Research ArticleNeuroscience

The acid-sensing ion channel ASIC1a mediates striatal synapse remodeling and procedural motor learning

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Sci. Signal.  07 Aug 2018:
Vol. 11, Issue 542, eaar4481
DOI: 10.1126/scisignal.aar4481

ASIC channels in motor learning

Members of the ASIC family of acid-sensing ion channels are abundant in various regions of the brain and mediate neuronal synaptic function. Yu et al. found that ASIC1a is present in neurons in the striatum. Manipulating ASIC1a expression in striatal neurons in mice revealed that this channel is critical for promoting the synaptic abundance and function of NMDA receptors in response to changes in extracellular pH. By activating a kinase-dependent transcriptional program that promotes the expression of NMDA receptor subunits, ASIC1a channels stimulate synaptic activity and dendritic spine maturation in the striatal neurons that facilitate motor learning. Mice lacking ASIC1a were slower to learn new motor coordination tasks. These findings may have implications for both neuronal development and neuronal disorders that affect the striatum and motor control.


Acid-sensing ion channel 1a (ASIC1a) is abundant in multiple brain regions, including the striatum, which serves as the input nucleus of the basal ganglia and is critically involved in procedural learning and motor memory. We investigated the functional role of ASIC1a in striatal neurons. We found that ASIC1a was critical for striatum-dependent motor coordination and procedural learning by regulating the synaptic plasticity of striatal medium spiny neurons. Global deletion of Asic1a in mice led to increased dendritic spine density but impaired spine morphology and postsynaptic architecture, which were accompanied by the decreased function of N-methyl-d-aspartate (NMDA) receptors at excitatory synapses. These structural and functional changes caused by the loss of ASIC1a were largely mediated by reduced activation (phosphorylation) of Ca2+/calmodulin-dependent protein kinase II (CaMKII) and extracellular signal–regulated protein kinases (ERKs). Consequently, Asic1a null mice exhibited poor performance on multiple motor tasks, which was rescued by striatal-specific expression of either ASIC1a or CaMKII. Together, our data reveal a previously unknown mechanism mediated by ASIC1a that promotes the excitatory synaptic function underlying striatum-related procedural learning and memory.


As the primary input nucleus to the basal ganglia, the dorsal striatum, which consists of caudate and putamen, forms a key part of the extrapyramidal motor system (1, 2). In addition to motor control, the dorsal striatum also mediates a particular form of learning called “procedural learning and memory,” in which stimulus-response associations or habits are incrementally acquired (3). This contrasts with “declarative learning,” which depends on the medial temporal lobe memory system and uses the hippocampus as a primary component. The striatum receives excitatory afferents from the cortex and thalamus and is densely innervated by midbrain dopamine neurons.

The excitatory striatal synapses are considered a key neural substrate for motor control and procedural memory, because they undergo activity-dependent synaptic plasticity that alters the transfer of information throughout basal ganglia circuits (4). To achieve this, synapses in medium spiny neurons (MSNs), which have densely spinous dendrites (5) and represent most neurons in the striatum, undergo remodeling characterized as the transformation of dendritic spine density and morphology in addition to changes in postsynaptic architecture and glutamate receptor function. Synaptic remodeling constitutes an adaptive mechanism essential for striatum-related motor control and learning. However, despite the compelling evidence for an association between synaptic remodeling and striatum-related motor learning (4), critical molecular determinants that mediate these processes are incompletely understood.

Proton-gated acid-sensing ion channels (ASICs) belong to the degenerin/epithelial Na+ channel (DEG/ENaC) superfamily (6) and include at least six isoforms: ASIC1a, ASIC1b, ASIC2a, ASIC2b, ASIC3, and ASIC4. ASIC1a is the dominant isoform in the central nervous system (79). In addition to mediating acid-evoked currents, ASIC1a also plays critical roles in synaptic plasticity in multiple brain regions, including the hippocampus (1013), the amygdala (14, 15), and the cortex (16). The loss of ASIC1a not only abolishes acid-evoked currents in neurons from these brain regions but also causes deficits in several forms of associative learning and memory. Ca2+-dependent signaling and function have been implicated as the homomeric ASIC1a, and heteromeric ASIC1a/2b channels are Ca2+-permeable (1719). Moreover, ASIC1a plays important roles in regulating long-term potentiation (LTP) at glutamatergic synapses in the hippocampus (1013) and the amygdala (14, 15) as well as the induction of long-term depression (LTD) in the insular cortex (16), possibly through detecting acute acidification in the synaptic cleft (14, 20). Although the roles of ASIC1a in hippocampus-dependent learning have been called into question (10, 11), Asic1a null mice show deficits in multiple forms of learning, such as amygdala-dependent fear learning and memory (14, 15, 21, 22), cerebellum-dependent eye-blink conditioning (10), and extinction learning of conditioned taste aversion (16).

ASIC1a is also abundantly expressed in the striatum, but its function there is not yet clear. It is known that ASIC1a exerts region-specific roles in regulating synaptic structure and function (23, 24). For example, whereas ASIC1a expression is positively correlated with dendritic spine density in the hippocampus (25), it is negatively correlated with spine density of MSNs in the nucleus accumbens, where the overexpression of ASIC1a suppresses cocaine-evoked plasticity (20). Here, we investigated ASIC1a function in the dorsal striatum, a region with predominant expression of the homomeric ASIC1a channels (17, 26). Using a combination of morphological, electrophysiological, and behavioral assays, we unveil a crucial role of ASIC1a in regulating excitatory synaptic structure and function of striatal MSNs, and its contribution to striatum-related motor coordination and learning.


ASIC1a is enriched in postsynaptic density fraction of mouse striatum

Previous studies revealing the function of ASIC1a largely focused on the cortex (16) and hippocampus (1013). However, ASIC1a is also abundant in the striatum (17, 26). To examine ASIC1a function in the striatum, we first systematically characterized the mRNA and protein expression levels as well as the subcellular distribution of ASIC1a in the mouse striatum. Compared to the cortex and hippocampus, the striatum of wild-type (WT) mice had greater expression of ASIC1a at both the mRNA (Fig. 1A) and protein (Fig. 1, B and C) levels. ASIC1a expression was absent in brain tissues obtained from the Asic1a knockout (KO) mice (Fig. 1B), confirming the specificity of the ASIC1a antibody. To ensure that the loss of ASIC1a did not cause up- or down-regulation of other ASIC isoforms, we also examined ASIC2a and found that Asic2a expression in the striatum was unaltered by Asic1a deletion (fig. S1), consistent with the previous study (26) showing that ASIC1a is the predominant form of ASICs in the mouse striatum.

Fig. 1 ASIC1a is enriched in the PSD fraction of the mouse striatum and is involved in regulating motor coordination and learning.

(A) Quantitative polymerase chain reaction (PCR) assessment of Asic1a mRNA amounts in the mouse cortex, hippocampus (Hipp), and striatum. n = 3 for each group. *P < 0.05 and ***P < 0.001, unpaired Student’s t test. (B and C) Representative immunoblots (B; −/−, striatum from an Asic1a KO mouse) and quantification (C) of ASIC1a protein abundance in the mouse cortex, hippocampus, and striatum. n = 3 for each group. *P < 0.05 and ***P < 0.001, unpaired Student’s t test. GAPDH, glyceraldehyde-3-phosphate dehydrogenase. (D) Enrichment of ASIC1a in the PSD fraction of the striatum shown by immunoblotting. GluN2B was used as a positive control. (E and F) Representative patch-clamp traces (E) and statistical analysis (F) of evoked EPSCs in dorsal striatal MSNs from WT (+/+) and Asic1a KO (−/−) mice before (WT, light gray; KO, pale red) and after (WT, black; KO, red) the application of CNQX (20 μM), d-APV (50 μM), and PTX (100 μM), without or with amiloride (500 μM) (WT, orange; KO, yellow). Arrows (E: left, WT, black; right, KO, red) indicate ASIC-dependent synaptic currents. n = 6 cells from three WT mice and 7 cells from three Asic1a KO mice. *P < 0.05, **P < 0.01, and ***P < 0.001, paired Student’s t test, comparison as indicated. ##P < 0.01, WT versus KO, unpaired Student’s t test. (G) Motor-related behaviors of WT and Asic1a null mice in the incremental fixed-speed rotarod learning test. n = 10 for each group. Two-way repeated-measures analysis of variance (ANOVA), main effect of genotype, F1,100 = 6.824, P = 0.011. *P < 0.05, unpaired Student’s t test. (H and I) Assessment of motor coordination, as time to fall (H) and distance traveled (I) in beam walking tests, in WT and Asic1a null mice. n = 7 to 9 for each group. Two-way repeated-measures ANOVA, main effect of genotype, F1,80 = 24.107, P < 0.001. *P < 0.05 and **P < 0.01, unpaired Student’s t test.

We further examined the subcellular localization of ASIC1a in the striatum and found it to be highly concentrated in the purified synaptosomes and postsynaptic density (PSD) fractions of the mouse striatum (Fig. 1D). This is consistent with previous findings in other brain regions, suggesting that ASIC1a is prominently enriched in postsynaptic structures (10, 21, 23, 25, 27, 28). Collectively, our data indicate that ASIC1a is abundant in the postsynaptic site of excitatory synapses in striatal neurons.

ASIC1a contributes to excitatory postsynaptic currents in MSNs of the dorsal striatum

To measure ASIC1a-mediated synaptic response in striatal neurons, we evoked excitatory postsynaptic currents (EPSCs) in MSNs of the dorsal striatum in mouse brain slices by stimulating cortical glutamatergic inputs under whole-cell voltage clamp mode. After blockade of α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA), N-methyl-d-aspartate (NMDA), and A-type γ-aminobutyric acid (GABAA) receptors with 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX; 20 μM), d-(−)-2-amino-5-phosphonopentanoic acid (d-APV; 50 μM), and picrotoxin (PTX; 100 μM), respectively, a small component of the EPSC remained and occurred in the same time frame as the AMPA-mediated EPSC in WT neurons. Amiloride (500 μM), the nonselective ASIC blocker, inhibited this component (Fig. 1, E and F), which was also largely absent in Asic1a KO neurons (Fig. 1, E and F). These findings are comparable with synaptic ASIC1a currents detected during normal synaptic transmission in other brain regions (14, 20, 29), supporting the idea that—as in other brain regions—ASIC1a contributes to an important portion of EPSCs in the dorsal striatum.

Loss of ASIC1a impairs motor coordination and learning

Because of the importance of the striatum in motor control, motor coordination, and motor learning (1, 2), we performed a series of motor-related behavioral tests on the Asic1a KO mice and their WT littermates. The basal locomotor activity of Asic1a KO mice was similar to WT, as shown by an open field test (fig. S2, A and B). The distance traveled over a period of 120 min was nearly identical (fig. S2B), and both genotypes showed similar short-term habituation within the 120-min test (fig. S2A). We further analyzed the footprints to characterize the primary locomotor gait (30, 31). The WT and Asic1a KO mice exhibited comparable front and hind stride lengths (fig. S2C), measured as the average distance of forward movement between each stride. The base widths, defined as the average distance between left and right footprints, were also similar between WT and Asic1a KO mice for both the front and hind paws (fig. S2D). Notably, the WT and Asic1a KO mice had nearly identical (close to zero) overlap between forepaw and hindpaw placement (fig. S2E), measured as the distance between the front and hind footprints on each side. Together, these suggest that Asic1a KO mice largely maintain normal locomotion and locomotor gait.

We next used an accelerating rotarod test (32) to evaluate the motor learning capability of the WT and Asic1a KO mice. The duration by which the mice maintained balance, meaning the latency to fall, was not significantly different between WT and Asic1a KO animals at all three acceleration speeds tested (0.12, 0.2, and 0.6 rpm/s) (fig. S2F). We then used an incremental fixed-speed rotarod learning protocol to assess their motor learning capability, which consists of the same test in five consecutive days (three trials each day). Notably, here, Asic1a KO mice showed significantly poorer performance than their WT controls (Fig. 1G). The difference between WT and Asic1a KO mice was significant during the first 2 days but diminished on the third day and thereafter (Fig. 1G). These results suggest that the loss of ASIC1a may slow down motor learning capability.

To determine whether fine motor coordination is impaired in the Asic1a KO mice, we assessed their performance on a balance beam test (33). Asic1a KO mice took significantly more time to cross the horizontal beams of various sizes and shapes (28-, 17-, and 11-mm round beams, as well as 28- and 12-mm square beams), with the exception of the 5-mm square beam, on which both genotypes performed poorly (Fig. 1H). To subject the animals to greater motor challenges (34), rather than resting or general moving, we used a modified inclined balance beam test, in which the animal had to crawl upward or downward on an inclined beam at a 40° angle. Similar to the horizontal beam tests, Asic1a KO mice spent significantly more time traversing the inclined balance beam than did WT controls, regardless of whether moving upward or downward (fig. S2G). The results of these behavioral tests indicate that the loss of ASIC1a may impair the ability to maintain coordination during a motor challenge.

To further characterize the motor learning capability, we trained the mice to cross the 5-mm square horizontal beams in five consecutive days and recorded the average distance the individual animal traveled on each day (two trials per day). Asic1a KO mice consistently traveled a shorter distance than their WT littermates (Fig. 1I). Whereas the WT mice learned to cross the whole distance of the beam in ~3 days, some of the Asic1a KO mice needed nearly 5 days of training to accomplish the task (Fig. 1I). Because Asic1a KO mice eventually attained the similar beam crossing distance as the WT animals, the poorer motor coordination and leaning capability to traverse the narrow beam likely reflect a specific motor deficiency rather than the lack of motivation to cross the beam or other unidentified factors. Together, these results suggest that ASIC1a is involved in motor coordination and motor learning, which represents a previously unidentified function in procedural learning and memory mediated by ASIC1a.

ASIC1a deficiency increases striatal spine density but decreases its maturation

To explore the mechanism(s), we first focused on the morphological and functional changes of excitatory synapses formed on the MSNs, because alterations in dendritic spine density and morphology have been implicated in motor-related function (4) and dysfunction (3537). In addition, previous studies showed opposite effects mediated by ASIC1a in regulating dendritic spine densities in the hippocampus (25) and the nucleus accumbens (20). We therefore examined whether and how loss of ASIC1a function could alter dendritic spine density and morphology in striatal MSNs. Golgi staining of MSNs in dorsal striatum slices (fig. S3) revealed an increase in dendritic spine density in Asic1a KO compared to their WT littermates (Fig. 2, A and B), similar to previous data obtained from ventral striatum (the nucleus accumbens) (20). Analysis of spine morphology showed that in Asic1a KO neurons, the spine head width was significantly decreased (Fig. 2C), without a significant change in spine length (Fig. 2D).

Fig. 2 Loss of ASIC1a increases dendritic spine density but disrupts spine maturation in dorsal striatum.

(A and B) Representative micrographs (A) and statistical analysis (B) of Golgi staining of striatal dendritic segments and spines of WT (Asic1a+/+) and Asic1a KO (Asic1a–/–) mice. Data are mean and distribution of 109 dendritic segments from eight WT mice and 66 dendritic segments from six KO mice. ***P < 0.001, unpaired Student’s t test. (C and D) Cumulative plots showing spine head width (C) and length (D) in MSNs from WT and Asic1a–/– mice. n = 2946 (C) and 2971 (D) spines from three WT mice and n = 2952 (C) and 2826 (D) spines from four KO mice. ***P < 0.001, two-sample Kolmogorov-Smirnov test. (E to G) Representative traces (E) and cumulative distribution plots of amplitudes (F) and frequency (G) of mEPSCs obtained by patch recordings from MSNs in dorsal striatal slices of WT (Asic1a+/+) and KO (Asic1a−/−) mice. Inset bar graphs show average for each; n = 26 cells from eight WT mice and 27 cells from six KO mice. (H to L) Quantification of different subtypes of spines in WT and KO MSNs. n = 24 dendritic segments from three mice for each genotype. *P < 0.05 and ***P < 0.001, unpaired Student’s t test.

To determine whether the changes in spine density and subtype distribution might alter basal glutamatergic synaptic transmission, we recorded miniature EPSCs (mEPSCs) in MSNs of dorsal striatum slices of WT and Asic1a KO mice. However, no significant difference was detected in either mEPSC frequency or mean amplitude (Fig. 2, E to G), suggesting that AMPA receptor (AMPAR)–mediated basal glutamatergic synaptic transmission was not affected by the loss of ASIC1a.

Upon more detailed analysis of spine morphology, we found that Asic1a KO neurons had significant increases in the percentages of stubby spines and filopodia, but not that of thin spines and mushroom spines (Fig. 2, H to L). The stubby spines and filopodia are thought to represent immature excitatory synapses, whereas the mushroom spines are thought to represent more mature synapses (38, 39). Therefore, it is possible that the loss of ASIC1a suppressed spine maturation (decrease number of mushroom spines) and resulted in an increase in the overall spine density in the dorsal striatum due to accumulation of immature spines (stubby spines and filopodia). To test this hypothesis, we analyzed the size of PSD in the dorsal striatum using electron microscopy. PSD is an electron-dense structure located at the spine head, which consists of densely packed ion channels, surface receptors, as well as cytoplasmic scaffolding and signaling proteins (40). The size and protein composition of PSD are dynamically regulated during activity- or experience-dependent remodeling of the synapse (4143), a structure thought to be important for learning and memory (44). Both the thickness and length of PSD were significantly reduced in Asic1a KO samples as compared to those from WT controls (Fig. 3, A to C), indicating a smaller size of postsynaptic architecture, which is consistent with our hypothesis that the loss of ASIC1a results in more immature excitatory synapses in the dorsal striatum.

Fig. 3 PSDs in striata of Asic1a null mice exhibit altered morphology and protein compositions.

(A) Representative electron micrographs of striatal neurons from WT (Asic1a+/+) and Asic1a KO (Asic1a–/–) mice depicting the synaptic contact and PSD. (B) Cumulative plots showing the PSD thickness in WT and Asic1a KO MSNs. n = 719 PSDs from three WT mice and 967 PSDs from four KO mice. ***P < 0.001, two-sample Kolmogorov-Smirnov test. (C) Cumulative plots showing PSD length in WT and Asic1a KO MSNs. n = 753 PSDs from three WT mice and 1100 PSDs from four KO mice. ***P < 0.001, two-sample Kolmogorov-Smirnov test. (D and E) Immunoblots of PSD and total proteins including GluN1, GluN2A, GluN2B, and PSD95, but not GluA1, GluA2, and PICK1, in striata prepared from WT (+/+) and Asic1a null (–/–) mice: (D) representative immunoblots; (E) statistical analysis of the results. n = 4 to 8 each group. *P < 0.05 and **P < 0.01, WT versus KO, unpaired Student’s t test.

In parallel with the morphological changes, our analyses of several major PSD proteins in the PSD fractions of striatal tissues also demonstrated significant decreases in the abundance of NMDA receptor (NMDAR) subunits GluN1, GluN2A, and GluN2B and its principal scaffold protein PSD protein 95 (PSD95), but not that of AMPAR subunits GluA1 and GluA2 or protein that interacts with C kinase 1 (PICK1) (Fig. 3, D and E). These changes were only found in the PSD fraction but not in the total striatal tissue, indicating a structure-related alteration in the postsynaptic distribution of NMDARs because of Asic1a deletion.

To determine the functional consequence, we recorded NMDAR- and AMPAR-mediated currents in striatal MSNs from WT and Asic1a KO mice (Fig. 4A). Whereas the AMPAR currents appeared to have similar amplitudes between the two genotypes, the NMDAR current was markedly smaller in Asic1a KO than in WT neurons. A significant decrease in the NMDAR/AMPAR ratio was detected in Asic1a null mice as compared to their WT littermates (Fig. 4B). This suggests a reduction in postsynaptic NMDAR activity in Asic1a KO neurons because—as described above—mEPSC frequency and amplitude (Fig. 2, E to G), as well as AMPAR abundance in the PSD (Fig. 3, D and E), were not changed by ASIC1a loss. Furthermore, plotting the input-output relationships of EPSC amplitudes against stimulating intensities revealed a significant reduction of synaptic responses mediated by NMDAR, but not AMPAR, in Asic1a KO neurons (fig. S4), validating the decrease in postsynaptic NMDAR, but not AMPAR, activity in Asic1a KO neurons.

Fig. 4 Asic1a null mice exhibit impaired NMDAR function.

(A and B) Representative traces (A) and statistical analysis (B) of NMDAR (top) and AMPAR (bottom) currents recorded with patch clamp from striatal MSNs in dorsal striatal slices prepared from WT (+/+; left) and Asic1a KO (−/−; right) mice. Data are NMDAR/AMPAR ratios for 14 cells from four WT mice and 12 cells from five Asic1a KO mice. Stimulation artifacts were removed for clarity. **P < 0.01, unpaired Student’s t test. (C and D) Locomotor behaviors of WT and Asic1a null mice in the open field test in response to intraperitoneal injection of NMDAR antagonist MK-801 (0.25 mg/kg, intraperitoneally at time 0). n = 16 for each group: (C) distance traveled of WT (Asic1a+/+) and Asic1a–/– mice in each 10-min interval within the 120-min test. Two-way repeated-measures ANOVA, main effects of genotype, F1,384 = 61.213, P < 0.001. *P < 0.05, **P < 0.01, and ***P < 0.001, Asic1a+/+ versus Asic1a–/–, unpaired Student’s t test. (D) Total distance traveled by Asic1a+/+ and Asic1a–/– mice during the entire 120-min test. ***P < 0.001, unpaired Student’s t test.

To verify the link between ASIC1a and NMDAR function in regulating the motor behavior, we used a behavioral model of hyperlocomotor response to NMDAR antagonists (45). We reasoned that if the behavioral phenotype seen in Asic1a KO mice was caused by a reduction of NMDAR function, then the KO mice would be more sensitive to NMDAR antagonists, such as MK-801, than WT animals. Intraperitoneal injection of MK-801 (0.25 mg/kg) induced a significantly greater increase in the locomotor activity of Asic1a KO than WT mice (Fig. 4, C and D). These data are consistent with the notion that synaptic NMDAR function is weakened in Asic1a null animals.

Striata of Asic1a null mice display impaired CaMKII and ERK signaling

Homomeric ASIC1a channels constitute the dominant ASIC subtype in striatal MSNs (26). These channels are thought to be Ca2+-permeable (17). In addition, NMDARs also mediate Ca2+ influx. Therefore, the loss of ASIC1a in striatal MSNs may lead to reduced Ca2+ influx due to the lack of Ca2+ entry through ASIC1a and reduced NMDAR activity. Ca2+ influx exerts its effects on dendritic spine remodeling (38, 39, 46) and functional synaptic plasticity (4) through the activation of downstream signaling pathways, including Ca2+-sensitive Ca2+/calmodulin-dependent protein kinase II (CaMKII) and the downstream mitogen-activated protein kinases (MAPKs) (4750). We therefore examined the amount of phosphorylated CaMKII (Fig. 5, A and B) and extracellular signal–regulated protein kinase 1 (ERK1) and ERK2 (ERK1/2; Fig. 5, C and D) in striatal tissues and found both to be significantly decreased in Asic1a KO mice compared to their WT controls. However, no significant change was observed in the phosphorylation of p38 MAPK (fig. S5, A and B) or c-Jun N-terminal kinase (JNK; fig. S5, C and D). The selective reduction in the CaMKII-ERK pathway, but not p38 or JNK, suggests a specific coupling of ASIC1a/NMDAR to ERK signaling.

Fig. 5 Striata of Asic1a null mice exhibit impaired activation of CaMKII-ERK signaling.

(A to D) Representative immunoblots and statistical analysis of phosphorylated (“p-”) and total protein abundance of CaMKII (A and B) and ERK1 and ERK2 (C and D) in striata from WT (+/+) and Asic1a null (−/−) mice. n = 4 to 6 for each group. *P < 0.05, **P < 0.01, and ***P < 0.001, unpaired Student’s t test. (E to H) Effects of treatment with acid (pH 6.0) on the phosphorylation of CaMKII (E and F) and ERK (G and H). (E and G) Representative immunoblots and analysis showing phosphorylated and total protein abundances of CaMKII (E and F) and ERK1 and ERK2 (G and H) in cultured striatal neurons prepared from WT mice and maintained at pH 7.4 only or exposed to a pH 6.0 external solution for 2 min without or with pretreatment (30 min) of PcTX1 (20 nM) or KN93 (10 μM), as indicated. n = 6 to 15 for each group. **P < 0.01 and ***P < 0.001, compared with pH 7.4 only; #P < 0.05, compared with pH 6.0 alone, unpaired Student’s t test. (I to L) Representative immunoblots and statistical analysis of the effects of the same acid treatment on the phosphorylation of CaMKII (I and J) and ERK (K and L) in cultured striatal neurons prepared from Asic1a null (Asic1a–/–) mice. n = 3 for each group.

Acid activates CaMKII-ERK signaling through ASIC1a in striatal neurons

The decreased CaMKII-ERK activity could potentially account for the reduced spine maturation (Fig. 2), reduced PSD size (Fig. 3, A to C), altered synaptic composition (Fig. 3, D and E), and further reduction of NMDAR function (Fig. 4) in the Asic1a KO striatum. If so, we would expect the activation of ASICs to increase the phosphorylation of CaMKII and ERK1/2. To test this possibility, we measured acid-induced phosphorylation of ERK and CaMKII in cultured striatal neurons and its dependence on ASIC1a. A brief acidic treatment (2 min at pH 6.0) significantly increased the phosphorylation of CaMKII and ERK1/2 (Fig. 5, E to H). These effects were largely attenuated by application of psalmotoxin 1 (PcTX1; 20 nM), a specific ASIC1a inhibitor (Fig. 5, E to H), suggesting that acid could enhance the CaMKII-ERK pathway in the striatal neurons through activation of ASIC1a. Conversely, the acid-induced kinase activities were abolished by the CaMKII inhibitor KN93 (Fig. 5, E to H), suggesting a signaling hierarchy order from ASIC1a to CaMKII and then to ERK1/2. Furthermore, the acid treatment failed to change CaMKII and ERK phosphorylation in striatal neuron cultures prepared from Asic1a null mice (Fig. 5, I to L), further demonstrating the involvement of ASIC1a in the acid-induced activation of CaMKII-ERK cascade. As controls, the phosphorylation of the other two MAPKs, JNK and p38, was not significantly altered by the acid treatment (fig. S5, E to H), suggesting that acidosis specifically up-regulates the ERK pathway.

Similar results were obtained in human embryonic kidney (HEK) 293 cells (fig. S6), which endogenously express ASIC1a, suggesting a general existence of acid-activated, ASIC1a-dependent ERK signaling cascade that is independent of any striatum-specific factor. Together with the findings that CaMKII-ERK activities are down-regulated in striata of Asic1a KO mice, these data establish a critical link between ASIC1a activity and CaMKII-ERK signaling, which may underlie the mechanism of ASIC1a in regulation of striatal synaptic remodeling.

Acute inhibition of ASIC1a in the striatum impairs motor learning

Thus far, the data collectively point to a critical role of ASIC1a in striatal synapse remodeling that depends on the CaMKII-ERK signaling cascade, implicating a possibility that the striatum is a site responsible for the effects of ASIC1a in motor coordination and learning. To test this hypothesis, we examined the effect of acute inhibition of ASIC1a in the horizontal beam walking tests. PcTX1 (20 pmol) or vehicle was injected bilaterally into the dorsal striata 30 min before each behavioral test. The administration of PcTX1 did not affect the time the mice spent to cross the horizontal beams of various sizes and shapes (28-, 17-, and 11-mm round beams, as well as 28- and 12-mm square beams; fig. S7A). On the 5-mm square beam, both the vehicle- and PcTX1-treated groups fell off from the beam before they finished crossing (fig. S7A). These results suggest that acute inhibition of ASIC1a channel activity in the dorsal striatum does not interfere with motor coordination. However, in the following motor learning tests on the 5-mm square beam, administration of PcTX1 30 min before the training test on each day for five consecutive days significantly attenuated motor learning compared with the administration of vehicle, as assessed by the distances the animals traveled on the beam (fig. S7B). These data suggest that, instead of being directly involved in motor coordination, continued ASIC1a activity in the striatum is critical for procedural learning of motor skills.

Reexpression of ASIC1a in the striata of Asic1a KO mice rescued defects in CaMKII-ERK signaling, spine density and structure, NMDAR function, and motor learning

To further validate the role of striatal ASIC1a in motor learning, we examined whether reintroducing ASIC1a into the dorsal striatum of adult Asic1a KO mice could rescue the behavioral deficits. To express ASIC1a specifically in dorsal striatal neurons, we created an adeno-associated virus (AAV) that contained the coding sequence of ASIC1a driven by the human synapsin I (hSynI) promoter (AAV-ASIC1a; Fig. 6A). AAV-ASIC1a or a control virus (AAV-GFP; Fig. 6A) was bilaterally injected into the dorsal striata of adult (8- to 10-week-old) Asic1a KO mice (Fig. 6B). We performed the experiments (biochemical assessment, electrophysiology, or behavioral tests) 4 weeks after viral injections, when the expression of the viral-encoded exogenous proteins in striatal tissues was evident (Fig. 6C and fig. S8). Notably, injection of AAV-ASIC1a, but not AAV-GFP, restored the phosphorylation levels of CaMKII (p-CaMKII) and ERK1/2 (p-ERK1/2) to those seen in WT striata (Fig. 6, C to E), strengthening our conclusion that ASIC1a in the dorsal striatum is involved in the activation of CaMKII-ERK cascade. Moreover, the reintroduction of ASIC1a in the striatum of adult Asic1a KO mice also largely corrected the alterations in spine density and maturation of the mutants (fig. S9). Notably, supporting the functional coupling between ASIC1a and NMDAR, the NMDAR/AMPAR ratio was normalized or even enhanced in samples from AAV-ASIC1a–injected mice (Fig. 6, F and G). These results indicate that the synaptic deficits caused by Asic1a gene deletion are plastic and reversible even at the adult age.

Fig. 6 Reintroduction of ASIC1a in the striatum rescued defects of CaMKII-ERK signaling, glutamate receptor function, motor coordination, and motor learning in Asic1a KO mice.

(A) Schematics of AAV vectors engineered to express a control construct [green fluorescent protein (GFP)] or ASIC1a. (B) Verification of AAV injection and expression, showing an example of AAV-mediated enhanced yellow fluorescent protein (EYFP) expression in the striatum. Scale bar, 1 mm. (C) Expression of ASIC1a protein (top) in striata of Asic1a null mice after the injection of AAV-ASIC1a, but not control (GFP), as shown by immunoblotting. A GFP antibody was used to detect both GFP from the AAV-GFP virus and EYFP from the AAV-ASIC1a virus (second panel from the top). Note that EYFP generated from the EYFP-2A-mASIC1a fusion protein by cleavage contained 21 extra amino acids of the 2A peptide and therefore migrated slower than just GFP. The fourth and sixth panels show that AAV-ASIC1a injection into the striata of Asic1a null (Asic1a–/–) mice also restored the phosphorylation levels of CaMKII and ERKs, respectively. (D and E) Statistic results for the ratios of phosphorylated/total CaMKII (D) and ERKs (E). n = 3 experiments for each group. *P < 0.05, unpaired Student’s t test. (F and G) Effects of AAV injection into the striata on the NMDAR/AMPAR ratio in MSNs. (F) Representative current traces of NMDAR (top) and AMPAR (bottom) currents, respectively. (G) Statistic results. n = 12 cells from three mice for each group. **P < 0.01 and ###P < 0.001, unpaired Student’s t test. (H and I) Effects of AAV injection into the striata on the performance in beam walking (H) and learning to cross the 5-mm square beam (I). (H) n = 8 to 10 for each group. *P < 0.05, **P < 0.01, and ***P < 0.001, compared with the Asic1a+/+ + AAV-GFP group. #P < 0.05 and ##P < 0.01, compared with the Asic1a–/– + AAV-GFP group. (I) n = 10 to 11 for each group. KO: two-way repeated-measures ANOVA, main effects of AAV, F1,110 = 26.706, P < 0.001.

Furthermore, the reintroduction of ASIC1a in the dorsal striata of adult Asic1a KO mice successfully rescued the behavioral impairments of the mutants, measured by horizontal beam walking tests, in terms of both motor coordination (Fig. 6H) and motor learning (Fig. 6I), as well as in traversing the inclined balance beam (fig. S10). These results not only strongly support the notion that the striatum is a major brain region in which ASIC1a exerts its role on motor-related behaviors but also demonstrate the reversibility of the behavioral deficits by restoration of ASIC1a expression in the dorsal striatum of the adult animal.

ASIC1a mediates striatal synapse remodeling and procedural motor learning through CaMKII-ERK signaling in the dorsal striatum

Finally, we examined whether ectopically increasing CaMKII/ERK activity in the dorsal striatum could correct the synaptic and behavioral deficits in Asic1a KO animals. AAV-CaMKII or a control virus (AAV-GFP; Fig. 7A), driven by the hSynI promoter, was bilaterally injected into dorsal striata of adult (8- to 10-week-old) Asic1a KO mice (Fig. 7B). Four weeks after the injection, the abundance of both total and phosphorylated CaMKII was increased in striatal tissues of AAV-CaMKII–injected mice (Fig. 7, B and C). The overexpression of CaMKII also increased the phosphorylation of ERK1/2 (Fig. 7, B and D) and the NMDAR/AMPAR ratio (Fig. 7, E and F). Furthermore, the overexpression of CaMKII in the dorsal striata of adult Asic1a KO mice improved the performance on the mutants in horizontal beam walking tests, in terms of both motor coordination (Fig. 7G) and motor learning (Fig. 7H). These results definitively suggest that the CaMKII-ERK pathway underlies the mechanism of ASIC1a regulation of synaptic NMDAR function in striatal MSNs that underlies procedural motor learning in mice. Collectively, our data demonstrate that ASIC1a in the dorsal striatum confers a critical role in synaptic remodeling via the CaMKII-ERK cascade, which, in turn, regulates procedural motor learning (Fig. 8).

Fig. 7 Overexpression of CaMKII in the striatum corrects the defects of CaMKII-ERK signaling, NMDAR function, and motor learning in Asic1a KO mice.

(A) Schematics of AAV vectors engineered to express a control construct (GFP) or CaMKII. (B) Verification of CaMKII overexpression and its effects on CaMKII and ERK phosphorylation in striata of Asic1a null mice: representative immunoblots. (C and D) Statistic results for the total and phosphorylation levels of CaMKII (C) and the ratios of phosphorylated/total ERKs (D). n = 4 to 5 experiments for each group. *P < 0.05 and **P < 0.01, unpaired Student’s t test. (E and F) Effects of AAV injection into the striata of Asic1a null mice on the NMDAR/AMPAR ratio in MSNs. (E) Representative current traces of NMDAR (top) and AMPAR (bottom) currents, respectively. (F) Statistic results. n = 10 cells from three AAV-GFP–injected mice and 7 cells from three AAV-CaMKII–injected Asic1a KO mice. **P < 0.01, unpaired Student’s t test. (G and H) Effects of AAV injection into the striata of Asic1a null mice on the performance on beam walking (G) and learning to cross the 5-mm square beam (H). (G) n = 7 to 8 for each group. *P < 0.05 and **P < 0.01, unpaired Student’s t test. (H) n = 7 to 8 for each group. Two-way repeated-measures ANOVA, main effects of AAV, F1,75 = 17.844, P < 0.001. *P < 0.05 and **P < 0.01, unpaired Student’s t test.

Fig. 8 Proposed mechanism for ASIC1a regulation of striatal synaptic remodeling and motor learning.

Postsynaptic ASIC1a channels are activated by drops in the pH in the synaptic cleft associated with striatal synaptic activities, leading to cation (such as Na+ or Ca2+) influx. The increase in intracellular Ca2+ activates the downstream CaMKII-ERK signaling pathway. Whereas CaMKII presumably contributes to actin dynamics to promote structural remodeling of dendritic spines and regulates postsynaptic distribution and function of NMDAR, ERK signaling presumably participates in the activity-dependent transcriptional regulation of a set of neuronal proteins, which, in turn, drive long-term synaptic plasticity. Together, the ASIC1a-CaMKII-ERK signaling cascade represents a novel molecular mechanism that facilitates synaptic remodeling in the striatum, which is important for procedural motor learning.


Here, we show that ASIC1a regulates striatal synaptic remodeling via the CaMKII-ERK signaling pathway and contributes to striatum-related procedural motor learning (Fig. 8). Notably, ASIC1a plays quite similar roles in synaptic remodeling in dorsal and ventral striata (that is, nucleus accumbens) (20) in that Asic1a KO MSNs in both regions exhibit increased spine densities, especially spines that represent immature excitatory synapses (for example, stubby spines) and decreases in the NMDAR/AMPAR ratio. However, in the nucleus accumbens, the loss of ASIC1a results in an increase in mEPSC frequency (20), which was not detected in the dorsal striatum. Because it is anticipated that the dorsal and ventral striata regulate different behaviors, it should not come as a surprise that, through synaptic remodeling, ASIC1a in the nucleus accumbens affects cocaine-seeking behaviors (20), whereas that in the dorsal striatum regulates procedural motor learning.

Mechanistically, MSNs in the nucleus accumbens have mostly PcTX1-insensitive and Ca2+-impermeable heteromeric ASIC1a/2a channels (20), whereas those in the dorsal striatum express mainly the PcTX1-sensitive homomeric ASIC1a channels that are Ca2+-permeable (26). This may explain why the loss of ASIC1a affected AMPAR function in the former but mainly NMDAR activity in the latter brain area. Notably, we demonstrated the critical involvement of the Ca2+-dependent CaMKII/ERK signaling pathway in mediating the ASIC1a-dependent effects in the dorsal striatum, including the maintenance of NMDAR activity and motor learning efficiency. It is well established that CaMKII and ERK coordinately regulate synaptic remodeling (38, 39), including an interplay with actin to regulate structural dynamics of dendritic spines (47, 48), an effect on postsynaptic distribution and function of NMDARs (5153), and activity-dependent transcriptional regulation of certain neuronal proteins that confer long-term synaptic plasticity (49, 50). Because of the heterogeneous nature of striatal neurons (5), more in-depth work is needed in the future to determine which of these cell types in the dorsal striatum might be important for the behavioral and physiological effects of ASIC1a observed here. Nevertheless, we speculated that ASIC1a in MSNs would play a more dominant role in the striatum-related functions, because our manipulations and the structural and functional analyses consistently sampled these subtypes of neurons.

Since the discovery of ASICs, the source of protons that cause their activation and the contributions of these channels in electrical and Ca2+ signaling in neurons under physiological and pathophysiological conditions have been subjects of debate. High-frequency stimulation of amygdalar synapses triggers massive release of transmitters along with protons to sufficiently acidify synaptic clefts, which causes ASIC1a activation and contributes to LTP (14). In the nucleus accumbens, the evoked EPSC contains an ASIC1a/2a component sensitive to manipulations that disrupt acidification and/or ASIC function (20). Our demonstration of an ASIC1a-dependent component in the evoked EPSC from mouse dorsal striatal MSNs is entirely consistent with the above findings, indicating ASIC1a activation as an inevitable consequence of normal synaptic transmission by protons co-released with neurotransmitters. Alternatively, glial cells may provide another proton source for the activation of neuronal ASIC1a (54). In either case, the proton-evoked activation of ASIC1a channels seems to couple to the CaMKII-ERK pathway particularly well in a context-independent manner, because it was detected not only in dorsal striatal MSNs but also in HEK-293 cells. Our behavioral data indicate that the ASIC1a-CaMKII-ERK cascade is activated in dorsal striatal MSNs during motor challenges, leading to synaptic remodeling, which underlies procedural motor learning (Fig. 8).

To this end, it is interesting that an acute application of PcTX1 into the dorsal striatum did not significantly alter motor performance on the balance beams, but its continued presence during motor challenges impaired motor learning. This would be consistent with the idea that it takes time from ASIC1a activation to synaptic remodeling because of the need for structural changes. The PcTX1 treatment differs from ASIC1a genetic ablation in that, for the latter, synaptic remodeling in response to routine physical activities and/or the 2-day preconditioning training on the 12-mm square beam was already impaired because of the loss of ASIC1a, and therefore, the mutant animals would perform more poorly than WT mice in the beam walking tests. This implies that motor learning is a continued process, to which ASIC1a constantly contributes. The findings that the motor learning deficits can be corrected in the mutant mice by reintroduction of ASIC1a or enhancing CaMKII expression or created by administration of ASIC1a inhibitor into dorsal striata of adult animals suggest that the ASIC1a-CaMKII-ERK–regulated synaptic remodeling is plastic and maintained throughout the life span of the animal, rather than a developmental function.

Noticeably, the loss of ASIC1a did not completely abolish striatal spine maturation and motor learning but attenuated or slowed down these processes, suggesting a modulatory action of ASIC1a to improve the efficiency of procedural learning. This modulatory function may be an important feature during evolution because it gives an edge for the animal to survival in the competing and often harsh environment. The modulatory rather than the absolutely essential role of ASIC1a in motor control may explain why the global Asic1a KO mice exhibit subtle motor dysfunction despite a robust impairment in procedural motor learning, although ASIC1a channels are also highly expressed in the cerebellum and spinal cord (17). Moreover, the finding that dorsal striatum–specific overexpression of ASIC1a was sufficient to correct the motor learning deficiency in global Asic1a KO mice was likely due to the predominant role of the dorsal striatum in procedural motor learning and memory (3, 55, 56). Finally, despite the obvious abnormalities in spine density and NMDAR function, Asic1a null animals did not exhibit marked motor dysfunction. This may be explained by the fact that cellular or protein activities including ASIC1a do not have to work at their full capacity to maintain a normal physiological function, because many physiological functions have spare capacity, which provides the “safety margin” for life.

Nonetheless, brain region– or circuit-specific characterization of molecular cascades such as the one identified here for ASIC1a in dorsal striatal synapse remodeling and procedural motor learning would no doubt provide novel mechanic insights into neuronal signaling during the physiological processes. The striatum represents the main input/output structure of the basal ganglia and plays a critical role in motor and cognitive functions (24). The identification of the ASIC1a-CaMKII-ERK signaling therefore adds to the list of pathways that regulate these functions. Pathologically, synaptic dysfunction in the striatum and its related circuits are associated with neurological disorders (5, 57) including Parkinson’s disease and Huntington’s disease, as well as psychiatric disorders such as obsessive-compulsive disorder. Incidentally, previous studies also implicated ASIC1a in the development of these neurodegenerative diseases (58, 59). Therefore, our demonstrations that Asic1a KO mice performed more poorly than the WT littermates in several motor-related behavioral tests and the deficiency was rescued by expressing ASIC1a or CaMKII in striata of adult mutant mice, together with the elucidation of the underlying mechanism for ASIC1a in striatal synaptic remodeling, raise a strong possibility that targeting ASIC1a in the striatum represents a viable therapeutic strategy to treat these striatum-related neurological disorders.



Animal procedures were carried out in accordance with the guidelines for the Care and Use of Laboratory Animals of Shanghai Jiao Tong University School of Medicine and approved by the Institutional Animal Care and Use Committee (Department of Laboratory Animal Science, Shanghai Jiao Tong University School of Medicine) (policy number DLAS-MP-ANIM. 01–05). All efforts were made to minimize animal suffering and to reduce the number of animals used. Mice were kept in a standard 12-hour light/dark cycle (lights on at 7:00 a.m.) at 21°C and 50 to 60% humidity and had access to food and water ad libitum except during tests. Asic1a+/+ and Asic1a–/– littermates (male; 12 to 16 weeks old) were derived from the Asic1a+/– intercrosses. Care of animals and experimental procedures were approved by the Animal Care and Use Committee of Shanghai Jiao Tong University School of Medicine. Mice were used in a randomized order during experiments, and the investigators were blind to the genotype. For behavioral tests, mice were acclimated to the behavior rearing room for at least 1 week and were habituated to the behavior testing room at least 1 hour before the test. All behavioral tests were performed during the light cycle.

Open field test

Both Asic1a+/+ and Asic1a–/– mice were placed in the center of a square Plexiglas open field apparatus (40 cm × 40 cm × 35 cm) and allowed to freely explore for 2 hours. Total distance traveled was quantified using the EthoVision video tracking system (Noldus Information Technology).

Footprint test

The footprint test was performed following the protocol described previously (32), with minor modifications. To obtain footprints, mouse paws were coated with nontoxic paint and the animal was allowed to walk along a narrow, paper-covered corridor, leaving a track of footprints. Once the footprints dried, measurements were taken from the prints manually. The stride length was measured as the average distance of forward movement between each stride. The base width was measured as the average distance between left and right front or hind footprints. The overlap between forepaw and hindpaw placement was measured as the distance between the front and hind footprints on the same side, which was used as a measure of the accuracy of foot placement and the uniformity of step alternation.

Rotarod test

A commercial accelerating rotarod (Ugo Basile) was used to assess motor performance. Both accelerating and incremental fixed-speed protocols were used. For the accelerating rotarod test, the mouse was first habituated to low rotation (4 rpm) for 30 s and then subjected to two trials of acceleration to 40 rpm within 5, 3, or 1 min (acceleration rate was 0.12, 0.2, and 0.6 rpm/s, respectively), with a 30-min interval between the consecutive tests to reduce stress and fatigue. For the incremental fixed-speed rotarod protocol, the mouse was placed on the rod and tested at the maximal speed of 40 rpm until it fell off or the maximal cutoff time (5 min) was reached. Mice were tested three times per day with a 30-min interval between consecutive trials. The time taken for the mouse to fall from the rotating rod was recorded.

Beam walking test

The beam walking test was performed based on the previous study (32), with minor modifications. Briefly, suspended wood beams of 1 m in length were used. Mice were first trained to traverse a medium (12 mm wide) square beam in two consecutive days (three trials per day). After this preconditioning training, most mice typically could traverse the 12-mm beam in 20 s. Then, the trained mice were tested on six beams with different shapes and diameters in 2 days, with round beams with diameters of 28, 17, and 11 mm on the first day and square beams with widths of 28, 12, and 5 mm on the second day. Two test trials were performed on each beam with a 10-s interval, and the average time the mouse spent to cross the middle section (80 cm in length) of the beam was recorded for analysis. The cutoff time was set to 60 s. Because most mice could not traverse the 5-mm square beam, the distance they walked on this beam was further tested and recorded for five consecutive days to evaluate the motor learning skill. The inclined beam was placed in a 40° angle, either upward or downward, above a cushion that served to protect animals that fell.

Drugs, antibodies, and solutions

All drugs and chemicals were purchased from Sigma-Aldrich, unless otherwise specified. The lysis buffer used for protein extraction contained 150 mM NaCl, 1% Triton X-100, 1 mM EDTA, 3 mM NaF, 1 mM β-glycerophosphate, 1 mM Na3VO4, 10% glycerol, and 20 mM tris-Cl (pH 7.4), supplemented with protease inhibitors and phosphatase inhibitors. The solution for acidic treatment of cultured striatal neurons contained 150 mM NaCl, 5 mM KCl, 1 mM MgCl2, 2 mM CaCl2, 10 mM glucose, and 10 mM Hepes (pH 6.0). Antibodies used in this study were as follows: ASIC1a (1:500; catalog no. sc-13905, Santa Cruz Biotechnology), p-CaMKII (1:500; catalog no. sc-12886, Santa Cruz Biotechnology), CaMKII (1:1000; catalog no. 3362, Cell Signaling Technology), GluN1 (1:1000; catalog no. PPS011B, R&D Systems), GluN2A (1:2000; catalog no. 07-632, Millipore), GluN2B (1:500; catalog no. MAB5220, Millipore), GluA1 (1:2000; catalog no. 3861-1, Epitomics), GluA2 (1:1000; catalog no. 3520-1, Epitomics), PSD95 (1:1000; catalog no. 2366-1, Epitomics), PICK1 (1:500; catalog no. sc-9539, Santa Cruz Biotechnology), p-ERK (1:1000; catalog no. 9106, Cell Signaling Technology), ERK (1:1000; catalog no. 9107, Cell Signaling Technology), p-P38 (1:1000; catalog no. 4511, Cell Signaling Technology), P38 (1:1000; catalog no. 8690, Cell Signaling Technology), p-JNK (1:1000; catalog no. 4668, Cell Signaling Technology), JNK (1:1000; catalog no. 9252, Cell Signaling Technology), GFP (1:1000; catalog no. sc-8334, Santa Cruz Biotechnology), GAPDH (1:1000; catalog no. KC-5G4, KangChen), β-actin (1:2000; catalog no. MAB1501, Chemicon), and α-tubulin (1:2000; catalog no. T8203, Sigma-Aldrich).

Real-time reverse transcription PCR

Striata, hippocampi, and cortices of Asic1a+/+ and Asic1a–/– mice were dissected, and total RNA was extracted using TRIzol reagent (Invitrogen). Four micrograms of total RNA was used as template for complementary DNA (cDNA) synthesis and amplification with the SuperScript III First-Strand Synthesis System (Invitrogen) according to the manufacturer’s instructions. The cDNA was diluted to an equal concentration of 100 ng/μl, and 100 ng of which was used for further PCR amplification. Real-time PCR was processed by the SYBR Premix Ex Taq kit (Takara) using the ABI PRISM 7000 Sequence Detection System (Applied Biosystems) with the following amplification conditions: 95°C for 5 min; 40 cycles of 95°C for 15 s, 60°C for 15 s, and 72°C for 31 s. The primers used were as follows: Asic1a, 5′-CACCTTCCCTGCCGTCACTC-3′ (forward) and 5′-GCCCTGCTCTGTCGTAGAACTCA-3′ (reverse); Asic2a, 5′-TCAGGAGCTAGAGTTTGAATGGG-3′ (forward) and 5′-AGAGTAGAGGTGTTGGCGAAGAT-3′ (reverse); Gapdh, 5′-TGTGATGGGTGTGAACCACGAGAA-3′ (forward) and 5′-CTGTGGTCATGAGCCCTTCCACAA-3′ (reverse).

Cell culture

Primary cultures of mouse striatal neurons were prepared as described previously (60). Briefly, postnatal day 0 C57BL/6 mice were anesthetized with halothane, and their brains were removed rapidly. The striata were dissected in ice-cold Ca2+- and Mg2+-free d-Hanks’ solution and then incubated in 0.05% trypsin at 37°C for 10 min. Cells were then dissociated by trituration with a fire-polished Pasteur pipette and plated in poly-d-lysine–coated 35-mm culture dishes (Corning) at a density of 1 × 106 cells per dish. Cultures were maintained in Neurobasal medium containing 2% B27 and 1% GlutaMAX supplements (Invitrogen) at 37°C for 12 days before experiments. Glial cell growth was suppressed by the addition of 5-fluoro-2-deoxyuridine (5 μg/ml) and uridine (12.5 μg/ml).

HEK-293 cells were cultured at 37°C in a humidified atmosphere of 5% CO2 and 95% air. The cells were maintained in Dulbecco’s modified Eagle’s medium supplemented with 1 mM l-glutamine, 10% fetal bovine serum, penicillin (50 U/ml), and streptomycin (50 μg/ml) (all from Invitrogen).

Preparation of protein samples

On culture day 12, striatal neurons were rinsed and incubated with pH 6.0 or pH 7.4 solutions. After 2 min at 37°C, cells were washed with phosphate-buffered saline and lysed in the lysis buffer. The resuspended lysates were incubated on ice for 30 min and centrifuged at 13,000g for 15 min at 4°C. The supernatants were collected for Western blotting analysis. For PcTX1 or KN93 blockage experiments, 20 nM PcTX1 (Peptide Institute) or 10 μM KN93 was first added into the culture medium 30 min before acidic treatment and then co-applied with the pH 6.0 solution.

To prepare protein samples from brain tissues, mice were killed by cervical dislocation and their cortices, hippocampi, or striata were dissected, homogenized, and lysed on ice for 30 min. Then, the lysates were centrifuged at 13,000g for 15 min at 4°C, and the supernatants were collected for further examination.

Preparation of PSD fraction

The purification of PSD fraction was performed on the basis of previous studies (61, 62), with modifications. Briefly, striatal tissue samples combined from four mice were homogenized in a buffer containing 1 mM MgCl2, 0.5 mM CaCl2, and 5 mM Hepes (pH 7.4) in the presence of protease inhibitors. The homogenate was centrifuged at 1400g for 10 min at 4°C. The resulting supernatant (S1) was saved, and the pellet was resuspended and centrifuged at 700g, resulting in the supernatant (S1′), which was then combined with S1 and centrifuged at 13,800g for 10 min at 4°C to obtain a crude membrane fraction in the pellet (P2 fraction). The P2 fraction was resuspended in 0.32 M sucrose, loaded onto a discontinuous sucrose gradient (from top, 0.85 M/1.0 M/1.2 M = 3 ml/3 ml/3 ml), and then centrifuged at 82,500g for 2 hours at 4°C using an SW 41 rotor (Beckman Coulter). The synaptosome fraction between 1 and 1.2 M sucrose was collected with a syringe needle and resuspended in a buffer containing 6 mM tris (pH 8.1) and 0.5% Triton X-100.

After 15-min treatment by Triton X-100 at 4°C, the suspension was centrifuged at 201,800g using a Type 90 Ti rotor (Beckman Coulter) for 1 hour at 4°C, and the final pellet (PSD) was resuspended with a buffer containing 0.2% SDS and protease inhibitors. The “One-Triton” PSD was used because of the limit in the amount of starting materials. PSD proteins were highly enriched in our preparation, in which presynaptic components, indicated by synaptophysin, were largely absent.

Western blotting

Protein samples from cell pellet, mouse brain, or purified PSD were separated by SDS–polyacrylamide gel electrophoresis and transferred to polyvinylidene difluoride filters. The filters were incubated overnight at 4°C with appropriate antibodies. After washing the primary antibodies, secondary antibodies conjugated with horseradish peroxidase were added to the filters, washed, and then visualized in enhanced chemiluminescence solution. The visualization was performed via the ImageQuant LAS 4000 mini Molecular Imaging System (GE Healthcare Life Sciences), and the ImageJ software (National Institutes of Health) was used for the analysis of optic density. The experiments were repeated for three to six times per group.

Golgi-Cox staining

Golgi-Cox staining was performed using an FD Rapid GolgiStain Kit (FD NeuroTechnologies). Briefly, unfixed mouse brain was rapidly removed, washed in distilled water, and immersed in the impregnation solutions according to the manufacturer’s instructions. Coronal sections of 150 μm were cut on a Leica CM 1950 cryostat (Leica Biosystems) and mounted on gelatin-coated microscope slides. After drying at room temperature (22° to 24°C), sections were stained, rinsed, dehydrated, cleared, and covered with cover glasses with Permount mounting medium. Sections were selected between bregma 1.10 mm and 0.38 mm.

Sections were then analyzed using a Zeiss LSM 510 confocal microscope (Carl Zeiss). Confocal z-stacks of MSN dendritic segments were acquired with a 63× oil immersion objective at 0.8-μm intervals. For each genotype, a total of 50 to 60 images were captured from the dorsal striata of three to four different mice. Dendritic spine morphology (spine density, spine head width, and spine length) was analyzed with the Image-Pro Plus 6.0 software (Media Cybernetics). The measurements were made from secondary dendrites that were 50 to 70 μm distal to the cell soma.

Electron microscopy

Electron microscopy samples were prepared at the Electron Microscopy Facility of Shanghai Jiao Tong University School of Medicine, as described previously (63). Images were acquired at a ×17,500 magnification using the CM-120 system (Philips). The measurements of striatal PSD thickness and length were performed using Image-Pro Plus 6.0 software (Media Cybernetics) by an observer who was blind to the genotypes of the samples.

Brain slice preparation and patch-clamp recordings

Experiments were performed on the striatal slices, as described previously (16), with minor modifications. Briefly, after decapitation, the mouse brain was quickly removed and placed in well-oxygenated (95% O2/5% CO2, v/v) ice-cold artificial cerebrospinal fluid (aCSF) containing 125 mM NaCl, 2.5 mM KCl, 12.5 mM d-glucose, 1 mM MgCl2, 2 mM CaCl2, 1.25 mM NaH2PO4, and 25 mM NaHCO3 (pH 7.35 to 7.45). Three coronal striatal slices (300 μm thick) were obtained at the level of the corpus callosum connection with a vibratome (VT 1000S, Leica) and incubated at 30 ± 1°C in oxygenated aCSF for at least 1 hour before being transferred to a recording chamber placed on the stage of a microscope (BX51WI, Olympus). The placement of individual slices was observed using an infrared–differential interference contrast video monitor. The slices were continuously perfused with well-oxygenated aCSF at room temperature during all electrophysiological studies. EPSCs were recorded from dorsal striatal MSNs with an Axon 200B amplifier (Molecular Devices), and the stimulations were delivered with a bipolar tungsten stimulating electrode placed in the corpus callosum to stimulate the cortical glutamatergic inputs to MSNs in the dorsal striatum. AMPAR-mediated EPSCs were induced by repetitive stimulations at 0.03 Hz, with the neuron voltage-clamped at −70 mV, except where indicated otherwise. The recording pipettes (3 to 5 megohms) were filled with a solution containing 132.5 mM cesium gluconate, 17.5 mM CsCl, 2 mM MgCl2, 0.5 mM EGTA, 10 mM Hepes, 4 mM Mg–adenosine 5′-triphosphate, and 5 mM QX-314 chloride (280 to 300 mosmol, pH 7.2 with CsOH). Access resistance was 15 to 30 megohms, and only cells with a change in access resistance of <20% were included in the analysis. The NMDAR-mediated EPSCs were recorded in the presence of CNQX (20 μM) and PTX (100 μM), with the neuron voltage clamp at +40 mV. The mEPSCs were obtained at −70 mV in the presence of tetrodotoxin (0.3 μM) and PTX (100 μM) without stimulation and analyzed using MiniAnalysis (Synaptosoft Inc.).

Generation and delivery of AAV vectors

EYFP protein was fused to the N terminus of mASIC1a (GenBank accession: NM_009597.1) by using “self-cleaving” 2A peptide, which can separate the two proteins apart during protein translation (64). The coding sequence for the EYFP-2A-mASIC1a fusion protein or CaMKII (GenBank accession: NM_177407.4) was subcloned to the pAAV-MCS vector and driven by hSynI promoter for neuronal-specific expression. This construct was then packaged into AAV2/8 chimeric virus with AAV8 capsids and AAV2 ITR (inverted terminal repeat) element. Once expressed in vivo, EYFP-2A and mASIC1a should be expressed separately, with an extra proline left at the N terminus of mASIC1a (65).

For virus injection, mice at the ages of 8 to 10 weeks were anesthetized and placed in a stereotaxic frame (RWD Life Science). AAV-ASIC1a [titer, 1.13 × 1012 viral genomes (vg)/ml] or AAV-GFP (titer, 1.98 × 1012 vg/ml) viruses were injected bilaterally into the dorsal striatum. The stereotaxic coordinates according to the mouse brain atlas (66) were as follows: anteroposterior, +0.4 mm; lateral, ±2.2 mm; dorsoventral, −3.5 mm. One injection (2 μl) of AAV-ASIC1a or AAV-GFP was made on each side using a microelectrode connected with a microinjector pump (KDS 310, KD Scientific) at a rate of 0.1 μl/min. Microelectrodes were left in situ for an additional 10 min to allow the injectate to diffuse. Mice were allowed to recover for 4 weeks before behavioral analysis, and the injection sites were examined at the end of the experiments by imaging brain slices from viral-infected mice under a fluorescence microscope.

Cannula implantation and local drug injection

Mice were anesthetized with 5% chloral hydrate and then fixed on a stereotaxic apparatus (RWD Life Science). Stainless steel guide cannulas (RWD Life Science) were bilaterally implanted into the target brain areas, and the tips of cannulas were at following coordinates: anteroposterior, +0.4 mm; lateral, ±2.2 mm; dorsoventral, −3.5 mm. The cannulas were fixed to the skull using acrylic cement and two skull screws. Stainless steel obturators (33 gauges) were inserted into guide cannulas to avoid obstruction until drug infusion. Animals were allowed to recover from surgery for 2 weeks before behavioral tests. Mice were handled and habituated to the infusion procedure several days before drug injection. During drug infusion, mice were briefly head-restrained, whereas the stainless steel obturators were removed and injection cannulas (33 gauges, RWD Life Science) were inserted into guide cannulas. Injection cannulas protrude 1.00 mm from the tips of guide cannulas. Infusion cannula was connected using PE20 tubing to a microsyringe driven by a microinfusion pump (KDS 310, KD Scientific). Drugs were infused bilaterally into the target brain areas at a flow rate of 0.15 μl/min. After finishing drug injection, the injection cannulas were left in place for 5 min to allow the solution to diffuse from the cannula tip. The stainless steel obturators were subsequently reinserted into guide cannulas, and the mice returned to their home cage for 30 min before behavioral tests. PcTX1 (10 μM in aCSF, 2 μl per side; Peptide Institute) or the vehicle only (aCSF) was microinfused into the dorsal striatum. The injection sites were examined at the end of the experiments, and animals with incorrect diffusion scope were excluded from the data analysis.

Data analysis

Experimenters were blind to the genotype until all data were collected and analyzed. All statistical analyses were performed using SPSS 11.5 (SPSS Inc.). Unpaired Student’s t test, one-way ANOVA test followed by Fisher’s least significant difference test, two-way ANOVA followed by Dunnett’s T3 post hoc test, and two-sample Kolmogorov-Smirnov tests were used for comparisons, as indicated in the figure legends. All summary data are means ± SEM, and P < 0.05 was considered to be significantly different.


Fig. S1. Effects of loss of ASIC1a in mice on Asic2a mRNA expression in different brain regions.

Fig. S2. Effects of Asic1a gene ablation in mice on basal locomotor activity and motor coordination in a series of behavioral paradigms.

Fig. S3. Demonstration of Golgi staining and analysis.

Fig. S4. Input-output relationships of AMPAR- and NMDAR-mediated synaptic currents in striatal neurons from WT and Asic1a KO mice.

Fig. S5. Effects of loss of ASIC1a on the activation of p38 or JNK signaling in striata.

Fig. S6. Effects of acidosis on the phosphorylation of ERK, p38, or JNK in HEK-293 cells.

Fig. S7. Effects of pharmacological inhibition of ASIC1a in the striatum on motor learning in the beam walking test.

Fig. S8. Representative images showing expression patterns of viral-encoded proteins (represented by EYFP) after AAV-ASIC1a injection into striata of Asic1a KO mice.

Fig. S9. Effects of reexpression of ASIC1a in striata of Asic1a null mice on spine density and maturation.

Fig. S10. Effects of reexpression of ASIC1a in striata of Asic1a null mice on the performance in the inclined beam walking test.


Acknowledgments: We thank J. A. Wemmie (University of Iowa) and M. J. Welsh (Howard Hughes Medical Institute, University of Iowa) for providing Asic1a KO mice. We also thank the Electron Microscopy Facility of Shanghai Jiao Tong University School of Medicine for the preparation of the electron microscopy samples and X. Gu, L. Han, and D. Tang for the technical assistance. Funding: This study was supported by grants from the National Natural Science Foundation of China (81730095, 91632304, 81771214, 31500671, and 31230028), the National Basic Research Program of China (2014CB910300), the Shanghai Committee of Science and Technology (18QA1402500), and the NIH (NS102452). Author contributions: T.-L.X., Z.Y., and W.-G.L. conceived the project and designed the experiments. Z.Y., Y.-J.W., D.-S.L., X.-L.S., Q.J., and Y.L. performed the research. Z.Y., Y.-J.W., Y.-Z.W., D.-S.L., S.Z., N.-J.X., and W.-G.L. analyzed the data. Z.Y., Y.-Z.W., W.-G.L., M.X.Z., and T.-L.X. wrote the manuscript. All authors read and approved the final manuscript. Competing interests: The authors declare that they have no competing interests. Data and materials availability: There are no restrictions on the data or materials used in this study. All data needed to evaluate the conclusions in the paper are present in the paper or the Supplementary Materials.
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