Molecularly distinct striatonigral neuron subtypes differentially regulate locomotion

Dong, Jie; Wang, Lupeng; Sullivan, Breanna T.; Sun, Lixin; Martinez Smith, Victor M.; Chang, Lisa; Ding, Jinhui; Le, Weidong; Gerfen, Charles R.; Cai, Huaibin

doi:10.1038/s41467-025-58007-x

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Published: 19 March 2025

Molecularly distinct striatonigral neuron subtypes differentially regulate locomotion

Nature Communications volume 16, Article number: 2710 (2025) Cite this article

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Abstract

Striatonigral neurons, traditionally known for promoting locomotion, comprise diverse subtypes with distinct transcriptomic profiles. However, their specific contributions to locomotor regulation remain incompletely understood. Using the genetic markers Kremen1 and Calb1, we demonstrate in mouse models that Kremen1⁺ and Calb1⁺ striatonigral neurons exerted opposing effects on locomotion. Kremen1⁺ neurons displayed delayed activation at locomotion onset but exhibited increasing activity during locomotion offset. In contrast, Calb1⁺ neurons showed early activation at locomotion onset and decreasing activity during locomotion offset. Optogenetic activation of Kremen1⁺ neurons suppressed ongoing locomotion, whereas activation of Calb1⁺ neurons promoted locomotion. Activation of Kremen1⁺ neurons induced a greater reduction in dopamine release than Calb1⁺ neurons, followed by a post-stimulation rebound. Conversely, activation of Calb1⁺ neurons triggered an initial increase in dopamine release. Furthermore, genetic knockdown of GABA-B receptor Gabbr1 in Aldh1a1⁺ nigrostriatal dopaminergic neurons (DANs) reduced DAN inhibition and completely abolished the locomotion-suppressing effect of Kremen1⁺ neurons. Together, these findings reveal a cell type-specific mechanism within striatonigral neuron subtypes: Calb1⁺ neurons promote locomotion, while Kremen1⁺ neurons terminate ongoing movement by inhibiting Aldh1a1⁺ DAN activity via GABBR1 receptors.

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Introduction

Striatal spiny projection neurons (SPNs) constitute approximately 95% of the neuronal population in the dorsal striatum and play a key role in motor learning, decision-making, and regulating voluntary movement^1,2. These SPNs can be broadly classified into two main subtypes: dopamine receptor D1 (Drd1)-expressing direct-pathway neurons (dSPNs), which project directly to the globus pallidus internus (GPi) and substantia nigra pars reticulata (SNr); and dopamine receptor D2 (Drd2)-expressing indirect-pathway SPNs (iSPNs), which project to the globus pallidus externus (GPe)^1,2. The dSPNs primarily project to the SNr and are also referred to as striatonigral neurons. It is well recognized that the dSPNs facilitate movement initiation and the iSPNs restrain unwanted movements².

SPNs can be further subdivided into two complementary compartments within the dorsal striatum, namely the patch (or striosome) and matrix compartments, which are characterized by distinct neurochemical markers and input-output connectivity^3,4,5,6. Each compartment contains both dSPN and iSPNs. The patch compartment is typically marked by the expression of μ-opiate receptor (MOR1) in clusters of adjacent SPNs in rodents^4,7, whereas the matrix compartment is labeled by Calbindin (CALB1) expression⁸. Advances in single-cell transcriptomics have identified additional genetic markers for dSPNs, iSPNs, and patch- and matrix-associated SPNs^9,10. Interesting, although patch markers are predominantly expressed in the patch compartment, some are also present among individual or small clusters of SPNs, referred to as “exopatch” neurons, scattered throughout the matrix compartment^10,11. Furthermore, any given patch marker typically label only a subset of patch-associated SPNs¹². Previous research has underscored the significance of patch SPNs in modulating mood, decision-making, and reward processing, while matrix SPNs are implicated in action selection¹³. However, the specific roles of molecularly defined patch-associated dSPNs in locomotion control remain poorly understood.

Both patch and matrix dSPNs innervate aldehyde dehydrogenase 1a1-positive (Aldh1a1⁺) DANs located in the ventral tier of substantia nigra pars compacta (SNc)¹⁴, which are particularly vulnerable to degeneration in Parkinson’s disease¹⁵. Notably, a portion of patch-associated dSPN axon terminals bundle up to form distinctive striosome-dendron bouquet structures, intertwining with the dendrites of Aldh1a1⁺ DANs that extend into the SNr^16,17. This anatomical arrangement likely underlies the more potent presynaptic inhibition of DANs by patch-associated dSPNs in contrast to other inhibitory inputs¹⁸. Moreover, patch-associated dSPNs exert a prolonged inhibitory influence on DAN neuronal activity via GABA-B receptors, crucially regulating the transition from tonic to burst firing in Aldh1a1⁺ nigrostriatal DANs, as evidenced by brain slice recordings^16,19. Despite these insights, the physiological and behavioral implications of this distinctive striatonigral circuit remain to be explored.

One major challenge to study patch-associated SPNs is the difficulty in definitively characterizing and manipulating these neurons, largely due to their less well-defined neurochemical organizations. In this study, we initially identified the Kremen1 gene as a specific molecular maker for patch-associated SPNs in the dorsal striatum. Subsequently, we generated a line of Kremen1^2A-Cre knock-in (KI) mice to investigate the anatomy of Kremen1-positive (Kremen1⁺) dSPN subpopulations and their functional significance in locomotor control compared to the Calb1-positive (Calb1⁺) matrix dSPNs.

Results

Kremen1 transcript is enriched in patch compartments in the dorsal striatum

Using published patch reporter Nr4a1-eGFP transgenic mice²⁰, we isolated the eGFP-containing and the adjacent tissues in the dorsal striatum by laser capture microdissection for bulk RNA sequencing NCBI accession: PRJNA870469. In comparison to various patch markers examined previously, such as Oprm1, Tshz1, Pdyn, Nr4a1, and Sepw1, the expression of Kremen1 mRNA exhibited substantially higher differentials in the patch versus matrix compartments (Fig. 1a, b). Consistently, Kremen1 was also identified as a patch marker by an independent single-nuclei transcriptomic study¹⁰. Further co-localization analyses with Drd1 and Drd2 using RNAscope in situ hybridization found that approximately 60% of Kremen1⁺ SPNs corresponded to dSPNs, while 35% were identified as iSPNs, resembling the ratio of total dSPNs versus iSPNs in the striatum (Fig. 1c–e). Moreover, Kremen1⁺ SPNs were predominantly distributed in the dorsal striatum (Fig. 1f). Within the striatum, Kremen1⁺ SPNs constituted 12.7% of total SPNs, 52% of SPNs within the patch compartments (defined by clusters of five or more adjacent Kremen1⁺ SPNs), and 8.2% of SPNs in the matrix compartments (Fig. 1g). Notably, the density of Kremen1⁺ SPNs was substantially higher in patch compartments compared to matrix compartments (Fig. 1h), although the total number of Kremen1⁺ SPNs in the patch compartments was slightly lower than that in the matrix compartments (Fig. 1i). These findings highlight Kremen1 as a useful genetic marker for identifying a distinct subpopulation of SPNs, with a subset forming patch-like clusters specifically in the dorsal striatum.

Fig. 1: Distribution of Kremen1+ SPNs in patch and matrix compartments. — **Fig. 1: Distribution of *Kremen1*⁺ SPNs in patch and matrix compartments.**

Tshz1 is also enriched in patch compartments^9,10. We further compared the distribution of Kremen1⁺ and Tshz1⁺ SPNs, especially dSPNs in the dorsal striatum, by RNAScope in situ hybridization. We found that Kremen1⁺ SPNs has a larger distribution in the patch than Tshz1⁺ SPNs (Supplementary Fig. 1a, b). There is only around 15-20% overlap between these two dSPN subpopulations (Supplementary Fig. 1c, d). These results suggest that Kremen1⁺ and Tshz1⁺ mark two largely different subtypes of patch SPNs in the dorsal striatum. Together, they accounted for approximately 70% of patch-associated SPNs (Supplementary Fig. 1e).

Kremen1 ^2A-Cre knock-in mice are useful for studying patch-associated SPNs

Given the enrichment of Kremen1 in patch SPNs, we generated a line of Kremen1^2A-Cre knock-in (KI) mice using CRISPR/Cas9-mediated gene editing (Supplementary Fig. 2). By crossing these mice with Ai14 reporter mice²¹, which express the red fluorescent protein tdTomato under Cre-dependent regulation, we observed distinct tdTomato signals scattered within the dorsal but not ventral striatum of Kremen1^2A-Cre;Ai14 mice (Fig. 2a). These signals co-localized with the widely used patch marker MOR1⁴ (Fig. 2a, b). Consistent with the RNAscope analyses (Fig. 1i), the number of tdTomato⁺ SPNs is comparable in the patch and matrix compartments, with the patch identified by MOR1 immunostaining (Supplementary Fig. 3). Apart from SPNs, tdTomato signals were also detected in pericytes, with no staining detected in the interneurons or glial cells of the dorsal striatum (Supplementary Fig. 4a, b). The Kremen1⁺ SPNs were restricted to the dorsal striatum but distributed throughout the entire dorsal striatum along the rostral to caudal and medial to lateral axes (Supplementary Fig. 5a, b). Beyond the dorsal striatum, tdTomato signals were also visible in the hippocampal regions (Fig. 2c, Supplementary Fig. 5b). Notably, the incoming tdTomato-positive dSPN axons exhibited distinct dendron-bouquet like structures¹⁷, forming connections with the dendrites of DANs in SN (Fig. 2d).

Fig. 2: Distinct distribution patterns of Kremen1+ SPNs in the dorsal striatum and their projections. — **Fig. 2: Distinct distribution patterns of *Kremen1*⁺ SPNs in the dorsal striatum and their projections.**

To investigate the projection pattern of Kremen1⁺ SPNs, we performed stereotaxic injections of adeno-associated viral (AAV) vectors co-expressing Cre-dependent tdTomato and synaptophysin-fused EGFP (sypEGFP) into the dorsal striatum of Kremen1^2A-Cre KI mice (Fig. 2e). Both tdTomato and sypEGFP signals were found within patch-like compartments in dorsal striatum (Fig. 2f). Additionally, sypEGFP specifically labeled the axon terminals of Kremen1⁺ SPNs in the GPe and entopeduncular nucleus (EPN), the mouse equivalent to GPi (Fig. 2g), as well as in the SNr and SNc (Fig. 2h). When similar AAV injections were performed in Calb1^IRES2-Cre mice, tdTomato and sypEGFP signals were observe in the matrix compartment, characterized by low MOR1 expression (Supplementary Fig. 6a, b). The axon terminals of Calb1⁺ SPNs were found projecting to the GPe, EPN, and SN (Supplementary Fig. 6c, d). In the SNr, a clear overlap was observed between the axon terminals of Calb1⁺ dSPNs and the dendrites of TH⁺ DANs (Supplementary Fig. 6d, e). Therefore, Kremen1^2A-Cre and Calb1^IRES2-Cre KI mice provide useful tools for studying the anatomy and functional roles of patch- or matrix-associated SPNs, respectively.

Distinct activity changes in Kremen1 ⁺ and Calb1 ⁺ dSPNs during locomotion onset and offset

To investigate the activity patterns of Kremen1⁺ and Calb1⁺ dSPNs during self-paced locomotion, we injected AAVs expressing the genetically encoded calcium indicator GCaMP8s (AAV1-FLEX-GCaMP8s) into the dorsal striatum and implanted an optic fiber in the SNr of the same hemisphere in Kremen1^2A-Cre or Calb1^IRES2-Cre mice²¹ (Fig. 3a, b). The mice were head-fixed and allowed to walk freely on a belt treadmill, where movement velocity signal was synchronized with a fiber photometry setup for recording calcium transients in the axon terminals of Kremen1⁺ or Calb1⁺ SPNs (Fig. 3c–e, Supplementary Fig. 7). Representative sample traces revealed that calcium transients in the axon terminals of both Kremen1⁺ and Calb1⁺ dSPNs correlated with velocity changes during locomotion bouts (Fig. 3f, g). However, distinct differences emerged when aligning their mean activity to locomotion onset and offset (Fig. 3h, i). At locomotion onset, Kremen1⁺ dSPNs reached their peak activity later compared to Calb1⁺ dSPNs (Fig. 3j). Notably, Calb1⁺ dSPN activity began to rise prior to locomotion onset, whereas Kremen1⁺ dSPN activity increased much later and mostly after locomotion onset (Fig. 3k). At locomotion offset, the peak activity timing of Kremen1⁺ dSPNs was delayed compared to Calb1⁺ dSPNs (Fig. 3l). Furthermore, the slope of activity preceding locomotion offset differed markedly between the two subtypes: Kremen1⁺ dSPNs exhibited an increasing trend (positive slope), while Calb1⁺ dSPNs displayed a decreasing trend (negative slope) (Fig. 3m). These distinct activity patterns highlight their differential roles in regulating motor activity, with Kremen1⁺ dSPNs potentially facilitating locomotion termination and Calb1⁺ dSPNs contributing to movement initiation. When comparing the locomotion speed of individual mice, the mean velocity at locomotion onset was slightly higher in Calb1^IRES2-Cre mice than in Kremen1^2A-Cre mice (Supplementary Fig. 8a–c). However, no significant differences in velocity were observed at locomotion offset, nor were there differences in the shape of velocity changes between the two groups at either locomotion onset or offset (Supplementary Fig. 8a, b, d). Nonetheless, we do not believe that mouse strain differences account for the distinct dynamic changes in Kremen1⁺ and Calb1⁺ dSPN activity during locomotion onset and offset.

Fig. 3: Activity of Kremen1+ and Calb1+ dSPNs during self-paced locomotion. — **Fig. 3: Activity of *Kremen1*⁺ and *Calb1*⁺ dSPNs during self-paced locomotion.**

Kremen1 ⁺ and Calb1 ⁺ dSPNs exert contrasting roles in locomotor control

To examine the causal influence of Kremen1⁺ dSPN activity on locomotion, we used optogenetic manipulations by introducing AAV vectors containing Cre-dependent optogenetic activator Channelrhodopsin-2 (AAV1-FLEX-ChR2) into the dorsal striatum of Kremen1^2A-Cre mice and implanted the optic fiber in the SN for light stimulation of Kremen1⁺ dSPN axon terminals (Fig. 4a, b). Optogenetic activation at 3 mW/20 Hz for 3 min led to a notable reduction in locomotion velocity, followed by a subsequent rebound in motor activity after the stimulation period (Fig. 4c, d). The decrease in walking speed likely resulted from both a reduction in velocity during each bout of ambulatory movement (Fig. 4e) and reduced locomotor activity, as evidenced by reduced movement frequency and shortened duration (Fig. 4f, g), as well as an increase in immobility frequency (Fig. 4h). This reduction in locomotion velocity was also observed at various stimulation parameters, including continuous 0.25 mW power intensities, 10 Hz stimulation frequency, and 10 s durations (Supplementary Fig. 9a–c). In contrast, optogenetic activation of Calb1⁺ dSPNs of Calb1^IRES2-Cre mice yielded opposite outcomes, resulting in a substantial increase in movement velocity, coupled with an increase of ambulatory speed and duration, as well as a decrease in immobility frequency and duration (Fig. 4i–p, Supplementary Fig. 9d–f). Taken collectively, these findings demonstrate that Kremen1⁺ and Calb1⁺ dSPNs exert contrasting roles in locomotor control.

Fig. 4: Optogenetics activation of Kremen1+ and Calb1+ dSPNs during locomotion. — **Fig. 4: Optogenetics activation of *Kremen1*⁺ and *Calb1*⁺ dSPNs during locomotion.**

State-dependent regulation of locomotion by Kremen1 ⁺ and Calb1 ⁺ dSPNs

To examine whether the effects of Kremen1⁺ and Calb1⁺ dSPN activation depend on locomotion states, we further analyzed the 10 s 3 mW/20 Hz stimulation dataset (Supplementary Fig. 9c, f), categorizing trials into quiescent or ambulatory states immediately preceding stimulation onset or after offset (Fig. 5a–d). Activation of Kremen1⁺ dSPNs immediately suppressed ongoing ambulatory movement but slightly increased movement when mice were in a quiescent state (Fig. 5a, c). Conversely, activation of Calb1⁺ dSPNs had no immediate effect on ongoing ambulatory movement but triggered ambulatory locomotion when mice were in a quiescent state prior to stimulation (Fig. 5b, d). Additionally, the termination of Kremen1⁺ dSPN activation led to increased locomotion velocity, while the termination of Calb1⁺ dSPN stimulation resulted in reduced movement speed (Fig. 5e, f). Therefore, these findings highlight the opposing roles of Kremen1⁺ and Calb1⁺ dSPNs in locomotion regulation, which are influenced by the locomotion state at the time of activation.

Fig. 5: Optogenetics activation Kremen1+ and Calb1+ dSPNs exerted state-dependent regulation of locomotion. — **Fig. 5: Optogenetics activation *Kremen1*⁺ and *Calb1*⁺ dSPNs exerted state-dependent regulation of locomotion.**

Kremen1 ⁺ iSPNs play limited role in locomotor control

To investigate the role of Kremen1⁺ iSPNs in the context of locomotion, we introduced AAV vectors containing optogenetic activators that were both Cre and Flp-dependent (AAV1-ConFon-ChR2) into the dorsal striatum of Kremen1^2A-Cre;A2a^2A-Flp double KI mice (Fig. 6a). Kremen1⁺ iSPNs project exclusively to the GPe and not to the EPN or SNr (Fig. 6b, c; Supplementary Fig. 10a). Light stimulation was applied at the axon terminals of Kremen1⁺ iSPNs situated in the GPe (Fig. 6b, c; Supplementary Fig. 10b). The high-power stimulation at 3 mW/20 Hz for 3 min resulted in a modest increase in average velocity and duration per bout of movement (Fig. 6d–h), while at low light power level with 0.25 mW constant stimulation we did not observe any notable alteration in locomotion velocity (Fig. 6i). These results underscore the limited role of Kremen1⁺ iSPNs in regulating locomotion. Indeed, chemogenetic activation of both Kremen1⁺ dSPNs and iSPNs in Kremen1^2A-Cre mice led to reduced locomotor activity (Supplementary Fig. 11), supporting a major role of Kremen1⁺ dSPNs in suppressing locomotion.

Fig. 6: Optogenetics activation of Kremen1+ iSPNs leads to a modest increase of locomotion. — **Fig. 6: Optogenetics activation of *Kremen1*⁺ iSPNs leads to a modest increase of locomotion.**

Kremen1 ⁺ and Calb1 ⁺ dSPNs differently regulate dopamine release

To investigate how Kremen1⁺ and Calb1⁺ dSPNs distinctly regulate locomotion at the circuit level, we examined the impact of dSPN activity on dopamine release in the dorsal striatum. Both Kremen1⁺ and Calb1⁺ dSPNs innervate nigrostriatal DANs and modulate their activity^14,16,18,22. Previous studies have shown that optogenetic stimulation of patch neurons can inhibit dopamine release^23,24. Using the genetically encoded dopamine sensor GRAB_rDA3m²⁵ and Cre-dependent ChR2, we assessed dopamine release in the dorsal striatum of Kremen1^2A-Cre (Fig. 7a, b) and Calb1^IRES2-Cre (Fig. 7a, c) mice through fiber photometry following optogenetic stimulation at the SNr for each mouse line. Activation of Kremen1⁺ dSPNs led to a gradual reduction in dopamine levels in a stimulation dose dependent manner, with stimulation frequency of 20 Hz and durations of 2, 5, and 15 seconds (Fig. 7d). In contrast, activation of Calb1⁺ dSPNs induced a biphasic change in dopamine release: an initial transient increase in dopamine signals at the onset of optic stimulation, followed by a subsequent reduction during the stimulation (Fig. 7d). The initial surge of dopamine release following Calb1⁺ dSPN activation was present across different stimulation frequencies and durations (Fig. 7e). However, the magnitude of dopamine release reduction was far less pronounced with Calb1⁺ dSPNs stimulation compared to Kremen1⁺ dSPNs (Fig. 7f). Additionally, only Kremen1⁺ dSPN stimulation resulted in a post-stimulation rebound of dopamine release (Fig. 7g), consistent with a previous observation of DANs rebound firing following patch dSPN stimulation in brain slice¹⁶. The same pattern of differential dopamine release regulation by Kremen1⁺ and Calb1⁺ dSPNs was also observed when stimulating the neurons at 10 Hz (Supplementary Fig. 12). These findings suggest that Kremen1⁺ and Calb1⁺ dSPNs regulate dopamine release differently, with Kremen1⁺ dSPN activation leading to a more potent inhibition of dopamine release.

Fig. 7: Kremen1+ and Calb1+ dSPNs differentially regulate dopamine release. — **Fig. 7: *Kremen1*⁺ and *Calb1*⁺ dSPNs differentially regulate dopamine release.**

Genetic knockdown of GABA-B receptor Gabbr1 in Aldh1a1 ⁺ DANs promotes locomotion

Considering that Aldh1a1⁺ nigrostriatal DANs are major recipients of striatal inputs in the SNc and receive prolonged inhibitory signals mediated by GABA-B receptors from patch dSPNs^16,19 and that GABBR1 is the primary GABA receptor in DANs (plotted from PRJNA775656²⁶, Supplementary Fig. 13a), we genetically deleted Gabbr1 in Aldh1a1⁺ nigrostriatal DANs. This was achieved through the delivery of Cre-dependent CRISPR/saCas9-Gabbr1sgRNA targeting AAVs into the SNc of Aldh1a1^CreERT2 mice¹⁴ (Fig. 8a). The gene targeting-vector specifically disrupted the expression of GABBR1 in Aldh1a1⁺ DANs, while sparing Aldh1a1-negative neurons (Fig. 8b–e, Supplementary Fig. 14). The resulting knockdown (KD) of Gabbr1 in Aldh1a1⁺ nigrostriatal DANs led to a significant increase in ambulatory velocity (Fig. 8f, g). These findings support an important role of GABA-B receptor-mediated inhibitory signaling in Aldh1a1⁺ DANs for regulating locomotor control.

Fig. 8: Knockdown of Gabbr1 receptor in Aldh1a1+ DANs leads to hyperactivity. — **Fig. 8: Knockdown of *Gabbr1* receptor in *Aldh1a1*⁺ DANs leads to hyperactivity.**

Kremen1 ⁺ dSPNs suppress locomotion through the GABA-B receptors in Aldh1a1 ⁺ DANs

To investigate the role of postsynaptic GABA-B receptors in the locomotor inhibition induced by Kremen1⁺ dSPNs, we selectively activated Kremen1⁺ dSPNs through optogenetics while simultaneously downregulating Gabbr1 in Aldh1a1⁺ nigrostriatal DANs of Kremen1^2A-Cre;Aldh1a1^CreERT2 double KI mice (Fig. 9a, b). Notably, the only Kremen1⁺ cells present in the SN are pericytes, which do not express GABBR1 (Supplementary Fig. 13b). Since baseline locomotion differed between Gabbr1-KD and control mice (Fig. 8f, g), we computed a velocity ratio normalized to each mouse’s baseline for group comparison. Remarkably, Gabbr1 KD in Aldh1a1⁺ nigrostriatal DANs completely abolished both the locomotion-suppressing effects during light stimulation and the post-stimulation rebound typically induced by Kremen1⁺ dSPN activation (Fig. 9c–e). In contrast, control mice showed a marked reduction in velocity during stimulation and a rebound effect after stimulation (Fig. 9c–e).

Fig. 9: Knockdown of Gabbr1 receptors in Aldh1a1+ DANs abolishes Kremen1+ dSPN-induced locomotion alterations. — **Fig. 9: Knockdown of *Gabbr1* receptors in *Aldh1a1*⁺ DANs abolishes *Kremen1*⁺ dSPN-induced locomotion alterations.**

To access whether GABA-B receptors are involved in locomotor regulation by Kremen1-negative dSPNs, we optogenetically stimulated Kremen1-negative dSPNs in Kremen1^2A-Cre;Aldh1a1^CreERT2 double KI mice under both control and Gabbr1-KD conditions (Fig. 9f, g; Supplementary Fig. 15). While dSPN stimulation substantially increased locomotion in both groups, the absence of GABBR1 in Aldh1a1⁺ nigrostriatal DANs did not alter the locomotor effects induced by Kremen1-negative dSPNs (Fig. 9h–j). Together, these findings demonstrate that Kremen1⁺ dSPNs suppress locomotor activity by modulating Aldh1a1⁺ nigrostriatal DAN activity via GABBR1 signaling.

Kremen1 ⁺ dSPNs regulate Aldh1a1 ⁺ DAN activity via GABBR1 receptors

To further confirm that Kremen1⁺ dSPNs mediate locomotor regulation by modulating Aldh1a1⁺ DAN activity through GABBR1 receptors, we used fiber photometry to selectively measure calcium transients in the axon terminals of Aldh1a1⁺ DANs during optical activation of Kremen1⁺ dSPNs under both control and Gabbr1 KD conditions (Fig. 10a, b). When stimulated at 20 Hz for 15 s, optical activation of Kremen1⁺ dSPNs resulted in a significantly smaller reduction in calcium transients in the axon terminals of Aldh1a1⁺ DANs in Gabbr1 KD mice compared to controls (Fig. 10c, d; individual traces, Supplementary Fig. 16). Additionally, under the same stimulation conditions, Gabbr1 KD abolished the rebound activity of Aldh1a1⁺ DANs following Kremne1⁺ dSPN activation (Fig. 10c, e. These findings underscore a critical role of Aldh1a1⁺ DANs in Kremen1⁺ dSPN-regulated locomotor control and establish GABBR1 receptors as an important mediator (Fig. 10f).

Fig. 10: Genetic knockdown of Gabbr1 receptor affects Kremen1+ dSPN-induced alterations in Aldh1a1+ DAN activity. — **Fig. 10: Genetic knockdown of *Gabbr1* receptor affects *Kremen1*⁺ dSPN-induced alterations in *Aldh1a1*⁺ DAN activity.**

Discussion

In this study, we utilized the distinct expression patterns of Kremen1 and Calb1 in dorsal striatum to investigate the functional roles of diverse dSPN subtypes in locomotor controls. We found that while the activation of Calb1⁺ dSPNs promoted locomotion, consistent with the traditional role of dSPNs, Kremen1⁺ dSPNs– a subset of which form patch-like clusters in the dorsal striatum– suppressed locomotion. This regulation of locomotion by Kremen1⁺ and Calb1⁺ dSPNs is state-dependent, varying with the locomotor context during their activation. Although dSPNs are traditionally known to promote locomotion by inhibiting inhibitory outputs from the SNr¹, our findings reveal that Kremen1⁺ dSPNs suppress locomotion through inhibition of Aldh1a1⁺ DANs via GABBR1-mediated signaling. These results underscore the complex regulation of locomotion by distinct dSPN subpopulations and highlight the value of molecular makers, combined with targeted neuronal manipulation and recording, in elucidating the specialized functions of different neuronal subtypes and circuits.

While prior studies suggest that patch-like dSPNs constrain locomotion^24,27,28,29, these findings do not necessarily generalize to the entire patch dSPN population. It is important to note that defining “patch” structures based on marker expression can be somewhat arbitrary. Here, we defined putative patch structures as groups of five or more adjacent Kremen1⁺ SPNs and manually delineated these clusters, which include both Kremen1⁺ and Kremen1-negative neurons. Approximately 70% of these circled neurons were either Kremen1⁺ or Tshz1⁺. Notably, patch markers are also expressed by a considerable number of neurons within matrix compartments^10,11,20. Although the density of Kremen1⁺ SPNs is significantly lower in matrix compartments than in patch compartments, the total number of Kremen1⁺ SPNs in the two compartments is comparable. Given this distribution, we cannot distinguish whether Kremen1⁺ dSPN axon terminals originate from patch or matrix compartments under the current experimental conditions. Therefore, we avoided using the terms “patch” or “matrix” to describe Kremen1⁺ and Calb1⁺ SPNs, referring to them instead as molecularly defined SPN subpopulations.

The role of GPi-projecting Tshz1⁺ dSPNs has also been studied in locomotion, with Tshz1⁺ patch dSPNs implicated in locomotor suppression during place-preference tasks²⁸. However, changes in movement velocity during valence contexts (e.g., place preference tests) may reflect alterations in active avoidance or approach choices rather than pure locomotion¹³. Moreover, SNr-projecting dSPNs, which are known for locomotion control, were not explicitly examined in prior studies²⁸. Our data reveal that Kremen1 and Tshz1 dSPNs represent largely different subtypes of patch-like dSPNs. While Kremen1⁺ dSPNs inhibit locomotion via SNr projections, the role of SNr-projecting Tshz1⁺ SPNs remains unclear. These findings emphasize the need for future studies to explicitly identify neuronal subtypes based on both molecular markers and projection targets.

Kremen1⁺ dSPN activity increases much later than Calb1⁺ dSPNs and mostly after locomotion onset, suggesting they do not initiate movement but instead modulate or terminate it. In contrast, Calb1⁺ dSPNs show elevated activity that precedes locomotion onset, consistent with a role in movement initiation. At the locomotion offset, Kremen1⁺ dSPNs exhibit elevated and upward-trending neuronal activity compared to Calb1⁺ dSPNs, suggesting that they play a role in terminating movement and facilitating state transitions from locomotion to quiescence. This aligns with the established principle of basal ganglia involvement in state transition regulation³⁰. Optogenetic activation of Kremen1⁺ dSPNs shortened movement bout duration and increased immobility, supporting their role in transitioning between locomotor states.

Aldh1a1⁺ nigrostriatal DANs are the primary recipients of monosynaptic inputs from the dorsal striatum and form reciprocal innervation with dSPNs¹⁴. This feedback loop likely facilitates the regulation of dopamine release and SPN activity during motor control and learning. Both patch and matrix dSPNs innervate Aldh1a1⁺ DANs¹⁴. Notably, Kremen1⁺ and Calb1⁺ dSPN axon terminals occasionally form striosome-dendron bouquet-like structures^16,17 with the DAN dendrites in the SNr, a configuration that likely influences dopamine release in the dorsal striatum and SNr^16,18. Consistent with these in vitro studies, our in vivo recordings reveal a more pronounced inhibition of dopamine release upon stimulating Kremen1⁺ dSPNs compared to the Calb1⁺ dSPNs, with a rebound dopamine release observed upon the cessation of Kremen1⁺ dSPN stimulation.

Activation of Calb1⁺ dSPNs initially increases dopamine release, likely due to disinhibition of DANs through inhibition of parvalbumin neurons (PVNs) in the SNr, which tonically inhibit DANs^31,32,33,34. Conversely, stimulation of Kremen1⁺ dSPNs results in greater dopamine suppression, followed by rebound release after stimulation cessation. These opposing effects on dopamine release align with their respective locomotor functions: Calb1⁺ dSPNs promote movement, while Kremen1⁺ dSPNs terminate it. The genetic knockdown of GABBR1 in Aldh1a1⁺ DANs abolished the locomotor suppression and post-stimulation rebound activity induced by Kremen1⁺ dSPNs, confirming the critical role of GABBR1-mediated signaling. Fiber photometry further showed that Gabbr1 KD in Aldh1a1⁺ DANs reduced the suppression of DAN activity and abolished the rebound response following Kremen1⁺ dSPN activation.

Our study demonstrates that molecularly distinct dSPN subpopulations play specialized roles in locomotion control. Calb1⁺ dSPNs promote movement by suppressing inhibitory outputs from SNr, whereas Kremen1⁺ dSPNs primarily inhibit locomotion by suppressing Aldh1a1⁺ DANs through GABBR1-mediated signaling. These findings emphasize the importance of molecular markers in dissecting neuronal circuit mechanisms and highlight the intricate interplay between distinct dSPN subpopulations in basal ganglia motor control.

Methods

Mouse work

All mouse studies were conducted in accordance with the guidelines approved by the Institutional Animal Care and Use Committees (IACUC) of the Intramural Research Program of the National Institute on Aging (NIA), NIH. All mouse lines were maintained as heterozygotes in a C57BL/6 J background. The Kremen1^2A-Cre KI mice were generated by Shanghai Model Organisms Inc. (Shanghai, China). The Aldh1a1^CreERT2 KI mice were generated as previously described¹⁴. The Nr4a1–eGFP (Stock No: 036737-UCD) transgenic mice were obtained from Mutant Mouse Resource & Research Centers (MMRRC). The Calb1^IRES2-Cre mice (Stock No: 028523) and Ai14 (Stock No: 007908) were obtained from the Jackson laboratory. The A2a^2A-Flp KI mice were generated by Dr. Gerfen and the Rodent Transgenic Core of National Institute of Mental Health (NIMH). Both females and males were used for all experiments. Mice used for viral injections were between 2 and 4 months of age. The mice were housed in a 12-hour-light/12-hour-dark cycle in groups of 2-5 animals and had ad libitum water and a regular diet. All the behavioral tasks were performed during the light cycle. Littermates were randomly assigned to different groups prior to experiments.

Generation of Kremen1 ^2A-Cre KI mice

The Kremen1^2A-Cre KI mice were generated using the CRIPSR/Cas9 approach in the C57BL/6 J strain by Shanghai Model Organisms Inc. A donor plasmid containing the 2 A ribosome skipping sequence, Cre DNA recombinase sequence, and flanking Kremen1 mouse genomic DNA sequence was constructed. This was used to insert to the 2A-Cre DNA fragment into exon 9 immediately after the stop codon of Kremen1 gene locus, guided by the gRNA with sequence GTGGGCTTCAGTCACTCACG AGG. One founder mouse was generated, and the correct genomic modification was confirmed by sequencing.

Laser capture microdissection and RNA sequencing

One-month-old Nr4a1-eGFP transgenic mice were euthanized with CO₂ followed by rapid decapitation. The brains were immediately dissected and preserved at −80 °C. Cryosectioning of the frozen brains was performed at −20 °C, and the sections were mounted onto PAN membrane frame slides (Applied Biosystems, Foster City, CA). The dorsal striatum was determined based on anatomic landmarks such as the corpus callosum, lateral ventricle and nucleus accumbens. Using the ArturusXT microdissection system with fluorescent illumination (Applied Biosystems), the eGFP-positive island-like structures within the dorsal striatum of Nr4a1-eGFP transgenic mice were carefully isolated onto LCM Macro Caps (Applied Biosystems) and designated as “patch”, while surrounding tissues of similar size were also isolated onto Macro Caps and designated as “matrix”.

Total RNA extraction and purification were executed from hundreds of caps using the PicoPure Isolation kit (Applied Biosystems), with subsequent genomic DNA clearance facilitated by RNase-free DNase (Qiagen) following the manufacturer’s protocols. Quantification of RNA was performed using a NanoDrop spectrophotometer (ThermoFisher), and RNA integrity was evaluated using the Bioanalyzer RNA 6000 pico assay (Agilent). Only RNA samples exhibiting high integrity, as indicated by ribosomal integrity numbers (RIN) of 7 or higher, were selected for subsequent patch and matrix RNAseq library preparation.

The cDNA libraries were prepared from the purified RNA using the TruSeq Stranded Total RNA LT library preparation kit (Illumina) following the manufacturer’s protocol. The quality of the libraries was assessed using the Bioanalyzer DNA 1000 assay (Agilent) before sequencing on an Illumina HiSeq 2000 platform. Fastq files were generated using the standard Illumina pipeline. Transcript abundance, annotated by Ensembl, was quantified using Salmon in a non-alignment-based mode, and gene-level counts were estimated utilizing the Tximport package (Bioconductor). Normalization of counts and subsequent data analysis were conducted following previously established procedures²⁶. The accession number of the striatal tissue RNA-seq is PRJNA870469.

RNA in situ hybridization and image analysis

RNA in situ hybridization (ACDBio, RNAscope) was used to detect the expression of Drd1, Drd2, Kremen1 and Tshz1 mRNAs in the dorsal striatum of adult C57BL/6 J mice. For tissue preparation, mice were anesthetized with CO₂ and rapidly decapitated. The brains were fresh-frozen on dry ice and stored at -80 °C before sectioning. Striatal sections (12 µm) were collected using a cryostat (Leica Biosystems) and stored at -80 °C until processed.

RNAscope was conducted according to the instructions of RNAscope Multiplex Fluorescent Reagent Kit v2 user manual. Probes against Drd1 (Cat. No. #401901), Drd2 (Cat. No. #406501), Kremen1 (Cat. No. #425771) and Tshz1 (Cat. No. #494291) were applied to brain sections. Fluorescent images were acquired using a laser scanning confocal microscope LSM 780 (Zeiss) with 20× or 40× lens. Bregma coordinates around 1.34 mm, 0.92 mm, 0.5 mm, 0.08 mm and -0.34 mm were selected for image analysis.

RNAscope images were analyzed using Imaris (v10.0.0, Bitplane, Belfast Northern Ireland, UK). Surfaces for the dorsal and ventral striatum were created using the Allen Brain Atlas as a reference. The spatial patch was defined by outlining Kremen1⁺ SPN clusters (a minimum of 5 SPNs), and the density of Kremen1⁺ SPNs within a spatial patch is at least 200 cells/mm². Surfaces for individual channels (i.e., Drd1, Drd2, Kremen1, Tshz1 and DAPI) were created with unique parameters, which were saved and applied to all images within the same batch. Minor adjustments were made in subsequent rounds of RNAscope to improve quantification accuracy. DAPI surfaces in the striatum were filtered by the mean intensity of Drd1 (dSPNs) and the mean intensity of Drd2 (iSPNs). These Drd1⁺ and Drd2⁺ cells were further filtered by the overlapped volume ratio to Kremen1 to quantify patch dSPNs and patch iSPNs, respectively. RNAscope analysis with Tshz1 was conducted in a similar manner, filtering Drd1⁺ and Kremen1⁺ cells by the overlapped volume ratio to the Tshz1 surface. All data points are presented as the average across striatal sections of both hemisphere for each mouse.

Due to the limitation of staining a maximum of three probes simultaneously using RNAscope V2 reagents, we stained different batches of brains with Kremen1, Drd1, and Drd2 probes for Fig. 1d, and with Kremen1, Tshz1, and Drd1 probes for Supplementary Fig. S1. The slightly difference in the numbers arise because the data were generated from two separate batches of brains and counted by different individuals.

Immunohistochemistry

Mice were anesthetized with pentobarbital and transcardially perfused with precooled PBS, followed by 4% paraformaldehyde (PFA) solution as described previously^14,35. Brains were stored in 4% PFA at 4 °C overnight and then transferred to 30% PBS buffered sucrose solution for at least 2 days before sectioning. Series of coronal sections (40 µm) were collected using a cryostat and stored at 4 °C in PBS. Sections were blocked in 10% normal donkey serum (Sigma-Aldrich) + 0.5% Bovine serum albumin (Sigma-Aldrich) + 0.3% Triton for 1 hr at room temperature (RT). Next, sections were incubated with the primary antibodies over one or two nights at 4 °C. Sections were then washed with PBS (3 × 10 min) and incubated with fluorescent secondary antibodies at RT for 1 hr. In some experiments, sections were further washed twice in PBS, incubated with DAPI (40, 6-diamidino-2-phenylindole, Invitrogen, D1306) (0.5 mg/ml in PBS) for 1 min. After washing with PBS (5 min), sections were mounted onto subbed slides, and coverslipped with mounting media (ProLong® Gold Antifade Mountant, Life technology). Images were taken using a laser scanning confocal microscope LSM 780 (Zeiss). The primary antibodies used for immunostaining included rabbit monoclonal anti-TH (Pel-Freez Biologicals, P40101; dilution 1:1000), mouse monoclonal anti-TH (ImmunoStar, 22941; dilution 1:1000), chicken polyclonal anti-TH (Aves Labs, TYH; dilution 1:500), rabbit anti-Mu-opioid receptor (MOR) (ImmunoStar, 24216; dilution 1:3000), chicken polyclonal anti-GFP (Aves Labs, GFP-1020; dilution 1:1000), mouse polyclonal anti-RFP (Rockland, 200-301-379; dilution 1:1000), guinea pig anti-Parvalbumin (Swant, GP72; dilution 1:1000), rabbit anti-Somatostatin-14 (T-4103, Peninsula Laboratories; dilution 1:1000), goat anti-ChAT (AB144P, Millipore; dilution 1:500), rabbit anti-Iba1 (Wako, 019-19741, 1:1000), rabbit anti-GFAP (Abcam, ab7260; dilution 1:2000), goat anti-CD13 (R&D Systems, AF2335; dilution 1:100), and mouse monoclonal anti-GABA_B (Abcam, ab55051, dilution 1:500). Appropriate fluorophore-conjugated secondary antibodies (Life Technologies) were used depending on the desired fluorescence colors.

Neuron counting in Gabbr1 KD mice

Coronal midbrain sections from Gabbr1 KD and Ctrl mice (40 μm thickness, approximately four sections per mouse spanning Bregma −2.70 mm to −3.80 mm) were stained with anti-TH, anti-ALDH1A1 and anti-GABBR1 antibodies for quantification. Images of the SN were captured as single layers by a laser scanning confocal microscope (LSM 780, Zesis) with × 20 objective lens. The percentage of GABBR1 receptor-positive neurons in ALDH1A1⁺/TH⁺ and ALDH1A1^-/TH⁺ dopaminergic neurons was qualified using ImageJ (v1.53t). The SNc region was manually delineated based on previously described method¹⁴.

Viral vectors

The following AAV vectors were purchased from Addgene (Watertown, MA, USA): AAV9-hSyn-DIO-hM4D(Gi)-mCherry (#44362), AAV9-hSyn-DIO-mCherry (#50459), AAV1-EF1a-double floxed-hChR2(H134R)-EYFP (#20298), AAV1-hSyn-hChR2(H134R)-EYFP (#26973), AAV8-hSyn-Con/Fon-hChR2(H134R)-EYFP (#55645), AAV8-hSyn-Con/Fon-EYFP (#55650), AAV1-Ef1a-DIO-EYFP (#27056), and AAV1-syn-FLEX-jGCaMP8s (#162377). AAV2/9-hSyn-DIO-GRAB-rDA3m were produced by BrainVTA (Gaithersburg, MD, USA). AAV9-FLEX-SaCas9-U6-sgGabbr1 was packaged by SignaGen Laboratories (Frederick, MD, USA). AAV9-CMV-NLS-SaCas9-3XHA-sNRPpa-U6-Basal-empty was packaged by Vigene Biosciences (Rockville, MD, USA).

Construction of pAAV-FLEX-SaCas9-U6-sgGabbr1

The reference sequence of Gabbr1 gene was retrieved from the UCSC genome browser database (http://genome.ucsc.edu/), and subsequent identification and alignment of exons were performed using the Mouse Genome Informatics (MGI) database. The most 5′ common coding exons were then selected, and the sequence was uploaded to the CRISPOR website (http://crispor.org) for identifying potential sgRNAs and PAM sequences. Three Gabbr1 sgRNAs were synthesized as short oligos (Eurofins Genomics) with a 5′ CACC- 3′ overhang on the forward primer, and a 5′ -AAAC 3′ overhang on the reverse primer, facilitating seamless integration into the pX601-AAV-CMV;NLS-SaCas9-NLS-3xHA-bGHpA;U6;BsaI-sgRNA vector, a gift provided by Dr. Feng Zhang (Addgene plasmid # 61591). Sanger sequencing confirmed the insertion of the Gabbr1 sgRNA using the primer TAACCACGTGAGGGCCTATTTC.

Mouse Neuro-2a (N2a) cell lines were cultured in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% FBS (HyClone) at 37 °C with 5% CO₂ incubation. Transfection of Cells with the plasmid pX601-AAV-CMV;NLS-SaCas9-NLS-3xHA-bGHpA;U6;sgGabbr1 vector was performed using Neuro-2a Transfection Kit (Altogen Biosystems), following the manufacturer’s protocol. Cells were then harvested for PCR-based identification of mutations induced by genome editing using the Guide-it Mutation Detection Kit (Cat. No. 631443). The sgGabbr1 with the sequence of CACCGCGCCACACTCCACAATCCCAC exhibited high efficiency in genome editing under SaCas9 and was thus selected for further integration into the pAAV-Flex-SaCas9-U6-sgDaglb vecgtor²⁶. The primers 5’-TAACCACGTGAGGGCCTATTTCCCATGATT and 5’-CGCTCGGTCCGAAAAATCTCGCCAACAAGTT were used to amplify the fragment containing sgGabbr1, which replaced sgDalgb in the plasmid pAAV-Flex-SaCas9-U6-sgDaglb at the PmlI and RsrII registration sites. Sanger sequencing confirmed the presence of sgGabbr1 in the resulting pAAV-Flex-SaCas9-U6-sgGabbr1 plasmid.

Stereotaxic injection and optic fiber implantation

The stereotaxic survival surgery was performed as previously described²⁶. All surgery were conducted under aseptic conditions, and body temperature was maintained using a heating pad. Briefly, adult mice (2-4 months old) were anesthetized with isoflurane (1-2%) and head-fixed in a stereotaxic frame (Kopf Instruments). A total volume of 500-700 nL of AAVs was injected unilaterally or bilaterally into the dorsal striatum with two locations per hemisphere (coordinates: 0.5 mm AP to Bregma, ±2.2 mm ML, −2.5 mm DV from dura surface; 1.3 mm AP from Bregma, ±1.8 mm ML, −2.5 mm DV from dura surface) or SN (coordinates: −3.1 mm AP from Bregma, ±1.5 mm ML, −3.9 mm DV from dura surface). The infusion of viruses was controlled by a stereotaxic injector (Stoelting) at a speed of 75 nL/min. After a 5-minute wait following the end of injection, the injector was slowly withdrawn. The scalp was then sutured, and the mice were returned to their home cages. All behavior experiments were performed at least 4 weeks after injection to allow time for full heterologous gene expression. For Aldh1a1^CreERT2 mice, 4-OHT was injected intraperitoneally at a dosage of 10 mg/kg bodyweight for five consecutive days, beginning one week after surgery, to induce Cre recombinase expression.

To prepare mice for optogenetics or fiber photometry experiments, we performed a second surgery to implant optic fibers at least 3 weeks after viral injection. For behavioral optogenetics experiments, optical fiber stubs (200 μm core, 0.39 NA, Thorlabs) were bilaterally implanted with the tips positioned over the SNc (−3.1 mm AP from Bregma, ±1.5 mm ML, −3.9 mm to −4.1 mm DV from dura surface) or GPe (−0.3 mm AP from Bregma, ±2.0 mm ML, −3.5 mm from dura surface). The optic fibers were secured in place with a thick layer of radiopaque adhesive cement (C&B METABOND, Parkell). Once dried, Vetbond tissue adhesive (3 M) was applied to seal the head incision with adhesive cement.

For simultaneous photometry recordings and optogenetics experiments, optic fiber stubs with a 200 μm core and 0.39 NA (Thorlabs) were unilaterally implanted over the SN ( − 3.1 mm AP from Bregma, 1.5 mm ML, −3.9 mm to −4.1 mm DV from dura surface) for optogenetic stimulation. In addition, optic fiber stubs with a 200 μm core and 0.5 NA (Plexon) were implanted over dorsolateral striatum (+1.0 mm AP from Bregma, 2.0 mm ML, −2.2 mm to −2.5 mm DV from the dura surface) on the same side for striatal rDA3m photometry recording during optogenetic stimulation.

For the treadmill fiber photometry experiment, optic fiber stubs with a 200 μm core and 0.5 NA (Thorlabs) were unilaterally or bilaterally implanted in the SN ( − 3.1 mm AP from Bregma, 1.5 mm ML, −4.0 mm to −4.3 mm DV from dura surface) to record the axon calcium signals of patch or matrix dSPNs. The optic fibers were fixed in place using adhesive cement. A titanium metal head-bar (Labeotech) was mounted on top of the adhesive cement rostral to the fiber stubs for head-restraint. The animals were allowed to recover for at least one week after the optic implantation before any optogenetics or fiber photometry experiments.

Open-field spontaneous locomotion

Locomotion in freely moving mice was measured using video tracing analyses. Mice were habituated for 30 min in the behavioral room, wherein a 20 W lamp was shielded in a box and positioned in the dark opposite to the testing area to provide a diffuse light source. For the video tracing test, mice were placed in a 50 cm × 50 cm gray opaque chamber for 30 min. Activity was recorded from overhead by a digital camera at a frame rate of 15 Hz. EthoVision XT software (Noldus) was used to track the mice and analyze the video for velocity, time, and distance traveled.

Chemogenetic manipulation

JHU37160-dihydrochloride (Hello Bio) was dissolved in water to achieve a stock concentration of 0.3 mg/mL and stored in small aliquots at -20 °C. Prior to each use, the working solution was freshly prepared by diluting the stock 10-fold with 0.9% saline. Mice received 0.3 mg/kg bodyweight dosage via intraperitoneal injection 30 minutes before behavioral tests.

Optogenetic stimulation

A LED light source with commutator (PlexBright, Plexon) was connected to a multimodal optic fiber patch cable (200 μm core, 0.39 NA, Plexon) through ceramic sleeves (Thorlabs) to the ferrule of the optic fiber stubs previously implanted in the mouse. Light power was calibrated using a power meter (PM100D, Thorlabs) to achieve the desired output measured at the tip of optic fiber. For ChR2 experiments, photo stimulation consisted of 5ms-width blue light pluses (465 nm, 3 mW) at different frequency or constant blue light (465 nm, 0.25 mW). Light pulses were generated by a TTL pulse generator (OPTG-4, Doric Lenses).

Optogenetics with open field

Mice were habituated in the testing room 30 min before testing, and the apparatus was cleaned with 50% ethanol between animals. Mice were placed in a 50 cm × 50 cm clear chamber and their activities were captured by top-view and side-view cameras (Logitech). After a 3 min exploration period, mice received either 3 min ON and 3 min OFF bilateral stimulation or 10 s ON and 1 min OFF bilateral stimulation. Video and LED light TTL were acquired and synchronized with Synapse software (Tucker-Davis Technologies). Distanced traveled and velocity during the acquisition period were calculated using EthoVision XT software. In Figs. 4 and 9, baseline or prior was identified as the 3 min period prior to 1^st stimulation, 1^st Off indicates 3 min period post 1^st stimulation, and 2^nd Off indicates 3 min period post 2^nd stimulation. Ambulation bouts were scored as periods of movement >2 cm/s lasting for > 0.5 s and separated by >0.5 s. Immobility bouts were scored as periods of <2% pixel change lasting for >0.5 s and separated by >0.5 s. For behavioral-state dependent effects of optogenetic stimulation experiment, mice received 10 s ON and 1 min OFF bilateral 20 Hz stimulation. Ambulation state was determined by movement velocity >2 cm/s in the entire 0.5 second immediately preceded either stimulation onset or stimulation offset. Quiescence (includes small movement) state was determined by movement velocity <1.5 cm/s in the entire 0.5 second immediately preceded either stimulation onset or stimulation offset. Due to kinematics of stimulation on behavior, quantification for effects of stimulation on velocity change used 1 s interval instead.

Optogenetics with fiber photometry

Mice were allowed to freely explore an open field chamber while receiving optical stimulation lasting 2 s, 5 s, or 15 s across approximately 10 trials per stimulation duration, with intervals of at least 45 s between trials. To prevent optic fiber patch cables from tangling, a pigtailed fiber-optic rotary joint (Doric Lens) was used to connect two multimodal optic fiber patch cables in the photometry light path. Real-time rDA3m or GCaMP8s signals and timing of TTL pulses were recorded and synchronized using Synapse software (Tucker-Davis Technologies). For data analysis, ΔF/F changes were normalized by subtracting each sample’s own baseline, defined as the average ΔF/F value during the 5 s preceding stimulation onset. The maximum reduction in dopamine or GCaMP amplitude was identified as the lowest ΔF/F value within 1 s interval prior to stimulus offset, representing the peak reduction in response to stimulation. Post-stimulation changes in rDA3m or GCaMP8s signals were calculated as the average ΔF/F during the 5 s interval spanning 4-9 s after stimulus offset, capturing the rebound phase. For Calb1^IRES2-Cre mice, the initial dopamine elevation was quantified as the average ΔF/F during the 0.5 s interval between 0.5 s and 1 s after stimulus onset.

Fiber photometry

Dopamine sensor or GCaMP8s fluorescence was measured using a locked-in amplifier system (Tucker-Davis Technologies, Model RZ10X with Synapse software). The photometry recordings were conducted with either green (GCaMP8s) or red (rDA3m) fluorescence. For GCaMP8s recordings, the blue LED (465 nm) was sinusoidally modulated at 330 Hz, and the UV LED (405 nm) was modulated at 211 Hz as an isosbestic control channel. For rDA3m recordings, the green LED (560 nm) was modulated at 410 Hz, and the UV LED was modulated at 211 Hz. The peak intensity of each LED was calibrated to 20–60 μW, measured at the distal end of the patch cable. Light emissions were filtered through a 6-port fluorescence mixing cube (Doric Lens) before being coupled to an optic patch cable (200μm core, 0.5 NA), affixed to the implanted optic fiber in each mouse. The emitted fluorescent signals were collected by integrated photosensors in RZ10X real-time signal processor equipped with lock-in amplifier. Transistor-transistor logic (TTL) signals were employed to timestamp onset times for each event of interest (e.g., stimulation onset, locomotion onset and offset), which were detected via the RZ10X in the Synapse software. Fiber photometry data was analyzed using custom MATLAB code. Demodulated 465 nm, 560 nm, and 405 nm recording traces were recorded at a sampling rate of 1k Hz.

For the analysis of rDA3m signals during optogenetic stimulation, to capture the large offset change caused by optogenetic stimulation, the demodulated photometry trace of rDA3m was normalized to compute ΔF/F. F was estimated as a referenced rDA3m signal, where a RANSAC ordinary least square linear regression was performed between the demodulated UV isosbestic reference signal and demodulated rDA3m signal to transform the reference signal to account for the differences in gain and offset between the two signals, as well as possible motion artefacts and long-term photo bleaching effects. Then the referenced rDA3m signal (F) was subtracted from the demodulated rDA3m signal to compute ΔF. The same reference approach was also applied to the analysis of GCaMP8s signals of Aldh1a1⁺ DANs during optogenetic stimulation.

To analyze the modulation of GCaMP8s activity change with locomotor behavior, photometry signal normalization followed a published study³⁶, using custom MATLAB code. Demodulated photometry traces of both GCaMP8s and UV isosbestic reference channels were pre-normalized by first computing ΔF/F0. F0 was estimated by calculating the 10th percentile of the raw photometry amplitude using a 5-sec sliding window to account for slow, correlated fluorescence changes, including photobleaching in both channels. ΔF was calculated as raw photometry amplitude subtracted its respective F0. Both channels were initially normalized with this procedure. An additional referencing procedure was performed to remove the effects of motion or mechanical artefacts from analysis. For this referencing procedure, the ΔF/F0 of the UV reference signal was low-pass filtered with a second-order Butterworth filter with a 3 Hz corner frequency. Then a RANSAC ordinary least squares regression between the filtered reference signal and normalized GCaMP8s signal was used to transform the reference signal to account for the differences in gain and offset between the two signals. Lastly, the transformed references trace was subtracted from the normalized GCaMP8s trace as the final ΔF/F. The transformed reference trace was then used as UV ΔF/F. Only the experiments where the maximum percentage of DF/F exceeded 1.5 and the GCaMP8s and the UV reference correlation was below 0.6 were included for further analysis. Z-score of ΔF/F was calculated with mean and standard deviation of the entire recording session.

Head-fixed voluntary treadmill with fiber photometry

The head-fixed locomotion was achieved with a compact low friction manually driven treadmill, originally designed by Janelia Research Campus of the HHMI, purchased from LABmaker (Berlin, Germany). Teflon belt movement was tracked by a rotary encoder, which sends 0–3.3 V analog output of speed and direction to an ADC port at TDT RZ10x to synchronize with photometry recordings. The speed was calibrated with 10 cm/s at 2.5 V and sampled at 1k Hz. The head fixture was achieved through head fixation system along with an implantable titanium head bar from Labeotech (Montreal, Canada). Mice started treadmill habituation training one week after recovering from head-bar implant surgery to get used to walking on treadmill while head was fixed. Each habituation session lasted 20 min and repeated at most twice a day for 3-4 days until mice were able to spontaneously initiate at least 20 walk bouts during a 20 min session. After habituation training, a 30 min fiber photometry recording session was performed for each individual mouse, while they walked on the treadmill with head fixation. No reward was provided neither during training nor during recording.

Movement versus rest time bins were defined with a 0.25 cm/s threshold on the velocity trace. Isolated movement periods with duration shorter than 0.5 s or movement periods with average velocity smaller than 0.5 cm/s were excluded for analysis. Because mice tended to walk slower on the treadmill, time bins were considered as rest periods only if they lasted longer than 0.8 s. A movement bout initiation time was defined as velocity cross the threshold at the end of the rest period, and a movement termination time was defined as the time velocity fell below the threshold followed by a rest period. Timestamp of maximum value of z-scored GCaMP activity during 1 s before and 1 s after locomotion onset or offset was calculate the time of maximum activity for the respective events. Activity slope during locomotion offset was computed as the coefficient of a least-square linear regression fit for the z-scored GCaMP activity during the interval of 0.5 s to 0 s before locomotion offset. and after locomotion offset. Timestamp of activity rise at locomotion onset was calculated with a 20-ms sliding window to compare with baseline activity which was defined as mean activity during an interval of -1s to -0.5 s preceding the locomotion onset. The 20-ms sliding window started at -0.5 s with a step of 2 ms towards locomotion onset. Mean activity of the sliding window then compared with the baseline activity across all trials (movement bouts) using a right-tailed pairwise t-test. An activity rise was defined as p < 0.05 in consecutive 5 steps, the center of the first sliding window was used as the activity rise time.

Statistics and reproducibility

For the fluorescent images as representative panels in the figures, no less than 3 mice from the same treatment group were checked for the validity. Data were analyzed by Prism 9 software (Graphpad) and custom code written in MATLAB (MathWorks). All statistical details for each experiment can be found in the corresponding figure legends. Data were presented as means and standard errors of the mean (SEM). We assessed the statistical significance using parametric t-test, one and two-way analysis of variance (ANOVA). Significance was considered for test statistics with a (^*) p-value of <0.05, (^**) p-value of <0.01, (^***) p-value of <0.001, (^****) p-value of <0.0001.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

The accession number of the striatal tissue RNA-seq data is PRJNA870469. The accession number of SNc RNA-seq data is PRJNA775656. All data generated or analyzed during this study are included in this published article. Source data are provided with this paper.

Code availability

The code used to process the fiber photometry data and optogenetics behavior analysis in this study has been deposited in GitHub [https://github.com/jdNIH/Kremen1_2025] and can also be accessed on Zendo [https://zenodo.org/records/14796086].

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Acknowledgements

This work is supported in part by the Intramural Research Programs of National Institute on Aging, NIH (HC, ZIA AG000944, AG000928), National Institute of Mental Health (CRG, ZIA MH002497-34), and National Nature Science Foundation of China (WDL, 32220103006). We thank Dr. Yulong Li of Peking University for providing the GRAB_rDA3m sensor, NIMH rodent behavioral core for assisting in behavioral tests, Dr. Sarah Hawes for sharing Calb1^IRES-Cre mice for the optogenetic study, Drs. Guohong Cui and Jingheng Zhou of National Institute of Environmental Health Sciences and members of Cai lab for their suggestions and technical assistance, and Drs. Da-Ting Lin of National Institute of Drug Abuse, Yangfeng Zhang of University of Exeter, and Jun Wang of Texas A & M University for their constructive feedback.

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Open access funding provided by the National Institutes of Health.

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Authors and Affiliations

Transgenic Section, Laboratory of Neurogenetics, National Institute on Aging, National Institutes of Health, Bethesda, MD, 20892, USA
Jie Dong, Lupeng Wang, Breanna T. Sullivan, Lixin Sun, Victor M. Martinez Smith, Lisa Chang & Huaibin Cai
Computational Biology Group, Laboratory of Neurogenetics, National Institute on Aging, National Institutes of Health, Bethesda, MD, 20892, USA
Jinhui Ding
Clinical Research Center on Neurological Diseases, the First Affiliated Hospital, Dalian Medical University, Dalian, Liaoning, 116011, China
Weidong Le
Institute of Neurology, Sichuan Academy of Medical Sciences-Sichuan Provincial Hospital, Medical School of University of Electronics & Technology of China, Chengdu, Sichuan, 610045, China
Weidong Le
Section on Neuroanatomy, National Institute of Mental Health, National Institutes of Health, Bethesda, MD, 20892, USA
Charles R. Gerfen

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Jie Dong
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Lupeng Wang
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Breanna T. Sullivan
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Lixin Sun
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Victor M. Martinez Smith
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Lisa Chang
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Jinhui Ding
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Weidong Le
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Charles R. Gerfen
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Huaibin Cai
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Contributions

H.C. conceived and designed experiments, wrote the manuscript, and prepared the figures with inputs from all authors. J.D. designed and performed stereotactic surgery, optogenetics, fiber photometry, histology, behavioral experiments and data analyses, prepared figures, wrote methods and figure legends, and wrote/edited the manuscript. L.W. designed and conducted fiber photometry experiments with head-fixed mice, performed data analyses, prepared figures and figure legends, wrote methods, and wrote/edited the manuscript. B.S. performed RNA in situ hybridization and data analyses, histology, mouse breeding, and behavioral experiments. L.S. contributed to the laser capture microdissection and RNA sequencing of the patch and matrix compartments, built the pAAV-FLEX-SaCas9-U6-sgGabbr1 vector, and wrote method for these experiments. V.M. contributed to RNAscope image analyses and histology. J.H.D. contributed to RNA sequencing data analyses. L.C. performed stereotactic surgery. W.L. contributed to data interpretation and edited the manuscript. C.R.G. provided A2a^Flp mice, contributed to data interpretation, and edited the manuscript. All authors read and approved the final manuscript.

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Correspondence to Huaibin Cai.

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Dong, J., Wang, L., Sullivan, B.T. et al. Molecularly distinct striatonigral neuron subtypes differentially regulate locomotion. Nat Commun 16, 2710 (2025). https://doi.org/10.1038/s41467-025-58007-x

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Received: 05 December 2024
Accepted: 10 March 2025
Published: 19 March 2025
Version of record: 19 March 2025
DOI: https://doi.org/10.1038/s41467-025-58007-x