The subthalamic nucleus (STN) is an element of cortico-basal ganglia-thalamo-cortical circuitry critical for action suppression. In Huntington's disease (HD) action suppression is impaired, resembling the effects of STN lesioning or inactivation. To explore this potential linkage, the STN was studied in BAC transgenic and Q175 knock-in mouse models of HD. At <2 and 6 months of age autonomous STN activity was impaired due to activation of KATP channels. STN neurons exhibited prolonged NMDA receptor-mediated synaptic currents, caused by a deficit in glutamate uptake, and elevated mitochondrial oxidant stress, which was ameliorated by NMDA receptor antagonism. STN activity was rescued by NMDA receptor antagonism or the break down of hydrogen peroxide. At 12 months of age approximately 30% of STN neurons had been lost, as in HD. Together, these data argue that dysfunction within the STN is an early feature of HD that may contribute to its expression and course.https://doi.org/10.7554/eLife.21616.001
The basal ganglia are a network of subcortical brain nuclei that are critical for action selection and central to the expression of several psychomotor disorders (Albin et al., 1989; Wichmann and DeLong, 1996). Information flow from the cortex to the output nuclei of the basal ganglia occurs via three major pathways. The so-called direct pathway through the striatum promotes movement and ‘rewarding’ behavior through inhibition of GABAergic basal ganglia output (Chevalier and Deniau, 1990; Kravitz et al., 2010; Kravitz and Kreitzer, 2012). In contrast, the indirect pathway via the striatum, external globus pallidus and subthalamic nucleus (STN) and the hyperdirect pathway through the STN suppress the same processes through elevation of basal ganglia output (Maurice et al., 1999; Tachibana et al., 2008; Kravitz et al., 2010; Kravitz and Kreitzer, 2012). Indeed, interruption of the indirect and hyperdirect pathways through lesion or inactivation of the STN is associated with elevated/involuntary movement, impulsivity and psychiatric disturbances such as hypomania and hyper-sexuality (Crossman et al., 1988; Hamada and DeLong, 1992; Baunez and Robbins, 1997; Bickel et al., 2010; Jahanshahi et al., 2015).
Huntington's disease (HD) is an autosomal dominant, neurodegenerative disorder caused by an expansion of CAG repeats in the gene (HTT) encoding huntingtin (HTT), a protein involved in vesicle dynamics and intracellular transport (Huntington’s Disease Collaborative Research Group, 1993; Saudou and Humbert, 2016). Early symptoms of HD include involuntary movement, compulsive behavior, paranoia, irritability and aggression (Anderson and Marder, 2001; Kirkwood et al., 2001). These symptoms have traditionally been linked to cortico-striatal degeneration, however a role for the STN is suggested by their similarity to those caused by STN inactivation or lesion. The hypoactivity of the STN in HD models in vivo (Callahan and Abercrombie, 2015a, 2015b) and the susceptibility of the STN to degeneration in HD (Lange et al., 1976; Guo et al., 2012) are also consistent with STN dysfunction.
Several mouse models of HD have been generated, which vary by length and species origin of HTT/Htt, CAG repeat length, and method of genome insertion. For example, one line expresses full-length human HTT with 97 mixed CAA-CAG repeats in a bacterial artificial chromosome (BAC; Gray et al., 2008), whereas Q175 knock-in (KI) mice have an inserted chimeric human/mouse exon one with a human polyproline region and ~188 CAG repeats in the native Htt (Menalled et al., 2012).
Increased mitochondrial oxidant stress exacerbated by abnormal NMDAR-mediated transmission and signaling has been reported in HD and its models (Fan and Raymond, 2007; Song et al., 2011; Johri et al., 2013; Parsons and Raymond, 2014; Martin et al., 2015). Several reports suggest that glutamate uptake is impaired due to reduced expression of the glutamate transporter EAAT2 (GLT-1) and/or GLT-1 dysfunction (Arzberger et al., 1997; Liévens et al., 2001; Behrens et al., 2002; Miller et al., 2008; Bradford et al., 2009; Faideau et al., 2010; Huang et al., 2010; Menalled et al., 2012; Dvorzhak et al., 2016; Jiang et al., 2016). However, others have found no evidence for deficient glutamate uptake (Parsons et al., 2016), suggesting that abnormal NMDAR-mediated transmission is caused by increased expression of extrasynaptic receptors and/or aberrant coupling to signaling pathways (e.g., Parsons and Raymond, 2014). The sensitivity of mitochondria to anomalous NMDAR signaling is likely to be further compounded by their intrinsically compromised status in HD (Song et al., 2011; Johri et al., 2013; Martin et al., 2015).
Although HD models exhibit pathogenic processes seen in HD, they do not exhibit similar levels and spatiotemporal patterns of cortico-striatal neurodegeneration. Striatal neuronal loss in aggressive Htt fragment models such as R6/2 mice does occur but only close to death (Stack et al., 2005), whereas full-length models exhibit minimal loss (Gray et al., 2008; Smith et al., 2014). Despite the loss and hypoactivity of STN neurons in HD and the similarity of HD symptoms to those arising from STN lesion or inactivation, the role of the STN in HD remains poorly understood. We hypothesized that the abnormal activity and progressive loss of STN neurons in HD may reflect abnormalities within the STN itself. This hypothesis was addressed in the BAC and Q175 KI HD models using a combination of cellular and synaptic electrophysiology, optogenetic interrogation, two-photon imaging and stereological cell counting.
Data are reported as median [interquartile range]. Unpaired and paired statistical comparisons were made with non-parametric Mann-Whitney U and Wilcoxon Signed-Rank tests, respectively. Fisher’s exact test was used for categorical data. p < 0.05 was considered statistically significant; where multiple comparisons were performed this p-value was adjusted using the Holm-Bonferroni method (adjusted p-values are denoted ph; Holm, 1979). Box plots show median (central line), interquartile range (box) and 10–90% range (whiskers).
STN neurons exhibit intrinsic, autonomous firing, which contributes to their role as a driving force of neuronal activity in the basal ganglia (Bevan and Wilson, 1999; Beurrier et al., 2000; Do and Bean, 2003). To determine whether this property is compromised in HD mice, the autonomous activity of STN neurons in ex vivo brain slices prepared from BACHD and wild type littermate (WT) mice were compared using non-invasive, loose-seal, cell-attached patch clamp recordings. 5–7 months old, symptomatic and 1–2 months old, presymptomatic mice were studied (Gray et al., 2008). Recordings focused on the lateral two-thirds of the STN, which receives input from the motor cortex (Kita and Kita, 2012; Chu et al., 2015). At 5–7 months, 124/128 (97%) WT neurons exhibited autonomous activity compared to 110/126 (87%) BACHD neurons (p = 0.0049; Figure 1A,B). The frequency (WT: 7.9 [5.2–12.6] Hz; n = 128; BACHD: 6.3 [1.4–10.2] Hz; n = 126; p = 0.0001) and regularity (WT CV: 0.27 [0.14–0.47]; n = 124; BACHD CV: 0.36 [0.20–0.80]; n = 110; p = 0.0012) of firing were reduced in BACHD neurons (Figure 1A,B). The distribution of firing frequency of WT neurons appears unimodal with a mode at ~6–8 Hz (Figure 1C), whereas the distribution of BACHD neurons is relatively bimodal with modes at ~0–2 Hz and ~8–10 Hz (Kolmogorov–Smirnov test, p = 0.0002; Figure 1C). This distribution suggests that BACHD neurons consist of a phenotypic population with compromised autonomous firing, and a non-phenotypic population with relatively normal autonomous firing. At 1–2 months 136/145 (94%) WT STN neurons were autonomously active versus 120/143 (84%) BACHD STN neurons (p = 0.0086). The frequency (WT: 9.8 [6.3–14.8] Hz; n = 145; BACHD: 7.1 [1.8–11.3] Hz; n = 143; p < 0.0001) and regularity (WT CV: 0.17 [0.13–0.26]; n = 136; BACHD CV: 0.23 [0.14–0.76]; n = 120; p = 0.0016) of firing were also reduced in BACHD neurons. Together, these data demonstrate that the autonomous activity of STN neurons in BACHD mice is impaired at both early presymptomatic and later symptomatic ages.
As described above, the majority of studies report that astrocytic glutamate uptake is diminished in the striatum in HD and its models. To test whether the STN of BACHD mice exhibits a similar deficit, EPSCs arising from the optogenetic stimulation of motor cortical inputs to the STN (as described by Chu et al., 2015) were compared in WT and BACHD mice before and after inhibition of GLT-1 and GLAST with 100 nM TFB-TBOA. STN neurons were recorded in ex vivo brain slices in the whole-cell voltage-clamp configuration using a cesium-based, QX-314-containing internal solution to maximize voltage control. Neurons were held at −40 mV and recorded in the presence of low (0.1 mM) external Mg2+ and the AMPAR antagonist DNQX (20 µM) to maximize and pharmacologically isolate the evoked NMDAR-mediated excitatory postsynaptic current (EPSC); analysis was performed on average EPSCs from 5 trials with 30 s recovery between trials (Figure 1D–H). The amplitude weighted decay time constant of the NMDAR EPSC was moderately but significantly prolonged in BACHD compared to WT neurons (WT: 38.1 [30.0–44.8] ms; n = 12; BACHD: 47.6 [38.7–55.9] ms; n = 11; p = 0.0455; Figure 1E–H). Subsequent application of TFB-TBOA increased the decay time constant of the NMDAR EPSC in STN neurons derived from WT (WT control: 39.0 [35.2–44.0] ms; WT TFB-TBOA: 50.2 [41.7–68.4] ms; n = 9; p = 0.0039; Figure 1E,H) but had no effect in BACHD neurons (BACHD control: 47.9 [38.9–59.4] ms; BACHD TFB-TBOA: 44.9 [34.7–52.2] ms; n = 10; p = 0.3223; Figure 1F,H). In control conditions, the amplitudes of EPSCs recorded from WT and BACHD neurons were similar (WT: 50.1 [34.7–61.0] pA; n = 12; BACHD: 45.6 [22.1–78.3] pA; n = 11; ph = 0.7399; Figure 2A) and there was no correlation between EPSC amplitude and the decay time constant in either group (WT: r2 = 0.16; n = 12; ph = 0.5871; BACHD: r2 = 0.10; n = 12; ph = 0.6686; Figure 2B). In order to increase spillover of glutamate from synaptic release sites, cortico-STN inputs were optogenetically stimulated 5 times at 50 Hz and the resulting compound NMDAR-mediated EPSC was compared in WT and BACHD STN neurons. Interestingly, the decay of compound NMDAR EPSCs under control conditions or following inhibition of glutamate uptake were not different in WT and BACHD mice (WT control: 79.0 [62.6–102.0] ms; n = 6; BACHD control: 65.2 [44.7–111.5] ms; n = 6; p = 0.4848; WT TFB-TBOA: 125.4 [106.8–146.6] ms; n = 6; BACHD TFB-TBOA: 108.3 [94.5–143.1] ms; n = 6; p = 0.6991; Figure 2C–E). Together, these data demonstrate that individual, but not compound, NMDAR-mediated cortico-STN synaptic EPSCs are prolonged in the BACHD model.
To test whether disrupted autonomous firing in BACHD is linked to NMDAR activation, brain slices from BACHD mice were incubated in control media or media containing the NMDAR antagonist D-AP5 (50 µM) for 3–5 hr prior to loose-seal, cell-attached recordings from STN neurons (Figure 3). D-AP5 treatment rescued autonomous firing in slices derived from 5–7 month old BACHD mice compared to untreated control slices (Figure 3A,B). The proportion of autonomously active neurons was greater in D-AP5 pre-treated slices (untreated: 18/30 (60%); D-AP5 treated: 29/30 (97%); p = 0.0011). The frequency (untreated: 1.0 [0.0–7.6] Hz; n = 30; D-AP5 treated: 13.2 [7.9–17.4] Hz; n = 30; p < 0.0001) and regularity (untreated CV: 0.43 [0.24–1.21]; n = 18; D-AP5 treated: CV: 0.13 [0.09–0.20]; n = 29; p < 0.0001) of autonomous firing were also greater in D-AP5 treated slices. In slices derived from 1–2 month old BACHD mice autonomous firing was also more prevalent in D-AP5 treated slices than in untreated slices (untreated: 10/30 (33%); D-AP5-treated: 27/30 neurons (90%); p < 0.0001) and the frequency of firing overall was greater (untreated: 0.0 [0.0–1.3] Hz; n = 30; D-AP5 treated: 8.7 [4.4–14.5] Hz; n = 30; p < 0.0001; Figure 3B). The regularity of autonomous firing was however not rescued (untreated CV: 0.61 [0.27–0.81]; n = 10; D-AP5-treated CV: 0.25 [0.14–0.69]; n = 27; p = 0.1368; Figure 3B). In slices from WT mice the rate (untreated: 10.4 [6.1–14.2] Hz; n = 27; D-AP5 treated: 5.6 [1.7–15.4] Hz; n = 27; p = 0.1683; Figure 3B), regularity (untreated: 0.18 [0.12–0.27]; n = 24; D-AP5 treated: 0.22 [0.12–0.46]; n = 22; p = 0.4785; Figure 3C) and incidence of firing (untreated: 24/27 (89%); D-AP5 treated 22/27 (81%); p = 0.7040; Figure 3D) were unaltered by D-AP5 treatment. Thus, prolonged blockade of NMDARs rescued autonomous firing in BACHD STN neurons but had no effect on autonomous activity in WT STN neurons. Together, these data demonstrate that NMDAR activation contributes to the disruption of autonomous activity in BACHD STN neurons.
NMDAR activation can elevate mitochondrial oxidant stress (Dugan et al., 1995; Moncada and Bolaños, 2006; Brennan et al., 2009; Nakamura and Lipton, 2011). To test whether STN neurons from BACHD mice exhibit increased mitochondrial oxidant stress, a mitochondria-targeted redox probe MTS-roGFP (Hanson et al., 2004) was virally expressed in 5–7-month-old BACHD mice and WT littermates (Figure 4A). 1–2 weeks later MTS-roGFP was imaged in brain slices under two-photon excitation with 890 nm light. Oxidant stress was estimated from the fluorescence of MTS-roGFP in individual neurons under baseline conditions relative to the fluorescence of MTS-roGFP under conditions of full reduction and oxidation in the presence of 2 mM dithiothreitol and 200 µM aldrithiol-4, respectively (Sanchez-Padilla et al., 2014). STN neurons were selected for analysis based on their appearance under two-photon, Dodt contrast imaging and were distinguishable from STN glial cells by their relatively large diameter (Figure 4A). STN neurons are reliably recorded when this selection strategy is employed to guide patch clamp recording (Atherton et al., 2008, 2010). MTS-roGFP imaging revealed that relative oxidant stress in BACHD STN neurons was elevated compared to WT (WT: 0.35 [0.18–0.42]; n = 23; BACHD: 0.40 [0.37–0.63]; n = 24; p = 0.0332; Figure 4B). In a separate experiment (performed 15 months later) to test whether NMDAR activation ex vivo contributed to elevated mitochondrial oxidant stress, brain slices from a different cohort of BACHD mice were treated for >3 hr in 50 µM D-AP5 prior to imaging. MTS-roGFP imaging confirmed that D-AP5-treated slices exhibited lower oxidant stress compared to untreated slices from the same mice (untreated: 0.39 [0.35–0.48]; n = 13; D-AP5-treated: 0.32 [0.24–0.42]; n = 17; p = 0.0445; Figure 4C). Thus, STN neurons from BACHD mice exhibit elevated mitochondrial oxidant stress, which can be reduced by antagonism of NMDARs.
NMDAR receptor-generated mitochondrial oxidant stress in BACHD may lead to the activation of KATP channels, which act as metabolic sensors and homeostatic regulators of excitability in multiple cell types (Nichols, 2006). STN neurons abundantly express all the molecular components of KATP channels including the Kir6.2 pore-forming subunit of the KATP channel (Thomzig et al., 2005) and the SUR1, SUR2A and SUR2B regulatory subunits (Karschin et al., 1997; Zhou et al., 2012). To determine whether KATP channels are responsible for impaired firing in BACHD mice, the effects of the KATP channel inhibitor glibenclamide (100 nM) on WT and phenotypic BACHD autonomous firing ex vivo were compared. Glibenclamide application increased both the rate (BACHD control: 5.7 [1.4–9.1] Hz; BACHD glibenclamide: 11.3 [9.2–13.3] Hz; n = 15; p = 0.0003) and regularity (BACHD control CV: 0.41 [0.16–1.43]; BACHD glibenclamide CV: 0.16 [0.12–0.32]; n = 14; p = 0.0001) of autonomous firing in 5–7-month-old BACHD STN neurons (Figure 5A,B). In contrast KATP channel inhibition had no effect on the firing of WT neurons (WT control frequency: 13.2 [8.4–19.6] Hz; WT glibenclamide frequency: 15.7 [9.7–18.4] Hz; n = 8; p = 0.4609; WT control CV: 0.18 [0.11–0.21]; WT glibenclamide CV: 0.14 [0.11–0.19]; n = 8; p = 0.1094).
To further examine the effects of KATP channels on autonomous firing, whole-cell current clamp recordings were obtained from 5–7-month-old BACHD mice and WT littermates (Figure 5C,D). Consistent with the hyperpolarizing and shunting effects of KATP channels, the interspike voltage trajectory was shallower in BACHD neurons compared to WT (WT: 413.8 [317.1–705.0] mV/s; n = 7; BACHD: 125.3 [59.2–298.0] mV/s; n = 7; ph = 0.0210). In addition, the rate, regularity and interspike voltage trajectory of autonomous firing of BACHD STN neurons recorded in this configuration increased following application of glibenclamide (BACHD control frequency: 7.7 [4.5–13.4] Hz; BACHD glibenclamide frequency: 15.1 [8.5–20.9] Hz; n = 7; p = 0.0156; BACHD control CV: 0.24 [0.13–0.35]; BACHD glibenclamide CV: 0.13 [0.10–0.15]; n = 7; p = 0.0313; BACHD control interspike voltage trajectory: 125.3 [59.2–298.0] mV/s; BACHD glibenclamide interspike voltage trajectory: 369.9 [156.7–474.0] mV/s; n = 7; ph = 0.0313; Figure 5C,D). In contrast inhibition of KATP channels did not alter the firing of WT neurons (WT control frequency: 14.7 [12.5–27.4] Hz; WT glibenclamide frequency: 16.8 [8.3–20.3] Hz; n = 7; p = 0.3750; WT control CV: 0.10 [0.07–0.18]; WT glibenclamide CV: 0.08 [0.06–0.16]; n = 7; p = 0.8125; WT control interspike voltage trajectory: 413.8 [317.1–705.0] mV/s; n = 7; WT glibenclamide interspike voltage trajectory: 361.9 [216.7–522.9] mV/s; n = 7; ph = 0.3750; Figure 5C,D).
Glibenclamide can also activate Epac2 and Rap1 (Zhang et al., 2009), which could rescue firing through a pathway that is independent of KATP channel inhibition. Therefore, gliclazide, a sulfonylurea that has no effect on Epac2/Rap1 signaling (Zhang et al., 2009; Takahashi et al., 2015), was applied to STN neurons of 5–7-month-old BACHD mice (Figure 6). In loose-seal, cell-attached recordings gliclazide (10 µM) increased both firing rate (control: 5.1 [2.0–8.0] Hz; gliclazide: 9.9 [4.6–13.8] Hz; n = 6; p = 0.0313; Figure 6) and regularity (control CV: 0.54 [0.31–0.88]; gliclazide CV: 0.29 [0.10–0.47]; n = 6; p = 0.0313; Figure 6) in phenotypic BACHD STN neurons. Together, these data argue that KATP channels are responsible for the impaired autonomous activity of STN neurons in the BACHD model.
As described above, 3–5 hr NMDAR antagonism with D-AP5 partially rescued autonomous activity in BACHD STN neurons. To determine whether this rescue was mediated through effects on KATP channels, glibenclamide was applied following this treatment. D-AP5 pre-treatment partially occluded the increases in the autonomous firing rate (BACHD glibenclamide Δ frequency: 4.3 [2.2–8.7] Hz, n = 15; D-AP5 pre-treated BACHD glibenclamide Δ frequency: 1.9 [0.7–3.2] Hz, n = 6; p = 0.0365) and regularity (BACHD glibenclamide Δ CV: −0.25 [−0.85– −0.13], n = 14; D-AP5 pre-treated BACHD glibenclamide Δ CV: −0.09 [−0.10– −0.03], n = 6; p = 0.0154) that accompany KATP channel inhibition. Thus, these observations are consistent with the conclusion that prolonged NMDAR antagonism partially rescued autonomous activity in BACHD STN neurons through a reduction in KATP channel-mediated firing disruption.
To further examine whether elevated NMDAR activation can trigger a homeostatic KATP channel-mediated reduction in autonomous firing in WT STN, brain slices from 2-month-old C57BL/6 mice were incubated in control media or media containing 25 µM NMDA for 1 hr prior to recording (Figure 7). NMDA pre-treatment reduced the proportion of autonomously firing neurons (untreated: 66/75 (88%); NMDA: 65/87 (75%); p = 0.0444) and the frequency (untreated: 14.9 [7.8–24.8] Hz; n = 75; NMDA: 5.2 [0.0–14.0] Hz; n = 87; ph < 0.0001) and regularity (untreated CV: 0.13 [0.08–0.25]; n = 66; NMDA CV: 0.24 [0.10–0.72]; n = 65; ph = 0.0150; Figure 7A–C) of autonomous activity relative to control slices. In a subset of neurons glibenclamide was applied to inhibit KATP channels. In neurons from untreated slices glibenclamide had no effect on firing rate (control: 16.6 [10.9–31.3] Hz; glibenclamide: 25.0 [16.3–32.8] Hz; n = 6; ph = 0.2188; Figure 7A–D) or regularity (control CV: 0.08 [0.07–0.37]; glibenclamide CV: 0.08 [0.06–0.09]; n = 6; ph = 0.3125; Figure 7A–D). However, in neurons from NMDA pre-treated slices glibenclamide application elevated firing rate (control: 3.3 [2.3–5.1] Hz; glibenclamide: 11.4 [10.8–24.4] Hz; n = 10; ph = 0.0078; Figure 7A–D) and regularity (control CV: 0.83 [0.25–1.03]; glibenclamide CV: 0.12 [0.07–0.16]; n = 8, ph = 0.0208; Figure 7A–D) to levels similar to that seen in untreated slices. Together, these data demonstrate that increased activation of STN NMDARs can lead to a persistent KATP channel-mediated homeostatic reduction in autonomous activity in STN neurons.
Mitochondrial oxidative phosphorylation generates superoxide, which may be dismuted by superoxide dismutase to produce H2O2 (Adam-Vizi, 2005). Superoxide and hydrogen peroxide can also be produced by NADPH oxidase (Brennan et al., 2009). Because KATP channels are activated by H2O2 (Ichinari et al., 1996; Avshalumov et al., 2005), we tested whether H2O2 contributes to KATP channel-mediated disruption of ex vivo autonomous activity in the BACHD model. The effect of a membrane permeable form of the enzyme catalase (polyethylene glycol-catalase), which breaks down H2O2, on the autonomous firing of STN neurons from 4–6-month-old BACHD mice was examined (Figure 8). Application of 250 U/ml catalase increased the rate (BACHD control: 3.4 [0.7–5.5] Hz; BACHD catalase: 10.8 [7.6–13.8] Hz; n = 11; ph = 0.0080; Figure 8A–C) and regularity (BACHD control CV: 1.0 [0.3–2.1]; BACHD catalase CV: 0.17 [0.12–0.21]; n = 11; ph = 0.0060; Figure 8A–C) of autonomous firing. Subsequent application of glibenclamide (100 nM) had no additional effect on firing rate (11.8 [8.2–14.4] Hz; n = 11; ph = 0.9658) or regularity (CV: 0.15 [0.12–0.23]; n = 11; ph = 0.4922; Figure 8A–C). Thus, these results suggest that H2O2 underlies KATP channel activation in BACHD STN neurons.
To test if the actions of H2O2 on autonomous firing are confined to its effects on KATP channels, these experiments were repeated in the presence of glibenclamide. As seen previously, application of glibenclamide increased firing rate (BACHD control: 5.2 [1.0–6.7] Hz; BACHD glibenclamide: 8.5 [7.2–11.6] Hz; n = 8, ph = 0.0156; Figure 8D) and regularity (BACHD control CV: 0.83 [0.27–1.30]; BACHD glibenclamide CV: 0.23 [0.15–0.58]; n = 8; ph = 0.0156; Figure 8D). Subsequent application of catalase had no additional effect on firing rate (8.7 [7.2–14.1] Hz; n = 8; ph = 0.6406; Figure 8D) but did produce a small but statistically significant increase in regularity (CV: 0.14 [0.11–0.21]; n = 8; ph = 0.0156; Figure 8D). In WT mice, catalase application did not change firing rate (WT control: 11.0 [10.5–14.2] Hz; WT catalase: 14.3 [11.3–18.3] Hz; n = 7; p = 0.0781; Figure 9) but lead to a small but statistically significant increase in regularity (WT control CV: 0.12 [0.10–0.23]; WT catalase CV: 0.11 [0.07–0.13]; n = 7; p = 0.0469; Figure 9). The effects of catalase on the frequency and regularity of firing in BACHD mice were greater than those observed in WT mice (frequency: p = 0.0154; CV: p = 0.0007; Figure 9). Together, these data suggest that suppression of autonomous activity of STN neurons in BACHD mice is largely mediated by the modulatory effect of H2O2 on KATP channels.
To test if the elevation of oxidant stress can result in KATP channel activation in WT STN neurons, glutathione peroxidase was inhibited with mercaptosuccinic acid (MCS) (Avshalumov et al., 2005). Following the application of 1 mM MCS both the rate (control: 12.0 [7.8–13.5] Hz; MCS: 9.0 [4.8–10.5] Hz; n = 11; ph = 0.0068; Figure 10) and regularity (control CV: 0.21 [0.15–0.22]; MCS CV: 0.30 [0.22–0.34]; n = 11; ph = 0.0137) of firing decreased. Subsequent application of glibenclamide rescued both firing rate (14.6 [10.3–19.2] Hz; n = 11; ph = 0.0020) and regularity (CV: 0.12 [0.12–0.17]; n = 11; ph = 0.0098; Figure 10). These data are also consistent with an action of H2O2 on STN KATP channels.
HD patients exhibit 20–30% STN neuron loss (Lange et al., 1976; Guo et al., 2012). Because mitochondrial oxidant stress and reactive oxygen species can trigger apoptotic pathways leading to cell death (Green and Reed, 1998; Bossy-Wetzel et al., 2008), the number of STN neurons in 12-month-old BACHD and WT mice were compared (Figure 11). The brains of BACHD mice and WT littermates were first fixed by transcardial perfusion of formaldehyde, sectioned into 70 µm coronal slices and immunohistochemically labeled for neuronal nuclear protein (NeuN). The total number of NeuN-immunoreactive STN neurons and the volume of the STN were then estimated using unbiased stereological techniques. Both the total number of STN neurons (WT: 10,793 [9,070–11,545]; n = 7; BACHD: 7,307 [7,047–9,285]; n = 7; p = 0.0262) and the volume of the STN (WT: 0.087 [0.084–0.095] mm3; n = 7; BACHD: 0.078 [0.059–0.081] mm3; n = 7; p = 0.0111; Figure 11A,B) were reduced in 12-month-old BACHD versus WT mice. The density of STN neurons was not different in BACHD and WT mice (WT: 121,248 [107,180–126,139] neurons/mm3; n = 7; BACHD: 115,273 [90,377–135,765] neurons/mm3; n = 7; p = 0.8048; Figure 11A,B). To determine whether the difference in cell number represents an early developmental abnormality or a progressive loss of adult neurons, the numbers of neurons in 2-month-old BACHD and WT mice were also compared. At 2-months-old, the total number of STN neurons (WT: 10,373 [9,341–14,414]; n = 7; BACHD: 10,638 [10,513–13,877]; n = 7; p = 0.7104; Figure 11C), the volume of the STN (WT: 0.098 [0.090–0.125] mm3; n = 7; BACHD: 0.085 [0.080–0.111] mm3; n = 7; p = 0.1649; Figure 11C) and STN neuronal density (106,880 [98,100–115,985] neurons/mm3; n = 7; BACHD: 124,844 [115,479–145,711] neurons/mm3; n = 7; p = 0.1282; Figure 11C) were not different in WT and BACHD mice. Together, these data demonstrate that between the ages of 2 months and 12 months BACHD mice lose approximately one third of their STN neurons compared to WT littermates.
STN neurons from BACHD mice exhibit perturbed autonomous firing that is caused by NMDAR activation/signaling leading to mitochondrial oxidant stress, H2O2 generation and KATP channel activation. Furthermore, STN neurons are progressively lost in BACHD mice. To determine whether these features are specific to the BACHD model or a more general feature of HD models, a subset of experiments were repeated in heterozygous Q175 KI mice (Figure 12). STN neurons from 6-month-old Q175 mice exhibited a severely reduced rate of autonomous activity (WT: 7.8 [1.9–14.7] Hz; n = 90; Q175: 0.0 [0.0–6.3] Hz; n = 90; p < 0.0001; Figure 12A,B), though the regularity of active neurons was unchanged (WT CV: 0.2 [0.1–0.6]; n = 77; Q175 CV: 0.4 [0.1–1.0]; n = 42; p = 0.1506; Figure 12A,B). Additionally, there was a large decrease in the proportion of active neurons in the Q175 STN (WT: 77/90 (86%); Q175: 42/90 (47%); p < 0.0001).
Inhibition of KATP channels with glibenclamide rescued both STN firing rate and regularity in Q175 and increased regularity only in WT (WT control frequency: 9.7 [5.4–13.5] Hz; WT glibenclamide frequency: 10.3 [7.4–15.4] Hz; n = 8; p = 0.1094; Q175 control frequency: 4.8 [3.5–6.2] Hz; Q175 glibenclamide frequency: 11.0 [9.3–13.6] Hz; n = 6; p = 0.0313; WT control CV: 0.19 [0.13–0.47]; WT glibenclamide CV: 0.11 [0.10–0.21]; n = 8; p = 0.0078; Q175 control CV: 0.45 [0.35–0.71]; Q175 glibenclamide CV: 0.15 [0.10–0.17]; n = 6; p = 0.03125; Figure 12C,D). Similar to BACHD, Q175 STN neurons recovered to WT-like firing rate following >3 hr pretreatment with D-AP5 (Q175 control: 4.6 [0.0–11.4] Hz; n = 45; Q175 D-AP5 treated: 11.6 [0.0–18.7] Hz; n = 45; p = 0.0144; Figure 12E,F), although the regularity (Q175 control CV: 0.16 [0.10–0.66]; n = 15; Q175 D-AP5 treated CV: 0.14 [0.09–0.32]; n = 12; p = 0.2884; Figure 12E,F) and proportion of active neurons (Q175 control: 30/45 (67%); Q175 D-AP5 treated: 33/45 (73%); p = 0.6460; Figure 12E,F) were unaltered. The 12-month-old Q175 STN (n = 7) exhibited a median 26% reduction in the total number of STN neurons with no effect on other parameters (WT: 8,661 [7,120–9,376] neurons; Q175: 6,420 [5,792–7,024] neurons; p = 0.0111; WT volume: 0.081 [0.074–0.087] mm3; Q175 volume: 0.079 [0.070–0.091] mm3; p = 0.6200; WT density: 109,477 [82,180–115,301] neurons/mm3; Q175 density: 88,968 [63,624–103,020] neurons/mm3; p = 0.2086; Figure 12G,H). Taken together, these data show that the STN exhibits similar dysfunction and neuronal loss in both the transgenic BACHD and Q175 KI mouse models of HD.
Dysfunction of the striatum and cortex has been extensively characterized in HD models, but relatively few studies have examined the extra-striatal basal ganglia. Here, we report early NMDAR, mitochondrial and firing abnormalities together with progressive loss of STN neurons in two HD mouse models. Furthermore, dysfunction was present in HD mice prior to the onset of major symptoms, implying that it occurs early in the disease process (Gray et al., 2008; Menalled et al., 2012). Cell death in the STN also preceded that in the striatum, as STN neuronal loss was observed at 12 months of age in both BACHD and Q175 mice, a time point at which striatal neuronal loss is absent but psychomotor dysfunction is manifest (Gray et al., 2008; Heikkinen et al., 2012; Smith et al., 2014; Mantovani et al., 2016). Together these findings argue that dysfunction within the STN contributes to the pathogenesis of HD.
Astrocytes appear to play a pivotal role in HD. Expression of mutant huntingtin in astrocytes alone is sufficient to recapitulate neuronal and neurological abnormalities observed in HD and its models (Bradford et al., 2009; Faideau et al., 2010). Furthermore, astrocyte-specific rescue approaches ameliorate some of the abnormalities observed in HD models (Tong et al., 2014; Oliveira et al., 2016). In the STN, inhibition of GLT-1 (and GLAST) slowed individual NMDAR EPSCs in WT but not BACHD mice and eliminated the differences in their decay kinetics, arguing that impaired uptake of glutamate by astrocytes contributed to the relative prolongation of NMDAR-mediated EPSCs in BACHD STN neurons. Interestingly, and in contrast to the striatum (Milnerwood et al., 2010), when spillover of glutamate onto extrasynaptic receptors was increased by train stimulation and inhibition of astrocytic glutamate uptake, the resulting compound NMDAR EPSC and its prolongation by uptake inhibition were similar in BACHD and WT mice, arguing against an increase in extrasynaptic STN NMDAR expression/function in BACHD mice. Slowing of astrocytic glutamate uptake has recently been shown to occur during increased presynaptic activity (Armbruster et al., 2016). Thus, train stimulation may have slowed glutamate uptake sufficiently to occlude/eliminate the differences in uptake that were observed in BACHD and WT STN neurons during single stimulation. Whether the modest differences in glutamate uptake that were observed here are sufficient to promote NMDAR-mediated dysfunction in HD STN neurons remains to be determined.
NMDARs play a key role in the abnormal activity of STN neurons in HD models. Antagonism of STN NMDARs in BACHD and Q175 brain slices rescued autonomous STN firing. Conversely, acute activation of STN NMDARs persistently disrupted STN firing in WT brain slices. If the relatively low level of glutamatergic transmission present ex vivo is sufficient to impair firing then this impairment is likely to be more severe in vivo where STN neurons are powerfully patterned by glutamatergic transmission arising from the cortex, thalamus, pedunculopontine nucleus and superior colliculus (reviewed by Bevan, 2017). Non-synaptic sources of extracellular glutamate, such as diffusion/release from astrocytes (Cavelier and Attwell, 2005; Lee et al., 2013) may also contribute to excessive NMDAR activation in HD mice.
Extended antagonism of NMDARs in BACHD slices also reduced mitochondrial oxidant stress in STN neurons. NMDAR activation can elevate ROS through a variety of Ca2+- and nitric oxide-associated signaling pathways and their actions on mitochondria, NADPH oxidase and antioxidant expression (Dugan et al., 1995; Moncada and Bolaños, 2006; Brennan et al., 2009; Nakamura and Lipton, 2011; Valencia et al., 2013). Although we saw no evidence of basal mitochondrial dysfunction that was not attributable to increased NMDAR function, there is considerable evidence that mutant huntingtin causes transcriptional dysregulation, which leads to defective mitochondrial quality control, an increase in the proportion of defective, ROS generating mitochondria and an increase in opening of the permeability transition pore (Milakovic and Johnson, 2005; Panov et al., 2002; Fernandes et al., 2007; Song et al., 2011; Chaturvedi et al., 2013; Johri et al., 2013; Martin et al., 2015). Thus, basal mitochondrial dysfunction could render HD STN neurons especially sensitive to NMDAR-mediated transmission and signaling.
Catalase rapidly restored autonomous firing in the BACHD model, an effect occluded by inhibition of KATP channels, arguing that H2O2, through its action on KATP channels is the major cause of firing disruption. H2O2 can act on KATP channels by decreasing their sensitivity to ATP (Ichinari et al., 1996), reducing the ratio of ATP to ADP (Krippeit-Drews et al., 1999), and/or modulating channel gating through a sGC-cGMP-PKG-ROS(H2O2)-ERK1/2-calmodulin-CaMKII signaling pathway (Zhang et al., 2014). H2O2 is likely to directly modulate STN KATP channels in HD mice because disrupted firing was also observed when STN neurons were recorded in the whole-cell configuration with patch pipettes containing exogenous ATP. Furthermore, H2O2 break down rapidly rescued activity, consistent with a direct action on KATP channels. H2O2-dependent modulation of KATP channels has been extensively characterized in midbrain dopamine neurons where it powerfully suppresses cellular excitability and synaptic transmission (Avshalumov et al., 2005; Bao et al., 2009). The activation of KATP channels in STN neurons may represent a form of homeostasis that suppresses firing when mitochondrial oxidant stress is high, limiting the possibility of oxidant damage and bioenergetic failure (Ray et al., 2012; Sena and Chandel, 2012).
In HD, chronic oxidant stress can lead to damage, such as lipid and protein peroxidation and nuclear/mitochondrial DNA damage, which profoundly impair cellular function and promote cell death (Perluigi et al., 2005; Browne and Beal, 2006; Acevedo-Torres et al., 2009). Consistent with the negative effects of such processes on neuronal viability, we observed progressive loss of STN neurons in both the BACHD and Q175 models. Furthermore, the level of neuronal loss at 12 months in the BACHD and Q175 models was similar to that observed in HD patients (Lange et al., 1976; Guo et al., 2012). The absence of neuronal loss in the cortex and striatum in the same models at an equivalent time point suggests that STN dysfunction and degeneration may be particularly influential in the early disease process. Although the STN is known to degenerate in HD, it is not clear why neuronal loss is ultimately less than that observed in the striatum at the end stage of the disease, despite the fact that dysfunction and degeneration occur earlier (at least in HD models). Future research will be required to determine whether subtypes of STN neurons exhibit selective vulnerability and/or whether the processes promoting their degeneration, e.g. cortical activation of STN NMDARs, ultimately wane.
As a key component of the hyperdirect and indirect pathways, the STN is critical for constraining cortico-striatal activity underlying action selection (Albin et al., 1989; Oldenburg and Sabatini, 2015). In the ‘classical’ model of basal ganglia function, degeneration of indirect pathway striatal projection neurons is proposed to underlie the symptoms of early stage HD (Albin et al., 1989). Here we show for the first time that STN dysfunction and neuronal loss precede cortico-striatal abnormalities in HD models. Thus, dysfunction and degeneration of cortical and striatal neurons occurs in concert with profound changes in other elements of the basal ganglia. Therapeutic strategies that target the STN may therefore be useful not only for treating the psychomotor symptoms of early- to mid-stage HD but also for influencing dysfunction and degeneration throughout the cortico-basal ganglia-thalamo-cortical circuit.
All animal procedures were performed in accordance with the policies of the Society for Neuroscience and the National Institutes of Health, and approved by the Institutional Animal Care and Use Committee of Northwestern University. Adult male hemizygous BACHD mice (RRID:IMSR_JAX:008197) and heterozygous Q175 mice (RRID:IMSR_JAX:027410), their WT litter mates, and C57BL/6 mice (Charles River Laboratories International, Inc., Wilmington, MA, USA) were used in this study.
Mice were anesthetized with 1–2% isoflurane (Smiths Medical ASD, Inc., Dublin, OH, USA). AAV vectors (serotype 9; ~1012–13 GC/ml) engineered to express hChR2(H134R)-eYFP under the hSyn promoter (University of Pennsylvania Vector Core, Philadelphia, PA, USA) or MTS-roGFP under the CMV promoter (Sanchez-Padilla et al., 2014) were injected under stereotaxic guidance (Neurostar, Tubingen, Germany). In order to express hChR2(H134R)-eYFP, AAV was injected bilaterally into the primary motor cortex (three injections per hemisphere; coordinates relative to bregma: AP, +0.6 mm, + 1.2 mm, and +1.8 mm; ML, + 1.5 mm, and −1.5 mm; DV, −1.0 mm; 0.3 μl per injection). In order to express MTS-roGFP, AAV was injected bilaterally into the STN (coordinates: AP, −2.06 mm; ML, +1.4 mm, and −1.4 mm; DV, −4.5 mm; 0.4 µl per injection). Brain slices were prepared from AAV-injected mice 10–14 days after injection.
Mice were lightly anesthetized with isoflurane, deeply anesthetized with ketamine/xylazine (87/13 mg/kg i.p.) and then perfused transcardially with ~10 ml of ice-cold sucrose-based artificial cerebrospinal fluid (sACSF) that contained 230 mM sucrose, 2.5 mM KCl, 1.25 mM NaH2PO4, 0.5 mM CaCl2, 10 mM MgSO4, 10 mM glucose, and 26 mM NaHCO3 equilibrated with 95% O2 and 5% CO2. The brain was removed, immersed in sASCF and 250 µm sagittal slices were cut with a vibratome (VT1200S; Leica Microsystems Inc., IL, USA). Slices were then transferred to a holding chamber, immersed in ACSF that contained 125 mM NaCl, 2.5 mM KCl, 1.25 mM NaH2PO4, 2 mM CaCl2, 2 mM MgSO4, 10 mM glucose, 26 mM NaHCO3, 1 mM sodium pyruvate, and 5 µM L-glutathione equilibrated with 95% O2 and 5% CO2, held at 35°C for 30–45 min, then maintained at room temperature.
Individual brain slices were placed in a recording chamber where they were perfused at 4–5 ml/min with synthetic interstitial fluid (SIF) at 35°C that contained 126 mM NaCl, 3 mM KCl, 1.25 mM NaH2PO4, 1.6 mM CaCl2, 1.5 mM MgSO4, 10 mM glucose and 26 mM NaHCO3 equilibrated with 95% O2 and 5% CO2. Somatic recordings were obtained under visual guidance (Axioskop FS2, Carl Zeiss, Oberkochen, Germany) using computer-controlled manipulators (Luigs and Neumann, Ratingen, Germany). Loose-seal cell-attached recordings were made using 3–5 MΩ impedance borosilicate glass pipettes (Warner Instruments, Hamden, CT, USA) that were filled with 140 mM NaCl, 23 mM glucose, 15 mM HEPES, 3 mM KCl, 1.5 mM MgCl2, 1.6 mM CaCl2 (pH 7.2 with NaOH, 300–310 mOsm/l). Whole-cell voltage clamp recordings were made using 3–5 MΩ pipettes filled with 135 mM CsCH3O3S, 10 mM QX-314, 10 mM HEPES, 5 mM phosphocreatine, 3.8 mM NaCl, 2 mM Mg1.5ATP, 1 mM MgCl2, 0.4 mM Na3GTP, 0.1 mM Na4EGTA (pH 7.2 with CsOH, 290 mOsm/l). Whole-cell current clamp recordings were made using 10–15 MΩ pipettes filled with 130 mM KCH3SO4, 3.8 mM NaCl, 1 mM MgCl2, 10 mM HEPES, 5 mM phosphocreatine di(tris) salt, 0.1 mM Na4EGTA, 0.4 mM Na3GTP, and 2 mM Mg1.5ATP (pH 7.3 with KOH; 290 mOsm/l). Electrophysiological records were acquired using a computer running Clampex 10 software (Molecular Devices, Palo Alto, CA, USA; RRID:SCR_011323) connected to a Multiclamp 700B amplifier (Molecular Devices) via a Digidata 1322 A digitizer (Molecular Devices). Data were low-pass filtered at 10 kHz and sampled at 50 kHz. Liquid junction potentials of 10 and 9 mV were accounted for in whole-cell voltage clamp and current clamp recordings respectively, and in voltage clamp recordings series resistance and membrane capacitance were corrected online. All recordings of autonomous action potential generation were made in the acute presence of 50 µM D-AP5, 20 µM DNQX, 10 µM GABAzine, and 2 µM CGP 55845 to block synaptic transmission.
MTS-roGFP-expressing neurons were imaged at 890 nm with 76 MHz pulse repetition and ~250 fs pulse duration at the sample plane. Two-photon excitation was provided by a G8 OPSL pumped Mira 900 F laser (Coherent, Santa Clara, CA, USA) and sample power was regulated by a Pockels cell electro-optic modulator (model M350-50-02-BK, Con Optics, Danbury, CT, USA). Images were acquired using an Ultima 2 P system running PrairieView 5 (Bruker Nano Fluorescence Microscopy, Middleton, WI, USA) and a BX51WI microscope (Olympus, Tokyo, Japan) with a 60 × 0.9 NA objective (UIS1 LUMPFL; Olympus). After baseline fluorescence had been measured, the maximum and minimum fluorescence were determined by the application of 2 mM dithiothreitol and then 200 µM aldrithiol-4 to fully reduce and oxidize the tissue, respectively. The relative oxidation at baseline, a measure of oxidative stress, was then calculated (Sanchez-Padilla et al., 2014).
Mice were lightly anesthetized with isoflurane, deeply anesthetized with ketamine/xylazine (87/13 mg/kg i.p.) and then perfused transcardially with ~5 ml of phosphate buffered saline (PBS) followed by 30 ml of 4% formaldehyde in 0.1 M phosphate buffer (pH 7.4). Brains were removed and post-fixed for 2 hr in 4% formaldehyde, then washed in PBS. Brains were blocked and 70 µm thick coronal sections containing the STN were cut using a vibratome (VT1000S; Leica). Sections were washed in PBS and incubated for 48 hr at 4°C in anti-NeuN (clone A60; MilliporeSigma, Darmstadt, Germany; RRID:AB_2298772) at 1:200 in PBS with 0.2% Triton X-100 (MilliporeSigma) and 2% normal donkey serum. Sections were then washed in PBS and incubated for 90 min at room temperature in Alexa Fluor 488 donkey anti-mouse IgG (1:250; Jackson Immunoresearch, West Grove, PA, USA; RRID:AB_2340846) in 0.2% Triton X-100 and 2% normal donkey serum. Then the sections were washed in PBS and mounted on glass slides in Prolong Gold anti-fade medium (Thermo Fisher Scientific, Waltham, MA, USA).
NeuN labeled sections were imaged using an Axioskop two microscope (Carl Zeiss) with a 100 × 1.3 NA oil immersion objective (Plan-Neofluar 1018–595; Carl Zeiss). Unbiased stereological counting of STN neurons in a single hemisphere was performed using the optical fractionator technique (West et al., 1991) as implemented in Stereo Investigator (MBF Bioscience, Williston, VT, USA; RRID:SCR_002526), using a counting frame of 50 µm × 50 µm × 8 µm and a grid size of 150 µm × 150 µm; all sections containing the STN were used for counting (~8 sections). STN volume was calculated from the sum of the areal extent of the STN on each section multiplied by the section thickness (70 µm). For all individual counts the Gundersen Coefficient of Error (CE) (Gundersen et al., 1999) was less than 0.1 (0.080 [0.075–0.090]), and the investigator performing the counting was blinded to the genotype of the mouse.
All drugs used in electrophysiology and imaging experiments were diluted to working concentration in SIF and bath applied. D-AP5, CGP 55845, DNQX, GABAzine (SR 95531), NMDA and gliclazide were purchased from Abcam (Cambridge, MA, USA). Glibenclamide, TFB-TBOA and DL-Dithiothreitol were purchased from Tocris Bioscience (Bristol, UK). Catalase (polyethylene glycol-catalase), aldrithiol-4 and MCS were purchased from Sigma-Aldrich (St. Louis, MO, USA).
Electrophysiological data were analyzed using routines running in Igor Pro 6 and 7 (Wavemetrics, Portland, OR, USA; RRID:SCR_000325) or matplotlib (Hunter, 2007; RRID:SCR_008624). The firing rate of STN neurons was calculated from 1 min of recording or 100 action potentials (whichever covered the longer time period). Imaging data were analyzed using FIJI (Schindelin et al., 2012; RRID:SCR_002285). Statistical analyses were performed in Prism 5 (GraphPad Software, San Diego, CA, USA; RRID:SCR_002798) or R (http://www.r-project.org/; RRID:SCR_001905, RRID:SCR_000432). In order to make no assumptions about the distribution of the data, non-parametric statistical tests were used, and data are reported as median [interquartile range]; outliers were not excluded from the analysis. An α-level of 0.05 was used for two-way statistical comparisons performed with the Mann-Whitney U test for unpaired data (represented with box plots), the Wilcoxon signed rank test for paired data (tilted line segment plots), Fisher's exact test for categorical data (bar plots) or the F-test for linear regression. Where datasets were used in multiple comparisons the p-value was adjusted to maintain the family-wise error rate at 0.05 using the Holm-Bonferroni method (Holm, 1979); adjusted p-values are denoted ph. Box plots show median (central line), interquartile range (box) and 10–90% range (whiskers). For the primary findings reported in the manuscript, sample sizes for Mann-Whitney and Wilcoxon tests were estimated to achieve a minimum of 80% power using formulae described by Noether (1987). The effect sizes used in these power calculations were estimated using data randomly drawn from uniform distributions (runif() function in R stats package). For Mann-Whitney tests, with a 50 percentile change in median between groups X and Y (the interquartile ranges of the groups don’t overlap) P(Y > X) ≈ 0.88 giving an estimation that at least 10 observations per group would be needed to achieve 80% power; for a 25 percentile change (the median of Y falls outside the interquartile range of X) P(Y > X) ≈ 0.72 and the estimated requirement is at least 27 observations per group. For Wilcoxon tests, if all pairs of observations show the same direction of change, P(X + X’ > 0)= 1 giving an estimation that at least 10 observations would be needed to achieve 80% power (note though that it is possible to show empirically that 6 observations gives 100% power in this case); if 90% of observations show the same direction of change, P(X + X’ > 0) ≈ 0.98 and the estimated requirement is at least 12 pairs of observations.
An overview of psychiatric symptoms in Huntington's diseaseCurrent Psychiatry Reports 3:379–388.https://doi.org/10.1007/s11920-996-0030-2
Glutamate clearance is locally modulated by presynaptic neuronal activity in the cerebral cortexJournal of Neuroscience 36:10404–10415.https://doi.org/10.1523/JNEUROSCI.2066-16.2016
Changes of NMDA receptor subunit (NR1, NR2B) and glutamate transporter (GLT1) mRNA expression in Huntington's disease--an in situ hybridization studyJournal of Neuropathology and Experimental Neurology 56:440–454.https://doi.org/10.1097/00005072-199704000-00013
Autonomous initiation and propagation of action potentials in neurons of the subthalamic nucleusThe Journal of Physiology 586:5679–5700.https://doi.org/10.1113/jphysiol.2008.155861
Mitochondria are the source of hydrogen peroxide for dynamic brain-cell signalingJournal of Neuroscience 29:9002–9010.https://doi.org/10.1523/JNEUROSCI.1706-09.2009
Bilateral lesions of the subthalamic nucleus induce multiple deficits in an attentional task in ratsEuropean Journal of Neuroscience 9:2086–2099.https://doi.org/10.1111/j.1460-9568.1997.tb01376.x
The subthalamic nucleusIn: H Steiner, K. Y Tseng, editors. Handbook of Basal Ganglia Structure and Function. Amsterdam, Netherlands: Academic Press, Elsevier. pp. 277–291.
Cognitive and neuropsychiatric effects of subthalamotomy for Parkinson's diseaseParkinsonism & Related Disorders 16:535–539.https://doi.org/10.1016/j.parkreldis.2010.06.008
NADPH oxidase is the primary source of superoxide induced by NMDA receptor activationNature Neuroscience 12:857–863.https://doi.org/10.1038/nn.2334
Oxidative damage in Huntington's disease pathogenesisAntioxidants & Redox Signaling 8:2061–2073.https://doi.org/10.1089/ars.2006.8.2061
Tonic release of glutamate by a DIDS-sensitive mechanism in rat hippocampal slicesThe Journal of Physiology 564:397–410.https://doi.org/10.1113/jphysiol.2004.082131
Mitochondrial diseases of the brainFree Radical Biology and Medicine 63:1–29.https://doi.org/10.1016/j.freeradbiomed.2013.03.018
Disinhibition as a basic process in the expression of striatal functionsTrends in Neurosciences 13:277–280.https://doi.org/10.1016/0166-2236(90)90109-N
N-methyl-D-aspartate (NMDA) receptor function and excitotoxicity in Huntington's diseaseProgress in Neurobiology 81:272–293.https://doi.org/10.1016/j.pneurobio.2006.11.003
The efficiency of systematic sampling in stereology--reconsideredJournal of Microscopy 193:199–211.https://doi.org/10.1046/j.1365-2818.1999.00457.x
Striatal neuronal loss correlates with clinical motor impairment in Huntington's diseaseMovement Disorders 27:1379–1386.https://doi.org/10.1002/mds.25159
Investigating mitochondrial redox potential with redox-sensitive green fluorescent protein indicatorsJournal of Biological Chemistry 279:13044–13053.https://doi.org/10.1074/jbc.M312846200
A simple sequentially rejective multiple test procedureScandinavian Journal of Statistics 6:65–70.https://doi.org/10.2307/4615733
Direct activation of the ATP-sensitive potassium channel by oxygen free radicals in guinea-pig ventricular cells: its potentiation by MgADPJournal of Molecular and Cellular Cardiology 28:1867–1877.https://doi.org/10.1006/jmcc.1996.0179
Parkinson's disease, the subthalamic nucleus, inhibition, and impulsivityMovement Disorders 30:128–140.https://doi.org/10.1002/mds.26049
PGC-1α, mitochondrial dysfunction, and Huntington's diseaseFree Radical Biology and Medicine 62:37–46.https://doi.org/10.1016/j.freeradbiomed.2013.04.016
Progression of symptoms in the early and middle stages of Huntington diseaseArchives of Neurology 58:273–278.https://doi.org/10.1001/archneur.58.2.273
Interference of H2O2 with stimulus -secretion coupling in mouse pancreatic beta-cellsThe Journal of Physiology 514:471–481..https://doi.org/10.1111/j.1469-7793.1999.471ae.x.
Morphometric studies of the neuropathological changes in choreatic diseasesJournal of the Neurological Sciences 28:401–425.https://doi.org/10.1016/0022-510X(76)90114-3
Impaired glutamate uptake in the R6 Huntington's disease transgenic miceNeurobiology of Disease 8:807–821.https://doi.org/10.1006/nbdi.2001.0430
Autophagy in Huntington disease and huntingtin in autophagyTrends in Neurosciences 38:26–35.https://doi.org/10.1016/j.tins.2014.09.003
Mitochondrial respiration and ATP production are significantly impaired in striatal cells expressing mutant huntingtinJournal of Biological Chemistry 280:30773–30782.https://doi.org/10.1074/jbc.M504749200
Nitric oxide, cell bioenergetics and neurodegenerationJournal of Neurochemistry 97:1676–1689.https://doi.org/10.1111/j.1471-4159.2006.03988.x
Sample size determination for some common nonparametric testsJournal of the American Statistical Association 82:645–647.https://doi.org/10.1080/01621459.1987.10478478
Proteomic analysis of protein expression and oxidative modification in r6/2 transgenic mice: a model of Huntington diseaseMolecular & Cellular Proteomics 4:1849–1861.https://doi.org/10.1074/mcp.M500090-MCP200
Fiji: an open-source platform for biological-image analysisNature Methods 9:676–682.https://doi.org/10.1038/nmeth.2019
Chronology of behavioral symptoms and neuropathological sequela in R6/2 Huntington's disease transgenic miceThe Journal of Comparative Neurology 490:354–370.https://doi.org/10.1002/cne.20680
Motor cortical control of internal pallidal activity through glutamatergic and GABAergic inputs in awake monkeysEuropean Journal of Neuroscience 27:238–253.https://doi.org/10.1111/j.1460-9568.2007.05990.x
Elevated NADPH oxidase activity contributes to oxidative stress and cell death in Huntington's diseaseHuman Molecular Genetics 22:1112–1131.https://doi.org/10.1093/hmg/dds516
Functional and pathophysiological models of the basal gangliaCurrent Opinion in Neurobiology 6:751–758.https://doi.org/10.1016/S0959-4388(96)80024-9
Harry T OrrReviewing Editor; University of Minnesota, United States
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Thank you for submitting your article "Early dysfunction and progressive degeneration of the subthalamic nucleus in mouse models of Huntington's disease" for consideration by eLife. Your article has been favorably evaluated by Huda Zoghbi (Senior Editor) and three reviewers, one of whom is a member of our Board of Reviewing Editors. The following individual involved in review of your submission has agreed to reveal their identity: Margaret Rice (Reviewer #2).
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This is a very interesting study that implicates enhanced NMDAR signaling, leading to increased mitochondrial oxidant production, including that of hydrogen peroxide which triggers increased KATP activity, and results in slower and more variable autonomous STN neuronal firing in two HD mouse models (BACHD and Q175). This is one of the first studies to explore pathophysiological changes occurring early in the STN of HD mouse models. Overall the reviews are very positive about this work and recognize its important contribution to understanding the early pathogenesis in the STN using tow HD mouse models.
Revise the Abstract, Discussion and text of Results by removing conclusions on the role of glutamate uptake in the enhanced NMDA receptor-mediated oxidative stress.
This body of work from Atherton et al. is a rather extensive study on the impact mutant Htt has on the STN. Involvement of the STN in HD has been debated over the years and this work, at least in mouse models of HD, provides the strongest evidence to data that the STN is a central aspect of HD pathology. A strength is that their observation/results were obtained using two full length Htt mouse models of HD. Starting at early stages of disease in these models they found that STN neurons had decreased intrinsic firing rate and related this to activation of KATP channels. This is followed by an elevated extra-synaptic NMDA activation by increased glutamate levels (results of decreased astrocyte uptake). Lastly, they show that at later stages, 12 months, there is a significant loss of STN neurons. Overall the data are strong making a convincible demonstration that the STN is involved in these models of HD. This is important as therapies for HD move to the clinic. This work provides a strong basis for the importance of assessing STN efficacy of treatments
This report describes a thoughtful and thorough series of studies that not only illuminate a critical role for subthalamic nucleus (STN) neurons in mouse models of Huntington's disease, but also point to underlying mechanisms, including impaired glutamate uptake, NMDA receptor activation, H2O2 generation, and consequent K-ATP channel opening that slows STN neuron firing. This discussion carefully considers these factors in terms of cause versus effect. For all of these reasons, the studies should advance the field. There are a few generally minor concerns; most concern how the data are presented and discussed.
1) The notion that the direct pathway promotes "thoughts" is not obviously supported by the references cited (Introduction, first paragraph); cognitive function and thinking are not interchangeable terms.
2) The difference in frequency between WT and BACHD mice (is mean or median reported in the text?) was relatively small (7.9 vs 6.3 Hz, or a decrease of 20%); however, the examples in Figure 1A imply a four-fold difference. More accurately representative records should be used. The data shown in Figure 3A better represent the means reported; however, the much lower range for the untreated cells from BACHD mice (1.0 Hz) than those from BACHD reported in Figure 1 (6.3 Hz) needs some explanation. Were the untreated cells also recorded after the 3-5 h period as for AP5 pre-incubation?
3) The control data for MTS-roGFP imaging in the STN of BACHD mice appear to differ between Figure 3B and C, with a lack of obvious difference between control BACHD in 3C and WT in 3B. Do statistical analyses indicate a difference between control BACHD data in 3C and WT, or between BACHD data in 3B and 3C? The meaning of "relative oxidation" as the y-axis title also needs clarification. How this was determined is explained in the Methods, but should also be noted briefly in the Results. As written, the text makes this murkier rather than clearer: "MTS-roGFP imaging revealed that the relative oxidant stress in BACHD STN neurons was elevated relative to WT."
4) Figure 6A shows data from the cell that had the greatest change with glicazide, not one representative of the mean (or median) response (6B). Similarly, 7B show a cell in NMDA with 3 spikes per 2 s, less than 2 Hz, whereas the lowest frequency in the range reported was 2.3 (subsection “NMDAR activation produces a persistent KATP channel-mediated disruption of autonomous activity in WT STN neurons”).
5) The results with exogenous NMDA do implicate NMDA-induced reactive oxygen species, but the more convincing experiment would have been to block NMDA receptors in BACHD mice (as in the MTS-roGFP imaging studies; Figure 4). They had an opportunity to test this in studies with Q175 mice (Figure 12), but although they show that glibenclamide and AP5 each lead to an increase in STN neurons firing rate in slices from Q175 mice, the key experiment of whether the effect of glibenclamide is absent when NMDA receptors are blocked by APV is not reported. This would strengthen the authors' conclusions, but is not essential.
6) It is not clear whether the decreased firing rate of STN neurons in slices from HD mice reflected inclusion of neurons that did not show autonomous activity (0 Hz). The frequency range for BACHD mice does not include 0 (subsection “The autonomous activity of STN neurons is disrupted in the BACHD model”), but the range for Q175 mice does (subsection “The STN of Q175 KI mice exhibits similar abnormalities to those observed in the BACHD model”).
H2O2Figuresthat Reviewer #3:
This is a very interesting study that implicates enhanced NMDAR signaling, leading to increased mitochondrial oxidative stress that triggers increased KATP activity, resulting in slowed, more variable autonomous STN neuronal firing in two HD mouse models (BACHD and Q175). This study includes a series of elegant experiments with results that convincingly demonstrate a mechanism for altered STN autonomous firing, implicating a pathological process independent of aberrant iSPN input to STN in HD.
One limitation, however, is that although the authors suggest this STN firing alteration underlies a variety of HD-related behaviors, this hypothesis is not tested. Moreover, they don't explore any in vivo recording to confirm altered STN firing or responsiveness to cortical and/or striatal iSPN input. As well, the link between these molecular alterations, aberrant autonomous firing, and STN neuronal loss is not established. The latter is complicated by the fact that the STN is part of the indirect pathway, which is impaired early at the level of striatal iSPNs (early loss of enkephalin and DARPP32, enhanced excitability, loss of LTP, increased extra-synaptic NMDAR signaling, etc.).
In addition, there are some specific concerns:
1) It seems counter-intuitive that by increasing the glutamate load (e.g., with pulse trains), a basal impairment in glutamate transport would no longer be observed; one would think it might instead be augmented. However, this could make sense in the context of a recent study by Armbruster et al. (2016), showing that train stimulation itself slows glutamate uptake, which might occlude a small difference in the NMDAR EPSC decay time constant, as the authors have postulated for the TFB-TBOA. The phenomenon reported by Armbruster et al. seems a more likely explanation for the difference in results, rather than what the authors suggest, that impaired glutamate uptake is restricted to the synapse. The latter seems unlikely since astrocytes ensheath more than one synapse.
2) Related to #1, the NMDAR component of the EPSC decay time constant can be quite variable (as shown in Figure 1G; also see Vicini et al., 1998), and other factors (e.g. differences in access resistance, space clamp, etc.) could potentially impact the accuracy of measuring decay time in the context of such small currents. Moreover, the difference between the mean decay time constant in wild-type vs. BAC HD STN neurons is barely significant (P=0.0455). To further test the idea that impaired glutamate uptake is responsible for slowed decay for single NMDAR EPSCs, the authors could try using the glutamate scavenger GPT to see if this can accelerate decay time and show a greater effect in HD than wild-type, and/or add dextran to the extracellular solution to slow diffusion – this should magnify any differences in transporter function between the genotypes (see Mahfooz et al. NBD 2016 for approach).
3) In general, the focus on impaired glutamate uptake as a determinant of enhanced NMDAR-mediated oxidative stress seems unsupported. First, the authors rely heavily on previous studies showing reduced mRNA and/or protein expression levels of GLT1, but this does not prove impaired glutamate uptake under physiological conditions. In fact, few studies have measured clearance of synaptically released glutamate in brain slice: of these, one study in striatum demonstrated no impairment of glutamate uptake/clearance in R6/2 or YAC28 mice (Parsons et al., 2016), while two others show either minimal impairment with prolonged (200 ms), high concentration (3 mM) glutamate pulses (Dvorzhak et al., 2016) or slowing only of the late component of the iGluSnFr response (> 2 sec after synaptic release; Jiang et al., 2016).
Second, the difference in NMDAR EPSC decay shown in the current study is very small and is eliminated in the setting of trains of 5 stimuli at 50Hz (see points 1 and 2); since the authors mention that cortical input to STN frequently occurs in bursts, it seems unlikely that the Glu uptake impairment they see for single EPSCs could be an important contributor to increased oxidative stress in HD vs. wild-type STN. These points should be addressed in the Discussion, and less emphasis placed on the role of impaired glutamate uptake, which is not directly measured in this study.
4) The authors show increased mitochondrial oxidation and altered firing rate in BAC HD STN that is rescued by APV – this implicates NMDAR activity in these downstream events. However, it could be because of altered signaling via the NMDAR complex, i.e. due to NMDAR association with different effector proteins, rather than a result of increased calcium influx via NMDARs. This could be tested by comparing intracellular calcium transients in STN neurons between the genotypes using GCaMP6.https://doi.org/10.7554/eLife.21616.034
- Jeremy F Atherton
- Mark D Bevan
- Eileen L McIver
- David L Wokosin
- Mark D Bevan
- Eileen L McIver
- David L Wokosin
- D James Surmeier
- Mark D Bevan
The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.
This study was funded by CHDI Foundation and by NIH NINDS Grants 2 R37 NS041280 and 2 P50 NS047085. We thank Drs. Vahri Beaumont (CHDI) and Ignacio Munoz-Sanjuan (CHDI) for their comments on the work and Sasha Ulrich, Danielle Schowalter, and Marisha Alicea for management of mouse colonies.
Animal experimentation: This study was performed in accordance with the policies of the Society for Neuroscience and the National Institutes of Health. All animals were handled according to approved Institutional Animal Care and Use Committee protocols (IS00001185) of Northwestern University. All procedures were performed under isoflurane or ketamine/xylazine anesthesia, and every effort was made to minimize suffering.
- Harry T Orr, Reviewing Editor, University of Minnesota, United States
© 2016, Atherton et al.
This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.