# Developmental alterations of indirect-pathway medium spiny neurons in mouse models of Huntington's disease

Margaux Lebouc, Léa Bonamy, Thibault Dhellemmes, Jakob Scharnholz, Quentin Richard, Gilles Courtand, Alexandre Brochard, Frédéric Martins, Marc Landry, Jérôme Baufreton, et al.

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Margaux Lebouc, Léa Bonamy, Thibault Dhellemmes, Jakob Scharnholz, Quentin Richard, et al. Developmental alterations of indirect-pathway medium spiny neurons in mouse models of Huntington's disease. Neurobiology of Disease, 2025, 208, pp.106874. 10.1016/j.nbd.2025.106874. hal-05361428

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# Developmental alterations of indirect-pathway medium spiny neurons in mouse models of Huntington's disease

<figure><img src="image_3.png" /></figure>

Margaux Lebouc<sup>a</sup>, Léa Bonamy<sup>a</sup>, Thibault Dhellemmes<sup>a</sup>, Jakob Scharnholz<sup>a</sup>, Quentin Richard<sup>a</sup>, Gilles Courtand<sup>b</sup>, Alexandre Brochard<sup>c</sup>, Frédéric Martins<sup>c</sup>, Marc Landry<sup>a</sup>, Jérôme Baufreton<sup>a, *, 1</sup>, Maurice Garret<sup>b, 1</sup>

## ARTICLE INFO

### Keywords:

- Striatal development

- Huntingtin

- Electrophysiology

- Neuronal excitability

- Glutamatergic transmission

- Neuronal morphology

## ABSTRACT

Huntington's disease (HD) is a complex neurodegenerative disorder with cognitive and motor symptoms that typically manifest in adulthood. However, embryonic brain development impairments leading to cortical defects in HD mutation carriers has been shown recently supporting a neurodevelopmental component in HD. Despite HD is primarily recognized as a striatal pathology, developmental alterations in this structure, particularly during the early postnatal period, remain poorly understood. To fill this gap, we examined striatal development in newborn R6/1 mice. We found that D2 receptor-expressing indirect-pathway medium spiny neurons (D2-MSNs) present in the matrix striatal compartment undergo early morphological and electrophysiological maturation. Altered electrophysiological properties were also observed in newborn CAG140 mice. Additionally, we also observed a D2-MSN-selective reduction in glutamatergic cortico-striatal transmission at the beginning of the second postnatal week as well as a reduced projection of D2-MSNs onto the GPe at birth in R6/1 mice. All these alterations were transient with the circuit normalizing after the second postnatal week. These results identify a compartment- and cell-type specific defect in D2-MSNs maturation, which can contribute in their latter vulnerability, as this cell-type is the first to degenerate in HD during adulthood.

## 1. Introduction

Huntington's disease (HD) is caused by a dominant mutation in the gene encoding the huntingtin protein (Htt), leading to progressive degeneration in both striatal and cortical structures. While the symptoms of HD typically manifest in adulthood (Ross et al., 2019; Tabrizi et al., 2022), neuroimaging studies have identified early morphological alterations in presymptomatic mutation carriers. These changes may not be fully explained by neurodegeneration alone, and instead suggest a potential neurodevelopmental origin (for review see (Nopoulos et al., 2007, 2011; Lee et al., 2012; Kubera et al., 2019; Mangin et al., 2020; Ratié and Humbert, 2024). Remarkably, studies on children carrying the HD mutation have demonstrated striatal alterations as early as six years old, further supporting the idea that neurodevelopmental processes are disrupted long before clinical onset (Van Der Plas et al., 2019;

Tereshchenko et al., 2020). Additionally, the study of fetal HD mutation carriers revealed abnormalities in cortical neural progenitor cells, further confirming the neurodevelopmental hypothesis of HD (Barnat et al., 2020).

These early alterations have also been observed in HD models, where mutated Htt is known to impact multiple aspects of cortical development, including cortical progenitor cell division, neuronal migration, dendritic maturation and glutamatergic transmission (McKinstry et al., 2014; Molina-Calavita et al., 2014; Barnat et al., 2017, 2020; Capizzi et al., 2022). Importantly, studies have shown that the expression of mutant Htt or reduction of normal Htt expression exclusively during developmental stages is sufficient to induce HD-like symptoms in adulthood, highlighting the long-lasting consequences of early developmental disturbances (Arteaga-Bracho et al., 2016; Molero et al., 2016).

<footnote><sup>a</sup> Univ. Bordeaux, CNRS, IMN, UMR 5293, F-33000 Bordeaux, France</footnote>

<footnote><sup>b</sup> Univ. Bordeaux, CNRS, INCIA, UMR 5297, F-33000 Bordeaux, France</footnote>

<footnote><sup>c</sup> Univ. Bordeaux, INSERM, Neurocentre Magendie, U1215, F-33000 Bordeaux, France</footnote>

<footnote>* Corresponding author.</footnote>

<footnote>E-mail address: jerome.baufreton@u-bordeaux.fr (J. Baufreton).</footnote>

<footnote n="1">¹ These authors share seniority on the study.</footnote>
Despite the well-established striatal pathology in adult human HD carriers and mouse models (Vonsattel et al., 1985; Pouladi et al., 2013), research on early striatal development remains limited. The striatum is divided into striosomes and matrix compartments predominantly composed of medium spiny neurons (MSNs), which are divided into two sub-populations: D1 dopamine receptor-expressing MSNs (D1-MSNs) and D2 receptor-expressing MSNs (D2-MSNs). During the first postnatal weeks, the striatum undergoes significant developmental changes, shaping the morphological and electrophysiological properties of striatal neurons as well as their connectivity with other brain regions (Tepper et al., 1998; Dehorter, 2011; Lieberman et al., 2018; Krajeski et al., 2019). Early alterations including delays in embryonic striatal neurogenesis have been observed in HD models (Molero et al., 2009; Lebouc et al., 2020). However, it remains unclear whether striatal neurons are

able to integrate properly into the circuit during the early postnatal period. The maturation of neuronal activity and glutamatergic transmission during this time appears essential for maintaining a functional circuit into adulthood (Braz et al., 2022).

The striatum being the primary site of HD pathology, establishing alterations during critical periods of striatal maturation is highly relevant to better understand HD pathophysiology. To address this question, we used R6/1 and CAG140 mice crossed with D1- or D2-GFP reporter mice to study striatal MSN development during the first two postnatal weeks, focusing on their electrophysiological and morphological properties as well as their synaptic connectivity. We report that mutant Htt induces developmental alterations selectively in D2-MSNs belonging to the matrix compartment of the striatum.

<figure><img src="image_4.png" /><figcaption>**Fig. 1.** D2-MSN density is reduced in the matrix of R6/1 mice. A. Illustration of immunohistochemistry images showing Ctip2 (yellow, a selective marker of MSNs) and MOR (cyan, used to label striosomes) in striatal sections of WT::D2-GFP and R6/1::D2-GFP mice. B. Quantification of striatal surface area in WT and HD mice. C. Proportions of matrix (yellow) and striosome (cyan) areas relative to the total striatal area. D. Total density of Ctip2+ MSNs in WT and HD mice. E. Representative images of the striosomal compartment showing GFP (magenta, D2-MSNs), Ctip2 (green, MSNs), and MOR (blue, striosomes). Quantification of D2-MSN (Ctip2+/GFP+) and D1-MSN (Ctip2+/GFP−) density within the striosomes over time. F. Representative images of the matrix compartment showing GFP (magenta, D2-MSNs), Ctip2 (green, MSNs), and MOR (blue, marking striosomes). Quantification of D1-MSN (Ctip2+/GFP−) and D2-MSN (Ctip2+/GFP+) density within the matrix over time. Scale bars: 500 µm (A) and 30 µm (E-F). (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>
## 2. Results

### 2.1. Selective reduction of matrix D2-MSNs in HD mice

To identify D1- and D2-MSNs, homozygous D1- and D2-GFP mice were crossed with R6/1 mice to generate WT and R6/1 littermates. Immuno-labeling was then performed to detect GFP expression, combined with Ctip2, a nuclear marker for all striatal MSNs, and μ-opioid receptor (MOR), a specific marker for striosomes (Pert et al., 1976; Tinterri et al., 2018).

Given previous evidence suggesting altered neurogenesis in the embryonic striatum of HD mice (Molero et al., 2009), we investigated whether the Htt mutation affects early postnatal striatal morphology. To address this, we first examined total striatal surface and the relative proportions of striosomes (MOR+) and matrix (MOR-) areas across six postnatal ages (P0, P3, P6, P8, P10 and P15, N = 3 per age and genotype). No significant differences were found between WT and R6/1 mice in either striatal surface or the relative proportions of matrix and striosomes (Fig. 1B, C). We then assessed the evolution of MSN density (Ctip2+) in WT and R6/1 mice across these same ages. As expected, MSN density decreased as striatal volume expanded. However, there was no significant difference in the total number of Ctip2+ cells between genotypes, indicating that the Htt mutation does not affect the overall number of MSNs during early development (Fig. 1D).

To refine our analysis, we next quantified the densities of D1-MSNs (Ctip2+ GFP-) and D2-MSNs (GFP+) within the matrix (MOR-) and striosomes (MOR+). Although no significant changes were observed in the striosomes (Fig. 1E), a marked reduction in D2-MSN density was detected specifically in the matrix at P8 in R6/1 mice suggesting a compartment- and subtype-specific alteration (Fig. 1F).

In R6/1::D2-GFP mice, we confirmed that the D2-GFP line reliably labels D2-MSNs, with approximately 45 % of neurons GFP+ at P0 (Fig. S1A-G). In contrast, in the R6/1::D1-GFP line (Fig. S2A-G), the percentage of D1-MSNs labeled with GFP among Ctip2+ MSNs was initially low during the earliest stages. Between P0 and P6, GFP labeling was mainly found within the striosomes consistent with prior studies showing low D1 receptor expression at embryonic and early postnatal stages (Rao et al., 1991; Caille et al., 1995; Jung and Bennett, 1996; Araki et al., 2007; Yang et al., 2021).

### 2.2. Altered electrophysiological maturation of D2-MSNs in HD mice

Adult MSNs are normally characterized by a hyperpolarized resting membrane potential and a high latency to spike (Gertler et al., 2008). Both *in vivo* and *ex vivo* recordings have shown that during development, MSNs are initially hyperexcitable, gradually developing their mature phenotype over the first postnatal weeks (Dehorter, 2011; Peixoto et al., 2016; Lieberman et al., 2018; Krajeski et al., 2019). To determine whether the Htt mutation impacts this maturation, we carried out *ex vivo* electrophysiological recordings in acute slices from WT or R6/1 mice at 4 postnatal stages: P0–2, P3–6, P7–10 and P15–16. The recorded neurons were filled with biocytin for post-hoc identification and reconstruction. To differentiate between recorded D1-MSNs and D2-MSNs in D1-GFP mice, we used the transcription factor Ebf1, specifically expressed by matrix D1-MSNs (Tinterri et al., 2018), and preproenkephalin (PPE) expressed by D2-MSNs (Lee et al., 1997; Krajeski et al., 2019) (Fig. 2A). As Ebf1 expression declines rapidly after P5 (Tinterri et al., 2018), we used PPE to identify D2-MSNs beyond P7 (Fig. 2A). In parallel, D1-MSNs and D2-MSNs in D2-GFP mice were identified based on GFP expression. Ctip2 staining confirmed that all recorded neurons were MSNs and not striatal interneurons. As no significant differences were found between D1-MSNs in D1- or D2-GFP mice, their data were pooled, as were those for D2-MSNs from both types of mice.

Both D1 and D2-MSNs from both genotypes exhibited reductions in excitability between P0 and P15 (Fig. 2, Fig. S3). Specifically, higher

current intensities were required to elicit spiking responses (Fig. 2B-C). Accordingly, input resistance and resting membrane potential decreased over this period (Fig. 2D-E), while rheobase and membrane capacitance increased (Fig. 2F-G) without statistical significance between both genotypes. While only 87 % of D2-MSNs from WT mice were able to fire upon current injection, and exhibited an immature pattern being unable to discharge at high frequencies, all D2-MSNs from R6/1 at P0–2 could generate high-frequency action potentials in response to high currents intensities (#, $F_{(10, 499)} = 6.182$; $P < 0.0001$; mixed-effects analysis; Fig. 2C). Additionally, D2-MSNs from R6/1 mice displayed a significantly more hyperpolarized membrane potential and higher membrane capacitance compared to WT mice (Em: $p = 0.0221$, Student's t-test; Fig. 2E; Cm: $p = 0.0055$, Student's t-test; Fig. 2G). Together, these data showed that at P0–2, D2-MSNs from R6/1 exhibited early signs of maturation. These differences were specific to the P0–2-time window, as no significant differences in discharge frequency or membrane parameters were observed from P3 onwards.

In adult MSNs, intrinsic excitability is regulated by various potassium channels, particularly inwardly rectifying potassium channels (Kir), which maintain the resting membrane potential toward the potassium equilibrium potential, contributing to the highly hyperpolarized resting membrane potential characteristic of MSNs (Nisenbaum and Wilson, 1995; Shen et al., 2007). As described in the literature, by injecting hyperpolarizing currents, we observed the progressive establishment of the Kir currents in WT mice (Fig. 2H)(Dehorter, 2011). During the first postnatal days, the potential/injected current curve was linear, reflecting the absence of Kir channels (Fig. 2I). As the mice aged, membrane potential hyperpolarization decreased with the increased in Kir channels expression. Interestingly, D2-MSNs from R6/1 mice exhibited significantly altered potential/injected current curve at P0–2, with reduced hyperpolarization responses to strong negative currents (#, $F_{(6, 321)} = 4.043$; $p = 0.0006$; mixed-effects analysis; Fig. 2I), suggesting early Kir channels expression. No significant differences were observed between genotypes from P3 onwards (Fig. 2I). These data support the conclusion that D2-MSNs from R6/1 mice exhibited abnormal early maturation of electrophysiological properties during the first postnatal days (P0–2), with these alterations normalizing within a few days. In contrast, when we compared the electrophysiological properties of D1-MSNs between the two genotypes no difference was observed (Fig. S3), suggesting a cell-type selective alteration restricted to D2-MSNs.

To further confirm these findings, we conducted the same electrophysiological characterization in the CAG140 mouse model of HD that is genetically distinct from R6/1 (Fig. S4). Similar to R6/1 mice, D2-MSNs from CAG140 mice displayed an early maturation at P0–2, including heightened action potential firing (Fig. S4B, C) and a non-linear response to hyperpolarizing current (Fig. S4H, I). Additionally, these alterations persisted longer in the CAG140 model, as changes in input resistance (Fig. S4D) and rheobase (Fig. S4E) were also observed at P3–6 before normalizing by P7–10.

### 2.3. Compartment-specific electrophysiological alterations in HD D2-MSNs

Having already observed global electrophysiological alterations in D2-MSNs from R6/1 mice at P0–2, we next sought to determine whether these changes were compartment-specific by retrospectively classifying the previously recorded neurons into striosomal and matrix neurons. To do this, we used the D1-GFP line where $GFP^+$ MSNs are clustered in striosomes (See Fig. S2). We were able to differentiate $GFP^-$ neurons located within $GFP^+$ clusters (identified as striosomal D2-MSNs) from those outside these areas (classified as matrix D2-MSNs). Striosomal neurons were further confirmed by their localization in DARPP-32-enriched regions, while matrix D2-MSNs were identified using Ebf1 and DARPP-32 markers (Fig. 3A, B).

Strikingly, no significant alterations were observed in the striosomal
<figure><img src="image_5.png" /><figcaption>**Fig. 2.** Maturation of the electrophysiological properties of D2-MSNs is impaired in R6/1 mice during the first postnatal days. A. Examples of recorded biocytin-filled neurons at 4 postnatal stages: P0–2, P3–6, P7–10 and P15–16. By performing immunohistochemistry against the marker for MSNs, Ctip2, and the specific markers for either D1-MSNs, Ebf1, or D2-MSNs, PPE, we were able to confirm the D2 nature of the recorded neurons. Recorded neurons are pointed out by an asterisk. B. Representative responses of D2-MSNs in WT mice (light blue) or R6/1 mice (dark blue) following injection of a depolarizing current. C. Mean discharge frequency/injected current curves obtained for each age. D-G. Main electrophysiological properties of D2-MSNs with membrane input resistance (Rin, D), resting membrane potential (Em, E), rheobase (F) and capacitance (Cm, G) plotted longitudinally. H. Representative responses of D2-MSNs in WT mice (light blue) or R6/1 mice (dark blue) following injection of a hyperpolarizing current. I. Potential/current curves obtained for each age. Scale bars: 25 and 10 µm. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>
<figure><img src="image_6.png" /><figcaption>**Fig. 3.** Electrophysiological properties of D2-MSNs from R6/1 mice are altered in a compartment-dependent manner during the first postnatal days. A-B. Examples of D2-MSNs neurons recorded in either striosomes (A; GFP-/DARPP-32+ domain) or matrix (B; GFP-/Ebf1-/DARPP-32- domain). C, F. Representative D2-MSNs responses from striosome (C) and matrix (F) in WT (light blue) and R6/1 (dark blue) mice following the injection of hyperpolarizing and depolarizing currents. D, G. Mean discharge frequency/injected current curves obtained for each condition. E, H. Membrane potential/current curves obtained for each condition. I-L. Main electrophysiological properties of D2-MSNs showing membrane input resistance (Rin, I), capacitance (Cm, J), rheobase (K) and resting membrane potential (Em, L) as a function of the striatal compartment. Abbreviations: Str = striosomes. Scale bars: 25 and 10 µm. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>

D2-MSNs of WT and R6/1 mice. Both evoked discharge frequency and the potential/injected current curve were comparable to WT mice (Fig. 3D, E). Similarly, none of the membrane parameters showed significant differences between the two genotypes (Fig. 3I-L).

In contrast, the previously described alterations were restricted to matrix D2-MSNs. Specifically, matrix neurons from R6/1 mice exhibited a higher evoked discharge frequency and reduced hyperpolarization in response to injected currents, along with a more hyperpolarized resting membrane potential compared to WT mice (Fig. 3F-H; **3 L**). Taken together, these findings further support the notion of an early maturation restricted to D2-MSNs of the matrix compartment.

We performed the same analysis for D1-MSNs and found no differences in excitability between WT and R6/1 mice in either striosomal or matrix compartments (Fig. S5). In summary, the early maturation defects observed in the electrophysiological properties of D2-MSNs from R6/1 mice at P0–2 are not only cell-specific but also compartment-specific, confined to the matrix, with these alterations normalizing by the end of the first postnatal week.

## 2.4. Dendritic maturation is faster in HD D2-MSNs

The postnatal period is also characterized by the progressive development of MSN morphology, with neurons extending their dendritic arbors and progressively invading the striatum (Krajeski et al., 2019). To examine the impact of Htt mutation on this maturation process, we performed three-dimensional reconstructions of biocytin-filled recorded neurons and traced their dendritic arborization (Fig. 4).

As expected, D2-MSNs from WT mice displayed significant dendritic growth, with arbors becoming longer and more elaborate over time, as evidenced by increases in both total dendritic length and the number of nodes (Fig. 4A, C, E). Scholl analysis further revealed a progressive rightward shift in the curves with age, indicating that dendrites extended into a broader area of the striatum. In contrast, although dendritic length in D2-MSNs from R6/1 mice did not differ significantly from that of WT mice at any developmental stage, abnormalities in dendritic complexity were evident at P0–2. Scholl analysis revealed a significant increase in the number of intersections in R6/1 mice compared to WT mice (# F(29, 580) = 2.257; *p* = 0.0002; repeated
<figure><img src="image_7.png" /><figcaption>**Fig. 4.** Maturation of the morphological properties of D2-MSNs is impaired in R6/1 mice during the first postnatal days. A. Examples of biocytin-filled D2-MSNs at different postnatal stages. B. Scholl analysis illustrating the number of intersections between the dendritic tree and concentric circles spaced 5 µm apart from the soma. C-E. Main features of the dendritic arborization with dendritic length (C), number of primary dendrites (D) and number of nodes (E). Scale bars: 50 µm.</figcaption></figure>

measures ANOVA; Fig. 4B). Similarly, there was a notable increase in dendritic nodes in R6/1 mice at P0–2 (Fig. 4E). These findings suggest that dendritic maturation in D2-MSNs from R6/1 mice is accelerated, with an early increase in branching that normalizes from P3 onward. Interestingly, this phenotype was specific to D2-MSNs, as D1-MSNs did not show any significant differences between WT and R6/1 mice (Fig. S6). Altogether, these observations reveal a cell-type-specific alteration in the morphological maturation of D2-MSNs in R6/1 mice, reinforcing the idea of that striatal circuit mature more rapidly in HD mice.

## 2.5. Cortico-striatal synaptic transmission is decreased in HD D2-MSNs

Synaptic excitatory afferents are rapidly established after birth and mature during the first postnatal weeks (Dehorter, 2011; Krajeski et al., 2019). To assess the maturation of glutamatergic transmission, we recorded spontaneous excitatory postsynaptic currents (sEPSCs) using whole-cell *ex vivo* electrophysiological recordings at P0–2, P3–6 and P7–10 (Fig. 5A). sEPSCs were recorded at –70 mV in the presence of GABA<sub>A</sub> and GABA<sub>B</sub> receptors antagonists to isolate excitatory currents and the glutamatergic nature of the recorded currents was confirmed by the addition of AMPA and NMDA receptor antagonists (not shown). In WT mice, the frequency of sEPSCs increased progressively with age (Fig. 5B), while the mean amplitude remained stable (Fig. 5C-D). No significant differences in EPSC frequency or amplitude were observed between genotypes from P0 to P6 (Fig. 5B-D). However, at P7–10, EPSC amplitudes were significantly reduced in R6/1 mice (*p* = 0.0299, Student's *t*-test; Fig. 5D).

Given that the cortico-striatal pathway represents the major excitatory input to the striatum and is particularly vulnerable in HD (Smith

et al., 2004; Voorn et al., 2004; Blumenstock and Dudanova, 2020), we questioned whether the alterations in spontaneous glutamatergic transmission could be attributed to changes in this pathway. To address this, we performed whole-cell patch-clamp recordings of D2-MSNs in the dorsal striatum and electrically stimulated cortical afferents *via* a bipolar electrode placed at the level of the corpus callosum (Fig. 5E). Paired-pulse ratio (PPR) analysis revealed no significant differences between genotypes, suggesting no changes in short-term glutamate release at cortico-striatal synapses (Fig. 5F-G). However, the NMDA/AMPA ratio was significantly increased in R6/1 mice (**p** = 0.0014, Mann-Whitney *U* test; Fig. 5H-I), driven by a selective reduction in AMPA currents (**p** = 0.0088, Mann-Whitney test; Fig. 5J), while NMDA responses remained unchanged (*p* = 0.9874, Student's *t*-test; Fig. 5K). No significant differences were observed in PPR, NMDA/AMPA ratio or receptor-specific currents in D1-MSNs between genotypes (Fig. S7). Together, these findings reveal a selective deficit in cortico-striatal transmission in D2-MSNs in R6/1 mice, characterized by reduced AMPA receptor-mediated currents during the second postnatal week.

## 2.6. The striato-pallidal projections are reduced in HD mice

Given that we observed alterations in dendritic arborization of D2-MSNs in HD mice, we next investigated whether their projections to the external globus pallidus (GPe) were also affected. To examine striato-pallidal projections, we performed whole brain clearing on D2-GFP mouse brains at postnatal days P0, P3, and P8 (Fig. 6A). This technique allowed us to quantify fluorescence intensity in the GPe as a proxy for axonal density (Fig. 6B). At P0, R6/1 mice exhibited a significant reduction in fluorescence intensity in the GPe/compared to WT mice (**p** = 0.0042, Student's *t*-test; Fig. 6D). This reduction was
<figure><img src="image_8.png" /><figcaption>**Fig. 5.** AMPA receptor mediated cortico- striatal transmission is reduced in D2-MSNs from R6/1 mice during the second postnatal week. A. Representative recordings of spontaneous excitatory postsynaptic currents (sEPSCs) imposed at a potential of –70 mV in WT (left, light blue) and R6/1 (right, dark blue) mice at P7–10. B. Mean frequency of sEPSCs as a function of age. C. Average traces of sEPSCs obtained for each condition. D. Mean amplitude of sEPSCs as a function of age. E. Schematic of the experiment showing the position of the stimulating electrode and the recording electrode. F. Representative recordings of evoked excitatory postsynaptic currents (eEPSCs) at a potential of –70 mV in WT (light blue) and R6/1 (dark blue) mice. G. Average paired pulse ratio (EPSC2/EPSC1). H. Representative recordings at an imposed potential of either –70 mV to record AMPA receptor-mediated currents or at +80 mV to record currents mediated by both AMPA and NMDA receptors. I. NMDA/AMPA ratio obtained by dividing the amplitude values of NMDA currents by those of AMPA currents. J-K. Amplitude values of AMPA (J) and NMDA (K) currents. All recordings were performed in presence of GABA<sub>A</sub> and GABA<sub>B</sub> receptors antagonists (5 µM GABAzine +1 µM CGP55845). (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>

specific to the GPe, as no differences were observed in the striatum or background fluorescence (Fig. S8). This defect was transient as the fluorescence intensity in the GPe normalized to WT levels by P3. Given that the number of cells was unchanged at P0, these findings suggest a transient reduction of striato-pallidal axonal projections in the GPe of HD mice. One hypothesis to explain the reduction in D2-MSN projections to the GPe is a direct impact of mHtt on axonal growth, as suggested by a recent study (Marquilly et al., 2021). Another possibility is a defect in BDNF expression levels, which have been reported to be reduced during the first post-natal weeks in the HdhQ-250 mouse model of HD (Jin et al., 2015).

2.7. *The locomotor activity is not altered in HD mice*

To further investigate whether these developmental alterations could influence locomotor activity, we examined movement patterns during the early postnatal period. Locomotion in mice is known to progress rapidly during the first postnatal week, beginning with minimal movement in the first few days, followed by a gradual transition from crawling to walking (Dehorter, 2011). To assess this progression, we performed an open field test, in which pups were placed in an arena for one minute (Fig. 7A). Their movements were tracked to measure both the total distance traveled and the distance covered in a straight line. No significant differences were found between WT and R6/1 mice in either the total distance traveled or the distance covered in a straight line
<figure><img src="image_9.png" /><figcaption>**Fig. 6.** Striato-pallidal projections are reduced in R6/1 mice. A. Whole-brain fluorescent images showing D2-MSNs (labeled by GFP) and their projections to the GPe, acquired from clarified brains at P0 and P8. B. Representative fluorescent images of the striatum and striato-pallidal projections to the GPe, based on a 300 µm projection, at P0 and P8 for WT and R6/1 mice. C. Graph depicting the evolution of GPe volume between P0 and P8 in WT and R6/1 mice. D. Quantification of mean fluorescence intensity per µm³ for striato-pallidal axons in the GPe at P0, P3 and P8. N = 3 animals per genotype. Scale bars: 1 mm (A) and 300 µm (B). Abbreviations: STR = striatum, GPe = external Globus pallidus.</figcaption></figure>

<figure><img src="image_10.png" /><figcaption>**Fig. 7.** Locomotor activity is normal in R6/1 mice. A. Photographs of representative postnatal mice at P1 and P9. Body motion of the pups in the open field during a 1-min test. B. Quantification of total distance traveled and distance traveled in straight line segments (cm). C. Mean righting reflex times measured at P1, P5 and P9. Data represent three litters with N = 15 animals per genotype.</figcaption></figure>

(Figs. 7B).

We also evaluated the righting reflex, a behavioral test that relies on vestibular function, proprioception, tactile feedback, and motor

coordination (Braz et al., 2022). Mice were placed on their backs, and the time taken to roll over and bring all four paws onto the surface was recorded (Fig. 7C). Righting times were comparable between WT and
R6/1 mice, indicating no significant differences between the genotypes (Fig. 7C). These results suggest that locomotor activity is not impaired in the HD model, at least during early postnatal development.

## 3. Discussion

HD progression and symptoms are recapitulated in transgenic mouse models. We utilized the widely studied R6/1 model due to its adult-onset and slow disease progression (Pouladi et al., 2013). Our findings reveal that mHtt disrupts the development of D2-MSNs by transiently altering their excitability, morphology, and cortico-striatal glutamatergic transmission during a critical postnatal window. Altered D2-MSN excitability is also observed in CAG140 newborn mice. These disruptions occur during a sensitive period when early neuronal activity is essential for shaping functional adult circuits. Both increases and decreases in neuronal activity during these critical windows can significantly alter connectivity and maturation (Bitzenhofer et al., 2021; Braz et al., 2022). This principle has been demonstrated across multiple neuro-developmental disorders, including fragile X syndrome, schizophrenia, and autism spectrum disorders, where early disruptions in activity are linked to long-term deficits in circuit functionality (Kirischuk et al., 2017; He et al., 2019; Ahmed et al., 2023). In the context of HD, the early alterations we observed in D2-MSNs suggest that mHtt interferes with activity-dependent processes critical for cortico-striatal circuit formation. These disruptions could impair the integration of D2-MSNs into the striatal network and compromise the establishment of functional cortico-striatal circuits in adulthood, potentially contributing to the progressive motor and cognitive deficits characteristic of HD. Matrix and striosome compartments are anatomically distinguishable at birth in mice, but they would undergo normal synaptic refinement until P7 in WT animals (Kokinovic et al., 2024; Ferrari et al., 2012). Given this dynamic developmental window, the loss of D2-MSN matrix identity could be attributed to intrinsic cellular alterations, disruptions in developmental processes, or both.

### 3.1. mHtt impacts the electrophysiological and morphological properties of matrix D2-MSNs

The postnatal period is marked by the progressive maturation of the electrophysiological properties of MSNs, as they acquire characteristics like those of adult neurons (Tepper and Trent, 1993; Dehorter, 2011; Krajeski et al., 2019). Comparing this maturation between WT and R6/1 mice, we observed that D2-MSNs showed abnormal hyperpolarization and increased capacitance during the first postnatal days, reflecting a decrease in excitability. Moreover, an inward rectifying current seems to be already present in these neurons, suggesting an earlier acquisition of the Kir channels involved in establishing the highly hyperpolarized resting potential of MSNs (Kita et al., 1984; Nisenbaum et al., 1996). Altered electrophysiological properties have also been observed in CAG140 neonates, including increased rheobase and altered driven activity. The CAG140 mouse model was developed in a different genetic context compared to the R6/1 model (Pouladi et al., 2013), suggesting that postnatal alterations in D2-MSNs are associated with the mutated huntingtin itself, rather than the specific mouse model used. In addition to these intrinsic excitability deficits, morphological changes were observed in D2-MSNs, including increased dendritic complexity and reduced axonal projections. These alterations are transient and limited to the first postnatal days, suggesting the development of compensatory mechanisms that contribute to achieving an allostatic state. Interestingly, transient alterations in electrophysiological properties and morphology of pyramidal neurons have also been observed during the first weeks after birth in the cortex of HD mice (Braz et al., 2022; Capizzi et al., 2022).

It is remarkable that alterations are preferentially observed in D2-MSNs, both during development and in adulthood, even though mHtt is ubiquitously expressed in both MSN subpopulations (Reiner et al.,

1988; Albin et al., 1992; Deng et al., 2004). In adulthood, decreased BDNF release at cortico-striatal synapses and decreased TrkB receptor expression in striatal neurons are known to play a crucial role in HD-related neurodegeneration (Ginés et al., 2006; Saudou and Humbert, 2016). Interestingly, TrkB receptor directly influences potassium channel activation (Carrillo-Reid et al., 2019; Rogalski et al., 2000), and this phenomenon has been shown to be impaired in a mouse model of HD (Carrillo-Reid et al., 2019). Moreover, it has been observed that during development TrkB receptor expression is predominantly found in D2-MSNs, with 98% of D2-MSNs expressing TrkB at P10, compared with only 18% of D1-MSNs (Baydyuk et al., 2011; Baydyuk and Xu, 2014). Although it is not known whether TrkB receptor expression is reduced during early development in HD mutation carriers, its dysregulation could promote an early onset of potassium conductances, disrupting D2-MSN excitability during the postnatal period.

### 3.2. Loss of matrix D2-MSNs identity in HD

Our findings further reveal that developmental alterations in HD mice are specific to matrix D2-MSNs. It is known that striosomal and matrix neurons have different afferents and efferents and are thus involved in distinct functions (Flaherty and Graybiel, 1994; Crittenden and Graybiel, 2011; Friedman et al., 2015). In the context of HD, these compartments are differentially implicated in disease symptoms, with matrix neurons contributing to motor deficits and striosomal neurons to psychiatric disturbances (Morton et al., 1993; Matsushima et al., 2023).

Interestingly, we observed that matrix D2-MSNs in HD mice exhibit electrophysiological properties resembling those of striosomal neurons, suggesting a reduced distinction between these compartments during development. This observation aligns with findings in adult HD mice, where the electrophysiological properties of striosomal and matrix cells were reported to be less distinct (Matsushima et al., 2023). They also revealed a loss of compartment-specific molecular signatures, with striosomal markers being overexpressed in matrix neurons and vice versa (Matsushima et al., 2023). This loss of identity was particularly pronounced in D2-MSNs, aligning with their preferential vulnerability to mHtt. Although these transcriptional changes have been documented in adulthood, our findings suggest that this loss of striosome/matrix identity might originate during early postnatal stages.

Altered neuronal migration could also contribute to these developmental changes. It is known that mHtt affects neuronal migration in the cortex (Barnat et al., 2017), and similar mechanisms could be at play in the striatum. For example, striosomal neurons migrate earlier than matrix neurons (Mason et al., 2005; Liao et al., 2008), and premature migration of matrix D2-MSNs could result in properties resembling those of striosomal neurons. This convergence between striosome and matrix identities may have significant implications for corticostriatal connectivity and circuit function. Investigating the developmental trajectories of these subpopulations, particularly during embryonic stages, could clarify the origins of these changes and their contribution to HD pathology.

### 3.3. mHtt impacts cortical glutamatergic transmission on D2-MSNs

Although cortical and thalamic afferents are partially present from birth, cortico-striatal synaptogenesis continues throughout the postnatal period (Dehorter, 2011; Peixoto et al., 2016; Krajeski et al., 2019). We observed a decrease in the amplitude of spontaneous excitatory post-synaptic currents (sEPSCs) in D2-MSNs of R6/1 mice from P7. As the cortico-striatal pathway is particularly altered in adulthood (Blumenstock and Dudanova, 2020), we specifically studied the responses evoked by electrical stimulation of cortico-striatal axons at P7-10. We observed a significant increase in the NMDA/AMPA ratio in D2-MSNs. This change in ratio is caused by a decrease in the amplitude of the currents induced by AMPA receptors, without a change in the currents induced by NMDA receptors suggesting a postsynaptic
reduction in AMPA receptors. Interestingly, similar observations have been made in the developing cortex of HQ111 mice (Braz et al., 2022). Taken together, our results indicate that mHtt induces a postsynaptic decrease in cortico-striatal glutamatergic transmission in D2-MSNs of HD mice during the second postnatal week. Several mechanisms can contribute to the reduction in AMPA currents in HD mice, including alterations in AMPA receptor transport and mobility on the surface, a reduction in receptor expression or phosphorylation. It is already known that in adulthood, mHtt reduces glutamatergic transmission by impacting the trafficking and surface diffusion of these receptors (Mandal et al., 2011; Zhang et al., 2018). Interestingly, treating HD pups with AMPAkine, a positive allosteric modulator of AMPA receptors, has been shown to not only restore the observed alterations but also to delay the appearance of major HD symptoms in adulthood (Braz et al., 2022). This suggests a critical role of glutamatergic synaptic transmission in the early dysfunction of the corticostriatal microcircuit in HD.

In conclusion, our study highlights that mHtt disrupts critical developmental processes, including excitability, morphology, and synaptic transmission, in D2-MSNs during a sensitive postnatal period. Although these alterations are transient, they likely initiate compensatory mechanisms that, over time, may become insufficient to sustain proper circuit function. This early vulnerability could contribute to the progressive striatal dysfunction characteristic of HD. Understanding these developmental disruptions not only sheds light on the mechanisms underlying circuit integration in HD but also opens avenues for therapeutic interventions aimed at preserving striatal circuit integrity and delaying disease progression.

# 4. Experimental model and subject details

## 4.1. Animals

Experiments were performed in two mouse models of HD: a model expressing a truncated form of the human huntingtin gene, the R6/1 model, and a knock-in CAG140 model. R6/1::D1-GFP mice were generated by crossing heterozygous R6/1 male with homozygous D1-GFP females. Resultant WT::D1-GFP or R6/1::D1-GFP mice were obtained in half proportion. R6/1::D2-GFP or CAG140::D2-GFP mice were generated by crossing heterozygous R6/1- or CAG140 males with homozygous D2-GFP females. These mice lines reveal D1- or D2-MSNs by the presence of green fluorescent protein (GFP). All animals were maintained in a 12/12 h light/dark cycle, in stable conditions of temperature and humidity, with access to food and water *ad libitum*. We verified the CAG repeat number in our R6/1 and CAG 140 lineages. Our R6/1 lineage exhibited a reduction in the CAG repeat number compared to the norm, with an average of 55 repeats instead of the expected 115 (Mangiarini et al., 1996). This extremely rare contraction has been described in the literature and leads from 1 year of age to mHtt aggregates similar to that found in human HD (Morton et al., 2019). The extension repeat number in our CAG140 lineage was 109. All experiments were approved by the local ethical committees and by the French Ministry of Research and APAFIS#14255.

## 4.2. Tissue preparation and immunohistochemistry

Mice (0–15 days) were deeply anaesthetized with isoflurane and then decapitated. Brains were extracted and post-fixed overnight in a solution of 4 % w/v PFA in 0.1 M phosphate buffered saline, pH 7.4 (PBS) at 4C. Next day, brains were washed twice in PBS and then stored in PBS-azide 0.03 % before being cut into 50 µm coronal sections on a vibratome (VT1000S; Leica Microsystems). For immunohistochemistry, sections were washed 3 times in PBS for 10 min. Sections were then incubated in blocking solution containing 4 % Normal Donkey Serum (NDS) in PBS with 0.3 % Triton X-100 for 1 h at room temperature with constant shaking. Finally, sections were incubated in primary antibodies diluted in 4 % NDS in PBS with 0.3 % Triton X-100 solution overnight at

4 °C. The following antibodies were used: rat anti-Ctip2 (1:1000, Abcam ab18465) and chicken anti-GFP (1:1000, Aves Lab GFP1010). Ctip2 is expressed by all MSNs but not interneurons (Arlotta et al., 2008). Next day, sections were rinsed three times in PBS for 10 min before incubation for two hours at room temperature with secondary antibodies. The following fluorescent probes were used: donkey anti-rat Alexa Fluor 647 (1:500, Jackson Laboratories), donkey anti-chicken Alexa Fluor 488 (1:500, Jackson Laboratories). Sections were rinsed 3 times for 10 min in PBS before mounting in ProLong Diamond Antifade Mountant (Invitrogen).

## 4.3. Image acquisition, analysis and quantification

Fluorescence images were acquired on confocal microscope (Carl Zeiss© LSM 900 or Zeiss Celldiscoverer 7) using ZEN software. For cell density analysis, three rostro-caudal levels of the striatum were defined using the anterior commissure as an anatomical landmark. A tile scan of the whole striatum on a single plane was then acquired using a x40 or x20 objectives. For cell density analysis, we manually delineated each striatal area to calculate the specific area of each level. Neuronal counting was then performed semi-automatically using the QuPath software (Bankhead et al., 2017). To determine the density of D1-MSNs and D2-MSNs, the number of GFP- or GFP+ neurons, respectively, identified in each striatum was divided by the area of the striatum.

## 4.4. Acute brain slices preparation and recording conditions

Coronal striatal slices were prepared from 0 to 16-day-old R6/1, CAG140 or non-carrier control mice. Briefly, animals were deeply anaesthetized with isoflurane and decapitated. The brains were removed from the skull and placed in ice-cold modified ACSF (mACSF), saturated with 95 % O₂ and 5 % CO₂, and containing (in mM): 230 sucrose, 10 MgSO₄·7H₂O, 2.5 KCl, 1.25 NaH₂PO₄·H₂O, 0.5 CaCl₂·H₂O, 26 NaHCO₃, et 10 D-glucose. Brains were sectioned into 300 µm-thick coronal slices by using a vibratome (VT-1200S; Leica Microsystems, Germany). Slices were then immediately transferred in standard ACSF containing (in mM): 126 NaCl, 2.5 KCl, 1.25 NaH₂PO₄·H₂O, 2 CaCl₂·H₂O, 2 MgSO₄·7H₂O, 26 NaHCO₃, 10 D-glucose, 5 L-glutathion and 1 sodium pyruvate (gassed with 95 % O₂/5 % CO₂). Slices were incubated 1 h at 32 °C in this solution and then maintained at room temperature in the same solution until recording. Striatal slices were transferred to a recording chamber, continuously perfused with oxygenated ACSF containing (in mM): 126 NaCl, 3 KCl, 1.25 NaH₂PO₄·H₂O, 1.6 CaCl₂·H₂O, 1.5 MgSO₄·7H₂O, 26 NaHCO₃, and 10 D-glucose (at 32 °C with a perfusion speed of 2 ml.min⁻¹). Striatal neurons were visualized using infrared gradient contrast video microscopy (E600FN, Eclipse workstation, Nikon, Japan) and a water-immersion objective (Nikon Fluor 60 X/1.0 NA). MSNs were distinguished from interneurons by their morphology and GFP-expressing neurons were identified in real time under epifluorescence (Nikon Intensilight C-HGFI). Recordings from individual neurons were made using patch pipettes (5–8 MΩ) pulled from thick-walled borosilicate glass capillaries (G150–4; Warner Instruments, Hamden, CT, USA) on a micropipette puller (P-97, Sutter Instruments, Novato, CA, USA). For recordings of the intrinsic properties, the patch pipette internal solution contained (in mM): 135 K-gluconate, 3.8 NaCl, 1 MgCl₂·6H₂O, 10 HEPES, 0.1 Na₄EGTA, 0.4 Na₂GTP, 2 MgATP and 5.3 biocytin. In current-clamp mode, the junction potential (+13 mV; JPCalc, Clampex 10) was corrected. For recordings of cortico-striatal transmission, the patch pipette internal solution contained (in mM): 132 CsMSO₃, 3.6 NaGlu, 1 MgCl₂·6H₂O, 10 HEPES, 0.1 Na₄EGTA, 5 TEA–Cl, 0.4 Na₂GTP, 2 MgATP, 5 QX-314 and 5.3 biocytin. The osmolarity and pH of the intrapipette solutions were adjusted at 290 mOsm and 7.2, respectively. The recordings were amplified using a Multiclamp 700B amplifier (Molecular Devices) and digitized at 20 kHz (Digidata 1320 A/1550B) using Clampex 10 acquisition software (Molecular Devices, Sunnyvale, CA, USA).
## 4.5. Stimulation and recording protocols

In voltage-clamp mode, the holding potential was set at -60 mV and successive voltage steps of -5 mV were performed to measure passive properties of the cells including input resistance and capacitance. In current-clamp mode, hyperpolarizing and depolarizing current steps were injected through the patch pipette to assess the active properties of MSNs (rheobase and spiking frequency). Current step ranges were for P0-2: -30 to +50 pA; for P3-6: -50 to +50 pA; for P7-10: -50 to +80 pA and P15-16: -200 to +250 pA, 1000-ms duration. Spontaneous post-synaptic currents (EPSCs) were recorded in voltage-clamp mode at a fixed membrane potential of -70 mV during 5 and 10 min. For these experiments, recordings were conducted in the presence of the GABA<sub>A</sub> antagonist SR95531/GABAzine (5 µM, Tocris Bioscience) and the GABA<sub>B</sub> antagonist CGP55845 (1 µM, Tocris Bioscience). To confirm the glutamatergic nature of the measured currents, antagonists of AMPA receptors (DNQX, 20 µM, Abcam) and NMDA receptors (D-AP5, 50 µM, Abcam) were used in some experiments. All pharmacological agents used in this study were prepared as stock solutions, aliquoted in distilled water, and stored at -20 °C. They were then diluted to experimental concentrations on the day of the experiment and perfused into the recording chamber. Activation of cortical afferents was achieved using a bipolar stimulation electrode placed just above the corpus callosum (stimulation intensity between 100 and 800 µA) connected to a constant current isolator (A.M.P.I., Israel). These experiments were conducted in the presence of GABAergic transmission blockers (5 µM GABAZine and 1 µM CGP55845). Double stimulations of 0.1 ms at 5 and 20 Hz were performed to measure the amplitude of EPSCs and assess the short-term plasticity of the cortico-striatal synapse determined through the paired-pulse ratio (PPR = Pulse2/Pulse1). For these experiments, neurons were recorded at a potential of -70 mV. To determine the relative contribution of AMPA/Kainate and NMDA receptors in cortico-striatal transmission in MSNs, synaptic responses to single stimulation of cortico-striatal neurons were recorded at -70 mV and +80 mV, respectively.

## 4.6. Analysis

Data analyses were performed using the pClamp 10.6 (Molecular devices) and OriginPro 7.0 (Microcal Software) softwares. Series resistance (Rs), membrane resistance (Rm) and membrane capacitance (Cm) were calculated in whole-cell voltage-clamp configuration by analysis of the transient currents in response to a -5 mV step of potential (ΔV). Cells with Rs exceeding 30 MΩ were excluded from analysis. The Rm was measured using Ohm's law (U = R*I) based on the steady-state current at the end of the potential step. The Cm was calculated using the following formula: Cm = Q/ΔV in which Q correspond to the area under the capacitive transient after removing leak currents. In current-clamp mode, action potential firing frequency and membrane potential were calculated for each injected current and F-I curve (frequency of action potential firing as a function of injected current) and I-V curve (membrane potential as a function of injected current) were constructed. The rheobase was also determined with progressive current steps of 10 ms. Electrically-evoked EPSCs amplitude were measured as the peak of inward current relative to the baseline holding current preceding the electrical pulse. To obtain the paired pulse ratio, the amplitude of the second EPSC was divided by the amplitude of the first EPSC (EPSC<sub>2</sub>/EPSC<sub>1</sub>). The AMPA/NMDA ratio was calculated from recordings of the same neuron held at -70 mV to observe AMPA currents or at +80 mV to observe NMDA currents. The AMPA receptor-mediated current corresponds to the amplitude of the peak from neuron held at -70 mV. The NMDA receptor-mediated current corresponds to the amplitude of the current, from neuron held at +80 mV, 50 ms after the electrical stimulation. Peak amplitude of AMPA receptor-mediated current was then divided by peak amplitude of NMDA receptor-mediated current to obtain the AMPA/NMDA ratio.

## 4.7. Cell identification and cell reconstruction

Following whole-cell electrophysiology, the striatal slices were fixed in 4% paraformaldehyde in PBS. Subsequently, sections were processed essentially as described above using rat anti-Ctip2 and Chicken anti-GFP. In addition, the following antibodies were used: rabbit anti-Ebf1 (1:500, Millipore AB10523), rabbit anti PPE (1:2000, LsBio LS-C23084), Rabbit anti-MOR (1:1000, Immunostar 24,216), Mouse anti-DARPP-32 (1:500, BD Biosciences 611,520). Ebf1 and PPE antibodies staining were improved through antigen retrieval by heating sections at respectively 65 °C for 1 h and 80 °C for 35 min in 10 mM sodium citrate (Target Retrieval Solution 10×, S1699; Aligent Technologies, Santa Clara (US); pH 6.1). Ebf1 and PPE are specific markers of respectively D1-MSNs (Garel et al., 1997) and D2-MSNs (Lee et al., 1997; Sharott et al., 2017; Krajeski et al., 2019). As Ebf1 is a transcription factor that progressively disappears during the first postnatal week, Ebf1 labelling was used to identify D1-MSNs from P0 to P4. PPE labelling was used to identify D2-MSNs from P5. Biocytin was revealed with Streptavidin-AF568 (1:2000, Invitrogen S11226).

Following multiple labeling, fluorescence images were acquired using a confocal microscope (Carl Zeiss© LSM 900) with the ZEN software. To identify biocytin-filled neurons, z-stack acquisitions were performed with a 2 µm step size using a 40× oil immersion objective. Images of the different immunohistochemical and biocytin labels were then overlaid using ImageJ (NIH) to determine the nature of the recorded MSNs. Additionally, z-stack acquisitions with a 0.15-µm step size were obtained using a 40× objective to image the entire dendritic arborization of the neurons. Once imaged, neurons were then reconstructed and analyzed using Neurolucida and Neuroexplorer software (MBF Bioscience, Delft, The Netherlands). Only neurons presenting a complete dendritic tree were included in the analysis. Scholl analysis was generated by calculating the number of intersections between the dendritic tree of neurons and each concentric circle spaced every 5 µm from the soma. Dendritic length, number of nodes, and number of primary dendrites were also measured.

## 4.8. Brain clearing

Whole mouse brains (P0, P3, P8) were cleared following the AdipoClear+ protocol (Chi et al., 2018) with modifications to optimize GFP amplification and tissue transparency. Brains were fixed in 4% PFA at 4 °C overnight, rinsed in PBS (3 × 1 h), and stored in PBS containing 0.03% sodium azide until use. Next, the samples were subjected to a series of methanol dehydration steps: 20%, 40%, 60%, 80% methanol in H₂O/0.1% Triton X-100/2% glycine/0.01% NaOH, 0.04% sodium azide (B1N buffer, pH 7), followed by 100% methanol. Each step was performed for 1 h at room temperature under constant agitation. Delipidation was performed with 100% dichloromethane (DCM; 3 × 1 h) and 100% methanol (2 × 1 h). Samples were bleached in methanol/H₂O₂ (5:1, with 30% H₂O₂) for 4 h at room temperature, then rehydrated in a reverse methanol gradient: 80%, 60%, 40%, 20% methanol in B1N buffer, each for 1 h, followed by an overnight wash in B1N buffer. Permeabilization was achieved using PTxwH buffer (PBS buffer containing 0.1% Triton X-100, 0.05% Tween-20, 2 µg/ml heparin, and 0.02% sodium azide) supplemented with 2.25% glycine and 2.5% DMSO. Samples were incubated with chicken anti-GFP (1:2000, AVES Labs) in PTxwH for 7 days at 37 °C with agitation, washed 12 times in PTxwH at 37 °C, for 1 h minimum, over a span of 3 days, and incubated with goat anti-chicken Alexa Fluor 647 (1:1000, Thermo Fisher) in PTxwH for 7 days. Dehydration was performed using a methanol gradient (20%, 40%, 60%, 80%, and 100%; 1 h each), followed by DCM/MeOH (2:1) overnight, and 100% DCM (2 × 1 h). Final clearing was achieved in dibenzyl ether (DBE; Sigma-Aldrich) for 4 h, and samples were stored at 4 °C in DBE until imaging. Samples were imaged using an UltraMicroscope II (Miltenyi Biotech) with a 12.6× magnification. 3D reconstructions and quantifications were performed with
Imaris (Bitplane). The external globus pallidus (GPe) was manually delineated, and the mean gray value was measured within this region. As a control, the mean gray value was also measured in the striatum, and noise from endogenous fluorescence was assessed for each brain and under all experimental conditions.

## 4.9. Motor behavior

Locomotor activity was assessed in R6/1 and WT mice from three different litters between P1 and P10. Each pup was tested once per day for 1 min in an open field test. The pups were placed in a transparent open field arena (18 cm × 12 cm). A camera positioned below the arena recorded the pup's trajectory. The acquired data were analyzed using DeepLabCut. The total distance traveled and the total distance covered along straight-line segments were derived from this trajectory.

## 4.10. Statistics

Statistical analysis was conducted using Prism 8 (GraphPad Software, USA). Data in the text are presented as mean ± standard error of the mean (SEM). For comparisons between two independent groups, an unpaired Student's *t*-test was performed if the data followed a normal distribution. Otherwise, a non-parametric Mann-Whitney test was used. For comparison of groups or F-I response curves, statistical analysis was conducted using two-way analysis of variance (ANOVA), followed by Sidak's multiple comparisons *post hoc* test. Data were considered significant for *p* < 0.05.

## CRediT authorship contribution statement

**Margaux Lebouc:** Writing – original draft, Methodology, Formal analysis, Data curation, Conceptualization. **Léa Bonamy:** Methodology, Formal analysis, Data curation. **Thibault Dhellemmes:** Methodology, Formal analysis, Data curation. **Jakob Scharnholz:** Formal analysis, Data curation. **Quentin Richard:** Formal analysis, Data curation. **Gilles Courtand:** Methodology, Formal analysis. **Alexandre Brochard:** Formal analysis, Data curation. **Frédéric Martins:** Formal analysis, Data curation. **Marc Landry:** Supervision, Methodology, Formal analysis. **Maurice Garret:** Writing – review & editing, Writing – original draft, Supervision, Methodology, Formal analysis, Data curation. **Jérôme Baufreton:** Conceptualization, Writing – review & editing, Writing – original draft, Supervision, Methodology, Formal analysis, Data curation.

## Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

## Acknowledgments

M.L. received a Ph. D fellowship from the Association Huntington France. This work was supported by recurrent funding from the University of Bordeaux and the CNRS. This study also received financial support from the French government in the framework of the University of Bordeaux's IdEx "Investments for the Future" program / GPR BRAIN_2030. We are also grateful to Dr. Sonia Garel for providing D2-GFP mice and to Dr. Sandrine Humbert, Dr. Emilie Pacary, and Dr. Nathalie Dehorter for helpful discussion. We also wish to thank Jeremie Teillon, who initiated the implementation of clearing approaches at the Bordeaux Imaging Center, facilitating its application in this study.

## Appendix A. Supplementary data

Supplementary data to this article can be found online at https://doi.

org/10.1016/j.nbd.2025.106874.

## Data availability

Data will be made available on request.

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