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1 Departement de Physiologie, Universite de Lausanne, Lausanne, Switzerland
2
Centre Botnar de Recherche Clinique, Departement de medecine interne, Centre Hospitalier Universitaire Vaudois, Lausanne, Switzerland
3Department of Biochemistry, School of Medical Sciences, University of Bristol, Bristol BS8 1TD, UK
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Abstract |
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(Received 13 June 2007;
accepted after revision 21 June 2007;
first published online 28 June 2007)
Corresponding author L. Pellerin: Departement de Physiologie, Rue du Bugnon 7 CH-1005 Lausanne, Switzerland. Email: luc.pellerin{at}unil.ch
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Introduction |
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Glucose is classically considered the main energy substrate of the brain (Sokoloff, 1989). In recent years, however, evidence has been provided for the notion that the brain can make use of alternative energy substrates under specific circumstances. For example, fatty acids provided by the diet readily cross the blood–brain barrier (BBB) and can contribute significantly to fulfil brain energy needs (Ebert et al. 2003). In addition to their direct utilization by the brain, fatty acids are converted in the liver to ketone bodies, i.e. β-hydroxybutyrate and acetoacetate, that can be also subsequently oxidized by brain cells. Under normal circumstances, the concentration of ketone bodies in the circulation is low and their utilization by the adult brain is considered of little physiological significance. But it can be reactivated during prolonged ketosis, diabetes and fasting (Cremer, 1982; Hawkins et al. 1986). Ketone bodies may thus become an important source of energy for the brain, depending of their blood concentration (for review, see Morris, 2005). Lactate is an additional energy substrate known to be used preferentially by neurons (Bouzier-Sore et al. 2003; Itoh et al. 2003). In addition to its formation from glucose within the brain parenchyma, notably by astrocytes (Pellerin, 2003), it was shown that elevation of blood lactate, as occurs during moderate exercise, is not only sufficient to promote its use by the brain (Dalsgaard et al. 2004) but it also reduces in parallel cerebral glucose consumption (Smith et al. 2003). Although the brain has the capacity to use alternative energy substrates in addition to glucose under particular circumstances, it is unknown whether an altered metabolic state (e.g. obesity) can lead to long-lasting brain adaptations that would pertain to both neuroenergetics and brain function.
Monocarboxylate transporters (MCTs) form a large family of proton-linked carriers that have the ability to transport lactate, pyruvate and the ketone bodies (Halestrap & Meredith, 2004; Pierre & Pellerin, 2005). In the central nervous system, three MCTs have been identified and their distribution determined at the cellular level. MCT1 is mostly expressed by endothelial cells and astrocytic processes in rodents (Gerhart et al. 1997; Pellerin et al. 1998; Leino et al. 1999; Hanu et al. 2000; Pierre et al. 2000; Baud et al. 2003) and in humans (Froberg et al. 2001; Chiry et al. 2006). MCT2 is the major neuronal transporter in the rodent brain (Pierre et al. 2002) while MCT4 is exclusively expressed in astrocytes (Bergersen et al. 2001; Rafiki et al. 2003; Pellerin et al. 2005). So far, there are few data reporting possible regulations of MCT expression in the adult brain under specific nutritional manipulations and none in relation to obesity.
In this study, we investigated the effect of a high fat diet (HFD), given up to 12 weeks, on levels of MCT1, MCT2 and MCT4 protein expression in the adult mouse brain. Moreover, we performed a parallel investigation using brains of mutant mice presenting a phenotype exhibiting common features (hyperinsulinaemia, hyperglycaemia, obesity) with mice fed a HFD. Our data reveal an up-regulation of all three monocarboxylate transporter isoforms in neurons from distinct brain areas compared to control mice fed with a standard diet. Such results suggest that the adult brain responds to peripheral metabolic changes with long-lasting adaptations.
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Methods |
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Six-week-old C57Bl6-Rj male mice were obtained from the Janvier Breeding Centre (Le Genest-Saint-Isle, France) and received 1 week prior to the experiments to acclimatize them to the animal facility. They were housed in a temperature-controlled environment with a 12: 12 h light–dark cycle and given free access to food and water. Animals were then placed randomly upon either a standard diet (SD, Provimi Kliba, Penthalaz, Switzerland. diet no. 34360.13, proteins 28%, carbohydrates 60% and fat 12%) or a fat-enriched diet (HFD, Harlan Teckland, Oxon, UK. Diet no. TD 93075, proteins 21%, carbohydrates 24%, fat 55%) during 2–12 weeks. Mice were weighed every 2 weeks and groups were killed after 2, 4, 6, 8, 10 and 12 weeks on either SD or HFD. Twelve hours prior to kill, animals were fasted. Five-week-old C57BL/KSJ@Rj-db (db/db) and C57BL/6J@Rj-ob (ob/ob) mice were obtained from Janvier Breeding Centre and housed 1 week with free access to food (standard diet) and water before being killed. At time of kill, the weight of db/db and ob/ob mice was 37.3 ± 1.6 g (n = 9) and 35.9 ± 1 g (n = 9), respectively. All experimental protocols were approved by the Cantonal Veterinary Office (Vaud, Switzerland).
Blood samples and biochemical assays
Prior to the dietary treatment, and then every 14 days, approximatively 500 µl of blood was taken by retro-orbitary puncture on 12 h-fasted animals and collected in heparinized tubes for glucose, insulin and β-hydroxybutyrate (BHB) plasmatic measurements. The last blood sample was collected by cardiac puncture, just prior to the animal kill. After centrifugation (2300g, 15 min), plasmas were frozen and stored at –80°C. Blood glucose was measured using the glucose oxidase method on a glucose analyser (Beckman Instruments). Plasmatic insulin was measured by the mouse insulin ELISA enzyme immunoassay (Mercodia, Uppsala, Sweden) and BHB levels were determined using D-3-hydroxybutyric acid colorimetric method (Boehringer Mannheim, Roche, Germany).
Immunohistochemistry
Under urethane anaesthesia (1.5 g kg–1 I.P.), animals were perfused intracardially with a heparinized solution of saline (25 IU ml–1 in 0.9% NaCl, during 2 min) followed by a freshly prepared solution of 4% paraformaldehyde in phosphate buffer (0.1 M, pH 7.4, for 15 min). Brains were removed and postfixed in 4% paraformaldehyde for 2 h, then cut on a vibratome (Leica VT1000M) to obtain coronal slices (40 µm thick), which were collected in phosphate buffer saline (PBS). Sections were rinsed several times in PBS and incubated with casein (0.5% in PBS) for 1 h to block non-specific sites. They were then incubated in rabbit antisera raised against either MCT1, or MCT2 (characterized in Pierre et al. 2000) or MCT4 (characterized in Wilson et al. 1998) diluted 1/500 in PBS-BSA (PBS containing 0.25% bovine serum albumin) overnight at 4°C in a humidified chamber. Sheep antirabbit immunoglobulins (Igs, 1/200, Jackson Immunoresearch Laboratories) followed by rabbit peroxidase–antiperoxidase complexes (1/500, Sigma) were used as immunolabels. The peroxidase reaction product was revealed with glucose oxidase-nickel-3,3'-DAB (Shu et al. 1988). For double labelling experiments, sections were incubated in a mixture of primary antibodies raised against one MCT (see above) and either against the neuronal marker NeuN (1/500, mouse monoclonal antibody, Chemicon International) or the glial fibrillary acidic protein (GFAP) (1/500, mouse monoclonal antibody, Sigma) overnight at 4°C. Immunofluorescence was revealed by incubation in a solution of second fluorescent antibodies (anti-rabbit Cy-3 and anti-mouse fluorescein isothiocyanate (FITC)-conjugated Igs, diluted 1/200, from Jackson Immunoresearch). Controls included omission of the primary serum or its replacement by non-immune rabbit serum. No specific staining was visible on such preparations. Sections were examined with a Zeiss axioplan2 microscope (Zeiss, Germany), using brightfield optics or epifluorescence. Pictures were acquired using a CCD camera (Axiocam, Zeiss, Germany) together with the 2.05 Axiovision software (Zeiss). They were also examined under a Leica SP5 AOBS confocal microscope using the Ar 488 nm and He/Ne 543 nm laser illumination. Frame size of the images was 512 x 512 pixels with 8-bit colour depth and stacks were made of images taken with 0.5–0.8 µm step size.
Western blotting
For Western blotting, we used extracts from whole brain or isolated cortex, hippocampus and cerebellum that were homogenized in a buffer of 0.32 M sucrose containing Hepes 1 mM, MgCl2 1 mM, NaHCO3 1 mM, phenyl-methyl-sulphonyl fluoride 0.1 mM, pH 7.4, in the presence of a complete set of protease inhibitors (Complete, Roche, Switzerland). Protein concentrations were determined by the method of Bradford (1976). Ten micrograms of proteins was heated at 95°C in SDS-PAGE sample buffer (62.5 mM Tris-HCl, 50 mM DTT, 2% SDS, 10% glycerol and 0.1% bromophenol blue) and loaded onto 10% polyacrylamide gels. After electrophoresis, samples were transferred to nitrocellulose membranes (Biorad Laboratories). Membranes were blocked in a blocking solution of Tris-buffered saline (TBS) containing 0.1% Tween, 10% non-fat milk and 1% BSA for 1 h. Membranes were incubated overnight at 4°C with a mixture containing either anti-MCT1 or anti-MCT2 antibody, diluted 1/1000 in TBS, 0.1% Tween, 1% milk (TBSTM) and anti-β-actin antibody (diluted 1/10000; Sigma). After several washes in TBSTM, they were incubated in a solution of IR-dye 800 anti-mouse IgG (diluted 1/5000, LI-COR Biosciences, Rockland) and Alexa Fluor 680 anti-rabbit IgG (Molecular Probes, Invitrogen detection technologies). Bound antibody signal quantification was done by infrared detection and analysis (Odyssey infrared detection system, LI-COR Biosciences). Statistics were performed using Student's t-test.
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Results |
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1.5-fold after 6 weeks, P < 0.001, Fig. 1B). Insulinaemia increased already after 2 weeks of diet in plasma of mice fed with HFD but enhancement was highly significant only after 8 weeks of HFD (3.5-fold, P < 0.01, Fig. 1C). Moreover, β-hydroxybutyrate (BHB) levels were significantly greater in plasma of mice fed with HFD than in SD-fed mice after 6 weeks of HFD and remained elevated up to 12 weeks (1.5-fold, P < 0.01, Fig. 1D), except at 8 weeks where the difference was not significant. Enhanced MCT expression in brains of mice fed with a high fat diet
Immunohistochemistry performed on brain sections from mice fed up to 12 weeks with a standard chow (SD) showed, in agreement with previous data (Pierre et al. 2000), that MCT1 immunoreactivity was very low throughout the brain. It occurred in the neuropile around immunonegative neuronal cell bodies and in endothelial cells forming microvessels (Fig. 2A). By contrast, feeding mice with HFD for 2 weeks only altered this pattern of expression since a very faint immunoreactivity for MCT1 appeared in neuronal somata throughout the cortex and also in the pyramidal layer of the hippocampus (Fig. 2B). Intensity of this neuronal immunolabelling increased with the duration of HFD feeding and reached a maximum between 6 and 10 weeks of HFD feeding (Fig. 2B–G). These changes were more pronounced in posterior sections, where the retrosplenial cortex is present but not the cingular cortex (not shown). After 8 weeks of HFD, cortical and hippocampal neurons displayed intense immunoreactivity for MCT1, occurring in their somata as well as in their processes (Figs 2Ec–Gc and 3B–D). The presence of immunopositive neuronal fibres was evident in cortical neurons as well as in pyramidal cells of CA1 and CA2 in the hippocampus, but not in CA3 (see Fig. 3C). Double labelling experiments using MCT1 and either NeuN, a neuronal marker, or GFAP, expressed by glial cells, confirmed that MCT1 immunoreactivity induced by HFD occurred specifically in neurons and not in astrocytes (Fig. 8Aa–c and D). HFD affected MCT1 distribution in other brain structures, such as the amygdala (Fig. 3E–F), as well as the auditory (not shown) and piriform cortices (Fig. 3G–H) where MCT1 immunolabelling occurred in cell bodies and proximal processes of many neurons (Fig. 3F and H). Increased neuronal MCT1 immunoreactivity was also observed in the striatum and the hypothalamus, but enhancement was less pronounced than in the cortex or the hippocampus (not shown).
As previously described (Pierre et al. 2002), MCT2 was expressed in the neuropile (associated with neuronal processes) in brain of mice fed with SD, but could not be detected in neuronal somata (Fig. 4Aa–c). It was also visible in capillaries (Fig. 4Aa). Two weeks of HFD triggered important changes in MCT2 expression since neuronal cell bodies became immunolabelled (Fig. 4B), as confirmed by double labelling experiments with NeuN (see Fig. 8Ba–c and E). Moreover, blood vessels became more intensely immunostained from 2 to 12 weeks of diet (Fig. 4Bc–Gc). Intensity of MCT2 labelling in neuronal somata increased gradually with time of diet, with a maximum reached after 6 weeks of HFD feeding, and had a tendency to decrease after 12 weeks of HFD (Fig. 4B–G). MCT2-immunopositive neurons were very abundant throughout the retrosplenial, somatosensory, auditory and piriform cortices (Fig. 5B and F), the hippocampus (Fig. 5D) and the amygdala (Fig. 5F), and to a lesser extent in the hypothalamus (Fig. 5G–H) and the striatum (not shown). Similarly to MCT1, MCT2-positive neuronal processes became visible after 6 weeks of diet.
MCT4 is considered to be a specific astrocytic transporter based on its previously established distribution (Bergersen et al. 1999; Bergersen et al. 2001; Rafiki et al. 2003; Pellerin et al. 2005). Consistent with these observations, MCT4 immunoreactivity in brain of mice fed with SD up to 12 weeks was found exclusively in astrocytes throughout the whole brain (Fig. 6A). After 2 weeks of HFD, neuron-like cells appeared immunostained for MCT4 in the mouse cortex, in addition to astrocytes (Fig. 6B). We performed double labelling experiments with NeuN or GFAP in order to identify the cell type(s) expressing MCT4 in brains of mice fed with HFD. As for both MCT1 and MCT2, MCT4 was expressed by neurons, as shown by superposition of MCT4 and NeuN immunostainings (Fig. 8Ca–c), but remained also visible in astrocytes since MCT4 and GFAP immunoreactivities partly colocalized (Fig. 8F). Neuronal cell bodies and processes that were immunostained for MCT4 displayed gradually a more intense immunoreaction with time of diet, while labelling of MCT4-positive astrocytes remained unchanged (Fig. 6C–G). Neuronal MCT4 expression induced by HFD was found mainly in the same brain structures where enhanced MCT1 and MCT2 expression was observed, i.e. the cortex (Fig. 7B), the amygdala (Fig. 7D) and the hippocampus (Fig. 7F). As compared to the other transporters, the changes in MCT4 expression were less convincing in the hypothalamus and the striatum (not shown).
Quantification of MCT1 and MCT2 expression induced by HFD
Since the increased expression of MCTs due to nutritional manipulations was stronger in rostral (containing retrosplenial, somatosensory and auditory cortex) than in frontal brain coronal sections, we performed brain dissection following the scheme shown in Fig. 9A, in order to distinguish what we termed cortex 1 (containing cingulate and motor cortex), cortex 2, including piriform, insular and entorhinal cortex, and cortex 3, composed of the somatosensory, the retrosplenial and the visual cortex. The effect of a 6 weeks of HFD on MCT expression was quantified using Western blots performed on extracts from different brain structures (Fig. 9B). We averaged results from two to three Western blots made on extracts obtained from different mice in order to perform statistical analysis. MCT1 protein level was slightly increased in cortex 1 of mice fed with HFD when compared to mice fed with SD (1.2-fold, P < 0.05, Fig. 9C). By contrast, the increase was highly significant in cortex 2 (1.4-fold, P < 0.01), cortex 3 (1.85-fold, P < 0.01) and hippocampus (1.45-fold, P < 0.01), while it was not affected in cerebellum (Fig. 9C). MCT2 expression was neither increased in cortex 1 nor in cerebellum of mice fed with HFD compared to mice fed with SD (Fig. 9C). By contrast, it was significantly enhanced in cortex 2 (1.8-fold, P < 0.001), cortex 3 (1.5-fold, P < 0.01) and hippocampus (1.44-fold, P < 0.01) (Fig. 9C).
Increased brain MCT expression in ob/ob or db/db mice fed with a standard diet
In order to distinguish between a direct effect of nutrients (e.g. fatty acids) on brain from other possible consequences of the resulting altered metabolic state induced by a high fat diet, we assessed MCT expression levels in brains of two mouse models presenting some common metabolic features (e.g. obesity) but fed with a standard diet. Six-week-old ob/ob or db/db mouse brains were investigated for MCT1, MCT2 and MCT4 expression levels. Immunolabelling using immunoperoxidase showed that the three MCTs were expressed in numerous neurons of the brain of both ob/ob and db/db mice (Fig. 10). Only cortical immunostaining is illustrated (Fig. 10) but many other brain structures were immunostained for all three MCTs, similarly to what was observed in the brain of mice fed with HFD (not shown). Double fluorescence immunolabelling using the neuronal marker NeuN confirmed neuronal expression of the three transporters (Fig. 10Ac–Fc). MCT1 and MCT2 protein levels were quantified in ob/ob and db/db versus wild-type (WT, C57Bl/6J) mice using Western blots performed on extracts from different brain structures. We averaged results from two Western blots made on extracts obtained from different mice in order to perform statistics (Fig. 11). In both cortex 1 and cerebellum, neither MCT1 nor MCT2 levels were significantly changed in ob/ob or db/db mice when compared to WT. By contrast, they increased significantly in cortex 2 of ob/ob mice (2-fold, P < 0.001 for MCT1; 2.5-fold, P < 0.001 for MCT2) and db/db mice (1.5-fold, P < 0.01 for MCT1; 3.8-fold, P < 0.001 for MCT2). A significant increase was also observed in cortex 3 of both ob/ob (1.3-fold, P < 0.01 for MCT1; 2.3-fold, P < 0.001 for MCT2) and db/db (1.3-fold, P < 0.01 for MCT1 and 2.3-fold, P < 0.001 for MCT2) mice as well as in the hippocampus (1.8-fold, P < 0.001 for MCT1 and 1.5-fold, P < 0.001 for MCT2 in ob/ob mice, and 1.8-fold, P < 0.001 for MCT1 and 1.7-fold, P < 0.001 for MCT2 in db/db mice).
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Discussion |
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The precise mechanism by which exposure to a high fat diet leads to enhanced MCT expression in brain remains uncertain for the moment. However, our observations that genetically induced obese mice (ob/ob) as well as genetically induced diabetic mice (db/db) fed on a standard diet exhibit a similar enhancement of MCT expression argue against a direct role of fatty acids. As an alternative possibility, hormonal alterations occurring as a consequence of exposure to a high fat diet and development of obesity could participate in the up-regulation of MCT expression in the central nervous system. Increased fat mass induced by a high fat diet, as occurs in obesity, leads to enhanced production and circulating levels of adiposity signals such as leptin and insulin (Benoit et al. 2004; Woods et al. 2004). Leptin was shown to activate several signal transduction mechanisms and induce gene expression in peripheral organs as well as in the central nervous system (Fruhbeck, 2006). However, in both the ob/ob mouse, which carries a mutation of the gene coding for leptin, and the db/db mouse, which has a mutated leptin receptor, the same alterations in MCT expression were present despite the disruption of the putative leptin signal. Thus, it is unlikely that leptin is the main signal causing enhanced cerebral MCT expression in mice fed with a high fat diet. Another feature commonly observed in obese animals is an elevation of circulating levels of insulin (Woods et al. 2004). Hyperinsulinaemia was not only observed in our mice fed with a high fat diet but it is also a characteristic of both ob/ob and db/db mice. Moreover, it is well known that insulin enters into the brain and modulates a number of cerebral functions, from energy and glucose homeostasis to learning and memory (Gerozissis, 2003; Plum et al. 2005). Interestingly, we have recently observed that insulin can enhance the expression of MCT2 in cultured cortical neurons (Chenal & Pellerin, 2006). Based on these observations, we believe that insulin represents a good candidate for the putative signal to mediate the changes of neuronal MCT expression in each obese mouse model we have investigated.
Monocarboxylate transporters have been implicated in the intercellular transfer of lactate in different organs including the brain (Brooks, 1986; Pellerin & Magistretti, 1994; Gladden, 2004). Moreover, as mentioned previously, they are involved in the import and the use of energy substrates such as lactate and the ketone bodies in particular metabolic situations. Exposure to a high fat diet sets a new metabolic environment for the brain that most likely leads to specific metabolic adaptations. Similarly, mice developing features of the metabolic syndrome following specific genetic mutations are likely to undergo cerebral modifications in response to such altered metabolic conditions. Although insulin can exert a certain influence on brain glucose metabolism, it might also have a significant impact on other aspects of neuroenergetics, including the use of alternative energy substrates such as monocarboxylates. In parallel with hyperinsulinaemia and hyperglycaemia, one common feature encountered with the development of obesity is the appearance of insulin resistance (Kahn et al. 2006). Such resistance to the action of insulin is well described for tissues like muscle or liver, but it was recently proposed that it might also affect the brain (Isganaitis & Lustig, 2005). Specific metabolic consequences of altered insulin sensitivity in the brain have not been explored but overexpression of monocarboxylate transporters by neurons as observed here could be a response to overcome brain insulin resistance and its impact on the utilization of energy substrates by brain cells. Such an observation might be of interest in relation to the suggestion that a relation exists between insulin resistance, metabolic deficits and the development of cognitive decline as observed in ageing and neurodegenerative diseases (Watson & Craft, 2006).
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References |
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