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Huntington's disease is caused by an expanded polyglutamine repeat in huntingtin (Htt), but the pathophysiological sequence of events that trigger synaptic failure and neuronal loss are not fully understood. Alterations in NMDA-type glutamate receptors (NMDARs) have been implicated, yet it remains unclear how the Htt mutation impacts NMDAR function and direct evidence for a causative role is missing. Here we show that mutant Htt re-directs an intracellular store of juvenile NMDARs to the surface of striatal neurons by sequestering and disrupting the subcellular localization of the GluN3A subunit-specific endocytic adaptor PACSIN1. Overexpressing GluN3A in wild-type striatum mimicked the synapse loss observed in Huntington's disease mouse models, whereas genetic deletion of GluN3A prevented synapse degeneration, ameliorated motor and cognitive decline, and reduced striatal atrophy and neuronal loss in the YAC128 model. Furthermore, GluN3A deletion corrected the abnormally enhanced NMDAR currents, which have been linked to cell death in Huntington's disease and other neurodegenerative conditions. Our findings reveal an early pathogenic role of GluN3A dysregulation in Huntington's disease, and suggest that therapies targeting GluN3A or pathogenic Htt-PACSIN1 interactions might prevent or delay disease progression.
Huntington's disease (HD) is a progressive neurodegenerative disorder with severe motor, cognitive and psychiatric disturbances that is caused by expansion of a polyglutamine repeat within the N-terminal region of Htt. Mutant Htt (mHtt) accumulates as oligomeric species and aggregates throughout the neuronal soma and dendrites1-3, triggering synaptic failure and loss later followed by death of subsets of striatal and cortical neurons. Limitations in understanding the mechanisms underlying Htt toxicity, especially at early stages, have been a major obstacle to finding a treatment.
A prevalent hypothesis is that mHtt engages in aberrant protein interactions, interfering with the function of key cellular components. But Htt interacts with a large network of proteins4,5 and discerning the pathogenic interactions has proven difficult. Nevertheless, a prominent group of Htt-interactors comprises proteins involved in clathrin-mediated endocytosis6,7, pointing towards altered protein trafficking as a key pathological mechanism.
One of these interactors, PACSIN1/syndapin1, works as an endocytic adaptor for neuronal NMDARs8. NMDARs play crucial roles in remodeling and maintaining excitatory synapses and their activity is altered in striatal medium-sized spiny neurons (MSNs) of transgenic mice expressing mHtt9-13. Because MSNs are the population first affected in HD and NMDAR dysfunction can be detected from early stages, this alteration has long been thought as a pathogenic trigger14. Intriguingly, PACSIN1 targets GluN3A subunits, which prevent premature synapse plasticity and stabilization during early stages of postnatal brain development, but are down-regulated in adult brains15,16. In addition, the binding affinity of PACSIN1 for Htt depends on the length of the polyglutamine expansion17, fulfilling a key criteria for pathogenic interactions18, and PACSIN1 gain-of-function suppresses mHtt toxicity in Drosophila screens4. Thus, we hypothesized that mHtt might interfere with the endocytic removal of GluN3A-containing NMDARs by PACSIN1, leading to age-inappropriate synapse destabilization during HD pathogenesis.
Here we confirm that mHtt binds and sequesters PACSIN1 away from its normal cellular locations, causing accumulation of juvenile GluN3A-containing NMDARs at the surface of striatal neurons. We then show that GluN3A levels are abnormally elevated across mouse models of HD and in human HD striatum, and that GluN3A overexpression in mice drives degeneration of afferent synapses onto MSNs. Importantly, suppressing GluN3A reactivation in corrected the early enhancement of NMDAR currents in MSNs from YAC128 mice, prevented both early-stage and progressive dendritic spine pathology, and ameliorated later motor and cognitive decline. Our results reveal a new mechanism that mediates NMDAR dysfunction and synapse loss in HD—dysregulation of the expression of NMDARs that contain juvenile GluN3A subunits by altered endocytic trafficking, and identify a potential safe target for pharmacological therapy.
We began by verifying the interaction between Htt and PACSIN14,17 with coimmunoprecipitation assays on striatal lysates from wild-type and YAC128 mice, which express full-length human HTT with 128 CAG repeats19. Although PACSIN1 interacted with both Htt variants, the interaction was stronger in YAC128 striatum (2.23 ± 0.3 fold increase in PACSIN1 bound to Htt relative to wild-type, P = 0.005, Fig. 1a). Coimmunoprecipitation assays from HEK293 cells co-transfected with PACSIN1 and GFP-tagged Htt exon-1 fragments that spanned the proline-rich domain that binds PACSIN117 and a normal or expanded polyglutamine tract (Httex117Q-GFP and Httex146Q-GFP) confirmed the polyglutamine dependence of the interaction, and additionally showed that exon-1 is sufficient for PACSIN1 binding (Fig. 1b,c). Specificity for Htt rather than the polyglutamine tract was demonstrated by experiments where PACSIN1 did not interact with either normal or expanded ataxin1, a polyglutamine repeat protein involved in spinocerebellar ataxia (Fig. 1c).
We next assessed whether binding to mHtt interferes with PACSIN1 localization or function in a cellular model of HD20. Cultured rat striatal neurons were transfected with Httex1 fragments, and PACSIN1 localization evaluated by immunofluorescence. Httex146Q-GFP formed detectable intracellular aggregates in ~ 25% of neurons within 24–48 h, whereas Httex117Q-GFP exhibited diffuse expression (Supplementary Fig. 1). PACSIN1 immunostaining was punctate and distributed evenly in somatodendritic compartments of untransfected and Httex117Q-GFP-expressing neurons (Fig. 1d), but was enriched in somatodendritic Htt aggregates in neurons expressing Httex146Q-GFP (Fig. 1d,e, Supplementary Fig. 1). A quantitative colocalization analysis21 yielded highly significant intensity correlation coefficient (ICQ) values for Httex146Q-GFP and PACSIN1 (0.26 ± 0.02, Psign test < 0.005). Lower ICQ values were obtained for Httex117Q-GFP (0.065 ± 0.01), consistent with the weaker binding in biochemical assays. Other endocytic proteins were not enriched in aggregates (Fig. 1f), ruling out non-specific sequestration.
Redistribution into mHttex1 aggregates was associated with reductions of endogenous PACSIN1 levels within their normal postsynaptic location, as quantified by decreases in dendrite-to-soma fluorescence intensity ratios and consistent with dendritic depletion of PACSIN1 in humans with HD17 (Supplementary Fig. 2a,b). Total PACSIN1 levels were unchanged (Supplementary Fig. 2c), indicating that reduced dendritic availability reflects altered subcellular distribution.
Because PACSIN1 controls the endocytic removal of GluN3A-containing NMDARs8, the stronger binding and/or subcellular sequestration by mHtt might be expected to interfere with ongoing endocytosis and affect plasma membrane expression. To test this, we co-transfected striatal neurons with HA-tagged GluN3A and Httex1 variants and measured surface HA-GluN3A levels 48 h later using fluorescence-based antibody uptake assays. Httex146Q-GFP expression resulted in elevated surface GluN3A (Fig. 2a,b) and concomitant increases in surface levels of the obligatory NMDAR subunit GluN1 (Fig. 2c), likely reflecting GluN1 assembly with GluN3A subunits which is required for their plasma membrane transport and localization22. Notably, increases in GluN3A were observed both in neurons with diffuse and aggregated Httex146Q-GFP expression (Fig. 2b) suggesting that aggregation is not required and that binding to soluble mHtt species may be sufficient to alter PACSIN1 function. Supporting specificity for mHtt, polyglutamine expansions in ataxin1 did not alter surface GluN3A (Fig. 2d). Because alterations in bulk endocytic recycling have been reported after mHtt expression in PC12 cells23, we analyzed transferrin uptake but found no alterations in neurons transfected with Httex146Q-GFP by 48 h, time when GluN3A levels were significantly elevated (Supplementary Fig. 2d,e). Nor did Httex146Q-GFP affect the surface levels of the AMPA receptor subunit GluA1 (Fig. 2e), which are highly reliant on endocytic recycling24. These results indicate that GluN3A accumulation is not due to a general impairment in neuronal endocytosis.
Knockdown of PACSIN1 using an anti-PACSIN1 short hairpin RNA (sh-PACSIN1, Supplementary Fig. 3) also increased surface GluN3A levels (Fig. 2f,g), demonstrating that PACSIN1 depletion can account for the mHtt-induced increase in surface GluN3A. We then asked whether exogenous supply of PACSIN1 could counteract the effect of mHtt. Indeed, co-transfection of PACSIN1 rescued the surface accumulation of GluN3A subunits, which returned to control levels in Httex146Q-GFP expressing neurons (Fig. 2a,h). These results show that loss of PACSIN1 function, likely due to mHtt binding, increases the surface expression of GluN3A-containing NMDARs in cultured striatal neurons.
GluN3A is highly expressed in the brain during early postnatal and juvenile stages (P8-P15 in mice and first years of life in humans) but expression declines afterwards25,26. Quantitative immunoblot analysis showed that GluN3A undergoes a sharper down-regulation in mouse striatum relative to other brain regions, with expression largely reduced into adulthood (Supplementary Fig. 4). In contrast, no developmental changes were observed in GluN2B, the predominant NMDAR subunit type in striatum (Supplementary Fig. 4). These results support a physiological role for GluN3A down-regulation in striatal NMDAR function and predict significant effects of disruption. To evaluate whether GluN3A dysregulation occurs in humans, we analyzed postmortem brain tissue from controls and HD individuals. GluN3A levels in control human putamen, a sub-region of the striatum, were low, but a significant increase in GluN3A was detected in affected individuals (Fig. 3a).
We next used the R6/1 and YAC128 mouse models of HD to evaluate the timing of GluN3A changes and association with pathogenic events. R6/1 mice, which express N-terminally truncated human HTT carrying 115 CAG repeats27, displayed a progressive increase in striatal GluN3A levels relative to wild-type starting around 12–16 weeks of age (Supplementary Fig. 5a). Because PACSIN1 is enriched at synapses where it mediates GluN3A removal8,28, we biochemically isolated synaptic plasma membranes (SPMs) to determine whether increased GluN3A levels reflect increased residency at synaptic compartments associated with changes in PACSIN1 localization. Indeed, a large increase in GluN3A levels (~2.5 fold) was detected in SPMs from 20 week-old R6/1 mice, matching a reduction in the synaptic abundance of PACSIN1 (Fig. 3b,c).
Similar results were obtained in 3 month-old YAC128 mice (Supplementary Fig. 5b, Fig. 3d-e). Increased GluN3A levels in striatal SPMs were paralleled by increases in GluN1 and GluN2B, suggesting that GluN3A forms heteromultimeric NMDAR complexes including these subunits (Fig. 3d), and were associated with reduced synaptic PACSIN1 levels (Fig. 3e). No changes in GluN2A or GluA2/3, presynaptic proteins such as synaptophysin, or the trafficking regulators Rab4 or Rab11 were detected (Fig. 3d,e). Further fractionation of striatal synaptic membranes showed increased GluN3A in both detergent-soluble (non-PSD) and insoluble (PSD) fractions, consistent with the subsynaptic localization pattern characteristic of GluN3A8 (Supplementary Fig. 6). Overall PACSIN1 protein expression was unchanged (0.93 ± 0.03 of wild-type, P = 0.19), corroborating our culture data and further indicating that lower synaptic levels of PACSIN1 reflect subcellular redistribution. At this early stage, GluN3A levels were not altered in brain regions affected later in HD such as cortex and hippocampus (Supplementary Fig. 5b). Immunolabeling further showed that GluN3A increases mainly occurred in DARPP-32 labeled MSNs (Supplementary Fig. 5c,d).
To test whether elevated GluN3A reflects a defect in endocytosis as predicted if subcellular sequestration of PACSIN1 by mHtt is the mechanism, striatal slices from control and YAC128 mice were incubated with the membrane-impermeable cross-linker BS3. This reagent cross-links surface receptors to form high molecular weight complexes that can be separated from unlinked intracellular receptors by electrophoresis, allowing a quantification of the internal receptor pool (Fig. 3f). These experiments revealed a significantly reduced fraction of intracellular GluN3A in YAC128 striatum but, again, no changes in GluA2/3 (Fig. 3f,g). We additionally looked for contributions of altered transcription but found no significant changes in GluN3A mRNA in either R6/1 or YAC128 striatum at times when synaptic and surface levels of GluN3A protein were increased by ~2 fold (Supplementary Fig. 5e). These results confirm that GluN3A is redistributed to the plasma membrane and synaptic compartments in mouse models of HD. They further show that striatal GluN3A dysregulation begins before or around the onset of motor or cognitive symptoms29-32, long preceding neuronal loss, and may thus be an early disease mechanism in HD pathophysiology. Increases in GluN3A preceding symptom onset were also observed in a third HD model, the HdhQ111 knock-in mice, where the phenotype is much delayed33 (Supplementary Fig. 5f).
Alterations in NMDAR function have been reported in MSNs of YAC128 mice by 1 month. The defect involves long-lasting responses to glutamate release and has been attributed to increased activation of extrasynaptic NMDARs13. Because: i) a distinguishing feature of GluN3A-containing receptors is their peri- and extrasynaptic localization8,34, and ii) GluN3A levels were increased early in synaptosomal subfractions from YAC128 striatum that are enriched in extrasynaptic plasma membrane receptors (non-PSD), we asked whether the electrophysiological defect could be explained by the elevated GluN3A levels. We crossed YAC128 mice with GluN3A−/− mice and compared NMDAR responses after intense paired pulse afferent stimulation in wild-type, YAC128 and YAC128 mice lacking GluN3A following the protocol by Milnerwood et al13. Isolated NMDAR responses were obtained by calculating the difference between responses recorded in the absence and then presence of the NMDAR blocker APV. No changes were detected in the AMPAR-mediated component obtained in APV, but large differences were detected in the decay kinetics of the isolated NMDAR-component (P < 0.005, YAC128 vs wild-type, Fig 3h,i), confirming previous observations13. Differences were largely abolished in YAC128 mice lacking GluN3A (Fig 3h,i), indicating that GluN3A is required for the mHtt-induced enhancement in NMDAR currents and consistent with the possibility that aberrant expression underlies the earliest events of HD pathophysiology.
Dendritic dystrophy and loss of glutamatergic spiny synapses are early degenerative changes in HD7,35,36 that might underlie symptom onset. Because enhancement of GluN3A function in the hippocampus drives synapse elimination16, we asked whether aberrant GluN3A expression could contribute to the synaptic pathology. We first used Golgi impregnation techniques to measure dendritic spines, sites where excitatory synapses form, in YAC128 mice at times when GluN3A levels were elevated (Fig. 4a). Spine numbers were decreased in dendrites of YAC128 MSNs by 3 months (13–15% decrease vs wild-type) and the decay worsened with age (Supplementary Fig. 7).
To test whether abnormal accumulation of GluN3A in adult MSNs could explain the progressive spine loss induced by mHtt, we measured spine densities in transgenic mice that overexpress GluN3A into adulthood (dt GFP-GluN3A16,34). MSNs from dt GFP-GluN3A displayed a pattern of spine loss remarkably similar to YAC128 mice (~11% decrease vs single-transgenic littermates by 3 months with larger spine losses at later stages, Fig. 4b,c).
We then reasoned that if the spine loss in MSNs was caused by enhanced GluN3A function, it should be prevented by lowering GluN3A levels. To test this, we re-examined spine densities in wild-type and YAC128 mice in the presence and absence of GluN3A. Lack of GluN3A prevented both the early reductions in spine numbers in YAC128 mice, and the progressive synaptic disconnection detected as a larger loss of MSN spines at later disease stages (Fig. 4d,e). In contrast, spine densities were not altered in MSNs from adult GluN3A−/− mice (Fig. 4d,e), indicating that blocking GluN3A expression targets the synaptic pathology in HD without affecting basal spine levels in controls. This result differs from the increased spine density reported in GluN3A−/− cortical neurons15 and likely reflects the lower GluN3A levels retained in adult striatum (Supplementary Fig. 4).
At higher resolution, electron microscopy revealed a marked loss of asymmetric glutamatergic synapses onto MSNs of YAC128 mice, which was likewise rescued by GluN3A deletion (Fig. 4f,g). In addition, postsynaptic densities were smaller in YAC128 mice (Fig. 4g), consistent with earlier studies where GluN3A overexpression decreased synapse size16, and the deficit was also rescued by suppressing GluN3A (Fig. 4g). These results show that diminished synaptic connectivity in MSNs from YAC128 mice begins at early stages, matching the timing of GluN3A dysregulation, and can be rescued by deleting GluN3A or replicated by transgenically overexpressing GluN3A.
We next assessed the impact of GluN3A on HD-like symptoms using striatum-dependent motor and cognitive tasks. YAC128 mice showed impaired motor learning by 10 months, as indicated by increased failures when learning a fixed-speed rotarod task, and impaired coordination as represented by a shorter latency to fall from an accelerating rotarod. Both deficits were improved in YAC128 mice lacking GluN3A (Fig. 5a,b). A vertical pole test, another measure of motor coordination, showed that YAC128 mice spent less time in the pole than wild-type, an effect that was again rescued by GluN3A deletion (Fig. 5c). Nevertheless, the hypokinetic phenotype exhibited by YAC128 mice in an open-field was still apparent in YAC128 mice lacking GluN3A (Fig. 5d). Neither weight gain nor loss of muscle tone was rescued by GluN3A deletion (Supplementary Fig. 8), ruling out that improvements in coordination resulted from reversal of these parameters.
Cognitive function was evaluated using a swimming T-maze task37. YAC128 mice exhibited a pronounced impairment during the learning phase, whereas YAC128 mice lacking GluN3A performed as well as wild-type (Fig. 5e,g). On day 4, the platform was switched to the opposite arm of the T-maze to probe reversal (strategy-shifting) learning. YAC128 required longer than wild-type to reach the new platform location in all trials, and the defect was rescued by GluN3A deletion (Fig. 5f,g).
To investigate whether abnormally increased GluN3A contributes to later striatal atrophy and MSN degeneration, we performed stereological analyses in 16 month-old mice, age when neuronal death can be reproducibly detected in the YAC128 model19. Suppressing GluN3A expression in YAC128 mice rescued striatal atrophy and reductions in NeuN- and DARPP-32-positive neurons (Fig. 6a,b,d). It also normalized DARPP-32 levels in surviving MSNs (Fig. 6c,d); DARPP-32 is a key component of MSN dopamine signaling cascades which is down-regulated in HD mouse models from early stages38.
Analogous analyses in control and dt GFP-GluN3A transgenic mice showed that GluN3A overexpression per se does not cause striatal atrophy or neuronal loss by 16 months of age (Fig. 6e,f), but mimics neuropathological features of HD including reductions in the size of MSNs39 (Fig. 6g) and in striatal DARPP-32 levels40 (single-transgenic, 100 ± 16; dt GFP-GluN3A, 61 ± 5, P < 0.05, Fig. 6h). Finally, we conducted experiments in a rat corticostriatal slice model41 to test whether increased GluN3A levels influence the susceptibility of MSNs to mHtt toxicity. Slice explants were co-transfected with expanded Httex1 (Httex173Q-GFP) along with GluN3A, and MSN degeneration was assessed by co-expression of yellow fluorescent protein. The number of surviving MSNs was decreased in brain slices co-transfected with mHtt and GluN3A relative to slices transfected with mHtt alone (Fig. 6i). Thus, increased GluN3A recapitulates features of HD neuronal dysfunction such as MSN shrinkage and disturbed signaling, and renders MSNs more vulnerable to mHtt toxicity.
We identify a novel early-stage disease mechanism that underlies NMDAR dysfunction and synapse loss in HD: subcellular sequestration of the endocytic adaptor PACSIN1 by mHtt leaves an overabundance of juvenile NMDARs containing GluN3A subunits in the plasma membrane and postsynaptic sites of striatal neurons. The new mechanism provides a missing link between two major phenomena previously known to be impaired by Htt mutations: defects in protein trafficking and impaired NMDAR transmission. GluN3A dysregulation targeted MSNs, the vulnerable population in HD striatum, and was an early feature across HD mouse models. Knocking-out GluN3A in HD mice corrected a full sequence of early-to-late pathophysiological events, demonstrating a critical role for GluN3A in disease pathogenesis in vivo: normalized NMDAR currents; fully prevented synapse degeneration; rescued motor and cognitive decline; and reduced striatal cell death.
The work extends previous findings pointing towards juvenile GluN3A subunits as a synapse destabilizing or pruning factor16, and demonstrates the pathogenic effects of aberrant reactivation in the adult brain of a pruning mechanism normally restricted to developmental stages. In the normal brain, PACSIN1-mediated retrieval of GluN3A from synapses contributes to receptor down-regulation and is thought to provide a critical signal for synaptic plasticity and growth and for robust information storage, by allowing the replacement of juvenile with mature NMDAR subtypes8,16. Factors that inhibit PACSIN1 function, such as dominant-negative variants of PACSIN18 or mHtt, increase surface and synaptic GluN3A yielding a higher proportion of small, immature synapses. Although the precise mechanisms by which mHtt alters PACSIN1 distribution remain unclear, our observations of reduced synaptic PACSIN1 in young YAC128 mice, months before aggregates appear42, and of surface accumulation of GluN3A in striatal neurons with no detectable Htt aggregates argue against a requirement for aggregation. This fits with current concepts linking soluble mHtt species, rather than aggregates, to cytotoxicity20,43,44. But PACSIN1 redistribution into microaggregates not visible by light microscopy cannot be ruled out, and the question of whether PACSIN1 binds to particular conformations of mHtt remains open.
Regarding stoichiometry of GluN3A-containing NMDARs in HD striatum, our biochemical fractionation results indicate the presence of GluN1 and GluN2B subunits. This is consistent with: biochemical evidence for assembly of GluN3A and GluN2B subunits in striatal tissue34; reports attributing the enhanced NMDAR responses in YAC128 mice to GluN2B-containing NMDARs13,45; and our finding that suppressing GluN3A corrects the alteration. GluN3A-containing subtypes are less anchored to postsynaptic densities than conventional NMDARs (GluN1/GluN2 heteromers) and GluN3A overexpression has been shown to impair the postsynaptic stabilization of GluN2B-containing NMDARs34, which might facilitate their diffusion towards extrasynaptic sites, providing a testable cell biological mechanism for the increased extrasynaptic GluN2B-mediated currents in YAC models13. Along with synapse loss and enhanced extrasynaptic NMDARs, elevated GluN3A could explain features of altered glutamatergic transmission that have been observed in presymptomatic HD mice and cannot be accounted for by the increased GluN2B-mediated currents without considering the presence of additional subunits in the receptor complex. These include anomalously reduced long-term potentiation46-48 and decreased magnesium sensitivity of NMDAR currents11,49; both have been reported in GluN3A overexpressing mice16.
An imbalance between synaptic and extrasynaptic NMDAR activity is thought to be critical for neurodegeneration because the two receptor populations signal to cell survival or death pathways respectively50, and increased extrasynaptic activity has been linked to cell death in HD13,43. Enhanced GluN3A expression could contribute to cell death by driving or aggravating this imbalance in two ways. First, it might trigger activation of death pathways by increasing extrasynaptic NMDAR localization. Second, MSNs receive dense glutamatergic input from cortical axons, and the synaptic disconnection driven by GluN3A could inhibit pro-survival signaling pathways coupled to synaptic NMDAR activation50,51. But GluN3A-containing subtypes flux less Ca2+ than other NMDAR subtypes22 and data from our own lab support a neuroprotective action of GluN3A overexpression in acute excitotoxic cell death assays34,52, which might explain the paradoxical resistance to acute excitotoxic insults observed in HD models53,54. Nonetheless, aberrant GluN3A expression over the much longer time course relevant to HD (i.e., decades versus minutes) could have deleterious effects due to inhibition of synaptotrophic NMDAR activity and/or chronically disturbed signaling by extrasynaptic NMDARs activated by ambient glutamate.
In summary, our work uncovers a previously unappreciated role for GluN3A dysregulation in HD and provides a rationale for the use of therapies targeting GluN3A or PACSIN1 early in the course of the disease to ameliorate cognitive or motor problems and/or halt disease progression. An advantage of targeting GluN3A (versus other NMDAR subunits) is that it is mostly lacking in adult brains, and would allow to selectively block a pathological trait without hampering normal synaptic function.
Methods and any associated references are available in the online version of the paper
Expression plasmids encoding Httex1 or ataxin variants fused to green fluorescent protein (GFP) were described previously20,55. HA-GluN1-1a was from T. Yamamoto and HA-GluA1 was from M. Ehlers. HA-GluN3A was obtained by replacing the GFP tag in GFP-GluN3A22. Myc-tagged and mCherry-tagged rat PACSIN1 were cloned in pRK5. A 59-bp encoding a 19-bp-long small hairpin RNA (shRNA) specific for rat and mouse PACSIN1 (sh-PACSIN1 target sequence: CCATAGAGTTCCAGACATA) or a scrambled control (sh-SCR: GATCAGTGCCAACTTAACA) were cloned into pSuper-GFP vector (OligoEngine, Seattle, WA).
Primary striatal neuronal cultures were prepared from E19 rat embryos8 and transfected at DIV8 using calcium phosphate. Experiments were performed 24–48 h later.
For immunofluorescence analysis of colocalization of endocytic proteins with Htt aggregates, Httex1-GFP-transfected neurons were fixed with 4% sucrose, 4% paraformaldehyde in phosphate-buffered saline (PBS), permeabilized with 0.1% saponin or TX-100, 1% bovine serum albumin and 2% normal goat serum in PBS, and incubated with primary antibodies followed by incubation with Cy3-conjugated secondary antibody (1:1000, Jackson ImmunoResearch). Mouse antibodies to PACSIN1 (1:200, 611811), alpha-adaptin (1:500, 610501) and EEA1 (1:250, E41120) from BD Transduction Labs, and to TfR (1:500, Zymed, 13-6800) were used. The specificity of the PACSIN1 antibody was tested by immunofluorescence in neurons transfected with two different PACSIN1 shRNAs. Images of Httex1-GFP transfected neurons were captured using a Coolsnap HQ camera (Photometrics) mounted on a Zeiss Axiovert 200 or a Zeiss LSM 510 confocal microscope using a 63×/1.4NA Plan-Apochromat objective.
Colocalization was quantified using a custom-made ImageJ plugin to apply the Intensity Correlation Analysis of Li et al21. Comparisons were made from pairs of single image planes through the soma or individual dendritic fragments after background subtraction in each channel. Regions of interest (ROIs) containing aggregates were drawn in somatic regions of mHtt-transfected neurons and fluorescence pixel intensities were quantified in matched ROIs in the green (GFP-Httex1) and red (Cy3, PACSIN1) channels. The difference from the mean for each pixel intensity (Ri-Rmean and Gi-Gmean where R and G are the red and green pixel intensities), and the product of the differences [PDM = (Ri-Rmean) × (Gi-Gmean)] were calculated. PDMs are positive when both and red pixel intensities vary in synchrony (i.e. both red and pixel intensities are either above or below their respective means). An intensity correlation quotient (ICQ) was then calculated that is equal to the ratio of the number of positive PDM values to the total number of pixels. The ICQ ranges between −0.5 to 0.5 (with random staining ICQ =0; dependent staining 0 < ICQ < 0.5; segregated staining 0 > ICQ > −0.5). Image sets were analyzed only in image regions free of pixel saturation, and pixel staining pairs with double 0-level intensity values were removed. Enrichment of PACSIN1 in aggregates was additionally quantified by comparing PACSIN1 intensity levels in aggregates versus a non-aggregate ROI (Supplementary Fig. 1).
To quantify dendritic PACSIN1 expression in Httex1-GFP transfected neurons, all individual dendritic segments in the focus plane were outlined and their average pixel intensities normalized to somatic intensity to obtain a dendrite-to-soma ratio for each transfected and untransfected neuron in the same field. For total PACSIN1 measurements, Httex1-GFP transfected neurons were outlined in the green channel, transferred to the red channel and average intensities measured after background subtraction were compared to untransfected neurons in the same field
Immunofluorescent analysis of surface receptor expression was as described8. Briefly, live transfected striatal neurons were incubated with an HA antibody (1:200, Covance, MMS-101P) for 30 min at 37 °C, fixed, blocked and surface HA-receptors detected with Cy3-conjugated secondary antibody. Neurons were then permeabilized, internal HA-receptors labeled with HA antibody and detected with Alexa647 secondary antibody (1:200, Invitrogen). Wide-field fluorescence images were acquired with a Zeiss 4× or 63× objective and a CoolSnap CCD camera and analyzed with Metamorph. For quantification, neurons were outlined in the GFP channel and outlines were transferred to the red and far red channel images to obtain average intensity measurements. Surface-to-internal ratios were calculated by dividing Cy3 intensity values by Alexa647 intensity values.
Striatal neurons were incubated with Alexa568-conjugated transferrin (Tf, 50 μg ml−1) in serum-free media for 10 min at 37 °C. Cells were then washed with serum-free medium at 10 °C, and incubated with holotransferrin (500 μg ml−1) in conditioned media to exchange the surface bound transferrin and selectively monitor the endocytosed fraction. After washing, neurons were fixed and remaining intracellular Alexa-Tf imaged. For quantification, fluorescent intensities of Alexa-Tf within three to four 50 μm dendritic segments of Httex1-GFP transfected neurons were measured for each neuron analyzed, averaged, and normalized to transferrin uptake values of untransfected neurons in the same field.
HEK293 cells transfected with myc-PACSIN1 and control or PACSIN1 shRNAs were lysed and PACSIN1 levels analyzed by immunoblotting with a myc antibody (1:1000, Roche, 1667149) and normalizing to GFP band (1:5000, Clontech, clone JL-8). sh-PACSIN1 was additionally tested in striatal neurons by transfection with calcium phosphate and quantification of total endogenous PACSIN1 levels.
Samples of putamen from humans with HD and controls were obtained from Banc de Teixits Neurològics (Servei Científico-Tècnics, Universitat de Barcelona, Spain) following ethical guidelines of the Declaration of Helsinki. Informed consent was obtained from all subjects under study. GluN3A expression was analyzed in post-nuclear supernatants by Western blot as described below. A rabbit antibody generated against a fragment of mouse GluN3A expanding Aas 1041–1152 was used; the specificity of the human GluN3A band was assessed by comparison with a mouse GluN3A band absent in GluN3A knockout mice and run in parallel in the same blot.
Three HD models were used in this study: transgenic R6/1 mice expressing exon1 of human HTT carrying 115 CAG repeats27, YAC128 mice (line 55 homozygotes) containing full-length human HTT with 128 CAG repeats19, and HdhQ111 mice, with targeted insertion of a 109 CAG repeat that extends the glutamine segment in mouse Htt to 111 residues56. HdhQ7/Q111 heterozygous males and females were intercrossed to generate HdhQ7/Q111 heterozygous and HdhQ7/Q7 control littermates. Age and genetic background matched wild-type mice were used as controls for the biochemical analysis in Fig. 3 and Supplementary Fig. 5&6, and for the initial spine analysis in Supplementary Fig. 7 (CBAxC57Bl6 littermates for R6/1, FVB/N mice for YAC128, HdhQ7/Q7 littermates for HdhQ7/Q111 mice). Transgenic mice expressing juvenile GluN3A levels through adulthood16 (dt GFP-GluN3A) and single transgenic littermates were used in spine density and neuropathological analyses. For rescue experiments, YAC128 mice (in a FVB/N background) were crossed to GluN3A−/− mice (in a C57Bl6 background, also known as Grin3a−/−15), yielding heterozygote mice that were then crossed to obtain wild-type, GluN3A−/−, YAC128 and YAC128 × GluN3A−/− mice. Littermate offspring from both sexes were used unless otherwise indicated. Electrophysiological, spine and neuropathological experiments were replicated in several mice cohorts to minimize potential confounding effects of genetic background and, in all cases, experimenters were blind to the genotype of the mice until analysis was completed. All procedures conformed to the European Community guidelines for the care and use of laboratory animals and were approved by the Ethical Committee of the University of Navarra (in accord with the Spanish Royal Decree 1201/2005).
For immunoprecipitation experiments, striatal tissue or transfected HEK293 cells were lysed in T-PER buffer (Pierce) supplemented with protease inhibitors (Roche Applied Science). Protein concentrations were measured using a BCA kit (Pierce). After preclearing, lysates were incubated with a GFP antibody (1:500, Synaptic Systems, 132002), monoclonal antibody to Htt (1:500, Chemicon, MAB2166) or ImmunoPure Rabbit or Mouse Gamma Globulin (Thermo Scientific) for 4 h at 4 °C. Antibody-bound proteins were separated by centrifugation after 2 h incubation with protein A/G Agarose beads (Pierce), washed extensively with T-PER, eluted with sample buffer (Biorad), resolved using Criterion XT 3–8% Tris-acetate precast gels (Biorad) and analyzed by immunoblotting using antibodies to PACSIN (gift from M. Plomann, 1:10000), GFP (1:5000, Clontech, clone JL-8) or Htt (1:1000, Sigma, H7540).
Total or synaptic plasma membranes (SPMs) from mouse striatal tissue were obtained by biochemical fractionation as described8. Further detergent extraction of striatal synaptic plasma membranes to yield an insoluble “postsynaptic density (PSD)-enriched” fraction and a soluble “non-PSD enriched” fraction (which includes peri- and extrasynaptic receptors) was conducted using the protocol by Milnerwood et al13. Protein samples were resolved by SDS-PAGE, transferred onto PVDF membranes, and detected by immunoblot (ECL Plus, Amersham) using antibodies to: GluN1 (1:2000, MAB363), synaptophysin (1:20000, MAB5256), GluN2A (1:500, 07-632) and GluA2/3 (1:1000, AB1506) purchased from Millipore; GluN2B (1:200, NeuroMab, clone N59120), GluN3A (1:1000), PACSIN1, Rab4 (1:1000, Stressgen, ADI-KAP-GF005), Rab11 (1:1000, 610657) and DARPP32 (1:1000, 611520) both from BD Transduction Labs. Band intensities were quantified on a phosphorimager (Storm 860, Amersham) using Image-Quant 5.0 software or on a GS-800 calibrated densitometer (BioRad).
To analyze levels of intracellular pool of glutamate receptor subunits we used the membrane impermeable cross-linker, bis(sulfosuccinimidyl)suberate (BS3, Pierce) as described previously57. Acute corticostriatal slices (270 μm thick) from age-matched YAC128 and FVB/N mice were allowed to recover for 45 min in Krebs buffer. Slices were then incubated 30 min at room temperature in freshly prepared BS3 (1 mg ml−1 in D-PBS) or D-PBS. A minimum of three slices per control or treatment groups were included for each animal. To quench remaining BS3, slices were washed three times in cold 0.1 M glycine in D-PBS and then in D-PBS. Striata were then dissected out and lysed in 0.32 M sucrose/HEPES 4 mM with protease inhibitors (Roche) and PMSF (Sigma). Intracellular pool of receptors was determined by Western blot (cross-linked surface receptor complexes are retained in the stacking gel), and normalized to total receptor (non-treated).
Mice were deeply anesthesized and perfused intracardially with 4% paraformaldehyde in 0.1 M phosphate buffer (PB, pH 7.4). Coronal brain sections (30 μm) were cut with a freezing sliding microtome and stained with antibodies to GluN3A (1:100), DARPP32 (1:1000) and NeuN (1:100, Millipore, MAB377). Antigen retrieval techniques were required to reveal endogenous GluN3A. GluN3A immunoreactivity was absent from the brain of GluN3A−/− mice, establishing the specificity of the antibody for immunohistochemistry. Binding of primary antibody was visualized with a biotin-conjugated secondary antibody followed by ABC kit (Pierce) and diaminobenzidine/H2O2, or with FITC and Cy3-conjugated secondary antibodies.
For stereological analysis, unbiased counting relative to genotype and condition was performed using the Computer Assisted Stereology Toolbox (CAST) software (Olympus Danmark A/S, Ballerup, Denmark) as described58. Briefly, striatal volume was obtained by the Cavalieri method. For counts of DARPP-32 and NeuN-positive neurons per striatum, we used the dissector counting procedure in coronal sections 30 μm thick, and spaced 240 μm apart. For estimating mean cellular/perikaryal volumes of neurons (so-called local volumes) with design-based stereology, the “nucleator” method was used.
Total RNA from mouse striatum was isolated using TRIzol Reagent (Life Technologies). First strand cDNA was synthesized from 1 μg of total RNA starting material, using Superscript III First-Strand Synthesis System for RT-PCR (Invitrogen). Quantitative real-time PCR (qPCR) was performed using a pre-designed TaqMan® Gene Expression Assay kit (Applied Biosystems), consisting of a pair of unlabeled forward and reverse amplification primers and a TaqMan® probe with a FAM™ dye label. Briefly, qRT-PCR was assayed in a total volume of 25 μl reaction mixture containing 5 μl of diluted cDNA, 12.5 μl of 2× Taqman Universal Master Mix, 1.25 μl of the 20× Taqman Gene Expression Assay and RNAse-free water. Taqman® probes for GluN3A (Mm 01341719_m1) and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (Mm 99999915_g1) were from Applied Biosystems. Probe Mm 01341719_m1 recognizes exons 2–3 of the mouse Grin3a gene. PCR thermal conditions included an initial 10 min at 95 °C, followed by 40 cycles of denaturation for 15 s at 95 °C and annealing/primer elongation for 1 min at 60 °C. All qPCR reactions were run in triplicate, in two independent experiments by using the Applied Biosystems 7300 RT-PCR system. Mean cycle threshold (Ct) values for each reaction were recorded for posterior data analyses. The relative RNA expression levels were calculated using GAPDH as a control: ΔCt=Ct (GAPDH) − Ct (GluN3A). The gene expression fold change, normalized to the GAPDH and relative to the control sample, was calculated as ΔCt.
Methods described for Figure 1 of ref 13 were followed almost exactly except the tip diameter of the stimulating electrode was larger (~20 μm); stimulus duration was 200 μS. The distance from stimulating to recording electrode was approximately 175 μm. Temperature was 23–25° C. The intracellular solution contained (in mM) 140 Cs-gluconate, 9 NaCl, 1 MgCl2, 0.2 CaCl2, 1 EGTA, 10 HEPES, 2 MgATP, 0.2 LiGTP, and 5 Q×314, and the extracellular solution contained 120 NaCl, 2.5 KCl, 2 CaCl2, 10 glucose, 1.25 NaH2PO4, 26 NaHCO3, nominally 5 μM MgCl2, and 20 μM glycine, 2 μM strychnine, 50 μM picrotoxin and 50 μM D-(–)-APV as indicated. Mice were 28–35 day old males, and experimenters were blind to genotype. Same results were obtained in two independent mice cohorts.
Fresh brain hemispheres were processed following the Golgi-Cox method59. The slides were randomly coded and the experimenter was blind to genotype during image acquisition and analysis. Bright-field images of Golgi-impregnated striatal MSNs were captured with a Nikon DXM 1200F digital camera attached to a Nikon Eclipse E600 light microscope (100× oil objective). Only fully impregnated MSNs with their soma found entirely within the thickness of the section, and with at least four orders of dendrites visible, were considered for analysis. Image z-stacks were taken every 0.5 μm and analyzed with ImageJ. Dendritic segments (> 20 μm long, average: 47.35 μm; mean range: 20–95 μm) were traced through different layers of the stack and spines counted. Spine density was calculated in 4–8 dendrites of each order per neuron and values averaged to obtain neuronal averages.
For electron microscopy mice were anesthetized and perfused with saline followed by 4% paraformaldehyde and 1.5% glutaraldehyde in PB. Then, coronal sections were cut at a thickness of 60 μm using a Leica vibration microtome through the striatum. After several washes in PB, sections were postfixed with osmium tetraoxide (1% in PB) and block-stained with uranyl acetate (1% in distilled water). Sections were then dehydrated followed by propylene oxide and flat-embedded on glass slides in Durcupan (Fluka). Striata were cut at 70 nm on an ultramicrotome (Reichert Ultracut E; Leica, Austria) and collected on 200-mesh copper grids. Staining was performed on drops of 1% aqueous uranyl acetate followed by Reynolds's lead citrate. Ultrastructural analyses were performed in a Jeol-1010 electron microscope; the genotype was unknown to the experimenter. For analysis of synapse number and size the dissector principle was applied and 15 fields of 64.392 μm2 at the striatum of each animal were randomly acquired.
Two independent mice cohorts (wild-type, GluN3A−/−, YAC128 and YAC128 × GluN3A−/−) were used. Cohort 1 was used for motor and open field assessment and included 10 month-old male mice. Cohort 2, used for testing cognitive function, included 12 month-old mice of both sexes. Body weight, muscular strength and open field activity were measured29. For rotarod learning assessment, mice were trained at a fixed speed of 10 rpm and subsequently tested with two trials/day spaced 1–2 h apart during three consecutive days. During this learning phase, mice falling from the rod were returned and the number of falls recorded until the addition of the latencies to fall reached a total time of 60 s per trial13,29. For the accelerating task, the rotarod was accelerated from 5 to 40 rpm over 5 min. The latency to fall was recorded for each of two trials and averaged. For the vertical pole test, a metal pole (1.5 cm in diameter × 50 cm long) wrapped with cloth tape was used. The mouse was placed in the center of the pole which was held in a horizontal position. The pole was then gradually lifted to a vertical position and the latency to fall measured. For the swimming T-maze test, a T-maze (dimensions: arms, 38 × 14 cm; platform, 10 × 14 cm) was filled with water to a depth of 7 cm and a submerged escape platform was located in the right arm of the maze. Mice were placed into the water at the base of the stem arm of the maze and learned the location of the escape platform. During the normal testing phase, mice received 4 trials/day spaced 45 min apart for 3 consecutive days. For reversal learning, the platform was switched to the left arm of the maze. Mice received 4 trials spaced 45 min apart. Times to reach the hidden platform were recorded.
Brain slice assays for Httex1Q73-induced neurodegeneration were as described41. Briefly, 250 μm thick corticostriatal brain slices from P10 Sprague-Dawley rat pups were prepared on a vibratome and placed in interface configuration over culture medium containing 15% heat-inactivated horse serum, 10 mM KCl, 10 mM HEPES, 100 U ml−1 penicillin/streptomycin, 1 mM MEM sodium pyruvate, and 1 mM L-glutamine in Neurobasal A (Invitrogen) under 5% CO2 at 32 °C. Rat pups were sacrificed in accordance with U.S. National Institutes of Health guidelines and under Duke University Medical Center Institutional Animal Care and Use Committee approval. Slices were biolistically transfected (Helios gene gun, Bio-Rad) with DNA plasmids encoding YFP, Httex1Q73, and/or GluN3A, all in gWiz (Genlantis). Four days after transfection, MSNs were identified by their location within the striatum and by their characteristic dendritic arborization, and scored as healthy if they exhibited continuous YFP fluorescence throughout a cell body of normal diameter, and ≥ 2 clear and unbroken primary dendrites that were ≥ 2 cell body diameters long.
We thank S. Finkbeiner for the Htt constructs, H. Zoghbi for the ataxin constructs, A. Zandueta, C. Rodríguez-Viña, F. Ballesteros, X. Remírez, and M. Montañana for excellent technical help, M. Galarraga for advice with image analysis, and M. Ehlers, M. Arrasate, T. Aragón, J.A. Esteban and B.D. Philpot for critical readings of the manuscript. Work was funded by the Unión Temporal de Empresas (UTE) project at the Centro de Investigación Médica Aplicada, Gobierno de Navarra, and Spanish Ministry of Science grants (SAF2010-20636 and CSD2008-00005 to I.P.O., BFU2009-12160 to J.F.W. and SAF2011-29507 to J.A.), grants from Hereditary Disease Foundation (to I.P.O. and D.C.L.), US National Institute of Health grants P01 HD29587, P01 ES016738, and R01 EY05477 (to S.A.L.), grants from Cure Huntington's Disease Initiative and Canadian Institutes of Health Research (to M.R.H.). M.R.H., a Killam University Professor, holds a Canada Research Chair in Human Genetics.
Author Contributions: S.M. performed and analyzed the biochemical experiments in cultured neurons, recombinant cells and YAC128 mice, the imaging experiments in cultured neurons, the spine morphology measurements, and collaborated with A.G. on neuropathological studies. A.G. performed and analyzed biochemical experiments in human, R6/1 and knock-in mice, and behavioral experiments. M.M.P. performed electrophysiological recordings. M.A.P. performed and analyzed behavioral experiments. R.M.T. contributed to mouse genotyping, and conducted biochemical fractionation and real-time PCR assays. J.M.H. and R.L. performed and analyzed electron microscopy studies. L.K. and D.C.L. performed and analyzed slice culture experiments. J.T.P. performed initial biochemical experiments in R6/1 mice. M.R.H. and R.K.G. provided the YAC128 mice. N.N. and S.A.L. provided the GluN3A−/− mice. M.W. made the GluN3A antibody used in biochemical and immunocytochemical analyses. J.A. designed and supervised experiments. J.F.W. designed and analyzed electrophysiological and fluorescence colocalization experiments, and contributed to the writing. I.P.O. conceived the study, designed experiments, analyzed data and wrote the paper.
Competing Interests Statement: The authors declare that they have no competing financial interests.