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First published online June 15, 2007
Journal of Experimental Biology 210, 2253-2266 (2007)
Published by The Company of Biologists 2007
doi: 10.1242/jeb.005116
Extreme anoxia tolerance in embryos of the annual killifish Austrofundulus limnaeus: insights from a metabolomics analysis
1 Department of Biology, Portland State University, PO Box 751, Portland, OR
97207-0751, USA
2 Department of Neurobiology, Pharmacology, and Physiology, The University
of Chicago, Chicago, IL 60637, USA
3 Department of Chemistry, Belknap Research Building, 2210 S. Brook Street,
University of Louisville, Louisville, KY 40208, USA
4 Hopkins Marine Station, Stanford University, Oceanview Boulevard, Pacific
Grove, CA 93950, USA
* Author for correspondence (e-mail: jpod{at}pdx.edu)
Accepted 5 April 2007
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-aminobutyrate (GABA), a potential
protector of neural function. It appears that the suite of characters
associated with normal development and entry into diapause II in this species
prepares the embryos for long-term survival in anoxia even while the embryos
are exposed to aerobic conditions. This is the first report of such extreme
anoxia tolerance in a vertebrate embryo, and introduces a new model for the
study of anoxia tolerance in vertebrates.
Key words: anaerobiosis, anoxia, development, fish, GABA, lactate, metabolic depression
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). The
physiology of anoxia tolerance has been examined in relatively few species of
vertebrates, notably the painted turtle Chrysemys picta and the
crucian carp Carassius carassius. Anaerobiosis in these organisms is
supported by a complex suite of traits that includes the storage of large
amounts of glycogen to support glycolytic production of lactate (turtles) or
ethanol (crucian carp), and elevated levels of -aminobutyrate (GABA) in
neural tissues (Lutz et al.,
1996). In both species, the single most critical factor for
survival of anoxia appears to be a severe reduction in metabolism (ATP
turnover), a state that is achieved via a coordinated downregulation
of energy producing and energy consuming processes
(Hand and Hardewig, 1996;
Jackson, 2000;
Krumschnabel, 2000;
Lutz and Nilsson, 2004).
In this study we investigated the anoxia tolerance of diapausing embryos of
the annual killifish Austrofundulus limnaeus. This species occurs in
ephemeral ponds in the Maracaibo Basin of Venezuela, which are a harsh and
highly variable habitat in which few aquatic vertebrates can survive.
Conditions in these ponds change radically during transitions from the rainy
season to the dry season. Rainy season ponds may remain inundated for several
weeks to several months with some ponds being semi-permanent
(Thomerson and Taphorn, 1992).
During the dry season, ponds may remain dry for several months and perhaps
years. A. limnaeus has evolved a life history strategy similar to
many aquatic invertebrate species, which includes the production of
drought-tolerant diapausing embryos
(Wourms, 1972b;
Podrabsky et al., 2001) that
are deposited into the muddy pond substrate
(Myers, 1952). Oxygen
limitation is often imposed by microbial activity in inundated soils, and the
habitat of A. limnaeus is no exception
(Podrabsky et al., 1998).
Consequently, embryos of A. limnaeus are likely to experience long
bouts of severe hypoxia or anoxia as a normal part of their developmental
environment.
Diapausing embryos of A. limnaeus share many metabolic
characteristics with other vertebrate facultative anaerobes such as freshwater
turtles and crucian carp. A significant metabolic depression (80–90%
reduction compared to developing embryos) accompanies diapause
(Podrabsky and Hand, 1999).
This metabolic depression is associated with a severe reduction in the rate of
protein synthesis (Podrabsky and Hand,
2000). Thus, these embryos are already in a state of metabolic
depression very similar to anoxia-induced quiescence even when incubated under
aerobic conditions. In fact, during aerobic development from fertilization
through entry into diapause II, there is a significant contribution of
anaerobic pathways to the overall metabolism of these embryos, as assessed by
calorimetry and respirometry (Podrabsky
and Hand, 1999). The present study was performed to further
characterize the predicted extreme anoxia tolerance of diapause II embryos of
A. limnaeus and identify the metabolic pathways that may provide this
species with its remarkable abilities to survive under extreme environmental
conditions.
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). Temperature
in the aquaria was regulated at 26°C. Embryos were collected twice weekly
to keep the spawning interval between 3 and 4 days, which has been shown to
produce the highest number of healthy embryos under laboratory conditions
(Podrabsky, 1999). All embryos
were incubated in the dark at 25°C in embryo medium containing 10 mg
l-1 gentamycin sulfate
(Podrabsky, 1999) unless
otherwise noted. These conditions are known to support entry of embryos into
diapause II, and exposure to a long-day photoperiod has been shown to break
diapause II (Podrabsky and Hand,
1999). On the day of exposure to anoxia, the embryos were treated
with two 5 min washes in 0.001% (w/v) hypochlorite bleach to reduce the
chances of bacterial and yeast growth during the long-term anoxia exposures
(Podrabsky, 1999).
Anoxia tolerance
Embryo medium containing 10 mg l-1 gentamycin sulfate was made
anoxic by purging the medium with high purity nitrogen gas for 20–30
min. Groups of 20 embryos were sealed into 20 ml glass screw-cap vials (caps
contained a silicon and Teflon septum) containing anoxic embryo medium. Just
prior to sealing the vials, sodium sulfite crystals were added to a final
concentration of 1.0 mg ml-1 to remove any residual oxygen and to
help protect against the reintroduction of oxygen. Vials were visually
inspected to ensure the absence of air bubbles. Calculations suggest that the
amount of sulfite added to each vial would be ten times the amount needed to
produce anoxic conditions from fully oxygenated medium. These levels were
non-toxic (see below). Anoxic vials were then sealed in a glass bell jar,
which was subsequently purged with nitrogen gas for 20 min. Anoxic conditions
were confirmed using a polarographic oxygen electrode (Model 1302;
Strathkelvin Instruments, Motherwell, UK) when each vial was opened for
processing. At each sampling time, a single vial of embryos from each
replicate was opened and the embryos were washed briefly in embryo medium, and
then placed in fresh, oxygenated embryo medium to recover from the anoxic
exposure. Survival of the embryos was assessed immediately and after 1 week of
aerobic recovery, using an inverted compound microscope. Embryos were scored
as alive if they were structurally intact and morphologically normal. Survival
after 1 week was assessed by the same criteria as the initial assessment with
the additional requirements of successful resumption of development (in
non-diapausing embryos) or normal cardiac function (in developing and
diapausing embryos). Mean lethal time (to 50% mortality; LT50)
estimates were calculated based on the number of embryos that were able to
survive and resume normal development or physiological function after 1 week
of aerobic recovery.
Anoxia tolerance was assessed in early developing and diapause II embryos
at 1, 2, 4, 8, 12, 16, 24 and 32 days post-fertilization (d.p.f.) and in
post-diapause II embryos at 4, 8 and 12 days post-diapause II (d.p.d.).
Embryos normally enter diapause II at 24 d.p.f. when incubated at 25°C.
Embryos were induced to break diapause synchronously by exposure to a long-day
photoperiod (14 h:10 h L:D) (Podrabsky and
Hand, 1999) at an incubation temperature of 30°C for 2 days.
After the embryos were observed to break diapause they were immediately
returned to conditions of 25°C in the dark. For all developmental stages,
three different spawning events were used to produce three replicates for
statistical analysis at all time points.
Toxicity of sodium sulfite
The potential toxicity of sodium sulfite during anoxic exposure was
assessed in 4 and 8 d.p.f. embryos. Embryos for these studies were treated as
described above, except that sodium sulfite was added to a final concentration
that ranged from 0.25 to 2.5 mg ml-1. For each developmental stage
a single batch of embryos was used to produce three replicates of 20 embryos
for each time point. At the concentrations used in this study, sodium sulfite
appears to have no toxic effect on the survival of embryos exposed to anoxia
(Figs S1 and S2 in supplementary material).
Preparation of metabolite extracts for gas chromatography–mass spectrometry analysis
During normoxic development embryos at 0, 4, 8, 12, 16, 20, 24 and 32
d.p.f. were flash frozen in liquid nitrogen and stored at –80°C
prior to extraction of metabolites. Embryos at 1, 2, 4, 8, 16, and 32 d.p.f.
were exposed to anoxia and sampled at times consistent with their anoxia
tolerance: 0.5 and 1 day of anoxia for 1 and 2 d.p.f. embryos, 7 and 21 days
for 4 d.p.f. embryos, and 21 and 60 days for 8, 16 and 32 d.p.f. embryos.
Embryos were removed from anoxia and the surviving embryos were quickly
counted and their mass determined prior to flash freezing. Embryos were
lyophilized in pre-tared microcentrifuge tubes and then pulverized using a
Teflon pellet pestle. The pulverized powder was then homogenized with 20
volumes of ice-cold 10% (w/v) trichloroacetic acid (TCA). The TCA homogenate
was then centrifuged at 14 000g for 30 min at 4°C to
pellet the protein and other TCA-insoluble materials. The supernatant was
carefully removed and retained. The pellet was then washed once with 20
volumes of ice-cold 10% TCA and centrifuged as described above. The two
supernatants were then pooled and lyophilized overnight to remove the TCA and
dehydrate the extract. The dried extract was then resuspended in water
(0.2–0.4 ml) and a small fraction of the rehydrated extract
(0.04–0.06 ml) was dispensed into a glass gas chromatography vial and
again lyophilized to dehydrate the sample [modified from Gradwell et al.
(Gradwell et al., 1998)].
Silylation gas chromatography–mass spectrometry analysis
The lyophilized TCA extract powder was derivatized with 0.025 ml of a 1:1
ratio mix of acetonitrile:
N-methyl-N-[tert-butyldimethylsily]trifluoroacetamide
(MTBSTFA). The vials were then sealed and sonicated at room temperature for 3
h. Following sonication, 0.5 µl of the derivatized extract was injected
into a Varian 3400 gas chromatograph (Varian Instruments, Palo Alto, CA, USA)
coupled with a Finnigan ITD 806 mass spectrometer (Finnegan MAT, San Jose, CA,
USA) according to the methods of Fan et al.
(Fan et al., 1993). A mixture
of 37 compounds was similarly silylated and analyzed by gas
chromatography–mass spectrometry (GC–MS) to serve as calibration
standards.
1H nuclear magnetic resonance spectroscopy
Perchloric acid extracts were prepared for three samples for analysis using
nuclear magnetic resonance (NMR) spectroscopy: diapause II, diapause II
embryos exposed to 75.5% relative humidity for 3 days, and 8 d.p.f. embryos.
These embryos were extracted twice in 20 vol of ice-cold 10% perchloric acid
(PCA). PCA was removed by precipitation with KOH (pH adjusted to between 6 and
7) and centrifugation of the precipitated materials at 14 000g
for 15 min (Fan et al., 1986).
The neutralized PCA extracts were then passed over a Chelex-100 column
(Bio-Rad, Hercules, CA, USA) to remove paramagnetic ions, lyophilized and
dissolved in D2O. One-dimensional and two-dimensional total
correlation spectroscopy (TOCSY) 1H NMR spectra were acquired at
25°C using a Varian Unity Plus NMR spectrometer operating at 11.75
Tesla.
Statistical analysis
All statistical analyses were performed using SPSS software (SPSS Inc.,
v.14.0.1, Chicago, IL, USA). LT50 was calculated for each time
series of anoxic exposures using probit regression analysis
(Chapman et al., 1995).
LT50 values determined for each developmental stage were compared
using analysis of variance (ANOVA, general linear models procedure).
Comparison of means for all data sets was accomplished using ANOVA (as above).
Unless noted otherwise, the Student–Neuman–Keul's
post-hoc test was applied to determine the differences among means.
Statistical significance was set for P<0.05 in all
comparisons.
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One-dimensional and two-dimensional TOCSY 1H-NMR
The NMR analysis (Figs S3, S4 in supplementary material) corroborates the
GC–MS data presented below, by confirming the identity of a number of
metabolites present in the embryo extracts. In addition, the NMR data revealed
major metabolites not observed by GC–MS, including glycinebetaine (GB),
phosphorylcholine (PC), and polysaccharides (broad resonances between 3.5 to
5.2 p.p.m. (Fig. S3 in supplementary material). These metabolites appeared to
accumulate in diapause II embryos, but their response to anoxia was not
investigated. GB is known to be a compatible solute involved in osmoregulation
whereas PC is important to phospholipid metabolism. The accumulation of
polysaccharides may reflect development of an energy reserve in the diapause
II embryos in preparation for anoxia exposure or dehydration. The roles of
these metabolites during diapause and exposure to anoxia warrants further
investigation.
Glycolytic metabolites
Only two glycolytic intermediates, glyceraldehyde 3-phosphate (G3P) and
lactate, were detected in this study, despite the ability of these methods to
identify several other potentially important metabolites such as pyruvate. The
concentration of G3P increased from around 0.5–2.0 mmol kg-1
embryo H2O during early development through diapause II in embryos
incubated aerobically (Fig. 4),
however this trend is not statistically significant (ANOVA, P=0.065).
During exposure to anoxia, G3P concentration increased significantly in 1, 16,
and 32 d.p.f. (ANOVA, P<0.014) but not in 2, 4 and 8 d.p.f.
embryos. During normoxic development lactate increased significantly (ANOVA,
P
). During exposure to anoxia, lactate was the most abundant
metabolic end-product accumulated and reached values between 30 and 40 mmol
kg-1 embryo H2O (Fig.
4). The rate of lactate accumulation during anoxia is inversely
correlated (r=–0.97, P0.000001) with the anoxia
tolerance of the embryos as estimated by the LT50
(Fig. 5). This pattern of
slower accumulation of metabolites in diapausing embryos is apparent in most
of the compounds analyzed in this data set and is likely correlated with the
metabolic rate of these embryos under anoxia.
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Citric acid cycle metabolites
Three citric acid cycle intermediates were identified in these samples:
succinate, malate and citrate (Fig.
6). Fumarate, 
-ketoglutarate, ß-hydroxybutyrate and
t-aconitate are all potentially quantifiable using the methods employed in
this study but were not detected. Succinate levels were undetectable during
normoxic development, but increased significantly (ANOVA, P<0.002)
to concentrations between 0.5–1.0 mmol kg-1 embryo
H2O during exposure to anoxia
(Fig. 6). Malate concentrations
increased by a small but statistically significant amount during normal
development (ANOVA, P=0.023), and tended to decrease during anoxia,
although this decrease is only significant (ANOVA, P=0.034) for 8
d.p.f. embryos (Fig. 6).
Citrate levels increased during normoxic development (ANOVA, P=0.042)
and declined significantly (ANOVA, P<0.042) in 1, 2 and 8 d.p.f.
embryos exposed to anoxia (Fig.
6). Overall, there appears to be an accumulation of succinate, a
small increase in malate, and little change in citrate concentrations in
embryos exposed to anoxia that possess a substantial anoxia tolerance.
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During normoxic development and during exposure to anoxia, the pool of free amino acids appears to be dominated (50–70%) by glutamate, alanine and lysine in most cases (Fig. 8). There is a significant increase in the concentration of all amino acids during normoxic development except for aspartate and methionine, which are found in very low concentrations (Figs 9, 10, 11, 12). Exposure to anoxia is accompanied by an increase in the total levels of essential free amino acids (Fig. 8). Changes in the concentration of individual amino acids during exposure to anoxia can be categorized into three major patterns. (1) A statistically significant increase in the concentration of nine amino acids (ANOVA P<0.05) occurs for at least one developmental stage after 60 days of anoxia; six of these are considered essential amino acids in fish (Fig. 9) and three of them are classified as non-essential (Fig. 10). These amino acids include, in order of abundance, alanine, valine, glycine, leucine, isoleucine, proline, phenylalanine, methionine and threonine. (2) A significant decrease in the concentration (ANOVA, P<0.05) of four amino acids: aspartate, asparagine, glutamate, glutamine, occurs after 60 days of exposure to anoxia in all treatments except for embryos at 32 d.p.f. (Fig. 11). (3) No significant change occurred in three amino acids during exposure to anoxia: serine and tyrosine (non-essential) and lysine (essential, Fig. 12). One trend that is obvious for all of the amino acids is an overall lower rate of accumulation, and often smaller total amount accumulated in embryos diapausing at 32 d.p.f. compared with those at 8 and 16 d.p.f.
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-aminobutryate
Concentrations of the inhibitory neurotransmitter -aminobutyrate
(GABA) rise to levels between 7 and 12 mmol kg-1 embryo
H2O during exposure to anoxia
(Fig. 13). This compound is
not found in embryos during normoxic development, and is only accumulated to
significant levels in embryos that have substantial anoxia tolerance.
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;
Jackson, 2000).
In vertebrates, the tissues most sensitive to oxygen deprivation are the
brain and heart (Lutz et al.,
1996; Lutz and Nilsson,
2004; Wasser,
1996; Giordano,
2005). Interestingly, the embryos of A. limnaeus that
exhibit the highest degree of anoxia tolerance are composed mainly of neural
tissues, and they contain a functional heart
(Wourms, 1972b;
Podrabsky and Hand, 1999).
Diapause II embryos can be observed `flexing' spontaneously and in response to
light. They also respond physiologically to light cues
(Podrabsky and Hand, 1999).
Many of the embryos establish a closed circulation with immature red blood
cells prior to entry into diapause II. During early post-diapause II
development, the differentiation and maturation of the neural and cardiac
tissues progresses, and includes the pigmentation of the retina, increased
volume of the brain, circulating red blood cells that express hemoglobin, as
well as the beginnings of the endoderm-derived internal organs such as the
liver and gut (Wourms, 1972a).
Thus, anoxia tolerance in these embryos is not simply a situation of
`undifferentiated' cells surviving anoxia. Rather, these embryos contain
functional neurons and cardiomyocytes that are incredibly resistant to
anoxia.
Anoxia-induced quiescence
When developing embryos of A. limnaeus are exposed to anoxia, they
stop developing and enter a state of anoxia-induced quiescence
(Fig. 3). Direct observation of
embryos during post-diapause II development indicates that these embryos
arrest development and experience a severe bradycardia or a complete cessation
of cardiac activity under anoxia
(Fergusson-Kolmes and Podrabsky, in
press). Developmental arrest has been reported for embryos of
other species of teleost fish in response to environmental anoxia and chemical
anoxia induced by mitochondrial inhibitors
(Crawford and Wilde, 1966;
Padilla and Roth, 2001).
However, in these instances survival is relatively short (1–2 days)
compared to the survival times reported here for embryos of A.
limnaeus.
Early post-diapause II embryos (4 d.p.d.) exhibit a tolerance of anoxia
that is equal to that of diapause II embryos
(Fig. 1). This result is
impressive, considering the increased metabolism and complexity associated
with further development in these embryos (see above). It appears that there
is a window of time during early post-diapause II development when the embryos
retain the ability to enter a profound state of anoxia-induced quiescence.
This is similar to the situation for anoxia-induced quiescence in
post-diapause embryos of the brine shrimp Artemia franciscana (e.g.
Hand and Gnaiger, 1988;
Clegg, 1997). Perhaps this
extended tolerance of anoxia reflects the ephemeral nature of these species'
habitats, where a return to low oxygen levels might occur after developmental
arrest is over. Thus, the evolution of diapause and anoxia-induced quiescence
appear to be linked in both of these distantly related species. This possible
example of convergent evolution is probably the result of high selection
pressures generated in harsh and highly unpredictable ephemeral habitats.
Recovery from anoxia
Embryos exposed to anoxia for 30–40 days are able to fully recover
from the anoxic insult (Fig.
2). However, after 60 days of anoxia, many of the embryos fail to
successfully resume development and eventually die. This failure to recover
may be due to metabolic failure associated with build-up of toxic amounts of
lactate (Fig. 4). An
alternative explanation is that the embryos suffer from damage caused by the
reintroduction of oxygen after long-term anoxia in a manner similar to
reperfusion injury associated with recovery from ischemia in mammalian tissues
(e.g. Gross and Auchampach,
2007). If a `reperfusion' of oxygen into the embryos is indeed a
source of damage after long-term anoxia, then comparing the metabolic changes
and gene expression patterns in embryos recovering from short-term and
long-term anoxia may lend insight into the mechanisms that induce reperfusion
injury.
General considerations for metabolite analysis in embryos
The levels of metabolites reported in this paper are from whole embryos.
These embryos contain several compartments that must be considered in the
interpretation of these data: the perivitelline space, the embryonic tissues,
and the yolk. The perivitelline space is filled with perivitelline fluid (PVF)
and contains about 50% of the total embryo water
(Podrabsky et al., 2001). This
compartment surrounds the embryonic tissues and yolk. The PVF is derived
partially from the contents of the cortical vesicles and is known in many
species of teleost fish to contain mostly large macromolecules that attract
water and cause osmotic swelling of the egg immediately following
fertilization (Eddy, 1974;
Shephard, 1987). If we assume
that the perivitelline space is devoid of the metabolites examined in this
study, then their effective concentrations in the embryo + yolk compartments
may be twice that reported, based on total embryo water content. The embryonic
tissues surround the central yolk mass, and for most of the embryonic stages
observed in this study constitute only 10–20% of the total embryo mass.
The embryonic tissues are separated from the yolk by the yolk syncytial layer,
which is a known barrier to diffusion of water and small organic molecules,
such as cryoprotectants, in freshwater fish embryos (e.g.
Hagedorn et al., 1997). In
addition, enzymes have been demonstrated to be distributed in a non-uniform
manner between the yolk and embryonic tissues in embryos of the loach
Misgurnus fossilis (Klyachko et
al., 1982). Thus, compartmentalization of metabolites and
metabolic function is a distinct possibility and without specific information
on where in the embryo/yolk/PVF system these compounds may be located we are
limited in our ability to interpret the results. However, this study does
identify the major metabolic pathways that contribute to anoxia tolerance in
these embryos, and our findings represent a first critical step in
mechanistically understanding the metabolic underpinnings of anoxia tolerance
in this species.
Anaerobic end-product accumulation
In general, animals that can survive long bouts of anoxia rely on
utilization of carbohydrates to fuel metabolism
(Hochachka and Somero, 2002).
This is true of both the crucian carp and freshwater turtle species, which
have large glycogen reserves used to fuel anaerobic metabolism
(Jackson, 2000;
Lutz and Nilsson, 2004).
Metabolism appears to be dominated by glycolytic production of lactate during
exposure of A. limnaeus embryos to anoxia
(Fig. 4B). Levels of glycogen
and glucose that are presumably available to support anaerobic metabolism
during early development and diapause II, have been previously reported for
A. limnaeus (Podrabsky and Hand,
1999). About 80–90% of the available glycosyl units are
stored as glycogen, and after 2 d.p.f. the average amount of glycosyl units
available for the production of lactate during early development and diapause
II is 36.2±1.0 nmol embryo-1 (mean ± s.e.m.), which
would yield about 72.4 nmol of lactate per embryo. This glycogen reserve is
similar to that of other fish species with similar size embryos such as loach
and medaka (Yurowitzky and Milman,
1973; Hishida and Nakano,
1954), and an order of magnitude lower than that of trout embryos
(Terner, 1968). Therefore, it
does not appear that accumulation of large glycogen stores can explain the
exceptional anoxia tolerance of this species.
The exact metabolic pathways that are used to support metabolism during
anoxia in embryos of A. limnaeus have not yet been confirmed with
biochemical studies. However, a simple analysis of end-products accumulated
suggests that available glycogen stores can support most, if not all, of the
accumulated compounds. The amounts of lactate, succinate and alanine produced
after 21 (4 d.p.f.) or 60 days (8, 16, 32 d.p.f.) of anoxia are presented in
nmol embryo-1 in Table
1. When these data are compared with the theoretical amount of
three to four carbon compounds that could be produced from glycogen and
glucose, it is apparent that exhaustion of carbohydrate stores may be a
limiting factor in the survival of embryos 4 d.p.f. and older. In contrast,
embryos at 1 and 2 d.p.f. die long before they run out of fuel to support
glycolytic metabolism (data not shown, but see
Fig. 4) perhaps due to an
inability to buffer the acidosis that likely accompanies anoxic metabolism [2
moles of H+ per mole of glycosyl unit
(Hochachka and Mommsen,
1983)], but no data are available for intracellular pH changes
associated with anoxia in this species. Rough calculations based on the mean
rate of lactate production in diapausing embryos (0.48±0.08 nmol
day-1) at 32 d.p.f. would suggest an exhaustion of available
glycosyl units after about 151 days of anoxia. When the survival experiments
were terminated at 90 days of anoxia, about 20% of the 32 d.p.f. embryos
survived the treatment, and thus it is possible that available glycogen stores
will ultimately limit survival of anoxia in these embryos as well. The strong
negative correlation between the rate of lactate production during anoxia and
survival of anoxia (Fig. 5)
reinforces the importance of metabolic rate depression to long-term survival
of anoxia in embryos of A. limnaeus.
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).
The role of these metabolites during anaerobiosis in A. limnaeus is
not yet understood, but a limited maintenance of Krebs cycle activity to
provide intermediates for other metabolic conversions (such as the production
of GABA) is a distinct possibility and has been reported in other vertebrate
anaerobes (Van der Boon et al.,
1992; van Raaij et al.,
1994). High levels of succinate relative to available
-ketoglutarate can also inhibit prolyl hydroxylase, which may lead to
stabilization and activation of hypoxia-inducible factor 1, thereby
enhancing glycolytic activity and oxygen-dependent gene expression
(Gottlieb and Tomlinson,
2005).
One additional metabolite that warrants discussion is citrate
(Fig. 6). Citrate increases in
concentration during early development and reaches levels around 3 mmol
kg-1 embryo H2O during diapause II, but these
concentrations do not tend to change during exposure to anoxia. This amount of
citrate is an order of magnitude higher than that found in embryos of loach
(Milman and Yurowitzky, 1967).
Citrate is known to be a potent inhibitor of phosphofructokinase with
inhibition occurring in the micromolar range (e.g.
MacDonald and Storey, 2004).
Given the above information on the rate of lactate production, we must
conclude that whole embryo levels of citrate probably do not reflect
cytoplasmic levels of citrate in these embryos. It is highly likely that a
large percentage of this citrate is partitioned in the yolk, but it may also
be sequestered in the mitochondria. Citrate has the potential to bind metal
ions of various types including calcium, magnesium and iron
(Salovaara et al., 2003) and
perhaps may be functioning as a chelating agent for the storage of metals in
the yolk. Citrate accumulation has also been implicated in redox balance and
may play a role in antioxidant defenses in some cells
(Mallet and Sun, 2003).
Moreover, citrate is the source of acetyl CoA in fatty acid biosynthesis, and
thus can serve as a reserve for new membrane synthesis during development. The
unique biochemistry of citrate and the high levels accumulated in diapausing
embryos of A. limnaeus suggest that this molecule may play a critical
role in the biochemistry of diapause.
Free amino acids
The free amino acid pool in teleost fish embryos is thought to be generated
by degradation of yolk proteins (Monroy et
al., 1961; Love,
1970; Srivastava et al.,
1995; Gunasekera et al.,
1999; Finn et al.,
2002). In embryos of A. limnaeus, the total protein
content of the embryos decreases significantly during early development
through diapause II (Podrabsky and Hand,
1999), which suggests liberation of amino acids from yolk protein
stores to fuel metabolism and biosynthesis during development, as observed in
other teleost embryos (Rønnestad
and Fyhn, 1993). Previous evaluation of the distribution of amino
acids between the embryonic tissues and non-embryonic compartments
(perivitelline space and yolk) indicates that a very large percentage of the
free amino acids (93–99%) are not found in the embryonic cells
(Podrabsky and Hand, 2000).
Thus, the most likely source for the increase in the essential amino acids
observed during normoxic development and after exposure to anoxia is protein
degradation of yolk proteins. However, we cannot exclude the possibilities
that degradation of proteins in the embryonic tissues, or reduced consumption
of amino acids due to arrest of protein synthesis do not contribute as well.
Although there is a dramatic ontogenetic increase in the total free amino acid
pool during early development, this still probably represents an insignificant
fraction of the osmotic pool at fertilization and only about 10–15% of
the total osmolality of the egg if we assume an internal osmolality of 290
mOsmol kg-1 embryo H2O
(Machado and Podrabsky,
2007).
The composition of the free amino acid pool in normoxic and anoxic embryos of A. limnaeus is quite divergent from their average representation in lipovitellin heavy chain I (LvH1) or in the precursor of this major yolk protein vitellogenin [Fig. 8; mean percentage of amino acids in LvH1 from three species as listed in GenBank: Krytolebias marmoratus (accession number AY279214), Fundulus heteroclitus (accession number Q90508) and Oncorhynchus mykiss (CAA63421.1)]. Similar percentages of amino acids are obtained if the whole vitellogenin protein is used, except for serine which increases due to its high representation in the phosvitin domain of the vitellogenin (data not shown). This altered profile of amino acids is likely the result of de novo synthesis of a few select amino acids such as glutamate and alanine, and selective sparing of others such as lysine. In fact, lysine is the only essential amino acid that increases substantially more than the others during diapause II in embryos under normoxic conditions, and can account for most of the additional increase in FAA observed in `steady-state' diapausing embryos at 32 d.p.f. compared with those just entering diapause II at 24 d.p.f. (Fig. 7). The role of elevated lysine during diapause II is currently unknown, but it is doubtful that it plays a major role during exposure to anoxia.
Glutamate, glutamine, aspartate and asparagine are all non-essential amino
acids of the `citric acid cycle superfamily' and can be transaminated into
citric acid cycle intermediates with relative ease within most cells
(Mathews and van Holde, 1990).
Glutamate and glutamine accumulate in large quantities in diapause II embryos
of A. limnaeus during normoxic development. These amino acids are
consumed during exposure to anoxia (Fig.
11), and thus their accumulation during early development and
diapause may be an important preparation for the metabolic changes associated
with exposure to anoxia. Similarly, decreases in brain tissue glutamate
concentrations have been documented for turtles and crucian carp (reviewed by
Lutz et al., 1996). The amino
acids are probably used as substrates for the production of GABA (see below).
However, even if 100% of these amino acids are shunted to the production of
GABA, there is still a substantial proportion of GABA which cannot be
accounted for (Table 2) that
could be produced from glycogen stores
(Table 1).
-aminobutyrate
The metabolic pathways that support -aminobutyrate (GABA) production
and degradation are well characterized in vertebrate nervous tissues. GABA is
typically synthesized from glutamate by the enzyme glutamate decarboxylase and
is degraded in post-synaptic neurons to succinate
(Turner and Whittle, 1983).
The large decreases in glutamate and glutamine observed in embryos exposed to
anoxia are probably associated with synthesis of GABA. The production of GABA
at the expense of glutamate would seem to be beneficial due to the decrease in
glutamate (an excitatory neurotransmitter), a consumption of protons, and the
increase in GABA as discussed above. The degradation pathways for GABA lead to
succinate, and thus perhaps the accumulation of succinate during anoxia may be
partially due to post-synaptic degradation of GABA.
The evidence available on the role of GABA during oxygen limitation
suggests that GABA is acting to limit excitotoxic neuronal cell death in
embryos of A. limnaeus exposed to anoxia. GABA is the most abundant
inhibitory neurotransmitter in the adult vertebrate central nervous system
(Turner and Whittle, 1983;
Martyniuk et al., 2005) and is
generally thought to protect the brain of a variety of organisms from
excitotoxic cell death during oxygen limitation
(Lutz and Milton, 2004;
Cheung et al., 2006). The best
studied receptors for GABA are the GABAA receptors, which are
ligand-gated chloride channels
(Stephenson, 1995). The action
of GABA via these receptors as excitatory or inhibitory is dependent
on the concentration of intracellular chloride ions
([Cl-i]), with low [Cl-i]
resulting in inhibition (Cheung et al.,
2006; Tyzio et al.,
2006). Recent reports indicate that regulation of
[Cl-i] via expression of the NKCC1
electroneutral cation–chloride cotransporter may play a critical role in
protecting neurons of a variety of organisms during exposure to hypoxia or
ischemia (Cheung et al., 2006;
Tyzio et al., 2006).
GABA has been shown to play important roles in the development of the
vertebrate central nervous system (CNS). During development and maturation of
the CNS, GABA is known to act as an excitatory neurotransmitter
(Tyzio et al., 2006). The
excitatory actions of GABA are due to a high [Cl-i]
early in development (Cherubini et al.,
1991) prior to the insertion of Cl transporters (see above) into
the neuronal membrane. Insertion of the Cl- transporters causes a
decrease in [Cl-i] and GABA then assumes an inhibitory
role. The accumulation of high levels of GABA in embryos of A.
limnaeus suggests that it is likely to be inhibitory at this stage of
development. Future studies might examine at what stage in development the
Cl- transporter appears in brain tissue.
In embryos of the teleost fish Gasterosteus aculeatus, GABA is the
first neurotransmitter to be expressed in the central nervous system and is
thought to play a role in nervous system patterning and in the establishment
of the early axonal scaffold (Ekström
and Ohlin, 1995). Presently, there is no information on the
development of GABAergic neurons in embryos of A. limnaeus. The lack
of detectable GABA in normoxic embryos may indicate that these neurons have
not yet developed or are very few in number in A. limnaeus embryos
entering diapause II. However, the large increases observed in GABA
concentration during exposure to anoxia could act as a signal to activate the
expression of genes involved in GABA-mediated signaling, as has been shown in
a variety of vertebrates including goldfish
(Martyniuk et al., 2005).
Indeed, in anoxic turtle brains a rise in GABA is accompanied by an increase
in GABAA receptors (Lutz and
Milton, 2004), which is thought to strengthen the inhibitory
effects of GABA. The complex interaction between GABA production and the
expression of GABA-related genes, and the pivotal role that GABA plays in the
development of the vertebrate CNS suggests that production of GABA during
anoxia may alter gene expression patterns that could have lasting effects on
the development of the CNS in embryos of A. limnaeus.
In addition to its role in the CNS, GABA is known to play a role in the
physiology of a number of organ systems, including the heart. GABAB
receptors are known to be expressed in mammalian cardiomyocytes, and
activation of these channels is thought to result in post-synaptic inhibition
and subsequent protection of cardiac tissue from ischemia
(Lorente et al., 2000). Thus,
GABA may have a role in supporting survival of cells during anoxia outside of
the CNS in embryos of A. limnaeus. A critical next step in
elucidating the role of GABA in the tolerance of anoxia in A.
limnaeus will be to establish the compartmentalization of GABA within the
embryo and establish which receptors (if any) are expressed in the various
tissues of the embryo.
Diapause embryos are `primed' for anoxia tolerance
Metabolic depression is supported in most organisms by a coordinated
downregulation of energy consuming and energy producing pathways
(Hand and Hardewig, 1996;
Hand, 1998;
Hochachka and Somero, 2002).
In general, protein synthesis and ion pumping are the two most energetically
expensive processes contributing to the basal metabolic rate of a cell
(Buttgereit and Brand, 1995;
Rolfe and Brown, 1997). A
common response in all animals that survive long-term anoxia is a
downregulation of protein synthesis, and a reduction in ion leakage, which
results in reduced ion motive ATPase activities
(Hand and Hardewig, 1996).
Entry into diapause II in embryos of A. limnaeus is associated with a
substantial decrease in metabolic rate as estimated by both oxygen consumption
and heat dissipation (Podrabsky and Hand,
1999). In addition, anaerobic metabolic pathways are thought to be
making a significant contribution to overall metabolism in diapause II
embryos, even though they are incubated under aerobic conditions
(Podrabsky and Hand, 1999).
Rates of protein synthesis are severely reduced in diapause II embryos
compared to developing embryos, which can account for about 30% of the overall
depression in metabolic rate (Podrabsky
and Hand, 1999). Recently, we have found that embryos of A.
limnaeus have very low Na+-K+-ATPase activity
during all of early development, and levels of this enzyme do not increase
substantially until about 8 d.p.d.
(Machado and Podrabsky, 2007).
Therefore, it is logical to hypothesize that ion homeostasis with the
environment may not be a significant metabolic demand in these embryos. Thus,
the two most important biochemical adjustments associated with metabolic
depression and survival of anoxia, reduction of rates of protein synthesis and
reduced ion pumping, have already been made in aerobically incubated
diapausing embryos. In addition, diapausing embryos of A. limnaeus
accumulate large stores of glutamate and glutamine during normal development,
which can be easily converted into GABA for protection of neural tissues.
Based on these observations, it appears that diapause II embryos of A.
limnaeus have already made the adjustments necessary to survive anoxia,
prior to any anoxic exposure, and are thus `primed' for survival of anoxia as
part of their normal developmental program. Importantly, this high tolerance
of anoxia is retained during early post-diapause II development and thus
embryos may break diapause and still survive long bouts of anoxia. The
ecological and evolutionary implications of `preparing' for anoxia are
intriguing and further exploration of this system seems likely to yield
interesting insights into the ecology and evolution of anoxia tolerance in
vertebrates.
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Acknowledgments |
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Footnotes |
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References |
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TOPSummaryIntroductionMaterials and methodsResultsDiscussionReferences |
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