ARTICLE
Auteur(s) : Hélène
Martin1, Béatrice Uring-Lambert2, Markus
Adrian3, Abdeslam Lahlou2, Alexandre
Bonet1, Céline Demougeot3, Sylvie
Devaux3, Pascal Laurant3, Lysiane
Richert1, Alain Berthelot3
1Laboratoire de toxicologie cellulaire, EA2SBP, UFR
des sciences médicales et pharmaceutiques, Besançon, France
2Laboratoire d’immunologie, Hôpitaux Universitaires,
Strasbourg, France
3Laboratoire de physiologie, EA2SBP, UFR des sciences
médicales et pharmaceutiques, Besançon, France
In most industrialized countries, hypomagnesaemia is frequent
among the general population [1, 2], since dietary behaviors have
changed with industrialized food. Indeed, food such as whole grains
and vegetables which are naturally magnesium (Mg)-rich, is largely
processed before being consumed, leading to an important loss of Mg
content. Consequently, a chronic dietary inadequacy of Mg is
commonplace. One other reason for the prevalence of hypomagnesaemia
is due to disturbances in the intestinal Mg absorption and/or to
increased renal Mg excretion. Hypomagnesaemia is well recognized as
an important human health problem, since Mg is involved in many
enzymatic reactions and hypomagnesaemia plays an important role in
the pathogenesis of numerous diseases, including ischemic heart
disease, sudden cardiac death, hypertension, stroke,
atherosclerosis and cancer (for review, see [3]).
Several in vivo and in vitro studies dealing with the effects of
Mg intake, have concluded that oxidative stress was involved in the
response to Mg deficiency in different tissues and organs [4-11].
This was evidenced either by an impairment of the defenses against
oxidative stress or by an accumulation of oxidation products which
was frequently associated with an increase in lipid peroxidation.
In our previous studies, we showed that extracellular Mg deficiency
has a negative effect on the survival of cultured rat and human
hepatocytes by inducing apoptosis involving oxidative stress;
however, supplementation of extracellular Mg did not reduce
spontaneous apoptosis occurring over time in hepatocyte cultures
[12, 13]. Other studies have evidenced that apoptosis was induced
in different rat tissues, following Mg deficiency [8, 14, 15].
However, most of the publications aiming to study Mg effects
consisted of short-term or acute exposure, which is not
representative of the in vivo chronic human situation. Moreover,
such experiments do not allow evaluation of the possible impact of
Mg intake on the ageing and mortality rate. Nevertheless, Mg
deficiency could have an important role in the acceleration of
cellular senescence. Indeed, epidemiological studies have suggested
that Mg intake may be beneficial in the prevention of ageing, since
Mg deficiency is a significant risk factor of ageing [16, 17].
Recently, Ferrè et al. have shown that Mg deficiency induced
senescent features in cultured human endothelial cells, the cdk
inhibitor p21 being upregulated [18]. Moreover, Killilea and Ames
have evidenced that Mg deficiency accelerated cellular senescence
in cultured human fibroblasts, by an increased p16(INK4a) and
p21(WAF1) protein expression and an increased telomere attrition
[19].
In the present study, we evaluated the effect of long-term
dietary Mg intake on the rate of oxidative stress, apoptosis and
ageing in rat livers. To address this issue, rats were fed diets
containing either a moderately deficient (0.15 g Mg/kg diet), a
standard (0.8 g Mg/kg diet) or a high (3.2 g Mg/kg diet) Mg dose
for two years, which corresponds to the average life time of this
species.
Materials and methods
In-life Experiment
All animals were treated in compliance with applicable guidelines
formulated by the European Union for the care and use of laboratory
animals (L 358-86/609/EEC). Male Sprague-Dawley rats four weeks old
were obtained from Charles River (St Germain sur l’Arbresle,
France). They were randomly divided into three groups and housed in
standard animal laboratory cages (4 rats per cage, 5 cages per
group) with free access to distilled water and food. They were kept
under constant temperature (23°C), constant humidity (50-60%), and
a daily 12h light-dark cycle. The rats were pair-fed a 0.15 g, 0.8
g or 3.2 g Mg/kg diet for 90 weeks. The synthetic diets contained
the following (%): casein 20, starch 40, sucrose 21, cellulose 6,
groundnut oil 2.5, corn oil 2.5, mineral mixture 7, and vitamin
mixture 1. Mg was given in the form of magnesium monooxide and Mg
concentrations of the diets were verified by atomic absorption
spectrophotometry (Perkin Elmer 3300, Saint Quentin en Yvelines,
France). At the end of the treatment period, the terminal body
weights were recorded. The animals were anesthetized with sodium
pentobarbital (40 mg/kg b.w.) and exsanguinated by abdominal artery
puncture. Blood samples were collected in heparinized tubes for
subsequent plasma Mg measurement by atomic absorption
spectrophotometry (Perkin Elmer 3300). The livers were rapidly
removed and weighed. Small parts of liver of approximately 1g were
washed in cold phosphate-buffer and immediately flash-frozen in
liquid nitrogen and kept frozen until used for analysis.
Oxidative stress measurements
Liver homogenates were prepared as previously described [20, 21].
All the procedures to quantify the different enzyme activities and
the reduced glutathione (GSH) content have been previously
described by Binda et al. [20] and Nicod et al. [21]. The enzyme
activities that we tested in the present study were the following:
catalase (CAT), superoxide dismutase (SOD), glutathione peroxidase
(GPx) and glutathione reductase (GR) activities. CAT activity was
expressed as EU/mg protein, SOD, GPx and GR activities were
expressed as nmol/min/mg protein. GSH content was expressed as nmol
per mg protein. For thiobarbituric acid reactive substances (TBARS)
concentration determination, liver homogenates were precipitated
with trichloacetic acid and centrifugated at 1360g for 15 min.
The supernatants were mixed with TBA reagent (0.067%) and the
mixtures were kept at 100°C for 15 min. The fluorescent
reaction product was extracted with n-butanol and the fluorescence
was measured in the organic phase using a fluorescence
spectrophotometer (excitation: 535nm and emission: 555 nm). TBARS
concentrations were calculated relative to a standard preparation
of 1,1,3,3-tetra-ethoxypropane and expressed as nmol
malondialdehyde (MDA) par mg protein.
Apoptosis measurement
Approximately 100 mg of frozen rat livers were homogenised in the
lysis buffer provided in the kit (Interchim, Montluçon, France).
After centrifugation at 15,000 rpm for 15 minutes at 4 °C, the
supernatants were collected and used to quantify caspase-3 activity
according to the manufacturer’s instructions. Standard curves were
obtained using 7-amino-4-methylcoumarin (AMC). Caspase-3 activity
was determined using a fluorescence spectrophotometer (excitation
at 340 nm and emission at 450 nm) and expressed as pmol AMC/min/mg
protein.
Protein concentration determination
The protein concentration of liver homogenates was evaluated by the
bicinchoninic acid protein assay kit, according to the
manufacturer’s instructions (Sigma Chemicals, St Louis, MO, USA)
and BSA was used as a standard.
Telomere length measurement
Hepatocytes from frozen rat livers were isolated by mechanical
disruption using a 35 μm Medicon (DAKO, Glostrup, Denmark)
followed by a cell filtration on a 50 μm filter (DAKO).
Hepatocytes were then mixed with 1301 cell line (2.106
cells of each type). The 1301 cell line [22] was used as an
internal control because it is near-tetraploid and has very long
telomeres (> 25 kb), and can therefore be distinguished from the
cell types used in the assay [23]. After DNA denaturation,
hybridization with the fluorescein (FITC)-conjugated peptide
nucleic acid (PNA) probe (DAKO) was performed overnight. Negative
control allowing the detection of cell autofluorescence was also
prepared by omitting the PNA probe in the mixture. After washing,
cells were stained with propidium iodide (PI) to identify G0/G1
cells and calculate the DNA index of hepatocytes and 1301 cell
line. Determination of telomere length was performed by flow
cytometry [23, 24]. FC500 flow cytometer (Beckman Coulter, USA) was
used to quantify FITC- and PI-staining cells. The Relative Telomere
Length (RTL) expressed in % was calculated according to the
manufacturer’s instructions (DAKO).
Statistical analysis
Statistical comparisons among experimental animal groups were
performed by one-way analysis of variance, using Tukey’s test. The
level of statistical significance was set at 0.05.
Results and discussion
With the aim of evaluating a chronic human in vivo situation of Mg
deficiency and/or supplementation, the present experiments were
designed to study the effects of either a Mg deficiency or a Mg
supplementation on the liver. Rats were therefore fed different
diets over their entire lifespan, i.e. approximately 2 years for
this species, with Mg dose levels that led respectively to diets
being Mg moderately deficient, standard and Mg supplemented.
As shown in table 1, the standard Mg
diet we used (0.8 g of Mg/kg diet) led to a plasma Mg concentration
of 0.7 ± 0.02 mmol/L, that defined the standard “Std” rat group.
The Mg moderately deficient diet (0.15 g of Mg/kg diet) we used,
which corresponds to “Def” group, resulting in a plasma Mg
concentration of 0.52 ± 0.03 mmol/L, has been preferred to a Mg
severe deficient diet for the maintenance of body weight gain of
animals during the 2 years of treatment. Moreover, this allowed us
to achieve low Mg plasma levels clinically relevant to
Mg-deficiency in humans. Finally, a Mg supplemented group (3.2 g of
Mg/kg diet), which corresponds to “Suppl” group, led to a plasma Mg
concentration of 0.86 ± 0.02 mmol/L. By using these experimental
conditions, we noted no differences in the body weights and liver
weights of the rats from the three groups at the end of the
treatment period (table 1). However, it
is noteworthy that a higher percentage of animal mortality was
observed in the lowest Mg diet, as compared to the other groups.
During the first 50 weeks of the experiment no death was noted in
the three animal groups, while at the end of experiments, the
percent of dead animals corresponded to 38%, 25% and 11% of the
initial animal number in Def, Std and Suppl groups, respectively
(figure 1). The
mortality trend was evaluated using a log-rank test and was found
significant (p = 0.011).
Oxidative stress was evaluated by the determination of several
enzyme activities: CAT, SOD, GPx and GR activities, as well as GSH
content. Figure
2 showed that Mg diets significantly impacted on GPx
activity, since a statistically significant lower GPx activity was
measured in the Def group. This result is in accordance with data
obtained after short-term exposures, showing a decrease in GPX
activity in rats kept on a Mg-deficient diet during 22 days or 8
weeks [25, 26]. In the present study, no other significant changes
between the three animal groups were noted on the other enzyme
activities tested (data not shown). Several studies reported the
impact of Mg deficiency on SOD and glutathione transferase gene
expression in rat thymocytes [8] and on CAT, SOD, GR and
glutathione S-transferase activities in red blood cells [26]. But
these studies were performed with acute exposure (i.e. no more than
22 days). Consequently, discrepancies between our results and other
published data could be explained by the duration of treatment
(short- vs long-term exposure). Moreover, we can hypothesize that
GPx activity revealed the most prominent alteration among
antioxidant enzymes, as earlier noted in studies dealing with the
effect of physical exercise [27, 28] and as already observed in our
laboratory. Finally, we can also suggest that a different pattern
could be obtained among various tissues, i.e. thymocytes or red
blood cells vs liver cells.
Lipid peroxidation, as assessed by TBARS concentration
measurement, was induced in rat liver by Mg deficiency, in
agreement with our in vitro data [12] and with several publications
[5, 6, 9], confirming that a chronic Mg deficiency induced
oxidative stress and subsequent production of oxidative compounds
in rat livers. However, no differences between the Mg-standard and
the Mg-supplemented groups were obtained. This is most probably not
related to the only marginal increase in plasma Mg concentration in
the latter group (0.86 ± 0.02 mmol/L plasma Mg) compared to animals
receiving a standard diet, since the same observation was made when
extracellular Mg was increased up to 2 mmol/L (as compared to
0.8 mmol/L for standard conditions) in primary cultures of rat
hepatocytes [12]. This suggests not only that rats under a chronic
Mg supplementation regimen regulate their blood Mg concentrations,
but also that hepatocytes regulate Mg uptake, therefore keeping
cell homeostasis constant.
Moreover, we observed a statistically significant activation of
caspase-3 in the Mg-deficient group, as compared to the Mg-standard
group. To our knowledge, this is the first demonstration that a
chronic Mg deficiency induced apoptosis in rat liver. This result,
in total agreement with our in vitro data on rat hepatocytes [12],
is also in accordance with other publications showing an induction
of caspase-3 in neutrophils [14] and in heart [15] of rat depleted
in Mg. The involvement of reactive oxygen species and related
secondary oxidant species such as lipid hydroperoxides are now well
documented in apoptosis [29-31]. Thus, the results of the present
in vivo study with livers from rats after a two-year chronic
Mg-deficient diet, confirm our previous in vitro observations in
rat hepatocytes cultured for 3 days in a Mg-deficient culture
medium [12], i.e. that extracellular Mg deficiency has a negative
effect on the survival of rat hepatocytes by inducing apoptosis,
probably as a result of an oxidative stress-related mechanism.
By using a long-term exposure that corresponds to the average
lifespan of rats, we were able to consider if Mg dietary intake and
subsequent plasma Mg concentration, can have a role in cellular
senescence. For this purpose, telomere shortening was measured as a
marker of ageing. Telomeres are unique DNA/protein structures that
contain noncoding TTAGGG repeats and telomere-associated proteins
[32]. They are present at the end of chromosomes as protective
chromosomal caps, since they protect against chromosomal end-to-end
fusions, which could lead to telomere dysfunction. In somatic
cells, telomere length is very heterogeneous but typically declines
with age, posing a barrier to tumor growth but also contributing to
loss of cells with age [33]. Telomeres have been reported to
shorten during ageing in various tissues [34]. To confirm these
data using rat liver cells, we completed our experiment with young
animals (four weeks old), which were fed only for one month with
the same standard diet rather than “old” rats. Relative Telomere
Length (RTL) was evaluated in hepatocytes by cytofluorimetric
analysis using a telomere specific FITC conjugated PNA probe. An
internal control of the analysis was added by using the 1301
leukemic cell line characterized by a stable telomere length [22].
RTL was expressed as a percentage of the length of the 1301 cells.
The mean RTL was 54.5 ± 1.1% and 57.5 ± 1.1% in “old” and “young”
animals, respectively. These data showed that telomere length was
decreased in old animals, fed a physiological Mg dose, as compared
to young animals, i.e. animals fed a physiological Mg dose for only
one month, confirming that telomere shortening well correlated with
ageing events. Moreover, in old animals, telomere length was
significantly reduced in Mg-deficient group, as compared to the
other animal groups. Indeed, the mean RTL was 46 ± 2.2, 54.5 ± 1.8
and 54.5 ± 1.1% in Def, Std and Suppl groups, respectively.
Accordingly, other authors reported that Mg deficiency induced
cellular senescence, in cultured human endothelial cells and
fibroblasts [18, 19]. This result is consistent with the higher
percentage of animal mortality that we observed with the
Mg-deficient group. Finally, this is in accordance with the
up-regulation of dyskeratosis congenita 1, dyskerin (dkc1)
expression we found in a recent work when Mg decreased in the diet
of animals [35], since it is known that DKC cells have a premature
telomere shortening, leading to accelerated ageing [36].
The present work should be linked to our recently published
observations on the impact of dietary Mg intake on the rat liver
transcriptome after a long-term exposure [35]. Among genes that
were differentially expressed following Mg deficiency, 32% of them
belonged to “the homeostasis family” that corresponded to genes
involved in oxidative stress, DNA damage, apoptosis and cellular
ageing. The present study therefore confirms on the protein level
the changes previously reported at the gene levels. More studies
are needed to assess underlying mechanisms involved in the
detrimental effect of Mg deficiency. However, recent studies
suggest that a systemic inflammation may play a role in the
pathophysiology of Mg deficiency. Indeed, Mg deficiency induced a
clinical inflammatory syndrome with excessive production of free
radicals (for review, see [37, 38]). Moreover, in humans, Mg
intakes below the recommended daily allowance are associated with
elevated C-reactive protein, a marker of inflammation, suggesting
that Mg deficiency may be involved in the development of the
low-grade chronic inflammatory syndrome [39, 40]. We demonstrate
for the first time that, while Mg supplementation had no beneficial
effect on the physiological decline in rat liver, a moderate
Mg-deficient diet was able to chronically accentuate cellular
ageing induced by oxidative stress and apoptosis processes. These
deleterious effects were associated with a higher mortality in
rats. It is noteworthy that these observations occurred with a
deliberately chosen moderate Mg deprivation, more relevant to
clinically low plasma levels.
Table 1 Effects of dietary magnesium on plasma Mg
concentration, body and liver weights in rat groups after 2 years
treatment.
|
Group
|
Mg diet (g Mg/kg diet)
|
Plasma Mg concentration (mmol/L)
|
Body weight (g)
|
Liver weight (g)
|
|
|
Mean
|
SEM
|
Mean
|
SEM
|
Mean
|
SEM
|
|
Def
|
0.15
|
0.52a
|
0.03
|
605.0a
|
23.9
|
15.6a
|
0.8
|
|
Std
|
0.8
|
0.70b
|
0.02
|
648.1a
|
20.4
|
16.0a
|
0.8
|
|
Suppl
|
3.2
|
0.86c
|
0.02
|
632.9a
|
16.9
|
16.1a
|
0.7
|
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