Long-lasting impact of sugar intake on neurotrophins and neurotransmitters from adolescence to young adulthood in rat frontal cortex

doi:10.21203/rs.3.rs-1622302/v1

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Long-lasting impact of sugar intake on neurotrophins and neurotransmitters from adolescence to young adulthood in rat frontal cortex

https://doi.org/10.21203/rs.3.rs-1622302/v1

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The detrimental impact of fructose, a widely used sweetener in industrial foods, was previously evidenced on various brain regions. Although adolescents are among the highest consumers of sweet foods, whether brain alterations induced by the sugar intake during this age persist until young adulthood or are rescued returning to a healthy diet remains largely unexplored. To shed light on this issue, adolescent rats were fed with a fructose-rich or control diet for 3 weeks. At the end of the treatment, fructose-fed rats underwent a control diet for a further 3 weeks until young adulthood phase, and compared with animals that received from the beginning the healthy control diet.

We focused on the consequences induced by the sugar on the main neurotrophins and neurotransmitters in the frontal cortex, as its maturation continues until late adolescence, thus being the last brain region to achieve a full maturity. We here outline that fructose intake induces inflammation and oxidative stress, alteration of mitochondrial function, changes of brain derived neurothrophic factor (BDNF) and neurotrophin receptors, synaptic proteins, acetylcholine, dopamine and glutamate levels, as well as increased formation of the glycation end-products Nε-carboxymethyllysine (CML) and Nε-carboxyethyllysine (CEL). Importantly, many of these alterations (BDNF, CML, CEL, acetylcholinesterase activity, dysregulation of neurotransmitters levels) persisted after switching to control diet, thus pointing out to the adolescence as a critical phase, in which extreme attention should be devoted to limit excessive consumption of sweet foods that can affect brain physiology also in the long term.

adolescent rat

Frontal cortex

fructose diet

brain derived neurotrophic factor

mitochondria

inflammation.

Fructose, a reducing monosaccharide present in fruit and honey, is the major component of the two most used sweeteners, namely sucrose and high fructose corn syrup (HFCS). In the last decades HFCS utilization has been grown, essentially because of its longer shelf life, cheaper production, and higher sweetness which increase the palatability of sugary beverages, baked and processed foods [1, 2]. The rise in the intake of diets rich in sweeteners, mostly sweetened beverages (fruit juices, alcopops and sport drinks) in young and adolescents is particularly troubling [3–6]. A diet rich in sweeteners leads to a marked increase in daily fructose consumption that, compared to a natural consumption of 16–24 g/day with fruits and honey, can reach 80 g/day, which represents the 17–20% of the daily caloric intake [7–9].

Excessive dietary sugar intake, both in early and later life, has been also associated with altered brain metabolism and behavioural functioning [10–13]. In this context, it is noteworthy that brain maturation, particularly in frontal cortex, continues approximately until early adulthood (24 years of age) [14–16], rendering this brain area particularly susceptible to developmental disruption due to early-life nutritional or environmental insults [17]. Indeed, the consumption of high-fat or high-sugar diets during adolescence was associated with both deficits in executive functioning and a reduced volume of the frontal cortical region in humans [18–21]. Therefore, the assessment of the impact of sugar exposure on brain-related outcomes is crucial, particularly during the critical windows of growth and development, when key features of brain tissue structure and function are establishing [22]. While several studies outlined the detrimental effect of fructose on hypothalamus or hippocampus [13, 23, 24], very little information is available on the impact of this sugar on frontal cortex. In this regard, we previously reported that, in both adolescent and adult rat, a short-term fructose intake may affect several processes, in fact redox homeostasis, autophagy, and the expression of synaptic markers were altered in fructose-fed rats compared to controls [25]. Furthermore, long-term fructose drinking was associated with neuroinflammation, altered insulin signaling and cognitive impairment in frontal cortex [26, 27].

Whether the alterations induced by fructose intake during adolescence (postnatal day 30–60, P30-P60) are rescued returning to a healthy diet, or persist until young adulthood is a critical issue that remains unexplored. Therefore, we focused on the frontal cortex of adolescent rats fed a fructose-rich (F) or control diet (C) for 3 weeks. After this period, to highlight whether the effects induced by the short-term fructose diet persist or are rescued, fructose-fed rats were fed a control diet for a further 3 weeks (FR) until young adulthood phase and compared with animals that received the control diet for the entire period (CR). Thus, we investigated putative damages induced by fructose intake on brain-derived neurotrophic factor (BDNF), nerve growth factor (NGF) and their specific receptors, which are crucial in modulating survival and development of the nervous system, synaptic function and plasticity, learning and memory [28, 29], as well as we evaluated the sugar impact on inflammation and oxidative stress. Moreover, a metabolomic analysis was accomplished to further clarify how the sugar influences the homeostasis of specific metabolites, particularly the main neurotransmitters, in the frontal cortex.

Materials

Bovine serum albumin fraction V (BSA), rabbit anti-human Haptoglobin IgG, amino acids, dopamine, acetylcholine, tyramine, γ-aminobutyric acid (GABA), DL-lysine-4,4,5,5-d₄ dihydrochloride (d4-lysine), formic acid, salts and buffers were purchased from Sigma-Aldrich (St. Louis, MO, USA). Nε-carboxymethyllysine (CML) and Nε-carboxyethyllysine (CEL) were obtained from Iris Biotech (Marktredwitz, Germany). Acetonitrile, water and ammonium formate of mass spectrometry grade were obtained from Merck (Darmstadt, Germany).

The dye reagent for protein titration was from Bio-Rad (Hercules, CA); polyvinylidene difluoride (PVDF) and nitrocellulose membranes were from GE Healthcare (Milan, Italy). Horseradish peroxidase (HRP)-conjugated secondary antibodies (goat anti-rabbit, GAR-HRP; goat anti-mouse, GAM-HRP) were from Immunoreagents (Raleigh, NC, USA). Fuji Super RX 100 film was from Laboratorio Elettronico Di Precisione (Naples, Italy).

Experimental design

All experiments were carried out with male Wistar rats (30 days old, P30) purchased from Charles River; Calco, Como, Italy). Housing, treatments and euthanasia of animals were performed as previously published [30]. The rats were divided into two groups, one fed a fructose rich diet (F group), the other one fed a control diet (C group) for 3 weeks. At the end of treatment, half of the rats from each group was euthanized, while the other half received a control diet (FR and CR groups) for further 3 weeks (P72). The composition of both control and fructose rich diet is reported in Supplementary Table 1.

The animals were then euthanized, and frontal cortex was harvested and dissected as previously described [25]. Mitochondrial oxygen consumption was immediately assessed in little aliquots of tissue. Pieces of each sample were fixed for immunofluorescence analysis, while the remaining samples were snap frozen in liquid nitrogen and stored at − 80°C for further analyses.

Mitochondrial analyses

Frontal cortex samples were homogenized (1:1000, w/v) in Mir05 medium containing 110 mM sucrose, 60 mM K-lactobionate, 20 mM Hepes, 20 mM taurine, 10 mM KH₂PO₄, 6 mM MgCl₂, 0.5 mM EGTA, and 0.1% w/v fatty acid-free BSA, pH 7.0.

Homogenates (2mg) were transferred into calibrated Oxygraph-2k (O2k, Oroboros Intruments, Innsbruck, Austria) 2-mL chambers. Oxygen polarography was performed at 37 ± 0.001°C (electronic Peltier regulation), and oxygen concentration (µM) and oxygen flux (pmol O₂ s⁻1 mL^− 1) were real-time recorded and corrected automatically for instrumental background by DatLab software (Oroboros Intruments, Innsbruck, Austria).

After addition of the homogenates, the O₂ flux was allowed to stabilize. A substrate, uncoupler, inhibitor titration (SUIT) protocol was applied to assess qualitative and quantitative mitochondrial changes [31]. After stabilization, leak respiration supported primarily by electron flow through complex I of the respiratory chain was evaluated by adding the substrates malate (0.5 mM), pyruvate (5 mM), and glutamate (10 mM). Electron transfer was coupled to phosphorylation by the addition of 2.5 mM ADP, assessing phosphorylating respiration with electron transfer supported by complex I. Succinate (10 mM) was added to the chamber to induce maximal phosphorylating respiration with parallel electron input from complexes I and II. Oligomycin (2.5 mM) was added to assess leak respiration when substrates and ADP were provided, but ATP synthase is inhibited. Maximum capacity of the electron transport chain was obtained by addition of the uncoupler carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP, 0.5 mM). Rotenone (0.5 µM) was added to inhibit complex I; hence, the maximal capacity supported by complex II alone was determined. Residual oxygen consumption was established by addition of the inhibitor antimycin A (2.5 mM) and the resulting value was subtracted from the fluxes in each run, to correct for non-mitochondrial respiration. All samples were run in duplicates and the mean was used for analysis.

Procedures to test mitochondrial integrity were routinely carried out at the beginning of each measurement, by evaluating the stimulating effect of 10 mM exogenous cytochrome c on mitochondrial respiration in the presence of complex I- linked substrates and ADP.

Metabolic parameters assay

The amount of fructose and uric acid in frontal cortex samples was measured by colorimetric enzymatic methods, using commercial kits according to the manufacturer’s instruction (Sigma Aldrich, St. Louis, MO, USA for fructose, and GS Diagnostics SRL, Guidonia Montecelio, Rome, Italy for uric acid).

Protein extraction

Aliquots of frontal cortex (about 50 mg) were homogenized in seven volumes (w/v) of cold RIPA buffer, as previously published [32], and protein concentration was titrated by the colorimetric Bio-Rad Bradford protein assay, using a commercial kit (Bio-Rad, Hercules, CA, USA), according to the manufacturer’s instruction. Then protein extracts were used for titrating the markers reported below by ELISA or Western blotting.

Analysis of tumor necrosis factor alpha (TNF-alpha)

TNF-alpha concentration was evaluated by sandwich ELISA in frontal cortex homogenates diluted 1:20 [33] using the TNF-alpha Duo-Set kit (R&D, DBA Italia). Data were reported as pg of TNF-alpha per mg of total proteins.

Western blotting

Aliquots (30 µg) of cortex proteins were resolved by electrophoresis, under denaturing and reducing conditions [34], on 12.5% (to quantify glucose transporter-5, Glut-5; glial fibrillary acidic protein, GFAP; synaptophysin; brain derived neurotrophic factor, BDNF; Nerve growth factor, NGF; extracellular signal-regulated kinase, Erk1/2) or 10% (post-synaptic density protein 95, PSD-95; synaptotagmin I; Peroxisome proliferator-activated receptor gamma coactivator 1-alpha, PGC-1α; Tropomyosin receptor kinase B, TrkB; Tropomyosin receptor kinase A, TrkA; Thyroxine hydroxylase, TH) polyacrylamide gels. Proteins were then blotted onto PVDF or nitrocellulose membrane [35], and the following washing and blocking steps were performed according to [36]. Immunodetection was carried out as follows:

Glut-5: rabbit anti Glut-5 IgG (Invitrogen, Carlsbad, CA, USA, 0.5 µg/mL in T-TBS containing 3% w/v BSA; overnight, 4°C), followed by goat anti-rabbit horseradish peroxidase-conjugated IgG (GAR-HRP; 1: 45,000 dilution in T-TBS containing 2% w/v BSA; 1 h, 37°C); GFAP: rabbit anti-human GFAP IgG (Cell Signaling, MA, USA; 1: 1,000 in T-TBS containing 1% v/v non-fat milk; overnight, 4°C), followed by GAR-HRP IgG (1:150,000 in 5% v/v non-fat milk; 1 h, 37°C); Synaptophysin: rabbit anti-Synaptophysin IgG (Merk Millipore, Milan, Italy; 1:150,000 in 3% w/v BSA; overnight, 4°C) followed by GAR-HRP IgG (1:40,000 dilution in 1% v/v non-fat milk; 1 h, 37°C); Synaptotagmin I: rabbit anti-Synaptotagmin I IgG (Cell Signaling; 1:1,000 dilution in 3% v/v non-fat milk; overnight, 4°C) followed by GAR-HRP IgG (1:200,000 dilution in 3% v/v non-fat milk; 1 h, 37°C); PSD-95: rabbit anti-PSD 95 IgG (Cell Signaling; 1:1,000 in 3% v/v non-fat milk; overnight, 4°C) followed by GAR-HRP IgG (1:70,000 dilution in 3% v/v non-fat milk; 1 h, 37°C); BDNF: rabbit anti-human BDNF IgG (Santa Cruz Biotechnology, CA, USA; 1:500 dilution in 0.25% v/v non-fat milk; overnight, 4°C), followed by GAR-HRP IgG (1:70,000 dilution in 1% v/v non-fat milk; 1 hour, 37°C); PGC-1α: rabbit anti-human PGC-1α IgG (Millipore, MA, USA; 1:2,000 dilution in 3% w/v BSA; overnight, 4°C), followed by GAR-HRP IgG (1:20,000 dilution in 3% w/v BSA; 1 h, 37°C); TrkB: rabbit anti-TrkB (Santa Cruz Biotechnology, CA, USA; 1:2000 dilution in 2% w/v BSA; overnight, 4°C) followed by GAR-HRP IgG (1: 140 000 dilution in 2% w/v BSA; 1 h, 37°C); pErk1/2: rabbit anti-pErk1/2 IgG (Cell Signaling; 1:1,000 dilution in 2% w/v BSA; overnight, 4°C) followed by GAR-HRP IgG (1:150,000 dilution in 2% w/v BSA; 1 h, 37°C); Erk 1/2: rabbit anti-Erk1/2 (Cell Signaling; 1:1,000 dilution in T-TBS containing 2% BSA; overnight, 4°C) followed by GAR-HRP IgG (1:170,000 dilution in 2% w/v BSA; 1 h, 37°C); TH: mouse anti-TH (Santa Cruz; 1:1,000 dilution in 5% v/v non-fat milk; overnight, 4°C) followed by goat anti-mouse (GAM-HRP) IgG (1:10,000 dilution in T-TBS; 1 h, room temperature); TrkA: rabbit anti-TrkA (Santa Cruz; 1:1,000 dilution in 5% v/v non-fat milk; overnight, 4°C) followed by GAR-HRP IgG (1:10,000 dilution in T-TBS; 1 h, room temperature); NGF mouse anti-NGF (Santa Cruz; 1:1,000 dilution in 5% v/v non-fat milk; overnight, 4°C) followed by GAM-HRP IgG (1:10,000 dilution in T-TBS; 1 h, room temperature).

For loading control, β-actin or vinculin was revealed after detection of each marker. To this aim, the membranes were stripped [37], and then treated with mouse anti-β-actin IgG (1:1,000 in 0.25% v/v non-fat milk; overnight, 4°C) followed by GAM-HRP IgG (1:30,000 in 0.25% v/v non-fat milk; 1 h, 37°C) or with mouse anti-vinculin IgG (Sigma Aldrich, Milan, Italy; (1:40,000 dilution in 5% v/v non-fat milk; overnight, 4° C) followed by GAM-HRP IgG (1:10,000 dilution in T-TBS; 1 h, room temperature).

The Excellent Chemiluminescent detection Kit (Cyanagen s.r.l., Bologna, Italy) was used for detection. Chemidoc or digital images of X-ray films exposed to immunostained membranes were used for densitometric analysis and quantification was carried out by Un-Scan-It gel software (Silk Scientific, UT, USA).

Haptoglobin (Hpt) evaluation

Hpt concentration in frontal cortex samples was measured by ELISA as previously reported [36]. Samples were diluted (1: 3,000, 1:10,000, 1:30,000) with coating buffer (7 mM Na₂CO₃, 17 mM NaHCO₃, 1.5 mM NaN₃, pH 9.6), and aliquots (50 µl) were then incubated in the wells of a microtiter plate (Immuno MaxiSorp; overnight, 4°C). Washing and blocking were carried out as previously reported [38], then the wells were incubated (1 h, 37°C) with 50 µl of rabbit anti-human haptoglobin, (1:500 in 130 mM NaCl, 20 mM Tris-HCl, 0.05% Tween, pH 7.4, containing 0.25% BSA), followed by 60 µl of GAR-HRP IgG (1:5000 dilution, 1 h, 37°C). Peroxidase-catalyzed color development from o-phenylenediamine was measured at 492 nm.

Evaluation of nitro-tyrosine levels, acetylcholinesterase (AChE) and monoamine oxidase (MAO) activities

Nitro-Tyrosine (N-Tyr) titration was carried out by ELISA in frontal cortex homogenates as previously described [39]. Samples were diluted (1:1,500, 1:3,000, 1:6,000) with coating buffer, and aliquots (50 µl) were then incubated in the wells of a microtiter plate (overnight, 4°C). After washing and blocking, the wells were incubated (1 h, 37°C) with 50 µl of rabbit anti-N-Tyr IgG (Covalab, distributed by VinciBiochem, Vinci, Italy; 1: 1000 dilution in T-TBS containing 0.25% w/v BSA) followed by 60 µl of GAR-HRP (1:9,000 dilution; 1 h, 37°C). Peroxidase-catalyzed color development from o-phenylenediamine was measured at 492 nm. Data were reported as OD per mg of total proteins.

The AChE activity was measured in frontal cortex samples as previously described [25]. Enzyme activity was expressed as nmol/min mg protein.

The monoamine oxidase (MAO) activity was measured spectrophotometrically following the conversion of benzylamine to benzaldehyde, as previously described [40].

Immunofluorescence analysis

Paraffin embedded sections of frontal cortex from all the groups were stained with the cAMP response element-binding protein (p-CREB) specific monoclonal antibody (p-CREB antibody, Cell Signaling, 1:1,000 in dilution in PBS containing 2% w/v BSA; overnight, 4°C), and DAPI (Sigma Aldrich, Saint Louis, MO, USA). For the analysis, images were acquired with x40 magnification and 3 random field/section per rat were analyzed using ImageJ (National Institutes of Health, Bethesda, MD, USA). Images were captured and visualized using a Nikon Eclipse E1000 microscope.

Liquid chromatography-electrospray-high-resolution tandem mass spectrometry (LC-MS/MS)

Amino acids and their derivatives were analyzed by liquid chromatography-electrospray-high-resolution tandem mass spectrometry (LC-MS/MS) as previously reported [41], with minor modifications. Frontal cortex samples (25 ± 10 mg) were dissolved in 0.390 mL of 0.1% formic acid along with 10 µL of 10 µg/mL lysine d-4; suspensions were accurately homogenized by using a stainless-steel disperser (IKA T10, Staufen, Germany, 3 passes, 30 s) in an ice bath. Supernatants (0.1 mL) were further purified by using 0.3 mL of 0.1% formic acid in acetonitrile by a protein precipitation and phospholipids removal cartridge (Phree, 1 mL, Phenomenex, Torrance, CA); eluates were collected and dried by using a centrifugal evaporator (SpeedVac, Thermo Fisher Scientific, Bremen, Germany). Dried samples were dissolved in 0.1 mL of a mixture acetonitrile: water: formic acid (50:49.9:0.1, v/v/v) and 5 µL injected into the LC-MS/MS system consisting in a linear ion trap with Orbitrap detector (LTQ Orbitrap XL) interfaced to an Ultimate 3000 RS (Thermo Fisher Scientific, Bremen, Germany). Analytes were separated through hydrophilic interaction chromatography in data dependent scan positive ions mode for identification and quantitation of amino acids, catecholamines, indole derivatives and polar markers of oxidation.

Chromatographic separation was achieved through a silica sulfobetaine zwitterionic modified HILIC column (100 x 2.1 mm, 1.7 µm, Syncronis HILIC, Thermo Fisher, Bremen, Germany) at 35°C. Mobile phases consisted in 0.1% formic acid in acetonitrile:water 95:5 (v/v, solvent A) and 0.1% formic acid in water (solvent B) both with 5 mM ammonium formate. Samples (4°C) were separated through the following gradient of solvent B (minutes/%B): (0/3), (3.5/3), (15.5/75), (17.5/75) at a flow rate of 0.25 mL/min. Electrospray interface (ESI) parameters were the following: spray voltage 5.0 kV, capillary voltage 21.0 V, capillary temperature 300°C, sheath gas flow and auxiliary gas flow were 25 and 4 arbitrary units, respectively. Profile data type were acquired in full scan FTMS mode (Fourier transformed) in the mass range 75–750 m/z. For data dependent scanning mode, MS/MS normalized collision energy was set to 20, activation Q 0.25, activation time 25 ms, with a 1 m/z isolation window, while a reject mass list was generated by injecting blank samples consisting in a mixture of acetonitrile:water:formic acid (50:49.9:0.1, v/v/v).

For compound identification, differential analysis and metabolic pathways, raw data were loaded in Compound Discoverer (v. 3.2, Thermo Fisher Scientific). The workflow included the identification of both expected and unknown metabolites; briefly, each node performed retention time alignment, expected compound detection, biotransformation, dealkylation and dearylation products formation. Resolution and isotope pattern matching with unknown compounds detection were used across all samples with a mass accuracy below 5 ppm. FISh (fragment ion searching) scoring was applied to all expected compounds with automatic fragment annotations based on targeted and untargeted compound chemical behavior outlined in Human Metabolome Database (https://hmdb.ca/). According to the background in blank samples, the procedure predicted elemental compositions for all unknown compounds, while quality control samples (QC, consisting in pooled samples spiked with amino acid standards) corrected signal intensities each 10 runs. Hierarchical clustering was obtained through filtering procedures based on technical replicates coefficient of variation (CV%), retention time and mass accuracy. The workflow included differential analysis (p-values, adjusted p-values, ratios, fold change), Euclidean distance and complete linkage method without data normalization to enhance differences among frontal cortex of groups fed different diets.

For targeted analyte quantitation, a calibration curve of the compounds listed in Supplementary Table 2 was built in the range 100–5000 ng/mL. Linearity and the responses of intraday and interday assays were monitored by using Xcalibur 2.1 with a mass accuracy fixed at 5 ppm (Thermo Fisher Scientific, Bremen).

Statistical analysis

Data were expressed as mean values ± SEM. The program GraphPad Prism 6 (GraphPad Software, San Diego, CA, USA) was used to verify that raw data had normal distribution and to perform one-way ANOVA followed by Bonferroni post-test. P < 0.05 was considered significant in the reported analyses.

Inflammation, oxidative stress and mitochondrial activity

To investigate whether the dietary treatment induces fructose transporter and metabolism, the protein expression of Glut-5, as well as the level of fructose and uric acid, one of the main products of fructose metabolism, were analyzed in frontal cortex after 3 weeks of a fructose-rich diet. The amount of Glut-5 was significantly higher (P < 0.05) in frontal cortex of F rats compared to C rats, while this increase disappeared in FR rats (Fig. 1a). Accordingly, a significant increase in the levels of fructose (P < 0.01) and uric acid (P < 0.01) was found in F rats compared to C rats, while no significant differences were found in FR rats compared to CR ones (Fig. 1b, c).

We then investigated the inflammatory status by measuring the protein expression of GFAP, a marker of astrogliosis, the pro-inflammatory cytokine TNF-alpha, and Hpt, a marker of inflammation very sensitive to nutritional changes [33, 36, 42]. As shown in Fig. 1d-f, an increase of inflammatory markers in frontal cortex was associated with the fructose-rich diet. As a matter of the fact, significantly higher levels of GFAP (P < 0.01; Fig. 1d), TNF-alpha (P < 0.01; Fig. 1e) and Hpt (P < 0.01; Fig. 1f) were found in frontal cortex of F rats compared to C rats. Importantly, these conditions were rescued after switching to a control diet as no difference in the levels of these markers was detected between CR and FR rats.

In line with previous results obtained in the hippocampus [30], fructose feeding was also associated with an increase of N-Tyr, the footprint of protein oxidative damage induced by peroxynitrite [43] in fructose-fed rats (P < 0.001; Fig. 1g). The observed condition of redox imbalance was corroborated by changes in mitochondrial activity induced by fructose-rich diet. In detail, F rats showed a significant decrease in leak respiration with complex I and II-linked substrates, in ADP-supported respiration with complex I or complex I- and II-linked substrates and in FCCP-stimulated respiration with complex II or complex I and II-linked substrates (Fig. 1l). The above mentioned mitochondrial dysfunction was reversed when rats were switched to a control diet. The analysis of PGC-1α was performed as a marker of mitogenesis, and no difference was observed (Fig. 1m).

We also detected higher levels of the advanced glycation end-products Nε-carboxymethyllysine (CML) and Nε-carboxyethyllysine (CEL) [44], both in free form, in frontal cortex of F rats compared to C rats (P < 0.01 and P < 0,05 respectively; Fig. 1h-i). Notably, after switching to the control diet, the differences in N-Tyr between CR and FR rats disappeared, while CML and CEL levels persisted higher in FR rats compared to C (P < 0.01).

Neurotrophins and synaptic proteins

The level of BDNF, a key cerebral factor involved in a wide range of neurophysiological processes such as neuronal survival, synaptic transmission and plasticity [45], was measured in the frontal cortex of all groups of rats. As shown in Fig. 2a, a significant fructose diet-dependent decrease of the mature form of BDNF was observed (P < 0.01). Notably, alterations of BDNF amount persisted after switching to a control diet, as displayed by FR rats compared to CR rats (P < 0.001). No difference was found in the levels of BDNF precursor (pro-BDNF), suggesting that changes in this neurotrophin were likely at post-translational level. The protein amount of TrkB, the high-affinity receptor of BDNF, was also analyzed, as its activation enhances synaptic plasticity, neuroprotection, and neurite outgrowth [45]. As shown in Fig. 2b, TrkB level was significantly lower in the frontal cortex of F rats compared to C ones (P < 0.05), but its level in FR rats returned to values comparable to CR rats. In line with the decrease of BDNF, a decrease in the activating phosphorylation of CREB (p-CREB) was found in both F rats compared to C, and FR compared to CR (P < 0.01; Fig. 2c), as assessed by immunofluorescence.

According to previous works in human, rat, and mouse brain, reporting that mature NGF is absent while pro-NGF is the predominant species [46, 47], mature NGF was not detectable in any frontal cortex samples analyzed. Moreover, no significant difference between the experimental groups was found in the levels of its precursor pro-NGF (Fig. 3a). Interestingly, TrkA level was increased by the fructose diet in F rats respect to C (P < 0.05), while no difference was detected in FR if compared to CR, after the sugar removal from the diet (Fig. 3b). Furthermore, no changes were observed in the amount of the low-affinity NGF receptor p75NTR (Fig. 3c). In addition, the phosphorylation degree of both Erk1 and Erk2 significantly increased in F compared to C rats (P < 0.01 and P < 0.001, respectively; Fig. 3d), with persistent increased levels in FR compared to CR rats (P < 0.01 and P < 0.001, respectively).

We further investigated the impact of fructose on the amounts of two pre-synaptic proteins, namely Synaptophysin and Synaptotagmin I, and the post-synaptic protein PSD-95, which plays a key role in synaptic plasticity [48]. The fructose-rich diet led to decreased levels of all the three proteins (P < 0.01), while the switch to control diet rescued their amount (Fig. 4a-c).

The effect of the short-term fructose diet on the activity of AChE, a pivotal enzyme involved in the regulation of cholinergic pathways [49], was also evaluated. As shown in Fig. 4d, fructose feeding resulted in a significant increase of AChE activity of F compared to C rats (P < 0.05). This increase was also found in FR rats compared to CR (P < 0.05). Notably, the activity of MAO, a central player in the modulation of monoamine neurotransmitters level [50] increased in F rats compared to C rats (P < 0.0001; Fig. 4e), with no significant change between FR and CR.

Metabolites and neurotransmitters

Polar hydrophilic metabolites were measured by using HILIC coupled to high-resolution tandem mass spectrometry according to the nature and duration of the intervention study; in particular, the C rat group was compared with the F one, while CR was matched to FR. Results are reported in the heat-maps in Fig. 5. Panel A highlighted a clear discrimination between fructose-diet (orange arrow) and control diet (blue arrow) through Euclidean distance between over- and down-represented analyte intensities in the normalized area range − 2.8 and 3.9, as related to the current ion associated with 89 molecules. In a similar way, polar hydrophilic metabolites differentiated rescue fructose-diet (light blue arrow, FR) and rescue control diet (blue arrow, CR) (Fig. 5b) when considering up to 272 over- and down-represented compounds. In the heat-map in panel A and according to metabolic pathways suggested in Metabolika node within Compound Discoverer, we identified a main cluster of metabolites and an increased abundance of aromatic amino acids, namely tyrosine, phenylalanine and tryptophan, along with sulphur compounds (cysteine and reduced glutathione) and other derivatives (Fig. 5c). In the second heat-map, we observed an increased abundance of some polar basic amino acids, such as lysine and CEL, secondary amides and other acid compounds, like aspartic acid derivatives and glutamic acid (Fig. 5d). In this view, we hypothesized that the metabolism of aromatic amino acids, lysine and glutamic acid can be the focus of high fructose diet. This hypothesis is in accordance with a previous observation [51], which reported that amino acids linked to tricarboxylic acid cycle, such glutamate and aspartate, undergo specific enrichment when young rats are fed with a high-fructose diet. Furthermore, a coherent alteration of the lysine catabolism was recently observed in diurnal metabolic pathways impacted by high-fat diet, suggesting that fructose intake can similarly exacerbate the alteration of amino acid metabolism [52].

Based on the above-reported untargeted analysis (Fig. 5), we then focused on the targeted quantification of various metabolites in all frontal cortex homogenates to highlight differences between C and F rats, and between CR and FR. Our major effort was to evaluate the quantitative impact of the fructose diet on the profile of the major neurotransmitters (Fig. 6). We observed that the concentration of acetylcholine (ACh), one of the main neuromodulators of the central nervous system regulating individual attention, learning, and memory [53], was significantly higher in F rats compared to C (P < 0.01), and this increase was rescued in FR (Fig. 6a). Conversely, the levels of dopamine, the neurotransmitter regulating the food eating reward circuit, motor activity and emotion [54, 55], were reduced in F rats compared to C ones (Fig. 6b; P < 0.001), and no difference between CR and FR groups was observed. Similarly, levels of both dopamine precursors tyrosine and tyramine decreased in F compared to C rats (P < 0.01 and P < 0.05, respectively; Fig. 6c-d), and returned to control values after the switch to the standard diet. On the other hand, no changes in the amounts of TH, the enzyme catalysing the rate-limiting step of catecholamine biosynthesis [56], were observed between F and C groups (Fig. 6e). Interestingly, the levels of the most common neurotransmitter in the CNS, namely glutamate [57], were lower in F group with respect to C one (Fig. 6f; P < 0.01). A different trend was observed after the switch to control diet, as FR showed higher glutamate levels compared to CR (Fig. 6f; P < 0.01). These results further demonstrated that fructose intake is associated with a dysregulation of glutamate metabolism, which is further observed, but with an opposite quantitative trend, after switching to control diet. Finally, GABA levels did not differ between C and F groups, while increased amounts were observed in FR compared to CR (Fig. 6g; P < 0.01).

Different lines of evidence have recently highlighted the fructose impact on brain metabolic alterations [13, 23], but, to our knowledge, poor information is available on the metabolic effects of this sugar on frontal cortex in adolescent animal models. More importantly, it remains unclear whether, during this critical phase of growth, the observed sugar-induced effects are limited exclusively to the period of increased intake or are persistent even when it is eliminated from the dietary regimen.

We here show that a short-term fructose diet was associated with increased levels of the fructose transporter Glut-5 in rat frontal cortex, compared to control, and a concomitant enhanced amount of fructose and uric acid, which were suggestive of an enhanced fructose metabolism therein. These modified protein and metabolite levels were paralleled by an evident inflammatory status, as deduced by the observed increased levels of TNF-alpha, GFAP and Hpt in frontal cortex of fructose-fed rats. A similar pattern was previously found in the hippocampus [30], thus showing that dietary fructose reaches the brain and is utilized in several districts. Augmented levels of uric acid were also reported to elicit oxidative stress [58–60] and consistently, we found a higher extent of oxidative damage to proteins (higher level of N-Tyr). Inflammation and oxidative stress are often related to a mitochondrial dysfunction [61, 62]. When we evaluated the mitochondrial respiratory functions in frontal cortex homogenates from fructose-fed rats, in a condition where mitochondria are maintained in a cellular context [39], we evidenced a corresponding general impairment of the mitochondrial oxidative capacity in all the experimental conditions we tested. This mitochondrial alteration was not linked to a lower organelle mass, since PGC-1α representation was not altered in all the experimental groups. All above-mentioned fructose-diet dependent effects on mitochondrial activity were reverted when this sugar was abolished from the dietary regimen.

Intake of fructose and the consequent altered glucose metabolism can lead to the formation of highly reactive intermediate products as α-dicarbonyls [63], that in turn can favour the formation of advanced glycation end-products, as CML and CEL [64]. Both compounds showed increased levels in frontal cortex of fructose-fed rats, in line with previous investigations showing that fructose can act as a potent glycating agent [65] and contribute to the accumulation of glyoxal and methylglyoxal, thus negatively impacting the brain functions [13, 25, 30, 66, 67]. Interestingly, the observed augmented levels of CML and CEL in fructose-fed rats were not restored to control values in FR group, suggesting the persistence of the effect of the ingested sugar, which might be hazardous for normal functioning of the brain.

Given the key role of neurotrophins in brain activity, we focused on the measurement of BDNF and NGF in frontal cortex of rat groups. In particular, we observed that the amount of BDNF, which has a fundamental role in synaptic transmission and plasticity [45], was lower in fructose-fed rats. This result is in line with previous findings obtained in adult rats experiencing a long- or short-term fructose intake [25, 27], highlighting this neurotrophin as a crucial target of fructose diet independently from the age. The BDNF receptor TrkB also decreased following fructose intake. Interestingly, the reduction of BDNF persisted in the young adult rats after the switching to the control diet. The early reduction of BDNF in adolescent rats and its persistent decrease later was corroborated by the observation of the corresponding lower levels of its downstream effector, namely p-CREB, both in F compared to C rats and in FR compared to CR counterparts. Since CREB is implicated in the transcription of genes essential for synaptic plasticity [68], the persistent reduction in its phosphorylated form, even after the interruption of the fructose feeding, was suggestive of a long term brain impairment.

The analysis of pro-NGF, TrkA, p75NTR, and Erk1/2 suggested that above-reported sugar-induced impairment of BDNF signaling might be compensated, at least in part, by the increase of TrkA and activation of its downstream transducers Erk1/2. This process is regulated by pro-NGF and critically depends on the balance of TrkA and p75NTR [69, 70]. This pathway is indeed committed to prevent apoptotic signals, with a positive effect on neuronal survival and neurite outgrowth [47].

The persistent impairment of BDNF-CREB signaling prompted us to investigate the issue of synaptic functioning. Our results showed that the fructose dietary treatment is associated with a decrease of pre- (synaptophysin, and synaptotagmin I) and post- (PSD-95) synaptic proteins. However, synaptic protein levels returned to values comparable to the controls in young adult rats switched to the standard diet. These findings suggest that the mechanisms underlying the regulation of these synaptic proteins might restore their homeostasis more rapidly than those involved in the maturation and activity of BDNF. To gain further insight into synaptic physiology, we also measured the activity of AChE and MAO in frontal cortex of various rat groups, as well as the levels of important neurotransmitters involved in synaptic transmission. In particular, increased activity of AChE and MAO was detected in fructose-fed rats. While increased MAO rescued after switching to standard diet, AChE increase persisted after fructose removal from the diet. In this context, it is worth mentioning that the enhanced AChE activity in the hippocampus and prefrontal cortex was previously proposed as an early event linked to hypercholesterolemia- or high-fat diet-induced alterations in cognitive function [71–73]. Notably, despite the enhanced AChE activity, we detected increased levels of ACh in fructose-fed rats. In this regard, previous studies showed that an abnormal intake of fructose provokes an immediate drop in the ATP/AMP ratio, a decreased acetyl-CoA carboxylase activity, and a consequent lowering of malonyl-CoA levels [74]. Accordingly, it can be hypothesized that the increased ACh levels after fructose intake might derive from the increased availability of acetyl CoA, which was in turn consequent to the decreased activity of acetyl-CoA carboxylase. In this sense, the higher AChE activity may represent an adaptive response to prevent prolonged ACh signaling. Indeed, a prolonged or enhanced ACh signaling above a homeostatic set point can heighten symptoms related to anxiety and depression, altering the optimal balance between cognition and mood regulation [75–78]. Results on the increase of ACh levels in F rats suggest that a short term fructose enriched diet may contribute to disrupt cholinergic signaling and predispose adolescents to altered states, such as anxiety disorders and depression, also corroborated by BDNF and dopamine reduction. Indeed, our metabolomic analysis also showed that fructose feeding is associated with a decrease of dopamine, which regulates different brain functions such as reinforcement processing, motivation, and attention in frontal cortex [79]. In this context, the observed dopamine decrease in F rats might be ascribed to both the corresponding enhanced MAO activity and the reduced amounts of tyrosine and tyramine. In agreement with this hypothesis, previous studies have reported that HFCS can impair dopamine function in mice in the absence of weight gain or increased fat consumption [10]. As a reduced dopamine function has been implicated in reduced energy expenditure [80, 81], changes in dopamine metabolism induced by HFCS were proposed to precede and possibly contribute to obesity in the long-term period [10]. Further, dysfunction in dopaminergic transmission has recently been associated with depression and accelerated brain aging [82]. Therefore, the herein reported diet-induced alteration of dopamine levels might contribute to prompt defects in learning and memory in the long-term.

Untargeted metabolomic also showed that glutamate levels decreased in fructose fed rats. A general alteration of some metabolic pathways related to specific aromatic or sulfur-containing amino acids, lysine and glutamic acid was also deduced from F vs C, and CR vs FR comparisons. Glutamate is the principal excitatory neurotransmitter involved in learning, memory and cognition, and any unbalance of its turnover may have severe consequences [57]. In line with our results, reduced glutamate levels were previously detected in the cortex of mice receiving a long term high-fat high-sucrose diet [83]. Of note, the switching to the control diet was associated with an increase of the neurotransmitter level in FR rats compared to CR counterparts. The here reported opposite differences, before and after switching to control diet, might be suggestive of a glutamatergic persistent dysregulation after fructose intake, which might prelude to long term dysfunction. Indeed, glutamatergic dysregulation has already been reported being an important contributor to different neurological pathologies, although the molecular basis is distinct for each disease [57]. As a matter of the fact, disorders of the glutamatergic system are associated with severe deficiency of information transmission in the neural network and impairment of cognitive processes [84].

ACh, dopamine and glutamate changes observed in fructose-fed rats were not paralleled by GABA alterations. Conversely, augmented concentration of this neurotransmitter was observed only in young adult rats after the sugar removal from the diet. Interestingly, increased GABA levels in astroglia were shown to occur in elderly humans, patients with Alzheimer’s disease (AD), and amyloid-depositing AD mouse models [85]. Hypertrophic GABAergic astrocytes have also been observed, in the cortex and hippocampus of AD mice, in the proximity of Aβ plaques [86]. Accordingly, the impairment in GABAergic tone was proposed to lead to microglial dysregulation, enhancing disease progression [87]. Based on what reported above, it could be hypothesized that the observed increase of GABA in FR rats might contribute to the induction of degenerative events. However, since higher levels of glutamate were also observed in FR rats, we cannot exclude that the concomitant increase of GABA amount may represent a compensatory mechanism to prevent possible dysfunction related to concomitant quantitative changes of its precursor.

In conclusion, this study demonstrates that the fructose feeding causes a perturbation of various biochemical machineries involved in brain metabolism and function, such as neurotrophins signaling and a consequent possible modification of excitatory/inhibitory neurotransmitter balance, which is essential for the proper functioning of the central nervous system (Fig. 7). Undoubtedly, these data also point out that adolescence represents a developmental window of vulnerability, in which extreme attention should be devoted to limit excessive consumption of industrial and processed sweet food, since it can impacts brain physiology not only immediately but also in the long term.

Acknowledgments

The authors wish to thank Dr. Emilia de Santis for skillful management of animal house.

Funding

This work was supported by grants from: a) University of Naples Federico II - Ricerca Dip 2020/2021 to LC; b) National Research Council - Premio DiSBA for the project DISCEFRU to MSS, and MUR - PON ARS-01-00783 for the project (ALIFUN) to AS.

Competing Interests

The authors have no competing interests to declare.

Author Contributions

Luisa Cigliano, Maria Stefania Spagnuolo, Arianna Mazzoli and Susanna Iossa contributed to the study conception and design. Material preparation was performed by Maria Stefania Spagnuolo, Arianna Mazzoli, Martina Nazzaro and Luisa Cigliano. Data collection and analysis were performed by Maria Stefania Spagnuolo, Arianna Mazzoli, Martina Nazzaro, Cristina Gatto, Antonio Dario Troise, Claudia Tonini, Mayra Colardo, Marco Segatto, Andrea Scaloni, Valentina Pallottini, Susanna Iossa. The first draft of the manuscript was written by Maria Stefania Spagnuolo and Luisa Cigliano, and all authors commented on previous versions of the manuscript. All authors read and approved the final manuscript.

Data Availability

All data supporting the findings of this study are available within the article and its supplementary information files.

Ethics approval

The study was conducted according to the guidelines of the Declaration of Helsinki, approved by “Comitato Etico-Scientifico per la Sperimentazione Animale” of the University of Naples Federico II and authorized by Italian Health Minister (4448/2019-PR).

Consent to participate

Not applicable

Consent to publish

Not applicable

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Long-lasting impact of sugar intake on neurotrophins and neurotransmitters from adolescence to young adulthood in rat frontal cortex

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Abstract

Figures

Introduction

Materials And Methods

Materials

Experimental design

Mitochondrial analyses

Metabolic parameters assay

Protein extraction

Analysis of tumor necrosis factor alpha (TNF-alpha)

Western blotting

Haptoglobin (Hpt) evaluation

Evaluation of nitro-tyrosine levels, acetylcholinesterase (AChE) and monoamine oxidase (MAO) activities

Immunofluorescence analysis

Liquid chromatography-electrospray-high-resolution tandem mass spectrometry (LC-MS/MS)

Statistical analysis

Results

Inflammation, oxidative stress and mitochondrial activity

Neurotrophins and synaptic proteins

Metabolites and neurotransmitters

Discussion

Declarations

References

Supplementary Files

Status:

Version 1