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Copper (Cu) is essential for proper brain development, particularly the cerebellum, and functions as a cofactor for enzymes including mitochondrial cytochrome c oxidase (CCO). Cu deficiency severely limits CCO activity. Augmented lactate in brain of Cu deficient (Cu−) humans and cerebella of Cu− rats is though to originate from impaired mitochondria. However, brain lactate may also originate from elevated blood lactate. The hypothesis that cerebellar lactate originates from elevated blood lactate in Cu− rat pups was tested. Analysis of Cu− and Cu adequate (Cu+) rat pups (experiment I) revealed blood lactate was elevated in Cu− rat pups and cerebellar lactate levels were closely correlated to blood lactate concentration. A second rat experiment (experiment II) assessed Cu− cerebellar lactate without the confounding factor of elevated blood lactate. Blood lactate levels of Cu− rat pups in experiment II were equal to those of controls; however, Cu− cerebellar lactate was still elevated, suggesting mitochondrial impairment by Cu deficiency. Treatment of rat pups with dichloroacetate (DCA), an activator of mitochondrial pyruvate dehydrogenase complex (PDC), lowered Cu− cerebellar lactate to control levels suggesting PDC inhibition is a site of mitochondrial impairment in Cu− cerebella. Results suggest Cu− cerebellar lactate originates from blood and cerebellum.
Copper (Cu) is a cofactor for a number of important enzymes and is an essential micronutrient for proper mammalian development (Prohaska 2006). In humans, deficiency of Cu occurs in Menkes’ disease, and leads to a number of pathologies including severe mental retardation and abnormal and slow brain development especially in cerebellum (Danks et al. 1972; Donsante et al. 2007;Menkes etal. 1962; Mercer, 1998). Rodent models of Menkes’ disease exhibit parallel symptoms (Prohaska and Wells 1974). Cu deficient (Cu−) rat cerebella exhibit blunted development including reduced myelination and synaptogenesis as well as motor-function disturbances (El Meskini et al. 2007; Everson et al. 1967; Penland and Prohaska 2004; Prohaska and Wells 1974; Prohaska 1981; Zimmerman et al. 1976). However, underlying causes of these symptoms have been poorly understood.
A prevalent explanation for the effect of Cu deficiency on brain development is impaired brain energy metabolism due to mitochondrial dysfunction (Mercer 1998). Cu is a co-factor for the mitochondrial enzyme cytochrome c oxidase (CCO), complex IV of the mitochondrial electron transport chain. CCO activity and protein levels are acutely sensitive to Cu deficiency (Cohen and Elvehjem 1934; Gybina and Prohaska 2006). Electron micrographs of Cu− rat brains have shown morphologically abnormal mitochondria (Prohaska and Wells 1975). Brain tissue and cerebrospinal fluid of Menkes patients exhibit elevated lactate levels, considered a classic sign of mitochondrial impairment (Kodama et al. 1999; Loyola and Dodson 1981; Munakata et al. 2005). Metabolite extracts of Cu− rat pup brains and cerebella also show greatly elevated lactate concentrations (Gybina and Prohaska 2008b; Prohaska and Wells 1975).
Elevated brain lactate levels can result from the failure of inhibited brain mitochondria to metabolize glycolytically generated pyruvate. High brain lactate concentrations, and associated acidosis, can have serious metabolic ramifications like inhibition of brain glucose metabolism (Kuschinsky et al. 1981; Miller et al. 1984; Thurston et al. 1983). Recent studies of Cu− rat cerebella with high lactate concentrations have shown evidence of such inhibition (Gybina and Prohaska 2008a). However, previous analyses of Cu− rats have not considered blood-to-brain lactate influx as one additional source of cerebellar lactate. Lactate can move bi-directionally across the blood brain barrier (BBB) down a lactate concentration gradient (Cremer et al. 1979; LaManna et al. 1993; Nemoto et al. 1974; Oldendorf 1971). Elevated blood lactate has previously been detected in Menkes patients (Loyola and Dodson 1981; Munakata et al. 2005; Rizzo et al. 2000). However, blood lactate levels and its effects on brain and cerebellar lactate concentrations have never been assessed in animal models of Cu deficiency.
Furthermore, previous assessments of Cu− brain mitochondrial dysfunction have not accounted for brain lactate accumulation. Cu− rat brain shows evidence of increased NADH to NAD+ ratio (NADH/NAD+), which can inhibit pyruvate dehydrogenase complex (PDC) via activation of pyruvate dehydrogenase kinase (PDK) (Prohaska and Wells 1975; Sugden and Holness 2003). PDC inhibition reduces mitochondrial pyruvate uptake and results in elevated lactate. However, PDC function has never been assessed in Cu− rat brain.
The present work evaluated the possibility that elevated cerebellar lactate may reflect elevated plasma lactate in Cu− animals. Cerebellar and plasma lactate concentrations were determined and correlations between them were investigated. Potential PDC inhibition in Cu− brain mitochondria was also evaluated by treating Cu− rat pups with dichloroacetate (DCA), a PDC activator, and characterizing subsequent cerebellar lactate concentrations.
Holtzman sperm-positive rats were purchased commercially (Harlan Sprague Dawley, Indianapolis, IN) and received either Cu-adequate (Cu+) or Cu-deficient (Cu−) dietary treatment consisting of a Cu−deficient modified AIN-76A diet (Teklad Laboratories, Madison, WI) that contained 0.34 mg Cu/kg by analysis (Experiment I) or 0.43 mg Cu/kg (Experiment II). Normal AIN-76A diet contains approximately 6 mg Cu/kg. All dams and offspring were fed the Cu− diet. Cu+groups drank water supplemented with cupric sulfate, 20 mg Cu/L, and Cu− groups drank deionized water (Gybina and Prohaska 2003).
Offspring in this perinatal model of Cu deficiency were sampled at postnatal age 24 (P24) and again at P30 after a 48 hour fast (Experiment I). Fasting (Experiment I) was initiated for Cu+ and Cu− rat pups at P28 and continued for 48 hours. During this period control (non-fasted) rats, Cu+ and Cu−, continued to feed ad libitum. Control pups were age, weight, and litter matched to their fasted Cu+ or Cu− counterparts. Both fed and fasted groups had ad libitum access to drinking water.
The Cu dietary paradigm was repeated for rats intended for DCA treatment (Experiment II), and rats were sampled at P23 to confirm Cu deficient status and determine plasma and cerebellar lactate levels. Pups were sampled again at P30 after DCA treatment. DCA (100 mg/kg) or saline, the control, was administered to rats by IP injection once daily starting at P25 and ending on P30, a total of six injections. Saline injected control pups were age, weight, and litter matched to their DCA Cu+ or Cu− counterparts. DCA solutions contained 25 mg/ml DCA in saline brought to pH 7 using NaOH.
No mortality was observed in Cu− rats during or after fasting or during or after DCA treatment. Animals appeared normal compared to their dietary counterparts. Each experiment (Experiment I and II) sampled rats from a minimum of four separate litters of each treatment group. All animals were maintained at 24°C with 55% relative humidity on a 12-h light cycle (0700-1900-h). All protocols were formally approved by the University of Minnesota Animal Care Committee.
To prevent changes in metabolite concentrations induced by anesthetics, animals were decapitated without anesthesia. To minimize stress experienced by animals from handling before decapitation, rats were handled daily from P0 to day of tissue collection. Upon decapitation, cerebella were immediately removed from the skull and frozen in liquid nitrogen. Only cerebella frozen in under 15 seconds from the time of decapitation were analyzed. Cerebellar extraction time did not differ between Cu+ and Cu− animals. Fast frozen cerebella were stored at −75°C until analysis. Remaining brain and a piece of liver tissue were collected for total Cu analysis. Upon decapitation, trunk blood was collected in heparinized tubes and spun immediately to obtain plasma, which was rapidly frozen and stored at −75°C until analysis.
Brains, without cerebella, were wet-digested with HNO3 (Trace Metal grade; Fisher Scientific, Pittsburgh, PA) and samples were analyzed for total Cu content by flame atomic absorption spectroscopy (Model 1100B, Perkin-Elmer, Norwalk, CT). Protein content of tissue samples was determined using a modified version of the Lowry method (Markwell et al. 1978).
Plasma was collected from heparinized blood and used to evaluate metabolite concentrations. Cerebellar metabolite extracts were prepared and analyzed according to Lowry et al. with some modifications (Lowry and Passonneau 1972). Briefly, fast frozen tissues were powdered under liquid nitrogen and then transferred to tubes, chilled on dry ice, and mixed with 5 volumes of 0.7 M HClO4. Tubes were then transferred to an −8°C alcohol bath, where samples and acid were mixed. This mixture was then rapidly homogenized using a tissue probe cooled to 4°C. Homogenates were spun and the supernatant neutralized with KHCO3 to yield the final cerebellar metabolite extract. Plasma and cerebellar extract metabolite concentrations were determined using enzymatic analyses as previously described (Lowry and Passonneau 1972). Plasma β-hydoxybutyrate concentrations were determined using instructions and kit purchased from Stanbio Laboratory (Boerne, TX).
Samples for western blot analysis of copper chaperone for superoxide dismutase (CCS) and CCO subunit IV (COX IV) were prepared by homogenizing fast frozen cerebella in 9 volumes phosphate buffered saline (PBS) pH 7.4 with 0.5% TritonX-100. Homogenates were spun at 15,000 × g for 15 minutes at 4°C and supernatant protein samples saved for analysis after an addition of protease inhibitors (Protease Inhibitor Cocktail, Sigma, St. Louis, MO). CCS and COX IV protein analysis was carried out by fractionation of 23μg protein on a 15% SDS-PAGE gel. Proteins were transferred to 0.2 μm nitrocellulose membranes and processed for immunoblotting as described elsewhere (Prohaska and Brokate 2001).
Cerebellar protein levels of CCS were evaluated using affinity purified rabbit anti-hCCS characterized previously, at a 1:1000 dilution (West and Prohaska 2004). COX IV protein levels were analyzed using mouse monoclonal anti-COX IV, at 1:4000 dilution (Molecular Probes, Eugene, OR). All membranes were blocked for a least an hour using 5% (w/v) albumin/Tris buffered saline (TBS) containing 0.1% Triton X-100, and incubated overnight with primary antibodies at 4°C. All secondary species-specific antibodies were diluted 1:10,000. SuperSignal West Pico chemiluminescent substrate (Pierce, Rockford, IL) was used for detection. Chemiluminescence was captured using high speed blue X-ray film (Lake Superior X Ray Inc., Duluth, MN) and densitometry was carried out using the FluorChemTM system (Alpha Innotech, San Leandro, CA).
Means ± SEM were calculated. Student’s unpaired two-tailed t-test was used to compare data between the two dietary treatments, α=0.05. Variance equality was evaluated by F-test. All data were processed using Microsoft Excel™. Data from fasting and DCA treatment studies were analyzed using one-way ANOVA (Student-Newman-Keuls multiple comparison). Correlation coefficients were calculated using Kaleidagraph™ analysis software. For comparative purposes immuno-blot data was normalized. A value of 1.0 was assigned to the mean pixel density of Cu+ samples then all, both Cu+ and Cu−, individual densities were recalculated before graphing.
Following induction of perinatal Cu deficiency (Experiment I), P24 Cu− rat pups exhibited lower body weight as compared to controls (76.9±1.1 g Cu+ vs 58.8±2.3 g Cu−; P<0.01). Cu deficiency also resulted in 93% lower Cu levels in the liver (6.15±0.89μg/g Cu+ liver vs 0.42±0.02μg/g Cu−; P<0.01) . Cu levels in the brain of Cu− rat pups were 81% lower (2.30±0.06μg/g Cu+ brain vs 0.44±0.04μg/g Cu−; P<0.01). The degree of Cu deficiency exhibited by these rats is very similar to Cu deficiency induced with previous work with this model (Gybina and Prohaska 2003; Gybina and Prohaska 2008b).
Evaluation of P24 cerebellar lactate showed it to be 225% higher in Cu− cerebella confirming previous observations that cerebellar lactate is elevated by Cu deficiency in rat pups (Fig. 1) (Gybina and Prohaska 2008b). Analysis revealed plasma lactate to be elevated in Cu− pups by 224%, bearing a striking similarity to the degree of elevation observed in Cu− cerebella (Fig. 1). Blood lactate concentration in Cu− rat pups, 11μmol/ml, exceeded the concentration of cerebellar lactate, 4.5μmol/g, indicating a concentration gradient favorable to lactate movement from blood to brain.
The possibility that blood lactate was responsible for observed Cu− cerebellar lactate was more thoroughly investigated. One approach to test the relationship between blood and cerebellar lactate in Cu− pups is to assess cerebellar lactate at various blood lactate concentrations. Elevated blood lactate in Cu− rats suggests mitochondrial impairment occurs in some Cu− tissues. During mitochondrial inhibition blood lactate is largely a bi-product of cellular anaerobic glycolysis and thus lactate production is heavily dependent on glucose availability. Therefore, blood lactate concentration was modulated by reducing blood glucose availability. This was accomplished by fasting Cu+ and Cu− rat pups.
P28 pups were fasted for 48 hours and assessed at P30 for markers of Cu deficiency and of fasting. Western blots of cerebellar homogenates were probed for the Cu deficiency markers COX IV and copper chaperone for superoxide dismutase (CCS), which confirmed Cu deficiency. COX IV protein levels were reduced by 68% in Cu− cerebella, while CCS, which increases in quantity when cellular Cu concentration is low (Prohaska et al. 2003), was 172% higher in P30 rat Cu− cerebellum, consistent with cerebellar Cu deficiency (Fig. 2A). Blood levels of β-hydroxybutyrate, a fasting marker, were significantly higher in fasted pups (Fig. 2B). Plasma glucose concentration was significantly lower in fasted rats (Fig. 2C). Cu+ fasted rat pups experienced 50% lower blood glucose levels and Cu− fasted pups had 31% lower blood glucose.
Plasma lactate levels were measured to determine whether lowered blood glucose in fasted Cu− pups was successful in modulating blood lactate concentrations. Plasma lactate concentration was significantly higher in fed Cu− animals compared to fed Cu+ controls (Fig. 3A). Fed Cu− pups displayed little variability in plasma lactate concentration. Fasting had no significant impact on blood lactate of Cu+ rat pups. Fasting lowered Cu− blood lactate to below fed Cu− control average in all but one animal (Fig. 3B and C), and the reduction trended towards significance (P=0.089). Fasting also introduced greater variability in Cu− pups as compared to Cu− fed controls (Fig. 3C) and was therefore successful in generating a wider range of blood lactate concentrations in Cu− pups.
As seen previously at P24, cerebellar lactate at P30 was significantly higher in fed Cu− pups compared to Cu+ pups (Fig. 3B). Fasting significantly reduced cerebellar lactate concentration in Cu− pups as compared to fed Cu− controls. Fasting had no impact on cerebellar lactate in Cu+ animals. Cu− rat pup cerebellar lactate concentrations were highly correlated to blood lactate concentration (R=0.99) (Fig. 3C).
Experimental data from fasted rat pups suggest that cerebellar lactate concentration is at least in part responsible for observed lactate concentration in Cu− rat pups. This evidence, however, did not clarify whether Cu− cerebellar mitochondrial impairment also contributed to cerebellar lactate concentrations. To answer this question, an additional rat experiment was conducted (Experiment II) in an attempt to evaluate Cu− cerebellar lactate concentration in the absence of the confounding variable of elevated blood lactate. Cu deficiency was induced, as evidenced by rat brain Cu levels (Fig. 4A). However, compared to the preceding animal experiment, as well as other past experiments, Cu deficiency induced in this rat experiment appeared to be less severe. Notably, no change in P23 animal body weights was observed (Fig. 4B) and Cu− blood lactate levels were not different from Cu+ controls (Fig. 4C). Importantly, a modest but significant 40% elevation in lactate was still detected in Cu− cerebella (Fig. 4D).
To further discern the source of metabolic impairment in Cu− cerebellar mitochondria, P25 rats were treated for five days with dichloroacetate (DCA), an activator of PDC. Western blots of control Cu+ and Cu− P30 cerebellar homogenates were probed for COX IV and CCS which confirmed Cu deficiency (Fig. 5A). Compared to saline injected Cu+ and Cu− pups, Cu+ and Cu− DCA treated animals exhibited more than 50% lower blood lactate (Fig. 5B), indicating that DCA was effective at increasing PDC activity in rat tissues. As observed at P23, P30 saline injected Cu− rat blood lactate levels were the same as Cu+ saline injected controls (Fig. 5B). Similar to P23 Cu− rats, saline injected P30 Cu− rats had cerebellar lactates that were significantly elevated (Fig. 5C). DCA treatment did not alter Cu+ cerebellar lactate levels. However, DCA treatment in Cu− pups reduced cerebellar lactate concentrations to Cu+ control levels. Together this data suggests that PDC is inhibited in Cu− cerebellar mitochondria in vivo.
Cu deficient Menkes patients exhibit Cu deficiency dependent augmented lactate levels in brain, cerebrospinal fluid, and blood (Kodama et al. 1999; Loyola and Dodson 1981; Munakata et al. 2005; Rizzo et al. 2000). High lactate levels in whole brain and cerebella are also observed in rodent models of Menkes disease (Gybina and Prohaska 2008b; Prohaska and Wells 1975). Cerebellum was the brain region focused upon in the current studies since this region is especially impacted in infants with Menkes disease (Danks et al. 1972). Also our recent studies in rodent models have focused on cerebellum. Blood lactate and its contribution to brain lactate has not been previously evaluated in Cu deficiency rodent models. Experiment I explored the possibility that blood lactate contributes to high lactate levels in Cu− rat pup cerebella. A typical level of Cu deficiency in pups was induced and was accompanied by lower tissue Cu as well as lower body weights; P24 Cu− pups were confirmed to have elevated cerebellar lactate. Cu− blood lactate concentration was also found to be elevated. This supported the possibility that cerebellar lactate may be a reflection of blood lactate levels in Cu− rats. Further evaluation of the cerebellar lactate at various blood lactate levels showed a strong correlation between cerebellar and blood lactate levels in Cu− rats. Together this data suggests that Cu− rat cerebellar lactate levels are influenced by blood lactate concentrations.
The contribution of blood lactate to elevated cerebellar lactate levels raised the question of whether all of the elevated lactate concentration in Cu− rat cerebella is attributable to augmented blood lactate levels rather then compromised Cu− cerebellar mitochondria. To answer the latter question, a second rat experiment (Experiment II) was conducted. Pups in experiment II were affected by Cu deficiency less than is typically seen. Lower tissue Cu levels confirmed Cu deficiency, but the Cu− pups did not experience lower body weights at P23 as would normally be seen. Blood lactate levels also remained unaffected, suggesting peripheral tissue Cu deficiency and consequent mitochondrial inhibition was not yet great enough to elevate blood lactate. However, despite the absence of elevated blood lactate, lactate levels in Cu− cerebella were still significantly higher. As blood lactate could not have contributed to the lactate increase observed in Cu− rat cerebella, these observations suggest some intrinsic mitochondrial impairment in Cu− cerebella contributes to cerebellar lactate levels.
DCA treatment further characterized the nature of mitochondrial impairment in Cu− cerebella. DCA is a powerful inhibitor of PDK, which inactivates PDC (Sugden and Holness 2003). By preventing PDC inhibition, DCA allows pyruvate to enter the citric acid cycle thereby preventing lactate accumulation. DCA is used clinically to lower body lactate concentration and has been shown to modulate brain lactate levels in rats (Henderson et al. 1997;Miller et al. 1990). DCA administration lowered blood lactate levels in both Cu+ and Cu− animals compared to dietary controls. Though DCA treatment did not significantly alter Cu+ cerebellar lactate levels, DCA treatment of Cu− pups was able to lower Cu− cerebellar lactate to Cu+ control levels. It is unlikely that lower blood lactate in Cu− DCA treated pups played a significant role in reducing cerebellar lactate because Cu+ animals experienced the same reduction in blood lactate as Cu− rats but only DCA treated Cu− pups experienced lower cerebellar lactates. Together this evidence points to significant PDC inhibition in Cu− cerebellar mitochondria and can explain the observed lactate accumulation.
Brain mitochondria appear to be quite sensitive to Cu deficiency. Mouse pup studies show that brain is the first tissue to show high lactate during Cu deficiency (Rusinko and Prohaska 1985). However up to now much of our understanding of Cu deficiency’s impact on function of brain mitochondria was based on in vitro respiratory studies (Prohaska and Wells 1975). Those studies showed that when glutamate was used as substrate in Cu− brain mitochondria, state 3 respiration was reduced by 30% and overall RCR by 60%; no changes were detected with succinate as a substrate. However, this previous evidence failed to explain the observed lactate accumulation in Cu− rat brain.
Our new evidence for PDC inhibition and mitochondrial dysfunction in vivo adds to and may help interpret previous data. Increased NADH/NAD+ activates pyruvate dehydrogenase kinase (PDK) which then inhibits PDC. The ability of DCA to attenuate cerebellar lactate levels supports previous observations of increased NADH/NAD+ in Cu− brain (Prohaska and Wells 1975). Increased NADH/NAD+ may also explain the disparate respiration results between glutamate and succinate. Mitochondrial glutamate is converted to α-ketoglutarate, which is then converted to succinyl-CoA and drives the generation of NADH from NAD+; this reaction may be inhibited in Cu− mitochondria due to increased NADH/NAD +. On the other hand, mitochondrial consumption of succinate is independent of NADH metabolism and therefore may remain unaffected. The increased NADH/NAD+ is likely a reflection of the mitchondrial electron transport chain’s inability to consume NADH due to CCO inhibition during Cu deficiency. Interestingly, despite evidence of mitochondrial dysfunction, ATP concentration in Cu− brain and cerebella is maintained at control levels, though the mechanism for this phenomenon is not known (Gybina and Prohaska 2008a;Gybina and Prohaska 2008b;Prohaska and Wells 1975).
Together, evidence presented in these studies shows that severe Cu deficiency generates high blood lactate and this exogenous lactate can affect the observed lactate levels in the Cu− rat cerebellum. However, lactate elevation also stems from within the cerebellar tissue itself and is likely due to cerebellar mitochondrial PDC inhibition. Future studies of Cu− brain metabolism will need to take into account lactate generated by peripheral Cu− tissues and its potential consequences for brain metabolism. Lactate can have inhibitory affects on glucose metabolism (Kuschinsky et al. 1981; Miller et al. 1984; Rovetto et al. 1975; Thurston et al. 1983), and there has been some evidence for inhibited glucose metabolism in Cu− rat pup cerebella (Gybina and Prohaska 2008a). Furthermore, the metabolism of the developing rat brain is complicated by the developmental shift from uptake and metabolism of a combination of glucose, lactate, and fuels prevalent in milk (ketones) to near exclusive glucose metabolism (Cremer et al. 1979; Lust et al. 2003; Nehlig 1999). Therefore, the precise effect on mitochondrial inhibition and high lactate levels on the metabolism of the developing rat brain remains to be established. Much more work remains to be done in the arena of metabolic disturbances in developing brain and dietary and genetic Cu deficiency.
We thank Margaret Boderius and Joshua Pyatskowit for their excellent technical assistance. This research was supported by NIH HD-039708 and the University of Minnesota Doctoral Dissertation Fellowship.