle. These data are
consistent with the concept of an intracellular lactate shuttle, first
proposed by Stainsby and Brooks (Stainsby
and Brooks, 1990), then later refined by Brooks
(Brooks, 2000) to include the
concept of direct mitochondrial uptake and metabolism of lactate. Although the
evidence for the latter component of the intracellular lactate shuttle is
conflicting, there is general agreement that some type of compartmentation of
lactate metabolism exists in skeletal muscle. Gladden
(Gladden, 2004), in a recent
review, puts forward the idea of `microcompartmentation' to explain the
different fates of extracellular and intracellular, glycolytically derived
lactate. In this model, the physical locations of mitochondria and glycolysis
in the cytosol are distinct, such that mitochondria are located primarily at
the periphery of the cell, close to the sarcolemmal membrane, the site of
extracellular lactate uptake via MCTs, and removed from the cytosolic
location of glycolysis. Thus, once extracellular lactate is transported into
the cell, it is readily converted to pyruvate via cytosolic LDH, which is then
transported into the mitochondria and oxidized while the glycolytically
derived lactate (intracellular) is localized closer to the glycogenic
machinery. Our current understanding of teleost white muscle architecture, in
which mitochondria are localized peripherally and the glycogen granules, and
presumably the glycolytic and glycogenic enzymes as well, are located between
the myofibrils (Sänger and Stoiber,
2001) is consistent with this concept of intracellular
compartmentalization of lactate metabolism. Certainly, the fact that the
presence of oxidizable extracellular substrates (e.g. lactate, glycerol and
palmitate) stimulates glycogenesis and reduces lactate clearance is consistent
with the notion that intracellular lactate is `spared' for glycogenesis and
the idea that the enzymatic machinery for glycogenesis is remote from the
mitochondria. Clearly, validation of this model awaits further
experimentation.
Whereas extracellular glycerol alone can support glycogenesis, its
contributions to both oxidative metabolism (its oxidation yields only about
0.4% of the ATP needed for glycogenesis;
Fig. 2C) and glycogenesis
(glycerol carbon contributed <0.01% to glycogen carbon;
Fig. 2B) were minimal. This
minimal use of glycerol as an oxidative or glycogenic substrate may reflect
low activities of glycerol kinase (GK), which catalyzes glycerol to glycerol
3-phosphate (Fig. 5#8) or an
enhanced role of the glycerol 3-phosphate shuttle in fish muscle. The greatest
impact of glycerol on muscle glycogenesis was its stimulatory effect on
glycogenesis and lactate use when tissues were incubated in glycerol plus
lactate. The effect was specific for glycerol on lactate, in that lactate did
not influence glycerol use (Fig.
1B,C vs Fig.
2B,C). The explanation for this stimulatory effect of glycerol on
lactate use and glycogenesis is not clear, but may be related to need for
cytosolic NAD+ to support the first step in lactate metabolism. The
glycerol 3-phosphate shuttle, present within mammalian muscle
(Fig. 5#8)
(Rasmussen et al., 2003),
reoxidizes cytosolic NADH to NAD+, potentially increasing the
availability of NAD+ for lactate metabolism
(Hettwer et al., 2002) and the
presence of extracellular glycerol may enhance the activity of the
shuttle.
Fatty acid oxidation has been shown to substantially contribute to fueling
glycogen re-synthesis during recovery from high-intensity exercise in rainbow
trout in vivo (Richards et al.,
2002a; Richards et al.,
2004). Fatty acids of extracellular origin can be used by muscle,
as fatty acid uptake has been shown to be carrier-mediated in trout muscle,
presumably by a fatty acid translocase that has yet to be characterized. Once
in the muscle, these fatty acids undergo ß-oxidation
(Fig. 5#9)
(Richards et al., 2004),
supplying the ATP necessary to support glycogenesis, again sparing
intracellular lactate for glycogenesis. Accordingly, in the presence of
palmitate, there is net glycogenesis and the amount of lactate cleared is
reduced. Clearly trout white muscle is able to take up and utilize fatty
acids, but the estimated contribution to fueling glycogenesis is somewhat less
than suggested in vivo. In vitro, the contribution of fatty acid
oxidation appears to be less important, as palmitate oxidation is only one
tenth that of lactate (Fig. 1C
vs Fig. 3B), and at
most, contributed to only 16% of the ATP required to support the associated
glycogenesis. Nonetheless, in the presence of palmitate, there was net
glycogenesis, an effect that was enhanced when tissues were incubated in
glycerol plus palmitate. Glycerol did not stimulate palmitate oxidation,
suggesting that the effects are independent of one another. Incubating tissues
in lactate plus glycerol plus palmitate had a negative impact on muscle
glycogenesis, but was without effect on either lactate or glycerol oxidation.
The slight inhibitory effect of lactate on palmitate oxidation is consistent
with the inhibitory effects of carbohydrate on fatty acid oxidation (e.g.
Richards et al., 2002) and the observation from the present study that lactate
is a preferred substrate for muscle oxidative metabolism. However, why the
combination of lactate plus palmitate should inhibit glycogenesis is not at
all clear. There are reports of palmitate having negative impacts on glycogen
synthesis and lactate use in mammalian muscle, but none are consistent with
the current observations. For example, in cultured skeletal muscle cells from
insulin-resistant diabetic humans, the products of palmitate oxidation have
been shown to inhibit fractional glycogen synthase activity presumably because
of fatty acid oxidation-induced production of glucosamines
(Mott et al., 2000), whereas
in the present study, palmitate alone actually stimulated glycogenesis.
Furthermore, the addition of extracellular palmitate to the isolated perfused
heart from diabetic rats decreases the rate of lactate uptake and oxidation
(Chatham et al., 1999), again,
not see in this study. The explanation for these negative interactive effects
of lactate, palmitate and glycerol on muscle glycogen metabolism is
elusive.
The results from this study have been very useful in dissecting out the
potential roles and interactions of various extracellular substrates in
fueling trout muscle glycogenesis, however, there is a fundamental difference
between these in vitro and in vivo observations that
deserves mention. Namely, glycogenesis is more rapid in vitro [up to
10 µmol g–1 within 1 h post-exercise
(Wang et al., 1997;
Frolow and Milligan, 2004)]
suggesting that mitigating factors exist in vivo that act to retard
metabolic recovery in muscle. Hormones, in particular cortisol, are absent in
theses in vitro preparations. In vivo, the elevation of
plasma cortisol levels, typically seen following exhaustive exercise, appears
to be inhibitory to glycogenesis
(Milligan, 2003) as no net
synthesis of glycogen is seen until cortisol levels begin to decline
(Pagnotta et al., 1994).
Similar trends were observed in vitro in muscle slice preparations
(Frolow and Milligan, 2004),
though the nature of cortisol's effects are unclear.
In conclusion, the results of the present study provide the basis for generating a comprehensive model describing muscle fuel use during glycogenesis. The hallmark of this model is that muscle exhibits a distinct preference for extracellular lactate, and lactate metabolism is compartmentalized, with the fate of lactate dependent upon whether it is of intracellular (glycolytically derived) or extracellular origin. There are admittedly unknowns in this model, but it provides a framework for further exploration of the metabolic functioning of trout white muscle.
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Acknowledgments |
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