Introduction

Lesch–Nyhan Disease (LND) is an innate metabolic error of purine metabolism caused by a severe deficiency of hypoxanthine–guanine phosphoribosyl-transferase (HPRT, EC 2.4.2.8) activity, leading to an increase in oxypurines levels, mainly hypoxanthine and uric acid [1,2,3]. Hypoxanthine is the major purine involved in the salvage pathway of purines in the brain, and studies show that it increases five times in cerebrospinal fluid of LND patients when compared to normal individuals [4,5,6]. It has been known that patients affected by this disease present prominent basal ganglia and dopaminergic alterations [7, 8]. There are reports showing that patients with LND can present cerebral palsy, cognitive and behavioral disturbances, motor dysfunction (spasticity, dystonia), self-mutilation behavior, and hyperuricemia [9,10,11,12,13]. Recently, studies showed that elevated levels of hypoxanthine lead to neuronal development aberrations probably by its action on adenosine transport and receptors, dysregulating some early neuronal differentiation gene expressions [14,15,16]. However, the physiopathologic mechanisms of LND are still not clear.

Previous reports show that hypoxanthine could lead to reactive oxygen species increase and exacerbate oxidative stress response [17,18,19]; moreover, these processes may be associated with the brain energy metabolism impairment, since the enzymes of the electron transport chain (ETC) are susceptible to reactive species [20]. Although the brain represents only 2% of the body weight, its energy requirements are very high, and solid regulatory mechanisms operate to provide proper delivery of energy substrates for the neuronal activity [21, 22]. Therefore, the brain is extremely sensitive to disruption of energy resources, being the most metabolically active organ in the human body. Mitochondria have a fundamental function in cellular energy metabolism. A portion of the free energy derived from the oxidation of food is transformed to ATP inside of the mitochondria, in a depending oxygen process. When oxygen rate is restricted, glycolytic products are metabolized directly in the cytosol by the less efficient anaerobic respiration, being independent of mitochondria. The mitochondrial ATP production relies on the ETC, composed of respiratory chain complexes I–IV [23,24,25]. Accordingly, the maintenance of ATP levels is very important for physiological neuronal activity. Impaired energy metabolism may trigger pro-apoptotic signaling (programmed cell death), oxidative stress, and damage to lipids, protein, and DNA [26], leading to conditions that affect the central nervous system (CNS) [27,28,29].

Since the mechanisms by which hypoxanthine affects the CNS still need to be further elucidated and to better comprehend the mechanisms involving hypoxanthine neurotoxicity, in the present study, we investigated the effect of the intrastriatal hypoxanthine administration on neuroenergetic parameters (activities of pyruvate kinase (PK), succinate dehydrogenase (SDH), complex II (CII), cytochrome c oxidase (COX), creatine kinase (CK), and ATP levels) and mitochondrial function (mitochondrial mass and membrane potential) in striatum of 60-day-old rats. Cell death parameters were also evaluated.

Material and Methods

Animals and Reagents

Sixty-day-old male Wistar rats were obtained from the Central Animal House of the Department of Biochemistry of the Universidade Federal do Rio Grande do Sul, Porto Alegre, Rio Grande do Sul, Brazil. Animals were maintained under a standard dark–light cycle at a room temperature of 22 ± 1 °C, with free access to a commercial protein chow and water. All the chemical reagents used for analysis were obtained from Sigma-Aldrich Co., St. Louis, MO, USA. The kit for determination of ATP levels was purchased from PerkinElmer, Waltham, MA, USA.

Ethics Statement

All experimental procedures involving animals were performed in accordance with the NIH Guide for the Care and Use of Laboratory Animals (no. 80-23, revised 1996) and a research project approved by the Ethics Committee of the Universidade Federal do Rio Grande do Sul, Brazil (no. 25717). All efforts were made to minimize the number of animals used and their suffering.

Experimental Model

The animals were anesthetized with ketamine and xylazine (75 and 15 mg/kg i.p., respectively). Based on stereotaxic measures, a 27 gauge 0.9-mm-diameter cannula was positioned and fixed in the rat right striatum as described by Biasibetti and colleagues [19]. The coordinates used for infant and young adult rats were based on Paxinos and Watson’s atlas [30] (coordinates relative from bregma: AP −0.5 mm, ML −2.5 mm, V − 2.5 mm from the dura). The animals were divided into two groups (n = 7 animals per group): (1) control (2 μL of 0.9% saline infusion) and (2) hypoxanthine (2 μL of 10 μM hypoxanthine infusion). Animals were decapitated 30 min after the saline or hypoxanthine infusion; the brain was dissected and the structure analyzed was striatum.

Tissue Preparation

For determination of pyruvate kinase activity, striatum were homogenized with a Teflon-glass homogenizer in five volumes of ice-cold SET buffer (0.32 M sucrose, 1 mM EGTA, 10 mM Tris–HCl), pH 7.4. Afterwards, the homogenate was centrifuged at 800×g and the pellet was discarded. Supernatant was centrifuged at 10,000×g for 10 min. The supernatant solution was collected and used for enzymatic assay. For SDH, CII, and cytochrome c oxidase activity determination, striatum was homogenized (1:20 w/v) in SET buffer (250 mM sucrose, 2 mM EDTA, 10 mM Trizma base, 50 UI/ml heparin), pH 7.4. The homogenates were centrifuged at 800×g for 10 min, and the supernatants were used for enzyme activity determination. For creatine kinase activity, the striatum was washed in SET buffer (0.32 M sucrose/1 mM EGTA/10 mM Tris–HCl; pH 7.4), chopped, and homogenized in the same SET buffer (1:10, w/v). Homogenate was centrifuged at 800×g for 10 min at 4 °C, keeping the supernatant, which was centrifuged at 10,000×g for 10 min at 4 °C. The supernatant of the second centrifugation was used to determine the activity of the CK cytosolic fraction. The sediment was resuspended in Tris–HCl buffer, pH 7.5, containing 100 mM MgSO4 to determine the activity of CK mitochondrial fraction.

Pyruvate Kinase Activity

Pyruvate kinase activity was assayed essentially as described by Leong and colleagues [31]. The incubation medium consisted of 0.1 M Tris–HCl buffer, pH 7.5, 10 mM MgCl2, 0.16 mM NADH, 75 mM KCl, 5.0 mM ADP, 7.0 U of l-lactate dehydrogenase, 0.1% (v/v) Triton X-100, and 10 μL of the mitochondria-free supernatant in a final volume of 0.5 ml. After 30 min of preincubation, the reaction was started by the addition of 1.0 mM phosphoenolpyruvate (PEP) and read by spectrophotometric method. Results were expressed as micromoles of pyruvate formed per minute per milligram of protein.

Succinate Dehydrogenase and Complex II Activities

Immediately before the assay, the samples were frozen and defrost three times. The activities of succinate/phenazine oxidoreductase (succinate dehydrogenase, SDH) and succinate-2,6-dichloroindophenol-oxidoreductase (complex II) were assayed by the method of Fischer and colleagues [32]. This method is based on the decrease in absorbance due to the reduction of 2,6-dichloroindophenol (DCIP) at 600 nm with 700 nm as reference wavelength (å = 19.1 mM−1 cm−1) in the presence of phenazine methosulfate (PMS). The reaction mixture consisting of 40 mM potassium phosphate, pH 7.4, 16 mM succinate, and 8 μM DCIP was preincubated with 40 to 80 μg homogenate protein at 30 °C for 20 min. Subsequently, for complex II activity, 4 mM sodium azide and 7 μM rotenone were added. The reaction was initiated by addition of 40 μM DCIP in a final volume of 1.2 ml and was monitored for 5 min.

Cytochrome c Oxidase Activity

The activity of COX was determined according to Rustin and colleagues [33]. Enzymatic activity was measured by following the decrease in absorbance due to oxidation of previously reduced cytochrome c at 550 nm with 560 nm as reference wavelength (ε = 19.1 mM−1 cm−1). The reaction buffer contained 10 mM potassium phosphate pH 7.0 and 0.6 mM n-dodecyl-β-d-maltoside. The reaction was initiated by the addition of 0.7 μg reduced cytochrome c and monitored for 10 min.

ATP Assay

Structures were homogenized in 1 ml of 0.1 M NaOH. Samples were assayed using the ATPlite Luminescence ATP detection assay system (PerkinElmer, Waltham, MA, USA), in accordance with Schmitz and colleagues [34]. Chemiluminescence was measured using the PerkinElmer Microbeta Microplate Scintillation Analyzer. ATP concentrations were calculated from a standard curve, normalized against wet tissue weights in grams, and expressed as micromoles per gram of tissue.

Creatine Kinase Activity

The reaction mixture consisted in the following medium: 65 mM Tris–HCl buffer (pH 7.5) containing 7 mM of phosphocreatine, 9 mM of MgSO4, and approximately 0.4 to 1.2 mg of protein in a final volume of 0.1 ml. The reaction was started by the addition of 3.2 mM of ADP plus 0.8 mM of reduced glutathione after 5 min of preincubation at 37 °C. After 10 min, the reaction was stopped by the addition of 1 mmol p-hydroxymercuribenzoic acid. The creatine formed was estimated according to the colorimetric method of Hughes [35]. The color was developed by the addition of 0.1 ml of 2% α-naphthol and 0.1 ml 0.05% diacetyl in a final volume of 1 ml, and read after 20 min at 540 nm. Results are expressed as micromoles of creatine per minute per milligram of protein.

Immunocontent of Cytochrome c Oxidase by Western Blotting

Samples were homogenized in a lysis solution containing 2 mM EDTA, 50 mM Tris–HCl, pH 6.8, and 4% SDS, and then dissolved in 25% (v/v) of a solution containing 40% glycerol, 5% mercaptoethanol, and 50 mM Tris–HCl, pH 6.8, and boiled for 5 min. Protein samples were separated by 10% SDS–PAGE (30 μg/lane of total protein) and transferred (Trans-Blot SD Semidry Transfer Cell, BioRad) to nitrocellulose membranes for 1 h at 15 V in transfer buffer (48 mM Trizma, 39 mM glycine, 20% methanol, and 0.25% SDS). The membranes were then washed for 10 min in Tris-buffered saline (TBS) (0.5 M NaCl, 20 mM Trizma, pH 7.5), and then incubated overnight at 4 °C in blocking solution containing the respective antibody cytochrome c oxidase (1:1000) and anti-β-actin (1:1000). The blot was then washed twice for 5 min with T-TBS and incubated for 2 h in antibody solution containing peroxidase-conjugated anti-mouse IgG or peroxidase-conjugated anti-rabbit IgG diluted 1:2000. The blot was washed again twice for 5 min with T-TBS and twice for 5 min with TBS and chemiluminescence developed using a kit (Immobilon Western Chemiluminescent HRP Substrate, Millipore) and detected by ImageQuant LAS 4000 (GE Healthcare Life Sciences).

Mitochondrial Mass and Membrane Potential Analyses

Striata were mechanically dissociated in PBS containing collagenase. To determine the mitochondrial mass and membrane potential (Δψ), dissociated cells were incubated with 100 nm MitoTracker Green and 100 nm MitoTracker Red (MTG and MTR, respectively, diluted from 1 mM stock solutions in DMSO), for 45 min, at 37 °C (in the dark). A relationship was established between MTG and MTR staining for accessing the rate of mitochondrial function [36]. Immediately after staining, cell suspensions were analyzed on a FACS Calibur flow cytometer (Becton–Dickinson, San Jose, CA), using red (670 nm long pass) and green (530 nm/30) filters. Controls stained with individual dyes were used to determine compensation parameters. For each sample, 10,000 events corresponding to intact cells (as gated in FSC vs. SSC plots) were analyzed.

Cell Death Analysis

Striata were mechanically dissociated in PBS containing collagenase. Cells were labeled by incubation with annexin V-FITC and propidium iodide (PI) in a binding buffer (apoptosis detection kit I-556547; BD Pharmingen) for 15 min at room temperature in the dark, according to the manufacturer’s instruction. Stained cells were acquired (10,000 for gated cells) on a FACS Calibur flow cytometer Becton–Dickinson, Franklin Lakes, NJ, USA, using green (530 nm/30) and red (670 nm long pass) filters. Controls stained with individual dyes were used to determine compensation parameters.

Protein Determination

Protein content of samples was determined using bovine serum albumin as standard, according to Lowry [37].

Statistical Analysis

Data were analyzed by Student’s t test for comparison of two means. All analyses were performed using the Statistical Package for the Social Sciences (SPSS) software in a compatible computer. Results are expressed as means ± SD, and differences were considered statistically significant when p < 0.05.

Results

We initially investigated the effect of hypoxanthine administration on the enzymatic activity of glycolytic pathway performed by the measurement of PK activity, a crucial enzyme of glycolysis pathway. Figure 1a shows no substantial difference in PK activity between the hypoxanthine and control groups (p > 0.05).

Fig. 1
figure 1

Effect of hypoxanthine intrastriatal administration on pyruvate kinase (a), succinate dehydrogenase (b), and complex II (c) activities in striatum of 60-day-old rats. Data are mean ± SD for six to seven animals in each group. Data from pyruvate kinase are expressed as μmol of pyruvate per mg of protein; data from succinate dehydrogenase and complex II are expressed as nmol per min per mg of protein. Different from control, *p < 0.05, **p < 0.01 (Student’s t test). Hpx hypoxanthine, SDH succinate dehydrogenase

Full size image

Further on tricarboxylic acid (TCA) cycle and respiratory chain pathway, we analyzed SDH and complex II activities. As seen in Fig. 1b, c, hypoxanthine injection was able to significantly increase the activity of SDH and complex II (p < 0.01, p < 0.05, respectively) when compared to controls. In addition, we analyzed cytochrome c oxidase, the rate-limiting enzyme of oxidative phosphorylation. Figure 2a, b shows that hypoxanthine significant reduces activity and immunocontent of this enzyme (p < 0.001, p < 0.01, respectively). Figure 2c shows that ATP levels were significantly decreased by hypoxanthine (p < 0.05).

Fig. 2
figure 2

Effect of hypoxanthine intrastriatal administration on cytochrome c oxidase activity (a) and immunocontent (b), as well as ATP levels (c) in striatum of 60-day-old rats. Data are mean ± SD for six to seven animals in each group. Data from cytochrome c oxidase activity are expressed as nmol per min per mg of protein; data from western blot are expressed as percentage of control. Uniformity of gel loading was confirmed with β-actin as standard. Data from ATP levels are expressed as μmol per gram of tissue. Different from control, *p < 0.05, **p < 0.01, ***p < 0.001 (Student’s t test). Hpx hypoxanthine

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Since CK system (cytosolic and mitochondrial isoenzymes) can contribute to the sustenance of ATP levels, we also evaluated the effect of hypoxanthine administration on this system. As illustrated in Fig. 3a, b, hypoxanthine significantly decreased the activities of cytosolic and mitochondrial CK (p < 0.001, p < 0.05, respectively).

Fig. 3
figure 3

Effect of hypoxanthine intrastriatal administration on cytosolic creatine kinase (a) and mitochondrial creatine kinase (b) activities in striatum of 60-day-old rats. Data are mean ± SD for six to seven animals in each group. Data are expressed as μmol of creatine per minute per mg of protein. Different from control, *p < 0.05, ***p < 0.001 (Student’s t test). Hpx hypoxanthine, CK creatine kinase

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To better elucidate the effect of hypoxanthine on mitochondrial mass and membrane potential (Δψ), we labeled striatal cells with the fluorescent dyes MTG and MTR, respectively. Hypoxanthine injection decreased the percentage of cells labeled with MTG (p < 0.01) and increase MTR labeling (p < 0.05). Therefore, the ratio between MTG and MTR was diminished (p < 0.001) in hypoxanthine group as compared to controls (Fig. 4a).

Fig. 4
figure 4

Effect of hypoxanthine intrastriatal administration on mitochondrial and cell death evaluation. Mitochondrial mass and membrane potential (a) and annexin-PI assay (b) in striatum of 60-day-old rats. Data are mean ± SD for six to seven animals in each group. Different from control, *p < 0.05, **p < 0.01, ***p < 0.001 (Student’s t test). Hpx hypoxanthine, MTG MitoTracker Green, MTR MitoTracker Red

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Given that mitochondria have a central role in cell viability and ATP can regulate cell death by apoptosis [38], we labeled striatal cells with annexin V and PI and analyzed these cell populations by flow cytometry to examine the effects of hypoxanthine administration on cell death parameters in striatum of 60-day-old rats. Hypoxanthine decreased the number of living cells (annexin V−/PI−) (p < 0.05), increasing apoptotic events (annexin V+/PI−) in striatum (p < 0.001) (Fig. 4b). Hypoxanthine administration did not alter the number of necrotic cells (annexin V−/PI+) in striatum of 60-day-old rats (p > 0.05) (Fig. 4b).

Discussion

Hypoxanthine, the major purine metabolite involved in the purine salvage pathway in the brain, is increased more than five times that its regular levels in plasma of patients with LND, which is originated by mutations in the HPRT1 gene, responsible for encoding the purines recycling enzyme HPRT [6, 9, 39]. A clinical study showed that hypoxanthine causes adenosine, dopamine, and serotonin receptors imbalance and alters various transcription factors related to early development of neuronal dopamine [14, 40]. In addition, preclinical studies showed that intrastriatal hypoxanthine administration induces oxidative stress, inflammation, and memory deficits in the brain of rats [18, 19, 41]. Considering that the hypoxanthine neurotoxicity and the physiopathology of LND still needs to be further elucidated, in the present study, we investigated the effect of intrastriatal hypoxanthine injection on neuroenergetic parameters, mitochondrial function, and cell death in striatum of rats.

Firstly, we investigated the effect of hypoxanthine administration on PK, which is a rate-regulating glycolytic enzyme that catalyzes the irreversible conversion of PEP to pyruvate [42]. Results showed that hypoxanthine did not alter PK activity in striatum of 60-day-old rats (Fig. 1a). We also evaluated the activity of SDH, an enzyme of the TCA cycle located in the inner mitochondrial membrane where it can directly transfer electrons into the ETC [24]. Results showed that hypoxanthine significantly increased SDH activity in striatum of 60-day-old rats. Inhibition of this enzyme is also observed in other disorders, such as hyperprolinemia, sarcosinemia, Huntington’s disease, Alzheimer’s disease, and multiple sclerosis [43,44,45,46,47].

We analyzed complex II of the ETC, which is an essential component of oxidative phosphorylation and is coupled to the synthesis of ATP from ADP and Pi [24]. Accordingly to SDH activity, we observed an increase in complex II activity (Fig. 1c), which catalyzes the oxidation of matrix succinate to fumarate, feeding electrons to the cytochrome complex. The regulation of complex II activity is not fully comprehended, despite the involvement of this enzyme in several conditions, recently emerging as an important point of investigation in cell signaling, cancer biology, immunology, cardiovascular conditions, and neurodegeneration [48]. It is well known that glucose is an essential and prevalent energy substrate for the adult brain under physiological condition; however, there are alternative substrates such as ketone bodies and amino acids that may also be used under certain circumstances [24]. Since PK is not altered in the present study, one possible explanation to increase in SDH and complex II could be by anaplerotic routes.

Since cytochrome c oxidase is a rate-limiting enzyme of the mitochondrial respiratory chain, we also analyzed the effect of hypoxanthine on activity and immunocontent of this enzyme. Results showed that hypoxanthine significantly decreased the activity and immunocontent of cytochrome c oxidase (Fig. 2a, b). Our results are similar to those found in preclinical and clinical studies showing that SDH activity was increased, whereas cytochrome c oxidase activity was decreased in cerebral cortex of mice exposed to chronic hypoxia [49] and in brain of patients with Alzheimer’s disease [46]. Additionally, some studies have shown that an increase in SDH activity could correlate with stimulation of anaerobic pathways coupled with reverse operation of TCA cycle towards the reduction of oxaloacetate to succinate when oxidative phosphorylation is impaired [50]. In addition, it has been shown that the increase in SDH and complex II activities and inhibition of cytochrome c oxidase activity can potentially lead to incomplete reduction of oxygen and consequently to increase free radical formation and reduce ATP synthesis [20, 51,52,53].

Accordingly with cytochrome c oxidase results, we also measured ATP levels and observed a decrease in this parameter in striatum of hypoxanthine group when compared to control (Fig. 2c). Thus, in the present study, we observed a reduction of the ETC flow (impaired oxidative phosphorylation) caused by hypoxanthine, resulting in lower ATP production, which is necessary to sustain energy requirement of cells. At cellular level, ATP reduction leads to neuron depolarization causing failure of membrane ion transport. Interventions for improving mitochondrial function by maintaining ATP levels may have solid influence in preventing neuronal dysfunction and loss [54]. Intracellular ATP levels lower than 10% of the initial value result in cell death [54], and in the present study, hypoxanthine decreased ATP levels in approximately 25%.

Coupling spatial intracellular ATP-producing and ATP-consuming is a fundamental process to cell function. CK is a major phosphotransfer system in cells with high-energy demand. CK is responsible for the relocation of ATP phosphoryl group from where it is produced (mostly in mitochondria) to where it is consumed (mainly in the cytosol) [55]. Hypoxanthine administration decreased cytosolic and mitochondrial CK activities (Fig. 3). Considering that some enzymes involved in energy metabolism require sulfhydryl groups and that previous studies of our group showed that hypoxanthine alters membrane protein thiol content and induces oxidative stress [17, 19], the present study showed a decreased in ATP availability, which could lead to a possible accumulation of debris and ultimately cell death by oxidative stress [56].

Mitochondrial dysfunction affects metabolism function, increases radical formation contributing to oxidative stress, and induces apoptosis, which may lead to cellular and ATP imbalance [25, 28]. Some evidence suggest that abnormalities in mitochondria are involved not only in aging, neurodegenerative diseases, cancer, and several other mitochondrial diseases [57, 58] but also in inborn errors of metabolism [43, 59, 60]. To better comprehend the effect of hypoxanthine on mitochondrial function, we analyzed mitochondrial mass and membrane potential (Δψ). Hypoxanthine administration decreased the percentage of cells labeled with MTG and increased MTR labeling when compared to controls (Fig. 4a). Mitochondrial membrane potential is considered a key indicator of mitochondrial function [61], which is in accordance with the increase in SDH and complex II activities, and modifications in these parameters suggest mitochondrial dysfunction [62].

Since mitochondrial damage can initiate signaling pathways of intrinsic/extrinsic apoptosis and regulated necrosis [63], cell redox state can also collaborate to apoptosis, and reactive species produced by mitochondria are involved in cellular damage [64], we analyzed the effect of hypoxanthine on striatal cells with annexin V and PI labeling. Figure 4b shows that hypoxanthine decreased the number of living cells (annexin V−/PI−), consequently increasing apoptotic events (annexin V+/PI−) in striatum. The number of necrotic cells (annexin V−/PI+) was not altered by hypoxanthine administration in striatum of 60-day-old rats.

In summary, the present results shed light into the mechanisms of hypoxanthine-induced neurotoxicity in striatum of 60-day-old rats. Intrastriatal hypoxanthine administration caused neuroenergetic impairment resulting in ATP depletion which can lead to mitochondrial dysfunction and further cell death by apoptosis. This study provides new enlightening basis of hypoxanthine neurotoxicity mechanisms on energetic and cell viability parameters, suggesting that this processes may be involved, at least in part, with disorders found in patients with LND.