Accepted Manuscript Brain Glucose Metabolism in an Animal Model of Depression Jan Detka, Anna Kurek, Mateusz Kucharczyk, Katarzyna Glombik, Agnieszka Basta-Kaim, Marta Kubera, Władysław Lasoń, Bogusława Budziszewska PII: DOI: Reference:
S0306-4522(15)00283-3 http://dx.doi.org/10.1016/j.neuroscience.2015.03.046 NSC 16156
To appear in:
Neuroscience
Accepted Date:
19 March 2015
Please cite this article as: J. Detka, A. Kurek, M. Kucharczyk, K. Glombik, A. Basta-Kaim, M. Kubera, W. Lasoń, B. Budziszewska, Brain Glucose Metabolism in an Animal Model of Depression, Neuroscience (2015), doi: http://dx.doi.org/10.1016/j.neuroscience.2015.03.046
This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
BRAIN GLUCOSE METABOLISM IN AN ANIMAL MODEL OF DEPRESSION
Jan Detka, Anna Kurek, Mateusz Kucharczyk, Katarzyna Glombik , Agnieszka Basta-Kaim, Marta Kubera, Władysław Lasoń, Bogusława Budziszewska
Department of Experimental Neuroendocrinology, Institute of Pharmacology, Polish Academy of Sciences, Smętna 12, PL 31-343 Kraków, Poland
Correspondence: Bogusława Budziszewska, e-mail: [email protected]
1
Abstract An increasing number of data support the involvement of disturbances in glucose metabolism in the pathogenesis of depression. We previously reported that glucose and glycogen concentrations in brain structures important for depression are higher in a prenatal stress model of depression when compared with control animals. A marked rise in the concentrations of these carbohydrates and glucose transporters were evident in prenatally stressed animals subjected to acute stress and glucose loading in adulthood. To determine whether elevated levels of brain glucose are associated with a change in its metabolism in this model, we assessed key glycolytic enzymes (hexokinase, phosphofructokinase and pyruvate kinase), products of glycolysis, i.e., pyruvate and lactate, and two selected enzymes of the tricarboxylic acid cycle (pyruvate dehydrogenase and α-ketoglutarate dehydrogenase) in the hippocampus and frontal cortex. Additionally, we assessed glucose-6-phosphate dehydrogenase activity, a key enzyme in the pentose phosphate pathway (PPP). Prenatal stress increased the levels of phosphofructokinase, an important glycolytic enzyme, in the hippocampus and frontal cortex. However, prenatal stress had no effect on hexokinase or pyruvate kinase levels. The lactate concentration was elevated in prenatally stressed rats in the frontal cortex, and pyruvate levels remained unchanged. Among the tricarboxylic acid cycle enzymes, prenatal stress decreased the level of pyruvate dehydrogenase in the hippocampus, but it had no effect on α-ketoglutarate dehydrogenase. Like in the case of glucose and its transporters, also in the present study, differences in markers of glucose metabolism between control animals and those subjected to prenatal stress were not observed under basal conditions but in rats subjected to acute stress and glucose load in adulthood. Glucose-6phosphate dehydrogenase activity was not reduced by prenatal stress but was found to be even higher in animals exposed to all experimental conditions, i.e. prenatal stress, acute stress, and glucose administration. Our data indicate that glycolysis is increased and the Krebs cycle is decreased in the brain of a prenatal stress animal model of depression. Key words: depression, prenatal stress, brain glucose metabolism, phosphofructokinase, lactate, pyruvate dehydrogenase
2
INTRODUCTION Depression is a complex mental illness with poorly recognized etiology. Growing evidence indicates that not only disturbed monoamine transmission is involved in the pathogenesis of this disease, but also hyperactivity of the hypothalamic-pituitary-adrenal (HPA) axis, excessive activation of proinflammatory cytokines, overstimulation with excitatory amino acids and a weaker action of neurotropic factors play an important role (Marsden et al., 2013; Zunszain et al., 2011). Experimental and clinical data support the role of stress and increased glucocorticoid action in the etiology and progression of major depression (Holsboer, 2000). The HPA axis hyperactivity, most likely due to the weakened inhibitory feedback mechanism, often occurs in depression and stress is a common trigger of a depressive episode. It has been shown that stress or excess glucocorticoid in experimental animals produces many of the behavioral, biochemical, functional and morphological characteristics of depression (Pariante and Lightman, 2008). Notably, particular stress factors acting during the perinatal period cause permanent changes in the HPA axis activity and increase the risk of depression (Barbazanges et al., 1996). Factors adversely affecting maternal environment, by persistent stimulation of the HPA axis activity, increases the risk of not only mental but also metabolic and cardiovascular diseases. Indeed, clinical data frequently show the co-occurrence of depression with obesity, metabolic syndrome, type-2 diabetes, hypertension and adverse coronary events (Murgatroyd et al., 2009; Bouwman et al., 2010). For example, depression increases the risk for diabetes by approximately 60%, and people with diabetes are twice as likely to have depression when compared with the general population (Nouwen et al., 2010). Available data indicate that the main reason for the coexistence of these diseases may be metabolic disturbances resulting from excessive glucocorticoid action (Detka et al., 2013). In fact, glucocorticoids, which are frequently used in the clinic, can evoke new-onset hyperglycemia in patients without diabetes and induce uncontrolled hyperglycemia in patients with diabetes (Baldwin and Apel, 2013). In experimental animals, glucocorticoids increase blood glucose levels and evoke peripheral insulin resistance at pharmacological doses and at the concentrations observed under stress (Li et al., 2013). In contrast to the well-defined peripheral metabolic effects of glucocorticoids, the central metabolic actions of these hormones and metabolic disturbances in depression are still poorly understood. Glucose is the main energy source for the adult brain, and its metabolism enables the proper synthesis and function of neurotransmitters, particularly glutamate and γ-aminobutyric acid (GABA), and the formation of appropriate levels of NADPH. Current data support the involvement of glucose metabolism dysfunction in the pathogenesis of depression; however, reports on the specific metabolic changes that occur in this disorder and the effect of glucocorticoids on brain glucose metabolism are conflicting. Previous studies of the metabolic activity in various brain areas of depressed patients did not provide clear results, most likely due to the diversity of depressive disorders (Hosokawa et al., 2009). Furthermore, experimental data are inconclusive. In peripheral tissues glucocorticoids inhibit insulinstimulated glucose uptake, down-regulate glucose metabolism, mainly process of glycolysis, and inhibit insulin signaling (Kuo et al., 2013). Some studies have shown that also in the 3
brain these hormones may exert similar effect on glucose uptake and metabolism (Osmanovic et al., 2010; Piroli et al., 2007). Such action may in turn lead to a reduction in energy production and disrupt the normal function and plasticity of brain cells. This theory is supported by studies showing that chronic corticosterone treatment decreases glucose metabolism in the rat parietotemporal cortex and hippocampus; in addition, adrenalectomy increases glucose metabolism in these brain regions (Hoyer and Lannert, 2008; Plaschke et al., 1996). The majority of animal models of depression are based on the long-term effect of various stress factors; however, none of these models have thoroughly examined glucose metabolism. Previous studies have shown that global cerebral glucose utilization is decreased in olfactory bulbectomized rat models of depression (Skelin et al., 2008), and rats subjected to a chronic unpredictable mild stress model of depression exhibit insulin resistance in the arcuate nucleus of hypothalamus (Pan et al., 2013). Some data do not support the hypothesis of reduced cerebral glucose metabolism in animal models of depression. For example, the Flinders Sensitive Line rat, a well-validated model of depression, exhibits increased glucose utilization in brain structures important in the pathogenesis of depression (Kanemaru and Diksic, 2009). Additionally, our previous studies demonstrated that glucose, glycogen and glucose transporter (GLUT-1 and GLUT-4) concentrations are higher in the hippocampus and frontal cortex in prenatally stressed rats when compared with control animals (Detka et al., 2014). These data and the results obtained with Flinders Sensitive Line rats indicate that brain metabolic changes in animal models of depression are complex and various stages of glucose metabolism (uptake, glycolysis, the Krebs cycle, and oxidative phosphorylation) in particular brain regions can be differentially disturbed. The aim of the present study was to determine if high glucose and glycogen levels in the hippocampus and frontal cortex in a prenatal stress model of depression result from only the intensity of glucose uptake or if this change requires a weakening of glycolysis and/or the Krebs cycle. Thus, we examined key glycolytic enzymes (hexokinase, phosphofructokinase and pyruvate kinase), products of glycolysis, i.e., pyruvate and lactate, and two selected enzymes of the Krebs cycle (pyruvate dehydrogenase and αketoglutarate dehydrogenase) in the hippocampus and frontal cortex, i.e., brain structures important for depression, using a prenatal stress model of depression. In addition, glucose-6phosphate dehydrogenase was examined because glucose metabolism in the pentose cycle is a major source of NADPH that can be used to remove excess of reactive oxygen species while oxidative stress plays a role in the pathogenesis of depression. Previously observed changes in glucose, glycogen and glucose transporters were stronger after exposure of prenatally stressed rats to adverse factors (acute stress, glucose load) in adulthood (Detka et al., 2014). For the current study, we assessed the concentrations of selected markers involved in glucose metabolism at basal conditions and in animals subjected to stress or administered glucose as adults. We chose to study this prenatal stress model of depression because the face, predictive and construct validities of this model are well defined and in addition to depression-like behavioral changes, hyperactivity of the HPA axis and increases in blood glucose levels have been described in this model (Koehl et al., 1999; Morley-Fletcher et al., 2003, 2004; Szymańska et al., 2009a). Additionally changes induced by prenatal stress appear gradually 4
and are present in adult animals, which allows for the determination of progressive responses to adverse factors. Our previous studies using this model showed also more potent glucocorticoid action in the brain (connected with changes in factors that regulate glucocorticoid receptor function), increased concentrations of glucose, glycogen, glucose transporters and corticosterone in the brain, and insulin resistance (Szymańska et al., 2009a, 2009b; Budziszewska et al., 2010; Detka et al., 2014).
EXPERIMENTAL PROCEDURES Animals Sprague-Dawley rats (200-250 g) were purchased from a licensed dealer. Rats were kept in an animal housing facility at a room temperature of 23°C and 12/12 h light/dark cycle (lights on at 07 a.m.) with food and water available ad libitum. All experiments were carried out according to the National Institutes of Health Guide for the Care and Use of Laboratory Animals and were approved by the Local Ethics Committee in Kraków, Poland. All efforts were made to minimize the number of animals used and their suffering. One week after arrival, vaginal smears from females were collected daily in order to determine estrous cycle phase. During proestrus, rats were placed with males for 12 h. Next, we tested vaginal smears for the presence of sperm. Pregnant females were than randomly assigned to the control and stress groups. Stress procedure Prenatal stress was performed as previously described by Morley-Fletcher et al. (2003, 2004). Briefly, pregnant rats were subjected daily to three stress sessions starting at 09.00, 12.00 and 17.00 h. During this time, rats were placed in plastic cylinders (7/12 cm) and exposed to a bright light for 45 min. Stress sessions were performed from day 14 of pregnancy until delivery. Control pregnant females were left undisturbed in their home cages. Twenty-one days after birth, male offspring from litters containing 10-14 pups with a comparable number of males and females were separated for experiments. One to 2 animals from each litter were randomly assigned to 1 of 8 designated experimental groups. Rats were housed in groups of five per cage under standard conditions until 3 months of age. Forced swimming test in rats The 3-month-old animals were individually subjected to two trials in which they were forced to swim in a cylinder (40 cm high, 18 cm in diameter) filled with water (25o C) up to a height of 15 cm. There was a 24-hour interval between the first and second trial. The first trial lasted for 15 min, and the second trial lasted for 5 min. The total duration of immobility and climbing time were measured throughout the second trial (Porsolt et al., 1978). Acute immobilization stress in adult animals Half of the animals from the control and prenatally stressed groups were subjected to acute immobilization stress. The rats were placed in plastic cylinders (7/12 cm) for 1 h. Oral glucose administration 5
Immediately after acute stress, half of the animals from each control and prenatally stressed group with and without acute immobilization stress were administered glucose (1 g/kg) by oral gavage as previously described (Dias et al., 2010). The remaining animals received water in place of glucose. Glucose or water was given in a volume of 2 ml/kg. Tissue collection Two hours after glucose administration, animals were killed under non-stress conditions by rapid decapitation. Brains were rapidly removed, and brain regions (hippocampus and frontal cortex) were dissected on ice-cold glass plates. The tissues were frozen on dry ice and stored at -80°C. Scheme showing the time course of the experiment is shown in Figure 1.
Biochemical analysis Lactate estimation Lactate was measured in the frontal cortex and hippocampus using a lactate fluorometric assay kit (Biovision, USA) according to the manufacturer's instructions. Briefly, fresh tissues were homogenized in 5 volumes of phosphate-buffered saline (PBS) and centrifuged at 20000 g for 20 min at 4°C. The supernatants were then deproteinized using perchloric acid (PCA) and a Deproteinizing Sample Detection Kit (Biovision, USA) according to the procedure provided by the manufacturer. Briefly, 250 µl of sample homogenates were mixed with 50 µl of ice-cold PCA, incubated on ice for 5 minutes, and then centrifuged at 20 000 g for 2 min at 4 °C. A 50 µl aliquot of each sample was transferred to 96-well plates in duplicate along with lactate standards (0; 0.2; 0.4; 0.6; 0.8, and 1.0 nmol/well). A total of 50 µl of reaction mix (46 µl lactate assay buffer, 2 µl probes, and 2 µl of enzyme mix) was added to each test sample and lactate standard. Plates were incubated for 30 min at room temperature. Fluorescence was measured at Ex/Em=535/590 in a fluorometer (Tecan Infinite 200 Pro). The lactate concentration in samples was calculated from a standard curve and expressed as nmol/mg protein. Pyruvate estimation Pyruvate was measured in the frontal cortex and hippocampus using a fluorometric pyruvate assay kit (Biovision, USA) according to the manufacturer's instructions. Briefly, fresh tissues were homogenized in 5 volumes of phosphate-buffered saline (PBS) and centrifuged at 20000 g for 20 min at 4°C. The supernatants were than deproteinized using perchloric acid (PCA) and a Deproteinizing Sample Detection Kit (Biovision, USA) according to the procedure provided by the manufacturer. Briefly, 250 µl of sample homogenates were mixed with 50 µl of ice-cold PCA, incubated on ice for 5 minutes, and then centrifuged at 20 000 g for 2 min at 4 °C. A 50 µl aliquot of each sample was transferred to 96-well plates in duplicate along with pyruvate standards (0; 0.2; 0.4; 0.6; 0.8 and 1.0 nmol/well). A total of 50 µl/well of reaction mix (47.6 µl pyruvate assay buffer, 0.4 µl pyruvate probes, and 2 µl enzyme mix) was added to each test sample and pyruvate standard. Plates were incubated for 30 min at room temperature. Fluorescence was measured at Ex/Em=535/590 in a fluorometer (Tecan Infinite 200 Pro). The pyruvate concentration in samples was calculated from a standard curve and expressed as nmol/mg protein.
6
Hexokinase The concentration of hexokinase in the frontal cortex and hippocampus was determined using an ELISA method (Wuhan EIAab Science Co., Ltd, China) according to the manufacturer’s instructions. Briefly, the tissues were homogenized in phosphate-buffered saline (PBS), shaken in an ice bath for 30 minutes and then centrifuged at 20 000 g for 20 min at 4 °C. An aliquot of 100 µl of each sample was transferred to pre-coated 96-well ELISA plates in duplicate along with hexokinase standards (0; 0.156; 0.312; 0.625; 1.25; 2.5; 5 and 10 ng/ml). The absorbance was measured at a wavelength λ=450 nm using a spectrophotometer (MultiskanSpectrum, ThermoLabsystems). The hexokinase concentration in samples was calculated from a standard curve and expressed as ng/mg protein. Phosphofructokinase The concentration of phosphofructokinase in the frontal cortex and hippocampus was measured using an ELISA method (USCN Life Science Inc., USA) according to the manufacturer’s instructions. Briefly, the tissues were homogenized in phosphate-buffered saline (PBS), shaken in an ice bath for 30 minutes and then centrifuged at 20 000 g for 20 min at 4 °C. An aliquot of 100 µl of each sample was transferred to pre-coated 96-well ELISA plates in duplicate along with phosphofructokinase standards (0; 1.56; 3.12; 6.25; 12.5; 25; 50 and 100 ng/ml). The absorbance was measured at a wavelength λ=450 nm using a spectrophotometer (MultiskanSpectrum, ThermoLabsystems). The concentration of phosphofructokinase was calculated from a standard curve and expressed as ng/mg protein. Pyruvate kinase The concentration of pyruvate kinase in the frontal cortex and hippocampus was determined using an ELISA method (Wuhan EIAab Science Co., Ltd, China) according to the manufacturer’s instructions. Briefly, the tissues were homogenized in phosphate-buffered saline (PBS), shaken in an ice bath for 30 minutes and then centrifuged at 20 000 g for 20 min at 4 °C. An aliquot of 100 µl of each sample was transferred to pre-coated 96-well ELISA plates in duplicate along with phosphofructokinase standards (0; 1.25; 2.5; 5; 10; 20; 40 and 80 Units/ml). The absorbance was measured at a wavelength λ=450 nm using a spectrophotometer (MultiskanSpectrum, ThermoLabsystems). The pyruvate kinase concentration in the samples was calculated from a standard curve and expressed as Units/mg protein. Isolation of mitochondria Concentration of pyruvate dehydrogenase and of α-ketoglutarate dehydrogenase was determined in mitochondrial fractions. Isolation of crude mitochondrial fraction from the frontal cortex and hippocampus was performed accordingly to the procedure described by Wernicke et al. (2010). Briefly, brain tissues, kept on ice were homogenized in motor-driven Teflon-glass homogenizer (9 strokes, 600 rpm) in 4 volumes of homogenization buffer, containing (5 mol/l HEPES/NaOH, pH 7.4, 320 mmol/l sucrose and 1 mmol/l Na+/EDTA with the addition of 0.5% protease inhibitor cocktail (Sigma P8340). After centrifugation in 1 300 × g (4 minutes, 4°C) supernatants were collected. Additionally, to increase the yield, pellet was washed twice with homogenization buffer and centrifuged at 1500 × g (4 minutes, 4°C). In order to collect the mitochondria, supernatants were combined, and centrifuged at 17 000 × g for 12 min at 4°C. After discarding of supernatants, the obtained pellets containing mitochondria were rapidly frozen and stored at –80°C. 7
Pyruvate dehydrogenase The concentration of pyruvate dehydrogenase 1 in the mitochondrial fraction of the frontal cortex and hippocampus was determined using an ELISA method (Wuhan EIAab Science Co., Ltd, China) according to the manufacturer’s instructions. An aliquot of 100 µl of each mitochondrial sample was transferred to pre-coated 96-well ELISA plates in duplicate along with phosphofructokinase standards (0; 0,312; 0.625; 1.25; 2.5; 5; 10; 20 and 40 ng/ml). The absorbance was measured at a wavelength λ=450 nm using a spectrophotometer (MultiskanSpectrum, ThermoLabsystems). The pyruvate dehydrogenase concentration in the samples was calculated from a standard curve and expressed as ng/mg protein. α-Ketoglutarate dehydrogenase The concentration of α-ketoglutarate dehydrogenase was measured in the mitochondrial fraction of the frontal cortex and hippocampus using a specific ELISA Kit (Wuhan EIAab Science Co., Ltd, China) according to the manufacturer’s instructions. An aliquot of 100 µl of each sample was transferred to pre-coated 96-well ELISA plates in duplicate along with αketoglutarate dehydrogenase standards (0; 0.075; 0.15; 0.31; 0.62; 1.25; 2.5 and 5 ng/ml). The absorbance was measured at a wavelength λ=450 nm using a spectrophotometer (MultiskanSpectrum, ThermoLabsystems). The concentration of α-ketoglutarate dehydrogenase in samples was calculated from a standard curve and expressed as ng/mg protein. Glucose-6-phosphate dehydrogenase activity The activity of glucose-6-phosphate dehydrogenase in the frontal cortex and hippocampus was determined using a colorimetric assay kit (TrinityBiotech, Ireland) according to the instructions provided by the manufacturer. A 10 µl aliquot of each sample was transferred in duplicate in 96-well plates and mixed with 50 µl of the Assay Solution. A total of 100 µl of the G-6-PDH Substrate Solution was added into sample wells. The absorbance was measured at a wavelength λ=340 nm using a MultiskanSpectrum spectrophotometer (ThermoLabsystems). The absorbance was measured immediately after addition of the substrate and after 5 minutes of incubation in order to determine enzymatic reaction kinetics. The activity of glucose-6-phosphate dehydrogenase was calculated and expressed as enzyme units/5 minutes/mg protein. Protein determination The protein content of the analyzed samples was determined using the method of Lowry et al. (1951). Statistical Analysis The data from the eight experimental groups, each containing 10 animals, are presented as the mean ± SEM, and the results were analyzed using the STATISTICA program. The homogeneity of the variance was first analyzed with a Levene’s test. The subsequent statistical analyses were performed by factorial analyses of variance (ANOVAs) to determine the effects of three factors (i.e., factor 1- prenatal stress, factor 2 - acute stress and factor 3 glucose). Next, Duncan’s post-hoc test was applied. A p-value of
S0306-4522(15)00283-3 http://dx.doi.org/10.1016/j.neuroscience.2015.03.046 NSC 16156
To appear in:
Neuroscience
Accepted Date:
19 March 2015
Please cite this article as: J. Detka, A. Kurek, M. Kucharczyk, K. Glombik, A. Basta-Kaim, M. Kubera, W. Lasoń, B. Budziszewska, Brain Glucose Metabolism in an Animal Model of Depression, Neuroscience (2015), doi: http://dx.doi.org/10.1016/j.neuroscience.2015.03.046
This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
BRAIN GLUCOSE METABOLISM IN AN ANIMAL MODEL OF DEPRESSION
Jan Detka, Anna Kurek, Mateusz Kucharczyk, Katarzyna Glombik , Agnieszka Basta-Kaim, Marta Kubera, Władysław Lasoń, Bogusława Budziszewska
Department of Experimental Neuroendocrinology, Institute of Pharmacology, Polish Academy of Sciences, Smętna 12, PL 31-343 Kraków, Poland
Correspondence: Bogusława Budziszewska, e-mail: [email protected]
1
Abstract An increasing number of data support the involvement of disturbances in glucose metabolism in the pathogenesis of depression. We previously reported that glucose and glycogen concentrations in brain structures important for depression are higher in a prenatal stress model of depression when compared with control animals. A marked rise in the concentrations of these carbohydrates and glucose transporters were evident in prenatally stressed animals subjected to acute stress and glucose loading in adulthood. To determine whether elevated levels of brain glucose are associated with a change in its metabolism in this model, we assessed key glycolytic enzymes (hexokinase, phosphofructokinase and pyruvate kinase), products of glycolysis, i.e., pyruvate and lactate, and two selected enzymes of the tricarboxylic acid cycle (pyruvate dehydrogenase and α-ketoglutarate dehydrogenase) in the hippocampus and frontal cortex. Additionally, we assessed glucose-6-phosphate dehydrogenase activity, a key enzyme in the pentose phosphate pathway (PPP). Prenatal stress increased the levels of phosphofructokinase, an important glycolytic enzyme, in the hippocampus and frontal cortex. However, prenatal stress had no effect on hexokinase or pyruvate kinase levels. The lactate concentration was elevated in prenatally stressed rats in the frontal cortex, and pyruvate levels remained unchanged. Among the tricarboxylic acid cycle enzymes, prenatal stress decreased the level of pyruvate dehydrogenase in the hippocampus, but it had no effect on α-ketoglutarate dehydrogenase. Like in the case of glucose and its transporters, also in the present study, differences in markers of glucose metabolism between control animals and those subjected to prenatal stress were not observed under basal conditions but in rats subjected to acute stress and glucose load in adulthood. Glucose-6phosphate dehydrogenase activity was not reduced by prenatal stress but was found to be even higher in animals exposed to all experimental conditions, i.e. prenatal stress, acute stress, and glucose administration. Our data indicate that glycolysis is increased and the Krebs cycle is decreased in the brain of a prenatal stress animal model of depression. Key words: depression, prenatal stress, brain glucose metabolism, phosphofructokinase, lactate, pyruvate dehydrogenase
2
INTRODUCTION Depression is a complex mental illness with poorly recognized etiology. Growing evidence indicates that not only disturbed monoamine transmission is involved in the pathogenesis of this disease, but also hyperactivity of the hypothalamic-pituitary-adrenal (HPA) axis, excessive activation of proinflammatory cytokines, overstimulation with excitatory amino acids and a weaker action of neurotropic factors play an important role (Marsden et al., 2013; Zunszain et al., 2011). Experimental and clinical data support the role of stress and increased glucocorticoid action in the etiology and progression of major depression (Holsboer, 2000). The HPA axis hyperactivity, most likely due to the weakened inhibitory feedback mechanism, often occurs in depression and stress is a common trigger of a depressive episode. It has been shown that stress or excess glucocorticoid in experimental animals produces many of the behavioral, biochemical, functional and morphological characteristics of depression (Pariante and Lightman, 2008). Notably, particular stress factors acting during the perinatal period cause permanent changes in the HPA axis activity and increase the risk of depression (Barbazanges et al., 1996). Factors adversely affecting maternal environment, by persistent stimulation of the HPA axis activity, increases the risk of not only mental but also metabolic and cardiovascular diseases. Indeed, clinical data frequently show the co-occurrence of depression with obesity, metabolic syndrome, type-2 diabetes, hypertension and adverse coronary events (Murgatroyd et al., 2009; Bouwman et al., 2010). For example, depression increases the risk for diabetes by approximately 60%, and people with diabetes are twice as likely to have depression when compared with the general population (Nouwen et al., 2010). Available data indicate that the main reason for the coexistence of these diseases may be metabolic disturbances resulting from excessive glucocorticoid action (Detka et al., 2013). In fact, glucocorticoids, which are frequently used in the clinic, can evoke new-onset hyperglycemia in patients without diabetes and induce uncontrolled hyperglycemia in patients with diabetes (Baldwin and Apel, 2013). In experimental animals, glucocorticoids increase blood glucose levels and evoke peripheral insulin resistance at pharmacological doses and at the concentrations observed under stress (Li et al., 2013). In contrast to the well-defined peripheral metabolic effects of glucocorticoids, the central metabolic actions of these hormones and metabolic disturbances in depression are still poorly understood. Glucose is the main energy source for the adult brain, and its metabolism enables the proper synthesis and function of neurotransmitters, particularly glutamate and γ-aminobutyric acid (GABA), and the formation of appropriate levels of NADPH. Current data support the involvement of glucose metabolism dysfunction in the pathogenesis of depression; however, reports on the specific metabolic changes that occur in this disorder and the effect of glucocorticoids on brain glucose metabolism are conflicting. Previous studies of the metabolic activity in various brain areas of depressed patients did not provide clear results, most likely due to the diversity of depressive disorders (Hosokawa et al., 2009). Furthermore, experimental data are inconclusive. In peripheral tissues glucocorticoids inhibit insulinstimulated glucose uptake, down-regulate glucose metabolism, mainly process of glycolysis, and inhibit insulin signaling (Kuo et al., 2013). Some studies have shown that also in the 3
brain these hormones may exert similar effect on glucose uptake and metabolism (Osmanovic et al., 2010; Piroli et al., 2007). Such action may in turn lead to a reduction in energy production and disrupt the normal function and plasticity of brain cells. This theory is supported by studies showing that chronic corticosterone treatment decreases glucose metabolism in the rat parietotemporal cortex and hippocampus; in addition, adrenalectomy increases glucose metabolism in these brain regions (Hoyer and Lannert, 2008; Plaschke et al., 1996). The majority of animal models of depression are based on the long-term effect of various stress factors; however, none of these models have thoroughly examined glucose metabolism. Previous studies have shown that global cerebral glucose utilization is decreased in olfactory bulbectomized rat models of depression (Skelin et al., 2008), and rats subjected to a chronic unpredictable mild stress model of depression exhibit insulin resistance in the arcuate nucleus of hypothalamus (Pan et al., 2013). Some data do not support the hypothesis of reduced cerebral glucose metabolism in animal models of depression. For example, the Flinders Sensitive Line rat, a well-validated model of depression, exhibits increased glucose utilization in brain structures important in the pathogenesis of depression (Kanemaru and Diksic, 2009). Additionally, our previous studies demonstrated that glucose, glycogen and glucose transporter (GLUT-1 and GLUT-4) concentrations are higher in the hippocampus and frontal cortex in prenatally stressed rats when compared with control animals (Detka et al., 2014). These data and the results obtained with Flinders Sensitive Line rats indicate that brain metabolic changes in animal models of depression are complex and various stages of glucose metabolism (uptake, glycolysis, the Krebs cycle, and oxidative phosphorylation) in particular brain regions can be differentially disturbed. The aim of the present study was to determine if high glucose and glycogen levels in the hippocampus and frontal cortex in a prenatal stress model of depression result from only the intensity of glucose uptake or if this change requires a weakening of glycolysis and/or the Krebs cycle. Thus, we examined key glycolytic enzymes (hexokinase, phosphofructokinase and pyruvate kinase), products of glycolysis, i.e., pyruvate and lactate, and two selected enzymes of the Krebs cycle (pyruvate dehydrogenase and αketoglutarate dehydrogenase) in the hippocampus and frontal cortex, i.e., brain structures important for depression, using a prenatal stress model of depression. In addition, glucose-6phosphate dehydrogenase was examined because glucose metabolism in the pentose cycle is a major source of NADPH that can be used to remove excess of reactive oxygen species while oxidative stress plays a role in the pathogenesis of depression. Previously observed changes in glucose, glycogen and glucose transporters were stronger after exposure of prenatally stressed rats to adverse factors (acute stress, glucose load) in adulthood (Detka et al., 2014). For the current study, we assessed the concentrations of selected markers involved in glucose metabolism at basal conditions and in animals subjected to stress or administered glucose as adults. We chose to study this prenatal stress model of depression because the face, predictive and construct validities of this model are well defined and in addition to depression-like behavioral changes, hyperactivity of the HPA axis and increases in blood glucose levels have been described in this model (Koehl et al., 1999; Morley-Fletcher et al., 2003, 2004; Szymańska et al., 2009a). Additionally changes induced by prenatal stress appear gradually 4
and are present in adult animals, which allows for the determination of progressive responses to adverse factors. Our previous studies using this model showed also more potent glucocorticoid action in the brain (connected with changes in factors that regulate glucocorticoid receptor function), increased concentrations of glucose, glycogen, glucose transporters and corticosterone in the brain, and insulin resistance (Szymańska et al., 2009a, 2009b; Budziszewska et al., 2010; Detka et al., 2014).
EXPERIMENTAL PROCEDURES Animals Sprague-Dawley rats (200-250 g) were purchased from a licensed dealer. Rats were kept in an animal housing facility at a room temperature of 23°C and 12/12 h light/dark cycle (lights on at 07 a.m.) with food and water available ad libitum. All experiments were carried out according to the National Institutes of Health Guide for the Care and Use of Laboratory Animals and were approved by the Local Ethics Committee in Kraków, Poland. All efforts were made to minimize the number of animals used and their suffering. One week after arrival, vaginal smears from females were collected daily in order to determine estrous cycle phase. During proestrus, rats were placed with males for 12 h. Next, we tested vaginal smears for the presence of sperm. Pregnant females were than randomly assigned to the control and stress groups. Stress procedure Prenatal stress was performed as previously described by Morley-Fletcher et al. (2003, 2004). Briefly, pregnant rats were subjected daily to three stress sessions starting at 09.00, 12.00 and 17.00 h. During this time, rats were placed in plastic cylinders (7/12 cm) and exposed to a bright light for 45 min. Stress sessions were performed from day 14 of pregnancy until delivery. Control pregnant females were left undisturbed in their home cages. Twenty-one days after birth, male offspring from litters containing 10-14 pups with a comparable number of males and females were separated for experiments. One to 2 animals from each litter were randomly assigned to 1 of 8 designated experimental groups. Rats were housed in groups of five per cage under standard conditions until 3 months of age. Forced swimming test in rats The 3-month-old animals were individually subjected to two trials in which they were forced to swim in a cylinder (40 cm high, 18 cm in diameter) filled with water (25o C) up to a height of 15 cm. There was a 24-hour interval between the first and second trial. The first trial lasted for 15 min, and the second trial lasted for 5 min. The total duration of immobility and climbing time were measured throughout the second trial (Porsolt et al., 1978). Acute immobilization stress in adult animals Half of the animals from the control and prenatally stressed groups were subjected to acute immobilization stress. The rats were placed in plastic cylinders (7/12 cm) for 1 h. Oral glucose administration 5
Immediately after acute stress, half of the animals from each control and prenatally stressed group with and without acute immobilization stress were administered glucose (1 g/kg) by oral gavage as previously described (Dias et al., 2010). The remaining animals received water in place of glucose. Glucose or water was given in a volume of 2 ml/kg. Tissue collection Two hours after glucose administration, animals were killed under non-stress conditions by rapid decapitation. Brains were rapidly removed, and brain regions (hippocampus and frontal cortex) were dissected on ice-cold glass plates. The tissues were frozen on dry ice and stored at -80°C. Scheme showing the time course of the experiment is shown in Figure 1.
Biochemical analysis Lactate estimation Lactate was measured in the frontal cortex and hippocampus using a lactate fluorometric assay kit (Biovision, USA) according to the manufacturer's instructions. Briefly, fresh tissues were homogenized in 5 volumes of phosphate-buffered saline (PBS) and centrifuged at 20000 g for 20 min at 4°C. The supernatants were then deproteinized using perchloric acid (PCA) and a Deproteinizing Sample Detection Kit (Biovision, USA) according to the procedure provided by the manufacturer. Briefly, 250 µl of sample homogenates were mixed with 50 µl of ice-cold PCA, incubated on ice for 5 minutes, and then centrifuged at 20 000 g for 2 min at 4 °C. A 50 µl aliquot of each sample was transferred to 96-well plates in duplicate along with lactate standards (0; 0.2; 0.4; 0.6; 0.8, and 1.0 nmol/well). A total of 50 µl of reaction mix (46 µl lactate assay buffer, 2 µl probes, and 2 µl of enzyme mix) was added to each test sample and lactate standard. Plates were incubated for 30 min at room temperature. Fluorescence was measured at Ex/Em=535/590 in a fluorometer (Tecan Infinite 200 Pro). The lactate concentration in samples was calculated from a standard curve and expressed as nmol/mg protein. Pyruvate estimation Pyruvate was measured in the frontal cortex and hippocampus using a fluorometric pyruvate assay kit (Biovision, USA) according to the manufacturer's instructions. Briefly, fresh tissues were homogenized in 5 volumes of phosphate-buffered saline (PBS) and centrifuged at 20000 g for 20 min at 4°C. The supernatants were than deproteinized using perchloric acid (PCA) and a Deproteinizing Sample Detection Kit (Biovision, USA) according to the procedure provided by the manufacturer. Briefly, 250 µl of sample homogenates were mixed with 50 µl of ice-cold PCA, incubated on ice for 5 minutes, and then centrifuged at 20 000 g for 2 min at 4 °C. A 50 µl aliquot of each sample was transferred to 96-well plates in duplicate along with pyruvate standards (0; 0.2; 0.4; 0.6; 0.8 and 1.0 nmol/well). A total of 50 µl/well of reaction mix (47.6 µl pyruvate assay buffer, 0.4 µl pyruvate probes, and 2 µl enzyme mix) was added to each test sample and pyruvate standard. Plates were incubated for 30 min at room temperature. Fluorescence was measured at Ex/Em=535/590 in a fluorometer (Tecan Infinite 200 Pro). The pyruvate concentration in samples was calculated from a standard curve and expressed as nmol/mg protein.
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Hexokinase The concentration of hexokinase in the frontal cortex and hippocampus was determined using an ELISA method (Wuhan EIAab Science Co., Ltd, China) according to the manufacturer’s instructions. Briefly, the tissues were homogenized in phosphate-buffered saline (PBS), shaken in an ice bath for 30 minutes and then centrifuged at 20 000 g for 20 min at 4 °C. An aliquot of 100 µl of each sample was transferred to pre-coated 96-well ELISA plates in duplicate along with hexokinase standards (0; 0.156; 0.312; 0.625; 1.25; 2.5; 5 and 10 ng/ml). The absorbance was measured at a wavelength λ=450 nm using a spectrophotometer (MultiskanSpectrum, ThermoLabsystems). The hexokinase concentration in samples was calculated from a standard curve and expressed as ng/mg protein. Phosphofructokinase The concentration of phosphofructokinase in the frontal cortex and hippocampus was measured using an ELISA method (USCN Life Science Inc., USA) according to the manufacturer’s instructions. Briefly, the tissues were homogenized in phosphate-buffered saline (PBS), shaken in an ice bath for 30 minutes and then centrifuged at 20 000 g for 20 min at 4 °C. An aliquot of 100 µl of each sample was transferred to pre-coated 96-well ELISA plates in duplicate along with phosphofructokinase standards (0; 1.56; 3.12; 6.25; 12.5; 25; 50 and 100 ng/ml). The absorbance was measured at a wavelength λ=450 nm using a spectrophotometer (MultiskanSpectrum, ThermoLabsystems). The concentration of phosphofructokinase was calculated from a standard curve and expressed as ng/mg protein. Pyruvate kinase The concentration of pyruvate kinase in the frontal cortex and hippocampus was determined using an ELISA method (Wuhan EIAab Science Co., Ltd, China) according to the manufacturer’s instructions. Briefly, the tissues were homogenized in phosphate-buffered saline (PBS), shaken in an ice bath for 30 minutes and then centrifuged at 20 000 g for 20 min at 4 °C. An aliquot of 100 µl of each sample was transferred to pre-coated 96-well ELISA plates in duplicate along with phosphofructokinase standards (0; 1.25; 2.5; 5; 10; 20; 40 and 80 Units/ml). The absorbance was measured at a wavelength λ=450 nm using a spectrophotometer (MultiskanSpectrum, ThermoLabsystems). The pyruvate kinase concentration in the samples was calculated from a standard curve and expressed as Units/mg protein. Isolation of mitochondria Concentration of pyruvate dehydrogenase and of α-ketoglutarate dehydrogenase was determined in mitochondrial fractions. Isolation of crude mitochondrial fraction from the frontal cortex and hippocampus was performed accordingly to the procedure described by Wernicke et al. (2010). Briefly, brain tissues, kept on ice were homogenized in motor-driven Teflon-glass homogenizer (9 strokes, 600 rpm) in 4 volumes of homogenization buffer, containing (5 mol/l HEPES/NaOH, pH 7.4, 320 mmol/l sucrose and 1 mmol/l Na+/EDTA with the addition of 0.5% protease inhibitor cocktail (Sigma P8340). After centrifugation in 1 300 × g (4 minutes, 4°C) supernatants were collected. Additionally, to increase the yield, pellet was washed twice with homogenization buffer and centrifuged at 1500 × g (4 minutes, 4°C). In order to collect the mitochondria, supernatants were combined, and centrifuged at 17 000 × g for 12 min at 4°C. After discarding of supernatants, the obtained pellets containing mitochondria were rapidly frozen and stored at –80°C. 7
Pyruvate dehydrogenase The concentration of pyruvate dehydrogenase 1 in the mitochondrial fraction of the frontal cortex and hippocampus was determined using an ELISA method (Wuhan EIAab Science Co., Ltd, China) according to the manufacturer’s instructions. An aliquot of 100 µl of each mitochondrial sample was transferred to pre-coated 96-well ELISA plates in duplicate along with phosphofructokinase standards (0; 0,312; 0.625; 1.25; 2.5; 5; 10; 20 and 40 ng/ml). The absorbance was measured at a wavelength λ=450 nm using a spectrophotometer (MultiskanSpectrum, ThermoLabsystems). The pyruvate dehydrogenase concentration in the samples was calculated from a standard curve and expressed as ng/mg protein. α-Ketoglutarate dehydrogenase The concentration of α-ketoglutarate dehydrogenase was measured in the mitochondrial fraction of the frontal cortex and hippocampus using a specific ELISA Kit (Wuhan EIAab Science Co., Ltd, China) according to the manufacturer’s instructions. An aliquot of 100 µl of each sample was transferred to pre-coated 96-well ELISA plates in duplicate along with αketoglutarate dehydrogenase standards (0; 0.075; 0.15; 0.31; 0.62; 1.25; 2.5 and 5 ng/ml). The absorbance was measured at a wavelength λ=450 nm using a spectrophotometer (MultiskanSpectrum, ThermoLabsystems). The concentration of α-ketoglutarate dehydrogenase in samples was calculated from a standard curve and expressed as ng/mg protein. Glucose-6-phosphate dehydrogenase activity The activity of glucose-6-phosphate dehydrogenase in the frontal cortex and hippocampus was determined using a colorimetric assay kit (TrinityBiotech, Ireland) according to the instructions provided by the manufacturer. A 10 µl aliquot of each sample was transferred in duplicate in 96-well plates and mixed with 50 µl of the Assay Solution. A total of 100 µl of the G-6-PDH Substrate Solution was added into sample wells. The absorbance was measured at a wavelength λ=340 nm using a MultiskanSpectrum spectrophotometer (ThermoLabsystems). The absorbance was measured immediately after addition of the substrate and after 5 minutes of incubation in order to determine enzymatic reaction kinetics. The activity of glucose-6-phosphate dehydrogenase was calculated and expressed as enzyme units/5 minutes/mg protein. Protein determination The protein content of the analyzed samples was determined using the method of Lowry et al. (1951). Statistical Analysis The data from the eight experimental groups, each containing 10 animals, are presented as the mean ± SEM, and the results were analyzed using the STATISTICA program. The homogeneity of the variance was first analyzed with a Levene’s test. The subsequent statistical analyses were performed by factorial analyses of variance (ANOVAs) to determine the effects of three factors (i.e., factor 1- prenatal stress, factor 2 - acute stress and factor 3 glucose). Next, Duncan’s post-hoc test was applied. A p-value of
