Hypometabolism, antioxidant defenses and free radical metabolism in the pulmonate land snail Helix aspersa
Oxyradical Research Group, Departamento de Biologia Celular, Universidade de Brasília, Brasília, DF, 70910-900, Brazil
* Author for correspondence (e-mail: hermes{at}unb.br)
Accepted 11 November 2002
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Summary |
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Key words: metabolic depression, free radical, glutathione, lipid peroxidation, carbonyl protein, Helix aspersa
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Introduction |
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Protein biosynthesis is a costly process, especially under hypometabolic
conditions. Thus, it would be expected that only enzymes relevant to the
maintenance of animal life would show increased activity (through
biosynthesis) during estivation. Hermes-Lima and Storey
(1995a) observed that, after
30 days of estivation, the activities of several antioxidant enzymes, mainly
catalase, selenium-dependent glutathione peroxidase (Se-GPX) and superoxide
dismutase (SOD), increase in the land snail Otala lactea. The
augmented endogenous antioxidant capacity during estivation was considered a
mechanism of preparation for the oxidative stress that accompanies arousal
(Hermes-Lima and Storey,
1995a
,b
;
Storey, 1996
;
Hermes-Lima et al., 1998
).
During arousal, there is a transitory increase in oxygen uptake, which may
create favorable conditions for an overgeneration of reactive oxygen species
(ROS). Indeed, in O. lactea, lipid peroxidation [as thiobarbituric
acid reactive substances (TBARS)] was significantly increased by 25% in
hepatopancreas during arousal (Hermes-Lima
and Storey 1995a
; Hermes-Lima
et al., 1998
). In the case of foot muscle no changes were observed
in TBARS during estivation and awakening
(Hermes-Lima and Storey,
1995a
).
These observations in land snails are analogous to the behavior of certain
antioxidant enzymes in garter snakes Thamnophis sirtalis parietalis,
leopard frogs Rana pipiens and goldfish Carassius auratus
during exposure to anoxia (Hermes-Lima and Storey,
1993a,
1996
;
Lushchak et al., 2001
).
Increased antioxidant enzyme activity during anoxia has been attributed to a
preparation against oxidative stress following reoxygenation
(Storey, 1996
; Hermes-Lima et
al., 1998
,
2001
;
Lushchak et al., 2001
;
Hermes-Lima and Zenteno-Savín,
2002
).
The aim of this study was to further characterize the changes occurring in several indicators of oxidative stress during estivation (20 days) and subsequent arousal of the land snail Helix aspersa. Lipid peroxidation (determined by two techniques), carbonyl proteins, glutathione (as GSH and GSSG) and the activities of glucose-6-phosphate dehydrogenase (G6PDH) and five antioxidant enzymes were quantified in H. aspersa hepatopancreas and foot muscle. The Se-GPX activity and GSH content were found to increase during estivation, and a complex process involving the formation and detoxification of lipid peroxidation products and protein oxidation products occurred during arousal.
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Materials and methods |
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Animals
Land snails Helix aspersa Muller 1774 were purchased from
Heliário Araras (Rio de Janeiro State, Brazil). The animals weighed
15-18 g and were kept in the laboratory at 25±1 °C in glass
containers with a 12:12 h light:dark cycle. The animals were sprayed with
dechlorinated water at 20 day intervals to induce arousal and were fed lettuce
sprinkled with ground chalk. No animals were used in experiments before
completing at least one cycle of 20 day estivation and arousal in the
laboratory. For sampling purposes, the snails were killed by breaking their
shells and the organs (foot muscle and hepatopancreas) quickly dissected out
and frozen in liquid nitrogen. Organ samples were stored at -75°C until
they were assayed. The deep-freeze storage period was no longer than 4
months.
Estivation/arousal experiments
Estivation was induced in the laboratory by removing water and food from
the containers. Within 1 day, the animals retracted inside their shells and
estivation was timed from that moment on. One group of snails was sampled
after 20 days of continuous dormancy. Another group was sprayed with water,
aroused and fed. The latter group was then also sampled after 24 h.
A temporal period of monitoring of arousal after 20 days of estivation was also carried out on another group of snails. After water and food were reintroduced, the length of arousal was timed from the moment the snails showed signs of activity (the foot emerging from the shell). This procedure was followed to account for the lack of perfect synchronism among individuals during arousal. Usually, 90% of the animals aroused within 5-10 min.
All the estivation experiments were conducted during June and July 1998, which corresponded to the dry winter season in Brasília, located in midwestern Brazil.
Preparation of extracts for enzyme assays
Tissue extracts were prepared using an Ultra-Turrax T8 (IKA Labottechnik;
Staufen, Germany) homogenizer. Samples of frozen tissue were quickly weighed
and then homogenized in ice-cold Buffer A (50 mmol l-1 potassium
phosphate buffer, pH 7.2, containing 0.5 mmol l-1 EDTA), in the
presence of 10 µmol l-1 phenylmethylsulfonyl fluoride (added
just before homogenization; stock solution was 1 mmol l-1, in
ethanol) at concentrations of 1:20 w/v for hepatopancreas and 1:15 w/v for
foot muscle. Samples were centrifuged in a Beckman centrifuge at 15 000
g for 15 min at 5°C. The supernatants (enzyme extracts)
were collected, stored on ice, and immediately used for enzyme assays at
25±1 °C. A preparatory enzyme extract desalting step by Sephadex
G-25 small-column filtration (Hermes-Lima
and Storey, 1993a; Willmore
and Storey, 1997a
) was omitted, since this procedure had no effect
on enzymatic activities, except for hepatopancreas SOD (75% loss of activity,
measured per mg of protein), hepatopancreas catalase and foot muscle SOD and
Se-GPX (25-50% loss of activity). This procedure was also omitted from the
determination of antioxidant enzyme activities of O. lactea
(Hermes-Lima and Storey,
1995a
).
Assays of antioxidant enzymes and G6PDH
The activity of catalase was quantified by the consumption of 10 mmol
l-1 H2O2 at 240 nm in Buffer A with 10 µl
of enzyme extract from hepatopancreas or 100 µl from foot muscle. Blanks
were run in the absence of H2O2
(Hermes-Lima and Storey,
1993a).
Total SOD activity (Mn- plus CuZn-SOD) was determined as previously
described (Hermes-Lima and Storey,
1995a) under the following assay conditions: 5 mmol l-1
EDTA, 2.5 mmol l-1 MnCl2, 0.25 mmol l-1 NADH,
4 mmol l-1 2-mercaptoethanol in 50 mmol l-1 potassium
phosphate buffer, pH 7.2. One SOD unit is defined as the amount of enzyme that
inhibits the superoxide-induced oxidation of NADH (monitored at 340 nm) by 50%
(IC50). Several 1 ml cuvettes were run for each sample, using
increasing amounts of enzyme extract (from 0 to 150 µl); these were plotted
as velocity versus amount of enzyme extract, and an IC50
value was obtained. Blanks were run in the absence of 2-mercaptoetanol.
Glutathione reductase (GR) activity was assayed by following the oxidation
of 0.25 mmol l-1 NADPH by 5 mmol l-1 GSSG in 1 ml of
Buffer A containing 75 µl of hepatopancreas enzyme extract or 150 µl of
foot muscle enzyme extract. Two blanks were run: one in the absence of GSSG
and another in the absence of enzyme extract
(Hermes-Lima and Storey,
1995a). The activity of Se-GPX (using H2O2
as the substrate that measures selenium-dependent GPX activity) was quantified
by a coupled-assay with GR-catalyzed oxidation of NADPH at 340 nm. First, the
basal consumption of 0.25 mmol l-1 NADPH was measured in 1 ml of
Buffer A containing 4 mmol l-1 azide, 5 mmol l-1 GSH,
1.5 i.u. ml-1 GR, and 50 µl of either hepatopancreas or foot
muscle enzyme extract. This background activity oxidized no more than 5-10% of
added NADPH. Next, 20 µl of H2O2 were added to a
final concentration of 0.2 mmol l-1. Blanks were run in the absence
of enzyme extract (Hermes-Lima and Storey,
1995a
).
The glutathione S-transferase (GST) activity was measured by following the
conjugation of 1 mmol l-1 GSH with 1 mmol l-1
1-chloro-2,4-dinitrobenzene (at 340 nm) in Buffer A containing 50 µl of
hepatopancreas or foot muscle enzyme extract. Two blanks were run: one in the
absence of GSH and the other in the absence of enzyme extract
(Hermes-Lima and Storey,
1993b).
G6PDH activity was determined as previously described by Lushchak et al.
(2001), using 100 µl of
enzyme extract from either hepatopancreas or foot muscle.
Glutathione measurements
Frozen tissue samples were homogenized (1:20 w/v) in ice-cold 5% w/v
sulfosalicylic acid (previously bubbled with nitrogen gas for 10 min), then
further bubbled with nitrogen gas for 10 s and centrifuged at 15 000
g in an Eppendorf microcentrifuge for 5 min. Supernatants were
removed and immediately used to measure total glutathione (GSH-eq=GSH+2 GSSG),
thus preventing any acid hydrolysis. GSH-eq was determined by following the
rate of reduction of DTNB by GSH at 412 nm and comparing this rate to a GSH
standard curve. The assay for GSH-eq was performed in 100 mmol l-1
potassium phosphate, pH 7.2, containing sample (20 µl for hepatopancreas
and 50 µl for foot muscle), 0.25 mmol l-1 NADPH and 0.6 mmol
l-1 DTNB. The absorbance at 412 nm was recorded up to
stabilization, after which GR was added (final concentration of 1 i.u.
ml-1) (Hermes-Lima and Storey,
1995a). The use of sulfosalicylic acid in sample preparation is
known to produce stable GSH-eq values over several hours
(Hermes-Lima and Storey,
1996
).
To quantify GSSG only, Griffith's method
(Griffith, 1980) was used,
with modifications (Hermes-Lima and
Storey, 1993a
). Briefly, 0.4 ml samples (extracts prepared in 5%
w/v sulfosalicylic acid) were mixed with 40 µl of 500 mmol l-1
2-vinylpyridine (prepared in ethanol), 0.4 ml of 500 mmol l-1
potassium phosphate buffer was then added and the pH adjusted to 7.0 with
NaOH. The GSH derivation was completed after 1 h incubation at room
temperature, after which GSSG alone was quantified as described for GSH-eq
determination. 30 µl samples were used for the measurements. A standard
curve of GSSG was done in the presence of ethanol to correct the latter's
inhibitory effect on the assay.
Assays for lipid peroxidation
Thiobarbituric acid reactive substances (TBARS) were quantified as an index
of lipid peroxidation (Hermes-Lima and
Storey, 1995a). Frozen samples were homogenized (1:20 w/v) in
ice-cold 1.1% phosphoric acid. Then, 0.4 ml of homogenate was mixed with 0.4
ml of 1% w/v thiobarbituric acid, 50 mmol l-1 NaOH, 0.1 mmol
l-1 butylated hydroxytoluene solution and 0.2 ml of 7% phosphoric
acid (all the solutions were kept on ice during manipulation to avoid side
reactions). Subsequently, samples (at approx. pH 1.5) were heated for 15 min
to 98°C and 1.5 ml of butanol then added. Finally, the tubes were
vigorously vortexed and centrifuged for 5 min in a benchtop centrifuge at 2000
g. The organic layers were removed and placed in glass
cuvettes. The thiobarbituric acid solution was replaced by 3 mmol
l-1 HCl for the blanks. Absorbances at 600 and 532 nm were
measured. The results were calculated so as to minimize background
interference: sample (A532A600) blank
(A532A600). Final TBARS values were expressed
using the extinction coefficient of 156 mmol l-1.
The spectrophotometric quantification of TBARS cannot be considered a
technique to determine malondialdehyde in tissues because the assay
overestimates the actual levels of malondialdehyde. However, it is considered
effective for comparative studies of oxidative stress since several other
thiobarbituric acid-reactive aldehydes are also products of lipid peroxidation
(Lapanna and Cuccurullo, 1993;
Hermes-Lima and Storey,
1995a
).
The xylenol orange assay for lipid hydroperoxides (FOX-reactive lipid
hydroperoxides) was performed as described by Hermes-Lima et al.
(1995). Frozen tissues were
homogenized at 1:20 w/v in high performance liquid chromatography (HPLC) grade
ice-cold methanol, centrifuged for 5 min in an Eppendorf microcentrifuge at 15
000 g and the supernatant retained. The assay mixture
contained 0.25 mmol l-1 FeSO4, 25 mmol l-1
sulfuric acid and 0.1 mmol l-1 xylenol orange to a final volume of
1 ml (the components were added in the order listed). This assay mixture was
incubated for 30 min. A supernatant sample (of hepatopancreas extract) was
then added and allowed to react for 5 h at room temperature before absorbance
was measured at 580 nm. A 15 µl sample of the supernatant was chosen
because it falls in the linear phase of the curve of supernatant volume
versus A580 (Ramos,
1999
). A 5 µl sample of 1 mmol l-1 cumene
hydroperoxide (5 µmol l-1 final concentration) was then added to
each cuvette and A580 remeasured after 30 min incubation. Blanks
were prepared by replacing tissue extracts with water. Lipid hydroperoxide
content was expressed in cumene hydroperoxide equivalents (CHE). This method
was not employed for foot muscle samples because linearity was not achieved in
the pre-tests of foot supernatant volume versus A580
(Ramos, 1999
).
Assay of carbonyl protein
Oxidative damage to proteins was quantified as carbonyl protein
(Stadtman and Levine, 2000).
Frozen samples were homogenized (1:20 w/v for both hepatopancreas and foot
muscle) in ice-cold 5% w/v sulfosalicylic acid and then centrifuged at 15 000
g in an Eppendorf microcentrifuge for 5 min. The supernatant
was removed and 0.5 ml of 2,4-dinitrophenyl-hydrazine (10 mmol l-1
in 2 mol l-1 HCl) solution was added to the pellet. The samples
were kept at room temperature for 1 h (the tubes were vigorously vortexed
every 10-15 min). Then, 0.5 ml of 20% w/v trichloracetic acid was added and
the tubes centrifuged for 3 min at 15 000 g. The supernatant
was again discarded and the excess 2,4-dinitrophenyl-hydrazine removed by
washing the pellet three times with 1 ml ethanol:ethyl acetate (1:1, v/v),
followed by vigorous vortexing and centrifuging for 3 min at 15 000
g. The pellet was dissolved in 6 mol l-1 guanidine
chloride and incubated for 15 min at 37°C. The maximum absorbance in the
range of 360-370 nm was recorded and the final carbonyl protein values
expressed using the extinction coefficient of 22 mmol l-1. Blanks
were prepared by replacing 2,4-dinitrophenyl-hydrazine with 2 mol
l-1 HCl. The samples were then read against the blanks.
Protein measurements and statistics
The protein concentration was measured by the classical Bradford method
with Coomassie Brilliant Blue G-250
(Bradford, 1976), using bovine
serum albumin as a standard. The values in all determinations were computed as
means ± S.E.M. A statistical analysis was performed by either unpaired
Student's t-test (indicated when used) or one-way analysis of
variance (ANOVA), followed by a one-tail Dunnett's test. The level of
statistical significance was taken as P<0.05.
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Results |
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Antioxidant enzymes
Catalase and SOD activities remained unaffected during 20 days estivation
in both hepatopancreas and foot muscle
(Table 1). Catalase activity in
hepatopancreas was 23-fold higher than in foot muscle of 24 h active H.
aspersa (P<0.01, t-test). However, SOD activity was
not significantly different when comparing the two organs.
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We also determined the time course of SOD and catalase activity of hepatopancreas during the arousal period: 20 days estivation (0 min), 15 min, 30 min, 2 h and 24 h later. No significant changes were detected in enzymatic activities (N=4, data not shown).
The activity of Se-GPX in 20-day estivating snails was significantly increased, by 391% and 290% in hepatopancreas and foot muscle, respectively, compared to 24 h aroused animals (Table 1). Se-GPX activity in hepatopancreas of 24 h active H. aspersa was not significantly different than in foot muscle of 24 h active animals.
No changes were observed in GR activity in hepatopancreas and foot muscle of H. aspersa during estivation (Table 1). GR activity in hepatopancreas of 24h active snails was 2.8-fold higher than in foot muscle (P<0.01, t-test).
The GST and G6PDH activities were also unchanged in hepatopancreas and foot muscle during estivation (Table 1). Moreover, the GST and G6PDH activities were also very similar when comparing hepatopancreas and foot muscle. This suggests that the two organs of H. aspersa have a similar capacity to deal with GST-catalyzed xenobiotic detoxification and to recycle NADPH, a substrate for GR.
Levels of GSH-eq and GSSG
The concentration of GSH-eq in hepatopancreas was significantly decreased
during the awakening process, diminishing from approximately 2900 nmol
g-1 wet mass during estivation to 1795 and 1585 nmol g-1
wet mass after 1 h and 24 h, respectively, of the awakening process
(Table 2).
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The levels of GSSG in hepatopancreas were statistically unaltered in the first moments of arousal (5-15 min; 300-350 nmol g-1 wet mass), followed by a progressive decrease to 180 nmol g-1 wet mass at 24 h (Table 2).
The concentrations of GSH-eq and GSSG in foot muscle of 24 h active snails were 1200 and 4.6 nmol g-1 wet mass, respectively. In contrast to hepatopancreas, no significant changes were observed for GSH-eq or GSSG in foot muscle during the estivationarousal cycle in H. aspersa (Table 2).
GSSG:GSH-eq ratio, lipid peroxidation and carbonyl protein
Arousal induced a significant increase (55%) in hepatopancreas GSSG:GSH-eq
ratio at 15 min compared to 20 day estivating snails (0 min arousal)
(Fig. 1). At 90-120 min, the
GSSG:GSH-eq ratio dropped to the same level observed during estivation. In the
case of foot muscle, an apparent increase in GSSG:GSH-eq ratio was observed at
15 min arousal, although this was non-significant.
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Lipid peroxidation measured as TBARS showed a complex time dependence in hepatopancreas during awakening (Fig. 2). The concentration of TBARS significantly decreased from 49 nmol g-1 wet mass at 20-day estivation (0 min arousal) to 30.7 nmol g-1 wet mass at 5 min arousal. At 30 min of awakening, TBARS rose significantly to 39.6 nmol g-1 wet mass, then gradually declined to 26.8 nmol g-1 wet mass at 24 h. Moreover, lipid hydroperoxides dropped to a very low level from 5.0 µmol CHE g-1 wet mass during estivation to 1.2 µmol CHE g-1 wet mass at 5 min arousal, remaining at this level for up to 24 h (Table 3).
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No significant changes were observed in TBARS during arousal in the case of foot muscle lipid peroxidation (Fig. 2). Lipid hydroperoxides were not measured in foot muscle of H. aspersa (see Materials and methods).
Protein oxidation, quantified as carbonyl protein
(Stadtman and Levine, 2000),
remained unchanged in hepatopancreas, but significantly reduced in foot muscle
during awakening (Table 3). Foot muscle carbonyl protein during estivation was 226 nmol g-1 wet
mass, falling significantly (by 26%) within 5 min of arousal and reaching 145
nmol g-1 wet mass at 24 h.
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Discussion |
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Although oxygen consumption was not measured in arousing H.
aspersa, a transient rise in oxygen uptake is a common phenomenon in
awakening estivators, including Pila ovata, O. lactea and Bulinus
nasutus snails (Coles,
1968; Herreid,
1977
). On resumption of normal breathing,
PO2 rises and stabilizes in tissues, while in
O. lactea oxygen consumption increases rapidly to a peak, reaching
levels at least twofold higher than control values and approximately sixfold
higher than consumption in the dormant state
(Hermes-Lima et al., 1998
;
Herreid, 1977
). Thus, with
this abrupt and rapid increase of oxygen consumption during arousal, internal
tissues would experience a transition from mild hypoxia (during estivation;
Barnhart, 1986
;
Pedler et al., 1996
) to
normoxia. Since it is known that the rate of production of
O2- and H2O2 at the mitochondrial
level in many biological systems is proportional to the oxygen tension
(Turrens et al., 1982
) and to
mitochondrial metabolic rate (Finkel and
Holbrook, 2000
), the rise in oxygen tension and consumption in
snail organs during arousal could result in a high production of ROS. Indeed,
Hermes-Lima and Storey (1995a
)
observed a transient increase in TBARS concentration and SOD activity in
hepatopancreas of arousing O. lactea. Moreover, certain antioxidant
enzymes were increased during estivation in O. lactea, possibly in
preparation for physiological oxidative stress during arousal. Increased
generation of ROS was also recently proposed for Arctic ground squirrels
Spermophilus parryii during arousal from hibernation
(Tøien et al., 2001
).
It has been proposed that ascorbate plays a relevant role in counteracting
oxidative stress in arousing squirrels.
Comparative and tissue-specific analysis of antioxidant enzymes
Hepatopancreas catalase activity was found to be similar in H.
aspersa (24h active), O. lactea (200i.u. mg-1
protein; Hermes-Lima et al.,
1998) and the mussel Mytilus edulis (260 i.u.
mg-1 protein; Livingstone et
al., 1992
), but it was one order of magnitude greater than in the
marine snail Litorina littorea (25 i.u. mg-1 protein;
Pannunzio and Storey, 1998
).
Moreover, hepatopancreas catalase activities of H. aspersa, O. lactea
and L. littorea were much greater than that found in foot muscle from
these species (1.5-8 i.u. mg-1 protein). Interestingly,
hepatopancreas catalase activity in H. aspersa was within the range
reported for the livers of goldfish, leopard frogs, wood frogs Rana
sylvatica, red-eared turtles Trachemys scripta elegans, garter
snakes and rats (70-550 i.u. mg-1 protein;
Pérez-Campo et al.,
1993
; Hermes-Lima et al.,
2001
), but tenfold lower than in desert spadefoot toads
Scaphiopus couchii (Grundy and
Storey, 1998
). SOD activity in both hepatopancreas and foot muscle
of 24h aroused H. aspersa was greater than that determined for O.
lactea (25 and 50 i.u. mg-1 protein for foot muscle and
hepatopancreas, respectively; Hermes-Lima
and Storey, 1995a
) and L. littorea (25-30 i.u.
mg-1 protein in hepatopancreas and foot muscle;
Pannunzio and Storey, 1998
).
Furthermore, hepatopancreas SOD activity in H. aspersa was higher
than that found in the liver of rats and several cold-blooded vertebrates
(10-80 i.u. mg-1 protein;
Pérez-Campo et al.,
1993
; Hermes-Lima et al.,
2001
). These observations indicate that SOD and catalase
activities in H. aspersa, albeit unchanged during the
estivationarousal cycle, have a high constitutive capacity for dealing
with O2- and H2O2.
Se-GPX activity was similar for 24h active H. aspersa and O.
lactea (4-14 mi.u. mg-1 protein in hepatopancreas and foot
muscle; Hermes-Lima et al.,
1998). However, hepatopancreas Se-GPX activity of H.
aspersa (24h active) was considerably lower than in the liver of rats and
several lower vertebrates (35-700 mi.u. mg-1 protein;
Hermes-Lima et al., 2001
).
During estivation, hepatopancreas Se-GPX activity of H. aspersa
reaches nearly 30 mi.u. mg-1 protein
(Table 1), indicating a
relevant capacity for detoxification of organic and inorganic peroxides,
especially important during the quick awakening period.
Hepatopancreas GR activity in H. aspersa was comparable to that
determined for O. lactea
(Hermes-Lima and Storey,
1995a) and L. littorea
(Pannunzio and Storey, 1998
),
and with the enzyme activity found in the livers of goldfish, leopard frogs,
wood frogs, spadefoot toads, red-eared turtles, garter snakes and rats (5-30
mi.u. mg-1 protein;
Pérez-Campo et al.,
1993
; Hermes-Lima et al.,
2001
). Foot muscle GR was also more active in H. aspersa
than in O. lactea (6 mi.u. mg-1 protein;
Hermes-Lima et al., 1998
) and
L. littorea (7 mi.u. mg-1 protein;
Pannunzio and Storey, 1998
).
Moreover, the GR activity of hepatopancreas and foot muscle was considerably
higher than that observed for Se-GPX activity of 24h active H.
aspersa in both organs. This is an interesting finding, since Se-GPX
normally displays a much higher activity than GR in most vertebrate species
(though not in O. lactea and L. littorea;
Hermes-Lima et al., 1998
;
Pannunzio and Storey, 1998
),
suggesting a major in situ capacity for GSH recycling in snails.
GST activity in hepatopancreas and foot muscle was comparable with that
observed in active O. lactea and L. littorea
(Hermes-Lima et al., 1998;
Pannunzio and Storey, 1998
),
and with the enzyme activity found in the liver of several cold-blooded
vertebrates and rats (400-800 mi.u. mg-1 protein;
Hermes-Lima et al., 2001
), but
lower than the liver enzyme activities of spadefoot toads and red-eared
turtles (1,500-2,000 mi.u. mg-1 protein;
Willmore and Storey, 1997a
;
Grundy and Storey, 1998
;
Hermes-Lima et al., 2001
).
G6PDH activity of H. aspersa was lower but within the same
order of magnitude than that observed in goldfish liver (250 mi.u.
mg-1 protein; Lushchak et al.,
2001
). These observations indicate that GR, GST and G6PDH are
functionally relevant enzymes in H. aspersa organs.
Comparative and tissue-specific analysis of glutathione and lipid
peroxidation
The drop in the GSH-eq concentration observed during the first hour of
arousal (Table 2) suggests that
GSH is consumed as a non-enzymatic antioxidant (which results in several GSH
oxidation products other than GSSG;
Halliwell and Gutteridge,
1999) or during GST-catalyzed conjugation with toxic metabolic
by-products. In addition, the 82% increase in the levels of hepatopancreas
GSH-eq in estivating snails when compared to 24h active animals also suggests
that enzymatic mechanisms of GSH synthesis are increased during
hypometabolism.
The concentration of hepatopancreas GSH-eq in H. aspersa was
comparable with that observed in O. lactea hepatopancreas (2,800 nmol
g-1 wet mass; Hermes-Lima et
al., 1998) and in the liver of goldfish, leopard frogs, wood
frogs, spadefoot toads, red-eared turtles and garter snakes (650-3500 nmol
g-1 wet mass; Hermes-Lima et
al., 2001
). GSH-eq in H. aspersa hepatopancreas was,
however, much higher than in the aquatic snails L. littorea and
Biomphalaria tenagophila (300-400 nmol g-1 wet mass,
Pannunzio and Storey, 1998
; S.
F. Arruda, M. V. R. Ferreira and M. Hermes-Lima, unpublished
observations).
The significantly higher levels of GSSG in hepatopancreas of estivating
snails in comparison with 24h active animals is possibly a reflection of the
increased GSH-eq concentration during estivation. The concentration of GSH
(GSH=GSH-eq 2 GSSG) in hepatopancreas of 24-h active snails was
estimated to be 1200 nmol g-1 wet mass, which is approximately 7
times the amount of GSSG. Comparatively, the GSH:GSSG ratio was 9 in
hepatopancreas of active O. lactea and 13, 15 and 17 in garter snake,
spadefoot toad and leopard frog liver, respectively (Hermes-Lima and Storey,
1993a,
1996
,
1995a
;
Grundy and Storey, 1998
).
The levels of foot muscle GSSG are extremely low compared to those found in
O. lactea (90 nmol g-1 wet mass;
Hermes-Lima and Storey, 1995a)
and L. littorea (30 nmol g-1 wet mass;
Pannunzio and Storey, 1998
),
suggesting that in vivo enzymatic oxidation of GSH to GSSG takes
place at very low rates during either estivation or the active state.
The levels of foot muscle TBARS were 34% and 50% lower than in
hepatopancreas of 24h active and estivating snails, respectively
(P<0.01, t-test). This is consistent with the lower
aerobic metabolic rates of foot muscle, which was also attested to by the very
low GSSG:GSH-eq ratio in this organ. Moreover, the levels of TBARS in H.
aspersa were comparable with those observed in O. lactea organs
(Hermes-Lima and Storey,
1995a). Lipid hydroperoxides (as CHE levels) in the hepatopancreas
of estivating snails were essentially the same as those determined in the
livers of golden-mantled ground squirrels Spermophilus lateralis and
red-eared turtles, but about 40% lower than in mouse liver
(Hermes-Lima et al., 1995
;
Willmore and Storey,
1997b
).
Antioxidant enzymes and GSH in estivating H. aspersa
The increase in foot muscle and hepatopancreas Se-GPX activity
(approximately four- and fivefold, respectively;
Table 1) and GSH-eq levels from
hepatopancreas (1.8-fold; Table
2) after 20 days of estivation indicate that H. aspersa's
antioxidant system responded to a cycle of estivationarousal. Moreover,
the activity of other antioxidant enzymes (catalase, SOD, GR and GST), as well
as G6PDH activity, was unchanged during estivation in both organs. These data
clearly show that H. aspersa either increases or preserves its
antioxidant defenses during metabolic depression.
Under hypometabolic conditions, it is imperative that only extremely
relevant biosynthetic ATP-consuming pathways remain active due to the high
energetic costs of protein biosynthesis (in carp hepatocytes, this accounts
for about 80% of energy demands; Pannevis
and Houlihan, 1992) (Hand and Hardening, 1996). This is the case
of the increase in Se-GPX activity (which may reflect the rise in enzyme
biosynthesis) in the estivating snails O. lactea
(Hermes-Lima and Storey,
1995a
) and H. aspersa
(Table 1). Such an increase in
Se-GPX activity would greatly improve the snails' ability to detoxify
H2O2 or organic peroxides, which could promote
(via Fenton-like reactions) lipid peroxidation and oxidative stress.
The increased activity of Se-GPX during estivation could be of key importance
in maintaining oxidative stress following arousal at controllable levels in
both hepatopancreas and foot muscle.
An increase in Se-GPX activity under metabolic depression was also observed
in goldfish brain and leopard frog heart after exposure to 8h (at 20°C)
and 30h (at 5°C) anoxia, respectively
(Hermes-Lima and Storey, 1996;
Lushchak et al., 2001
).
Moreover, Se-GPX activity increased on sub-zero freezing (a hypometabolic
condition that imposes ischemia on internal organs;
Storey, 1996
) in garter snakes
(Hermes-Lima and Storey,
1993a
) and wood frogs
(Joanisse and Storey, 1996
)
and under severe dehydration in leopard frogs (50% loss of body water after
92h, at 5°C, causing ischemia in internal organs;
Hermes-Lima and Storey, 1998
).
Moreover, Se-GPX activity from the hepatopancreas of the freshwater snail
B. tenagophila (control activity, 10 mi.u. mg-1 protein)
increased by 1.4-fold after 24h exposure to underwater anoxia at 27°C
(Ferreira and Hermes-Lima,
1997
; Hermes-Lima and
Zenteno-Savín, 2002
). These results, overall, suggest that
Se-GPX is a highly relevant antioxidant defense for the biochemical adaptation
against oxidative stress following hypometabolism and/or ischemia in
non-mammalian animals (see Hermes-Lima and
Zenteno-Savín, 2002
).
Glutathione is another endogenous antioxidant whose concentration increases
during estivation in H. aspersa. The increase in hepatopancreas
GSH-eq after 20 days estivation (mostly as GSH; see
Table 2) might be caused by
increased ATP-dependent biosynthesis and/or by decreased biotransformation of
GSH (discussed above). A rise in GSH-eq was also observed in hepatopancreas
and foot muscle of the marine snail L. littorea after 6 days of
underwater anoxia at 5°C (Pannunzio
and Storey, 1998) and in skeletal muscle of garter snakes after
10h anoxia at 5°C (Hermes-Lima and
Storey, 1993a
). Both the antioxidant properties of GSH itself
(against hydroxyl radicals and peroxynitrite;
Halliwell and Gutteridge,
1999
) and the effect that high GSH substrate levels can have on
the in situ activities of glutathione-utilizing enzymes, may be
important in dealing with oxidative stress conditions.
Oxidative stress: estivating versus arousing land
snails
Both the increased Se-GPX activity
(Table 1) and GSH-eq
concentration (Table 2) in
estivating H. aspersa could be of importance in minimizing oxidative
damage during arousal. One line of evidence for oxidative stress during
arousal was the transient increase in the hepatopancreas GSSG:GSH-eq ratio in
the first moments of awakening. Since the increase in GSSG:GSH-eq is
considered a relevant marker of oxidative stress and of the redox state of
cells (Schafer and Buettner,
2001), it is possible that overgeneration of
H2O2 occurs in hepatopancreas during awakening. The
metabolism of peroxides via Se-GPX, and possibly via the
peroxidase activity of GST (Prohaska,
1980
; Grundy and Storey,
1998
), measured in this work as part of the total-GST activity, is
assumed to increase GSSG production, which would explain the transient
increase in GSSG:GSH-eq ratio during arousal. It is also possible that the
decrease in hepatopancreas GSSG concentration from estivating/awakening to the
fully active state (24 h; see Table
2) is caused by the export of GSSG from the hepatopancreas
cells.
In the case of lipid peroxidation, we observed a complex behavior in TBARS
concentration during awakening (Fig.
2) and a rapid decrease in the levels of FOX-reactive lipid
hydroperoxides (CHE levels) within 5 min of arousal
(Table 3). The higher levels of
lipid peroxidation during estivation may indicate, at first glance, that
increased rates of ROS formation (relative to active animals) take place
during hypometabolism. However, reduced oxygen consumption in mitochondria
during estivation (a 50% reduction, measured in isolated hepatopancreas cells
of H. aspersa; Bishop and Brand,
2000) would ensure less production of oxygen free radicals.
Moreover, non-mitochondrial respiration is also suppressed by 64% during
estivation (Brand and Bishop, 2000). Such non-mitochondrial oxygen uptake
might be caused by the P450 system and by soluble oxidases, which can also be
a source of O2- and/or H2O2.
On the other hand, the intermittent oxygen uptake experienced by land
snails during estivation, which occurs every 20-50 h in O. lactea (a
condition wherein snails also hyperventilate;
Storey, 2002), might induce
quick bursts of ROS formation. Such a ROS formation might be higher than basal
estivation rates and could be a relevant source of ROS for oxidative damage to
lipids and proteins. Furthermore, it is also possible that the low metabolic
rates during estivation could decrease the rate of detoxification of
byproducts of lipid peroxidation, thus inducing their accumulation. This
situation would reverse when animals arouse and by-products of lipid
peroxidation products are metabolized. An increase in lipid peroxidation
products during estivation was observed in several organs of desert spadefoot
toads (burrowed in soil for 2 months at 21°C)
(Grundy and Storey, 1998
). In
this case, a concomitant reduction in endogenous enzymatic antioxidant
potential and increase in GSSG:GSH-eq ratio was also observed after 2 months
of estivation, indicating that spadefoot toads cope with oxidative stress
conditions during periods of hypometabolism
(Grundy and Storey, 1998
;
Hermes-Lima et al., 2001
).
The rise in TBARS concentration from 5 min to 30 min arousal (see
Fig. 2) suggests that ROS
overgeneration (in comparison with rates in active animals) also takes place
during recovery from estivation. This coincides with the awakening period,
when the GSSG:GSH-eq ratio is increased in hepatopancreas (see
Fig. 1). Thus, we propose that
arousal may induce two independent events in hepatopancreas: (i) an increase
in the detoxification rates of lipid peroxidation by-products, and (ii) an
increase in mitochondrial formation of ROS due to fast recovery of oxidative
metabolic rates. Interestingly, goldfish also experience a physiological
increase in lipid peroxidation (measured as conjugated dienes) in liver and
brain during post-anoxic reoxygenation
(Lushchak et al., 2001).
The accumulation of carbonyl protein in foot muscle during estivation
(Table 3) may be related to the
rates of protein oxidation by ROS (and by aldehydes formed from lipid
peroxidation; Stadtman and Levine,
2000) and proteasome-mediated recycling of oxidized proteins. It
is possible that the oxidized protein recycling mechanism in foot muscle is
diminished during the hypometabolic condition of estivation, causing
accumulation of carbonyl protein. Arousal may activate protein turnover
processes immediately to full rates, which could explain the quick decrease in
carbonyl protein levels in foot muscle within 5 min. The reason why carbonyl
protein increases during estivation in foot and not in hepatopancreas is yet
to be studied. It is possible that a more efficient oxidized protein recycling
mechanism is present in the hepatopancreas than in foot muscle.
Perspectives and conclusion
In conclusion, we observed that the land snail H. aspersa
increases its antioxidant capacity during estivation as a possible strategy to
minimize the effects of ROS generation following arousal. The transient
increase in TBARS concentration and the GSSG:GSH-eq ratio in hepatopancreas
strongly suggest that a physiological process of oxidative stress occurs
during arousal in H. aspersa, similar to that observed in the case of
awakening O. lactea (Hermes-Lima
and Storey, 1995a; Hermes-Lima
et al., 1998
) and post-anoxic goldfish
(Lushchak et al., 2001
).
Most studies reveal that ROS overgeneration and oxidative stress are
associated with the post-hypoxic/ischemic phase of the
hypoxia/ischemia-reperfusion process
(Storey, 1996;
Halliwell and Gutteridge,
1999
; Hermes-Lima et al.,
2001
). Be that as it may, the increased levels of lipid
hydroperoxides (and GSSG; see Hermes-Lima
et al., 1998
) in hepatopancreas during estivation could be a
triggering factor for the activation of signaling pathways leading to the
activation of GSH and Se-GPX biosynthesis and/or maintenance of other
antioxidant enzyme activities.
The increased levels of markers of oxidative damage during estivation
(TBARS, lipid hydroperoxides and carbonyl proteins) might be a consequence of
low levels of ROS formation associated with decreased rates of detoxification
of oxidative damage products. These moderate and tolerable levels of ROS
formation during estivation could also signal increased Se-GPX activity in
both hepatopancreas and foot muscle (there are examples in the literature
where low doses of H2O2 induce the activity of Se-GPX
and/or other antioxidant defenses; see
Halliwell and Gutteridge,
1999). Thus, snails would be protected against ROS overgeneration
during arousal. The idea of mild oxidative stress during hypometabolism as a
trigger mechanism to induce a preparation for arousal-induced stress was
recently proposed by Carey and co-authors
(2000
) in the case of
hibernating 13-lined ground squirrels Spermophilus
tridecemlineatus.
Furthermore, the mildly hypoxic environment of the internal organs of
estivating snails could also activate O2-sensing-related
transcriptional factors, such as hypoxia inducible factor 1 (HIF-1), which
have been associated with adaptive changes in mammalian cell metabolism (under
severe hypoxia), including the increased expression of proteins and enzymes
that respond to hypoxic stress (Wenger,
2000; Semenza,
2001
). No study, to date, links HIF-1 with the regulation of
antioxidant enzymes. In any case, it is tempting to propose this alternative
explanation, whereby HIF-1 would be linked to the upregulation of Se-GPX, and
interestingly, Se-GPX activation has been reported in HepG2 cells under
hypoxia (Ehleben et al.,
1997
).
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