Download Life in the slow lane: molecular mechanisms of estivation

Comparative Biochemistry and Physiology Part A 133 (2002) 733–754
Review
Life in the slow lane: molecular mechanisms of estivation夞
Kenneth B. Storey*
Institute of Biochemistry, Carleton University, 1125 Colonel By Drive, Ottawa, Ontario, Canada K1S 5B6
Received 13 January 2002; received in revised form 4 April 2002; accepted 10 April 2002
Abstract
Estivation is a state of aerobic hypometabolism used by organisms to endure seasonally arid conditions, often in desert
environments. Estivating species are often active for only a few weeks each year to feed and breed and then retreat to
estivate in sheltered sites, often underground. In general, estivation includes a strong reduction in metabolic rate, a
primary reliance on lipid oxidation to fuel metabolism, and methods of water retention, both physical (e.g. cocoons) and
metabolic (e.g. urea accumulation). The present review focuses on several aspects of metabolic adaptation during
estivation including changes in the activities of enzymes of intermediary metabolism and antioxidant defenses, the effects
of urea on estivator enzymes, enzyme regulation by reversible protein phosphorylation, protein kinases and phosphatases
involved in signal transduction mechanisms, and the role of gene expression in estivation. The focus is on two species:
the spadefoot toad, Scaphiopus couchii, from the Arizona desert; and the land snail, Otala lactea, a native of the
Mediterranean region. The mechanisms of metabolic depression in estivators are similar to those seen in hibernation and
anaerobiosis, and contribute to the development of a unified set of biochemical principles for the control of metabolic
arrest in nature.
䊚 2002 Elsevier Science Inc. All rights reserved.
Keywords: Scaphiopus couchii; Otala lactea; Signal transduction; Gene expression; Reversible phosphorylation; Riboflavin binding
protein; Metabolic rate depression; Urea effects on enzymes
1. Introduction
Estivation is a state of aerobic torpor that is
probably best defined as a survival strategy for
dealing with arid conditions, but is also typically
夞 This paper was originally presented at ‘Chobe 2001’; The
Second International Conference of Comparative Physiology
and Biochemistry in Africa, Chobe National Park, Botswana
– August 18–24, 2001. Hosted by the Chobe Safari Lodge
and the Mowana Safari Lodge, Kasane; and organised by
Natural Events Congress Organizing ([email protected]).
*Corresponding author. Tel.: q1-613-520-3678; fax: q1613-520-2569.
E-mail address: kenneth [email protected]
(K.B. Storey).
associated with a lack of food availability and
frequently with high environmental temperatures
(Pinder et al., 1992; Abe, 1995; Land and Bernier,
1995). Estivation occurs widely in both vertebrates
and invertebrates. Among vertebrates the list
includes various air-breathing fish (prominently
the lungfish), amphibians (frogs, toads, salamanders) and reptiles (lizards, crocodilians, turtles) as
well as some small mammals (Bemis et al., 1987;
Etheridge, 1990; Kennett and Christian, 1994;
Abe, 1995; Land and Bernier, 1995; Wilz and
Heldmaier, 2000). The physiology and biochemistry of estivation has been studied in greatest
detail in two groups—anuran amphibians and pulmonate land snails. Estivation can be short term
1095-6433/02/$ - see front matter 䊚 2002 Elsevier Science Inc. All rights reserved.
PII: S 1 0 9 5 - 6 4 3 3 Ž 0 2 . 0 0 2 0 6 - 4
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K.B. Storey / Comparative Biochemistry and Physiology Part A 133 (2002) 733–754
(Wilz and Heldmaier, 2000), but is more often
employed to allow organisms to survive through a
long dry season that is incompatible with active
life. Animals often estivate for 9–10 months of
the year, but there are numerous examples of
continuous estivation stretching for 2 or more
years. Spadefoot toads (Scaphiopus couchii) of
the American southwest are a good example of
anuran estivators. They may be active above
ground for less than 20 nights of the year (Tocque
et al., 1995). Their initial emergence from underground burrows is triggered by the first torrential
rains of the summer. With this stimulus, toads
quickly dig out from underground, males immediately set up breeding choruses, and mating and
egg-laying occur within the first night in rainwater
ponds. Adults then become ravenous eaters and
over the next few days replenish the fuel reserves
needed to sustain them for many more months
underground.
Animal adaptations for life in stressful environments typically include adjustments at multiple
levels—behavioral, physiological and biochemical—and estivation is no different. The critical
elements for long-term survival during estivation
are water retention and sufficient fuel reserves. At
a behavioral level, species seek sheltered locations
in which to estivate that help them to conserve
body water, minimize their exposure to the elements, and hide them from predators. For example,
snails shelter in crevices, under logs, etc., lungfish
burrow into the mud of drying streams, and toads
dig deep underground. Fish often curl into a Ushape and anurans assume a crouched posture with
limbs drawn in under the body; these postures
reduce the surface area for evaporative water loss.
Estivators also express physiological adaptations
that defend the body from water loss during
dormancy. Because water is lost during breathing
and also across the skinyepithelium, animals typically enter estivation with large reserves of body
water that can be drawn upon to keep tissues
hydrated. Anurans, for example, have a huge
reserve of water in the bladder that is slowly
resorbed over time to replace tissue water lost by
evaporation. Evaporative water loss during
breathing is minimized by apnoic breathing patterns; for example, pulmonate land snails may
breath only 2–3 timesyh and show patterns of
discontinuous CO2 release and O2 uptake (Barnhart and McMahon, 1987). Evaporative water loss
across the integument is also reduced by physical
or chemical barriers. Snails secrete a mucus epiphragm to cover the aperture of the shell (Barnhart,
1983), lungfish secrete a cocoon of dried mucus
(Bemis et al., 1987), and several frog species
produce cocoons made of multiple layers of shed
skin (Loveridge and Withers, 1981; Pinder et al.,
1992). Water loss is also retarded by colligative
means by elevating the osmolality of body fluids
via the production of high concentrations of solutes. Urea is used for this purpose by various fish,
amphibian and snail species (Rees and Hand,
1993; Land and Bernier, 1995; Withers and Guppy,
1996); for example, in spadefoot toads (that do
not make cocoons) urea can rise to ;300 mM in
blood after several months of estivation (McClanahan, 1967). Ultimately, however, estivating
species also show high tolerance for tissue dehydration; estivating, arboreal and freeze tolerant
species all top the list of desiccation tolerant
anurans (Hillman, 1980; Shoemaker, 1992; Storey
and Storey, 1996). Spadefoot toads endure the loss
of as much as 60% of their total body water
(representing ;50% of their body mass) after
several months of natural estivation (McClanahan
1967) and, with bladders drained prior to experimental dehydration, they still endured the loss of
45% of body water (Hillman, 1980).
To deal with metabolic fuel supply during dormancy, estivation is prefaced by the laying down
of large reserves of endogenous fuels. For vertebrate species, energy metabolism during estivation
is based primarily on the aerobic oxidation of
lipids together with some protein catabolism and
a relatively small contribution by carbohydrates.
In spadefoot toads, for example, the net contributions to the total energy budget over the estivating
season were calculated as 72% from fatty acid
oxidation, 23% from protein, and 5% from carbohydrate (Jones, 1980). Protein use changes over
time, being low during the early weeks of estivation but rising as the water potential of the soil
declines and the demand for urea synthesis rises.
However, carbohydrate catabolism appears to be
of major importance for land snail estivation (Livingstone and de Zwaan, 1983). Fuel use over 7
months of estivation was monitored in two species
of mountain snails, Oreohelix strigosa and O.
subrudis (Rees and Hand, 1993). Polysaccharide
was the primary metabolic fuel for the initial 2–4
months of estivation and when this was depleted,
net protein catabolism began; a low rate of lipid
catabolism was maintained throughout.
K.B. Storey / Comparative Biochemistry and Physiology Part A 133 (2002) 733–754
Conservation of fuel reserves is a key issue for
long-term survival and this results from one of the
most important adaptations that support estivation:
metabolic rate depression. Metabolic rate during
estivation is low, typically only approximately 10–
30% of the corresponding resting metabolic rate
in aroused individuals at the same temperature
(Herreid, 1977; Bemis et al., 1987; Pinder et al.,
1992; Pedler et al., 1996). The greater the reduction in metabolic rate, the longer the time that a
fixed reserve of fuel can sustain basal metabolism.
Part of the reduction in metabolic rate comes from
the cessation of digestion and a lack of voluntary
movements, and part is due to reduced rates of
breathing and heart beat as well as apnoic
breathing patterns and their consequent effects on
pH and oxygen consumption in oxy-conforming
species. However, a substantial component is due
to a coordinated reduction in the rates of energy
turnover in tissues. For example, in vitro incubation studies with isolated tissues from both frogs
and snails have shown intrinsic metabolic depression that is sustained for several hours after tissue
excision (Guppy et al., 1994). This intrinsic component accounted for approximately 30% of the
overall metabolic depression observed when isolated mantle tissues from control vs. estivating
land snails, Helix aspersa, were compared; the
remaining 70% was attributed to extrinsic factors
(PO2, pH) that changed during estivation (Pedler
et al., 1996). Comparable studies with isolated
cells from H. aspersa hepatopancreas attributed
approximately half of the metabolic depression to
intrinsic factors (Guppy et al., 2000). Bishop and
Brand (2000) showed that metabolic depression
included proportional reductions in the rates of
three components of oxygen consumption: nonmitochondrial respiration, mitochondrial respiration driving ATP turnover, and mitochondrial
respiration driving proton cycling. Non-mitochondrial respiration decreased as oxygen tension was
lowered and was responsible for the oxygen-conforming behavior of the cells. This implicated
intrinsic factors in the suppression of mitochondrial
oxygen consumption during estivation. Using metabolic control analysis, Bishop et al. (2002)
assessed the influences on mitochondrial respiration rate that resulted in an approximate two-thirds
reduction in respiration rate in hepatopancreas cells
from estivating, compared with control, snails.
Mitochondrial respiration was divided into two
blocks: mitochondrial membrane potential produc-
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ers and mitochondrial membrane potential consumers, with membrane potential (Dcm) as the
intermediate connecting the blocks. Approximately
75% of the decrease in mitochondrial respiration
during estivation could be attributed to changes in
the Dcm producers (i.e. the kinetics of substrate
oxidation), with only 25% of the response attributed to changes in the Dcm consumers (i.e. ATP
turnover) (Bishop et al., 2002; Curtis et al., in
press). These data suggest the key importance of
controls that decrease substrate oxidation and ATP
production (rather than those that decrease ATP
use) in the development and regulation of the
hypometabolic state.
Factors contributing to intrinsic metabolic rate
depression include suppressed rates of fuel catabolism, ion channel arrest, and reduced rates of
protein synthesis (Churchill and Storey, 1989;
Storey and Storey, 1990; Rees and Hand, 1991;
Guppy et al., 1994). For example, analysis of
changes in the levels of glycolytic intermediates
during estivation in land snails, as well as estivation-induced changes in the kinetic properties of
enzymes in estivators, indicated glycolytic rate
depression during long-term estivation (Churchill
and Storey, 1989; Brooks and Storey, 1997). Suppression of oxidative metabolism was also indicated by cytochrome c oxidase (CCO) activities in
mitochondria from hepatopancreas of estivating
snails, Cepaea nemoralis, that were reduced by
84% (although total mitochondrial protein was
unaffected) (Stuart et al., 1998). Membrane-bound
proteins such as CCO are responsive to changes
in their phospholipid environment and the selective
suppression of CCO or other enzymes could come
from a change in the composition of mitochondrial
phospholipids. Notably, the cardiolipin content of
hepatopancreas mitochondrial membranes fell by
83% during estivation (Stuart et al., 1998) and
several studies have shown a direct dependence of
CCO activity on the presence of cardiolipin in the
mitochondrial inner membrane (Robinson, 1993).
Protein synthesis is also suppressed in estivating
animals. Estimates from in vitro studies with liver
slices from control vs. estivating Australian desert
frogs (Neobratrachus centralis) indicated a 67%
decrease in the rate of protein synthesis in estivating animals which accounted for 52% of the
metabolic depression in liver (Fuery et al., 1998).
The present article reviews some recent advances in our understanding of the biochemistry of
estivation with a particular focus on recent studies
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K.B. Storey / Comparative Biochemistry and Physiology Part A 133 (2002) 733–754
in our lab that address the role of reversible
phosphorylation in enzyme control, the signal
transduction mechanisms that mediate changes in
metabolism, adaptive changes to activities and
properties of selected enzymesyproteins, and the
role of gene expression in providing new protein
variants for specific roles in the hypometabolic
state.
2. Coordinated control by reversible protein
phosphorylation in metabolic rate depression
Key to the long-term maintenance of a hypometabolic state such as estivation are biochemical
mechanisms that strongly reduce net ATP turnover.
The transition to the hypometabolic state must be
regulated in order to achieve a coordinated suppression of many metabolic functions and set up a
new balance between ATP-producing and ATPutilizing reactions in cells. The need for such
coordinated control has been amply illustrated by
many studies of the injurious effects of hypothermia or hypoxia in mammals (summarized by
Hochachka, 1986). When ATP production in mammals is disrupted by a drop in temperature or
limited oxygen availability, cellular ATP levels fall
rapidly and membrane potential difference soon
collapses because ATP-dependent ion pumps (that
move ions against their concentration gradient)
can no longer keep pace with the opposing facilitated flow of ions through ion channels. When
membrane potential difference is lost, a variety of
destructive events are unleashed that quickly
become irreversible; many of these are Ca2qmediated and arise due to an uncontrolled influx
of Ca2q into the cytoplasm when depolarization
opens voltage-gated channels.
Organisms that incorporate facultative hypometabolism into their lifestyle (e.g. estivators, hibernators, anoxia-tolerant species) must implement a
suite of control mechanisms that both coordinate
an overall steep suppression of metabolic rate and
adjust the relative rates of many cell functions.
Thus, whereas rates of energy-expensive functions
such as ion pumping or protein synthesis are all
strongly suppressed in hypometabolic systems,
their percentage inhibition can vary widely in line
with their relative importance in the hypometabolic
state. In hibernating ground squirrels, for example,
the rate of protein synthesis in brain was reduced
to only 0.04% of the mean rate in euthermia
(Frerichs et al., 1998), whereas the activities of
sodiumypotassium ATPase were reduced by 40–
60% in most tissues and unaffected in heart (MacDonald and Storey, 1999).
The biochemical mechanism(s) used for metabolic suppression in estivation need to be powerful
yet easily reversible to allow a quick return to
normal metabolism during arousal. Indeed, estivators return to active life with great speed. Spadefoot toads are instantly alert even when they are
dug out of earth-filled tubs as gently as possible
and when estivating land snails, Otala lactea, are
misted with water the foot can emerge from the
shell in as little as 5 min. Thus, estivation is not a
dormancy in the sense that organisms are unresponsive or must undergo major metabolicydevelopmental changes in order to return to active life,
and this further emphasizes the fact the mechanisms of metabolic suppression in estivators must
be (a) rapidly reversible, and (b) require very
little de novo protein synthesis and reorganization
of metabolism.
One of the regulatory mechanisms that fits these
requirements is reversible protein phosphorylation
and in a variety of studies we have shown that
this mechanism plays a key role in the regulation
of multiple metabolic functions in estivation, hibernation, and anaerobiosis in both vertebrate and
invertebrate systems (Storey and Storey, 1990;
Storey, 1996, 1997, 1998; Brooks and Storey,
1997). Large changes to the activity state of many
enzymes and functional proteins can be made via
the addition or removal of covalently bound phosphate through the action of protein kinases and
protein phosphatases. The virtually onyoff control
of glycogen phosphorylase (GP) that is provided
by this mechanism is a classic example, and a
coordinated regulation of the rate of carbohydrate
catabolism in hypometabolic states is often supplied by reversible phosphorylation controls on
key regulatory enzymes including GP, 6-phosphofructo-1-kinase (PFK), pyruvate kinase (PK) and
pyruvate dehydrogenase (PDH).
2.1. Reversible phosphorylation control of enzymes
in land snails
The first demonstration that reversible phosphorylation was involved in metabolic control during
estivation came from studies of O. lactea. In a
series of studies we showed that carbohydrate
catabolism is suppressed in a coordinated fashion
in estivating snails via reversible phosphorylation
K.B. Storey / Comparative Biochemistry and Physiology Part A 133 (2002) 733–754
control over enzymes including GP, PFK, PK and
PDH (reviewed by Brooks and Storey, 1997).
Protein phosphorylation changes the charge distribution on enzymes so that the phosphorylated and
dephosphorylated forms of enzymes are separable
by isoelectrofocusing or ion exchange chromatography. Isofocusing trials provided evidence that
estivation alters the phosphorylation state of O.
lactea enzymes. The isoelectric point (pI) of both
PFK and PK was 5.85 in foot muscle of control
snails, but rose to 6.75 for PFK and 6.20 for PK
in muscle from estivating animals. By contrast, the
pI for aldolase, which is not regulated by reversible
phosphorylation, was 4.84 and 4.82 in the two
states.
Kinetic analysis of enzymes from control vs.
estivating snails documented both a stable modification of enzyme properties during estivation and
showed that kinetic properties during estivation
were consistent with a less active enzyme form.
Thus, PK from foot or mantle of estivating snails
showed changes, compared with controls, that
included a significant reduction in enzyme affinity
for both substrates, PEP and ADP (S0.5 values rose
by 18–83%) and an increase in enzyme sensitivity
to inhibitors, alanine and ATP (I50 values dropped
by 33–58%, except for a rise in I50 alanine of
mantle PK) (Whitwam and Storey, 1990). Similar
effects were seen for PK from foot muscle of H.
aspersa (Fields, 1992) and ventricle of H. lucorum
(Michaelidis and Pardalidis, 1994). The changes
in O. lactea PK properties during estivation were
very similar to those that resulted from anoxia
exposure of the snails and were qualitatively the
same as the anoxia-induced phosphorylation of PK
that occurs in various marine molluscs (Brooks
and Storey, 1997). Thus, it appears that the same
controls over PK activity are used in estivation
and anoxia to facilitate overall glycolytic rate
suppression.
An analysis of the time course of changes in
the I50 for ATP by foot PK showed that this
parameter dropped from aerobic to estivating values between 12 and 48 h after food and water
were withdrawn, but rebounded within 10 min
when 22 day estivated snails were aroused by
misting with water. The mechanism of estivationinduced changes in PK properties was confirmed
from in vitro incubation studies. Stimulation of
protein kinase activities in foot muscle extracts
from control snails resulted in a reduction in the
I50 value for alanine of PK, but did not affect the
737
enzyme from estivating snails whereas treatment
with alkaline phosphatase elevated the I50 value of
estivator PK, but did not affect the control enzyme
(Whitwam and Storey, 1990). Hence, the enzymes
in control vs. estivating snails were identified as
the low and high phosphate forms of PK, respectively. The identity of the protein kinase involved
in PK control during estivation or anoxia in O.
lactea remains to be determined. Levels of cyclic
AMP drop in both foot and hepatopancreas over
the early hours of estivation (Brooks and Storey,
1996) and the percentage of protein kinase A
present as the active catalytic subunit remains
constant or decreases in estivation (Brooks and
Storey, 1994a). Therefore, a role for protein kinase
A in the suppression of PK activity in estivation
seems unlikely (Brooks and Storey, 1994a). However, cyclic GMP levels rise during the early hours
of estivation (and during anoxia exposure) in O.
lactea and this implicates protein kinase G in the
stress-induced suppression of PK activity in snails
(Brooks and Storey, 1996).
Similar results were seen when PFK was
assessed in control vs. estivating snails (Whitwam
and Storey, 1991). Stable modifications to fructose-6-phosphate substrate affinity and inhibitory
activator constants were found that were consistent
with a less active enzyme form in estivating
animals (Table 1). Furthermore, in vitro incubation
studies indicated that estivation-induced changes
in enzyme properties were the result of PFK
phosphorylation so that, as with PK, the enzymes
in control vs. estivating snails were identified as
the low and high phosphate forms of PFK, respectively. PDH, the enzyme that gates the entry of
carbohydrate into the tricarboxylic acid cycle, is
also regulated by reversible phosphorylation during
estivation in O. lactea. PDH occurs in active
(dephosphorylated) and inactive (phosphorylated)
forms. The percentage of PDH present in the active
a form decreased from approximately 98% in
controls to approximately 60% in estivating animals over the first 25–30 h, but rebounded to
control values within 1 h when snails were aroused
(Brooks and Storey, 1992).
Oppositely directed controls on PFK and fructose-1,6-bisphosphatase (FBPase) are often used
to promote unidirectional flux through glycolysis
or gluconeogenesis depending on tissue metabolic
requirements. In vertebrate liver, for example, both
enzymes are regulated by fructose-2,6-bisphosphate (F2,6P2) and AMP, but these metabolites
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K.B. Storey / Comparative Biochemistry and Physiology Part A 133 (2002) 733–754
Table 1
Kinetic properties of 6-phosphofructo-1-kinase and fructose1,6-bisphosphatase from Otala lactea hepatopancreas: control
vs. 22 day estivation
Control
Estivated
PFK
Activity, mmolyminyg wet wt.
S0.5 fructose-6-P, mM
Ka AMP, mM
Ka fructose-2,6-P2, mM
I50 citrate, mM
0.49"0.04
2.4"0.2
300"10
0.70"0.09
46"3.5
0.78"0.05a
3.2"0.1a
230"10a
2.3"0.4a
24"0.5a
FBPase
Activity, mmolyminyg wet wt.
S0.5 fructose-1,6-P2, mM
I50 AMP, mM
I50 fructose-2,6-P2, mM
I50 CaCl2, mM
0.45"0.04
1.04"0.48
53.5"3.8
0.40"0.06
0.54"0.04
0.53"0.11
1.13"0.14
56.6"10.6
0.33"0.01
0.62"0.02
a
Significantly different from the corresponding control, P0.05.
Data are means"S.E.M., ns3 for FBPase and ns4–10 for
PFK. Data for PFK are from Whitwam and Storey (1991) and
the preparation of hepatopancreas extracts for FBPase assay
was also as described in that paper. Assay conditions for FBPase were 50 mM imidazole buffer, pH 7.2, 0.2 mM NADP, 5
mM MgCl2 with varying F1,6P2 for S0.5 determinations, and 7
mM F1,6P2 for I50 determinations. I50 values are defined as the
inhibitor concentration that reduces enzyme velocity by 50%
and were determined from plots of Vo yVi vs. wIx.
activate PFK and inhibit FBPase. Phosphorylation
by cAMP-dependent protein kinase also has opposite effects on the two liver enzymes, inhibiting
PFK but activating FBPase. During estivation,
however, the rates of both glycolysis and gluconeogenesis should be suppressed as part of the
overall metabolic depression, and so we hypothesized that phosphorylation-mediated suppression
of PFK in tissues of estivating O. lactea (discussed
above) would not be met by an opposite activation
of FBPase. Indeed, this seems to be the case.
Contrary to the results for PFK, an analysis of the
kinetic properties of FBPase from hepatopancreas
of control vs. estivating snails showed no significant differences in a range of enzyme properties
including maximal activity, Km for substrate, and
I50 values for inhibitors between the two states
(Table 1).
However, activities of both PFK and FBPase
may be strongly influenced by changes in the
levels of the allosteric effector, fructose-2,6-bisphosphate (F2,6P2), during estivation. Levels of
F2,6P2 decreased by 11-fold in hepatopancreas
during estivation, from 0.56"0.20 mM in controls
to 0.05"0.01 mM in 22 day estivated snails
(Whitwam and Storey, 1991). Reduced F2,6P2
could potentiate FBPase activity (and gluconeogenesis in general) during estivation by releasing
enzyme inhibition by this metabolite whereas,
oppositely, the reduced F2,6P2 levels, coupled with
the 3.5-fold rise in Ka F2,6P2 for PFK (Table 1),
would greatly lower the influence of this activator
on PFK activity during estivation. High F2,6P2
typically mediates a high glucose signal in cells
that potentiates the use of carbohydrate for anabolic purposes (Hue and Rider, 1987). Low
F2,6P2, by contrast, potentiates glucose export in
two ways: (a) by inhibiting PFK so that glycogenolysis can be directed into glucose output; and (b)
by activating FBPase to promote gluconeogenesis.
Indeed, F2,6P2 levels are reduced in liver-like
tissues in multiple situations where glycolysis is
inhibited andyor metabolic rate is suppressed
including hypoxia, anoxia, starvation, freezing and
responses to various hormones (Hue and Rider,
1987; Storey, 1988; Vazquez-Illanes and Storey,
1993). Gluconeogenesis is typically important in
states of aerobic dormancy such as estivation and
hibernation because it utilizes inputs of glycerol
(from triglyceride catabolism) or amino acids
(from protein breakdown) to provide a sustained
low production of glucose that is needed to fuel
selected tissues. Hence, changes in tissue F2,6P2
levels may be key to shifting the relative flux
through the PFK vs. FBPase loci during estivation
in favor of FBPase and gluconeogenesis.
Estivation also had a marked effect on the
enzyme that is responsible for F2,6P2 synthesisy
degradation. This enzyme is bifunctional. Acting
as a 6-phosphofructo-2-kinase (PFK-2) it synthesizes F2,6P2, but in the reverse direction it acts as
a fructose-2,6-bisphosphatase (F2,6Pase) to catabolize F2,6P2. Phosphorylation of the mammalian
liver enzyme by cAMP-dependent protein kinase
activates the F2,6Pase function and inactivates the
PFK-2 activity (Hue and Rider, 1987). Analysis
of the PFK-2 activity in O. lactea foot and hepatopancreas is shown in Table 2. Major changes in
the kinetic properties of foot muscle PFK-2 were
noted during estivation. The enzyme from 22 day
estivated snails showed a Km for fructose-6-P that
was 3.1-fold higher than that of the control enzyme
whereas the Km ATP dropped to 42% of the control
value. Both of these stable changes in enzyme
properties indicate of a covalent modification of
PFK-2 during estivation, probably as a result of
enzyme phosphorylation. These estivation-induced
K.B. Storey / Comparative Biochemistry and Physiology Part A 133 (2002) 733–754
changes in properties would reduce the enzyme’s
PFK-2 activity and probably enhance its F2,6Pase
activity. The effect of estivation on Km fructose-6P would particularly influence enzyme function
for levels of this substrate drop in vivo in foot
during estivation (from 16 mM in controls to 7
mM in 22 day estivating snails) whereas ATP
levels in foot were substantially higher than the
enzyme Km value in both cases (830–1160 mM)
(Churchill and Storey, 1989). In hepatopancreas a
different effect of estivation on PFK-2 was seen.
Maximal activity of the enzyme was reduced by
38% during estivation, without changes in enzyme
kinetic properties (Table 2). This suggests a possible down-regulation enzyme synthesis during
estivation that could be responsible for the reduced
levels of F2,6P2 in hepatopancreas. Effects of
estivation on PFK-2 in both tissues are consistent
with a starved state and similar responses by
mammalian liver PFK-2 characterizes carbohydrate sparing during starvation in mammalian liver
(Hue and Rider, 1987).
2.2. Reversible phosphorylation control of enzymes
in spadefoot toads
Reversible phosphorylation control also plays a
major role in the control of metabolic rate depression in estivating toads. Cowan and Storey (1999)
evaluated estivation-associated changes in PK and
PFK in toad skeletal muscle. Isoelectrofocusing
revealed that two forms of each enzyme were
present in toad muscle and that their proportions
changed substantially after 2 months of estivation
at 15 8C. For both enzymes, estivation led to a
strong increase in the proportion of total activity
in peak II (pIs6.2–6.4), compared with peak I
(pIs5.3–5.4) (Fig. 1). For PK the distribution
739
Fig. 1. Pyruvate kinase elution in two peaks after isoelectric
focusing of spadefoot toad leg muscle extracts. (a) Extract
from control toads, (b) extract from 2 month estivated toads.
PK activity is expressed as a percentage of the total activity
applied to the column. From Cowan and Storey (1999).
was 23% in Peak I and 77% in Peak II in muscle
extracts from control toads, whereas after estivation the proportions were 13% in Peak I and 87%
in Peak II. For PFK, approximately 50% of activity
was in each peak in extracts from control toads
but this changed to ;25% in Peak I and 75% in
Peak II in estivators. In vitro incubation of muscle
extracts with 32P-ATP resulted in strong radiolabeling of the enzymes in Peak I, but not Peak II.
This provided evidence that the basis of the estivation-associated change in enzyme distribution
was reversible protein phosphorylation and, furthermore, that the effect of estivation was to
increase the proportion of the dephosphorylated
(Peak II) enzymes in muscle. Reverse phase HPLC
also confirmed that the subunits of PK and PFK
found in Peak I contained 32P. Evidence that
cAMP-dependent protein kinase A (PKA) was the
enzyme responsible for the phosphorylation of
peak I enzymes came from in vitro incubations in
the presence vs. the absence of the PKA inhibitor,
PKA-I (5-24), followed by immunoprecipitation
Table 2
Properties of 6-phosphofructo-2-kinase from Otala lactea: effect of 22 day estivation
Foot muscle
Activity, nmolyminyg wet wt.
Km fructose-6-P, mM
Km ATP, mM
a
Hepatopancreas
Control
Estivated
Control
Estivated
0.60"0.08
8.77"2.53
73.2"7.80
0.48"0.04
27.4"4.56a
30.5"6.90a
0.92"0.09
10.7"1.00
590"55
0.57"0.05a
9.90"1.00
650"100
Significantly different from the corresponding control value, P-0.05.
Tissue extracts were prepared and PFK-2 assayed as in Vazquez-Illanes and Storey (1993), except that the supernatant from fractionation with 12% wyv polyethylene glycol 8000 was used for assay with optimal substrate conditions of 1 mM fructose-6-P (q3
mM glucose-6-P), and 2.5 or 5 mM Mg.ATP for foot or hepatopancreas. Data are means"S.E.M., ns3 for Km values, ns10 for
activity.
740
K.B. Storey / Comparative Biochemistry and Physiology Part A 133 (2002) 733–754
of PK or PFK with specific antibodies. The amount
of radiolabeled PK in immunoprecipitates was 36fold higher when extracts were incubated with 32PATP and PKA alone, compared with incubations
that also contained PKA-I; the comparable value
for PFK was 3.2-fold.
Kinetic analysis also showed significant differences in properties of the two forms of toad
skeletal muscle PK and PFK. For PK, the Km for
phosphoenolpyruvate and the Ka for fructose-1,6bisphosphate of the Peak II (low phosphate)
enzyme were 1.6-fold and 2.4-fold higher, respectively, than the values for the Peak I (high phosphate) form (Cowan and Storey, 1999). These
differences suggest that Peak II PK is the less
active enzyme form, and coupled with the shift to
predominantly the Peak II form during estivation,
the data are consistent with a suppression of PK
activity in estivating muscle. Phosphorylation of
vertebrate skeletal muscle PFK typically occurs
during exercise and the phospho-enzyme shows
increased binding to myofibrils in active muscle
(Foe and Kemp, 1982). Therefore, the increased
content of the Peak II, low phosphate, form of
PFK in muscle of estivating, inactive toads is also
consistent with metabolic suppression during estivation. Peak II PFK also showed reduced sensitivity to inhibition by Mg:ATP (I50 50% higher)
compared with Peak I PFK, suggesting that the
enzyme in estivating muscle is less tightly regulated by cellular adenylate status than in control
toads. Data for both enzymes are consistent with
less active enzyme forms and overall metabolic
rate depression during estivation.
3. Signal transduction
3.1. Protein kinase A
Given the prominent role of reversible phosphorylation in enzymatic control during estivation, it
was obvious that protein kinases and protein phosphatases must play a role in transitions to and
from the hypometabolic state during estivation.
Effects of estivation on the activities of seriney
threonine protein kinases and phosphatases have
recently been evaluated in spadefoot toad organs
(Cowan et al., 2000). The reduced content of Peak
I enzymes (both PK and PFK) during estivation
suggested a reduced activity of PKA during estivation and indeed, the organ-specific responses of
PKA during estivation were consistent those pre-
Table 3
Effect of estivation on protein kinases A and C in spadefoot
toad tissues: the percentage of PKA present as the active catalytic subunit (PKAc) and the percentage of PKC present as
the active, membrane-bound enzyme
% PKAc
Control
Brain
Liver
Muscle
Heart
Kidney
57.0"8.0
97.0"5.0
80.0"6.0
38.0"3.0
82.0"2.0
% PKC membrane
bound
Estivated
a
25.0"3.0
62.0"8.0a
47.7"1.0a
20.0"5.0a
62.0"4.0a
Control
Estivated
45.8"1.6
46.6"5.7
29.6"5.6
62.2"1.4
70.8"12.0
21.5"3.2a
16.4"2.4a
29.8"3.9
65.6"1.1
18.7"4.7a
a
Significantly different from the corresponding control value, P-0.05. From Cowan et al. (2000).
Note that total PKA activity did not change in any tissue
during estivation whereas total PKC activity changed only in
liver and kidney. Data are means"S.E.M., ns4.
dictions. Table 3 shows that the percentage of the
enzyme present as the free catalytic subunit
(%PKAc), the active form, was strongly reduced
in all organs tested during estivation. A common
role for PKA is the activation of fuel catabolism,
frequently as a response to hormones such as
adrenaline or glucagon; for example, PKA mediates the activation of both glycogen phosphorylase
and hormone sensitive lipase to increase the breakdown of carbohydrate and lipid stores, respectively.
Therefore, the reduced %PKAc in toad organs
during estivation is consistent with lower rates of
fuel catabolism and other PKA-mediated processes
during dormancy. However, total PKA activity was
unaltered during estivation (Cowan et al., 2000)
and this maintains organ sensitivity to PKA-mediated signals, so that fuel catabolism and metabolic
rate can be rapidly re-elevated during the arousal
process.
3.2. Protein kinase C
Ca2q-activated, phospholipid-dependent protein
kinase C (PKC) mediates intracellular responses
to multiple events including cell proliferation, differentiation, exocytosis, and neural activity (Kikkawa et al., 1989). The enzyme is distributed
between membrane-bound and cytosolic forms and
in response to extracellular stimuli that elevate
phosphatidylserine and Ca2q, PKC is translocated
from inactive cytosolic pools to the plasma
membrane where it becomes active. Hence, a key
parameter in the analysis of PKC activity is the
K.B. Storey / Comparative Biochemistry and Physiology Part A 133 (2002) 733–754
percentage that is membrane-bound. Analysis of
the percentage of membrane-bound PKC in toad
organs showed a strong reduction in this parameter
in all toad tissues tested during estivation (Table
3). In addition, total PKC activity also decreased
by 57% in liver during estivation (Cowan et al.,
2000). The net effect then, was a reduction in the
amount of active membrane-bound PKC during
estivation, to 50% of the control value in brain
and kidney and to just 13% in liver. This low
amount of active PKC is consistent with the
reduced organ metabolic rate of estivation and
suggests that various PKC-regulated metabolic
processes are suppressed in the hypometabolic
state. Furthermore, the similar responses of PKA
and PKC suggest that suppression of protein
kinase-mediated signal transduction pathways is
important for the induction and maintenance of
the hypometabolic state during estivation.
3.3. Protein phosphatases 1 and 2
The responses by protein phosphatases to metabolic stress are typically opposite to those of their
protein kinase partners. However, an analysis of
estivation effects on the activities of protein phosphatase (PP) types 1, 2A and 2C in toad organs,
showed only a few changes in phosphatase activities in the hypometabolic state (Cowan et al.,
2000). PP-1 was quantified in dilute, trypsintreated extracts to release any enzyme bound to
inhibitor proteins and give a measure of total PP1 activity and also in untreated extracts to measure
the free, active enzyme (Price et al., 1986). Total
PP-1 activity (in trypsin-treated extracts) and the
percentage of free, active enzyme (in untreated
extracts) changed in four organs of estivating
toads, but changes in these two parameters were
generally oppositely directed so that the net change
in active PP-1 was low in liver, heart and gut
(Cowan et al., 2000). However, active PP-1 in
kidney fell from 1.16 Uymg in control toads to
0.33 Uymg in estivated animals. What is striking
about these results is that whereas PKA and PKC
activities were uniformly suppressed in toad
organs, PP-1 activities did not show an opposite
rise but remained constant or were reduced. PP2A activity was also constant (brain, skeletal
muscle, lung, kidney) or reduced by 29–43%
(liver, heart, gut) in estivating toads, whereas PP2C activity decreased by 50% in heart, was unaffected in three organs, and rose in brain, muscle
741
and gut of estivating animals (Cowan et al., 2000).
The conclusion from this analysis is that the
normally oppositely-directed responses by seriney
threonine kinases and phosphatases are abandoned
during estivation, and both arms of these signal
transduction mechanisms are suppressed in
estivation.
3.4. Tyrosine kinases and phosphatases
Multiple signal transduction systems are present
in cells and all can potentially contribute to the
support of estivation. The possible roles of protein
tyrosine kinases (PTKs) and protein tyrosine phosphatases (PTPs) in organismal responses to environmental stress have received little consideration
to date. However, a new study with spadefoot
toads provides some initial evidence that PTKs
and PTPs have roles to play in estivation.
Protein phosphorylation on tyrosyl residues was
first identified approximately 20 years ago (Witte
et al., 1980; Hunter and Sefton, 1980) and reversible control of proteins via the actions of PTKs
and PTPs is now known to be involved in many
cell functions (Chen et al., 1987; Chou et al.,
1987). Both PTKs and PTPs exist in multiple
forms and in two broad classes: receptor
(membrane-associated) and non-receptor (or nontransmembrane) enzymes. Among PTKs, the
receptor types, such as the EGF, PDGF and insulin
receptor tyrosine kinases, have attracted the most
attention but cytosolic PTKs have also been found
in all mammalian tissues tested (Elberg et al.,
1995). Elevated PTK activities have been associated with multiple cellular responses including cell
differentiation in rat lung (Srivastava, 1985), hypotonic cell swelling of cardiac myocytes (Sadoshima et al., 1996), and tumor growth (Rosen et al.,
1986).
To assess PTK and PTP responses to estivation,
we chose an initial approach that broadly surveyed
PTK and PTP classes in control vs. 2 month
estivated toads by monitoring four parameters in
multiple tissues: changes in total enzyme activities,
changes in enzyme distribution between soluble
and insoluble fractions, changes in enzyme elution
patterns off ion exchange chromatography and, for
PTPs only, changes enzyme response to different
phospho-substrates (Cowan and Storey, 2001).
Analysis of these parameters for PTKs revealed
the following: (1) total PTK activity decreased
significantly by 27–52% in liver, lung and skeletal
742
K.B. Storey / Comparative Biochemistry and Physiology Part A 133 (2002) 733–754
Table 4
Effect of estivation on total protein tyrosine kinase and protein tyrosine phosphatase activities in organs of spadefoot toads, Scaphiopus
couchii
PTK
PTP
Active
Heart
Liver
Lung
Muscle
1499"171
3217"214
3224"293
2359"152
Estivated
(57%)
(58%)
(88%)
(71%)
a
2488"291 (66%)
1841"154a (60%)
1540"85a (71%)
1720"175a (63%)
Active
Estivated
12.5"2.2 (33%)
260"42.2 (37%)
67.9"14.6 (46%)
60.0"6.4 (46%)
21.8"2.2 (54%)a
118"12.8 (44%)a
57.0"10.9 (56%)
85.7"12.7 (72%)
a
Activities are nanomoles of phosphate transferred per minute per gram wet weight, ns3–4, measuring phosphate transfer onto a
poly Glu–Tyr substrate for PTK and phosphate removal from the peptide, ENDpYINASL, for PTP. Values in brackets show the mean
percentage of the total activity that is soluble. Modified from Cowan and Storey (2001).
Significantly different from the corresponding value in active toads, P-0.05.
muscle during estivation, but rose by 66% in heart
(Table 4). (2) The distribution of PTKs between
soluble, cytoplasmic (13 000=g supernatant) and
insoluble, membrane-associated (Triton-extracted)
forms shifted substantially in some organs during
estivation; for example, PTKs in lung decreased
from 88% soluble in controls to 71% soluble in
estivated toads (Table 4). (3) Changes in PTK
activities resulted from effects on both the soluble
and insoluble enzymes in heart and liver, but were
due to changes in soluble enzymes only in lung
and muscle (Fig. 2). (4) Three major peaks of
PTK activity were identified in both soluble and
insoluble fractions of skeletal muscle via DEAE
Sephadex chromatography and distinct shifts in
their elution positions were seen for one peak in
the soluble fraction, and two in the insoluble
fraction when extracts from control vs. estivated
toads were compared (Cowan and Storey, 2001).
A stable change in the elution position of an
enzyme on ion exchange is indicative of a covalent
modification that alters protein charge and this
could suggest that PTKs are themselves subject to
reversible protein phosphorylation during estivation. Thus, it is apparent that PTKs also respond
to estivation in toad organs, that their activities
may be modified by multiple mechanisms (altered
activities, solubleyinsoluble distribution, possible
reversible phosphorylation), and that they may be
involved in regulating responses by various metabolic processes in the hypometabolic state.
Analysis of PTP activities in spadefoot toad
tissues revealed that the same types of control
mechanisms applied to them (Cowan and Storey,
2001). For example, total PTP activity changed in
some organs (increased by 75% in heart, decreased
Fig. 2. Effect of estivation on the activities of protein tyrosine kinases in (a) soluble and (b) insoluble (membrane-associated, Tritonextracted) fractions of tissues from active (gray bars) and 2 month estivated (black bars) spadefoot toads. Data are Uymg protein,
means"S.E.M., ns4, one unit being the amount of enzyme that catalyzes the transfer of one nanomole of Pi to the poly Glu–Tyr
(4:1) substrate per minute at 30 8C. a—Significantly different from the corresponding control value by the Student’s t-test, P-0.05.
Modified from Cowan and Storey (2001).
K.B. Storey / Comparative Biochemistry and Physiology Part A 133 (2002) 733–754
by 55% in liver) during estivation (Table 4), as
did the distribution of PTPs between soluble and
insoluble fractions (e.g. soluble PTP activities rose
in skeletal muscle and heart and insoluble PTPs
decreased). Evidence of enzyme covalent modification came again from shifts in the elution position of two of the four peaks of soluble PTP
activity, and all three peaks of insoluble PTP
activity in skeletal muscle of estivating, vs. control,
toads. Furthermore, differential effects of estivation
on PTP activities were also revealed by assays
with two different peptide substrates: (a) a general
PTP substrate (ENDpYINASL) that represents a
highly conserved motif (DYINA) in the catalytic
domain of most PTPs and seems to be the better
substrate for soluble PTPs; and (b) the peptide
DADEpYLIPQQG which corresponds to the autophosphorylation site found in EGFR (a receptor
PTP), and is a substrate for PTPs that contain the
SH2 (src-homology) binding site. Assays revealed
substrate-specific differences in both the distribution of PTPs between soluble and insoluble fractions during estivation, and in the ion exchange
elution profiles of the enzymes (Cowan and Storey,
2001). For example, when assayed with ENDpYINASL, soluble PTP activity in heart rose during
estivation by 71%, whereas insoluble activity
decreased by 36%. However, just the opposite
occurred when heart PTP was assayed with
DADEpYLIPQQG; soluble PTP activity decreased
by 75% whereas insoluble activity rose three-fold.
These results for heart as well as differential
responses to the two substrates in most other
tissues provided evidence that the receptor and
non-receptor PTPs respond differently during
estivation.
Hence, current data suggest that changes in
PTKs and PTPs have an impact on estivation and
provide a starting point for future studies. Whether
this is a positive impact in stimulating metabolic
changes that support estivation or a case that PTK
or PTP mediated processes are suppressed in the
hypometabolic state, remains to be determined as
does the identification of specific PTK or PTP
types and their specific actions in estivation. The
data for heart are particularly interesting, however,
in that the elevated total activities of PTK and
PTP in heart contrast with the opposite effects of
estivation in suppressing the activities of seriney
threonine kinases and phosphatase in heart (PKA,
PP-2A, PP-2C were all reduced in heart; Cowan
et al., 2000). Hence, the influence of seriney
743
threonine kinases and phosphatases is apparently
reduced during estivation in heart, whereas that of
the tyrosine kinases and phosphatases is enhanced.
4. Estivation and metabolic enzymes
Estivation is a complex phenomenon and several
influences may be at work to readjust metabolism
for optimal function over the long months of
dormancy. One of these is starvation and the
pattern of stored fuel use during estivation, typically changes to include a major reliance on the
oxidation of body lipid reserves, a low rate of
gluconeogenesis from glycerol or amino acids to
maintain glucose supply for selected organs, and a
gradual rise in protein catabolism as the demand
for urea synthesis increases. Thus, for estivating
toads the general expectations associated with the
starved state during estivation would be suppression of glycolysis and fatty acid synthesis, and
increased protein catabolism, gluconeogenesis,
ketone body metabolism, and lipid oxidation.
Another influence on metabolic reorganization during estivation may be metabolic arrest. Whereas
some enzymes are regulated via reversible phosphorylation to achieve the hypometabolic state
during estivation (see above), and some may be
influenced by changes in the lipid environment in
which they reside (Stuart et al., 1998), others may
be controlled by synthesis or degradation with the
resulting changes in the maximal activities of
selected enzymes contributing to the suppression
of different metabolic pathways. Further influences
on metabolism during estivation probably also
include: (a) dehydration stress may demand changes in selected enzymesypathways in addition to
the known elevation of urea cycle enzymes in
response to waterysalt stress in anurans (Balinsky,
1981); (b) maturation of reproductive tissues during estivation may require metabolic adjustments
in selected tissues. To assess the extent of metabolic reorganization that occurs during estivation,
we analyzed changes to the enzymatic make-up of
spadefoot toad organs during estivation, focusing
on two groups—enzymes of intermediary energy
metabolism and enzymes of antioxidant defense.
4.1. Enzymes of intermediary metabolism
The maximal activities of a variety metabolic
enzymes representing glycolysis, gluconeogenesis,
and amino acid, fatty acid, ketone body, and
744
K.B. Storey / Comparative Biochemistry and Physiology Part A 133 (2002) 733–754
adenylate metabolism were assessed in three tissues (brain, liver, and skeletal muscle) of active
toads vs. those that were subsequently estivated
for 2 months (Cowan et al., 2000). Estivation had
little effect on enzymes in brain with activities of
17 out of 23 enzymes unaffected by estivation.
Effects in brain included reduced activities during
estivation for two enzymes of glycolysis, PFK and
lactate dehydrogenase (LDH), as well as creatine
kinase (CK) (activities fell by 25, 40 and 25%,
respectively). By contrast, the activity of glutamate
dehydrogenase, a key enzyme of amino acid
metabolism, rose by 3.2-fold whereas glucose-6phosphate dehydrogenase activity increased by
70%, and b-hydroxybutyrate dehydrogenase rose
by 50%. These enzymatic changes in brain are
consistent with suppressed glycolysis and
increased ketone body and amino acid catabolism
during estivation and hence, are primarily related
to a reorganization of fuel use.
Toad liver also showed relatively few changes
to the activities of metabolic enzymes during
estivation with only nine out of 25 enzymes
affected. The maximal activities of eight enzymes
of carbohydrate, amino acid, ketone body and
phosphagen metabolism decreased during estivation (by 25–60%), whereas activity of the gluconeogenic enzyme, malic enzyme (ME), increased
by 2.4-fold (Fig. 3a) (Cowan et al., 2000). The
percentage of GP present in the active a form also
decreased significantly in liver of estivated toads
from 27.6"3.33% in controls to 16.8"1.97% in
estivation (Cowan et al., 2000). All of these
changes in liver enzymes are again consistent with
changes in patterns of fuel use and with a starved
state during estivation. Reduced activities of GP
and three other glycolytic enzymes are consistent
with carbohydrate sparing, and the effects on GP
are also those that are expected from the reduced
percentage of PKA present as the active catalytic
subunit in liver of estivating animals. Only one
metabolic enzyme showed a pronounced increase
in activity in liver during estivation and that was
ME, which has an anaplerotic role in the provision
of 4-carbon units (oxaloacetate, derived from malate) to the tricarboxylic acid cycle for condensation with acetyl-CoA. During lipid oxidation
(providing 2-carbon acetyl-CoA inputs only), the
supply of oxaloacetate can be rate-limiting and
malate synthesis using pyruvate derived from a
low rate of carbohydrate or amino acid catabolism
keeps the TCA cycle primed with 4-carbon units
to allow efficient lipid oxidation. Hence, a 2.4fold increase in ME activity during estivation
would increase the potential for malate synthesis
from pyruvate when the organ switches to fatty
acids as a primary fuel during estivation.
In skeletal muscle, however, the activities of
most metabolic enzymes changed significantly during estivation, with only three out of 27 unaffected.
Of these, the activities of 14 enzymes increased
whereas 10 decreased (Cowan et al., 2000). Notably, as in brain and liver, these effects were
independent of changes in tissue water content and
hence, indicate specific and widespread reorganization of muscle metabolism during estivation.
Contrary to the results for the two other organs,
activities of four enzymes of glycolysis (PFK, PK,
aldolase, and glyceraldehyde-3-phosphate dehydrogenase) increased during estivation, although a
reduced activity of hexokinase and no change in
GP argue against an increased rate of carbohydrate
utilization. It is not clear what the purpose of these
changes could be. One possibility might be that
an enhanced glycolytic capacity, coupled with
elevated CK (producing quick ATP from creatine
phosphate) and citrate synthase (increasing the
capacity for carbohydrate entry into the Krebs
cycle), could develop during estivation in anticipation of the high levels of muscular work that
will be needed upon emergence from estivation,
to support the frenzied breeding that is characteristic of this species. Such a proposition, however,
will require detailed measurements of muscle enzymatic make-up both seasonally and over estivationyarousal cycles.
Amino acid oxidation also appeared to be facilitated in muscle of estivating toads with elevated
activities of serine dehydratase, glutamate-pyruvate
transaminase and glutamate dehydrogenase
(GDH). Elevated GDH, also seen in brain, suggests an increased capacity for NHq
4 production in
support of urea synthesis by liver. GDH activities
in toad liver did not change during estivation but,
notably, they were very high to start with, 18-fold
higher than in muscle (Cowan et al., 2000). These
changes in amino acid metabolizing enzymes correlate with a suite of changes in free amino acid
levels in tissues of estivating toads (Cowan et al.,
2000). In muscle, glutamate, alanine and valine
were elevated by almost two-fold during estivation
whereas glutamine was reduced by nearly half.
The same pattern was seen in brain and liver
(except for glutamine in liver which increased),
K.B. Storey / Comparative Biochemistry and Physiology Part A 133 (2002) 733–754
745
Fig. 3. Maximal activities of selected enzymes in liver of active vs. 2 month estivated spadefoot toads, S. couchii. (a) Metabolic
enzymes measured in active toads (gray bars) vs. toads that were subsequently allowed to estivate for 2 months (black bars). Abbreviations: GAPDH, glyceraldehyde-3-phosphate dehydrogenase; PK, pyruvate kinase; LDH, lactate dehydrogenase; IDH, NADP-dependent isocitrate dehydrogenase; GPT, glutamate-pyruvate transaminase; CK, creatine kinase; BDH, b-hydroxybutyrate dehydrogenase;
FBP, fructose-1,6-bisphosphatase; ME, malic enzyme (from Cowan et al., 2000). (b) Enzymes of antioxidant defense in toads estivated
for 2 months vs. active toads that were aroused from estivation and sampled 10 days later. Abbreviations are: GST, glutathione-Stransferase; GR, glutathione reductase; Se-GPX, selenium-dependent glutathione peroxidase; Tot-GPX, total glutathione peroxidase;
CAT, catalase; SOD, superoxide dismutase. Compiled from data in Grundy and Storey (1994). Data are means"S.E.M., ns3–4 for
metabolic and ns4–9 for antioxidant enzymes. Values for LDH and CAT should be multiplied by 10. *—All values for estivated toads
are significantly different from their corresponding value in active aroused toads, P-0.05.
K.B. Storey / Comparative Biochemistry and Physiology Part A 133 (2002) 733–754
746
Table 5
Free amino acid levels in three organs of land snails, Otala lactea: effects of 45 h anoxia exposure under a 95% N2y5% CO2 atmosphere
or 22 days of estivation at 21 8C
Foot
Control
Aspartate
Glutamate
Asparagine
Glutamine
Arginine
Alanine
Valine
Total AA
1.49"0.47
1.16"0.32
1.99"0.32
6.53"0.77
2.32"0.28
0.75"0.10
0.72"0.14
15.9"4.28
Hepatopancreas
Anoxic
1.04"0.11
2.30"0.34
0.34"0.04a
2.25"0.36a
3.80"0.20b
2.16"0.24a
0.28"0.03b
13.5"1.48
Estivated
2.10"0.41
4.17"0.49a
0.23"0.01a
0.92"0.19a
2.63"0.40
0.80"0.11
0.33"0.08b
13.7"3.19
Control
1.74"0.15
2.24"0.41
0.55"0.08
2.99"0.39
0.80"0.07
0.71"0.09
0.70"0.07
15.0"1.37
Mantle
Anoxic
Estivated
a
0.42"0.05
2.17"0.56
0.16"0.04a
1.17"0.11a
1.02"0.12
0.63"0.17
0.33"0.07a
8.89"1.43b
a
1.06"0.13
8.34"1.09a
0.25"0.04a
0.97"0.19a
0.53"0.18
0.94"0.24
0.43"0.01b
16.0"1.71
Control
Anoxic
Estivated
0.97"0.17
1.23"0.27
0.47"0.07
1.15"0.20
0.41"0.05
0.50"0.19
0.30"0.03
7.42"0.98
0.83"0.11
3.82"0.68a
0.35"0.06
2.62"0.44b
2.76"0.28a
2.06"0.24a
0.31"0.06
15.4"1.10a
1.90"0.24a
4.71"0.31a
0.14"0.05b
1.00"0.25
1.67"0.27a
0.78"0.20
0.41"0.05
12.4"1.18b
a
Significantly different from the corresponding control, P-0.01.
P-0.05.
Data are in micromoles per gram wet weight, means"S.E.M., ns3–4. Levels of 13 other amino acids did not change significantly
under either experimental condition. Amino acids were quantified by HPLC as per Churchill and Storey (1992).
b
so it appears that these changes may be characteristic of some anuran tissues when poised for urea
synthesis from protein catabolism.
Free amino acid levels were also affected by
estivation (22 days) in O. lactea tissues as shown
in Table 5. These data are compared with the
effects of anoxia stress (45 h under a nitrogen gas
atmosphere) because molluscs have well-known
amino acid responses to anoxia. As in toads, a
major and consistent response to estivation in O.
lactea was a pronounced increase in glutamate by
3.6–3.8-fold in all organs. Correlated with this,
although not stoichiometrically equivalent, was a
large decrease in glutamine in two organs with
levels falling to 14 and 32% of control values in
foot and hepatopancreas, respectively. Glutamine
conversion to glutamate releases an ammonium
ion, which may be used for urea biosynthesis.
Asparagine also decreased consistently in all three
tissues during estivation and both of its products,
NHq
and aspartate, are substrates of the urea
4
cycle. Valine also decreased in foot and hepatopancreas. Substantive changes in free amino acids
during anoxia exposure in O. lactea were somewhat different and included significant increases
in arginine and alanine in foot muscle and mantle.
Alanine is a well-known metabolic end product of
anaerobic metabolism in molluscs (Hochachka and
Somero, 1984), and arginine is the product of the
hydrolysis of the phosphagen, arginine phosphate,
which indicates that snails deplete this energy
reserve under low oxygen stress. Arginine also
rose by four-fold in the mantle of estivating snails,
which indicates that some tissues may also be
under energy stress during estivation. As in estivation, asparagine and glutamine also decreased in
anoxia in foot and hepatopancreas whereas aspartate, which is a good anaerobic substrate in marine
molluscs, decreased only in hepatopancreas.
Hence, substantial differences in amino acid
metabolism were seen in the responses to estivation and anoxia, which indicate that free amino
acids have different roles to play in each
phenomenon.
4.2. Enzymes of antioxidant defense
The effect of estivation on enzymes of antioxidant defense has been analyzed in both toads and
snails (Hermes-Lima and Storey, 1995; Grundy
and Storey, 1998; Hermes-Lima et al., 2001). In
toads, the activities of antioxidant enzymes were
generally significantly lower in 2 month estivating
animals than in active toads sampled 10 days after
arousal (Grundy and Storey, 1998). Fig. 3b illustrates this for liver; activities of five enzymes were
70–100% higher in active toads than in estivating
animals. The exception to this was superoxide
dismutase (SOD) which showed an activity 50%
lower in active toads. Results for other organs
were not as consistent as for liver, but in general,
for the six enzymes quantified in five other tissues
(heart, kidney, muscle, lung, gut) there were 12
instances where activities were higher in active
toads than in estivators, and only 4 where the
opposite was true (with no change in 14 cases).
In particular, glutathione-S-transferase (GST)
activity was significantly higher in active toads in
K.B. Storey / Comparative Biochemistry and Physiology Part A 133 (2002) 733–754
all organs except skeletal muscle (Grundy and
Storey, 1998). The rate of reactive oxygen species
generation and the accumulation of oxyradical
damage products correlates positively with oxygen
consumption in various species (Adelman et al.,
1988). Hence, the above data showing generally
lower activities of antioxidant enzymes in organs
of estivating toads are consistent with the significantly reduced rates of organ oxygen consumption
during estivation, as well as a reduced load of
xenobiotics to process due to the starved state.
Antioxidant defenses are clearly adaptable enzyme
systems that respond to the oxyradical load
imposed upon them.
By contrast, a different situation occurred in
estivating O. lactea. Activities of catalase (CAT),
SOD, glutathione peroxidase and GST were generally higher in hepatopancreas and foot muscle of
estivating snails than in active snails (HermesLima and Storey, 1995). It was postulated that
these adjustments to antioxidant defenses would
minimize damage due to a burst of reactive oxygen
species generation, occurring as a result of a rapid
increase in oxygen consumption over the early
minutes of arousal (Hermes-Lima et al., 1998,
2001). Indeed, levels of lipid peroxidation damage
products peak within 20 min when snails are
aroused which suggests an initial burst of oxyradical production (Hermes-Lima and Storey, 1995).
Periods of high oxygen uptake also occur intermittently during estivation; oxygen uptake rises by
up to 5-fold over short periods every 20–50 h
during which snails also hyperventilate and release
CO2 (Barnhart and McMahon, 1987). As opposed
to the continuous pattern of oxygen uptake seen
in active snails, this pattern of discontinuous oxygen uptake during estivation could also cause short
bursts of oxyradical production intermittently during estivation. Because snails may need to sustain
continuous estivation for over a year or more,
good antioxidant defenses may be critical to preventing or minimizing the accumulation of oxyradical damage products during prolonged estivation.
5. Urea effects on enzymesyproteins from
estivators
Many estivating species accumulate urea in their
body fluids to help retard the loss of body water
(Rees and Hand, 1993; Land and Bernier, 1995;
Withers and Guppy, 1996). Concentrations can
often reach several hundred millimolar; for exam-
747
ple, values of 200–300 mM have been measured
in spadefoot toads after several months of estivation (Jones, 1980; Grundy and Storey, 1998). Urea
is well known for its ability to denature proteins,
and researchers first became interested in the
potentially negative effects of high urea on body
proteins from studies of elasmobranch fish. These
maintain permanently high plasma and tissue urea
concentrations (400–500 mM) in order to keep
body fluids isosmotic with seawater. Multiple studies of urea effects on elasmobranch enzymes
revealed various disruptive effects of high urea on
the properties of selected enzymes (summarized in
Hochachka and Somero, 1984). However, these
effects were largely counteracted by the effects of
other osmolytes, the methylamines, that are also
present in elasmobranch body fluids. At the natural
ratio of 2:1 ureaymethylamines, net enzyme function was unperturbed.
Because of the situation in sharks, we wondered
whether or not spadefoot toad proteins would be
sensitive to urea, particularly in light of the fact
that there is little evidence of the presence of
counteracting solutes in estivating anurans. In spadefoot toads there is no osmolality gap that could
be filled by such solutes (McClanahan, 1967), and
three species of Australian frogs that accumulated
high urea during estivation (100–200 mM)
showed little or no accumulation of methylamines
(such as trimethylamine oxide or betaine that are
prominent in elasmobranchs) or other potential
counteracting solutes (Withers and Guppy, 1996).
Fuery et al. (1997) reported effects of urea on
pyruvate binding by LDH from Australian desert
frogs, but interestingly, 400 mM urea had less
effect on the Km for pyruvate of M4-LDH from
urea-accumulating anurans than it did on the
enzyme from non-accumulators which suggests an
adaptation for tolerance of high urea by the
enzyme of the estivating species.
In recent studies, we analyzed the effects of
urea on selected enzymes involved in intermediary
metabolism and antioxidant defense in spadefoot
toad organs and compared these with the effects
of high salt (KCl) (Grundy and Storey, 1994,
1998). The colligative effect of high urea helps to
retain water in toad tissues, but viewed from
another perspective the accumulation of urea also
minimizes the increase in the ionic strength of
body fluids that would otherwise occur as estivators dehydrate. Among vertebrates, amphibians
have the highest tolerance for variation in body
748
K.B. Storey / Comparative Biochemistry and Physiology Part A 133 (2002) 733–754
Fig. 4. Effect of urea and KCl on maximal activities of
enzymes from spadefoot toad skeletal muscle. Bars are: control
(light gray), q300 mM urea (dark gray), q200 mM KCl
(black). Enzymes are: PK, pyruvate kinase; PFK, phosphofructokinase; IDH, NAD-dependent isocitrate dehydrogenase;
and GDH, glutamate dehydrogenase. Data are expressed relative to control activities and are means"S.E.M., ns3–4. *—
Significantly different from the corresponding control, P0.01. Compiled from data in Grundy and Storey (1994).
water content and body fluid ionic strength, an
adaptation necessitated by their highly water-permeable skin (Hillman, 1988). However, although
anurans can endure high ion levels that would kill
mammals, there is undoubtedly also an upper limit
to their tolerance.
Fig. 4 shows the effect of 300 mM urea and
200 mM KCl on the maximal activities of PK,
PFK, GDH and NAD-dependent isocitrate dehydrogenase (IDH) from spadefoot toad skeletal
muscle (Grundy and Storey, 1994). Urea had no
effect on three enzymes, but lowered GDH activity
to 65% of control values. By contrast, 200 mM
KCl inhibited all four enzymes with a particularly
strong effect on GDH activity, which was reduced
to just 11% of control values. Similar responses to
urea and KCl were also seen when these same
enzymes were assessed in skeletal muscle of leopard frogs (Rana pipiens), a non-estivating species
(Grundy and Storey, 1994). Urea and KCl effects
on the kinetic properties of PK and PFK from
spadefoot toad muscle were also analyzed. Neither
300 mM urea nor 200 mM KCl changed the Km
values for substrates of either enzyme but calculated I50 values (inhibitor concentration reducing
activity by 50%) were instructive. For PK assayed
at low, near physiological levels of PEP (0.04
mM), the I50 value for KCl was ;300 mM
whereas I50 urea was ;1300 mM. For PFK the
I50 KCl was ;250 mM, whereas I50 urea was ;2
M. Hence, spadefoot toad enzymes are clearly
much more sensitive to high ion concentrations
than they are to high urea and, in the absence of
urea accumulation, it is likely that cellular ionic
strength during estivation would rise to a level
that could be deleterious both to individual
enzymes and to the integrated functioning of various metabolic pathways. Hence, a key consequence of the accumulation of urea is that it
minimizes the elevation of cellular ionic strength
that would otherwise occur, and thereby reduces
or prevents metabolic disruption by high ionic
strength.
Urea and KCl effects were also assessed during
the study of antioxidant enzymes in spadefoot
toads (Grundy and Storey, 1998). Again, 300 mM
urea had little or no effect on enzyme activities;
of three antioxidant enzymes assessed wGST, glutathione reductase (GR), CATx in six organs from
both control and estivating spadefoot toads, only
three instances were identified where 300 mM urea
significantly reduced enzyme maximal activity.
Urea reduced GR activity in kidney and GST in
lung of control (but not estivated) toads by 19–
20%, and urea lowered CAT activity in heart of
estivated toads by 26%. By contrast, the addition
of 200 mM KCl reduced GST and CAT activities
by 51–55% in all tissues of both control and
estivated toads, and also reduced GR activity in
liver by 25–40%. Hence, these data again support
the idea that a key function of urea accumulation
in toad tissues is to minimize the elevation of
cellular ionic strength that would otherwise occur
and hence, limit the deleterious effects of high
ions on enzyme function.
5.1. Fatty acid binding protein (FABP)
Vertebrate estivators derive most of their energy
needs from the aerobic oxidation of lipid reserves
and so adaptations of proteins involved in lipid
catabolism could be important for long term survival. Fatty acid binding proteins (FABPs) are 14
kDa cytoplasmic proteins that mediate the intracellular trafficking of fatty acids between three
points: plasma membrane; intracellular triglyceride
droplets; and mitochondria. In a recent study we
purified and analyzed the FABP from skeletal
muscle of S. couchii (Stewart et al., 2000). Little
is known about amphibian FABPs; only two other
studies have reported on the amphibian liver isoform (Schleicher and Santome, 1996; Baba et al.,
K.B. Storey / Comparative Biochemistry and Physiology Part A 133 (2002) 733–754
1999). Like FABPs from most other sources (Paulussen and Veerkamp, 1991), spadefoot toad muscle FABP had a molecular weight of ;14 kDa, a
blocked N-terminus (due to N-acetylation) and
bound fatty acids with a 1:1 stoichiometry. The
Kd values for oleate and the fluorescent probe, cisparinarate, were 1.04"0.08 and 1.15"0.16 mM,
respectively, within the typical range for FABPs
from other sources. Compared with FABP from
other species, no outstanding differences were
noted for the toad protein that might specifically
serve estivation.
Effects of urea on the binding of cis-parinarate
were also assessed. Urea at 200 mM increased the
binding of the fluorescent probe by ;60%, which
suggests that fatty acid binding and transport by
FABP would not be impeded as cytoplasmic urea
levels rose in estivating toads (Stewart et al.,
2000). High levels of urea (1 M) also had no
negative effect on the protein. By contrast, the
addition of 200 mM KCl showed a trend (not
significant) to reduce probe binding to FABP.
Again, as suggested previously, this suggests that
high urea itself is not detrimental to estivator
proteins, and that one of the functions of urea
accumulation is to mitigate the negative effects on
proteins of the very high ion concentrations that
would otherwise develop in the cytoplasm as
organs dehydrated.
6. Estivation and gene expression
Modern techniques in molecular biology are
having a huge impact in many areas of biology
and in the field of biochemical adaptation are
finally providing the means to do thorough surveys
that identify all of the key genesyproteins that are
involved in stress and adaptation to environmental
insults. Techniques such as cDNA library screening
and differential display PCR have recently identified a wide range of genes and their protein
products that are responsive to stresses including
anoxiayhypoxia, osmotic stress, chilling and freezing in a variety of species and that support a range
of hypometabolic states including anaerobiosis,
hibernation, and diapause (Andrews et al., 1998;
Gracey, 1999; Cannon et al., 1999; Storey, 1999,
2001; Stears and Gullans, 2000; Denlinger, 2001).
Furthermore, the recent advent of cDNA microchipsyarrays with their opportunity to evaluate
responses to stress by hundreds or even thousands
of genes simultaneously, is causing an explosion
749
of information in mammalian fields that is now
beginning to filter into comparative biochemistry.
For example, cDNA array screening has recently
been used in our lab to assess gene responses to
hibernation in ground squirrels. Using a rat gene
array we found good cross-reaction between species for most of the over 500 genes assessed and,
more importantly, were able to compare relative
mRNA transcript levels for genes involved in a
wide range of metabolic pathwaysyfunctions in
euthermic vs. hibernating states (Eddy and Storey,
in press). Arrays based on the Drosophila or
Xenopus genomes will soon bring this technology
to many other groups of animals.
Relatively little is known to date about the gene
and protein responses that underlie estivation. Various studies have focused on specific enzymatic
systems. For example, it has been well known for
many years that one of the protein synthesis
responses to water or salt stresses in anuran
amphibians, including estivating species, is an
increase in the activities of the enzymes involved
in urea biosynthesis (Balinsky, 1981). Various of
the estivation-linked changes in the activities of
enzymes involved in intermediary energy metabolism or antioxidant defenses in toads (discussed
above) must also be based on estivation (or arousal)-stimulated changes in gene expression and
protein synthesis. Furthermore, evidence for estivation-specific proteins has been found in land
snails, O. lactea (Brooks and Storey, 1995). Snails
injected with 35S-methionine, then allowed to enter
estivation and sampled 2 days later showed strong
synthesis of a 50 kDa protein in hepatopancreas
that was not labeled in control snails, whereas
labeled proteins of 70 and 30 kDa appeared in
foot muscle after 2 days of estivation. By contrast,
snails that were first estivated for 14 days, then
injected with 35S-methionine and sampled 1 day
later while still estivating showed labeling of a 91
kDa protein in hepatopancreas. These results not
only document new protein synthesis during estivation but show tissue- and time-specific differences in the proteins produced. Furthermore, these
results for estivation can be contrasted with comparable studies that compared 35S-labeling patterns
in control vs. anoxic-exposed O. lactea; these
found no change in protein labeling patterns under
anoxia (Brooks and Storey, 1994b). However,
whereas protein-labeling studies do show that new
protein types are synthesized during estivation,
750
K.B. Storey / Comparative Biochemistry and Physiology Part A 133 (2002) 733–754
their downfall is that protein identity remains
elusive.
A recent study used cDNA library screening to
provide the first identification of a gene that is upregulated during estivation in liver of spadefoot
toads. Screening of a cDNA library made from
liver of 2 month estivated toads with probes made
from liver of active (control) toads vs. 2 month
estivated toads revealed one strong positive result
(Storey et al., 1999). The insert of the clone
contained a complete open reading frame coding
for a protein of 235 amino acids that was identified
as riboflavin binding protein (RfBP). RfBP is a
phosphoglycoprotein monomer that is synthesized
by liver of female vertebrates and secreted into
the plasma, where it binds plasma riboflavin and
loads the vitamin into the yolk of avian or reptilian
eggs or the fetus of mammals (White and Merrill,
1988). Our study was the first confirmation that
RfBP also occurs in amphibians. The putative
amino acid sequence of toad RfBP showed 50%
identity with the sequences of the chicken or turtle
proteins and essential structural features found in
chicken RfBP were conserved. These included 18
cysteine residues forming nine disulfide bridges,
two asparagine glycosylation sites and six tryptophan residues that appear to act in ligand binding
(Blankenhorn, 1978; Monaco, 1997). However,
toad RfBP differed in one key way from the
protein of higher vertebrates and that was the lack
of a highly phosphorylated region near the Cterminus that in chicken RfBP contains eight
phosphoserines interspersed with glutamate residues. This forms the recognition site for the carrier
that facilitates transport of the vitamin-loaded protein into the yolk (Sooryanarayana et al., 1998).
Toad RfBP showed only three serine residues in
this region, spaced by two glutamate residues.
Hence, recognition and binding of RfBP to
amphibian oocytes could be quite different than it
is in birds and reptiles.
The reasons for RfBP up-regulation during estivation are still unclear and have not been experimentally tested. However, we can suggest two
possibilities. One is that RfBP production may be
related to the maturation of eggs in females so
that females are prepared for the explosive breeding that occurs within hours after emergence. A
second possibility is that RfBP might function in
the adult estivator to ‘cache’ riboflavin within the
body over the 9–10 months of estivation when the
toads neither eat, defecate or urinate. Hence, any
riboflavin released as the result of catabolic processes in cells during estivation, including the
significant wasting of skeletal muscle mass that
occurs (S. couchii loses ;17% of its body protein
reserves during estivation) (Jones, 1980) can be
cached. This could be an important conservation
strategy for key nutrients in these desert animals.
In this regard, the results of the northern blots that
analyzed the organ distribution of RfBP mRNA
transcripts were very interesting. RfBP was strongly up-regulated in liver during estivation, but no
transcripts were detected in brain, gut, heart or
kidney. Surprisingly, however, RfBP transcripts
were found in skeletal muscle of estivating toads,
but not in controls, a finding that could be consistent with the above idea of vitamin caching in
tissues that experience significant wasting during
estivation.
7. Perspectives
Studies of the biochemical and molecular mechanisms underlying estivation are well advanced on
some fronts. For example, we have a good understanding of fuel metabolism, urea production, antioxidant defenses, glycolytic regulation and the role
of reversible protein phosphorylation as a major
control mechanism in metabolic arrest. We also
have strong data that link the participation of
multiple types of protein kinases to the molecular
events involved in estivation, although as yet, the
targets of action for most of these kinases have
not yet been elucidated. Much of the data on
estivation parallels information that has come out
of other systems of animal metabolic arrest (e.g.
hibernation, anoxia tolerance) and together with
those data, support the idea that there are unifying
molecular mechanisms of metabolic arrest that are
broadly expressed across phylogenetic lines. For
example, the use of reversible protein phosphorylation for the coordinated suppression of multiple
metabolic processes (e.g. fuel metabolism, ion
pump and channel arrest, protein synthesis) in
hypometabolic states is now well-established in
systems of anoxia tolerance and hibernation. The
parallels between phosphorylation control of glycolytic enzymes in estivating land snails and in
anoxic marine molluscs confirm the importance of
this mechanism in the regulation of fuel catabolism
in estivators, and imply that other aspects of
metabolic suppression and reorganization during
estivation will likely also prove to be regulated
K.B. Storey / Comparative Biochemistry and Physiology Part A 133 (2002) 733–754
via reversible phosphorylation. For example, the
suppression of protein synthesis in hibernating
mammals and anoxic marine snails that has recently been linked to phosphorylation of specific ribosomal proteins, the eukaryotic initiation factor-2
(eIF-2) and eukaryotic elongation factor-2 (eEF2) (Frerichs et al., 1998; Chen et al., 2001; Larade
and Storey, 2002). For eEF-2 in hibernators, this
result was achieved by an ;50% higher activity
of eEF-2 kinase in tissues from hibernating animals
and a corresponding 20–30% decrease in the
activity of protein phosphatase 2A (which opposes
eEF-2 kinase), as a result of a 50–60% increase
in the levels of the specific inhibitor of PP2A,
IPP2A
. It could be predicted that parallel studies of
2
the controls on protein synthesis in estivators will
produce similar results, and such studies will be
important for confirming the importance of the
suppression of protein biosynthesis as an integral
part of metabolic rate depression.
Considerable research remains to be done on
other aspects of metabolic regulation in estivation.
Thus, whereas it is obvious that protein kinases
and phosphatases have a central regulatory role to
play in the phenomenon, there is virtually no
information to date on the upstream signals (extraor intracellular) that stimulate these signal transduction cascades and no more than a few of the
downstream targets of kinaseyphosphatase action
are known. The possible involvement of hormones
or other extracellular signaling molecules remains
to be investigated. For example, there is growing
evidence that mammalian hibernation may involve
a ‘hibernation induction trigger’ that is opioid in
nature (Horton et al., 1998), and the possibility of
a similar blood-borne hormonal factor in vertebrate
estivators should to be explored.
Much more research also needs to be done on
the role of gene expression in estivation. It is
probable that estivation involves selected changes
in the expression of a variety of proteins as is also
proving to be the case in hibernation and anoxia
tolerance (for reviews see Storey, 1999, 2001;
Larade and Storey, in press). Our initial analysis
of differential gene expression in estivating spadefoot toads made a comparison between aroused
and 2 month estivated animals (Storey et al.,
1999), but it can be hypothesized that that major
changes in gene expression are much more likely
to be associated with the early stages of entry into
estivation (andyor during the arousal from estivation). Hence, such studies should be redone to
751
monitor changes in gene expression over the transition period while animals are sinking into the
hypometabolic state. The advent of cDNA array
screening will provide a major aid to this search.
We have recently used human 19K cDNA arrays
to screen for anoxia-induced changes in gene
expression in the hepatopancreas of the marine
snail Littorina littorea (Larade and Storey, in
press), and freeze-induced changes in liver of the
freeze-tolerant wood frog, Rana sylvatica (unpublished). Although heterologous probing is generally plagued by low cross-reactivity (e.g. L. littorea
cDNA cross-reacted with only 18.35% of the
human cDNA sequences), the potential for gene
discovery is still enormous (18.35% of 19 000
genes means the responses of 3500 genes were
still assessed). Of these cross-reacting genes,
almost 11% (or 385 genes) showed putative upregulation by two-fold or more during anoxia.
These included selected protein phosphatases and
kinases, MAP kinase-interacting factors, translation factors, antioxidant enzymes, and nuclear
receptors (Larade and Storey, unpublished data).
Although many of these candidate genes may not
hold up to scrutiny by techniques such as Northern
blotting that are needed to confirm their upregulation, we nonetheless have gained a much
broader view of the potential gene expression
responses that may play significant roles in anoxia
survival, including genes representing areas of cell
function that have never before been considered
as having a role in estivation or metabolic rate
depression. Arrays are already available for the
Drosophila and C. elegans genomes and these may
represent effective alternatives for analyzing estivation-induced gene expression in land snails, and
arrays currently being developed for the Xenopus
genome will be effective for studies of estivating
anurans. Overall then, there are still many exciting
avenues of research to pursue before we will gain
a full understanding of the molecular regulation of
estivation.
Acknowledgments
Thanks to R. Whitwam, J. Duncan and T.
Churchill for their contributions to previously
unpublished data that are presented (O. lactea
FBPase, PFK-2, and amino acids) and to other
recent members of my lab including S.P.J. Brooks,
M. Hermes-Lima, J. Grundy, K. Cowan, J. McDonald, and E. Dent for their studies of estivation
752
K.B. Storey / Comparative Biochemistry and Physiology Part A 133 (2002) 733–754
that are reviewed here. Thanks to J.M. Storey for
help in the preparation of the manuscript.
References
Abe, A.S., 1995. Estivation in South American amphibians
and reptiles. Braz. J. Med. Biol. Res. 28, 1241–1247.
Adelman, R., Saul, R.L., Ames, B.N., 1988. Oxidative damage
to DNA: relation to species, metabolic rate and life span.
Proc. Natl. Acad. Sci. USA 85, 2706–2708.
Andrews, M.T., Squire, T.L., Bowen, C.M., Rollins, M.B.,
1998. Low-temperature carbon utilization is regulated by
novel gene activity in the heart of a hibernating mammal.
Proc. Natl. Acad. Sci. USA 95, 8392–8397.
Baba, K., Abe, T.K., Tsunasawa, S., Odani, S., 1999. Characterization and primary structure of fatty acid-binding protein
and its isoforms from the liver of the amphibian, Rana
catesbeiana. J. Biochem. (Tokyo) 125, 115–122.
Balinsky, J.B., 1981. Adaptation of nitrogen metabolism to
hyperosmotic environment in Amphibia. J. Exp. Zool. 215,
335–350.
Barnhart, M.C., 1983. Gas permeability of the epiphragm of a
terrestrial snail. Physiol. Zool. 56, 436–444.
Barnhart, M.C., McMahon, B.R., 1987. Discontinuous carbon
dioxide release and metabolic depression in dormant land
snails. J. Exp. Biol. 128, 123–138.
Bemis, W.E., Burggren, W.W., Kemp, N.E., 1987. In: Bemis,
W.E., Burggren, W.W., Kemp, N.E. (Eds.), The Biology and
Evolution of Lungfish. Alan R. Liss, New York.
Bishop, T., Brand, M.D., 2000. Processes contributing to
metabolic depression in hepatopancreas cells from the snail
Helix aspersa. J. Exp. Biol. 203, 3603–3612.
Bishop, T., St-Pierre, J., Brand, M.D., 2002. Primary causes
of decreased mitochondrial oxygen consumption during
metabolic depression in snail cells. Am. J. Physiol. 282,
R372–R382.
Blankenhorn, G., 1978. Riboflavin binding in egg white
flavoprotein: the role of tryptophan and tyrosine. Eur. J.
Biochem. 82, 155–160.
Brooks, S.P.J., Storey, K.B., 1992. Properties of pyruvate
dehydrogenase from Otala lactea: control of enzyme activity
during estivation. Physiol. Zool. 65, 620–633.
Brooks, S.P.J., Storey, K.B., 1994. Metabolic depression in
land snails: in vitro analysis of protein kinase involvement
in pyruvate kinase control in isolated Otala lactea tissues.
J. Exp. Zool. 269, 507–514.
Brooks, S.P.J., Storey, K.B., 1994. Patterns of protein synthesis
and phosphorylation during anoxia in the land snail Otala
lactea. Can. J. Zool. 72, 856–862.
Brooks, S.P.J., Storey, K.B., 1995. Evidence for aestivation
specific proteins in Otala lactea. Mol. Cell. Biochem. 143,
15–20.
Brooks, S.P.J., Storey, K.B., 1996. Protein kinase involvement
in land snail aestivation and anoxia-protein kinase A kinetic
properties and changes in second messenger compounds
during depressed metabolism. Mol. Cell. Biochem. 156,
153–161.
Brooks, S.P.J., Storey, K.B., 1997. Glycolytic controls in
estivation and anoxia: a comparison of metabolic arrest in
land and marine molluscs. Comp. Biochem. Physiol. A 118,
1103–1114.
Cannon, B., Jacobsson, A., Nedergaard, J., 1999. Influence of
cold on gene expression in mammals. In: Storey, K.B. (Ed.),
Environmental Stress and Gene Regulation. BIOS Scientific
Publishers, Oxford, pp. 109–123.
Chen, W.S., Lazar, C.S., Poenie, M., Tsien, R.Y., Gill, G.N.,
Rosenfeld, M.G., 1987. Requirement for intrinsic protein
tyrosine kinase in the immediate and late actions of the
EGF receptor. Nature 328, 820–823.
Chen, Y., Matsushita, M., Nairn, A.C., et al., 2001. Mechanisms for increased levels of phosphorylation of elongation
factor-2 during hibernation in ground squirrels. Biochemistry
40, 11565–11570.
Chou, C.K., Dull, T.J., Russell, D.S., et al., 1987. Human
insulin receptors mutated at the ATP-binding site lack
protein tyrosine kinase activity and fail to mediate postreceptor effects of insulin. J. Biol. Chem. 262, 1842–1847.
Churchill, T.A., Storey, K.B., 1989. Intermediary energy
metabolism during dormancy and anoxia in the land snail
Otala lactea. Physiol. Zool. 62, 1015–1030.
Churchill, T.A., Storey, K.B., 1992. Natural freezing survival
by painted turtles Chrysemys picta marginata and C. p.
bellii. Am. J. Physiol. 262, 530–537.
Cowan, K.J., MacDonald, J.A., Storey, J.M., Storey, K.B.,
2000. Metabolic reorganization and signal transduction during estivation in the spadefoot toad. Exp. Biol. Online 5, 1.
Cowan, K.J., Storey, K.B., 1999. Reversible phosphorylation
control of skeletal muscle pyruvate kinase and phosphofructokinase during estivation in the spadefoot toad, Scaphiopus
couchii. Mol. Cell. Biochem. 195, 173–181.
Cowan, K.J., Storey, K.B., 2001. Tyrosine kinases and phosphatases in the estivating spadefoot toad. Cell Physiol.
Biochem. 11, 161–172.
Curtis, R.K., Bishop, T., Brand, M., in press. Control analysis
of metabolic depression. In: Storey, K.B. and Storey, J.M.
(Eds.), Cell and Molecular Responses to Stress, vol. 3,
Sensing, Signalling and Cell Adaptation. Elsevier Press,
Amsterdam.
Denlinger, D.L., 2001. Time for a rest: programmed diapause
in insect. In: Storey, K.B. (Ed.), Molecular Mechanisms of
Metabolic Arrest. BIOS Scientific Publishers, Oxford, pp.
155–167.
Elberg, G., Li, J., Leibovitch, A., Shechter, Y., 1995. Nonreceptor cytosolic protein tyrosine kinases from various rat
tissues. Biochim. Biophys. Acta 1269, 299–306.
Eddy, S.F. and Storey, K.B., in press. Dynamic use of cDNA
arrays: heterologous probing for gene discovery and exploration of organismal adaptation to environmental stress. In:
Storey, K.B. and Storey, J.M., (Eds.), Cell and Molecular
Responses to Stress, vol. 3. Elsevier Press, Amsterdam.
Etheridge, K., 1990. Water balance in estivating sirenid salamanders (Siren lacertina). Herpetologica 46, 400–406.
Fields, J.H., 1992. The effects of aestivation on the catalytic
and regulatory properties of pyruvate kinase from Helix
aspersa. Comp. Biochem. Physiol. B 102, 77–82.
Foe, L.G., Kemp, R.G., 1982. Properties of phospho and
dephospho forms of muscle phosphofructokinase. J. Biol.
Chem. 257, 6368–6372.
Frerichs, K.U., Smith, C.B., Brenner, M., et al., 1998. Suppression of protein synthesis in brain during hibernation
involves inhibition of protein initiation and elongation. Proc.
Natl. Acad. Sci. USA 95, 14511–14516.
K.B. Storey / Comparative Biochemistry and Physiology Part A 133 (2002) 733–754
Fuery, C.J., Attwood, P.V., Withers, P.C., Yancey, P.H., Baldwin, J., Guppy, M., 1997. Effects of urea on M4-lactate
dehydrogenase from elasmobranchs and urea-accumulating
Australian desert frogs. Comp. Biochem. Physiol. B 117,
143–150.
Fuery, C.J., Withers, P.C., Hobbs, A.A., Guppy, M., 1998. The
role of protein synthesis during metabolic depression in the
Australian desert frog, Neobatrachus centralis. Comp.
Biochem. Physiol. A 119, 469–476.
Gracey, A.Y., 1999. Environmentally induced gene expression:
poikilotherm responses to cold. In: Storey, K.B. (Ed.),
Environmental Stress and Gene Regulation. BIOS Scientific
Publishers, Oxford, pp. 81–94.
Grundy, J.E., Storey, K.B., 1994. Urea and salt effects on
enzymes from estivating and non-estivating amphibians.
Mol. Cell. Biochem. 131, 9–17.
Grundy, J.E., Storey, K.B., 1998. Antioxidant defenses and
lipid peroxidation damage in estivating toads, Scaphiopus
couchii. J. Comp. Physiol. B 168, 132–142.
Guppy, M., Fuery, C.J., Flanigan, J.E., 1994. Biochemical
principles of metabolic depression. Comp. Biochem. Physiol. B 109, 175–189.
Guppy, M., Reeves, D.C., Bishop, T., Withers, P., Buckingham,
J.A., Brand, M.D., 2000. Intrinsic metabolic depression in
cells isolated from the hepatopancreas of estivating snails.
FASEB J. 14, 999–1004.
Hermes-Lima, M., Storey, K.B., 1995. Antioxidant defenses
and metabolic depression in a pulmonate land snail. Am. J.
Physiol. 268, R1386–R1393.
Hermes-Lima, M., Storey, J.M., Storey, K.B., 1998. Antioxidant defenses and metabolic depression. The hypothesis of
preparation for oxidative stress in land snails. Comp. Biochem. Physiol. B 120, 437–448.
Hermes-Lima, M., Storey, J.M., Storey, K.B., 2001. Antioxidant defenses and animal adaptation to oxygen availability
during environmental stress. In: Storey, K.B., Storey, J.M.
(Eds.), Cell and Molecular Responses to Stress—Protein
Adaptations and Signal Transduction, Vol. 2. Elsevier Press,
Amsterdam, pp. 263–287.
Herreid, C.F., 1977. Metabolism of land snails (Otala lactea)
during dormancy, arousal and activity. Comp. Biochem.
Physiol. A 56, 211–215.
Hillman, S.S., 1980. Physiological correlates of differential
dehydration tolerance in anuran amphibians. Copeia 1980,
125–129.
Hillman, S.S., 1988. Dehydrational effects on brain and cerebrospinal fluid electrolytes in two amphibians. Physiol. Zool.
61, 254–259.
Hochachka, P.W., 1986. Defense strategies against hypoxia and
hypothermia. Science 231, 234–241.
Hochachka, P.W., Somero, G.N., 1984. Biochemical Adaptation. Princeton University Press, Princeton, NJ.
Horton, N.D., Kaftani, D.J., Bruce, D.S., et al., 1998. Isolation
and partial characterization of an opioid-like 88 kDa hibernation-related protein. Comp. Biochem. Physiol. B 119,
787–805.
Hue, L., Rider, M.H., 1987. Role of fructose-2,6-bisphosphate
in the control of glycolysis in mammalian tissues. Biochem.
J. 245, 313–324.
Hunter, T., Sefton, M.B., 1980. Transforming gene product of
Rous sarcoma virus phosphorylates tyrosine. Proc. Natl.
Acad. Sci. USA 77, 1311–1315.
753
Jones, R.M., 1980. Metabolic consequences of accelerated urea
synthesis during seasonal dormancy of spadefoot toads,
Scaphiopus couchii and Scaphiopus multiplicatus. J. Exp.
Zool. 212, 255–267.
Kennett, R., Christian, K., 1994. Metabolic depression in
estivating long-neck turtles (Chelodina rugosa). Physiol.
Zool. 67, 1087–1102.
Kikkawa, U., Kishimoto, A., Nishizuka, Y., 1989. The protein
kinase C family: heterogeneity and its implications. Annu.
Rev. Biochem. 58, 31–44.
Land, S.C., Bernier, N.J., 1995. Estivation: mechanisms and
control of metabolic suppression. In: Hochachka, P.W.,
Mommsen, T.P. (Eds.), Biochemistry and Molecular Biology
of Fishes, Vol. 5. Elsevier Science, Amsterdam, pp.
381–412.
Larade, K., Storey, K., 2002. Reversible suppression of protein
synthesis in concert with polysome disaggregation during
anoxia exposure in Littorina littorea. Mol. Cell. Biochem.
232, 121–127.
Larade, K. and Storey, K.B. 2002. A profile of the metabolic
responses to anoxia in marine invertebrates. In: Storey, K.B.
and Storey, J.M. (Eds.), Cell and Molecular Responses to
Stress, vol. 3, Sensing, Signalling and Cell Adaptation.
Elsevier Press, Amsterdam.
Livingstone, D.R., de Zwaan, A., 1983. Carbohydrate metabolism in gastropods. In: Wilbur, K.M. (Ed.), The Mollusca,
vol. 1. Academic Press, New York, pp. 177–242.
Loveridge, J.P., Withers, P.C., 1981. Metabolism and water
balance of active and cocooned African bullfrogs Pyxicephalus adspersus. Physiol. Zool. 54, 203–214.
MacDonald, J.A., Storey, K.B., 1999. Regulation of ground
squirrel Naq Kq-ATPase activity by reversible phosphorylation during hibernation. Biochem. Biophys. Res. Commun.
254, 424–429.
McClanahan, L., 1967. Adaptations of the spadefoot toad,
Scaphiopus couchii, to desert environments. Comp. Biochem. Physiol. 20, 73–79.
Michaelidis, B., Pardalidis, T., 1994. Regulation of pyruvate
kinase from the ventricle of the land snail Helix lucorum L.
during early and prolonged estivation and hibernation.
Comp. Biochem. Physiol. B 107, 585–591.
Monaco, H.L., 1997. Crystal structure of chicken riboflavinbinding protein. EMBO J. 16, 1475–1483.
Paulussen, R.J.A., Veerkamp, J.H., 1991. Intracellular fattyacid-binding proteins: characteristics and function. In: Hilderson, H.J. (Ed.), Subcellular Biochemisty: Intracellular
Transfer of Lipid Molecules, Vol. 16. Plenum Press, New
York, pp. 175–226.
Pedler, S., Fuery, C.J., Withers, P.C., Flanigan, J., Guppy, M.,
1996. Effectors of metabolic depression in an estivating
pulmonate snail (Helix aspersa): whole animal and in vitro
tissue studies. J. Comp. Physiol. B 166, 375–381.
Pinder, A.W., Storey, K.B., Ultsch, G.R., 1992. Estivation and
hibernation. In: Feder, M.E., Burggren, W.W. (Eds.), Environmental Biology of the Amphibia. University of Chicago
Press, Chicago, pp. 250–274.
Price, D.J., Tabarini, D., Li, H., 1986. Purification, subunit
composition and regulatory properties of type I phosphoprotein phosphatase from bovine heart. Eur. J. Biochem. 158,
635–645.
754
K.B. Storey / Comparative Biochemistry and Physiology Part A 133 (2002) 733–754
Rees, B.B., Hand, S.C., 1991. Regulation of glycolysis in the
land snail Oreohelix during estivation and artificial hypercapnia. J. Comp. Physiol. 161, 237–246.
Rees, B.B., Hand, S.C., 1993. Biochemical correlates of
estivation tolerance in the mountainsnail Oreohelix (Pulmonata: Oreohelicidae). Biol. Bull. 184, 230–242.
Robinson, N.C., 1993. Functional binding of cardiolipin to
cytochrome c oxidase. J. Bioenerg. Biomembr. 25, 153–163.
Rosen, N., Bolen, J.B., Schwartz, A.M., Cohen, P., DeSeau,
V., Israel, M.A., 1986. Analysis of pp60c-src protein kinase
activity in human tumor cell lines and tissues. J. Biol.
Chem. 261, 13754–13759.
Sadoshima, J., Qiu, Z., Morgan, J.P., Izumo, S., 1996. Tyrosine
kinase activation is an immediate and essential step in
hypotonic swelling-induced ERK activation and c-fos gene
expression in cardiac myocytes. EMBO J. 15, 535–546.
Schleicher, C.H., Santome, J.A., 1996. Purification, characterization, and partial amino acid sequencing of an amphibian
liver fatty acid binding protein. Biochem. Cell Biol. 74,
109–115.
Shoemaker, V.H., 1992. Exchange of water, ions and respiratory gases in terrestrial amphibians. In: Feder, M.E., Burggren, W.W. (Eds.), Environmental Biology of the Amphibia.
University of Chicago Press, Chicago, pp. 125–150.
Sooryanarayana, S.S., Adiga, P.R., Visweswariah, S.S., 1998.
Identification and characterization of receptors for riboflavin
carrier protein in the chicken oocyte. Role of the phosphopeptide in mediating receptor interaction. Biochim. Biophys.
Acta 1382, 230–242.
Srivastava, A.K., 1985. Stimulation of tyrosine protein kinase
activity by dimethyl sulfoxide. Biochem. Biophys. Res.
Commun. 126, 1042–1047.
Stears, R.L., Gullans, S.R., 2000. Transcriptional response to
hyperosmotic stress. In: Storey, K.B., Storey, J.M. (Eds.),
Cell and Molecular Responses to Stress—Environmental
Stressors and Gene Responses, Vol. 1. Elsevier, Amsterdam,
pp. 129–139.
Stewart, J.M., Claude, J.F., MacDonald, J.A., Storey, K.B.,
2000. The muscle fatty acid binding protein of spadefoot
toad (Scaphiopus couchii). Comp. Biochem. Physiol. B 125,
347–357.
Storey, K.B., 1988. Mechanisms of glycolytic control during
facultative anaerobiosis in a marine mollusc: tissue-specific
analysis of glycogen phosphorylase and fructose-2,6-bisphosphate. Can. J. Zool. 66, 1767–1771.
Storey, K.B., 1996. Metabolic adaptations supporting anoxia
tolerance in reptiles: recent advances. Comp. Biochem.
Physiol. B 113, 23–35.
Storey, K.B., 1997. Metabolic regulation in mammalian hibernation: enzyme and protein adaptations. Comp. Biochem.
Physiol. A 118, 1115–1124.
Storey, K.B., 1998. Survival under stress: molecular mechanisms of metabolic rate depression in animals. S. Afr. J.
Zool. 33, 55–64.
Storey, K.B., 1999. Stress-induced gene expression in freezetolerant and anoxia-tolerant vertebrates. In: Storey, K.B.
(Ed.), Environmental Stress and Gene Regulation. BIOS
Scientific Publishers, Oxford, pp. 1–23.
Storey, K.B., 2001. Turning down the fires of life: metabolic
regulation of hibernation and estivation. In: Storey, K.B.
(Ed.), Molecular Mechanisms of Metabolic Arrest. BIOS
Scientific Publishers, Oxford, pp. 1–23.
Storey, K.B., Storey, J.M., 1990. Facultative metabolic rate
depression: molecular regulation and biochemical adaptation
in anaerobiosis, hibernation, and estivation. Q. Rev. Biol.
65, 145–174.
Storey, K.B., Storey, J.M., 1996. Natural freezing survival in
animals. Ann. Rev. Ecol. Syst. 27, 365–386.
Storey, K.B., Dent, M.E., Storey, J.M., 1999. Gene expression
during estivation in spadefoot toads, Scaphiopus couchii:
up-regulation of riboflavin binding protein in liver. J. Exp.
Zool. 284, 325–333.
Stuart, J.A., Gillis, T.E., Ballantyne, J.S., 1998. Compositional
correlates of metabolic depression in the mitochondrial
membranes of estivating snails. Am. J. Physiol. 275,
R1977–1982.
Tocque, K., Tinsley, R., Lamb, T., 1995. Ecological constraints
on feeding and growth of Scaphiopus couchii. Herpetol. J.
5, 257–265.
Vazquez-Illanes, M.D., Storey, K.B., 1993. 6-Phosphofructo2-kinase and control of cryoprotectant synthesis in freeze
tolerant frogs. Biochim. Biophys. Acta 1158, 29–32.
White, H.B., Merrill, A.H., 1988. Riboflavin-binding proteins.
Ann. Rev. Nutr. 8, 279–299.
Whitwam, R.E., Storey, K.B., 1990. Pyruvate kinase from the
land snail Otala lactea: regulation by reversible phosphorylation during estivation and anoxia. J. Exp. Biol. 154,
321–337.
Whitwam, R.E., Storey, K.B., 1991. Regulation of phosphofructokinase during estivation and anoxia in the land snail
Otala lactea. Physiol. Zool. 64, 595–610.
Wilz, M., Heldmaier, G., 2000. Comparison of hibernation,
estivation and daily torpor in the edible dormouse, Glis glis.
J. Comp. Physiol. B 170, 511–521.
Withers, P.C., Guppy, M., 1996. Do Australian desert frogs
co-accumulate counteracting solutes with urea during aestivation? J. Exp. Biol. 199, 1809–1816.
Witte, O.N., Dasgupta, A., Baltimore, D., 1980. Abelson
murine leukaemia virus protein is phosphorylated in vitro
to form phosphotyrosine. Nature 283, 826–831.