ABSTRACT Huntington’s disease has been associated with a failure in energy metabolism and oxidative damage. Ascorbic acid is a powerful antioxidant highly concentrated in the brain where it

acts as a messenger, modulating neuronal metabolism. Using an electrophysiological approach in R6/2 HD slices, we observe an abnormal ascorbic acid flux from astrocytes to neurons, which is

responsible for alterations in neuronal metabolic substrate preferences. Here using striatal neurons derived from knock-in mice expressing mutant huntingtin (STHdhQ cells), we study ascorbic

acid transport. When extracellular ascorbic acid concentration increases, as occurs during synaptic activity, ascorbic acid transporter 2 (SVCT2) translocates to the plasma membrane,

GLUTATHIONE–ASCORBATE METABOLISM INDUCES TRANSIENT DOPAMINE INCREASE AND MOTOR DYSFUNCTION Article Open access 28 October 2024 PS421 HUNTINGTIN MODULATES MITOCHONDRIAL PHENOTYPES AND

CONFERS NEUROPROTECTION IN AN HD HIPSC MODEL Article Open access 25 September 2020 PYROPTOTIC CELL DEATH IN THE R6/2 MOUSE MODEL OF HUNTINGTON’S DISEASE: NEW INSIGHT ON THE INFLAMMASOME

Article Open access 31 July 2020 INTRODUCTION Huntington’s disease (HD) is a progressive, autosomal dominant, neurodegenerative disorder1. The disease is caused by an expanded polyglutamine

(polyQ) stretch in the IT15 gene, resulting in major cell loss in the striatum2. The _IT15_ gene codes for a protein called huntingtin (Htt). Wild-type (WT) Htt has an important role in the

intracellular transport of vesicles, organelles and trafficking of proteins to the cell surface3,4. Expansion of a glutamine stretch within the Htt protein to >40 repeats (mHtt) appears

to confer a dominant toxic property that is deleterious to neurons and detrimental to normal Htt biological activities. Defects in energy metabolism have been observed in presymptomatic and

symptomatic subjects5,6,7,8,9. This deregulation in brain energy metabolism makes it reasonable to suspect the presence of increased amounts of oxidative species. Indeed, oxidative damage is

evident, as well as impaired superoxide dismutase (SOD) activity8 and a decrease in ascorbic acid levels10,11. Ascorbic acid is an important antioxidant that is concentrated in the brain12.

It plays a fundamental role in the modulation of neuronal metabolism (for reviews, refer to the studies by Castro _et al._13 and Beltrán _et al._14). During synaptic activity, ascorbic acid

is released into the extracellular space15,16,17,18 and is taken up by neuronal cells19. This uptake of ascorbic acid occurs via a specific neuronal ascorbic acid transporter, the

sodium-dependent vitamin C transporter, SVCT220. Intracellular ascorbic acid is able to sustain an adequate energy supply that induces uptake of monocarboxylates such as pyruvate and

lactate21. It is also used to maintain redox balance in the cell12. R6/2 is a transgenic mouse model that expresses exon 1 protein of human Htt, which contains ~150 CAG repeats. These mice

develop a neurological phenotype that mimics the disease characteristics including many of the motor-control deficits that are characteristic of HD22,23 and show a pronounced and sustained

loss of ascorbic acid in the extracellular fluid. This only occurs during periods of behavioural activation. Rather than demonstrating a deficit in brain ascorbic acid, these mice seem

unable to maintain adequate levels of ascorbic acid during behavioural activation. Thus, a failure in ascorbic acid uptake may be closely linked with metabolic failure and redox imbalance in

HD. In this study, we demonstrate that a failure in ascorbic acid flux from astrocytes to neuronal cells occurs and that this phenomenon precedes the onset of HD-like symptoms in mice. The

failure is accompanied by an alteration in neuronal metabolic energy substrate preferences. We also show a failure in neuronal ascorbic acid uptake in an HD mouse and cellular models. In

cells that express mutant Htt (mHtt), Huntingtin-associated protein 1-SVCT2 (HAP1-SVCT2) colocalization is decreased and the neuronal ascorbic acid transporter, SVCT2, fails to reach the

plasma membrane. Ascorbic acid is an essential protector against oxidative damage in brain neurons and modulates neuronal metabolism during synaptic activity. Therefore, early failure in

ascorbic acid release could lead to metabolic failure and early impairment in the maintenance of synaptic transmission in basal ganglia. Such a failure in ascorbic acid uptake in symptomatic

HD subjects could promote neuronal death. RESULTS GLIAL-NEURON ASCORBIC ACID FLUX IS IMPAIRED IN R6/2 MICE Ascorbic acid flux from astrocytes to neurons during synaptic activity is

essential to protect neurons against oxidative damage and for modulation of neuronal metabolism, thus permitting optimal ATP production. According to our previous data, ascorbic acid

inhibits glucose consumption and stimulates lactate uptake during glutamatergic synaptic activity (ascorbic acid metabolic switch19,21). We were interested in determining whether the

ascorbic acid metabolic switch functions in R6/2 transgenic mice that carry HD gene, as HD mice have been shown to be unable to maintain adequate levels of ascorbic acid during behavioural

activation22,24 and HD has also been associated with a failure in brain energy metabolism. In our studies of mice striatal slices, we have shown that intracellular ascorbic acid inhibits the

ability of glucose to sustain excitatory postsynaptic currents (EPSCs) when neuronal lactate uptake is inhibited (Fig. 1, Table 1). EPSCs were evoked by stimulating the corticostriatal

pathway (Fig. 1a). After glucose deprivation, glucose application produced a fast recovery of synaptic responses (Fig. 1b), as much in WT mice as in both presymptomatic and symptomatic R6/2

mice (Fig. 1c, control). In the presence of external 100 μM alpha-cyano-4-hydroxycinnamic acid (4-CIN), an inhibitor of the neuronal monocarboxylate transporter MCT2, glucose was able to

restore EPSCs only in presymptomatic (80.0±14.1%) and symptomatic (72.3±16.4%) R6/2 mice (Fig. 1c, 4-CIN). Thus, in WT mice, glucose is not able to sustain neural function when

monocarboxylate uptake is inhibited (27.8±12.0% and 44.8±10.6% for EPSC amplitudes after a period of glucose deprivation in presymptomatic and symptomatic WT slices, respectively). In

presymptomatic and symptomatic R6/2 mice, 4-CIN did not affect the ability of glucose to sustain EPSCs. Given these findings, our experiments strongly suggest that the ANLS (astrocyte-neuron

lactate shuttle) is necessary to maintain adequate ATP levels in healthy cells, a normal mechanism that is impaired in HD neurons. When extracellular ascorbic acid was applied, we observed

a strong decrease in the amplitude of the EPSCs after a period of glucose deprivation in presymptomatic R6/2 slices (Fig. 1c, 4-CIN/Asc). In symptomatic R6/2 slices, we saw no significant

decrease in the amplitude of EPSCs after glucose deprivation (Fig. 1c, 4-CIN/Asc). So, ANLS would seem to function in presymptomatic R6/2 slices by over-supplementation of extracellular

ascorbic acid. This effect was reversed when ascorbic acid uptake was inhibited (by including an anti-SVCT2 antibody in the recording pipette; Fig. 1c, 4-CIN/Asc/anti-SVCT2). This antibody

inhibits ascorbic acid uptake by binding to the endofacial side of SVCT219. In conclusion, the ANLS and the ascorbic acid metabolic switch do not function in the striatum of presymptomatic

R6/2 mice due to the fact that ascorbic acid release from glial cells seems to be decreased in these mice. The ANLS and ascorbic acid metabolic switch is also impaired in symptomatic R6/2

mice, though in these mice, this dysfunction appears to arise from a failure in ascorbic acid uptake rather than from a decreased release of ascorbic acid from glial cells. To investigate

whether the failure in ascorbic acid uptake may be related to a deficiency in SVCT2 expression, we analysed mRNA and protein levels in samples obtained from mice striata. RT–qPCR analyses

showed increased levels of mRNA coding for SVCT2 in symptomatic R6/2 mice, while mRNA levels were not affected in presymptomatic R6/2 mice (Fig. 1d,e). Relative levels of SVCT2 were also

increased in symptomatic R6/2 mice as seen by immunofluorescence analysis (Fig. 1f). In presymptomatic R6/2 mice, SVCT2 expression did not seem to be altered (Fig. 1d–f). Thus, before the

onset of HD-like symptoms, SVCT2 expression and function should be normal. After the onset of HD-like symptoms, ascorbic acid uptake seems to be impaired, although SVCT2 expression is

increased. ASCORBIC ACID TRANSPORT IN CELLULAR MODEL OF HD IS IMPAIRED To visualize the possibility that neuronal ascorbic acid transport is impaired in a cellular model of HD, we studied

SVCT2 expression and function in STHdhQ7 and STHdhQ111 cell lines. Obtained from huntingtin (Htt) mouse knock-in embryos, STHdhQ is a cell line of immortalized striatal neurons that

accurately express endogenous levels of normal and mutant huntingtin (Htt), and are so dubbed HD cells. WT STHdhQ7 cells express Htt containing 7 glutamine repeats, while mutant STHdhQ111

cells express Htt containing 109 glutamine repeats25. To simplify, we can refer to WT and mutant cells as Q7 and Q111 cells respectively. As for symptomatic R6/2 mice, qPCR analyses showed

an increased mRNA coding for SVCT2 in Q111 cells (Fig. 2a). We apparently found increased SVCT2 levels with western blot analysis (Fig. 2b,c, Supplementary Figs S1 and S2); however, this was

not significant. Immunofluorescence analyses showed strong immunoreactive spots throughout the cell body (Fig. 2d). Immunoreactivity seemed to be more abundant in Q111 than Q7 cells. Both

Q7 and Q111 cells showed ascorbic acid uptake. To demonstrate that ascorbic acid transport was specific, we investigated the sodium dependence of uptake. Ascorbic acid transport was measured

using 1 min uptake assays in the presence of the Na+,K+-ATPase inhibitor, ouabain and the non-related inhibitor cytochalasin B (Cyto B, Fig. 2e). Incubation with ouabain decreased ascorbic

acid uptake by 65.4 and 69.1% in Q7 and Q111 cells, respectively. Additional assays were performed to determine kinetic parameters (Fig. 2f,g). Q7 cells showed an apparent Km of 118.6 μM, a

value comparable to the _K_m described for SVCT220,26,27. Surprisingly, the apparent _K_m for Q111 cells (55.5 μM) was lower than that for Q7 cells, implying an increased affinity for

ascorbic acid in HD cells. The higher affinity for ascorbic acid in Q111 cells was negatively compensated with a low _V_max (_V_max=269.1 pmoles per 106 cells × min for Q7 cells and 110.5 

pmoles/106cell × min for Q111 cells). A decrease in _V_max could be interpreted as a decrease in the number of transporters at the plasma membrane or as a decrease in the efficiency of the

transporter (kcat). With biotinylation assays, no changes were seen in SVCT2 levels at the plasma membrane on comparing Q7 and Q111 cells (Fig. 4e,f). Therefore, SVCT2 expression is

increased in Q111 cells, but the number of transporters at the plasma membrane remains the same as for Q7 cells, implying a decrease in SVCT2 efficiency. MUTANT HUNTINGTIN IMPAIRS SVCT2

TRANSLOCATION Synaptic activity is expensive in energetic terms. The continual production of energy and the metabolism of neurotransmitters require antioxidants to maintain redox balance. As

synaptic activity is known to induce an increase in extracellular ascorbic acid concentration, we measured 14C-ascorbic acid uptake in Q cells preincubated with non-radioactive ascorbic

acid (Fig. 3). 14C-ascorbic acid uptake in preincubated Q7 cells increased by ~200% (Fig. 3a). This effect was not significant for Q111 cells, with an increased uptake of ~150% (Fig. 3a). We

also studied 14C-ascorbic acid uptake stimulation induced by preincubation with non-radioactive ascorbic acid in the non-neuronal cell line, HEK293, which expresses endogenous SVCT2 (ref.

21). The effect of preincubating with ascorbic acid was dose- and time-dependent and was reversible (Fig. 3b). We reasoned that an increase in 14C-ascorbic acid uptake is possible due to an

increased number of transporters at the plasma membrane or an increase in uptake efficiency. We tested the first hypothesis using total internal reflection fluorescence (TIRF) microscopy in

both Q cells and cultured striatal neurons expressing SVCT2-EGFP. We observed a significant increase in fluorescence corresponding to SVCT2-EGFP at the plasma membrane after preincubation

with ascorbic acid in Q7 cells (Fig. 4b,d) and in cultured striatal neurons (Fig. 4a,c). However, no changes were observed in fluorescence intensity for Q111 cells (Fig. 4b,d) and cultured

striatal neurons that expressed mHtt-DsRED (Fig. 4a,c). TIRF results were supported by biotinylation (Fig. 4e,f, Supplementary Figs S3 and S4) and immunofluorescence assays (Fig. 5a). It

would seem therefore that SVCT2 is translocated to the plasma membrane in response to an increase in extracellular ascorbic acid concentration. SVCT2-plasma membrane colocalization is weakly

observed in cells expressing mtHtt, implying that SVCT2 translocation is Htt-dependent. Under resting conditions (in which the extracellular ascorbic acid concentration is 0–200 μM) SVCT2

is located mainly at intracellular compartments and shows colocalization with early endosome markers (Fig. 5). We measured the mobile fraction (Mf) of SVCT2-EGFP with fluorescence recovery

after photobleaching (FRAP) experiments in Q cells, striatal neurons and in HEK293 cells. Mf is represented by molecules that contribute to the fluorescence recovery. The FRAP recovery curve

does not always reach the original level of fluorescence intensity. The plateau level is often lower than the original fluorescence intensity because some bleached molecules at the region

of interest are immobile and they do not contribute to the fluorescence recovery. SVCT2 present in vesicles should be more mobile than SVCT2 present at the plasma membrane. Indeed, after

preincubation with ascorbic acid, the Mf for SVCT2 decreases in Q7, in cultured striatal neurons and in HEK293 cells (Fig. 6). Less significant changes in Mf were observed in Q111 cells,

while no changes in Mf were observed in cultured striatal neurons expressing mHtt (Fig. 6), which suggests that SVCT2 remains contained within vesicular intracellular compartments. The

weaker effect observed in Q111 cells could be due to differences in mHtt expression. Q111 cells are derived from knock-in embryos, while neuronal cells are overexpressing mHtt. CYCLING OF

SVCT2 IS ALTERED IN CELLS EXPRESSING MHTT The idea that transporters revolve in a cycle to and from the plasma membrane is not new. Constitutive endocytosis and recycling of the glutamate

transporter EAAC1 occurs to and from the plasma membrane28. It is thus possible that SVCT2 may be cycling to and from the plasma membrane so that when extracellular ascorbic acid

concentration increases, SVCT2 becomes ‘trapped’ at the cell surface. If our idea is true, we reason that if we are able to manipulate the rates of endocytosis and exocytosis, then we might

be able to manipulate the amount of SVCT2 at the plasma membrane. In other words, if we decrease the rate of endocytosis or increase the rate of exocytosis we should see an increase in the

amount of SVCT2 at the plasma membrane. To try and confirm this, we used TIRF microscopy in conjunction with biotinylation and kinetic assays using cytochalasin D (Cyto D) and phenylarsine

oxide (PAO). Cyto D inhibits actin polymerization and in this way is known to inhibit clathrin-independent endocytosis29 and exocytosis30. PAO is known to inhibit clathrin-dependent

endocytosis by reacting with vicinal sulfhydryls to form stable ring structures31. TIRF microscopy (TIRFM) experiments in the presence of Cyto D (1 μg ml−1) and ascorbic acid (1 mM) gave no

indication of an increased amount of SVCT2-EGFP at the plasma membrane (Fig. 7). A similar negative result was observed for endogenous SVCT2 using immunofluorescence assays and biotinylation

assays (Fig. 8b,c). Treatment with Cyto D alone revealed a decreased amount of SVCT2 at the cell surface (Fig. 8b,c). This decrease in SVCT2 was accompanied by a decrease in ascorbic acid

uptake (Fig. 8a). Preincubation with extracellular ascorbic acid did not reverse the effects of Cyto D (Fig. 8a). Cells treated with PAO (50 μM) showed an increased amount of SVCT2 at the

plasma membrane (Fig. 8b,c, Supplementary Figs S5 and S6) and increased ascorbic acid uptake (Fig. 8a). Treatment with extracellular ascorbic acid and PAO showed an increase in ascorbic acid

uptake without significant changes in the amount of SVCT2 at the plasma membrane. As SVCT2 is a sodium-dependent transporter (co-transport of ascorbate and Na+), we were curious to see

whether the ascorbic acid effect was dependent on the presence of the ascorbate anion or the sodium cation. To do this, we performed experiments in Na+-free buffers, replacing Na+ with

choline, an inert cation. TIRFM assays were suggestive of a slight decrease in SVCT2 at the plasma membrane in the presence of ascorbic acid in Na+-free buffer (Fig. 7, Na+-free). This

observation was supported by uptake analyses that showed a decrease in 14C-ascorbic acid uptake in control cells and in cells pretreated with extracellular non-radioactive ascorbic acid

(Fig. 8a, Na+-free). Therefore, our results suggest that SVCT2 is constitutively cycling between the plasma membrane and endocytic compartments (early endosomes) and that extracellular

ascorbic acid ‘traps’ SVCT2 at the plasma membrane (by a decrease in the rate of clathrin-dependent endocytosis). Ascorbic acid may also induce translocation of SVCT2 to the plasma membrane

(by an increase in the rate of exocytosis) as observed in 14C-ascorbic acid uptake experiments in the presence of PAO and extracellular non-radioactive ascorbic acid (Fig. 8a, PAO). SVCT2

translocation to the plasma membrane was altered in cells expressing mHtt, as shown by immunofluorescence imaging in Q7 and Q111 cells, with GLUT3-mCherry as a plasma membrane marker (Fig.

5). In the absence of extracellular ascorbic acid, SVCT2 is mainly located at early endosomes (Figs 5 and 9). Htt is an essential integrator of vesicular trafficking, in which several

Htt-binding partners (Htt-associated proteins) are involved3. Considering that Htt is determinant in the positioning of endosomes32, we studied SVCT2 colocalization with early endosomes,

Htt, HAP1 (Fig. 9a) and Htt-associated protein 40 (HAP40, Fig. 9b). In Q7 cells, SVCT2 colocalizes with HAP1 and early endosomes (using EEA1 as early endosome marker) into subcellular

structures that are similar to vesicles or endosomes (Fig. 9a). However, cells expressing mHtt (Q111 cells) show a decrease in SVCT2-HAP1 colocalization with no apparent change in SVCT2-EEA1

colocalization (Fig. 9a). We saw no apparent change in SVCT2-HAP40 and SVCT-Htt (Fig. 9b) colocalization when we compared Q7 with Q111 cells. HAP1 has been linked to microtubule-associated

vesicular trafficking3 and to delivery of plasma membrane proteins to the cell surface4. Our results suggest that SVCT2 is being recycled constitutively. The loss of Htt function induces a

decrease in vesicular motility and SVCT2-HAP1 mislocalization, thus disrupting SVCT2 translocation to the plasma membrane. DISCUSSION The brain is an expensive organ in energetic terms.

Monocarboxylates are essential for sustaining neuronal energy metabolism during synaptic activity33,34. HD is characterized by a failure in brain energy metabolism. Positron emission

tomography studies have demonstrated marked reductions in metabolic rates in the basal ganglia8 and in the cerebral cortex of symptomatic HD patients7,35. In the present study, we have shown

that an alteration in neuronal substrate energetic preferences occurs in presymptomatic and symptomatic brain slices from R6/2 mice. The observed alteration seems to be related to a

deficiency in ascorbic acid flux between astrocytes and neurons and to a deficiency in the ascorbic acid metabolic switch13. According to the ascorbic acid metabolic switch, astrocytes

release ascorbic acid in response to glutamatergic activity. Ascorbic acid is taken up by neurons via the SVCT2. Neuronal intracellular ascorbic acid modulates energetic substrate

preferences, favouring lactate consumption as well as contributing to the antioxidant capacity of these cells. The failure we observe in R6/2 slices may be associated with a failure to

obtain neuronal energetic substrate and thus to an inability to adequately sustain synaptic activity, which may produce a decrease in striatum output that alters the function and integration

capacity in the circuitry of basal ganglia. The observed failure was reversed by the addition of exogenous ascorbic acid in brain slices from presymptomatic mice. These findings are

consistent with observations made by Rebec _et al._10,22. HD models, R6/2 and knock-in mice, show a pronounced and sustained loss of extracellular fluid that appears only during periods of

behavioural activation. Both models show an inability to maintain adequate levels of ascorbic acid during behavioural activation. Treatment with ascorbic acid has been shown to significantly

attenuate the neurological motor signs of HD in R6/2 mice24. The upregulation of SVCT2 expression we observed in symptomatic mice may be an adaptive mechanism to compensate for decreased

extracellular ascorbic acid levels during brain activation. STHdhQ cells are a good model for the studying the cellular and molecular mechanisms of HD. With respect to ascorbic acid

transporters, Q111 cells show an increase in SVCT2 expression, as do symptomatic R6/2 mice. Neuronal SVCT2 has an apparent Km of 103 μM (ref. 20) comparable to that observed here for

ascorbic acid transport in Q7 cells. The observed great affinity for ascorbic acid transport in Q111 cells may be an adaptive response mechanism when transporter trafficking is impaired.

Extracellular ascorbic acid concentration under resting conditions has been shown to be in the order of 200–300 μM. Thus, considering the _K_m for SVCT2 ascorbic acid, transporters should be

saturated and insensitive to changes in extracellular substrate concentrations. SVCT2 translocation to the plasma membrane in response to an increase in extracellular ascorbic acid

concentration explains how ascorbic acid flux from astrocytes to neurons is successful during brain activation. According to our data, under resting conditions, SVCT2 should be located

mainly in intracellular compartments that colocalize with early endosome markers. So, SVCT2 should be continually cycling between the plasma membrane and intracellular compartments. The idea

of a transporter cycling to and from the plasma membrane and with trafficking modulated by its substrate is not a new idea. Such a mechanism has been described for the glutamate

transporter, EAAC1 (ref. 28), the dopamine transporter, DAT (ref. 36), the GABA transporter, GAT137 and the siderophore iron transporter, Sit1 (ref. 38). If SVCT2 is cycling constantly,

translocation of SVCT2 to the plasma membrane may arise as a result of changes in the rate of transporter exocytosis and endocytosis. We proposed that extracellular ascorbic acid ‘traps’

SVCT2 at the plasma membrane, such that an increase of SVCT2 at the cell surface may result from a decreased rate of endocytosis or an increased rate of exocytosis. Our experiments with PAO

strongly support this hypothesis. We found no significant change on comparing experiments with PAO and with PAO plus ascorbic acid, implying that SVCT2 is constitutively internalized in a

clathrin- and dynamin-dependent manner. A similar internalization method has been described for the glutamate transporter, EAAC1 (ref. 28). However, we should not overlook the fact that

extracellular ascorbic acid induces an intrinsic activation of the transporter (for example, altering kcat by inducing conformational changes). In Q7 cells, we observed a 200% increase in

14C-ascorbic acid uptake after preincubation with non-radioactive ascorbic acid, while Q111 cells showed a 150% increase. These data are not consistent however with the results obtained from

TIRF microscopy. This discrepancy may be attributed to the fact that radioactive assays are more sensitive than imaging methods. Nevertheless, we must not discount the possibility that

intrinsic activation may be occurring. Expression of mHtt induces a decrease in SVCT2-HAP1 colocalization. HAP1 facilitates molecular interactions with microtubule motor proteins such as

kinesin39,40. HAP1 interacts with the p150glued subunit of dynactin, associated with dynein molecular motor proteins that confer movement of vesicles through microtubules41. Alterations in

trafficking of proteins to neuronal membranes are common in HD and other neurological and psychiatric disorders4,42. Indeed, GABAA receptor delivery to synapses is Htt- and HAP-1-dependent

and occurs with the participation of the HAP-1-KIF5 complex4. Furthermore, regarding the association between Htt and actin filaments, HAP40, an effector of Rab5, has been reported to form a

complex with huntingtin, thus favouring an association between endosomes and actin filaments43. We saw no apparent change in SVCT2-HAP40 colocalization in Q7 compared with Q111 cells. In

fact, only slight colocalization was seen in all cases. However, the possibility that a SVCT2–HAP40 interaction may exist should not be discounted. We conclude that neuronal ascorbic acid

uptake is impaired in a mouse model of HD. When the glutamatergic corticostriatal pathway is activated, astrocytes remove glutamate from the synaptic cleft (Fig. 10a). Glutamate uptake in

astrocytes stimulates lactate and ascorbic acid release from these cells. The increase in extracellular ascorbic acid induces SVCT2 translocation to the plasma membrane by inhibiting

endocytosis. Thus, neuronal cells can efficiently uptake ascorbic acid through SVCT2. Neuronal intracellular ascorbic acid stimulates lactate uptake favouring optimal energy production and

maintaining redox balance. Synaptic activity produces oxidant species that oxidize ascorbic acid to dehydroascorbic acid (DHA). DHA is released from neurons and taken up by astrocytes.

Astrocytes reduce DHA to ascorbic acid through glutathione-dependent reductases. In the presymptomatic stages of HD-like disease in mice, astrocytes do not release ascorbic acid efficiently

(Fig. 10b). In fact, over-supplementation with exogenous ascorbic acid is enough to produce an adequate ascorbic acid uptake. During the symptomatic stages of HD-like disease in mice,

exogenous ascorbic acid is not able to induce SVCT2 translocation to the plasma membrane as this process is Htt-dependent (Fig. 10b). Our results thus demonstrate that ascorbic acid

homeostasis is altered before the onset of overt HD-like behavioural symptoms in an HD mouse model. METHODS ANIMALS All Wistar rats (females, 6–12 weeks old) were obtained from the

Biosecurity Laboratory Facility for Animal Experimentation, Veterinary Science Faculty, Universidad Austral de Chile, Valdivia, Chile). All experiments were conducted in accordance with the

Chilean Government’s Manual of Bioethics and Biosafety (CONICYT: The Chilean Commission of Scientific and Technological Research, Santiago, Chile) and according to the guidelines established

by the Animal Protection Committee of the Universidad Austral de Chile. All R6/2 mice (male and female, 3–4 and 6–8 weeks old) used were obtained from the breeding colony maintained at the

University of California, Los Angeles (UCLA). Offspring were obtained from mouse pairings consisting of either male R6/2 hemizygote × female WT (strain B6CBAF1) or male WT × female WT with

an ovarian transplant from a R6/2 hemizygote (obtained from the Jackson Laboratory, Bar Harbor, ME). All experimental procedures were carried out in accordance with the National Institutes

of Health _Guide for the Care and Use of Laboratory Animals_ and were approved by the Institutional Animal Care and Use Committee at UCLA. Animals were maintained in a 12:12 light:dark cycle

environment and supplied with commercial food pellets and water _ad libitum._ Studies were performed on two age groups of R6/2 mice and with age-matched WT control mice based on development

of the overt motor phenotype. The first group of animals was tested before the appearance of overt behavioural symptoms at 3–4 weeks23. The second group was tested after the appearance of

the full behavioural phenotype at 8–12 weeks23. Efforts were made to minimize the number of animals used for experimental purposes. SLICE PREPARATION Mice were anaesthetized with isoflurane

(1-chloro-2,2,2-trifluoroethyl difluoromethyl ether) for decapitation. Brains were removed into ice-cold low-Ca2 oxygenated artificial cerebrospinal fluid (ACSF: 130 mM NaCl, 5 mM MgCl2, 1 

mM CaCl2, 3 mM KC, 1.25 mM NaH2PO4, 26 mM NaHCO3 and 10 mM glucose) and coronal slices (350 μm) were prepared using a microslicer. Slices containing striatum and overlying cortex were

maintained at room temperature in an incubation chamber filled with ACSF that was bubbled continuously with 95% O2—5% CO2 for 1 h before being transferred to a laminar flow, thin-layer

submersion recording chamber44,45. ELECTROPHYSIOLOGICAL ASSAYS Recordings were made from medium-sized spiny neurons from the striatum using the whole-cell patch-clamp configuration in

voltage-clamp. Medium-sized spiny neuronss were visualized in the dorsolateral striatum using infrared illumination with differential interference contrast optics (IR-DIC microscopy) and

identified by their somatic size and basic membrane properties. A −70 mV holding potential was maintained during the experiment. Electrophysiological data were acquired at 10 KHz. Recording

electrodes consisting of borosilicate glass had an impedance of 5–7 MΩ. The pipette internal solution was: 131 mmol l−1 potassium gluconate, 1 mmol l−1 EGTA, 1 mmol l−1 MgCl2, 2 mmol l−1

ATP-K2, 0.3 mmol l−1 NMDAGTP-Na, 6 mmol l−1 KCl, 1 mmol l−1 NaCl and 5 mmol/l HEPES (pH 7.3) osmolarity 290 mOsm l−1. Postsynaptic currents were evoked by a tungsten electrode placed into

the corpus callosum to stimulate the corticostriatal pathway. Stimulation pulses were 0.1–0.2 ms in duration of varying amplitude and were applied every 200 ms. pCLAMP 8 software (Axon

Instruments) was used to generate stimuli and for data display, acquisition and storage19,46. Drugs were superfused with gassed ACSF (extracellular substances) or were included within the

patch pipette (intracellular substances). UPTAKE ASSAYS Uptake assays were performed in 400 μl of incubation buffer (IB contains 15 mM hepes (pH 7.4), 135 mM NaCl, 5 mM KCl, 1.8 mM CaCl2,

0.8 mM MgCl2) containing 0.1±0.4 μCi of 1±14C-L-ascorbic acid (8.2 mCi mmol−1, Dupont NEN). Uptake was halted by washing cells with ice-cold IB containing 0.2 mM HgCl2. Cells were dissolved

in 200 μl of lysis buffer (10 mM Tris–HCl (pH 8.0), 0.2% SDS) and the incorporated radioactivity was measured by liquid scintillation spectrometry47. LIVE CELL IMAGING ANALYSIS For imaging

studies, cells were grown on coverslips coated with poly-L-lysine (Sigma). The cells were transfected using Lipofectamine 2000 (Invitrogen). Lipofectamine–DNA complexes were made by

incubating DNA plasmids and cationic lipids in Opti-MEM Reduced-Serum Medium (Invitrogen) at room temperature. TIRF microscopy analysis: cells expressing SVCT2-EGFP were visualized using an

objective-based TIRF microscope. A 473-nm solid-state diode laser was focused onto a single-mode optical fibre and transmitted via the rear illumination port of an Olympus IX70 inverted

microscope. Laser light was reflected from a double dichroic mirror (Z488/532rpc; Chroma Technology), passed through a high-numerical aperture objective ( × 60, NA 1.45 (oil immersion)

Olympus) and was totally internally reflected by the glass–water interface. Under these experimental conditions, fluorescent objects were within 150 nm of the glass–water interface48.

Fluorescence recovery after photobleaching: cells expressing SVCT2-EGFP were visualized with an Olympus Fluoview 1,000 confocal microscope. Spot areas of 0.18±0.06 μm2 were bleached using

argon-ion gas lasers, at wavelengths of 488 and 515 nm at 100%. The experimental data was corrected for any bleaching that may occur during recording, normalized to a pre-bleach fluorescence

intensity Fi=1, and fitted to the equation: In this equation, F0 is the fluorescence intensity immediately after photobleaching and F∞ is the intensity at the asymptote of the fluorescence

recovery after bleaching. The theoretical fit is adjusted to the characteristic recovery time, the post-bleach fluorescence intensity immediately after photobleaching, F0, and the

fluorescence intensity at time infinity, F∞, after photobleaching49. Mf is defined as the fraction of molecules that recover during the time-course of the experiment. Mf was calculated from

data obtained from the normalized recovery curves: BIOTINYLATION OF SURFACE PROTEINS Biotinylation assays were performed as described by Zambrano _et al._50 Briefly, cells (1 × 108) were

photolabelled with Sulfo-NHS-SS-Biotin (Thermo Fisher Scientific, Rockford, IL, USA). Cells were washed three times in ice-cold PBS (pH 8.0) to remove any contaminating proteins and were

resuspended at a concentration of 20 × 106 cells per ml in PBS (pH 8.0), with the addition of 80 μl of 10 mM Sulfo-NHS-SS-Biotin per ml of reaction volume. Cells were incubated at room

temperature for 30 min and again washed three times with ice-cold PBS (pH 8.0). The labelled cells were homogenized in 200 μl of buffer A (0.3 mmol l−1 sucrose, 3 mmol l−1 dithiothreitol

(DTT), 1 mmol l−1 EDTA, 100 μg ml−1 phenylmethylsulfonyl fluoride (PMSF) with Triton X-100 and then precipitated by continuous mixing with 40 μl of Immobilized NeutrAvidin Protein (Thermo

Fisher Scientific, Rockford, IL, USA) for 2 hours at room temperature. The precipitates were washed four times in 0.3% (_w_/_v_) Triton X-100 and twice in PBS. The resin-bound complex was

boiled in SDS–PAGE sample buffer. Samples were resolved in 10% (_w_/_v_) SDS–polyacrylamide gels and western blot analysis was performed. QUANTITATIVE REVERSE TRANSCRIPTION REAL-TIME PCR The

total RNA of the samples was extracted with the EZNA Total RNA Kit II (Omega Bio-Tek, Norcross, GA, USA) according to the manufacturer’s protocol. RNA quantification and purity were

analysed with a spectrophotometer. cDNA was synthesized using the Maxima Universal First Strand cDNA Synthesis Kit (Thermo Fisher Scientific, Rockford, IL, USA) and PCR was performed using

the Maxima SYBR Green qPCR Master Mix (Thermo Fisher Scientific). The following primers were used to analyse SVCT2 expression, sense: 5′-ATGATGGGTATCGGCAAGAACA-3′ and antisense:

5′-GCTCCGTGTCCTCGTTGTC-3′ (expected product size: 166 bp). WESTERN BLOT ANALYSIS Total protein extracts were obtained from R6/2 mice striatum or cell cultures. Biotinylated proteins were

obtained from cell cultures (see below). Dissected striata were homogenized in buffer A, 2 μg ml−1 pepstatin A, 2 μg ml−1 leupeptin and 2 μg ml−1 aprotinin) and sonicated three to five times

for 10 s at 4 °C. Proteins were resolved by SDS–PAGE (70 μg per lane for total protein extract) in a 10% (_w_/_v_) polyacrylamide gel, transferred to polyvinylidene difluoride membranes

(0.45-μm pore; Amersham Pharmacia Biotech., Piscataway, NJ, USA) and probed with anti-SVCT2 (1:1,000, SC-9927, Santa Cruz Biotechnology, Santa Cruz, CA, USA), anti-TfR (1:1,000, 13-6800,

Invitrogen, Carlsbad, CA, USA) and anti-b actin antibodies (1:1,000, sc-81178, Santa Cruz Biotechnology, Santa Cruz, CA, USA). The reaction was developed using anti-goat, anti-mouse HRP

conjugated antibodies (1:10,000, PA1-86326 and 31430, Thermo Scientific, Rockford, IL, USA) and the enhanced chemiluminescence western blot method50 (Amersham Biosciences, Pittsburgh, PA,

USA). IMMUNOFLUORESCENCE ANALYSIS Cells grown on poly-L-lysine-coated coverslips were fixed with HistoChoice Tissue Fixative (Sigma-Aldrich). Mice brains were dissected and fixed immediately

by immersion in 4% paraformaldehyde in 0.1 M phosphate buffer (pH 7.4). Samples were dehydrated in graded alcohol solutions, embedded in paraffin and incubated overnight with anti-SVCT2

antibodies (1:100, SC-9927, Santa Cruz Biotechnology), anti-MCT1 (1:100, AB1286, Chemicon International, Inc., Temecula, CA, USA), anti-HAP1 (1:100, MAB5516, Chemicon International, Inc.,

Temecula, CA, USA), anti-HAP40 (1:100, AB5872, Chemicon International, Inc), anti-Htt (1:100, MAB2166, Chemicon International, Inc.) and/or anti-EEA1 (1:100, sc-33585, Santa Cruz

Biotechnology). Samples were washed and incubated with anti-rabbit, anti-goat, anti-mouse or anti-chicken IgG-Alexa Fluor 488, IgG-Alexa Fluor 568 or IgG-Alexa Fluor 633 (1:300, A-11055,

A-21202, A-11039, A11057, A21070, Invitrogen) and propidium iodide (1.7 μg ml−1, Sigma-Aldrich) and then subsequently washed and mounted. An inverted Olympus FluoView 1000 confocal

microscope was used to examine the samples47. CELL CULTURES Cell lines: The hybridoma cell line (NG108, ATCC Number: HB-12317™) was grown in DMEM-F12 (United States Biological, Swampscott,

MA, USA) containing 10% foetal bovine serum (FBS: Hyclone, Logan, UT, USA) with HAT media supplement (final concentration: 0.1 mM hypoxanthine, 400 nM aminopterin, 0.016 mM thymidine;

Invitrogen), 50 U ml−1 penicillin, 50 mg ml−1 streptomycin, 50 ng ml−1 amphotericin B and 2 mM L-glutamine (Nalgene, Rochester, NY, USA). The human embryonic kidney cell line (HEK293, ATCC

number: CRL-1573) and mouse striatal STHdhQ7 and STHdhQ111 cells (Q7 and Q111 cells; Trettel _et al._25) were grown using DMEM (United States Biological) containing 10% fetal bovine serum

(FBS: Hyclone, Logan, UT, USA), 50 U ml−1 penicillin, 50 mg ml−1 streptomycin, 50 ng ml−1 amphotericin B and 2 mM L-glutamine (Nalgene, Rochester, NY, USA). Primary cultures: Striatal

neurons were obtained from 17-day-old embryos of Wistar rats, as described previously19. Adult female rats were anaesthetized using isoflurane (1-chloro-2,2,2-trifluoroethyl difluoromethyl

ether), embryo forebrains were removed and the striatum dissected. Tissue was digested with 0.12% trypsin (_w_/_v_, Gibco Co., Rockville, MD, USA) in 0.1 M phosphate buffer (PBS: pH 7.4,

osmolarity 320 mOsm) and mechanically disrupted with a fire-polished Pasteur pipette. Cells were plated at 0.3 × 106 cells per cm2 in plates onto coverslips coated with poly-L-lysine (mol.

wt>350 kDa, Sigma-Aldrich Corp. St Louis, MO, USA). After 20 min, floating cells were removed and attached cells were cultured for 5–7 days in neurobasal medium (Gibco) supplemented with

B27 (Gibco), 50 U ml−1 penicillin, 50 mg ml−1 streptomycin, 50 ng ml−1 amphotericin B and 2 mM L-glutamine (Nalgene). PLASMIDS Plasmid constructs, SVCT2-EGFP, encoding full-length SVCT2

fused at the C termini to EGFP (enhanced green fluorescent protein, Godoy _et al._, 2007)52 were kindly provided by Dr J.C. Vera (Departamento de Fisiopatología, Universidad de Concepción,

Concepción, Chile). Plasmid constructs, mHtt-DsRED, encoding Htt amino acids 1–588 containing 138 glutamine repeats fused to DsRED51 were kindly provided by Dr C. Hetz (Instituto de Ciencias

Biomédicas, Universidad de Chile, Santiago, Chile). STATISTICAL ANALYSES Statistical comparison between two or more groups of data was respectively performed using Student’s _t_-test or

analysis of variance (ANOVA) followed by the Bonferroni post-test. Linear and non-linear regressions were calculated using SigmaPlot v9.0 software (only correlations >0.9 were accepted).

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dependence on bivalent cations. _J. Biol. Chem._ 282, 615–624 (2007). Article  CAS  PubMed  Google Scholar  Download references ACKNOWLEDGEMENTS We gratefully acknowledge the technical

assistance of A. Riveros. This work was supported by Chilean FONDECYT grants 1110571 (M.A.C.), 1110906 (S.B.) and 1110508 (I.I.C.), Chilean FONDEF grant D111I1007 (C.H.), Chilean Ring

Initiative ACT1109 (C.H.), Millenium Institute P09-015-F (C.H.), CONICYT interantional grant USA2013-0003 (C.H.), CONICYT grant 7912010006 (R.L.V.), international SfA grant (M.A.C.),

DID-UACh Research Grant from the Universidad. AUTHOR INFORMATION Author notes * Aníbal I. Acuña, Magdalena Esparza and Carlos Kramm: These authors contributed equally to this work AUTHORS

AND AFFILIATIONS * Instituto de Bioquímica y Microbiología, Facultad de Ciencias, Universidad Austral de Chile, Campus Isla Teja s/n, Valdivia, 5090000, Chile Aníbal I. Acuña, Magdalena

Esparza, Carlos Kramm, Felipe A. Beltrán, Alejandra V. Parra, Ilona I. Concha & Maite A. Castro * Centro de Investigación Sur-Austral en Enfermedades del Sistema Nervioso (CISNe),

Universidad Austral de Chile, Campus Isla Teja s/n, Valdivia, 5090000, Chile Aníbal I. Acuña, Magdalena Esparza, Carlos Kramm, Felipe A. Beltrán, Alejandra V. Parra, Sebastián Brauchi & 

Maite A. Castro * Escuela de Graduados, Facultad de Ciencias, Universidad Austral de Chile, Campus Isla Teja s/n, Valdivia, 5090000, Chile Aníbal I. Acuña, Felipe A. Beltrán & Carlos A.

Toro * Intellectual and Developmental Disabilities Research Center, Semel Institute for Neuroscience and Human Behavior, Brain Research Institute, The David Geffen School of Medicine, 760

Westwood Plaza, University of California Los Angeles, Los Angeles, 90095-1759, California, USA Carlos Cepeda & Michael S. Levine * Instituto de Fisiología, Facultad de Medicina,

Universidad Austral de Chile, Campus Isla Teja s/n, Valdivia, 5090000, Chile Carlos A. Toro & Sebastián Brauchi * Instituto de Ciencias Biomédicas, Universidad de Chile, Avda,

Independencia 1027, Santiago, Chile René L. Vidal * Neurounion Biomedical Foundation, Independencia 1027, Santiago, Chile Claudio Hetz Authors * Aníbal I. Acuña View author publications You

can also search for this author inPubMed Google Scholar * Magdalena Esparza View author publications You can also search for this author inPubMed Google Scholar * Carlos Kramm View author

publications You can also search for this author inPubMed Google Scholar * Felipe A. Beltrán View author publications You can also search for this author inPubMed Google Scholar * Alejandra

V. Parra View author publications You can also search for this author inPubMed Google Scholar * Carlos Cepeda View author publications You can also search for this author inPubMed Google

Scholar * Carlos A. Toro View author publications You can also search for this author inPubMed Google Scholar * René L. Vidal View author publications You can also search for this author

inPubMed Google Scholar * Claudio Hetz View author publications You can also search for this author inPubMed Google Scholar * Ilona I. Concha View author publications You can also search for

this author inPubMed Google Scholar * Sebastián Brauchi View author publications You can also search for this author inPubMed Google Scholar * Michael S. Levine View author publications You

can also search for this author inPubMed Google Scholar * Maite A. Castro View author publications You can also search for this author inPubMed Google Scholar CONTRIBUTIONS The work

presented here was carried out in collaboration between all authors. A.I.A performed FRAP and TIRFM experiments in HEK293 cells, analysed the data, interpreted the results and helped in the

manuscript preparation. M.E. carried out experiments in HEK293 cells (uptake, biotinylation and colocalization assays). C.K. performed FRAP, TIRFM and colocalization assays in Q cells.

F.A.B. carried out sample collection from R6/2 mice and performed qPCR and histological experiments using samples from R6/2 mice. A.P. carried out biotinylation and western blot assays using

proteins from Q cells and R6/2 mice. C.C. and M.S.L. supervised and helped in data interpretation of electrophysiological experiments. C.C and M.A.C. performed electrophysiological

experiments. C.T. and S.B. supervised and helped in data interpretation and design of TIRFM experiments. R.V. and C.H. supervised and helped in all experiments using STHdhQ cells. S.B. and

I.I.C. co-designed and discussed analyses. M.A.C. carried out kinetic analyses, designed the project, organized the entire research and wrote the manuscript. CORRESPONDING AUTHOR

Correspondence to Maite A. Castro. ETHICS DECLARATIONS COMPETING INTERESTS The authors declare no competing financial interests. SUPPLEMENTARY INFORMATION SUPPLEMENTARY INFORMATION

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license, visit http://creativecommons.org/licenses/by-nc-nd/3.0/ Reprints and permissions ABOUT THIS ARTICLE CITE THIS ARTICLE Acuña, A., Esparza, M., Kramm, C. _et al._ A failure in energy

metabolism and antioxidant uptake precede symptoms of Huntington’s disease in mice. _Nat Commun_ 4, 2917 (2013). https://doi.org/10.1038/ncomms3917 Download citation * Received: 27 May 2013

* Accepted: 12 November 2013 * Published: 13 December 2013 * DOI: https://doi.org/10.1038/ncomms3917 SHARE THIS ARTICLE Anyone you share the following link with will be able to read this

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