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Edited* by Robert G. Shulman, Yale University School of Medicine, New Haven, CT, and approved July 23, 2013 (received for review April 1, 2013)
Abstract
It has been reported that chronic and acute alcohol exposure decreases cerebral glucose metabolism and increases acetate oxidation. However, it remains unknown how much ethanol the living brain can oxidize directly and whether such a process would be affected by alcohol exposure. The questions have implications for reward, oxidative damage, and long-term adaptation to drinking. One group of adult male Sprague–Dawley rats was treated with ethanol vapor and the other given room air. After 3 wk the rats received i.v. [2-13C]ethanol and [1, 2-13C2]acetate for 2 h, and then the brain was fixed, removed, and divided into neocortex and subcortical tissues for measurement of 13C isotopic labeling of glutamate and glutamine by magnetic resonance spectroscopy. Ethanol oxidation was seen to occur both in the cortex and the subcortex. In ethanol-naïve rats, cortical oxidation of ethanol occurred at rates of 0.017 ± 0.002 µmol/min/g in astroglia and 0.014 ± 0.003 µmol/min/g in neurons, and chronic alcohol exposure increased the astroglial ethanol oxidation to 0.028 ± 0.002 µmol/min/g (P = 0.001) with an insignificant effect on neuronal ethanol oxidation. Compared with published rates of overall oxidative metabolism in astroglia and neurons, ethanol provided 12.3 ± 1.4% of cortical astroglial oxidation in ethanol-naïve rats and 20.2 ± 1.5% in ethanol-treated rats. For cortical astroglia and neurons combined, the ethanol oxidation for naïve and treated rats was 3.2 ± 0.3% and 3.8 ± 0.2% of total oxidation, respectively. 13C labeling from subcortical oxidation of ethanol was similar to that seen in cortex but was not affected by chronic ethanol exposure.
The liver is the major organ for the oxidation of ethanol (Etoh) (1, 2), followed by the stomach and other organs. Acetate (Ac) generated from Etoh by the liver is consumed by the brain (3, 4), where it replaces a significant portion of cerebral glucose metabolism in humans and animals (5⇓⇓–8), either decreasing glucose consumption directly or compensating for glucose consumption decreased by some other effect. However, the brain may also oxidize Etoh (9⇓⇓⇓⇓–14), and that capacity is important with respect to several perspectives. The first step of Etoh oxidation generates acetaldehyde (AA), which is toxic, reactive, and potentially carcinogenic. AA is aversive systemically, as has been observed from the unpleasant effects of disulfuram, an inhibitor of AA dehydrogenase, but it has been shown to be rewarding in parts of the brain (12, 15, 16) especially in the posterior ventral tegmental area (17⇓–19) by activating dopamine neurons (20⇓–22). The liver maintains circulating levels of AA at levels of 20–155 µM (23, 24), and AA has been reported not to penetrate the blood–brain barrier or to penetrate very slowly (23, 25). Thus, if the brain can oxidize Etoh, then intracerebral AA may mediate behavioral, neurochemical, and toxic effects of Etoh in the brain, possibly playing a role in the development of alcohol dependence (6, 16, 26).
AA is difficult to measure in living systems. The concentrations of AA in liver, blood, and brain that occur with drinking lie in the micromolar range (23, 24), in contrast to millimolar levels of Etoh and Ac, and AA rapidly disappears after sample collection due to its volatility. EtOH oxidation and the formation of AA have been measured in brain homogenates (11, 27⇓⇓–30) and cell cultures (31). In a novel approach, Zimatkin and Buben made the measurement in living brain by intraventricular perfusion (9) of anesthetized rats with 180 mM Etoh and measured AA concentrations rising to ∼50 µM in the perfusate (23). When the authors perfused the brain with 85–90 mM Etoh, they again observed AA rise in the perfusate with no measurable AA in the blood (9). These studies provide powerful evidence that Etoh oxidation and AA production occur in the living brain. However, cerebral generation of AA is believed to be extremely slow, and Etoh oxidation has not been quantified directly in the awake brain. Neither have the consequences of chronic alcohol exposure on intracerebral oxidation been reported.
It has proven difficult to discriminate by isotopic means the intracerebral oxidation of Etoh from cerebral consumption of Etoh-derived Ac produced in the liver. When hamsters or rats are infused with 14C/13C–Etoh (32), the liver generates 14C/13C–Ac, whose consumption by the brain yields isotopic labeling that is indistinguishable from that of the Etoh. In this study, a combination of [1, 2-13C2]acetate (Ac12) and [2-13C]Etoh (Etoh2) provided positional information that differentiated the metabolism of Ac from that of Etoh (Fig. 1). This strategy was used in Etoh-naïve rats and rats exposed to vapor to quantify intracerebral Etoh oxidation and assess the impact of chronic Etoh exposure on intracerebral oxidation of Etoh.
Fig. 1.
Model for cerebral metabolism of [1, 2-13C2]Ac and [2-13C]Etoh showing incorporation of 13C into the astroglial and neuronal TCA cycles. The 13C in [2-13C]Etoh is represented by a filled black circle, and the 13C in [2-13C]Ac generated by the liver from [2-13C]Etoh is represented by a filled gray circle. The 13C in [1,2-13C]Ac is represented by two circles marked with an X. The fate of those carbons can be tracked by their colors in their products. Ac inside the astroglia is a mixture of [1,2-13C]Ac transported from the blood (two circles with Xs) and [2-13C]Ac that came from the blood and from intracerebral oxidation of [2-13C]Etoh (half gray/half black). Because neurons import negligible quantities of Ac, the only Ac inside the neuron is from [2-13C]Etoh (filled black circle). The 13C enters the TCA cycle in the neuron, where it arrives in the large neuronal pool of Glu. The Glu is released and taken up by astroglia, where it mixes with the small astroglial pool and is converted to Gln, which flows to the neuron. α-KG, α-ketoglutarate; AA, acetaldehyde; Ac, acetate; Etoh, ethanol; Gln, glutamine; Glu, glutamate; OAA, oxaloacetate; Succ, succinate.
Results
Plasma.
Throughout the 2-h infusion, Ac and Etoh were the only substrates whose labeling was detected above natural abundance in the plasma; no labeling appeared for glucose, lactate, and β-hydroxybutyrate. By 30 min after the start of the infusion, blood Etoh2 levels rose to 25.5 ± 0.9 mM and 23.2 ± 1.0 mM (117 and 107 mg/dL) and rose slightly by 2 hr to 28.5 ± 0.9 and 28.1 ± 1.5 mM (131 and 129 mg/dL), respectively, with no differences between the groups (Fig. 2A).
Fig. 2.
The concentrations (mM) of ethanol (A) and acetate (B), the total 13C enrichment (%) of acetate (C), and the ratio between [2-13C]acetate (Ac2) and [1, 2-13C2]acetate (Ac12) in the blood (D), for naïve (10 rats) and ethanol treated (nine rats) groups.
The concentration of Ac increased within 5 min of the start of the coinfusion of Ac12 and Etoh2, with the treated animals having more Ac (4.75 ± 0.17 mM) than the naïve group (4.09 ± 0.24 mM) (P = 0.04). The treated group kept a constant level, whereas Ac in the naïve rats decreased to 3.51 ± 0.20 mM at 15 min and stayed constant thereafter (3.66 ± 0.18 mM) (Fig. 2B). The difference in plasma Ac may be associated with metabolic tolerance, an ability of exposed rats to metabolize ethanol more quickly (33, 34), if the rate of acetate clearance is elevated in exposed rats, although this study was not designed to address that question.
Fig. 3A shows the 13C spectra from the plasma, with the Ac12 doublet around the Ac2 singlet. The total 13C enrichment of Ac (Ac2 + Ac12) increased to ∼85% after 5 min for the Etoh-treated and naïve groups and stayed constant during the infusion (Fig. 2C). To use the steady-state Eq. 1 from Materials and Methods, the concentration of Ac and the ratio Ac2/Ac12 had to be maintained constant for the final hour of the procedure, which was achieved here (Fig. 2D). The combination of rapid metabolism of Etoh and the Ac arriving via infusion led to the appearance of 13C-Ac by the first measured plasma time point, 5 min (Fig. 2C). Then the ratio Ac2/Ac12 rose to a stable value after 30 min of infusion (Fig. 2D).
Fig. 3.
13C-NMR spectra of (A) plasma and (B) brain extracts, showing the disproportionately large singlet/doublet ratios of brain glutamate and glutamine relative to plasma acetate. Ac, acetate; Ac1,2, double resonance of [1,2-13C]Ac; Ac2s, singlet resonance of [2-13C]Ac; Ac2Total, sum of all resonances of Ac labeled at carbon 2; Etoh, ethanol; Gln, glutamine, with analogous notations; Glu, glutamate; Glu3,4, doublet resonance of [3,4-13C]Glu; Glu4s, singlet resonance of [4-13C]Glu; Glu4,5, doublet resonance of [4,5-13C]Glu; Glu4Total, sum of all resonances of Glu labeled at carbon 4.
Total Enrichments of Brain Glu4 and Gln4.
The 13C enrichments of Glu4 and Gln4 for cortex and subcortex obtained after 2 hr of infusion are shown in Fig. 4 A and B. Total enrichments of Gln4 in the Etoh-treated group was significantly higher than the naïve group in both regions (P = 0.03 and 0.02 for cortex and subcortex, respectively), demonstrating that after chronic Etoh exposure, the oxidation of Ac and/or Etoh was increased. The total enrichment of Glu4 in the naïve and treated groups did not differ significantly (P = 0.4 and 0.5 for cortex and subcortex, respectively).
Fig. 4.
The 13C labeling in Glu and Gln in the cortex and subcortex for naïve (10 rats) and ethanol-treated (nine rats) groups. (A) The percentage of carbon in cortical Glu and Gln labeled in different patterns: total C4 labeling (Glu and Gln4Total), C4 only (Glu4s, Gln4s), C3,4 (Glu34, Gln34), and C4,5 (Glu45, Gln45). (B) Analogous to A, but for subcortex. (C) The percentage of carbon in Glu and Gln that came from Etoh and Ac. Glu-Ac2, Glu 13C content from [2-13C]Ac; Glu-Ac12, Glu 13C content from [1,2-13C]Ac; Glu-Eth, Glu 13C content from Etoh. The abbreviations for Gln follow the same pattern. (D) Analogous to C but for subcortex. *P < 0.05, **P < 0.01.
Isotopomer Enrichment Patterns of Brain Glu4 and Gln4.
As a result of spin–spin splitting between the C3 and C4 carbons and the C4 and C5 carbons, it was feasible to differentiate metabolism of Ac12, which labels Gln and Glu always at both C4 and C5, from that of Ac2, which never labels Gln and Glu at C5. Resonances for Gln45 and Glu45 were distinguished from those of Gln4, Glu4, Gln34, and Glu34 (Fig. 3B). The concentrations of Gln345 and Glu345 were too low to be detected, effectively zero by comparison with the other isotopomers, so only the C4, C34, and C45 labeling were considered for the analysis. The ratio of the Glu4 singlet resonance (Glu4s) plus the Glu34 doublet relative to the Glu45 doublet is visibly greater than the ratio of the Ac2 singlet (Ac2s) to the Ac12 doublet, indicating the intracerebral oxidation of Etoh. Gln also has a singlet and Gln34 doublet resonances that are disproportionately large compared with the Gln45 doublet, also consistent with Etoh oxidation.
Fig. 4 A and B shows that Gln4s and Gln34 were significantly more labeled after chronic Etoh treatment for cortex (P = 0.02 and 0.001, respectively) and subcortex (P = 0.04 and 0.04, respectively), and the isotopomer analysis allows the source to be ascribed to Ac2 or Etoh2, as follows.
Quantification of the Intracerebral Etoh Oxidation.
Fig. 4 C and D shows the 13C enrichments from Ac12, Ac2, and Etoh2 in the cortex and subcortex regions, based on the isotopomers observed for Glu4 and Gln4. GlnEtoh2 was significantly greater (P = 0.003) after Etoh chronic treated in cortex, and although GlnAc2 was also higher, the treatment effect did not reach significance (P = 0.8). The results indicate significant intracerebral oxidation of Etoh and that the oxidation is increased by chronic exposure to Etoh.
The observed enrichments were used in Eqs. 5 and 6 together with values for rates of the cortical astroglial and neuronal tricarboxylic acid (TCA) cycles and Glu–Gln cycling in the cortex of awake rats (35) (Fig. 5). In astroglia of Etoh-naïve rats, Etoh was oxidized at a rate of 0.017 ± 0.002 µmol/min/g and in neurons 0.014 ± 0.003 µmol/min/g, corresponding to 12.3 ± 1.4% of astroglial oxidation and 1.7 ± 0.4% of neuronal oxidation, respectively (Table 1). The cortical Etoh oxidation rate for neurons and astroglia combined, 0.031 ± 0.003 µmol/min/g, corresponds to 3.2 ± 0.3% of the global brain oxidation of all substrates. Chronic alcohol exposure increased the astroglial Etoh oxidation to 0.028 ± 0.002 µmol/min/g (P = 0.001), corresponding to 20.2 ± 1.5% of astroglial oxidation, with an insignificant effect on neuronal Etoh oxidation (P = 0.1). TCA cycle and Glu–Gln cycling rates from three other publications (36⇓–38) were also used with Eqs. 5 and 6 to estimate the rates of cortical Etoh oxidation; in all cases astroglial oxidation of Etoh was found to be elevated significantly (Table S1). In the subcortex, 13C labeling from Etoh oxidation was detected, with no significant impact of Etoh exposure (Fig. 4D). Due to the heterogeneity of the subcortex and its metabolic rates (39), the absolute rates of Etoh oxidation were not estimated. However, the percentages of 13C labeling in the subcortical brain are similar to those in the cortex (Fig. 4 C and D), which suggests that the average Etoh oxidation over the subcortical brain comprises a similar fraction of total oxidation.
Fig. 5.
The cortical oxidation rates of ethanol in astroglia, neurons, and whole brain, calculated using published rates of VtcaN, VtcaA, and Vcycle, for naïve (10 rats) and ethanol-treated (nine rats) groups. **P < 0.01.
View this table:
Table 1.
The contribution of ethanol to oxidation in neurons, astroglia, and their sum relative to the TCA cycle in awake rats (35)
Discussion
Metabolic Fate of Etoh in the Brain.
The first step of Etoh oxidation is its conversion to AA by mitochondrial alcohol dehydrogenase, catalase, or cytochrome P450. Alcohol dehydrogenase has been reported to have very low activity in the brain but appears to be heterogeneously distributed, occurring in some types of neurons (40, 41). Cytochrome P450 also has been shown to metabolize Etoh in the brain in neurons and astroglia, depending on the brain regions (42), but at a rate of 0.00051 µmol/min/g (43), significantly more slowly than the rate of 0.012 µmol/min/g reported for catalase in brain homogenates (calculated from the 3.5–7.1 nmol/mg protein/h (11), using 0.1 g protein/g brain tissue (44)). Furthermore, inhibition of brain catalase activity in mice inhibits many of the pharmacologic effects of ethanol (45), and inhibition or stimulation of catalase activity in brain homogenates, respectively, reduces or increases the formation of AA (14, 15). Thus, evidence suggests that catalase is likely to be a more major contributor to Etoh oxidation in the brain. Inhibition of brain catalase activity in mice inhibits many of the pharmacologic effects of Etoh (45), and inhibition or stimulation of catalase activity, respectively, reduces or increases the formation of AA in brain homogenates (15). Although catalase is found in neurons and astroglia (46, 47), it occurs to a greater extent in neurons (48). From those previous reports of activity in vitro, one can predict a total rate of AA production of ∼0.01 µmol/min/g from alcohol dehydrogenase, catalase, and cytochrome P450 combined, much slower than indicated by the present data in vivo. A possible explanation is the presence of potential cofactors and other environmental conditions that lead to greater activity in vivo than is measured in other preparations. For example, in the presence of glucose oxidase, the activity of catalase for Etoh significantly rises to ∼10-fold (15). In this context, the faster oxidation rates observed in the living rat brain (0.031 and 0.036 µmol/min/g for naïve and treated groups, respectively) are not unreasonable.
The second stage of Etoh metabolism is the conversion of AA to Ac via AA dehydrogenase, which is located in astroglia and neurons in many brain regions (41, 49). The ability of the brain to metabolize aldehydes has long been known (22). It has been shown that the aldehyde-oxidizing capacity of the brain fraction of capillaries and astroglia cells is 5–6 times higher than that of the neurons (49, 50), providing ample capacity to carry out the second step of Etoh oxidation. The rates of oxidation in neurons and glia result from the balance of the latter’s higher catalase activity and the slower neuronal aldehyde conversion to Ac.
Ac is considered to be an astroglia-specific substrate, due to a specific Ac-uptake mechanism in astrocytes (51). However, the activity of acetyl-CoA synthetase is greater in neurons than in astrocytes (51). Therefore, a lack of Ac transport into neurons prevents them from oxidizing Ac from the blood, but one would predict that neurons can oxidize Ac that is derived from Etoh intraneuronally.
14C- and 11C- 13C–labeled Etoh have been used to study the fate of Etoh in the brain (10, 52⇓–54). One difficulty has been that isotopically labeled Etoh quickly becomes labeled Ac, whose intracerebral oxidation could not be distinguished from that of Etoh. Xiang and Shen (52) avoided this limitation by using saturation transfer. They detected no exchange between Etoh and AA in the brain, with a detection threshold of 0.001 s−1 for the rate constant from Etoh to AA. Given our Etoh levels of ∼28 mM, that saturation transfer detection threshold corresponds to a rate of 1.7 µmol/min/g, which is far greater than our finding of 0.037–0.087 μmol/min/g.
Intracerebral Etoh Oxidation After Chronic Etoh Exposure.
It has been shown that the brain consumes Ac and, when Etoh is given to rodents (4⇓–6) and humans (3, 7, 8, 55), decreases its consumption of glucose and increases its use of Ac. Three weeks of Etoh vapor exposure were associated with increased intracerebral Etoh oxidation, at a blood level of ∼28 mM (130 mg/dL). Pharmacokinetic modeling suggested that the change occurred primarily in astroglial Etoh oxidation; the percentage of oxidation supplied by Etoh was higher than in neurons and increased after Etoh exposure.
At least three mechanisms may explain how ethanol oxidation could be increased. (i) Chronic Etoh treatment reduces Mg2+ in the brain (56, 57). Exposure of brain cells to 10–25 mM EtOH in vitro reduces intracellular Mg2+ 15–45% within 10 min in astrocytes and CA1 pyramidal neurons (58). Mg2+ deficiency has been associated with increased catalase activity and H2O2 production in vitro (59), which might be a potential mechanism for the increase of the oxidation of Etoh in astroglia. (ii) Although alcohol dehydrogenase normally has negligible activity in the brain (11), its activity has been reported to increase after chronic Etoh treatment (60). (iii) Cytochrome P450 has also been reported to be induced by a moderate dose of 1 g of ethanol in rats weighing 250–300 g (43), with increases of 150–500% in rats treated with 10% ethanol in drinking water for 30 d (61).
Significance of Intracerebral Etoh Oxidation.
The current study shows that Etoh can provide energy for the brain, not only from Ac, which was generated in the liver, but also from the directly oxidation in the brain. The small but significant contribution of Etoh to brain oxidation might provide some energetic reward. When Etoh is oxidized directly by the brain, AA is produced within the brain, where it may serve as a pharmacologic reward (62), despite the fact that AA is aversive in the rest of the body. Another likely impact of intracerebral metabolism of Etoh is cumulative damage due to AA toxicity and oxidative damage from free radical generation (63).
Limitations.
This study shows that intracerebral oxidation of Etoh occurs and that its rate is increased in astroglia by chronic Etoh treatment. However, the sensitivity of the technique was insufficient to measure oxidation in particular regions of interest, such as the posterior ventral tegmental area (19, 62). A further limitation pertains to the kinetic modeling. The measurements were conducted at an isotopic steady state, which means that the data contain only relative rates: that is, rates of ethanol oxidation relative to the rates of other pathways. Therefore, the absolute rates of ethanol oxidation in glia and neurons, in μmol/min/g, were estimated assuming values for the ratios Vcycle/VtcaN and Vcycle/VtcaA. A factor to note is that one purpose of this study was to test whether chronic exposure to Etoh could affect intracerebral metabolism of Etoh; it did, but whether such a mechanism is associated with eventual dependence or not is an important question that remains to be investigated.
Conclusions
The liver oxidizes most Etoh in the body (1, 2), but there remains some question of how much, if any, Etoh the brain can oxidize directly (9⇓⇓⇓–13). However, the quantification of direct intracerebral Etoh oxidation has been encumbered by hepatic conversion of isotopically labeled Etoh to abundant Ac that circulates in the blood and is readily consumed by the brain. In this experiment, the combination of i.v. infusion of both Ac12 and Etoh2 allowed the discrimination of the oxidation of Ac from that of Etoh. The results showed that Etoh can be oxidized by the brain to provide energy, supplying ∼12% of astroglial oxidation and ∼2% of neuronal oxidation, and that 3 wk of exposure to Etoh vapor was associated with an increase in astroglial Etoh consumption from 12% to 20% of total astroglial oxidation.
Materials and Methods
Strategy for Isotopic Labeling and Estimation of Etoh Oxidation Rates.
After the 13C-labeled Ac crosses the blood–brain barrier, it is metabolized exclusively by astroglia (Fig. 1) (51) and labels Glu and Gln (36⇓–38, 51, 64). When Ac12 is infused, it enters the brain and yields [4, 5-13C2]Glu and Gln in the first turn of the TCA cycle and [3, 4, 5-13C3]Glu and Gln in the second and subsequent turns (Fig. 1). The total C45 and C345 labeling is proportional to the percent enrichment (PE) of Ac12 in the blood, with a proportionality constant kAc = ([4, 5-13C2]Glu + [3, 4, 5-13C3]Glu)/Ac12. When [2-13C]Etoh is infused simultaneously, the liver quickly converts some of it to [2-13C]acetate (Ac2). In the brain, Ac2 generates [4-13C] and [3, 4-13C2]Glu and Gln, following the same proportion kAc observed from the C45+C345-glutamate labeled by Ac12. The present study was designed with the intention that Etoh2 would be the only singly labeled substrate in the blood besides Ac2, so any C4 and C45 labeling of glutamate beyond that expected from the proportion kAc must come from Etoh2. The 13C-labeling of Gln from Etoh2 was according to:
where PE is the 13C enrichment of Gln at the numbered positions. An equivalent expression exists for Glu.
Etoh oxidation rate was evaluated using steady-state isotopic balance equations:
N and A denote neurons and astroglia; Vcycle, VtcaA, and VtcaN are the rates of the glutamate–glutamine neurotransmitter cycle and the astroglial and neuronal TCA cycles, respectively; and CMRetN and CMRetA are the cerebral metabolic rates of Etoh in neurons and astroglia. fEtoh2 and fAc2 represent the fractional isotopic enrichments of Etoh2 and Ac2, after subtraction of the 1.1% natural abundance enrichment.
Defining 
yields the following expressions for the rates of neuronal and astroglial Etoh oxidation:
where 
and 
. The values of CMRetN and CMRetA can be obtained using published rates for Vcycle, VtcaN, and VtcaA, and the total oxidation rate of Etoh (CMRettot) is the sum of CMRetN and CMRetA.
Animal Procedures.
All experiments were carried out in accordance with protocols approved by the Yale Animal Care and Use Committee. Nineteen adult male Sprague–Dawley rats (Charles River Labs, Inc.) were maintained on a 12 h/12 h light–dark cycle, and food (Teklad Global 18% Protein Rodent Diet 2018, Harlan Laboratories) and water were available ad libitum. They were allowed a minimum of 2 d to acclimate before the start of the experiments. To familiarize the rats with human interaction and minimize stress on the day of the infusion, they were handled daily until the final day. Room air or Etoh was administered to rats in vapor chambers constructed in-house (65), applying the Etoh vapor 8 h/d for 3 wk. At the start of the exposure period, the naïve rat (n = 10) weighed 156.9 ± 1.8g , and the treated rats (n = 9) weighed 155.5 ± 1.5 g (P = 0.572). The initial vapor level was set at 18∼20 mg/L and gradually increased to 23∼25 mg/L over the first week. This inhalation model is associated with minimal hepatic dysfunction (66). To monitor the blood Etoh concentration, 200 μL of blood was drawn from the saphenous vein of one rat in each cage each day after 7 h of the day’s exposure. With three rats housed per cage, each rat was tested once every 3 d. The blood samples were centrifuged to obtain plasma, and the Etoh concentration measured (Analox GM7, Analox Instruments, Inc.). During the exposure periods, the blood Etoh concentrations reached 150∼200 mg/dL, levels that have been shown to induce depressive- and anxiety-like behaviors in rats (67) and slurred speech, double vision, and difficulty walking in humans (68). After 3 wk of Etoh exposure, the weight of the treated rats (301.0 ± 2.8 g) was 3% higher than that of the naïve rats (310.7 ± 2.7 g) (P = 0.033).
On the day of infusion, anesthesia was induced with 3% (vol/vol) isoflurane and lowered to 1.5–2.5% for 1–2 min to catheterize one lateral tail vein with a 23-G needle connected with PE-50 tubing (Instech Laboratories, Inc.) (35). The rats then recovered for 30 min, moving and walking freely in the cage before the infusions. Etoh2 (up to 1.373 mg/kg over 2 h) was infused through the tail catheter using a pharmacologically modeled infusion protocol (69). Ac12 (up to 322 mg/kg over 2 h) was coadministered by a variable rate infusion over 2 h (2 M dissolved in water, pH = 7.0) according to the following rate schedule: 0–15 s, 1.875 mmol/min/kg; 15 s–4 min, 0.265 mmol/min/kg; 4–14.5 min, 0.15 mmol/min/kg; and 14.5–120 min, 0.075 mmol/min/kg. During the administration, 100 μL blood samples were collected from the saphenous vein at the time points 0, 5, 15, 30, 60, 90, and 120 min; centrifuged at 18,000 × g for 10 min at 2–4 °C; and frozen for later measurement of plasma Etoh2, Ac2, and Ac12. After the infusion, the rats were euthanized with focused microwave irradiation (35, 70). The fixed brain was removed and dissected into cortex and subcortex and frozen for later NMR analysis.
Preparation of Plasma Samples for NMR Study.
The frozen samples were thawed and centrifuged at 18,000 × g for 5 min. The supernatant was withdrawn and mixed with water to total volume of 150 μL, and an additional 60 μL D2O and 240 μL buffer with 12.5 mM Pi (90 mM K2HPO4 and 35 mM KH2PO4) were added, with 5 mM sodium formate as a concentration and chemical shift reference for 1H-NMR and 150 μL 10 mM [2-13C]glycine as a chemical shift reference for 13C-NMR. The mixture was transferred into a 5 mL NMR tube for NMR analysis.
Preparation of Brain Samples for NMR Study.
The frozen brain samples were weighed and placed into a 7 mL homogenizer with 300 μL 0.1 M HCl/mEtoh in a wet ice bath and ground. We added 3 mL ice-cold 60% (vol/vol) Etoh with 150 μL 10 mM [2-13C]glycine as an internal standard, and the mixture was homogenized. The homogenizer was washed with 2 mL ice-cold 60% Etoh, and the total mixture was centrifuged at 18,000 × g for 30 min at 2–4 °C. The supernatant was withdrawn, frozen with liquid nitrogen, and lyophilized for NMR analysis.
The lyophilized extracts were dissolved with 60 μL D2O and 540 μL phosphate buffer. The solution was centrifuged at 18,000 × g for 30 min, and the supernatant was collected for [1H-13C]NMR and 13C-NMR analysis.
NMR Analysis for the Brain Tissue Extracts and Plasma Samples.
The measurements were performed at 11.75T on a Bruker vertical bore NMR spectrometer operating at 500.13 MHz for 1H and 125.76 MHz for 13C. 1H-NMR was used to measure the concentrations and total 13C-enrichments of energy substrates in the plasma. 1H-Observed/13C-Edited (POCE) NMR was used to measure the brain concentrations and total 13C-enrichments of Glu and Gln C4 (71). Polarization transfer was used to observe the 13C spectrum to measure plasma Ac12 distinctly from Ac2 and to distinguish brain Glu and Gln C4 and C34 from Glu and Gln C45 and C345.
For 1H- and POCE-NMR, the magnetic field homogeneity on each sample was optimized with an automated 3D filed mapping algorithm capable of adjusting up to fifth-order spherical harmonics.
1H-NMR.
Spectra were obtained with 12 μs hard-pulse nonselective excitations, with free induction decays acquired as 16,384 complex points. Total concentrations were determined relative to the added formate, and the 13C enrichments of Ac were calculated from the ratio of the areas of the satellite peaks relative to the entire integral of the signal for Ac2.
POCE.
Spectra were obtained with the POCE (71), using an 8-ms echo time, 20-s repetition time, 12-ppm sweep width, and 32,768 complex data points. Briefly, the heteronuclear editing method consists of the acquisition of two spin-echo measurements, one with a broadband inversion pulse applied at the 13C frequency and the other without the inversion pulse. The difference between the spectra represents protons bound to 13C (at twice the true intensity), whereas the subspectrum without the inversion pulse represents the protons for 13C-labeled plus the unlabeled compounds (i.e., the total concentrations). The total 13C enrichments of Glu4 and Gln4 were half of the respective ratios between the Glu4 and Gln4 signals in the difference spectrum and subspectrum without the inversion pulse.
13C-NMR.
The pulse sequence Insensitive Nuclei Enhanced by Polarization Transfer (INEPT) (72⇓–74) was involved, and 32,768 complex points were acquired after manual adjustment of the magnetic field homogeneity. The enrichments of C4, C34, C45, and C345 of Glu and Gln were calculated according to:
where AGln4 is the area of the Gln C4 singlet resonance, AGln34 is the area of the doublet resonance that represents [3, 4-13C2]Gln, and so on for the relevant combinations of labeling. The analogous calculations were performed for fGlu4+34 and fGlu45+345.
Statistical Analysis.
The significance of differences between means of measured parameters in different treatment groups was assessed using two-tailed t tests, assuming unequal variances. All means are presented ± SEM.
Acknowledgments
The authors thank Professor Sean O’Connor of Indiana University–Purdue University Indianapolis for providing the pharmacokinetic model and guidance in its use to derive the Etoh infusion protocol, and Golam Chowdhury for guidance on the Ac infusion protocol. The research was supported by National Institutes of Health (NIH) Grants R21 AA018210 (to G.F.M.), R21 AA019803 (to G.F.M.), and P30 NS052519, and National of Natural Science Foundation of China 21105116 (to J.W.).
Footnotes
- ↵1To whom correspondence should be addressed. E-mail: graeme.mason{at}yale.edu.
Author contributions: J.W. and G.F.M. designed research; J.W., H.D., and X.M. performed research; J.W., R.A.d.G., K.L.B., and G.F.M. contributed analytic tools; J.W., H.D., L.J., and G.F.M. analyzed data; and J.W., L.J., and G.F.M. wrote the paper.
The authors declare no conflict of interest.
↵*This Direct Submission article had a prearranged editor.
This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1306011110/-/DCSupplemental.
References
Oxidation of ethanol by rat brain
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