Oxidative Stress Accompanies HIF1-Dependent Impairment of Glucose Metabolism in the Hippocampus of Adult Rats That Survived Prenatal Severe Hypoxia

Hypoxia is defined as a condition of reduced oxygen supply and can appear at the whole organism, tissue, and cellular levels. Violation of the 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 during postnatal development [3‒6]. Meanwhile, malfunctions in oxygen supply are also extremely dangerous in the process of fetal development and in the early postnatal period [7‒13]. Neurodevelopmental disorders caused by hypoxia and asphyxia occur in approximately 2% of term infants and 60% of premature infants [14, 15]. The consequences of perinatal hypoxia may be revealed in infants who remain invisible in the process of growing up and manifest only in early or late adulthood [9, 16‒18]. Perinatal hypoxia results in a variety of effects depending on the stage of development in which exposure occurs [7, 19, 20]. Therefore, episodes of prenatal hypoxia during different periods of prenatal development may subsequently cause schizophrenia, depression-like disorders, addiction, autism, and neurodegenerative diseases [19, 21‒25].

Understanding the patterns of brain development in humans and rodents makes it possible to model the pathologies of prenatal development in vivo. In rats, 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 extra-hypothalamic structures of the brain during lifespan. This led to disruption of glucocorticoid-negative feedback in the offspring and predetermined the development of a chronic depressive-like state associated with reduced plasticity of reactions to external stressors [26, 27]. In mature organisms, damaging effects lead the brain structures to malfunction, and the pathological effects in the prenatal period cause perturbations in the 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 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) and glucose metabolism (glucose transporter 1, almost all glycolysis enzymes, and lactate dehydrogenase [LDH]) [30‒34]. It also regulates the expression of many other genes that are required during chronic hypoxia [33, 35]. However, studies on post-hypoxic reoxygenation have shown that HIF1 has a maladaptive effect, disrupting the integrity of the blood-brain barrier, causing cerebral edema, and triggering apoptosis [36‒39]. In particular, in experiments on rodents, we showed the participation of this transcription factor in the processes of post-hypoxic cell death due to inhibition of the pentose phosphate pathway (PPP) of glucose metabolism and the resulting oxidative stress [40]. We also demonstrated a mechanism for HIF1-dependent suppression of PPP in human cells [41]. In addition, similar to 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 aimed to perform a comparative analysis of HIF1α protein expression, HIF1-dependent activity of anaerobic glycolysis, and the PPP, 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 hypoxia, SH) or psycho-emotional stress (“learned helplessness” [LH] model), which are widely described factors that increase HIF1α protein expression and HIF1-dependent metabolism [40, 43].

Wistar rats from the Collective Usage Center “Biocollection of laboratory mammals of different taxonomic affiliation” of the Pavlov Institute of Physiology of RAS were used in experiments. The 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 (protocol no. 08/02 of August 02, 2022).

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

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

To create PSH, we used a flow-type hypobaric chamber at a temperature of 20°–25°C, in which atmospheric pressure was gradually reduced to 180 Torr (5% of normobaric oxygen equivalent to 11,000 m above sea level) for 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 sessions. The mortality rate in the hypobaric chamber is approximately 15%. Intact control females were also placed in the hypobaric chamber for 3 h on 14th, 15th, and 16th days of pregnancy; however, there was no hypoxic exposure (sham procedure).

The pups were weaned from their mothers at the age of 30 days. After weaning, the rats were housed in 60 × 30 × 20 cm cells, including 3 control and PSH rats. Each group of rats used in the experiment consisted of randomly selected rats born to different dams to minimize litter bias. The 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%. In further experiments, we used adult male offspring with active spermatogenesis from the control and PSH groups at the age of 3 months. All animal experiments were performed between 9:00 a.m. and 11:00 a.m.

The method was based on the standard paradigm commonly used to induce a depressive-like state of “learned helplessness” (LH), which provides a reliable animal model of depression [43, 45]. To induce LH, 3-month-old control and PSH rats were exposed for 1 h to inescapable treatment with 60 asynchronous electric shocks (1 mA, 60 Hz, 15 s) in a small chamber with a conductive floor (shown in Fig. 1). Animals were treated under such conditions for three consecutive days (d1–d3; shown in Fig. 1) with an interval of 24 h between sessions.

One trial of severe hypoxia (SH) (180 Torr, 5% normobaric oxygen or 11,000 m above sea level, 3 h) reported in our previous studies was used on PSH and control 3-month-old adult rats to model a severe hypoxic injury (shown in Fig. 1).

To collect brain samples for immunohistochemistry, 3-month-old intact control and PSH rats were sacrificed using guillotine. The brain samples were submerged in a fixative solution (28 mL Fine Fix + 72 mL 96% ethanol; Milestone, Italy) for 24 h following a standard histological protocol. Dehydration of the tissues was achieved by treatment with 70% ethanol for 1.5 h, 96% ethanol:100% isopropyl alcohol (80:20) for 3 h, and 100% isopropyl alcohol for 3 h at 60°C. The samples were then immersed in liquid paraffin (twice, for 1 h) at 56°C and sectioned. Coronal sections of the brain (7 µm thick) 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 alcohol (96% → 96% → 96% → 70% for 5 min in each solution), and unmasked by boiling them in 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 7 days after the last LH or sham procedure (d10; shown in Fig. 1) or 1 day after the SH or sham procedure (d2; shown in Fig. 1). After decapitation, samples of the HPC were dissected and frozen in liquid nitrogen.

The protein expression levels of HIF1α in the CA1 HPC region were analyzed 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 (PBS) (0.1 M PBS, pH 7.4), the sections were incubated with biotinylated secondary antibodies (1:200, AK-5001-NB, Vectastain ABC kit with anti-rabbit antibodies; Vector Laboratories, USA) and the ABC complex (1:100) for 30 min each. Diaminobenzidine (DAB Substrate Kit for Peroxidase; Vector Laboratories, USA) was used as the chromogen. The sections were dehydrated, mounted, and assessed using an image analysis system consisting of a light microscope (Jeneval; Carl Zeiss) and digital camera (Baumer CX05c; Baumer Optronic). The number of HIF1α-positive cells was quantified at a length of 400 μm in the CA1 HPC region using the ImageJ software. Four slices from each brain were analyzed and averaged, with one field measured per slice.

To obtain total protein extracts for Western blotting, 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 and centrifuged for 10 min at 14,000 g, and supernatants were collected. The total protein content in the samples was measured using the Pierce^TM Rapid Gold BCA Protein Assay Kit (Thermo Fisher Scientific, USA) following the manufacturer’s protocol. Samples containing equal amounts of total protein were boiled for 10 min at +70°C with a ×3 Laemmli buffer.

Proteins were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis and transferred to the PVDF membranes (Thermo Fisher Scientific, USA). After blocking for 1 h in PBS containing 5% skim milk, the membranes were incubated in PBS with rabbit anti-HIF1α (1:2,000, AF1009; Affinity Biosciences, USA), anti-glucose-6-phosphate dehydrogenase (G6PD) (1:2,000, DF6444; Affinity Biosciences, USA), and anti-β-Tubulin (1:5,000, ab179513; Abcam, UK) primary antibodies for 2 h at room temperature.

The membranes were then washed thrice with PBST (TBS with 0.1% Tween 20) and incubated in PBS with HRP-conjugated anti-rabbit (1:5,000, 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 using a Clarity ECL chemiluminescence kit (Bio-Rad, USA) with a ChemiScope 6000 Imaging System (Clinx Science Instruments, China). Protein levels were quantified using ImageJ software (NIH, Bethesda, MD, USA) and normalized to β-Tubulin. Full images of the Western blotting results are presented in the online supplementary materials 1 (for all online suppl. material, see https://doi.org/10.1159/000535326) (S1 for LH experiments; S2 for severe hypoxia experiments).

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°C and centrifuged at 10,000 g for 10 min to isolate the cytosolic proteins. The assay procedures were conducted following the manufacturer’s protocol, and the absorbance was measured at 450 nm using a spectrophotometric microplate reader (CLARIOstar PLUS; BMG LABTECH, Germany). The amount of pyruvate generated during the reaction was determined by using a standard pyruvate curve. LDH activity was calculated as pmol of pyruvate generated per minute per milligram of total protein. Here and in the other biochemical tests below, the 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.

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°C and centrifuged at 10,000 g for 10 min to isolate the lactate-containing supernatant. The assay procedures were conducted following the manufacturer’s protocol, and the absorbance was measured at 530 nm using a spectrophotometric microplate reader (CLARIOstar PLUS, BMG LABTECH, Germany). The amount of lactate was quantified using a standard curve, calculated, and expressed as nanomoles of lactate per milligram of total protein.

The NADPH/NADP total ratio was analyzed using a colorimetric assay (E-BC-K803-M; Elabscience, USA), in accordance with the manufacturer’s protocol. Dissected HPC samples were homogenized with extraction buffer and centrifuged at 12,000 g for 10 min 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 10,000 g for 10 min to remove the precipitate. The assay procedures were conducted following the manufacturer’s protocol, and the absorbance was measured at 450 nm using a spectrophotometric microplate reader (CLARIOstar PLUS; BMG LABTECH, Germany). The total amount of NADPH or NADP was quantified using a standard curve, calculated as pmol per mg of total protein, and expressed as NADH/NADP total ratio.

Reduced glutathione (GSHred) 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), homogenized in 50 mM Tris-HCl (pH 7.4) containing 150 mM NaCl, 1 mM EDTA, 1% Triton X100 at +4°C, and centrifuged at 10,000 g for 10 min to isolate the GSH-containing supernatant. The assay procedures were conducted following the manufacturer’s protocol, and the absorbance was measured at 405 nm using a spectrophotometric microplate reader (CLARIOstar PLUS; BMG LABTECH, Germany). The amount of GSHred was quantified using a standard curve, calculated, and expressed as nanomoles of GSHred per milligram of total protein.

Malonic dialdehyde (MDA) levels were analyzed using a colorimetric assay (E-BC-K025-M; Elabscience, USA) according to the manufacturer’s protocol. Dissected HPC samples were washed and homogenized with PBS (0.01 M, pH 7.4) at +4°C and centrifuged at 10,000 g for 10 min to isolate the MDA-containing supernatant. The assay procedures were conducted following the manufacturer’s protocol, and the absorbance was measured at 532 nm using a spectrophotometric microplate reader (CLARIOstar PLUS; BMG LABTECH, Germany). The amount of MDA was quantified using a standard curve, calculated, and expressed as nanomoles of MDA per milligram of total protein.

Statistical analysis was performed using Prism 10 (GraphPad, Inc.). For all quantification analyses, one-way ANOVA criterion was used, followed by Tukey’s honest significance test. Statistical significance was set at p <0.05. The results are expressed as mean ± standard error of the mean. For immunohistochemistry and Western blotting, the mean and standard error of the mean were recalculated as % of the control, taken as 100%.

In the hippocampal CA1 region of intact PSH rats, we observed an increase in HIF1α protein levels compared with control animals (shown in Fig. 2a, b; ANOVA F [1, 11] = 48.1139, p = 0.00004; control vs. PSH, p = 0.001, Tukey’s HST). The immunohistochemistry results 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 was mainly detected in astrocytes (shown in Fig. 2b).

Western blotting results confirmed that in the HPC of intact PSH rats, HIF1α protein levels were increased in comparison with control animals (shown in Fig. 2c, e, f, h; ANOVA F [1, 9] = 92.7109, p = 0.00001; control vs. PSH, p = 0.001, Tukey’s HST). Upregulation of HIF1α protein expression in the HPC of intact PSH rats was accompanied by an increase in both LDH activity (shown in Fig. 3a; ANOVA F [1, 9] = 17.4660, p = 0.003; control vs. PSH, p = 0.003, Tukey’s HST) and lactate levels (shown in Fig. 3b; ANOVA F [1, 9] = 24.6388, p = 0.001; control vs. PSH, p = 0.001, Tukey’s HST) compared with the intact control.

In the HPC of both control and PSH rats in response to LH, there was an increase in HIF1α protein levels in comparison with the intact control and PSH rats (shown in Fig. 2c, e; ANOVA F [3, 19] = 17.1117, p = 0.00003; control vs. control + LH, p = 0.001, control vs. PSH + LH, p = 0.001, PSH vs. control + LH, p = 0.04, PSH vs. PSH + LH, p = 0.001, Tukey’s HST). The upregulation of HIF1α protein expression in the HPC of control rats after LH was accompanied by increase in lactate levels to levels of intact PSH (shown in Fig. 3b; ANOVA F [3, 19] = 9.6077, p = 0.0007; control vs. control + LH, p = 0.03, PSH vs. control + LH, p = 0.15, Tukey’s HST), but not in LDH activity (shown in Fig. 3a; ANOVA F [3, 19] = 9.2453, p = 0.0009; control vs. control + LH, p = 0.3, PSH vs. control + LH, p = 0.04, Tukey’s HST), in comparison with the intact control. In HPC of PSH rats, LH did not affect LDH activity (shown in Fig. 3a; ANOVA F [3, 19] = 9.2453, p = 0.0009; control vs. PSH + LH, p = 0.005, PSH vs. PSH + LH, p = 0.8, Tukey’s HST) or lactate levels (shown in Fig. 3b; ANOVA F [3, 19] = 9.6077, p = 0.0007; control vs. PSH + LH, p = 0.03, PSH vs. PSH + LH, p = 0.14, Tukey’s HST) compared with the intact PSH.

We also found increased amounts of HIF1α in the HPC of control and PSH rats after SH treatment in comparison with the intact control and PSH rats (shown in Fig. 2f, h; ANOVA F [3, 19] = 68.0716, p = 0.000000003; control vs. control + SH, p = 0.001, control vs. PSH + SH, p = 0.001, PSH vs. control + SH, p = 0.02, PSH vs. PSH + SH, p = 0.001, Tukey’s HST). Moreover, an increase in HIF1α protein levels in the HPC of PSH + SH rats was also observed in comparison with control + SH rats (control + SH vs. PSH + SH, p = 0.005, Tukey’s HST). In the control + SH group, upregulation of HIF1α protein expression in the HPC was accompanied by an increase in both LDH activity (shown in Fig. 3a; ANOVA F [1, 9] = 18.3286, p = 0.003; control vs. control + SH, p = 0.002, Tukey’s HST) and lactate levels (shown in Fig. 3b; ANOVA F [1, 9] = 10.9604, p = 0.01; control vs. control + SH, p = 0.01, Tukey’s HST) compared with the intact control. In the HPC of PSH rats, SH did not affect LDH activity (shown in Fig. 3a; ANOVA F [1, 10] = 0.5217, p = 0.4884) or lactate levels (shown in Fig. 3b; ANOVA F [1, 10] = 0.1266, p = 0.7213) in comparison with intact PSH.

In the HPC of intact PSH rats, we observed a decrease in G6PD protein levels in comparison with control animals (shown in Fig. 2d, e, g, h; ANOVA F [1, 9] = 22.2878, p = 0.001; control vs. PSH, p = 0.002, Tukey’s HST). The downregulation of G6PD protein expression in the HPC of PSH rats was accompanied by a decrease in the NADPH/NADP total ratio (shown in Fig. 3c; ANOVA F [1, 9] = 7.1853, p = 0.03; control vs. PSH, p = 0.03, Tukey’s HST).

The HPC of control rats showed a decrease in G6PD protein levels in response to LH compared to the intact control to levels of intact PSH (shown in Fig. 2d, e; ANOVA F [3, 19] = 8.5821, p = 0.001; control vs. control + LH, p = 0.01, PSH vs. control + LH, p = 0.9, Tukey’s HST). In the HPC of PSH rats, LH did not affect G6PD protein levels compared to the intact PSH (shown in Fig. 2d, e; ANOVA F [3, 19] = 8.5821, p = 0.001; control vs. PSH + LH, p = 0.001, PSH vs. PSH + LH, p = 0.53, Tukey’s HST). After the SH episode, there was an increase in G6PD protein levels in the HPC of PSH rats to levels of the intact control (shown in Fig. 2g, h; ANOVA F [3, 19] = 4.9035, p = 0.01; control vs. control + SH, p = 0.29, control vs. PSH + SH, p = 0.55, PSH vs. control + SH, p = 0.01, PSH vs. PSH + SH, p = 0.04, Tukey’s HST).

After both LH and SH in the HPC of control rats, we observed a decrease in the NADPH/NADP total ratio to levels of the intact PSH, in comparison with the intact control (shown in Fig. 3c; for LH, ANOVA F [3, 19] = 6.1023, p = 0.005; control vs. control + LH, p = 0.01, PSH vs. control + LH, p = 0.9, Tukey’s HST; for SH, ANOVA F [3, 19] = 12.7096, p = 0.0001; control vs. control + SH, p = 0.001, PSH vs. control + SH, p = 0.9, Tukey’s HST). In the HPC of PSH rats, LH and SH did not affect the NADPH/NADP total ratio, in comparison with the intact PSH (shown in Fig. 3c; for LH, ANOVA F [3, 19] = 6.1023, p = 0.005; control vs. PSH + LH, p = 0.025, PSH vs. PSH + LH, p = 0.9, Tukey’s HST; for SH, ANOVA F [3, 20] = 12.7096, p = 0.0001; control vs. PSH + SH, p = 0.001, PSH vs. PSH + SH, p = 0.9, Tukey’s HST).

We found that impairment in G6PD protein expression and NADPH levels (shown in Fig. 2d, e, g, h, 3c) in the HPC of PSH rats was accompanied by a decrease in the GSHred levels (shown in Fig. 3d; ANOVA F [1, 9] = 63.4514, p = 0.00005; control vs. PSH, p = 0.001, Tukey’s HST) and an increase in lipid peroxidation detected by the amount of MDA (shown in Fig. 3e; ANOVA F [1, 9] = 12.1721, p = 0.008; control vs. PSH, p = 0.01, Tukey’s HST) in comparison with the intact control. After both LH and SH in the HPC of control rats, we observed a decrease in the GSHred levels (shown in Fig. 3d; for LH, ANOVA F [3, 19] = 45.4572, p = 0.00000004; control vs. control + LH, p = 0.001, PSH vs. control + LH, p = 0.76, Tukey’s HST; for SH, ANOVA F [3, 20] = 9.0128, p = 0.0009; control vs. control + SH, p = 0.003, PSH vs. control + SH, p = 0.9, Tukey’s HST), but no increase was found in the lipid peroxidation, detected by MDA levels (shown in Fig. 3e; for LH, ANOVA F [3, 19] = 8.1000, p = 0.0017; control vs. control + LH, p = 0.67, PSH vs. control + LH, p = 0.02, Tukey’s HST; for SH, ANOVA F [3, 20] = 6.4439, p = 0.004; control vs. control + SH, p = 0.9, PSH vs. control + SH, p = 0.09, Tukey’s HST) compared to the intact control.

After LH in the HPC of PSH rats, we found changes neither in the GSHred (shown in Fig. 3d; for LH, ANOVA F [3, 19] = 45.4572, p = 0.00000004; control vs. PSH + LH, p = 0.001, PSH vs. PSH + LH, p = 0.82, Tukey’s HST) nor in MDA levels (shown in Fig. 3e; ANOVA F [3, 19] = 8.1000, p = 0.0017; control vs. PSH + LH, p = 0.027, PSH vs. PSH + LH, p = 0.58, Tukey’s HST) in comparison with the intact PSH rats. There were no changes in GSHred levels in the HPC of PSH rats after SH (shown in Fig. 3d; ANOVA F [3, 20] = 9.0128, p = 0.0009; control vs. PSH + SH, p = 0.005, PSH vs. PSH + SH, p = 0.84, Tukey’s HST) in comparison with the intact PSH rats, although MDA levels decreased (shown in Fig. 3e; ANOVA F [3, 20] = 6.4439, p = 0.004; control vs. PSH + SH, p = 0.76, PSH vs. PSH + SH, p = 0.003, Tukey’s HST).

Oxygen is critical for aerobic organisms. Deviations in oxygen concentration significantly reduce the efficiency of energy metabolism and limit vital activities of organisms, thereby causing death [46, 47]. Therefore, signaling aimed at adapting to changes in oxygen accessibility is among the most important cellular processes [1, 30‒32, 35]. The HIF1 transcription factor, which is a heterodimer of HIF1α and HIF1β proteins, is well known as a key factor in adaptation to oxygen deficiency [1, 34, 35]. HIF1α is a regulated subunit that accumulates under hypoxic conditions, whereas HIF1β is a constitutively expressed subunit [30‒32]. In addition to prolyl hydroxylase-associated activity, there are oxygen-independent pathways for stabilizing HIF1α, as well as mechanisms of regulation at the transcriptional level [34, 48‒50]. HIF1 is 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 tissue adaptation to chronic hypoxia [30‒35].

Epigenetic programming during fetal development is a very important and complex process that determines the balance of gene expression specific to cells and tissues, plasticity during their life, and adaptation to permanently changing environmental conditions [28, 51]. A lack of oxygen supply is extremely dangerous for the developing fetal brain [8, 9, 13, 16]. The impact of stressors (e.g., hypoxic stress) during the period of HPC maturation not only has a direct damaging effect on the fetus [25, 52], but also can cause stable disturbances in epigenetic modifications and associated gene expression throughout life [18, 26, 53]. A significant increase in the levels of HIF1α protein in the brain of 21-day-old fetus was previously shown in a model of antenatal hypoxia [10], which indicated a hypoxic condition in 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 from neurons in terms of mitochondrial metabolism and a high rate of glycolysis [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 are determined by HIF1-dependent gene expression [54‒59]. However, we have shown that the increase in HIF1α content in the HPC of adult rats that survived prenatal hypoxia occurs predominantly in neuronal cells. Taking into account the high importance of the adequate expression of this transcription factor to ensure the efficient functioning of the energy metabolism of the brain, deviations in HIF1α expression patterns can affect gene expression and lead to disorders of the nervous system.

Our data on LDH activity and lactate levels indicated that the increased expression of HIF1α in the HPC of PSH rats was 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 PPP, which is a source of NADPH, widely known to be necessary to maintain the cellular antioxidant potential [60‒62]. The delicate balance between these pathways is critical for maintaining both energy demands and antioxidant functions [62]. The 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 can directly inhibit PPP by decreasing G6PD expression [40, 41, 64]. Indeed, in this study, we showed that a stable increase in HIF1α expression in the HPC of rats that survived PSH was accompanied by a decrease in the protein expression of G6PD and a decrease in the amount of the PPP product NADPH. Such 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, the suppression of PPP may cause oxidative stress [40, 62]. A decrease in the efficiency of NADPH production mediates a wide range of adverse conditions, such as disruption of mixed-function oxidases, a decrease in the efficiency of steroid hormone biosynthesis, and a decrease in the rate of recovery of thioredoxins and glutathione, which are reflected in both a defect in individual cell functions and violation of the integrative regulation of the whole organism [61, 62, 65, 66]. Both upregulated anaerobic glycolysis and decreased G6PD activity caused by HIF1-associated perturbations in the HPC of PSH rats led to a decrease in the efficiency of glutathione reduction, which was also accompanied by an increase in free lipid peroxidation.

It is interesting to note that the HIF1-associated metabolic rearrangements 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 1 day after an episode of severe hypoxia or 7 days after an episode of emotional stress (model of “LH”), there was an increase in the protein expression of HIF1α, which was accompanied by an increase in the activity of anaerobic glycolysis and a decrease in the PPP-associated processes 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 did not affect the activity of glycolysis and PPP compared to that in intact PSH rats. In response to emotional stress, the protein expression of G6PD in the HPC of control animals decreased to the level of intact and stressed PSH rats, whereas the severe hypoxic stress only caused an increase in G6PD expression in PSH to control levels, probably through an Nrf2-dependent way [67, 68].

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

Thus, the obtained results show significant HIF1-dependent metabolic changes in the HPC of adult animals as a result of prenatal hypoxia. Having survived an episode of severe hypoxia in the prenatal period, the adult rats demonstrated an increased content of the transcription factor HIF1α in the hippocampal neurons, which was accompanied by a stable imbalance between glycolysis and NADPH production associated with the PPP, thereby resulting in the development of oxidative stress.

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

Animal experiments were performed according to the Guidelines for Reporting Animal Research. This study protocol was reviewed and approved by the local Ethics Committee of the Pavlov Institute of Physiology (protocol no. 08/02 of August 02, 2022).

The authors declare no conflict of interest.

The work has been supported by the Ministry of Science and Higher Education of the Russian Federation (Agreement no. 075-15-2020-921 of November 13, 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.”

O.V. designed experiments; O.V., V.S., S.P., and E.T. performed experiments and wrote the manuscript; O.V. and V.S. analyzed data. All authors have read and approved the final manuscript.

Data are not publicly available due to ethical reasons. Further inquiries can be directed to the corresponding author.