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Metabolic recovery in goldfish: A comparison of recovery from severe hypoxia exposure and exhaustive exercise
Comparative Biochemistry and Physiology, Part C 148 (2008) 332–338
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Comparative Biochemistry and Physiology, Part C j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / c b p c
Metabolic recovery in goldfish: A comparison of recovery from severe hypoxia exposure and exhaustive exercise☆ Milica Mandic, Gigi Y. Lau, Manu M.S. Nijjar, Jeffrey G. Richards ⁎ Department of Zoology, The University of British Columbia, Vancouver, British Columbia, V6T 1Z4 Canada
A R T I C L E
I N F O
Article history: Received 9 March 2008 Received in revised form 28 April 2008 Accepted 29 April 2008 Available online 10 May 2008 Keywords: Glycogen Ethanol Lactate Liver Muscle Anoxia Carassius auratus
A B S T R A C T Severe hypoxia exposure and exhaustive exercise in goldfish both elicit a strong activation of substrate-level phosphorylation with the majority of the metabolic perturbations occurring in the white muscle. Approximately half of the muscle glycogen breakdown observed during severe hypoxia exposure was accounted for by ethanol production and loss to the environment, which limited the extent of muscle glycogen recovery when animals were returned to normoxic conditions. Ethanol production in goldfish is not solely a response to anoxia/hypoxia exposure however, as a transient increase in ethanol production was observed during the early stages of recovery from exhaustive exercise. These data suggest that ethanol production is a ubiquitous “anaerobic” end product, which accumulates whenever metabolic demands exceed mitochondrial oxidative potential. Exhaustive exercise and hypoxia exposure both caused a 7 to 8 μmol g− 1 wet mass increase in muscle [lactate] and the rates of recovery following these perturbations were similar. The rates of muscle PCr and pHi recovery after hypoxia exposure and exhaustive exercise were similar with levels returning to controls values within 0.5 h. Surprisingly, liver [glycogen] was not depleted during exposure to severe hypoxia, however, during recovery from both hypoxia and exercise dramatically different responses in liver [glycogen] were noted. During the early stages of recovery, liver [glycogen] transiently increased to high levels after exhaustive exercise, while during recovery from hypoxia there was a transient decrease in liver glycogen over the same time frame. Overall, this points to the liver playing a dramatically different role in facilitating recovery from exercise compared with hypoxia exposure. © 2008 Elsevier Inc. All rights reserved.
1. Introduction Energy production during exposure to severe hypoxia and exhaustive exercise occurs primarily via substrate-level phosphorylation (phosphocreatine (PCr) hydrolysis and glycolysis), which compensates for ATP demands that exceed oxidative capacity. During severe hypoxia exposure, goldfish suppress metabolic rate by up to 60% (Van Waversveld et al., 1989) and rely on large liver and muscle glycogen reserves to support substrate-level phosphorylation to maintain cellular energy balance (Lutz and Nilsson, 1997). In contrast, ATP demands in muscle during exhaustive exercise increase beyond the oxidative capacity and thus substrate-level phosphorylation must be activated to meet the energetic demands of muscle contraction (Richards et al., 2002a). Although the metabolic reasons for the activation of substrate-level phosphorylation differ during exposure to
☆ Contribution to the Special Issue of CBP on Chinese Comparative Biochemistry and Physiology presented at or related to the International Conference of Comparative Physiology, Biochemistry and Toxicology and the 6th Chinese Comparative Physiology Conference, October, 10–14, 2007, Zhejiang University, Hangzhou, China. ⁎ Corresponding author. Tel.: +1 604 822 2381; fax: +1 604 822 2416. E-mail address: [email protected] (J.G. Richards). 1532-0456/$ – see front matter © 2008 Elsevier Inc. All rights reserved. doi:10.1016/j.cbpc.2008.04.012
severe hypoxia and exhaustive exercise, the end metabolic profiles observed in muscle are similar with low [glycogen] and [PCr] and high [lactate] and metabolic [H+] (van den Thillart et al., 1989; Richards et al., 2002a, 2003, 2007). During recovery from hypoxia and exhaustive exercise, pathways must be activated to resynthesize ATP, PCr, and glycogen. To this end, the early portions of recovery are characterized by a period of excess O2 consumption (Scarabello et al., 1991; van den Thillart and Verbeek, 1991; van Ginneken et al., 1995), which in part represents a stimulation of oxidative phosphorylation to support the energetic costs of recovery. Generally, muscle [PCr] returns to resting values quickly during recovery from both hypoxia exposure and exhaustive exercise (within 1 to 2 h if not sooner), but recovery of muscle [lactate] and [glycogen] requires a longer period (N2 h; Richards et al., 2002b). Lactate is preferentially retained in muscle during recovery from exercise or hypoxia exposure (Richards et al., 2007) and there is accumulating evidence to suggest that muscle glycogenesis occurs in situ and relies on lactate as the substrate (Schulte et al., 1992). Members of the family Cyprinidae, especially the crucian carp (Carassius carassius L.) and the common goldfish (Carassius auratus L.), have long been recognized as unique among vertebrates for their
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incredible anoxia tolerance. The common goldfish, for example, can survive complete anoxia at room temperature (20 °C) for many hours (van den Thillart et al., 1983) and as environmental temperature decreases, survival can be extended to weeks or possibly even months (Blazka, 1958; Walker and Johanson, 1977). Prolonged anoxia survival at cold temperatures is likely related to three main factors: 1. the storage of high concentrations of fermentable substrates (e.g. liver [glycogen] of ∼ 800 μmol g− 1 wet mass; van den Thillart et al., 1980a), 2. the suppression of whole-animal metabolic rate (Van Waversveld et al., 1989), and 3. the ability to convert anaerobically derived lactate to ethanol and CO2 (Shoubridge and Hochachka, 1980). Ethanol production by the crucian carp and goldfish begins with the movement of lactate from other tissues to red and white skeletal muscle, where lactate is converted to pyruvate via lactate dehydrogenase. Pyruvate is then decarboxylated to acetaldehyde and CO2 via an inefficient or modified pyruvate dehydrogenase complex (van Waarde et al., 1993; Lutz and Nilsson, 1997). Typically, PDH converts pyruvate to acetyl-CoA with acetaldehyde as a bound intermediate, but during anerobiosis acetaldehyde “leaks” out of the PDH complex, diffuses into the cytoplasm and is converted to ethanol by alcohol dehydrogenase. Ethanol then freely diffuses across the gills, limiting lactate accumulation. Loss of ethanol to the environment, however, can represent a substantial loss of carbon, especially over months of anoxia exposure, and this may seriously impact the ability of the animal to recover in a normoxic environment. Few studies have examined the effects of exhaustive exercise on Cyprinids, and those that have mostly focused on the non-ethanol producing common carp (Cyprinus carpio; Driedzic and Hochachka, 1976; Sugita et al., 2001; van Ginneken et al., 2004) and no studies, to our knowledge, have examined whether ethanol production occurs during “anaerobic” exhaustive exercise. For the most part, the metabolic profiles observed in carp white muscle during exhaustive exercise are similar to those observed in other fish (e.g. trout), albeit the magnitude of the effects are smaller. Huber and Guderley (1993) noted significant accumulation of lactate during ∼ 30 min of exhaustive exercise in goldfish, but did not measure whether lactate was converted to ethanol. It is likely, however, that even if lactate conversion to ethanol occurs, due to the rapid rate of glycolysis necessary to support muscle contraction, lactate would be preferentially accumulated over ethanol. The objectives of this study were to compare the recovery profiles of white muscle [ATP], [PCr], [pHi], and glycogen in goldfish after exposure to 10 h of severe hypoxia at 10 °C or a bout of exhaustive exercise. We predicted that severe hypoxia exposure would lead to ethanol production and the loss of carbon to the environment and as a result, muscle glycogen recovery after hypoxia exposure would be impaired relative to exhaustive exercise where no or limited ethanol production was predicted to occur. Specifically, we examined the rate of ethanol production in goldfish during and after severe hypoxia exposure and exhaustive exercise to determine the relative amounts of ethanol produced and its relation to glycogen recovery. Changes in liver glycogen were also examined to gain insight into the potential role of the liver in fuelling metabolic recovery from hypoxia exposure and exhaustive exercise. 2. Materials and methods 2.1. Animal care Adult goldfish (Carassius auratus L.; ∼27 ± 1 g, 11.6 ± 0.4 cm; mean ± SD) were purchased from a local supplier (Delta Aquatics, Richmond, B.C., Canada) and held under flow-through conditions in 750 L tanks supplied with well-aerated, dechlorinated City of Vancouver tapwater (∼ 10 °C) for at least 1 month before experimentation. During holding, fish were fed daily with commercial goldfish flakes, but food was withheld 24 h before experimentation.
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2.2. Experimental protocols 2.2.1. Exhaustive exercise and recovery To exercise goldfish to exhaustion, four size-matched fish were placed into a Brett-style swimming respirometer supplied with wellaerated, dechlorinated tapwater at 10 °C. Fish were allowed 30 min of recovery from transfer and water velocity was maintained at b1 body lengths s− 1 during this period. Following the recovery period, fish were exhaustively exercised using a similar protocol to that described by Dobson and Hochachka (1987). Briefly, the swimming protocol began by increasing the water velocity to the maximum speed at which the fish swam in a burst-and-glide pattern (∼7 to 8 body length s− 1). This velocity was maintained until each fish fatigued and fell to the back of the swim tunnel. As each fish fatigued, it was manually returned to the current at which point the fish typically regained position and began swimming in a burst-and-glide pattern. This procedure was continued until the fish could no longer regain position in the tunnel and would not perform burst-and-glide swimming even at reduced speeds. The average time to exhaustion was ∼15 min. The objective of the chosen exercise protocol was to swim a fish to the point at which it could no longer perform white muscle powered burst style swimming. At exhaustion, fish were then randomly assigned to one of the following treatments: exhausted or 0.5-, 1-, 2-, 4-, 8-, 12-, 16-, or 32-h recovery and eight fish were sampled per treatment. To sample exhausted fish, individuals were immediately removed from the swim tunnel without air exposure and anesthetized with an overdose of benzocaine (0.5 g L− 1). At complete anesthesia (b1 min) fish were removed from the water, patted dry, and weighed. A crosssection of the trunk musculature was then taken posterior to the dorsal fin and immediately freeze-clamped between two aluminum blocks cooled in liquid N2. Liver was also dissected and frozen in liquid N2. All samples were stored at –80 °C for later analysis. Tissue sampling from a single fish took less than 1 min. For recovery samples, exhausted fish were placed into 1 L exposure chambers with mesh sides and placed randomly into one of two, 250 L aquarium containing well-aerated City of Vancouver tapwater at 10 °C. Two water pumps were situated in each aquarium to ensure adequate mixing of the water. Exposure chambers were designed to fit smoothly into basins that were slightly larger than the chambers and allowed the fish to be removed from the recovery tank without inducing agitation or exposing the fish to air. At the predefined recovery times, a chamber containing a fish was removed from a tank and an overdose of benzocaine (0.5 g L− 1) was added to the basin. Muscle and liver were sampled according to the procedures outlined above. A total of eight fish were sampled at each time point, with four samples from each recovery tank. Resting control samples were obtained by placing goldfish into the individual, 1 L exposure chambers and submerging the chambers into the same aquaria used for recovery. After 24 h in the aquarium, resting control samples were obtained as described above. 2.2.2. Anoxia exposure and recovery The day before experimentation, goldfish were transferred into individual exposure chambers and placed into one of two, 250 L aquarium containing well-aerated tapwater at 10 °C (identical setup to that used for exercise recovery). After a 24 h acclimation period, a random chamber containing a fish was removed from a tank and an overdose of benzocaine (0.5 g L− 1) was added to the each basin. At complete anesthesia (b1 min) the fish was removed and tissues sampled as described above. Four exposure chambers were sampled from each aquarium for a total of eight fish per sampling time. Following normoxia sampling, severe hypoxia was induced by bubbling N2 into the aquaria containing the exposure chambers and covering the surface with plastic. Water [O2] decreased from normoxia to b0.1 mg O2 L− 1 over the first 1.5 h of bubbling and then was maintained at b0.1 mg O2 L− 1 for 10 h. Water O2 was monitored inside the exposure chambers using a dissolved oxygen meter (DO6
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dissolved oxygen meter; Oaktron). At 10 h hypoxia exposure, four fish from each aquarium were randomly selected and tissues sampled as described above. Following tissue sampling at 10 h exposure to severe hypoxia, the water was vigorously aerated and returned to normoxic levels (N8 mg O2 L− 1) within 20 min and four fish from each aquaria were sacrificed and tissues sampled at 0.5-, 1-, 2-, 4-, 8-, 12-, 16-, and 32-h recovery as described above. A total of eight fish, four from each aquarium, were sampled at each time point. 2.2.3. Whole animal ethanol production To determine whole-animal ethanol production rates during severe hypoxia exposure and recovery from hypoxia or exhaustive exercise, we monitored ethanol production in goldfish using closed respirometry. Briefly, to determine whole-animal ethanol production rates during hypoxia exposure, individual fish were placed into size-matched respirometers (1 g to 10 mL of water) and allowed to acclimate to the respirometer for 24 h while the respirometer received flow-through well-aerated water at a rate of 50 mL min− 1. The small fish mass to volume ratio was required to ensure that ethanol production could be monitored. Respirometers were maintained at 10 °C by placing them in a wet table supplied with temperature regulated water. After the acclimation period, the water flow into each respirometer was stopped and the water O2 lowered to b0.1 mg O2 L− 1 over a 1 h period by bubbling N2 gas into the respirometer. Once water O2 was at the desired level, the respirometer was sealed and maintained for 10 h. Water samples (200 μL) were taken at time-zero (immediately after sealing the respirometer) and every 2 h during the 10 h anoxia exposure via a three-way stopcock and a gas-tight syringe. A larger water sample (∼1 ml) was taken at 10 h and analyzed for O2 to ensure water O2 was b0.1 mg O2 L− 1. At the end of the respirometry period, fish were removed, weighed, and returned to a separate stock tank. Water samples were analyzed for ethanol as described below and ethanol flux rates were calculated as μmol ethanol g− 1 of fish h− 1. Ethanol production rates were also determined during recovery from hypoxia and exhaustive exercise. For hypoxia recovery, fish were exposed to 10 h severe hypoxia as described above and at time zero of recovery, the respirometer was opened, flushed heavily with wellaerated dechlorinated tapwater at 10 °C, and an airline inserted to maintain water O2. The respirometers were then kept open to the atmosphere with aeration and no water flow for the 32 h recovery. Water samples (200 μL) were taken immediately after flushing and at 1-, 2-, 4-, 8-, 24-, and 32-h during recovery for ethanol determination. Ethanol flux rates were determined from the difference in water [ethanol] before and after each flux period and corrected for respirometer volume, fish weight, and time. Similar protocols were used to determine the ethanol production rates following exhaustive exercise. Briefly, individual fish were exercised to exhaustion as described above and immediately placed into the same well-aerated respirometers used for the severe hypoxia trial and water samples taken over the same time intervals as described above for hypoxia recovery. To control for the potential loss of ethanol from the water to the atmosphere during our recovery trials, control trials were performed where a respirometer without a fish water was spiked with 1 mM ethanol and held under identical conditions to those used for recovery (e.g. with aeration and at 10 °C). Water [ethanol] was determined over the same time intervals as in the recovery experiment. In these control trials, there were no changes in [ethanol] over 32 h, therefore the results obtained are not due to ethanol loss to the atmosphere. 2.3. Analytical protocols Frozen muscle and liver were broken into pieces in an insulated mortar and pestle cooled in liquid N2 and stored at −80 °C for metabolite extraction. A portion of the frozen muscle (∼100 mg) was ground into a fine powder under liquid N2 and used for intracellular pH (pHi) measurements as described by Pörtner et al. (1991) using a
Fig. 1. Ethanol production rates by goldfish during 10 h exposure to severe hypoxia (b0.1 mg O2 L− 1). Values are means ± SEM (n = 8). Bars with different letters are significantly different (P ≤ 0.05).
thermostatted Radiometer G297/G2 capillary microelectrode with PHM71 acid-base analyzer. For the extraction of metabolites, pieces of frozen muscle or liver (∼100 mg) were weighed into borosilicate tubes containing 1 mL of icecold 1 M HClO4 and immediately homogenized at the highest speed of a Polytron homogenizer for 30 s at 0 °C. Homogenates were transferred to 1.5 mL microcentrifuge tubes, centrifuged for 5 min at 20,000 g at 4 °C, and the supernatant neutralized with 3 M Tris-base. Muscle and liver extracts were assayed immediately for ethanol using a commercial kit (Diagnostic Chemicals Ltd, PEI, Canada). Muscle extracts were also analyzed for ATP, CrP, and lactate, while liver extracts were analyzed for lactate using methods described in Bergmeyer (1983). For determination of muscle and liver glycogen, ∼20 mg of tissue was digested in 1 mL of 30% KOH at 100 °C. Glycogen was isolated as described by Hassid and Abraham (Hassid and Abraham, 1957) and free glucose determined after digestion with amyloglucodisase (Bergmeyer, 1983) and was reported as μmol glucosyl units g− 1 wet mass. 2.4. Data presentation and statistical analysis All data are presented as means ± SE (n = 8 at each data point). Statistical analysis consisted of two-way analysis of variance (ANOVA) with treatment (severe hypoxia and exhaustive exercise) and time as independent variables. Where necessary, data were log transformed in order to meet the assumptions of normality and homogeneity of variance. When interaction terms were not significant, post-hoc analysis was performed among groups using the Holm–Sidak comparison method to a defined control group. If the interaction terms were significant the data were separated and analyzed independently using a one-way ANOVA. Rate constants for recovery were determined on mean data using a monoexponential function. All statistical decisions were based on P ≤ 0.05. 3. Results No mortality was observed during any experimental trial. Approximately ∼ 15% of the goldfish lost equilibrium during the severe hypoxia exposure, but all recovered and survived through to their pre-assigned sampling time. 3.1. Whole animal ethanol production There was an increase in whole-animal ethanol production rate over the first 4 to 6 h of severe hypoxia exposure (b0.1 mg O2 L− 1) then ethanol production leveled off (Fig. 1). The total ethanol production
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during the 10 h exposure to hypoxia was 12.1 μmol g− 1 of fish. During the first 4 h of recovery from severe hypoxia exposure, there was a sustained efflux of ethanol from the goldfish followed by rates of ethanol production that could not be distinguished from zero (Fig. 2A). Ethanol production at rest was negligible and below the detection limit of our assay. During recovery from exhaustive exercise, ethanol was released from the fish to the water over the first 1 to 2 h of recovery and followed immediately by a period between 2 and 4 h recovery that was characterized by the loss of ethanol from the respirometer. At recovery times greater than 4 h, ethanol production rates were not distinguishable from zero (Fig. 2B). 3.2. White muscle There was no significant effect of treatment (hypoxia exposure or exhaustive exercise) or time on white muscle [ATP] (Fig. 3A). There was no significant effect of treatment on white muscle [PCr], but there was a significant effect of time (Fig. 3B). White muscle [PCr] decreased in response to both hypoxia exposure and exhaustive exercise and returned to values that were not different than controls by 0.5 h recovery. There was a significant effect of treatment and time on white muscle pHi (Fig. 3C). White muscle pHi differed in the resting/ normoxic groups of both treatments (Fig. 3C) and decreased in response to hypoxia exposure and exhaustive exercise. However, the magnitude of the decrease in white muscle pH was larger (0.31 pH units) after exhaustive exercise compared with that observed after 10 h hypoxia exposure (0.25 pH units). In addition, the rate of recovery
Fig. 3. White muscle ATP (A), PCr (B), and pHi (C) in goldfish at rest (R) and during 32 h recovery from severe hypoxia exposure (unfilled circles) or exhaustive exercise (filled circles). Vertical gray bar represents either 10 h severe hypoxia exposure or 15 min of exhaustive exercise. Values are means ± SEM (n = 8 for each point). Within each treatment (hypoxia or exercise), significant differences from resting are indicated by an asterisk (P ≤ 0.05).
Fig. 2. Ethanol production rates by goldfish during recovery from 10 h severe hypoxia (A) or 15 min exhaustive exercise (B). Time zero indicates the start of recovery. Positive values for ethanol production indicate ethanol release to water while negative values indicate ethanol loss from the water. Significant differences from zero are indicated by an asterisk (P ≤ 0.05).
was faster after hypoxia exposure compared with exhaustive with recovery rate constants of 1.35 h− 1 and 0.65 h− 1, respectively. Exposure to severe hypoxia and exhaustive exercise caused significantly different responses in muscle [glycogen] (two-way ANOVA; Fig. 4A). White muscle [glycogen] decreased as a result of hypoxia exposure and remained depressed compared with resting/ normoxic goldfish over the first 2 h recovery returning to values that were not significantly different from controls between 4 and 12 h with the exception of 16 h recovery, which was lower than the normoxic controls. In contrast, exhaustive exercise and recovery had no significant effect on white muscle [glycogen] (Fig. 4A). There was no effect of treatment on white muscle [lactate], but in both groups there was a significant effect of time (Fig. 4B). White muscle [lactate] increased to roughly equivalent levels as a result of hypoxia exposure and exercise and returned to values equivalent to the resting/normoxic controls over the same time period. There was a significant effect of treatment and time on white muscle [ethanol] (Fig. 4C). White muscle [ethanol] increased ∼40 fold during 10 h severe hypoxia exposure and remained elevated for the first 2 h of recovery followed by a return to levels that were not elevated compared with control fish. In contrast, exhaustive exercise had no effect on white muscle [ethanol], but during recovery,
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4. Discussion
Fig. 4. White muscle glycogen (A), lactate (B), and ethanol (C) in goldfish at rest (R) and during 32 h recovery from severe hypoxia exposure (unfilled circles) or exhaustive exercise (filled circles). See Fig. 3 caption for more details.
muscle [ethanol] increased gradually to levels that were significantly elevated at 16 h compared with control fish. 3.3. Liver There was a significant effect of treatment and recovery time on liver [glycogen] (Fig. 5A). Severe hypoxia exposure itself had no effect on liver glycogen, but within 0.5 h recovery, liver [glycogen] decreased by ∼ 35%, remained depressed compared with control samples until 2 h recovery and returned to values that were not significantly different than controls by 4 h recovery. Exhaustive exercise alone also had no effect on liver [glycogen]; however, in contrast to the severe hypoxia treatment, liver [glycogen] increased transiently during the first 2 h of recovery only to return to values that were not different than controls by 4 h recovery. Severe hypoxia exposure and exhaustive exercise yielded different responses in liver [ethanol], with a significant effect of treatment and time on liver [ethanol] (Fig. 5B). Liver [ethanol] increased ∼ 7.7 fold during severe hypoxia exposure and remained elevated compared with controls for 2 h recovery, subsequently returning to values that were not elevated compared with control fish. In contrast, there was no significant effect of exhaustive exercise or recovery on liver [ethanol].
Goldfish have long been recognized as unique among vertebrates for their ability to produce ethanol and CO2 as the major fermentative products of anaerobic metabolism (Shoubridge and Hochachka, 1980). The ethanol-producing pathway in goldfish is primarily isolated to the skeletal muscle due to the high tissue-specific alcohol dehydrogenase expression (Shoubridge and Hochachka, 1980; Mourik et al., 1982) and it is believed that lactate produced in other tissues during periods of anaerobiosis are shuttled to the skeletal muscle for ethanol biosynthesis. Ethanol is a small, highly diffusible molecule that once produced is released to the water via the gills, preventing self-intoxication and lactate buildup. Ethanol production, in combination with the large liver and muscle glycogen reserves, and the ability to undergo metabolic rate depression are thought to be the primary “adaptive” traits explaining how goldfish can survive weeks to months of complete anoxia. As pointed out by Shoubridge and Hochachka (1980), the disadvantage of ethanol production during anaerobiosis is that it is wasteful of carbon and may negatively impact the ability of goldfish to recover from metabolic insult. In the present study, we compared the metabolic responses in muscle and liver of goldfish before and after exposure to severe hypoxia or exhaustive exercise, as well as during recovery. We hypothesized that during hypoxia exposure, glycogen conversion to ethanol would predominate, while during a short bout of exhaustive exercise, glycogen conversion to lactate would predominate. Indeed substantial ethanol production was observed during the 10 h exposure to severe hypoxia (Fig. 1) as well as during the first 4 h recovery (Fig. 2A), which limited the extent of muscle glycogen recovery. However direct comparisons between the rates of glycogen recovery after hypoxia exposure and exhaustive exercise are not possible because goldfish do not preferentially deplete glycogen during exhaustive exercise. Despite this difficulty, comparisons of recovery from hypoxia exposure and exhaustive exercise in goldfish have yielded three novel findings. First, ethanol appears to be a ubiquitous metabolic end-product when tissue lactate levels are elevated either by severe hypoxia exposure or exhaustive exercise. Second, ethanol is not only released to the environment, but also appears to be taken up from the environment during recovery from exhaustive exercise. Third, differential responses in hepatic glycogen were observed during recovery from hypoxia exposure compared with exhaustive exercise, suggesting that the liver may play different roles during recovery from these two metabolic insults. The initial goal of our experiment was to reduce white muscle [glycogen] in goldfish to the same level by exposure to 10 h severe hypoxia exposure or exhaustive exercise and examine the rates of metabolic recovery. We successfully reduced white muscle [glycogen] by ∼ 60% (Fig. 4A) with hypoxia exposure, but we were unable to cause a significant decrease in white muscle glycogen with exhaustive exercise. The general picture of how fish white muscle powers exhaustive exercise suggests an almost complete depletion of all available fuels (ATP, PCr, and glycogen) and the accumulation of high levels of metabolic waste (e.g. lactate and H+); however, it must be acknowledged that this metabolic picture of exercise is primarily based on studies performed on athletic salmonids (Wang et al., 1994; Milligan, 1996; Richards et al., 2002a). Responses to exercise are heavily influenced by fish species and choice of exhaustive exercise protocol (Moyes and West, 1995), with salmonids being among the most willing to exhaust metabolic fuels. In goldfish however, complete exhaustion was characterized by a large reduction in white muscle [PCr] (Fig. 3B), tissue acidification (Fig. 3C), only modest accumulation of [lactate] (Fig. 4B) and no significant drop in muscle [glycogen] (Fig. 4A). Numerous preliminary trials were performed using several exercise protocols (manual chasing and various swim tunnel protocols) and rarely was substantial glycogen depletion noted. Indeed, the majority of studies that have examined the effects of intense exercise
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Fig. 5. Liver glycogen (A) and ethanol (B) in goldfish at rest (R) and during 32 h recovery from severe hypoxia exposure (unfilled circles) or exhaustive exercise (filled circles). See Fig. 3 caption for more details.
on muscle metabolism of Cyprinids, suggest that the metabolic paradigm associated with fish exercise does not hold true (Driedzic and Hochachka, 1976; van Ginneken et al., 2004). The common carp (Cyprinis carpio), for example, showed only a ∼13 μmol g− 1 increase in lactate during an exhaustive exercise protocol (Sugita et al., 2001), which is comparable to the present study, but far lower than that obtained using a similar protocol on trout (e.g. up to 30 μmol g-1 in trout; Richards et al., 2002b). Furthermore, Huber and Guderley (1993) demonstrated in goldfish that exhaustive exercise yielded limited muscle lactate accumulation (∼ 13 μmol g− 1 wet mass). Although exhaustive exercise only caused a non-significant decrease in white muscle [glycogen] of the ∼8 μmol glucosyl units g− 1 wet mass (Fig. 4A) this decrease was more than sufficient to account for the ∼ 8 μmol g− 1 wet mass accumulation of lactate. The reasons for muscle exhaustion remain unknown, but in our experiments the profound drop in pHi (Fig. 3C) or depletion of muscle [PCr] (Fig. 3B) are the likely causes of exhaustion. During hypoxia exposure, goldfish are well known for their ability to reduce metabolic rate by up to 60% (measured as heat dissipation; Van Waversveld et al., 1988, 1989), thus reducing whole animal ATP turnover and extending survival. Despite this impressive ability to reduce whole animal ATP turnover, an activation of substrate-level phosphorylation, primarily glycolysis using stored glycogen as a substrate, is still necessary to supply ATP and maintain cellular energy balance. The two major glycogen stores available to support whole animal metabolism during hypoxia exposure are located in the liver and white muscle. Surprisingly, there was no effect of 10 h severe hypoxia exposure on liver [glycogen] in the present experiment, which was contrary to expectation and to that observed by other authors (e.g. van den Thillart et al., 1980b). van den Thillart et al. (1980b) showed a transient decreases in liver [glycogen] during hypoxia exposure in goldfish and a more severe depletion during exposure to anoxia. The possible reasons for the lack of liver glycogen mobilization could be due to the relatively short duration of the hypoxia exposure as well as
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the fact that we did not expose our goldfish to complete anoxia, but rather severe hypoxia (b0.1 mg O2 L− 1). Independent of the reasons why, our data strongly support the notion that muscle glycogen stores are sacrificed preferentially over liver glycogen stores to support whole animal glycolysis. White muscle glycogen decreased by 19.5 μmol glucosyl units g− 1 wet mass (a 60% decrease compared with normoxic animals; Fig. 4A) and the majority of this glycogen depletion can be accounted for by lactate and ethanol accumulation. Our ethanol excretion rates are comparable to those reported by Johnston and Bernard (1983) for crucian carp exposed to anoxia at 15 °C with a maximum rate of ∼ 3 μmol g− 1 h− 1, but both the present study and others (Johnston and Bernard, 1983) report slightly higher excretion rates than van den Thillart (1983; maximum noted ∼ 1 μmol g− 1 h− 1). In the present study, white muscle glycogen is the primary substrate for whole ethanol production during hypoxia and we can nearly fully account for all the ethanol production by the depletion in white muscle glycogen. Assuming 60% of a goldfish is white muscle (estimate for salmonids; Moyes and West, 1995) then the observed muscle glycogen depletion, on a whole-animal basis is equivalent to 292.5 μmol glucosyl units (assuming an average fish mass of 25 g). Accumulated muscle lactate accounts for ∼19% of the glycogen depletion (7.3 μmol g− 1 muscle; 109.5 μmol total in 15 g of tissue), while the total ethanol production and accumulation in the tissues (assuming the measured muscle ethanol concentrations are equivalent to all cellular concentrations because of the high lipid solubility of ethanol) accounts for an additional 31% of the glycogen depletion. Furthermore, if we assume that the rapid efflux of ethanol observed during the first 4 h of recovery represent efflux of ethanol accumulated during the 10 h hypoxia exposure, then this accounts for an additional 18% of the glycogen depletion (107 μmol). In total, ethanol production accounts for a minimum of 50% of the total glycogen depleted during severe hypoxia exposure and we can account for 70% of the glycogen depletion by the accumulated lactate and ethanol. The remaining 30% of unaccounted for glycogen depletion maybe due to preferential accumulation of ethanol in different tissues or errors in our estimates of the percent of body weight occupied by muscle. During the recovery period, white muscle PCr (Fig. 3B) and pHi (Fig. 3C) recovered quickly (within 0.5 h) following hypoxia exposure and exhaustive exercise, while muscle lactate recovery took N12 h to return to resting/normoxic level following exhaustive exercise and hypoxia exposure (Fig. 4B). Differences in the rate of white muscle pHi recovery following hypoxia exposure and exhaustive exercise were likely due to the differences in magnitude of the pHi disturbance between the two treatments and the degree of perturbation brought about by the treatments. Lactate disappearance after exercise was not associated with increases in muscle glycogen, suggesting that the accumulated muscle lactate may, at least in part, be the substrate for the transient increase in whole animal ethanol production observed from 1 to 2 h recovery (Fig. 2B). It is surprising however, that only a modest increase in white muscle [ethanol] was observed (and the only significant increase occurred at 16 h recovery; Fig. 4C) suggesting that the tissue-specific source of ethanol may not be white muscle, but rather red muscle. Red muscle alcohol dehydrogenase activity is about 4 times higher than white muscle (Mourik et al., 1982), therefore lactate may be shuttled from the white muscle to the red for conversion to ethanol for subsequent release to the environment. Our ethanol flux measurements obtained after exhaustive exercise suggests that ethanol may be taken up from the environment and utilized by the fish during recovery. It is intriguing to speculate that the ethanol disappearance from the respirometer observed between 2 and 4 h recovery from exercise may represent uptake for use as a carbon substrate for glycogen synthesis or for ATP production. For either to occur, the pyruvate dehydrogenase complex would need to accept acetaldehyde as a substrate and facilitate its movement to either acetylCoA, for ATP production, or reversed to pyruvate, for use in glycogenesis.
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Further research into the regulation of the pyruvate dehydrogenase complex in goldfish muscle during recovery is needed to determine whether these possibilities are feasible. At this time however, our measurements of ethanol flux during recovery must be regarded as preliminary and await confirmation. If ethanol is a dynamic metabolite that can serve as a substrate for metabolism, then recovery of fish in an ethanol rich environment should enhance the rate of recovery. This is unlikely to be of a major consequence in natural populations of goldfish as ethanol is unlikely to accumulate in their environment to levels that would enhance recovery, but it remains an intriguing biochemical mystery worthy of further investigation. Liver glycogen stores responded in a dramatically different fashion during the early potions of recovery from severe hypoxia versus exhaustive exercise (Fig. 5A). During the very early portion of hypoxia recovery (0 to 0.5 h), mobilization of liver glycogen and release of glucose into the blood may be directly related to the early recovery of muscle glycogen observed between 1 and 2 h recovery or to facilitate recovery of other organs. Unlike exercise, which primarily affects muscle, hypoxia exposure affects the whole body and therefore organs not examined here (e.g. brain, heart, and gills). Thus during recovery, enhanced glucose release to the blood may be required to facilitate the recovery of other metabolically active tissues. The recovery of liver glycogen between 1 and 4 h is more of a mystery. Either the glucose released during the early portion of recovery is resynthesized into glycogen later on, or alternate carbon sources are used to resynthesize the glycogen. It remains intriguing to speculate that accumulated ethanol, may in part, account for some, albeit a potentially small fraction of the glycogen recovery after exhaustive exercise or hypoxia exposure. Partitioning of radiolabelled ethanol during recovery may help to address this question. In contrast to the liver responses observed during recovery from hypoxia, liver glycogen transiently increased by 1.7 fold during the first 1 to 2 h recovery from exhaustive exercise (Fig. 5A) suggesting that the liver is sequestering carbohydrates from the blood and other tissues, although again, this occurs in a transient fashion with almost compete recovery to control values within 4 h recovery. Overall, the transient changes in liver glycogen during the early phase of recovery may simply point to an enhanced total body turnover where liver serves as a temporary source or storage reserve for whole body carbohydrates. Acknowledgements This work was funded by a Natural Sciences and Engineering Research Council (NSERC) of Canada Discovery Grant to JGR. MM was supported by an NSERC Post Graduate Scholarship. References Bergmeyer, H.U., 1983. Methods of Enzymatic Analysis. Academic Press, New York. Blazka, P., 1958. The anaerobic metabolism of fish. Physiol. Zool. 31, 117–128. Dobson, G.P., Hochachka, P.W., 1987. Role of glycolysis in adenylate depletion and repletion during work and recovery in teleost white muscle. J. Exp. Biol. 129, 124–140. Driedzic, W.R., Hochachka, P.W., 1976. Control of energy metabolism in fish white muscle. Am. J. Physiol. 230, 579–582. Hassid, W., Abraham, S., 1957. Chemical procedures for analysis of polysaccharides. Methods Enzymol. 3, 34–37.
Huber, M., Guderley, H., 1993. The effect of thermal acclimation and exercise upon the binding of glycolytic enzymes in muscle of the goldfish Carassius auratus. J. Exp. Biol. 175, 195–209. Johnston, I.A., Bernard, L.M., 1983. Utilization of the ethanol pathway in carp following exposure to anoxia. J. Exp. Biol. 104, 73–78. Lutz, P.L., Nilsson, G., 1997. Contrasting strategies for anoxia brain survivial — glycolysis up or down. J. Exp. Biol. 200, 411–419. Milligan, C.L., 1996. Metabolic recovery from exhaustive exercise in rainbow trout. Comp. Biochem. Physiol. A 113, 51–60. Mourik, J., Raeven, P., Steur, K., Addink, A.D.F., 1982. Anaerobic metabolism of red skeletal-muscle of goldfish, Carassius auratus (L) — mitochondrial produced acetaldehyde as anaerobic electron-acceptor. FEBS Letts. 137, 111–114. Moyes, C.D., West, T.G., 1995. In: Hochachka, P.W., Mommsen, T.P. (Eds.), Exercise metabolism of fish. . Molecular Biology of Fishes, vol. 4. Elsevier Science, Amsterdam, pp. 367–392. Pörtner, H., Boutlier, R., Tang, Y., Toews, D., 1991. The use of fluoride and nitriloacetic acid in tissue acid-base physiology. II. Intracellular pH. Respir. Physiol. 81, 255–275. Richards, J.G., Heigenhauser, G.J.F., Wood, C.M., 2002a. Glycogen phosphorylase and pyruvate dehydrogenase transformation in white muscle of trout during highintensity exercise. Am. J. Physiol. 282, R828–R836. Richards, J.G., Heigenhauser, G.J.F., Wood, C.M., 2002b. Lipid oxidation fuels recovery from exhaustive exercise in white muscle of rainbow trout. Am. J. Physiol. 282, R89–R99. Richards, J.G., Heigenhauser, G.J.F., Wood, C.M., 2003. Exercise and recovery metabolism in the Pacific spiny dogfish (Squalus acanthias). J. Comp. Physiol. B 173, 463–474. Richards, J.G., Wang, Y.S., Brauner, C.J., Gonzalez, R.J., Patrick, M.L., Schulte, P.M., Choppari-Gomes, A.R., Almeida-Val, V.M., Val, A.L., 2007. Metabolic and ionoregulatory responses of the Amazonian cichlid, Astronotus ocellatus, to severe hypoxia. J. Comp. Physiol. B 177, 361–374. Scarabello, M., Heigenhauser, G.J., Wood, C.M., 1991. The oxygen debt hypothesis in juvenile rainbow trout after exhaustive exercise. Respir. Physiol. 84, 245–259. Schulte, P.M., Moyes, C.D., Hochachka, P.W., 1992. Integrating metabolic pathways in post-exercise recovery of white muscle. J. Exp. Biol. 166, 181–195. Shoubridge, E.A., Hochachka, P.W., 1980. Ethanol: novel end product of vertebrate anaerobic metabolism. Science 209, 308–309. Sugita, T., Shimeno, S., Nakano, N., Hosokawa, H., Masumoto, T., 2001. Response of enzyme activities and metabolic intermediate concentrations to a long burst of exercise and following resting in muscle and the hepatopancreas of carp. Fish Sci. 67, 904–911. van den Thillart, G., Verbeek, R., 1991. Anoxia-induced oxygen debt of goldfish (Carassius auratus L). Physiol. Zool. 64, 525–540. van den Thillart, G., Kesbeke, F., van Waarde, A., 1980a. Anaerobic energy metabolism of goldfish, Carassius auratus (L): influence of hypoxia and anoxia on phosphorylated compounds and glycogen. J. Comp. Physiol. 136, 45–52. van den Thillart, G., Kesbeke, F., van Waarde, A., 1980b. Anaerobic energy-metabolism of goldfish, Carassius auratus (L): influence of hypoxia and anoxia on phosphorylated compounds and glycogen. J. Comp. Physiol. 136, 45–52. van den Thillart, G., Van Berge-Henegouwen, M., Kesbeke, F., 1983. Anerobic metabolism of goldfish, Carassius auratus (L.): ethanol and CO2 excretion rates and anoxia tolerance at 20, 10, and 5 °C. Comp. Biochem. Physiol. A 76, 295–300. van den Thillart, G., van Waarde, A., Muller, H., Erklens, C., Addink, A., Lugtenberg, J., 1989. Fish muscle energy metabolism measured by in vivo 31P-NMR during anoxia and recovery. Am. J. Physiol. 256, R922–R929. van Ginneken, V., van den Thillart, G., Addink, A., Erkelens, C., 1995. Fish muscle energy metabolism measured during hypoxia and recovery: an in vivo 31P-NMR study. Am. J. Physiol. 268, R1178–1187. van Ginneken, V., Boot, R., Murk, T., van den Thillart, G., Balm, P., 2004. Blood plasma substrates and muscle lactic-acid response after exhaustive exercise in common carp and trout: indications for a limited lactate-shuttle. Anim. Biol. 54, 119–130. van Waarde, A., van den Thillart, G., Verhagen, M., 1993. Ethanol formation and pH regulation in fish. In: Hochachka, P.W., Lutz, P.L., Sick, T., Rosenthal, M., van den Thillart, G. (Eds.), Surviving hypoxia: Mechanisms of control and adaptation. CRC Press, Boca Raton, Fl., pp. 157–170. Van Waversveld, J., Addink, A.D., Van den Thillart, G., Smit, H., 1988. Anaerobic heat production measurements: a new perspective. J. Exp. Biol. 138, 529–533. Van Waversveld, J., Addink, A.D.F., van den Thillart, G., 1989. Simultaneous direct and indirect calorimetry on normoxic and anoxic goldfish. J. Exp. Biol. 142, 325–335. Walker, S.C., Johanson, P.H., 1977. Anaerobic metabolism in goldfish Carassius auratus. Can. J. Zool. 55, 1304–1311. Wang, Y., Heigenhauser, G.J.F., Wood, C.M.,1994. Integrated responses to exhaustive exercise and recovery in rainbow trout white muscle: acid–base, phosphogen, carbohydrate, lipid, ammonia, fluid volume and electrolyte metabolism. J. Exp. Biol. 195, 227–258.
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Comparative Biochemistry and Physiology, Part C j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / c b p c
Metabolic recovery in goldfish: A comparison of recovery from severe hypoxia exposure and exhaustive exercise☆ Milica Mandic, Gigi Y. Lau, Manu M.S. Nijjar, Jeffrey G. Richards ⁎ Department of Zoology, The University of British Columbia, Vancouver, British Columbia, V6T 1Z4 Canada
A R T I C L E
I N F O
Article history: Received 9 March 2008 Received in revised form 28 April 2008 Accepted 29 April 2008 Available online 10 May 2008 Keywords: Glycogen Ethanol Lactate Liver Muscle Anoxia Carassius auratus
A B S T R A C T Severe hypoxia exposure and exhaustive exercise in goldfish both elicit a strong activation of substrate-level phosphorylation with the majority of the metabolic perturbations occurring in the white muscle. Approximately half of the muscle glycogen breakdown observed during severe hypoxia exposure was accounted for by ethanol production and loss to the environment, which limited the extent of muscle glycogen recovery when animals were returned to normoxic conditions. Ethanol production in goldfish is not solely a response to anoxia/hypoxia exposure however, as a transient increase in ethanol production was observed during the early stages of recovery from exhaustive exercise. These data suggest that ethanol production is a ubiquitous “anaerobic” end product, which accumulates whenever metabolic demands exceed mitochondrial oxidative potential. Exhaustive exercise and hypoxia exposure both caused a 7 to 8 μmol g− 1 wet mass increase in muscle [lactate] and the rates of recovery following these perturbations were similar. The rates of muscle PCr and pHi recovery after hypoxia exposure and exhaustive exercise were similar with levels returning to controls values within 0.5 h. Surprisingly, liver [glycogen] was not depleted during exposure to severe hypoxia, however, during recovery from both hypoxia and exercise dramatically different responses in liver [glycogen] were noted. During the early stages of recovery, liver [glycogen] transiently increased to high levels after exhaustive exercise, while during recovery from hypoxia there was a transient decrease in liver glycogen over the same time frame. Overall, this points to the liver playing a dramatically different role in facilitating recovery from exercise compared with hypoxia exposure. © 2008 Elsevier Inc. All rights reserved.
1. Introduction Energy production during exposure to severe hypoxia and exhaustive exercise occurs primarily via substrate-level phosphorylation (phosphocreatine (PCr) hydrolysis and glycolysis), which compensates for ATP demands that exceed oxidative capacity. During severe hypoxia exposure, goldfish suppress metabolic rate by up to 60% (Van Waversveld et al., 1989) and rely on large liver and muscle glycogen reserves to support substrate-level phosphorylation to maintain cellular energy balance (Lutz and Nilsson, 1997). In contrast, ATP demands in muscle during exhaustive exercise increase beyond the oxidative capacity and thus substrate-level phosphorylation must be activated to meet the energetic demands of muscle contraction (Richards et al., 2002a). Although the metabolic reasons for the activation of substrate-level phosphorylation differ during exposure to
☆ Contribution to the Special Issue of CBP on Chinese Comparative Biochemistry and Physiology presented at or related to the International Conference of Comparative Physiology, Biochemistry and Toxicology and the 6th Chinese Comparative Physiology Conference, October, 10–14, 2007, Zhejiang University, Hangzhou, China. ⁎ Corresponding author. Tel.: +1 604 822 2381; fax: +1 604 822 2416. E-mail address: [email protected] (J.G. Richards). 1532-0456/$ – see front matter © 2008 Elsevier Inc. All rights reserved. doi:10.1016/j.cbpc.2008.04.012
severe hypoxia and exhaustive exercise, the end metabolic profiles observed in muscle are similar with low [glycogen] and [PCr] and high [lactate] and metabolic [H+] (van den Thillart et al., 1989; Richards et al., 2002a, 2003, 2007). During recovery from hypoxia and exhaustive exercise, pathways must be activated to resynthesize ATP, PCr, and glycogen. To this end, the early portions of recovery are characterized by a period of excess O2 consumption (Scarabello et al., 1991; van den Thillart and Verbeek, 1991; van Ginneken et al., 1995), which in part represents a stimulation of oxidative phosphorylation to support the energetic costs of recovery. Generally, muscle [PCr] returns to resting values quickly during recovery from both hypoxia exposure and exhaustive exercise (within 1 to 2 h if not sooner), but recovery of muscle [lactate] and [glycogen] requires a longer period (N2 h; Richards et al., 2002b). Lactate is preferentially retained in muscle during recovery from exercise or hypoxia exposure (Richards et al., 2007) and there is accumulating evidence to suggest that muscle glycogenesis occurs in situ and relies on lactate as the substrate (Schulte et al., 1992). Members of the family Cyprinidae, especially the crucian carp (Carassius carassius L.) and the common goldfish (Carassius auratus L.), have long been recognized as unique among vertebrates for their
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incredible anoxia tolerance. The common goldfish, for example, can survive complete anoxia at room temperature (20 °C) for many hours (van den Thillart et al., 1983) and as environmental temperature decreases, survival can be extended to weeks or possibly even months (Blazka, 1958; Walker and Johanson, 1977). Prolonged anoxia survival at cold temperatures is likely related to three main factors: 1. the storage of high concentrations of fermentable substrates (e.g. liver [glycogen] of ∼ 800 μmol g− 1 wet mass; van den Thillart et al., 1980a), 2. the suppression of whole-animal metabolic rate (Van Waversveld et al., 1989), and 3. the ability to convert anaerobically derived lactate to ethanol and CO2 (Shoubridge and Hochachka, 1980). Ethanol production by the crucian carp and goldfish begins with the movement of lactate from other tissues to red and white skeletal muscle, where lactate is converted to pyruvate via lactate dehydrogenase. Pyruvate is then decarboxylated to acetaldehyde and CO2 via an inefficient or modified pyruvate dehydrogenase complex (van Waarde et al., 1993; Lutz and Nilsson, 1997). Typically, PDH converts pyruvate to acetyl-CoA with acetaldehyde as a bound intermediate, but during anerobiosis acetaldehyde “leaks” out of the PDH complex, diffuses into the cytoplasm and is converted to ethanol by alcohol dehydrogenase. Ethanol then freely diffuses across the gills, limiting lactate accumulation. Loss of ethanol to the environment, however, can represent a substantial loss of carbon, especially over months of anoxia exposure, and this may seriously impact the ability of the animal to recover in a normoxic environment. Few studies have examined the effects of exhaustive exercise on Cyprinids, and those that have mostly focused on the non-ethanol producing common carp (Cyprinus carpio; Driedzic and Hochachka, 1976; Sugita et al., 2001; van Ginneken et al., 2004) and no studies, to our knowledge, have examined whether ethanol production occurs during “anaerobic” exhaustive exercise. For the most part, the metabolic profiles observed in carp white muscle during exhaustive exercise are similar to those observed in other fish (e.g. trout), albeit the magnitude of the effects are smaller. Huber and Guderley (1993) noted significant accumulation of lactate during ∼ 30 min of exhaustive exercise in goldfish, but did not measure whether lactate was converted to ethanol. It is likely, however, that even if lactate conversion to ethanol occurs, due to the rapid rate of glycolysis necessary to support muscle contraction, lactate would be preferentially accumulated over ethanol. The objectives of this study were to compare the recovery profiles of white muscle [ATP], [PCr], [pHi], and glycogen in goldfish after exposure to 10 h of severe hypoxia at 10 °C or a bout of exhaustive exercise. We predicted that severe hypoxia exposure would lead to ethanol production and the loss of carbon to the environment and as a result, muscle glycogen recovery after hypoxia exposure would be impaired relative to exhaustive exercise where no or limited ethanol production was predicted to occur. Specifically, we examined the rate of ethanol production in goldfish during and after severe hypoxia exposure and exhaustive exercise to determine the relative amounts of ethanol produced and its relation to glycogen recovery. Changes in liver glycogen were also examined to gain insight into the potential role of the liver in fuelling metabolic recovery from hypoxia exposure and exhaustive exercise. 2. Materials and methods 2.1. Animal care Adult goldfish (Carassius auratus L.; ∼27 ± 1 g, 11.6 ± 0.4 cm; mean ± SD) were purchased from a local supplier (Delta Aquatics, Richmond, B.C., Canada) and held under flow-through conditions in 750 L tanks supplied with well-aerated, dechlorinated City of Vancouver tapwater (∼ 10 °C) for at least 1 month before experimentation. During holding, fish were fed daily with commercial goldfish flakes, but food was withheld 24 h before experimentation.
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2.2. Experimental protocols 2.2.1. Exhaustive exercise and recovery To exercise goldfish to exhaustion, four size-matched fish were placed into a Brett-style swimming respirometer supplied with wellaerated, dechlorinated tapwater at 10 °C. Fish were allowed 30 min of recovery from transfer and water velocity was maintained at b1 body lengths s− 1 during this period. Following the recovery period, fish were exhaustively exercised using a similar protocol to that described by Dobson and Hochachka (1987). Briefly, the swimming protocol began by increasing the water velocity to the maximum speed at which the fish swam in a burst-and-glide pattern (∼7 to 8 body length s− 1). This velocity was maintained until each fish fatigued and fell to the back of the swim tunnel. As each fish fatigued, it was manually returned to the current at which point the fish typically regained position and began swimming in a burst-and-glide pattern. This procedure was continued until the fish could no longer regain position in the tunnel and would not perform burst-and-glide swimming even at reduced speeds. The average time to exhaustion was ∼15 min. The objective of the chosen exercise protocol was to swim a fish to the point at which it could no longer perform white muscle powered burst style swimming. At exhaustion, fish were then randomly assigned to one of the following treatments: exhausted or 0.5-, 1-, 2-, 4-, 8-, 12-, 16-, or 32-h recovery and eight fish were sampled per treatment. To sample exhausted fish, individuals were immediately removed from the swim tunnel without air exposure and anesthetized with an overdose of benzocaine (0.5 g L− 1). At complete anesthesia (b1 min) fish were removed from the water, patted dry, and weighed. A crosssection of the trunk musculature was then taken posterior to the dorsal fin and immediately freeze-clamped between two aluminum blocks cooled in liquid N2. Liver was also dissected and frozen in liquid N2. All samples were stored at –80 °C for later analysis. Tissue sampling from a single fish took less than 1 min. For recovery samples, exhausted fish were placed into 1 L exposure chambers with mesh sides and placed randomly into one of two, 250 L aquarium containing well-aerated City of Vancouver tapwater at 10 °C. Two water pumps were situated in each aquarium to ensure adequate mixing of the water. Exposure chambers were designed to fit smoothly into basins that were slightly larger than the chambers and allowed the fish to be removed from the recovery tank without inducing agitation or exposing the fish to air. At the predefined recovery times, a chamber containing a fish was removed from a tank and an overdose of benzocaine (0.5 g L− 1) was added to the basin. Muscle and liver were sampled according to the procedures outlined above. A total of eight fish were sampled at each time point, with four samples from each recovery tank. Resting control samples were obtained by placing goldfish into the individual, 1 L exposure chambers and submerging the chambers into the same aquaria used for recovery. After 24 h in the aquarium, resting control samples were obtained as described above. 2.2.2. Anoxia exposure and recovery The day before experimentation, goldfish were transferred into individual exposure chambers and placed into one of two, 250 L aquarium containing well-aerated tapwater at 10 °C (identical setup to that used for exercise recovery). After a 24 h acclimation period, a random chamber containing a fish was removed from a tank and an overdose of benzocaine (0.5 g L− 1) was added to the each basin. At complete anesthesia (b1 min) the fish was removed and tissues sampled as described above. Four exposure chambers were sampled from each aquarium for a total of eight fish per sampling time. Following normoxia sampling, severe hypoxia was induced by bubbling N2 into the aquaria containing the exposure chambers and covering the surface with plastic. Water [O2] decreased from normoxia to b0.1 mg O2 L− 1 over the first 1.5 h of bubbling and then was maintained at b0.1 mg O2 L− 1 for 10 h. Water O2 was monitored inside the exposure chambers using a dissolved oxygen meter (DO6
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dissolved oxygen meter; Oaktron). At 10 h hypoxia exposure, four fish from each aquarium were randomly selected and tissues sampled as described above. Following tissue sampling at 10 h exposure to severe hypoxia, the water was vigorously aerated and returned to normoxic levels (N8 mg O2 L− 1) within 20 min and four fish from each aquaria were sacrificed and tissues sampled at 0.5-, 1-, 2-, 4-, 8-, 12-, 16-, and 32-h recovery as described above. A total of eight fish, four from each aquarium, were sampled at each time point. 2.2.3. Whole animal ethanol production To determine whole-animal ethanol production rates during severe hypoxia exposure and recovery from hypoxia or exhaustive exercise, we monitored ethanol production in goldfish using closed respirometry. Briefly, to determine whole-animal ethanol production rates during hypoxia exposure, individual fish were placed into size-matched respirometers (1 g to 10 mL of water) and allowed to acclimate to the respirometer for 24 h while the respirometer received flow-through well-aerated water at a rate of 50 mL min− 1. The small fish mass to volume ratio was required to ensure that ethanol production could be monitored. Respirometers were maintained at 10 °C by placing them in a wet table supplied with temperature regulated water. After the acclimation period, the water flow into each respirometer was stopped and the water O2 lowered to b0.1 mg O2 L− 1 over a 1 h period by bubbling N2 gas into the respirometer. Once water O2 was at the desired level, the respirometer was sealed and maintained for 10 h. Water samples (200 μL) were taken at time-zero (immediately after sealing the respirometer) and every 2 h during the 10 h anoxia exposure via a three-way stopcock and a gas-tight syringe. A larger water sample (∼1 ml) was taken at 10 h and analyzed for O2 to ensure water O2 was b0.1 mg O2 L− 1. At the end of the respirometry period, fish were removed, weighed, and returned to a separate stock tank. Water samples were analyzed for ethanol as described below and ethanol flux rates were calculated as μmol ethanol g− 1 of fish h− 1. Ethanol production rates were also determined during recovery from hypoxia and exhaustive exercise. For hypoxia recovery, fish were exposed to 10 h severe hypoxia as described above and at time zero of recovery, the respirometer was opened, flushed heavily with wellaerated dechlorinated tapwater at 10 °C, and an airline inserted to maintain water O2. The respirometers were then kept open to the atmosphere with aeration and no water flow for the 32 h recovery. Water samples (200 μL) were taken immediately after flushing and at 1-, 2-, 4-, 8-, 24-, and 32-h during recovery for ethanol determination. Ethanol flux rates were determined from the difference in water [ethanol] before and after each flux period and corrected for respirometer volume, fish weight, and time. Similar protocols were used to determine the ethanol production rates following exhaustive exercise. Briefly, individual fish were exercised to exhaustion as described above and immediately placed into the same well-aerated respirometers used for the severe hypoxia trial and water samples taken over the same time intervals as described above for hypoxia recovery. To control for the potential loss of ethanol from the water to the atmosphere during our recovery trials, control trials were performed where a respirometer without a fish water was spiked with 1 mM ethanol and held under identical conditions to those used for recovery (e.g. with aeration and at 10 °C). Water [ethanol] was determined over the same time intervals as in the recovery experiment. In these control trials, there were no changes in [ethanol] over 32 h, therefore the results obtained are not due to ethanol loss to the atmosphere. 2.3. Analytical protocols Frozen muscle and liver were broken into pieces in an insulated mortar and pestle cooled in liquid N2 and stored at −80 °C for metabolite extraction. A portion of the frozen muscle (∼100 mg) was ground into a fine powder under liquid N2 and used for intracellular pH (pHi) measurements as described by Pörtner et al. (1991) using a
Fig. 1. Ethanol production rates by goldfish during 10 h exposure to severe hypoxia (b0.1 mg O2 L− 1). Values are means ± SEM (n = 8). Bars with different letters are significantly different (P ≤ 0.05).
thermostatted Radiometer G297/G2 capillary microelectrode with PHM71 acid-base analyzer. For the extraction of metabolites, pieces of frozen muscle or liver (∼100 mg) were weighed into borosilicate tubes containing 1 mL of icecold 1 M HClO4 and immediately homogenized at the highest speed of a Polytron homogenizer for 30 s at 0 °C. Homogenates were transferred to 1.5 mL microcentrifuge tubes, centrifuged for 5 min at 20,000 g at 4 °C, and the supernatant neutralized with 3 M Tris-base. Muscle and liver extracts were assayed immediately for ethanol using a commercial kit (Diagnostic Chemicals Ltd, PEI, Canada). Muscle extracts were also analyzed for ATP, CrP, and lactate, while liver extracts were analyzed for lactate using methods described in Bergmeyer (1983). For determination of muscle and liver glycogen, ∼20 mg of tissue was digested in 1 mL of 30% KOH at 100 °C. Glycogen was isolated as described by Hassid and Abraham (Hassid and Abraham, 1957) and free glucose determined after digestion with amyloglucodisase (Bergmeyer, 1983) and was reported as μmol glucosyl units g− 1 wet mass. 2.4. Data presentation and statistical analysis All data are presented as means ± SE (n = 8 at each data point). Statistical analysis consisted of two-way analysis of variance (ANOVA) with treatment (severe hypoxia and exhaustive exercise) and time as independent variables. Where necessary, data were log transformed in order to meet the assumptions of normality and homogeneity of variance. When interaction terms were not significant, post-hoc analysis was performed among groups using the Holm–Sidak comparison method to a defined control group. If the interaction terms were significant the data were separated and analyzed independently using a one-way ANOVA. Rate constants for recovery were determined on mean data using a monoexponential function. All statistical decisions were based on P ≤ 0.05. 3. Results No mortality was observed during any experimental trial. Approximately ∼ 15% of the goldfish lost equilibrium during the severe hypoxia exposure, but all recovered and survived through to their pre-assigned sampling time. 3.1. Whole animal ethanol production There was an increase in whole-animal ethanol production rate over the first 4 to 6 h of severe hypoxia exposure (b0.1 mg O2 L− 1) then ethanol production leveled off (Fig. 1). The total ethanol production
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during the 10 h exposure to hypoxia was 12.1 μmol g− 1 of fish. During the first 4 h of recovery from severe hypoxia exposure, there was a sustained efflux of ethanol from the goldfish followed by rates of ethanol production that could not be distinguished from zero (Fig. 2A). Ethanol production at rest was negligible and below the detection limit of our assay. During recovery from exhaustive exercise, ethanol was released from the fish to the water over the first 1 to 2 h of recovery and followed immediately by a period between 2 and 4 h recovery that was characterized by the loss of ethanol from the respirometer. At recovery times greater than 4 h, ethanol production rates were not distinguishable from zero (Fig. 2B). 3.2. White muscle There was no significant effect of treatment (hypoxia exposure or exhaustive exercise) or time on white muscle [ATP] (Fig. 3A). There was no significant effect of treatment on white muscle [PCr], but there was a significant effect of time (Fig. 3B). White muscle [PCr] decreased in response to both hypoxia exposure and exhaustive exercise and returned to values that were not different than controls by 0.5 h recovery. There was a significant effect of treatment and time on white muscle pHi (Fig. 3C). White muscle pHi differed in the resting/ normoxic groups of both treatments (Fig. 3C) and decreased in response to hypoxia exposure and exhaustive exercise. However, the magnitude of the decrease in white muscle pH was larger (0.31 pH units) after exhaustive exercise compared with that observed after 10 h hypoxia exposure (0.25 pH units). In addition, the rate of recovery
Fig. 3. White muscle ATP (A), PCr (B), and pHi (C) in goldfish at rest (R) and during 32 h recovery from severe hypoxia exposure (unfilled circles) or exhaustive exercise (filled circles). Vertical gray bar represents either 10 h severe hypoxia exposure or 15 min of exhaustive exercise. Values are means ± SEM (n = 8 for each point). Within each treatment (hypoxia or exercise), significant differences from resting are indicated by an asterisk (P ≤ 0.05).
Fig. 2. Ethanol production rates by goldfish during recovery from 10 h severe hypoxia (A) or 15 min exhaustive exercise (B). Time zero indicates the start of recovery. Positive values for ethanol production indicate ethanol release to water while negative values indicate ethanol loss from the water. Significant differences from zero are indicated by an asterisk (P ≤ 0.05).
was faster after hypoxia exposure compared with exhaustive with recovery rate constants of 1.35 h− 1 and 0.65 h− 1, respectively. Exposure to severe hypoxia and exhaustive exercise caused significantly different responses in muscle [glycogen] (two-way ANOVA; Fig. 4A). White muscle [glycogen] decreased as a result of hypoxia exposure and remained depressed compared with resting/ normoxic goldfish over the first 2 h recovery returning to values that were not significantly different from controls between 4 and 12 h with the exception of 16 h recovery, which was lower than the normoxic controls. In contrast, exhaustive exercise and recovery had no significant effect on white muscle [glycogen] (Fig. 4A). There was no effect of treatment on white muscle [lactate], but in both groups there was a significant effect of time (Fig. 4B). White muscle [lactate] increased to roughly equivalent levels as a result of hypoxia exposure and exercise and returned to values equivalent to the resting/normoxic controls over the same time period. There was a significant effect of treatment and time on white muscle [ethanol] (Fig. 4C). White muscle [ethanol] increased ∼40 fold during 10 h severe hypoxia exposure and remained elevated for the first 2 h of recovery followed by a return to levels that were not elevated compared with control fish. In contrast, exhaustive exercise had no effect on white muscle [ethanol], but during recovery,
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4. Discussion
Fig. 4. White muscle glycogen (A), lactate (B), and ethanol (C) in goldfish at rest (R) and during 32 h recovery from severe hypoxia exposure (unfilled circles) or exhaustive exercise (filled circles). See Fig. 3 caption for more details.
muscle [ethanol] increased gradually to levels that were significantly elevated at 16 h compared with control fish. 3.3. Liver There was a significant effect of treatment and recovery time on liver [glycogen] (Fig. 5A). Severe hypoxia exposure itself had no effect on liver glycogen, but within 0.5 h recovery, liver [glycogen] decreased by ∼ 35%, remained depressed compared with control samples until 2 h recovery and returned to values that were not significantly different than controls by 4 h recovery. Exhaustive exercise alone also had no effect on liver [glycogen]; however, in contrast to the severe hypoxia treatment, liver [glycogen] increased transiently during the first 2 h of recovery only to return to values that were not different than controls by 4 h recovery. Severe hypoxia exposure and exhaustive exercise yielded different responses in liver [ethanol], with a significant effect of treatment and time on liver [ethanol] (Fig. 5B). Liver [ethanol] increased ∼ 7.7 fold during severe hypoxia exposure and remained elevated compared with controls for 2 h recovery, subsequently returning to values that were not elevated compared with control fish. In contrast, there was no significant effect of exhaustive exercise or recovery on liver [ethanol].
Goldfish have long been recognized as unique among vertebrates for their ability to produce ethanol and CO2 as the major fermentative products of anaerobic metabolism (Shoubridge and Hochachka, 1980). The ethanol-producing pathway in goldfish is primarily isolated to the skeletal muscle due to the high tissue-specific alcohol dehydrogenase expression (Shoubridge and Hochachka, 1980; Mourik et al., 1982) and it is believed that lactate produced in other tissues during periods of anaerobiosis are shuttled to the skeletal muscle for ethanol biosynthesis. Ethanol is a small, highly diffusible molecule that once produced is released to the water via the gills, preventing self-intoxication and lactate buildup. Ethanol production, in combination with the large liver and muscle glycogen reserves, and the ability to undergo metabolic rate depression are thought to be the primary “adaptive” traits explaining how goldfish can survive weeks to months of complete anoxia. As pointed out by Shoubridge and Hochachka (1980), the disadvantage of ethanol production during anaerobiosis is that it is wasteful of carbon and may negatively impact the ability of goldfish to recover from metabolic insult. In the present study, we compared the metabolic responses in muscle and liver of goldfish before and after exposure to severe hypoxia or exhaustive exercise, as well as during recovery. We hypothesized that during hypoxia exposure, glycogen conversion to ethanol would predominate, while during a short bout of exhaustive exercise, glycogen conversion to lactate would predominate. Indeed substantial ethanol production was observed during the 10 h exposure to severe hypoxia (Fig. 1) as well as during the first 4 h recovery (Fig. 2A), which limited the extent of muscle glycogen recovery. However direct comparisons between the rates of glycogen recovery after hypoxia exposure and exhaustive exercise are not possible because goldfish do not preferentially deplete glycogen during exhaustive exercise. Despite this difficulty, comparisons of recovery from hypoxia exposure and exhaustive exercise in goldfish have yielded three novel findings. First, ethanol appears to be a ubiquitous metabolic end-product when tissue lactate levels are elevated either by severe hypoxia exposure or exhaustive exercise. Second, ethanol is not only released to the environment, but also appears to be taken up from the environment during recovery from exhaustive exercise. Third, differential responses in hepatic glycogen were observed during recovery from hypoxia exposure compared with exhaustive exercise, suggesting that the liver may play different roles during recovery from these two metabolic insults. The initial goal of our experiment was to reduce white muscle [glycogen] in goldfish to the same level by exposure to 10 h severe hypoxia exposure or exhaustive exercise and examine the rates of metabolic recovery. We successfully reduced white muscle [glycogen] by ∼ 60% (Fig. 4A) with hypoxia exposure, but we were unable to cause a significant decrease in white muscle glycogen with exhaustive exercise. The general picture of how fish white muscle powers exhaustive exercise suggests an almost complete depletion of all available fuels (ATP, PCr, and glycogen) and the accumulation of high levels of metabolic waste (e.g. lactate and H+); however, it must be acknowledged that this metabolic picture of exercise is primarily based on studies performed on athletic salmonids (Wang et al., 1994; Milligan, 1996; Richards et al., 2002a). Responses to exercise are heavily influenced by fish species and choice of exhaustive exercise protocol (Moyes and West, 1995), with salmonids being among the most willing to exhaust metabolic fuels. In goldfish however, complete exhaustion was characterized by a large reduction in white muscle [PCr] (Fig. 3B), tissue acidification (Fig. 3C), only modest accumulation of [lactate] (Fig. 4B) and no significant drop in muscle [glycogen] (Fig. 4A). Numerous preliminary trials were performed using several exercise protocols (manual chasing and various swim tunnel protocols) and rarely was substantial glycogen depletion noted. Indeed, the majority of studies that have examined the effects of intense exercise
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Fig. 5. Liver glycogen (A) and ethanol (B) in goldfish at rest (R) and during 32 h recovery from severe hypoxia exposure (unfilled circles) or exhaustive exercise (filled circles). See Fig. 3 caption for more details.
on muscle metabolism of Cyprinids, suggest that the metabolic paradigm associated with fish exercise does not hold true (Driedzic and Hochachka, 1976; van Ginneken et al., 2004). The common carp (Cyprinis carpio), for example, showed only a ∼13 μmol g− 1 increase in lactate during an exhaustive exercise protocol (Sugita et al., 2001), which is comparable to the present study, but far lower than that obtained using a similar protocol on trout (e.g. up to 30 μmol g-1 in trout; Richards et al., 2002b). Furthermore, Huber and Guderley (1993) demonstrated in goldfish that exhaustive exercise yielded limited muscle lactate accumulation (∼ 13 μmol g− 1 wet mass). Although exhaustive exercise only caused a non-significant decrease in white muscle [glycogen] of the ∼8 μmol glucosyl units g− 1 wet mass (Fig. 4A) this decrease was more than sufficient to account for the ∼ 8 μmol g− 1 wet mass accumulation of lactate. The reasons for muscle exhaustion remain unknown, but in our experiments the profound drop in pHi (Fig. 3C) or depletion of muscle [PCr] (Fig. 3B) are the likely causes of exhaustion. During hypoxia exposure, goldfish are well known for their ability to reduce metabolic rate by up to 60% (measured as heat dissipation; Van Waversveld et al., 1988, 1989), thus reducing whole animal ATP turnover and extending survival. Despite this impressive ability to reduce whole animal ATP turnover, an activation of substrate-level phosphorylation, primarily glycolysis using stored glycogen as a substrate, is still necessary to supply ATP and maintain cellular energy balance. The two major glycogen stores available to support whole animal metabolism during hypoxia exposure are located in the liver and white muscle. Surprisingly, there was no effect of 10 h severe hypoxia exposure on liver [glycogen] in the present experiment, which was contrary to expectation and to that observed by other authors (e.g. van den Thillart et al., 1980b). van den Thillart et al. (1980b) showed a transient decreases in liver [glycogen] during hypoxia exposure in goldfish and a more severe depletion during exposure to anoxia. The possible reasons for the lack of liver glycogen mobilization could be due to the relatively short duration of the hypoxia exposure as well as
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the fact that we did not expose our goldfish to complete anoxia, but rather severe hypoxia (b0.1 mg O2 L− 1). Independent of the reasons why, our data strongly support the notion that muscle glycogen stores are sacrificed preferentially over liver glycogen stores to support whole animal glycolysis. White muscle glycogen decreased by 19.5 μmol glucosyl units g− 1 wet mass (a 60% decrease compared with normoxic animals; Fig. 4A) and the majority of this glycogen depletion can be accounted for by lactate and ethanol accumulation. Our ethanol excretion rates are comparable to those reported by Johnston and Bernard (1983) for crucian carp exposed to anoxia at 15 °C with a maximum rate of ∼ 3 μmol g− 1 h− 1, but both the present study and others (Johnston and Bernard, 1983) report slightly higher excretion rates than van den Thillart (1983; maximum noted ∼ 1 μmol g− 1 h− 1). In the present study, white muscle glycogen is the primary substrate for whole ethanol production during hypoxia and we can nearly fully account for all the ethanol production by the depletion in white muscle glycogen. Assuming 60% of a goldfish is white muscle (estimate for salmonids; Moyes and West, 1995) then the observed muscle glycogen depletion, on a whole-animal basis is equivalent to 292.5 μmol glucosyl units (assuming an average fish mass of 25 g). Accumulated muscle lactate accounts for ∼19% of the glycogen depletion (7.3 μmol g− 1 muscle; 109.5 μmol total in 15 g of tissue), while the total ethanol production and accumulation in the tissues (assuming the measured muscle ethanol concentrations are equivalent to all cellular concentrations because of the high lipid solubility of ethanol) accounts for an additional 31% of the glycogen depletion. Furthermore, if we assume that the rapid efflux of ethanol observed during the first 4 h of recovery represent efflux of ethanol accumulated during the 10 h hypoxia exposure, then this accounts for an additional 18% of the glycogen depletion (107 μmol). In total, ethanol production accounts for a minimum of 50% of the total glycogen depleted during severe hypoxia exposure and we can account for 70% of the glycogen depletion by the accumulated lactate and ethanol. The remaining 30% of unaccounted for glycogen depletion maybe due to preferential accumulation of ethanol in different tissues or errors in our estimates of the percent of body weight occupied by muscle. During the recovery period, white muscle PCr (Fig. 3B) and pHi (Fig. 3C) recovered quickly (within 0.5 h) following hypoxia exposure and exhaustive exercise, while muscle lactate recovery took N12 h to return to resting/normoxic level following exhaustive exercise and hypoxia exposure (Fig. 4B). Differences in the rate of white muscle pHi recovery following hypoxia exposure and exhaustive exercise were likely due to the differences in magnitude of the pHi disturbance between the two treatments and the degree of perturbation brought about by the treatments. Lactate disappearance after exercise was not associated with increases in muscle glycogen, suggesting that the accumulated muscle lactate may, at least in part, be the substrate for the transient increase in whole animal ethanol production observed from 1 to 2 h recovery (Fig. 2B). It is surprising however, that only a modest increase in white muscle [ethanol] was observed (and the only significant increase occurred at 16 h recovery; Fig. 4C) suggesting that the tissue-specific source of ethanol may not be white muscle, but rather red muscle. Red muscle alcohol dehydrogenase activity is about 4 times higher than white muscle (Mourik et al., 1982), therefore lactate may be shuttled from the white muscle to the red for conversion to ethanol for subsequent release to the environment. Our ethanol flux measurements obtained after exhaustive exercise suggests that ethanol may be taken up from the environment and utilized by the fish during recovery. It is intriguing to speculate that the ethanol disappearance from the respirometer observed between 2 and 4 h recovery from exercise may represent uptake for use as a carbon substrate for glycogen synthesis or for ATP production. For either to occur, the pyruvate dehydrogenase complex would need to accept acetaldehyde as a substrate and facilitate its movement to either acetylCoA, for ATP production, or reversed to pyruvate, for use in glycogenesis.
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Further research into the regulation of the pyruvate dehydrogenase complex in goldfish muscle during recovery is needed to determine whether these possibilities are feasible. At this time however, our measurements of ethanol flux during recovery must be regarded as preliminary and await confirmation. If ethanol is a dynamic metabolite that can serve as a substrate for metabolism, then recovery of fish in an ethanol rich environment should enhance the rate of recovery. This is unlikely to be of a major consequence in natural populations of goldfish as ethanol is unlikely to accumulate in their environment to levels that would enhance recovery, but it remains an intriguing biochemical mystery worthy of further investigation. Liver glycogen stores responded in a dramatically different fashion during the early potions of recovery from severe hypoxia versus exhaustive exercise (Fig. 5A). During the very early portion of hypoxia recovery (0 to 0.5 h), mobilization of liver glycogen and release of glucose into the blood may be directly related to the early recovery of muscle glycogen observed between 1 and 2 h recovery or to facilitate recovery of other organs. Unlike exercise, which primarily affects muscle, hypoxia exposure affects the whole body and therefore organs not examined here (e.g. brain, heart, and gills). Thus during recovery, enhanced glucose release to the blood may be required to facilitate the recovery of other metabolically active tissues. The recovery of liver glycogen between 1 and 4 h is more of a mystery. Either the glucose released during the early portion of recovery is resynthesized into glycogen later on, or alternate carbon sources are used to resynthesize the glycogen. It remains intriguing to speculate that accumulated ethanol, may in part, account for some, albeit a potentially small fraction of the glycogen recovery after exhaustive exercise or hypoxia exposure. Partitioning of radiolabelled ethanol during recovery may help to address this question. In contrast to the liver responses observed during recovery from hypoxia, liver glycogen transiently increased by 1.7 fold during the first 1 to 2 h recovery from exhaustive exercise (Fig. 5A) suggesting that the liver is sequestering carbohydrates from the blood and other tissues, although again, this occurs in a transient fashion with almost compete recovery to control values within 4 h recovery. Overall, the transient changes in liver glycogen during the early phase of recovery may simply point to an enhanced total body turnover where liver serves as a temporary source or storage reserve for whole body carbohydrates. Acknowledgements This work was funded by a Natural Sciences and Engineering Research Council (NSERC) of Canada Discovery Grant to JGR. MM was supported by an NSERC Post Graduate Scholarship. References Bergmeyer, H.U., 1983. Methods of Enzymatic Analysis. Academic Press, New York. Blazka, P., 1958. The anaerobic metabolism of fish. Physiol. Zool. 31, 117–128. Dobson, G.P., Hochachka, P.W., 1987. Role of glycolysis in adenylate depletion and repletion during work and recovery in teleost white muscle. J. Exp. Biol. 129, 124–140. Driedzic, W.R., Hochachka, P.W., 1976. Control of energy metabolism in fish white muscle. Am. J. Physiol. 230, 579–582. Hassid, W., Abraham, S., 1957. Chemical procedures for analysis of polysaccharides. Methods Enzymol. 3, 34–37.
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