Oxidative stress accompanies HIF1-dependent impairment of glucose metabolism in the hippocampus of adult rats survived prenatal severe hypoxia

doi:10.21203/rs.3.rs-3248640/v1

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Oxidative stress accompanies HIF1-dependent impairment of glucose metabolism in the hippocampus of adult rats survived prenatal severe hypoxia

https://doi.org/10.21203/rs.3.rs-3248640/v1

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Many socially significant diseases are associated with disorders of prenatal development. Previously, we have shown the pathological role of hypoxia inducible factor HIF1 in post-hypoxic reoxygenation. This study aims to investigate the effect of prenatal severe hypoxia (PSH) on HIF1α protein expression as well as on HIF1-dependent activity of the pentose phosphate pathway (PPP) and anaerobic glycolysis in the hippocampus (HPC) of the offspring reached adulthood.

We showed that PSH causes a stable increase in the content of HIF1α protein in the HPC which was accompanied by an increase in the efficacy of anaerobic glycolysis. This was testified by increased LDH activity and lactate concentration. At the same time, the amounts of G6PD, NADPH and also reduced glutathione decreased in the HPC of PSH rats, whereas the concentration of an oxidative stress marker, MDA, exceeded the control values.

In a series of experiments using the model of emotional stress "learned helplessness" or the model of severe hypoxic stress, it was shown that in the HPC of control rats there was an increase in the amount of HIF1α in response to stress, which was also accompanied by more efficient anaerobic glycolysis and decreased efficacy of the PPP similar to the intact PSH rats. In the PSH rats, in turn, emotional stress resulted even in higher HIF1α levels without affecting glycolysis and PPP.

Therefore, the increased content and activity of the transcription factor HIF1α in the HPC of adult rats exposed to prenatal hypoxia leads to the imbalance between glycolysis and the PPP which is accompanied by oxidative stress.

prenatal hypoxia

HIF1

glycolysis

pentose phosphate pathway

oxidative stress

Hypoxia is defined as the condition of reduced oxygen supply and can appear at either whole-organism or tissue- and cellular-level. A violation in oxygen supply underlies the pathogenesis of many socially significant diseases [1, 2]. In particular, brain strokes and diseases of the cardiovascular system are the most dangerous pathologies associated with hypoxic episodes within postnatal development [3–6]. Meanwhile, malfunctions of oxygen supply are also extremely dangerous in the process of fetal development and in the early postnatal period [7–13]. So, neurodevelopmental disorders caused by hypoxia and asphyxia occur in about 2% of term infants and almost 60% of premature infants [14, 15]. The consequences of perinatal hypoxia may be revealed in infants remaining invisible in the process of growing up, and manifest only in the early or late adulthood [9, 16–18]. Perinatal hypoxia results in variety of effects depending on the stage of development, in which the exposure occurs [7, 19, 20]. So, the episodes of prenatal hypoxia in different periods of prenatal development may subsequently cause schizophrenia, depressive-like disorders, addictions, autism and also neurodegenerative diseases [19, 21–25].

Knowledge of the patterns of brain development in humans and rodents makes it possible to model the pathologies of prenatal development in vivo. We previously showed that a fetus, upon glucocorticoid stimulation caused by maternal body reaction to hypoxia exposure on the 14th-16th days of embryogenesis, demonstrated significant rearrangements of the glucocorticoid system activity in the extrahypothalamic structures of the brain. It led to disruption of the glucocorticoid negative feedback in the offspring and predetermining the development of a chronic depressive-like state associated with reduced plasticity of reactions to external stressors [26, 27]. Whereas in a mature organism damaging effects lead the brain structures to malfunction, the pathological effects in the prenatal period cause perturbations in epigenetic tuning of the signaling systems of a developing organism [9, 21, 28, 29]. However, it is still not completely clear whether the changes observed during pregnancy associated with hypoxia/ischemia, are due to the hypoxic factor itself or they are merely the outcomes of a nonspecific stress response of a mother.

One of the main regulators of the molecular response to hypoxia is hypoxia-inducible factor-1 (HIF1) [1, 30]. This transcription factor controls the expression of target genes involved in vascularization (e.g., vascular endothelial growth factor - VEGF) and glucose metabolism (glucose transporter 1 - GLUT1, almost all glycolysis enzymes and lactate dehydrogenase) [30–34]. It also regulates many other genes whose expression is required in chronic hypoxia [33, 35]. However, in the studies on posthypoxic reoxygenation, it was shown that HIF1 is able to have a maladaptive effect, disrupting the integrity of the blood-brain barrier, to cause cerebral edema, and triggering apoptosis [36–39]. In particular, in experiments on rodents, we have shown the participation of this transcription factor in the processes of post-hypoxic cell death due to inhibition of the pentose phosphate pathway of glucose metabolism (PPP) and resulting oxidative stress [40]. We also proved the existence of a mechanism of HIF1-dependent suppression of PPP in human cells [41]. In addition, like other researchers we have shown that, in the models of prenatal hypoxia, HIF1α expression increases after birth, which if maintained throughout life, can cause significant disturbances in brain energy metabolism [10, 42].

Therefore, this study was aimed at a comparative analysis of HIF1α protein expression, HIF1-dependent activity of anaerobic glycolysis and the pentose phosphate pathway, as well as the ratio of pro- and antioxidant processes in the hippocampus (HPC) of adult intact control and prenatally hypoxic rats both under normal conditions, as well as in response to damaging hypoxia (a model of severe hypobaric hypoxia, SH) or psycho-emotional stress (“learned helplessness” model, LH), which are widely described factors to increase HIF1α protein expression and HIF1-dependent metabolism as well [40, 43].

Animals

Wistar rats from the Collective Usage Centre “Biocollection of laboratory mammals of different taxonomic affiliation” of the Pavlov Institute of Physiology of RAS were used in experiments. Experimental procedures were performed in compliance with The Guidelines for Reporting Animal Research [44] and approved by the Ethical Committee for the Use of Animal Subjects at the Pavlov Institute of Physiology.

Pups were weaned from their mothers at the age of 30 days. After weaning, the rats were housed in cells 60 × 30 × 20 cm in size, with 6 animals in each. Each rat group consisted of randomly selected rats born by different dams to minimize litter bias. Rats received food and water ad libitum and were kept on a 12:12-h dark: light cycle at room temperature with a constant humidity of approximately 60%. All animal experiments were performed between 9 and 11 a.m.

During all the experiments, the allocation of the groups was blinded to the researchers. Animals were identified by random numbers, which were revealed to the researchers only after completing the experiments and data analysis.

Prenatal severe hypobaric hypoxia

Prenatal severe hypobaric hypoxia (PSH), reported in our previous studies, was used as a reliable model of maternal stress response during pregnancy [18, 22, 26, 27, 42]. The animals used in the experiments were born either from naïve females (Control) or females exposed to three sessions of severe hypobaric hypoxia on the 14th, 15th, and 16th days of pregnancy (PSH group) (Fig. 1).

For creating PSH, we used a flow-type hypobaric chamber at a temperature of 20° to 25°C in which atmospheric pressure was gradually reduced to 180 Torr (5% of normobaric oxygen equivalent to 11,000 m above sea level) during 20 min. Pregnant dams were treated under such conditions for 3 h for 3 consecutive days (E14-E16) with an interval of 24 h between the sessions. The mortality rate in the hypobaric chamber was around 15%.

Intact control females were also placed in the hypobaric chamber for 3 h on the 14th, 15th, and 16th days of pregnancy, but there was no hypoxic exposure (sham procedure).

In further experiments, we used adult male offspring with active spermatogenesis from the Control and PSH groups at the age of 3 months.

Learned Helplessness

Our method was based on the standard paradigm commonly used to induce a mental state of learned helplessness, which provided a reliable animal model of depression [43, 45]. In order to induce learned helplessness (LH), 3-month-old control and PSH rats were exposed during 1 h to inescapable treatment by 60 asynchronous electric shocks (1 mA, 60 Hz, 15 s) in a small chamber with a conductive floor (Fig. 1). Intact control and PSH rats were also placed in the chamber without electrical stimulation for 1 h (sham procedure). Animals were treated under such conditions for 3 consecutive days (d1-d3, Fig. 1) with an interval of 24 h between the sessions.

Severe hypobaric hypoxia

One trial of severe hypobaric hypoxia (SH) (180 Torr, 5% of normobaric oxygen or 11 000 m above sea level, 3 hours) reported in our previous studies was used on PSH and control 3-month-old adult rats to model a severe hypoxic injury.

Sample preparation

To collect the brain samples for immunohistochemistry, 3-month-old intact control and PSH rats were sacrificed by a guillotine. The brain samples were submerged in a fixative solution (28 ml Fine Fix + 72 ml 96% ethanol, Milestone, Italy) for 24 hours following a standard histological protocol. Dehydration of the tissues was achieved by treatment with 70% ethanol for 1.5 hours, 96% ethanol: 100% isopropyl alcohol (80:20) for 3 hours and 100% isopropyl alcohol for 3 hours at 60°C. The samples were then immersed in liquid paraffin (twice, for 1 hour) at 56°C and sectioned. The 7 µm thick coronal sections of the brain were prepared using a rotary microtome (Reichert, Austria) at about − 2.80 mm from the bregma. Sections were mounted onto poly-L-lysine covered slides, deparaffinized in xylol (twice, for 5 min), rehydrated in alcohols (96% → 96% → 96% → 70% for 5 min in each solution) and un-masked by boiling them in the citrate buffer solution (pH 6.0).

To collect the brain samples for western blotting and colorimetric analysis, intact or stressed control and PSH rats were sacrificed by a guillotine seven days after the last LH or sham procedure (d10, Fig. 1) or one day after SH or sham procedure (d2, Fig. 1). After decapitation samples of the hippocampus (HPC) were dissected and frozen in liquid nitrogen.

Immunohistochemistry

The analysis of the protein expression levels of HIF1α in the CA1 HPC region was performed by immunohistochemistry.

After incubation in blocking serum (AK-5001-NB, Vectastain ABC kit, Vector Laboratories, USA), brain sections were incubated overnight at 4°C with primary rabbit polyclonal antibodies against HIF1α (1:50, AF1009, Affinity Biosciences, USA). After several washes with phosphate buffered saline (0.1 M PBS, pH 7.4), the sections were incubated with biotinylated secondary antibodies (BA-9200 anti-mouse antibodies for synaptophysin or AK-5001-NB, Vectastain ABC kit with anti-rabbit antibodies for GFAP, Vector Laboratories, USA) and the ABC complex for 30 min each. Diaminobenzidine (DAB substrate kit for peroxidase, Vector Laboratories, USA) was used as a chromogen. The sections were dehydrated, mounted, and assessed by an image analysis system consisting of a light microscope (Jeneval, Carl Zeiss) and digital camera (Baumer CX05c, Baumer Optronic). The average optical density of HIF1α-positive cells was quantified in a length of 400 µm in the CA1 HPC region with ImageJ software. Four slices from each brain were analyzed and averaged, with one field per slice being measured. All parameters were expressed as % of levels in control rats.

Western blotting

To obtain total protein extracts for western blotting, the HPC samples were homogenized in 50 mM Tris-HCl (TBS, pH 8.0) containing 150 mM NaCl, 1% Triton X100 and a cocktail of protease and phosphatase inhibitors (SB-G2006, SB-G2007, Servicebio, China). Homogenates were incubated on a shaker for 30 min at + 4°C, centrifuged for 10 minutes at 14 000g, and supernatants were collected. Total protein content in the samples was measured using Pierce^™ Rapid Gold BCA Protein Assay Kit (Thermo Scientific, USA) following the manufacturer’s protocol. Samples, containing equal amounts of total protein were boiled for 10 minutes at + 70°C with a 3x Laemmli buffer.

The proteins were separated by SDS-PAGE (sodium dodecyl sulfate-polyacrylamide gel electrophoresis) and transferred to the PVDF membranes (Thermo Scientific, USA). After blocking for 1 h in PBS with 5% skimmed milk, the membranes were incubated in PBS with rabbit anti-HIF1α (1:2000, AF1009, Affinity Biosciences, USA), anti-G6PD (1:2000, DF6444, Affinity Biosciences, USA), anti-β-Tubulin (1:5000, ab179513, Abcam, UK) primary antibodies for 2 h at room temperature.

The membranes were then washed three times with PBST (TBS with 0.1% Tween 20) and incubated in PBS with HRP-conjugated anti-rabbit (1:5000, E-AB-1003, Elabscience, USA) secondary antibodies for 1 h at room temperature. The membranes were then washed twice with PBST. Immunoreactive protein bands were visualized by a Clarity ECL chemiluminescence kit (Bio-Rad, USA) with the ChemiScope 6000 Imaging System (Clinx Science Instruments, China).

The protein levels were quantified using the ImageJ software (NIH, Bethesda, MD, USA) and normalized to β-Tubulin.

Measurement of lactate dehydrogenase activity

LDH activity was analyzed using a colorimetric assay (E-BC-K046-M, Elabscience, USA). Dissected HPC samples were washed and homogenized with PBS (0.01 M, pH 7.4) at + 4℃ and centrifuged at 10 000g for 10 min to isolate the cytosolic proteins. Assay procedures were conducted following the manufacture’s protocol and the absorbance measured at 450 nm with a spectrophotometric microplate reader (CLARIOstar PLUS, BMG Labtech, Germany). The amount of pyruvate generated in the reaction was determined using a pyruvate standard curve. LDH activity was calculated as pmoles of pyruvate generated per minute per mg of total protein. Total protein in the samples was measured using a Pierce Rapid Gold Bicinchoninic Acid Protein Assay Kit (Thermo Fisher Scientific, Waltham, MA, USA) according to the manufacturer’s protocol.

Measurement of lactate levels

Lactate levels were analyzed using a colorimetric assay (E-BC-K044-M, Elabscience, USA). Dissected HPC samples were washed and homogenized with PBS (0.01 M, pH 7.4) at + 4℃ and centrifuged at 10 000g for 10 min to isolate the lactate containing supernatant. Assay procedures were conducted following the manufacturer’s protocol and the absorbance measured at 530 nm with a spectrophotometric microplate reader (CLARIOstar PLUS, BMG Labtech, Germany). The amount of lactate was quantified using a standard curve, calculated and expressed as nmol of lactate per mg of total protein. Total protein in the samples was measured using a Pierce Rapid Gold Bicinchoninic Acid Protein Assay Kit (Thermo Fisher Scientific, Waltham, MA, USA) according to the manufacturer’s protocol.

Measurement of NADPH levels

NADPH/NADP total ratio was analyzed using a colorimetric assay (E-BC-K803-M, Elabscience, USA) in accordance with manufacture protocol. Dissected HPC samples were homogenized with the extraction buffer and centrifuged at 12 000g for 10 minutes to isolate the NADPH/NADP⁺ containing supernatants. For NADPH only detection supernatants were additionally heated at 60°C for 30 min to decompose NADP⁺, cooled on ice and spun at 10 000g for 10 minutes to remove the precipitate. Assay procedures were conducted following the manufacturer’s protocol and the absorbance measured at 450 nm with a spectrophotometric microplate reader (CLARIOstar PLUS, BMG Labtech, Germany). The amount of NADPH or NADP total was quantified using a standard curve, calculated as pmol per mg of total protein and expressed as NADH/NADP total ratio. Total protein in the samples was measured using a Pierce Rapid Gold Bicinchoninic Acid Protein Assay Kit (Thermo Fisher Scientific, Waltham, MA, USA) according to the manufacturer’s protocol.

Measurement of reduced glutathione levels

Reduced glutathione levels were analyzed using a colorimetric assay (E-BC-K030-M, Elabscience, USA). Dissected HPC samples were washed with PBS (0.01 M, pH 7.4) and homogenized in 50 mM Tris-HCl (pH 7.4) containing 150 mM NaCl, 1mM EDTA, 1% Triton X100 at + 4℃ and centrifuged at 10 000g for 10 min to isolate the GSH containing supernatant. Assay procedures were conducted following the manufacturer’s protocol and the absorbance measured at 405 nm with a spectrophotometric microplate reader (CLARIOstar PLUS, BMG Labtech, Germany). The amount of GSH^red was quantified using a standard curve, calculated and expressed as nmol of GSH^red per mg of total protein. Total protein in the samples was measured using a Pierce Rapid Gold Bicinchoninic Acid Protein Assay Kit (Thermo Fisher Scientific, Waltham, MA, USA) according to the manufacturer’s protocol.

Measurement of malonic dialdehyde levels

Malonic dialdehyde levels were analyzed using a colorimetric assay (E-BC-K025-M, Elabscience, USA) in accordance with manufacturer’s protocol. Dissected HPC samples were washed and homogenized with PBS (0.01 M, pH 7.4) at + 4℃ and centrifuged at 10 000g for 10 min to isolate the MDA containing supernatant. Assay procedure were conducted following the manufacturer’s protocol and the absorbance measured at 532 nm with a spectrophotometric microplate reader (CLARIOstar PLUS, BMG Labtech, Germany). The amount of MDA was quantified using a standard curve, calculated and expressed as nmol of MDA per mg of total protein. Total protein in the samples was measured using a Pierce Rapid Gold Bicinchoninic Acid Protein Assay Kit (Thermo Fisher Scientific, Waltham, MA, USA) according to the manufacturer’s protocol.

Statistical analysis

Statistical analysis was performed using Prism 9.4 (GraphPad, Inc) software. Evaluation of the statistical significance was performed using the nonparametric Kruskal-Wallis test. The results were considered as significant at p < 0.05. As a post-hoc procedure, the Dunn’s test was used at a significance level of α = 0.05. The results were expressed as mean ± standard error of the mean (SEM). For Western blotting mean and SEM were then recalculated as % of the control taken as 100%.

The effects of prenatal hypoxia on the expression of HIF1α and HIF1 -dependent anaerobic glycolysis in the HPC in normal adulthood and after learned helplessness or severe hypoxia

In the hippocampal CA1 region of intact PSH rats we observed an increase in the HIF1α protein levels in compare with control animals (Fig. 2a, 2b). The results of immunohistochemistry additionally showed that in the CA1 region of the HPС of PSH rats, the localization of increased HIF1α protein expression was predominantly neuronal, whereas in control animals HIF1α mainly could be detected in astrocytes (Fig. 2b).

Western blotting results confirmed that in the HPC of intact PSH rats HIF1α protein levels increased in comparison with control animals (Fig. 3a, 3c, 3d). Upregulation of the HIF1α protein expression in the HPC of intact PSH rats was accompanied by an increase in both the LDH activity (Fig. 4a) and lactate levels (Fig. 4b) in comparison with intact control.

In the HPC of both control and PSH rats in response to LH there was an increase in the HIF1α protein levels in comparison with the intact control and PSH (Fig. 3a, 3c). An upregulation of the HIF1α protein expression in the HPC of control rats after LH was accompanied by an increase in the lactate levels (Fig. 4b), but not in the LDH activity, compared with the intact control. In the HPC of PSH rats LH did not affect LDH activity (Fig. 4a) or lactate levels (Fig. 4b).

We also found increased amount of HIF1α in the HPC of control and PSH rats after SH treatment in comparison with intact control and PSH (Fig. 3a, 3d). Moreover, an increase in the HIF1α protein levels in the HPC of PSH + SH rats was also observed in comparison with control + SH. In the control + SH group upregulation of the HIF1α protein expression in the HPC was accompanied by an increase in both the LDH activity (Fig. 4a) and lactate levels (Fig. 4b) compared to the intact control. In the HPC of PSH rats SH did not affect the LDH activity (Fig. 4a) and the lactate levels (Fig. 4b) in comparison with the intact PSH.

The effects of prenatal hypoxia on the activity of the pentose phosphate pathway in the HPC in normal adulthood and after learned helplessness or severe hypoxia

In the HPC of intact PSH rats we observed a decrease in the G6PD protein levels in comparison with control animals (Fig. 3b, 3c, 3d). Downregulation of the G6PD protein expression in the HPC of PSH rats was accompanied by a decrease in the NADPH/NADP total ratio (Fig. 5a).

The HPC of both control and PSH rats showed a decrease in the G6PD protein levels in response to LH compared to the intact control, but did not in comparison with the intact PSH (Fig. 3b, 3c). There was an increase in the G6PD protein levels in the HPC of both control and PSH rats in response to SH compared to the intact control (for control + SH) and PSH (for control + SH and PSH + SH) (Fig. 3b, 3d). After both LH and SH in the HPC of control and PSH rats we observed a decrease in NADPH/NADP total ratio (Fig. 5a) compared to intact control, but not PSH.

The effects of prenatal hypoxia on the redox state in the HPC in normal adulthood and after learned helplessness or severe hypoxia

We found that impairment in PPP efficiency (Fig. 3b, 3c, 3d, 5a) in the HPC of PSH rats was accompanied by a decrease in the GSH^red levels (Fig. 5b) and an increase in the lipid peroxidation detected by the amount of MDA (5c) in comparison with intact control. Both LH and SH decreased the hippocampal GSH^red levels (Fig. 5b), but did not increase lipid peroxidation levels (Fig. 5c), compared to the intact control. After LH in the HPC of PSH rats we found no changes in both the GSH^red (Fig. 5b) and MDA levels (Fig. 5c) in comparison with the intact PSH rats. There were no changes in GSH^red levels in the HPC of PSH rats after SH (Fig, 5b) in comparing to intact PSH, although MDA levels were decreased (Fig, 5c).

Oxygen is critical for life of aerobic organisms. Deviations in oxygen concentration significantly reduce an efficiency of energy metabolism and limit normal functioning causing death [46, 47]. Therefore, the signaling aimed at adaptation to changes of oxygen accessibility is among the most important cellular processes [1, 30–32, 35]. The HIF1 transcription factor, which is a heterodimer of the HIF1α and HIF1β proteins, is well known as a key factor in adaptation to oxygen deficiency [1, 34, 35]. HIF1α, being a regulated subunit, accumulates under hypoxic conditions, while HIF1β is expressed constitutively [30–32]. In addition to the prolyl hydroxylase associated activity, there are oxygen-independent pathways of stabilizing HIF1α, as well as mechanisms of regulation at the transcriptional level [34, 48–50]. HIF1 is reasonably recognized as a master regulator of adaptation to hypoxia affecting the expression of vascular endothelial growth factor, glucose transporters, anaerobic glycolysis enzymes, including LDH, erythropoietin cytokine, and products of many other genes, which are necessary for functioning in conditions of chronic hypoxia [30–35].

Hypoxia is among the most damaging stressors for the developing fetal brain [8, 9, 13, 16]. Epigenetic programming during fetal development is a very important and complexly organized process that determines the balance of gene expression specific to cells and tissues and determines a plasticity during their life and adaptation to permanently changing environmental conditions [28, 51]. The impact of stressors (e.g., hypoxic stress) during the period of HPC maturation not only has a direct damaging effect on a fetus [25, 52], but also can cause intracellular processes, which are stable throughout life, associated with a steady change in the patterns of epigenetic modifications [18, 26, 53]. A significant increase in the levels of HIF1α protein in the brain of a 21-day-old fetus was previously shown on a model of antenatal hypoxia [10], which indicated a hypoxic condition in the fetal brain tissues. Our data show that prenatal hypoxia causes a prolonged increase in the content and activity of HIF1α, which persists not only in newborn animals [42] but also in adults.

The role of astrocytic expression of HIF1α under normoxia in the organization of energy metabolism cooperation between neurons and astrocytes is well known [54–56]. Thus, astrocytes differ in terms of mitochondrial metabolism and a high rate of glycolysis compared to neurons [57, 58]. Most of the glucose entering the glycolytic pathway in astrocytes is subsequently released into the extracellular space in the form of lactate and sent to neurons [58, 59]. The glycolytic nature of astrocytes and their preference for lactate production and secretion is determined by HIF1-dependent gene expression [54–59]. However, we have shown that the increase in the HIF1α content in the HPC of adult rats that survived prenatal hypoxia occurs predominantly in neuronal cells. Taking into account the highest importance of the adequate expression of this transcription factor to ensure the efficient functioning of the energy metabolism of the brain, the deviations in HIF1α expression patterns can affect gene expression and lead to disorders of the nervous system.

Our data on the LDH activity and the amount of lactate indicate that the increased expression of HIF1α in the HPC of PSH rats is accompanied by an intensification of anaerobic glycolysis. It is important to note that an increase in glucose consumption in neurons through the glycolytic pathway has a detrimental effect on the PPP, which is a source of NADPH, widely known as necessary to maintain the cellular antioxidant potential [60–62]. The delicate balance between these pathways is critical for maintaining both energy demands and providing antioxidant functions [62]. Induction of glycolysis in neuronal cells has been reported to induce a PPP deficiency-mediated state of oxidative stress and cell death [63].

In addition to limiting the effectiveness of PPP through the induction of glycolysis, HIF1α is able to directly inhibit PPP through a decrease in G6PD expression [40, 41, 64]. Indeed, in this study, we show that a stable increase in HIF1α expression in the HPC of rats that survived PSH is accompanied by both a decrease in the protein expression of G6PD and a decrease in the amount of the PPP product NADPH. Such a reprogramming of metabolism due to the activity of the PPP in the processes of antioxidant protection and anabolism as the source of NADPH, is an important homeostatic response during the period of prolonged tissue hypoxia, when there is a need to increase the efficiency of catabolism [33]. However, in the absence of oxygen deficiency, suppression of the PPP can be a serious pathogenetic factor [40, 62]. A decrease in the efficiency of the PPP mediates a wide range of adverse conditions, such as disruption of the work of mixed-function oxidases, a decrease in the efficiency of steroid hormones biosynthesis and a decrease in the rate of recovery of thioredoxins and glutathione, which are together reflected in both a defect in individual cell functions and in violation of the integrative regulation of the whole organism [61, 62, 65, 66]. Both up-regulated anaerobic glycolysis and decreased efficiency of the PPP caused by HIF1-associated perturbations in the HPC of PSH rats led the efficiency of glutathione reduction to fall, which was also accompanied by an increase in free lipid peroxidation compared to control rats.

It is interesting to note that the HIF1-associated rearrangements of metabolism in the HPC of PSH rats are characterized by low plasticity to external stressors, for which the effect of increased expression of HIF1α has been shown [40, 43]. Thus, in the HPC of control animals one day after an episode of severe hypoxia or 7 days after an episode of emotional stress (model of “learned helplessness”) there is an increase in the protein expression of HIF1α, which is accompanied by an increase in the activity of anaerobic glycolysis and a decrease in the efficiency of the PPP to values observed in intact PSH animals. At the same time, an additional increase in the amount of HIF1α in the HPC of PSH rats in response to stressors does not affect the activity of glycolysis and the PPP compared to intact PSH rats. Whereas in response to emotional stress the protein expression of G6PD in the HPC of control animals decreases to the level of intact and stressed PSH rats, hypoxic stress causes an increase in G6PD expression in both control and PSH, probably in an Nrf2 dependent way [67, 68].

It should be noted that the HIF1-associated changes of hippocampal energy metabolism in control animals were short-term after a stressful episode and not accompanied by an increased lipid peroxidation at each time point studied. In the PSH group, in contrast, a permanent imbalance between glycolysis and PPP in the HPC was the established readout. Nevertheless, PSH animals responded to severe hypoxia with a decrease in the amount of MDA. It allows us to consider the alterations in the genetic program, which occur after hypoxic stress in the prenatal period, as rather an adaptive conditioning to damaging environmental stimuli during postnatal life. But this adaptability comes at the cost of serious adverse long-term consequences, including premature brain aging [18, 26].

Thus, the results obtained suggest that the HIF1-dependent rearrangements of brain metabolism appear to play a significant role in the pathogenesis of prenatal hypoxia. Having survived an episode of severe hypoxia in the prenatal period, the adult rats demonstrate an increased content of the transcription factor HIF1α in the hippocampal neurons accompanied by a stable imbalance between glycolysis and the pentose phosphate pathway which results in development of oxidative stress.

G6PD - glucose-6-phosphate dehydrogenase

GSH^red – reduced glutathione

HPC - hippocampus

HIF1 - hypoxia-inducible factor-1

LDH – lactate dehydrogenase

LH – «learned helplessness» model of emotional stress

MDA – malonic dialdehyde

NADPH/NDAP⁺/NADP total – reduced/oxidized/total nicotinamide adenine dinucleotide phosphate

PPP - pentose phosphate pathway

PSH - prenatal severe hypoxia

SH - severe hypobaric hypoxia

Acknowledgments

The authors are deeply grateful to Elena Axenova and Elena Lavrova for their excellent technical assistance in experiments on animal models.

Author Contribution

OV – designed experiments; OV, VS, SP, ET – performed experiments; OV, VS – analyzed data; OV, VS, ET - wrote the manuscript. All authors read and approved the final manuscript.

Funding

The work has been supported by the Ministry of Education and Science of the Russian Federation (Agreement no. 075-15-2020-921 of 13.11.2020) in the framework of the Project of World-class research center Pavlov Center “Integrative Physiology to Medicine, High-Tech Healthcare and Technologies of Stress Resistance”, section “Biological and Social Basis of Inclusion”.

Data Availability

The data that support the findings of this study are available from the corresponding author upon reasonable request.

Ethics Approval

Animal experiments were carried out according to The Guidelines for Reporting Animal Research [44]. Experimental protocols were approved by the local ethical committee at the Pavlov Institute of Physiology.

Consent to participate

Not applicable.

Consent for publication

Not applicable.

Conflict of interest

The authors declare no conflict of interest.

Semenza GL (2014) Oxygen sensing, hypoxia-inducible factors, and disease pathophysiology. Annu Rev Pathol 9:47–71. 10.1146/annurev-pathol-012513-104720
Darby IA, Hewitson TD (2016) Hypoxia in tissue repair and fibrosis. Cell Tissue Res 365(3):553–562. 10.1007/s00441-016-2461-3
Adav SS, Sze SK (2020) Hypoxia-Induced Degenerative Protein Modifications Associated with Aging and Age-Associated Disorders. Aging Dis 11(2):341–364. 10.14336/AD.2019.0604
Chen PS, Chiu WT, Hsu PL, Lin SC, Peng IC, Wang CY, Tsai SJ (2020) Pathophysiological implications of hypoxia in human diseases. J Biomed Sci 27(1):63. 10.1186/s12929-020-00658-7
Reed GW, Rossi JE, Cannon CP (2017) Acute myocardial infarction. Lancet 389(10065):197–210. 10.1016/S0140-6736(16)30677-8
Tuo QZ, Zhang ST, Lei P (2006) Mechanisms of neuronal cell death in ischemic stroke and their therapeutic implications. Med Res Rev 42(1):259–305. 10.1002/med.21817
Golan H, Huleihel M (2006) The effect of prenatal hypoxia on brain development: short- and long-term consequences demonstrated in rodent models. Dev Sci 9:338–349. 10.1111/j.1467-7687.2006.00498.x
Li Y, Gonzalez P, Zhang L (2012) Fetal stress and programming of hypoxic/ischemic-sensitive phenotype in the neonatal brain: mechanisms and possible interventions. Prog Neurobiol 98(2):145–165. 10.1016/j.pneurobio.2012.05.010
Xiong F, Zhang L (2013) Role of the hypothalamic–pituitary–adrenal axis in developmental programming of health and disease. Front Neuroendocrinol 34:27–46. 10.1016/j.yfrne.2012.11.002
Gonzalez-Rodriguez PJ, Xiong F, Li Y, Zhou J, Zhang L (2014) Fetal hypoxia increases vulnerability of hypoxic–ischemic brain injury in neonatal rats: Role of glucocorticoid receptors. Neurobiol Dis 65:172–179. 10.1016/j.nbd.2014.01.020
Giussani DA (2016) The fetal brain sparing response to hypoxia: physiological mechanisms. J Physiol 594(5):1215–1230. 10.1113/JP271099
Millar LJ, Shi L, Hoerder-Suabedissen A, Molnár Z (2017) Neonatal Hypoxia Ischaemia: Mechanisms, Models, and Therapeutic Challenges. Front Cell Neurosci 8:11:78. 10.3389/fncel.2017.00078
Piešová M, Mach M (2020) Impact of perinatal hypoxia on the developing brain. Physiol Res 69(2):199–213. 10.33549/physiolres.934198
Graham EM, Ruis KA, Hartman AL et al (2008) A systematic review of the role of intrapartum hypoxia-ischemia in the causation of neonatal encephalopathy. Am J Obstet Gynecol 199:587–295. 10.1016/j.ajog.2008.06.094
Kurinczuk JJ, White-Koning M, Badawi N (2010) Epidemiology of neonatal encephalopathy and hypoxic–ischaemic encephalopathy. Early Hum Dev 86:329–338. 10.1016/j.earlhumdev.2010.05.010
Warner MJ, Ozanne SE (2010) Mechanisms involved in the developmental programming of adulthood disease. Biochem J 427(3):333–347. 10.1042/BJ20091861
Langley-Evans SC, McMullen S (2010) Developmental origins of adult disease. Med Princ Pract 2010;19(2):87–98. doi: 10.1159/000273066
Vetrovoy O, Stratilov V, Nimiritsky P, Makarevich P, Tyulkova E (2021) Prenatal Hypoxia Induces Premature Aging Accompanied by Impaired Function of the Glutamatergic System in Rat Hippocampus. Neurochem Res 46(3):550–563. 10.1007/s11064-020-03191-z
Brandon A, Cui X, Luan W, Ali AA, Pertile RAN, Alexander SA, Eyles DW (2022) Prenatal hypoxia alters the early ontogeny of dopamine neurons. Transl Psychiatry 12(1):238. 10.1038/s41398-022-02005-w
Camm EJ, Cross CM, Kane AD, Tarry-Adkins JL, Ozanne SE, Giussani DA (2021) Maternal antioxidant treatment protects adult offspring against memory loss and hippocampal atrophy in a rodent model of developmental hypoxia. FASEB J 35(5):e21477. 10.1096/fj.202002557RR
Wang B, Zeng H, Liu J, Sun M (2021) Effects of Prenatal Hypoxia on Nervous System Development and Related Diseases. Front Neurosci 15:755554. 10.3389/fnins.2021.755554
Stratilov VA, Vetrovoy OV, Tyulkova EI (2022) Prenatal Hypoxia Affects Nicotine Consumption and Withdrawal in Adult Rats via Impairment of the Glutamate System in the Brain. Mol Neurobiol 59(7):4550–4561. 10.1007/s12035-022-02866-8
Kolevzon A, Gross R, Reichenberg A (2007) Prenatal and perinatal risk factors for autism: a review and integration of findings. Arch Pediatr Adolesc Med 161(4):326–333. 10.1001/archpedi.161.4.326
Howell KR, Pillai A (2014) Effects of prenatal hypoxia on schizophrenia-related phenotypes in heterozygous reeler mice: a gene × environment interaction study. Eur Neuropsychopharmacol 24(8):1324–1336. 10.1016/j.euroneuro.2014.05.011
Nalivaeva NN, Turner AJ, Zhuravin IA (2018) Role of Prenatal Hypoxia in Brain Development, Cognitive Functions, and Neurodegeneration. Front Neurosci 12:825. 10.3389/fnins.2018.00825
Vetrovoy O, Tyulkova E, Stratilov V, Baranova K, Nimiritsky P, Makarevich P, Rybnikova E (2021) Long-term effects of prenatal severe hypoxia on central and peripheral components of the glucocorticoid system in rats. Dev Neurosci 42:145–158. https://doi.org/10.1159/000512223
Vetrovoy O, Stratilov V, Lomert E, Tyulkova E (2023) Prenatal Hypoxia-Induced Adverse Reaction to Mild Stress is Associated with Depressive-Like Changes in the Glucocorticoid System of Rats. Neurochem Res 48(5):1455–1467. 10.1007/s11064-022-03837-0
Chen M, Zhang L (2011) Epigenetic mechanisms in developmental programming of adult disease. Drug Discovery Today 16:1007–1018. 10.1016/j.drudis.2011.09.008
Tyul’kova EI, Vataeva LA, Stratilov VA, Barysheva VS, Vetrovoy OV (2020) Peculiarities of DNA and Histone H3 Methylation in the Hippocampus and Neocortex of Rats Subjected to Pathological Treatments during the Prenatal Period. Neurochem J 14:64–72. 10.1134/S1819712420010195
Semenza GL (2011) Regulation of metabolism by hypoxia-inducible factor 1. Cold Spring Harb Symp Quant Biol 76:347–353. 10.1101/sqb.2011.76.010678
Semenza GL (2000) HIF-1 and human disease: one highly involved factor. Genes Dev 14:1983-91. PMID: 10950862
Semenza GL (2001) Hypoxia-inducible factor 1: oxygen homeostasis and disease pathophysiology. Trends Mol Med 7:345–350. 10.1016/s1471-4914(01)02090-1
Chavez J, Agani F, Pichiule P, LaManna J (2000) Expression of hypoxia-inducible factor-1alpha in the brain of rats during chronic hypoxia. J Appl Physiol 89(5):1937–1942. 10.1152/jappl.2000.89.5.1937
Dengler VL, Galbraith M, Espinosa JM (2014) Transcriptional regulation by hypoxia inducible factors. Crit Rev Biochem Mol Biol 49(1):1–15. 10.3109/10409238.2013.838205
Zepeda AB, Pessoa A Jr, Castillo RL, Figueroa CA, Pulgar VM, Farías JG (2013) Cellular and molecular mechanisms in the hypoxic tissue: role of HIF-1 and ROS. Cell Biochem Funct 31(6):451–459. 10.1002/cbf.2985
Yang XS, Yi TL, Zhang S, Xu ZW, Yu ZQ, Sun HT, Yang C, Tu Y, Cheng SX (2017) Hypoxia-inducible factor-1 alpha is involved in RIP-induced necroptosis caused by in vitro and in vivo ischemic brain injury. Sci Rep 7(1):5818. 10.1038/s41598-017-06088-0
Sun Y, Chen X, Zhang X, Shen X, Wang M, Wang X, Liu WC, Liu CF, Liu J, Liu W, Jin X (2017) β2-Adrenergic Receptor-Mediated HIF-1α Upregulation Mediates Blood Brain Barrier Damage in Acute Cerebral Ischemia. Front Mol Neurosci 10:257. 10.3389/fnmol.2017.00257
Chen C, Hu Q, Yan J, Yang X, Shi X, Lei J, Chen L, Huang H, Han J, Zhang JH, Zhou C (2009) Early inhibition of HIF-1alpha with small interfering RNA reduces ischemic-reperfused brain injury in rats. Neurobiol Dis 33(3):509–517. 10.1016/j.nbd.2008.12.010
Page S, Raut S, Al-Ahmad A (2019) Oxygen-Glucose Deprivation/Reoxygenation-Induced Barrier Disruption at the Human Blood-Brain Barrier is Partially Mediated Through the HIF-1 Pathway. Neuromolecular Med 21(4):414–431. 10.1007/s12017-019-08531-z
Vetrovoy O, Sarieva K, Lomert E, Nimiritsky P, Eschenko N, Galkina O, Lyanguzov A, Tyulkova E, Rybnikova E (2020) Pharmacological HIF1 Inhibition Eliminates Downregulation of the Pentose Phosphate Pathway and Prevents Neuronal Apoptosis in Rat Hippocampus Caused by Severe Hypoxia. J Mol Neurosci 70(5):635–646. 10.1007/s12031-019-01469-8
Vetrovoy OV, Nimiritsky PP, Tyulkova EI, Rybnikova EA (2021) Transcription Factor HIF1 Negatively Regulates the Content of Glucose-6-Phosphate Dehydrogenase in HEK293T Cells. Cell Tiss Biol 15:181–188. 10.1134/S1990519X21020127
Vetrovoy OV, Nimiritsky PP, Tyulkova EI, Rybnikova EA (2020) The Content and Activity of Hypoxia-Inducible Factor HIF1α Increased in the Hippocampus of Newborn Rats That Were Subjected to Prenatal Hypoxia on Days 14–16 of Embryogenesis. Neurochem J 14:286–289. 10.1134/S1819712420030125
Baranova KA, Mironova VI, Rybnikova EA, Samoilov MO (2010) Characteristics of the transcription factor HIF-1α expression in the rat brain during the development of a depressive state and the antidepressive effects of hypoxic preconditioning. Neurochem J 4:35–40. 10.1134/S1819712410010071
Kilkenny C, Browne WJ, Cuthill IC, Emerson M, Altman DG (2010) Improving bioscience research reporting: the ARRIVE guidelines for reporting animal research. PLoS Biol 8(6):e1000412. 10.1371/journal.pbio.1000412
Henkel V, Bussfeld P, Möller HJ, Hegerl U (2002) Cognitive-behavioral theories of helplessness/hopelessness: valid models of depression? Eur Arch Psychiatry Clin Neurosci 252(5):240–249. 10.1007/s00406-002-0389-y
Sendoel A, Hengartner MO (2014) Apoptotic cell death under hypoxia. Physiol (Bethesda) 29(3):168–176. 10.1152/physiol.00016.2013
Lenihan CR, Taylor CT (2013) The impact of hypoxia on cell death pathways. Biochem Soc Trans 41(2):657–663. 10.1042/BST20120345
Brocato J, Chervona Y, Costa M (2014) Molecular responses to hypoxia-inducible factor 1α and beyond. Mol Pharmacol 85(5):651–657. 10.1124/mol.113.089623
Vanderhaeghen T, Beyaert R, Libert C (2021) Bidirectional Crosstalk Between Hypoxia Inducible Factors and Glucocorticoid Signalling in Health and Disease. Front Immunol 12:684085. 10.3389/fimmu.2021.684085
Lee KE, Simon MC (2015) SnapShot: Hypoxia-Inducible Factors. Cell 19;163(5):1288–1288.e1. doi: 10.1016/j.cell.2015.11.011
Murgatroyd C, Spengler D (2011) Epigenetics of early child development. Front Psychiatry 2:16. 10.3389/fpsyt.2011.00016
Amakhin DV, Soboleva EB, Postnikova TY, Tumanova NL, Dubrovskaya NM, Kalinina DS, Vasilev DS, Zaitsev AV (2022) Maternal Hypoxia Increases the Excitability of Neurons in the Entorhinal Cortex and Dorsal Hippocampus of Rat Offspring. Front Neurosci 16:867120. 10.3389/fnins.2022.867120
Zhuravin IA, Dubrovskaya NM, Vasilev DS, Kozlova DI, Kochkina EG, Tumanova NL, Nalivaeva NN (2019) Regulation of Neprilysin Activity and Cognitive Functions in Rats After Prenatal Hypoxia. Neurochem Res 44(6):1387–1398. 10.1007/s11064-019-02796-3
Turner DA, Adamson DC (2011) Neuronal-astrocyte metabolic interactions: understanding the transition into abnormal astrocytoma metabolism. J Neuropathol Exp Neurol 70(3):167–176. 10.1097/NEN.0b013e31820e1152
Vicente-Gutiérrez C, Jiménez-Blasco D, Quintana-Cabrera R (2021) Intertwined ROS and Metabolic Signaling at the Neuron-Astrocyte Interface. Neurochem Res 46(1):23–33. 10.1007/s11064-020-02965-9
Fernandez AM, Martinez-Rachadell L, Navarrete M, Pose-Utrilla J, Davila JC, Pignatelli J, Diaz-Pacheco S, Guerra-Cantera S, Viedma-Moreno E, Palenzuela R, Ruiz de Martin Esteban S, Mostany R, Garcia-Caceres C, Tschöp M, Iglesias T, de Ceballos ML, Gutierrez A, Torres Aleman I (2022) Insulin regulates neurovascular coupling through astrocytes. Proc Natl Acad Sci USA 119(29):e2204527119. 10.1073/pnas.2204527119
Bonvento G, Bolaños JP (2021) Astrocyte-neuron metabolic cooperation shapes brain activity. Cell Metab 33(8):1546–1564. 10.1016/j.cmet.2021.07.006
Mächler P, Wyss MT, Elsayed M, Stobart J, Gutierrez R, von Faber-Castell A, Kaelin V, Zuend M, San Martín A, Romero-Gómez I, Baeza-Lehnert F, Lengacher S, Schneider BL, Aebischer P, Magistretti PJ, Barros LF, Weber B (2016) In Vivo Evidence for a Lactate Gradient from Astrocytes to Neurons. Cell Metab 23(1):94–102. 10.1016/j.cmet.2015.10.010
Bélanger M, Allaman I, Magistretti PJ Brain energy metabolism: focus on astrocyte-neuron metabolic cooperation. Cell Metab 14(6):724–738. doi: 10.1016/j.cmet.2011.08.016
Ulusu NN, Sahilli M, Avci A, Canbolat O, Ozansoy G, Ari N, Bali M, Stefek M, Stolc S, Gajdosik A, Karasu C (2003) Pentose phosphate pathway, glutathione-dependent enzymes and antioxidant defense during oxidative stress in diabetic rodent brain and peripheral organs: effects of stobadine and vitamin E. Neurochem Res 28(6):815–823. 10.1023/a:1023202805255
Fernandez-Fernandez S, Almeida A, Bolaños J (2012) Antioxidant and bioenergetic coupling between neurons and astrocytes. Biochem J 443:3–11. 10.1042/BJ20111943
Tang BL (2019) Neuroprotection by glucose-6-phosphate dehydrogenase and the pentose phosphate pathway. J Cell Biochem 120(9):14285–14295. 10.1002/jcb.29004
Herrero-Mendez A, Almeida A, Fernández E, Maestre C, Moncada S, Bolaños JP (2009) The bioenergetic and antioxidant status of neurons is controlled by continuous degradation of a key glycolytic enzyme by APC/C-Cdh1. Nat Cell Biol 11(6):747–752. 10.1038/ncb1881
Guo S, Miyake M, Liu KJ, Shi H (2009) Specific inhibition of hypoxia inducible factor 1 exaggerates cell injury induced by in vitro ischemia through deteriorating cellular redox environment. J Neurochem. 2009;108(5):1309-21. doi: 10.1111/j.1471-4159.2009.05877.x
Dhur A, Galan P, Hercberg S (1989) Effects of different degrees of iron deficiency on cytochrome P450 complex and pentose phosphate pathway dehydrogenases in the rat. J Nutr 119:40–47. 10.1093/jn/119.1.40
Stincone A, Prigione A, Cramer T, Wamelink MM, Campbell K, Cheung E, Olin-Sandoval V, Grüning NM, Krüger A, Tauqeer Alam M, Keller MA, Breitenbach M, Brindle KM, Rabinowitz JD, Ralser M (2015) The return of metabolism: biochemistry and physiology of the pentose phosphate pathway. Biol Rev Camb Philos Soc 90(3):927–963. 10.1111/brv.12140
Spagnuolo MS, Bergamo P, Crescenzo R, Iannotta L, Treppiccione L, Iossa S, Cigliano L (2020) Brain Nrf2 pathway, autophagy, and synaptic function proteins are modulated by a short-term fructose feeding in young and adult rats. Nutr Neurosci 23(4):309–320. 10.1080/1028415X.2018.1501532
Kovac S, Angelova PR, Holmström KM, Zhang Y, Dinkova-Kostova AT, Abramov AY (2015) Nrf2 regulates ROS production by mitochondria and NADPH oxidase. Biochim Biophys Acta 1850(4):794–801. 10.1016/j.bbagen.2014.11.021

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Oxidative stress accompanies HIF1-dependent impairment of glucose metabolism in the hippocampus of adult rats survived prenatal severe hypoxia

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Abstract

Figures

Introduction

Materials and Methods

Animals

Prenatal severe hypobaric hypoxia

Learned Helplessness

Severe hypobaric hypoxia

Sample preparation

Immunohistochemistry

Western blotting

Measurement of lactate dehydrogenase activity

Measurement of lactate levels

Measurement of NADPH levels

Measurement of reduced glutathione levels

Measurement of malonic dialdehyde levels

Statistical analysis

Results

Discussion

Abbreviations

Declarations

References

Additional Declarations

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