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First published online May 8, 2007
Journal of Experimental Biology 210, 1786-1797 (2007)
Published by The Company of Biologists 2007
doi: 10.1242/jeb.004499
Characterization of circannual patterns of metabolic recovery from activity in Rana catesbeiana at 15°C
Department of Integrative Physiology University of Colorado, Boulder, CO 80309-0354, USA
* Author for correspondence (e-mail: Ann.Petersen{at}colorado.edu)
Accepted 7 February 2007
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Summary |
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Key words: season, lactate, exercise, metabolism, ectotherm, bullfrog, blood pH
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Introduction |
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TOPSummaryIntroductionMaterials and methodsResultsDiscussionAppendixReferences |
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Studies on carbohydrate metabolism in amphibians have established a basic
pattern of accumulation and clearance of lactate following activity for these
animals. Following forced activity, under standard laboratory conditions
(20–25°C, 12 h:12 h L:D photoperiod), amphibians accumulate lactate
rapidly. For example, plasma lactate peaks almost immediately following
exercise in Rana catesbeiana
(Hutchison and Miller, 1978
;
Putnam, 1979) (for a review,
see Bennett, 1982). Lactate
production in the muscle follows a similar pattern, but clearance continues
long after oxygen consumption has returned to resting values (for a review,
see Gleeson, 1991). Muscle
lactate utilization in ranids follows a pattern established for other
ectotherms, with most intramuscular lactate recycled to glycogen
(Fournier and Guderley, 1992).
Data from a radioisotope tracer study in Bufo americanus indicates
that less than 1.5% of labeled lactate injected into an exercising toad is
detected as expired CO2
(Withers et al., 1988), which
is surprising given the relatively high aerobic scope of toads compared to
ranid amphibians (Carey, 1979;
Putnam, 1979;
Putnam and Bennett, 1983). To
our knowledge no radioisotope data exist demonstrating the metabolic fate of
lactate in a ranid. Overall, the available literature suggests a metabolic
recovery strategy in amphibians of both intracellular production and
sequestration of lactate, as well as uptake of circulating plasma lactate,
mainly by muscle tissue, which is subsequently used to replenish glycogen
stores (Putnam, 1979;
Withers et al., 1988;
Fournier and Guderley,
1992).
With this basic pattern established for carbohydrate metabolism under a
single set of conditions, it is important to consider that most amphibians,
especially those ranids that inhabit far northern regions of North America,
rarely actually exist under such homogenous conditions as those in the
laboratory. Species such as the American Bullfrog (Rana catesbeiana)
that successfully inhabit a range from British Columbia and Nova Scotia south
into Mexico (Lannoo, 2005)
must acclimatize to significant seasonal climate changes. Activity in the form
of migrations, hunting and predator avoidance must occur throughout the year
under a variety of ambient conditions. Active swimming by Rana
pipiens in water temperatures very close to freezing in a Canadian stream
has been reported (Cunjack,
1986), and Cunjack suggests that the over-wintering frogs may
change microhabitats frequently in order to seek out water with higher oxygen
content. Given that the metabolism of ectothermic vertebrates fluctuates with
thermal environment, properly allocating, for example, lactate to hepatic
rather than intramuscular glycogen stores, is exceedingly important when
environmental temperature varies widely. Very little is known about regulation
of carbohydrate metabolism in response to changes in season or environmental
conditions, and almost nothing is known about lactate metabolism alteration,
specifically. In this study, we provide insight into this question, using the
bullfrog as a model species that experiences significant seasonal changes in
its environment.
It has already been established that several aspects of carbohydrate
metabolism do fluctuate seasonally in amphibians. Examples of metabolic
parameters that change seasonally in amphibians include metabolite levels
(Byrne and White, 1975), oxygen
consumption (Bícego-Nahas et al.,
2001), and intrinsic skeletal muscle properties
(Girgenrath and Marsh, 2003).
Circulating levels of (and sensitivity to) epinephrine, glucagon, and insulin
also change with the seasons in frogs
(Hanke and Neumann, 1972;
Farrar and Frye, 1977;
Schlaghecke and Blum, 1981).
Hepatic glycogen levels in Rana esculenta were found to be 6.5x
higher in winter frogs than summer frogs, with a correlated increase of
activity in the enzymes that control glycogen deposition
(Scapin and Di Giuseppe,
1994).
The functional significance of circannual rhythms has been widely
speculated on, and may include preemption of freezing conditions
(King et al., 1995),
protection of glycogen stores for spring emergence from hibernation
(Scapin and Di Giuseppe,
1994), and support of glycolytic versus oxidative
metabolism in potentially hypoxic over-wintering habitats (Tattersall and
Boutlier, 1997). It is not known if lactate metabolism is similarly seasonally
sensitive. It is also not known if post-exercise glucose homeostasis,
gluconeogenesis and glycogenesis respond to time of year.
The purpose of this study was twofold. First, to trace the fate of lactate post-exercise, and create a basic profile of carbohydrate metabolism in the American bullfrog Rana catesbeiana, including measurements of extracellular acid–base balance, oxygen consumption, and blood and tissue metabolite profiles. Second, we sought to characterize the metabolic response to activity in the American bullfrog at a seasonally neutral temperature (15°C) in winter (January) and summer (June) months, in order to determine if `seasonal' changes in carbohydrate metabolism are a result of an underlying circannual component, or simply a response to an acute environmental condition, such as temperature or photoperiod. We predict that glycogen stores will be increased in winter months even when the animals are acclimated to a seasonally neutral temperature, and that lactate and glucose metabolism (at rest and in response to activity), will be sensitive to season even in the absence of change in environmental condition.
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Materials and methods |
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Cannulation
Frogs were submerged in neutralized 0.75% tricaine methanesulfonate
(MS-222, Sigma, St Louis, MO, USA; cat. no. A5040) solution until fully
anaesthetized (approximately 10–20 min). A 2.5–5.0 cm parasagittal
incision was made to expose the abdominal cavity directly below the heart,
with care taken to avoid (1) severing the mid-ventral vein or (2) damaging the
sternal cartilage. A cannula (3-French Soft PU intravascular tubing,
bpULD-T30, Instech Solomon, San Antonio, TX, USA) was inserted 5–6 mm
into the left systemic arch. The left systemic arch was chosen because it
supplies the abdominal viscera rather than more hypoxia-sensitive organs such
as the kidney, which are supplied by the right systemic arch
(Gilbert, 1965). The cannula
was flushed with 0.015% heparin in 0.9% saline, and secured to the vessel
using 000 silk surgical sutures. The cannula was run subcutaneously and exited
the cavity caudo-dorsally. All surgeries were completed within 2 h. Animals
were excluded from experimentation if approximately >1 ml of blood loss
occurred during surgery. Frogs were allowed to recover from surgery for 24 h
prior to experimentation based on the results of the following control
experiments.
Control experiments
In order to ensure that our cannulation procedure and serial blood sampling
were not affecting metabolism, we performed two initial experiments on resting
animals. Four adult male Rana catesbeiana were chronically
cannulated. Blood was sampled via the cannula at 0, 6, 24 and 30 h
following the surgery and measured for pH using a BMS 3 MK2 analysis system
(Radiometer, Copenhagen, Denmark) with a glass capillary electrode. Samples
were also analyzed for lactate concentration
(Gleeson, 1985). In a separate
experiment designed to test the effect of serial sampling on lactate and pH
levels, four adult Rana catesbeiana were chronically cannulated and
allowed to recover for 24 h. Blood was sampled (250 µl) via the
cannula at rest and then every 60 min for 240 min, totaling six separate 250
µl blood samples (equal or greater than the total volume of blood removed
in the following experiments). Whole blood samples were analyzed for pH and
plasma samples were analyzed for lactate levels as previously described. Data
were statistically compared utilizing a one-way ANOVA, Fisher's PLSD to test
for significant (P
0.05) differences between treatment groups.
This statistical analysis was used to compare all treatment groups in this
study, with the exception of oxygen consumption
(
O2) and carbon dioxide
production (CO2) data,
which were analyzed utilizing unpaired t-tests at individual time
points.
To ensure the most accurate approximations of organ mass in Rana catesbeiana, we dissected six adult males (mass= 178.0±2.8 g) and recorded the mass of the individual organ (blotted free of blood) for use in calculations of the tracer portion of the study.
Experiments
Frogs were allowed to rest in individual containers undisturbed for 240 min
on the day of experimentation, with
O2 monitored
continuously. At the end of the initial rest period, animals were injected
with 1 µCi (winter) or 2 µCi (summer) of 50 µCi 0.5
ml–1 [U-14C]L-lactic acid sodium salt
(MP Biomedicals, Irvine CA, USA). The higher dosage used in summer was to
improve the absolute amount of radioactivity captured in respiratory
CO2 samples. All isotopic data are expressed either as percentage
of injected dose or are divided by the specific activity of the lactate pool
and expressed as µmol lactate converted g–1 tissue 4
h–1, thus negating any effect of variable dosage on either
data interpretation or the reported data themselves.
Following 10 min of equilibration time post-injection, experimental animals
were hopped on a small animal treadmill for 2 min. Control animals were
allowed to continue resting with no exercise for 2 min. Blood was sampled
via the cannula from both groups 12 min after injection (time=0).
Depending on treatment group, animals were either sacrificed immediately
post-exercise (PE), or allowed to recover for 240 min (REC 4), and then
sacrificed. The control group was rested for 240 min (REST) and then
sacrificed. Blood was sampled at rest, and every 60 min from time=0 and
immediately preserved in six volumes of ice-cold 6.0% HClO4. Whole
blood was also rapidly collected from the cannula and analyzed for pH using a
radiometer BMS 3 MK2 analysis system with a glass capillary electrode,
calibrated to measure pH at 15°C. Respiratory gas exchange was
continuously measured throughout recovery using an open flow respirometry
system. Downstream air was dried using Drierite desiccant (W. A. Hammond
Drierite Company Ltd, Xenia, OH, USA) before entering the analyzer system.
Samples were then analyzed by an Anarad, Inc. AR-411 carbon dioxide analyzer
and an Applied Electrochemistry S-3A O analyzer.
O2 and
CO2 (ml
g–1 h1 STPD; standard temperature and
pressure dry) were calculated as per Withers
(Withers, 1977) and recorded
using LABVIEW data acquisition. All expired CO2 was then trapped in
24 ml of 1:3 ethanolamine:methylcellusolve, collected and replaced every 30
min, and later analyzed for 14C activity by scintillation counting.
At the end of the 240 min experiment, animals were sacrificed via rapid
decapitation using a small animal guillotine. Legs, liver and heart (heart in
summer only) were quickly dissected out and dropped into liquid nitrogen. The
rest of the carcass was blended in four volumes ice cold 6.0% HClO4
and the homogenate saved for later determination of 14C activity.
All methods were approved by the University of Colorado Internal Animal Care
and Use Committee.
Metabolite determination
Acidified whole blood samples were kept frozen at –70°C for <6
months, thawed and centrifuged (7200 g for 10 min). The
supernatant was analyzed for lactate
(Gleeson, 1985) and glucose
concentrations (Bergmeyer and Bernt,
1974). Tissues were kept frozen at –70°C for <6
months, and then homogenized in six volumes ice cold 6.0% HClO4.
Homogenates were then centrifuged at 7200 g for 10 min, and
the supernatant analyzed for lactate and glucose as above. Metabolite values
for all the tissues were corrected for extracellular spaces of 10%, 26% and
30%, for muscle (Ling and Kromash,
1967), liver (Devireddy et
al., 1999) and heart (Armstrong
et al., 1969), respectively. Glycogen content of tissues was
analyzed by ethanol precipitation of glycogen, subsequent breakdown with
amyglucosidase (Keppler and Decker,
1974) followed by determination of glucose levels as referenced
above.
14C activity
Supernatant from blood and tissue samples, as well as trapped
CO2 in ethanolamine and methycellusolve solution, were analyzed for
activity of injected 14C. Aliquots (100 µl) of blood and tissue
samples were separated into lactate and glucose using a Dowex ion exchange
column as described in Donovan and Gleeson
(Donovan and Gleeson, 2006).
These samples, as well as aliquots of the precipitated glycogen were then
analyzed using scintillation counting for 14C activity. Samples (1
ml) of trapped CO2 were mixed with 3 ml of Scintiverse
Scintillation cocktail (Fisher Scientific, Houston, TX, USA) and aqueous
samples (plasma, tissue) were mixed with 3.5 ml of Scintisafe Plus 50%
Scintillation cocktail (Fisher Scientific, Houston, TX, USA). Samples were
analyzed using a Wallac Model 1204 scintillation counter with internal quench
correction. For ease of comparison, all data presented and discussed are
expressed in terms of percentage of counts returned.
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Results |
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Blood pH, oxygen consumption, and metabolites – summer
Following activity in the `summer' frogs, both
O2 and
CO2 immediately increased
three to four times but steadily returned to resting levels within 70 min.
Peak O2 following
activity reached 0.12 ml O2 g–1
h–1 and peak
CO2 was 0.33 ml
CO2 g–1 h–1
(Fig. 2). Blood pH dropped
rapidly following exercise (Fig.
3A) and was inversely proportional to plasma accumulation of
lactate (Fig. 3B). Blood pH
continually recovered from the initial acidosis until reaching resting levels
within 240 min, rising on average 0.1 pH units h–1
(Fig. 3A). Within the same time
period, blood lactate remained elevated 2–3 times above resting levels,
with clearance in the 240 min of recovery equal to about 30% of lactate
produced by activity (Fig. 3B).
Resting blood glucose levels remained relatively stable over the 4 h
experiment, and did not differ significantly at any time point from the
average blood concentration of 0.72±0.1 mmol l–1
(P>0.05). Blood glucose concentration progressively increased
following exercise and peaked at 1.6 mmol l–1 at 180 min of
recovery, and was significantly elevated from resting values at all sampling
intervals following exercise (Fig.
3C).
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Tissue metabolites
Intracellular lactate concentrations in the gastrocnemius muscle, the liver
and the heart, increased significantly following activity as would be expected
(P<0.05, Fig. 4A).
Following exercise, lactate levels in the gastrocnemius muscle increased from
resting levels of 9.2±3.5 mmol l–1 to an average of
nearly 40.0 mmol l–1. Muscle lactate levels were
statistically indistinguishable from resting levels by 240 min of recovery
(P>0.05). This is in contrast to the pattern of incomplete lactate
clearance during recovery period measured in both heart and liver tissues
(Fig. 4A). Liver cells were
found to have an average resting lactate concentration of 4.2±1.3 mmol
l–1. Immediately following 2 min of hopping, liver lactate
levels doubled, and remained significantly elevated throughout the duration of
recovery. Heart tissue followed a similar pattern, with low resting lactate
concentrations (2.0±0.1 mmol l–1) that rose
immediately following activity to approximately 10 mmol l–1.
Heart lactate levels remained significantly elevated from resting values over
the 240 min of recovery (P<0.05,
Fig. 4A).
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Fate of [14C]L-lactate
Tissue and gas analyses accounted for 50–70% of the injected
[14C]lactate. Of this fraction, approximately 60–70% of the
[14C]lactate was converted to another form during the 240 min of
recovery (Fig. 5,
Table 2 and Appendix). The
measured pattern of lactate dispersal was the same regardless of whether the
frog rested for 240 min or was exercised and allowed to recover for 240 min
(P>0.05, Fig. 5).
Of that 60%, the majority (40–45%) of lactate cleared was
gluoconeogenically converted to glycogen stores in the muscle and the liver,
whereas another 10% was gluconeogenically converted to glucose in tissue and
blood glucose pools. Approximately 10% of the lactate was oxidized
(Fig. 5). Five- to tenfold as
much de novo glucose and glycogen was deposited in skeletal muscle
(per gram) as was deposited in liver (Fig.
6).
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Seasonal changes in metabolic profile
Most parameters measured did not change significantly based on season alone
when temperature and photoperiod were held constant
(Table 2).
There was more lactate carbon incorporated into liver glycogen than liver glucose, although this trend was only significant in the summer (Fig. 6, Appendix). The gastrocnemius muscle converted the majority of its lactate into glycogen in the winter (P<0.05), but equally into both glucose and glycogen in the summer. The muscle appeared to demonstrate an overall increased gluconeogenic and glyconeogenic function in summer, converting up to two to three times more lactate to glycogen and glucose than in winter, although this trend was not significant (Fig. 6). Lactate deposition in liver was not influenced by season. Activity did not significantly increase either liver or muscle conversion of lactate to glucose or glycogen in either season (Fig. 6, Table 2, Appendix).
Tissue glucose
Of all parameters examined, levels of free intracellular glucose
demonstrated the most significant fluctuations based solely on season.
Although all three tissues (liver, heart, muscle) demonstrated an identical
response to activity (immediate increases in glucose that were not reduced to
resting values within 4 h of activity, P<0.05,
Fig. 7), tissues from January
experiments contained two to four times as much glucose at all sampling
periods as those collected in June (Table
2, Fig. 7). Muscle
and heart glucose levels remained significantly elevated for the duration of
the 4 h recovery, while liver samples returned to resting levels in both
seasons.
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O2 and CO2O2 of
frogs in winter over 240 min was significantly higher than that of frogs in
summer following exercise (P<0.05;
Fig. 2,
Table 2), although no
significant difference was found between seasons at individual 30 min
intervals (Fig. 2). Resting
O2 was not different
between seasons (P>0.05; Fig.
2, Table 2). The
average CO2 of winter
frogs was not different from summer frogs (P>0.05) when averaged
over the 4 h of recovery. However, the peak
CO2, as well as the
CO2 averages at 20, 30
and 40 min of recovery, were found to be significantly higher in summer frogs
(P<0.05; Fig.
2).
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Discussion |
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TOPSummaryIntroductionMaterials and methodsResultsDiscussionAppendixReferences |
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),
lizards (Gleeson and Dalessio,
1989) and fish (Milligan and
Girard, 1993). The increase in both tissue and blood lactate
levels are typical of an animal performing power sprint type exercise,
supported mainly by glycolytic white muscle
(Putnam and Bennett,
1983).
The intra–extracellular concentration gradient for lactate over the
course of recovery is not typical for an ectotherms.
Fig. 3B clearly shows a lactate
clearance pattern in which blood lactate levels remain significantly elevated
for the entire course of recovery, despite the complete clearance of excess
lactate from the skeletal muscle tissue
(Fig. 4A). In contrast to these
findings, blood and tissue lactate levels reported by Fournier et al.
(Fournier et al., 1994) in
Rana pipiens demonstrate what could be considered to be a pattern of
lactate clearance typically found in reptiles and mammals, in which
extracellular levels remain equal to or lower than intracellular levels for
the duration of recovery (for reviews, see
Gleeson, 1991;
Gleeson, 1996). Similar to our
findings in bullfrogs, a lactate `reversed-gradient' following exercise
recovery was observed in Rana temporaria, but only at hibernating
temperatures (>7°C) in frogs submerged for several months
(Tattersall and Boutilier,
1999). Although the bullfrogs in our study were acclimated to a
temperature of 15°C, it is unlikely that this temperature was cold enough
to elicit a hibernation response. However, when we repeated this experiment at
25°C the concentration gradient was abolished (Petersen et al., 2006),
indicating that temperature may play a role in creating this gradient.
Although monocarboxylate transporter (MCT) isoforms have not been
identified in amphibian skeletal muscle, data from Mason et al.
(Mason et al., 1986) suggest
that lactate export from frog sartorius muscle is driven by lactate/proton
co-transport. Evidence for MCT 1 and MCT 4 transporter proteins has been
established in fish (Wang et al.,
1997; Laberee and Milligan,
1999) and in lizards (Donovan
and Gleeson, 2001). Assuming that amphibian skeletal muscle also
relies on a lactate/H+ co-transport system, low temperature may be
affecting lactate uptake by reducing transporter protein activity in the
muscle and other tissues. Although changes in extracellular pH associated with
the large lactate load in the blood could also alter transporter kinetics,
blood pH recovers to resting value within the 240 min of recovery while
extracellular lactate remains elevated
(Fig. 3A,B). Therefore, it is
unlikely that alterations in the intra–extracellular pH gradient are
driving the exclusion of lactate from the cells. Amphibians may employ an
active lactate transporter system in order to maintain this
reversed-concentration gradient (Donohoe
and Boutilier, 1998), but to our knowledge, no additional evidence
has been reported to support this hypothesis. Limited perfusion may also limit
the muscle's ability to take up lactate or glucose. Amphibian studies have
clearly demonstrated that the combination of cold submergence (5°C) and
hypoxia elicit shunting of blood flow away from vessels feeding muscle beds
(Boutilier et al., 1986;
Pinder et al., 1992). White
muscle tissue specifically is under-perfused at low temperatures in rainbow
trout (Barron et al., 1987).
More studies are necessary in order to determine if the muscle is limited in
protein-facilitated metabolite uptake, or if reduced perfusion is responsible
for the gradient observed in this study.
Plasma and muscle glucose increased significantly and remained elevated
throughout the course of recovery (Fig.
3C and Fig. 4B).
The hyperglycemic plasma response to activity is indicative of the
exercise-induced catecholamine activation reported in Rana pipiens
(Fournier et al., 1994). Yet
this is an unusual pattern for ectotherms. In studies on lizards sprinted for
15 s (Donovan and Gleeson,
2006) or sprinted for 5 min
(Gleeson and Dalessio, 1989),
blood glucose levels remain unchanged. Post-exercise increases in blood
glucose are reported to be non-existent
(Pagnotta and Milligan, 1991),
or minimal (Wang et al., 1994)
in rainbow trout. Frogs on the other hand, appear to follow a more `mammalian'
pattern, of significant increases in blood glucose in response to activity
(Fig. 4B), coinciding with and
presumably due to catecholamine release
(Fournier et al., 1994).
Mammals restore normoglycemia within 1 h of epinephrine clearance
(Sigal et al., 1994), whereas
significant hyperglycemia was persistent over the 4 h of recovery in this
study (Fig. 3C) and that of
Fournier et al. (Fournier et al.,
1994).
It is unknown whether extended, elevated glucose post-activity confers any
functional benefit to recovery in frogs, or may simply be a consequence of low
metabolism. However, the very presence of elevated extracellular glucose
levels is intriguing, because it suggests that hepatic and renal tissues may
be more involved in blood glucose homeostasis and exercise fuel provisioning
than previously suggested (Fournier and
Guderley, 1992; Fournier and
Guderley, 1993; Fournier et
al., 1994; Pagnotta and Milligan, 1994).
Previous research on Rana pipiens demonstrated negligible lactate
uptake or glucose efflux by the liver
(Fournier and Guderley, 1992;
Fournier and Guderley, 1993).
In fact, after these authors removed the liver from the frog, post-exercise
lactate removal rates and blood glucose levels were not significantly
different from those of intact animals
(Fournier and Guderley, 1993).
The increases in circulating glucose (Fig.
4B) might suggest that the liver or possibly the kidneys are
breaking down glycogen or gluconeogenically converting triglycerides or
protein to glucose. In humans, the kidneys have been shown to be responsible
for up to 40% of systemic glucose appearance in response to epinephrine
(Stumvoll et al., 1995), a
finding that could reconcile the conundrum of normal plasma glucose levels in
hepatectomized frogs. Liver is the only tissue examined that returned to
resting glucose values by the end of the trial
(Fig. 4B), further suggesting
possible export of glucose from this tissue. Concomitant elevated hepatic
lactate levels are additionally indicative of Cori-cycling. 240 min after
cessation of exercise, free glucose in the gastrocnemius muscle remained
elevated, and the heart was higher in glucose than immediately following
exercise (Fig. 4B), suggesting
continued uptake of extracellular glucose.
However, hepatic glycogen reserves were not measurably changed in response
to exercise (Fig. 4C), arguing
against this interpretation. An intriguing pathway, that frogs are exporting
glucose from muscle utilizing glucosidic pathways, has been proposed
(Fournier and Guderley, 1993).
This hypothesis has not yet been thoroughly tested. The liver of some frogs,
albeit in response to freezing temperatures, is nonetheless extraordinarily
capable of exporting glucose (Storey,
1987). Our findings of [14C]glucose in the blood
suggest that the liver is involved in vivo in post-activity glucose
production. The role of the liver as a glucose source in frogs remains
enigmatic.
Limitations of tracer methodology
Tissue lactate exchange with the pyruvate pool can theoretically limit the
interpretation of the isotopic data
(Stanley and Lehman, 1988).
This exchange may be more problematic in the resting state than during
recovery, when the net flux through LDH strongly favors net reduction in the
lactate pool size (Gleeson and Delassio, 1989). Under resting conditions,
lactate and pyruvate pools are probably closer to equilibrium. As a result,
label incorporation into glucose, glycogen, or CO2 does not reflect
net lactate removal as much as it reflects lactate pool turnover. Given these
limitations we have interpreted resting animal data with caution.
O2, CO2 and pH: summer
Both O2 and
CO2 increased immediately
following activity, and returned to resting levels within 100 min of recovery
(Fig. 2). Withers et al.
(Withers et al., 1988) found
that toads exercised for 10 min also demonstrated a rapid return to resting
oxygen consumption within 1 h, in spite of the persistence of blood lactate
elevation. In our study,
CO2 returned to resting
levels approximately 80 min before respiratory acidosis was significantly
alleviated (Fig. 2). This
pattern is indicative of an acid–base buffering system that depends on
the buffering capacity of bicarbonate (for a review, see
Reeves, 1977), and consistent
with more recent explanations of buffering from mineralized tissues such as
the endolymphatic sac (Warren and Jackson,
2005), and renal and cutaneous proton excretion (for a review, see
Boutilier et al., 1992).
The post-activity fate of lactate: summer
The majority of the [14C]lactate metabolized was converted to
glycogen or glucose following exercise or rest. Unexpectedly, exercise had no
effect on the metabolic fate of lactate
(Fig. 5). We might have
expected an increase in oxidation of lactate concomitant to increased overall
aerobic metabolism following exercise. Our findings in terms of the fate of
lactate are in keeping with those reported in toads
(Withers et al., 1988) and
lizards (Gleeson and Delassio, 1989) that the majority of lactate produced
following activity is stored as glycogen, with <20% of lactate being
oxidized. This `carbon-recycling' pattern post-exercise is now well
established for most ectothermic vertebrates (for reviews, see
Gleeson, 1991;
Gleeson, 1996). Based on label
incorporation, both muscle and liver demonstrated glyconeogenic and
gluconeogenic capacity (Fig.
6). The liver, as previously reported in lizards (Gleeson and
Delassio, 1989), fish (Milligan and Gerard, 1993) and frogs
(Fournier and Guderley, 1992)
was found to be of lesser importance to whole body activity recovery than the
skeletal muscle. Our data support this
(Fig. 6), as the per gram
tissue glyconeogenic and gluconeogenic rate of label incorporation in the
muscle tissue exceeded that of the liver by 5–10 times; however, the
liver does appear to contribute to lactate recycling to some extent.
Seasonal changes in carbohydrate metabolism
Bullfrogs demonstrated surprisingly little change in metabolic profile
between seasons when acclimated to a single temperature. Glycogen levels
(Fig. 4C,D) were not different
between summer and winter after only a 2-week acclimatization period. These
data are in contrast to the findings of Scapin and Di Giuseppe
(Scapin and Di Giuseppe, 1994)
who demonstrated a sixfold increase in hepatic glycogen levels in Rana
escualenta housed in an outdoor terrarium over the course of the year.
This is an impressive glycogen loading ability considering that the maximum
glycogen `supercompensation' thought possible in rat tissue is twice resting
values, and this is only achieved via extreme training regimens and
creatine phosphate supplementation (Nelson
et al., 2001) or treatment with anabolic steroids
(Cunha et al., 2005). Our
findings in Rana catesbeiana suggest that previously reported extreme
seasonal glycogen production and storage must be correlated to acute
temperature rather than a chronic change between seasons.
No significant difference was found to exist between summer and winter
frogs in terms of resting or post-exercise lactate and pH levels. Likewise,
circulating glucose levels were not impacted by season alone. However, both
liver and muscle glucose levels were significantly higher in January animals
at rest and following exercise (Fig.
7). This is an interesting finding in the light of the large body
of literature describing the use of glucose by over-wintering frogs as a
cryoprotectant (Storey and Storey,
1984; Costanzo et al.,
1993; Steiner et al.,
2000). However, the mechanism by which glucose accumulates in
tissues has clearly been established in some ranids to be triggered not by
winter, or even cold temperature, but only by temperatures below zero
(Storey and Storey, 1984;
Costanzo et al., 1993).
Therefore, it is unlikely that the 15°C exposure reported here is
eliciting a cryoprotective response. However, older literature has reported
that such glucoregulatory hormones as epinephrine
(Farrar and Frye, 1977) and
insulin are more effective in amphibians in the fall (Hanke and Neumman, 1972)
and circulate in higher concentrations in winter
(Schlaghecke and Blum, 1981).
It is therefore plausible that heightened sensitivity to and/or concentrations
of insulin could be responsible for the increased tissue glucose levels found
in winter animals (Fig. 7).
The isotope incorporation data suggest that gluconeogenic and glyconeogenic
function of the gastrocnemius muscle are increased in summer
(Fig. 6). It would make sense
that in summer months, when breeding and feeding occur, to have increased
gluconeogenic capacity to support substrate homeostasis. This increased
conversion of labeled lactate to glucose and glycogen in the muscle does not
translate to higher intracellular glucose or glycogen levels at this time of
year, even in response to activity. Therefore, this increased capacity may be
representative of an overall fuel substrate change. Muscle may rely
predominantly on breakdown of stored fat bodies in winter, but switch to
carbohydrate sources in the summer. Our data do, in fact, suggest a higher
respiratory exchange ratio (RER) value in summer frogs post-exercise
(Fig. 2), possibly indicating
that more carbohydrate is being metabolized in this season. Donohoe and
Boutilier (Donohoe and Boutilier,
1998) report that wild caught frogs in January maintain resting
respiratory quotient (RQ), reflective of lipid metabolism, at least for the
first 45 days of submergence in the laboratory. The seasonal differences
reported here in CO2 and
O2 could be due to
seasonal changes in bicarbonate buffering capacity. Another possibility
besides changes in bicarbonate buffering capacity is that mineralized tissue
buffering capacity is altered between seasons. Warren and Jackson
(Warren and Jackson, 2005)
have recently demonstrated that Rana pipiens utilized both bone and
the endolymphatic sacs as sinks for increased proton load following exercise.
Changes in the capacity of this system could certainly lead to alteration in
levels of CO2 expired. Future studies should address questions of
whole body buffering capacity by season.
In conclusion, Rana catesbeiana appear to recycle lactate accumulated during exercise into glycogen stores in a manner similar to other amphibians, fish and lizards. Our data suggest an interesting reversed concentration gradient of extracellular to intracellular lactate post-exercise at 15°C that could be due to changes in transporter kinetics. In terms of seasonal changes in carbohydrate metabolism, it appears that intracellular glucose level is the only parameter of those examined that is altered by season alone. Lactate metabolism dose not appear to be altered seasonally. Based on this study, we conclude that previously reported `seasonal' changes in metabolic profiles in ranid amphibians are likely due to an acute environmental condition rather than a chronic seasonal alteration of metabolism.
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