Abstract
Synaptic plasticity relies on new protein synthesis in dendrites that involves the selective translation of specific mRNAs. This requires a tight control of mRNA levels in dendrites. Consistently, RNA translation and degradation pathways have been recently linked to neurodevelopmental and neuropsychiatric diseases, suggesting a role for RNA regulation in synaptic plasticity and cognition. Despite being the only RNA regulatory pathway that is associated with multiple mental illnesses, the Nonsense-Mediated mRNA Decay (NMD) pathway presents an unexplored regulatory mechanism for synaptic function and plasticity. NMD is a highly conserved and selective RNA degradation pathway that exerts its function in a cell- and spatiotemporally-specific manner. Here, we show that neuron-specific disruption of NMD in adulthood attenuates learning, memory, hippocampal LTP, and potentiates perseverative/repetitive behavior. While it is known that local translation of specific mRNAs in dendrites enables synaptic plasticity, the tightly-controlled mechanisms that regulate local quantity of specific mRNAs remains poorly understood. We report that the NMD pathway operates within dendrites to regulate GluR1 surface levels. Specifically, NMD modulates the internalization of GluR1 and promotes its local synthesis in dendrites. We identified AMPK as a mechanistic substrate for NMD that contributes to the NMD-mediated regulation of GluR1 by limiting total GluR1 levels. These data establish that NMD regulates synaptic plasticity, cognition, and local protein synthesis in dendrites, providing fundamental insight into the neuron-specific function of NMD within the brain.
Introduction
NMD is a process that regulates protein expression though degradation of specific mRNAs [1–3]. NMD was first identified as an mRNA surveillance pathway that degrades mRNAs undergoing premature termination of translation [4–6]. These mRNAs are detected during the first round of translation. In most mRNAs, all exon-junction complexes (EJCs), a group of proteins bound to mRNA after splicing, are upstream of the termination codon. When a ribosome encounters an EJC downstream of a termination codon, the termination codon is perceived as premature, and the mRNA is considered nonsense. This initiates NMD, recruiting core Up-Frameshift (UPF) proteins - namely UPF1, UPF2, and UPF3B - and ultimately leads to mRNA degradation [7, 8]. NMD thus prevents the production of truncated proteins with a dominant-negative function. Recently, it has been shown that NMD also targets physiological mRNAs that naturally have NMD-inducing features but code for functional proteins [9, 10]. A common event that leads to NMD is alternative splicing, since alternatively spliced exons or introns typically create in-frame termination codons followed by EJCs [11, 12]. Intra-axonal translation of such an mRNA coupled to NMD controls a switch in receptor expression and thereby regulates axon guidance [11]; suggesting that mRNA turnover is a key player in local protein synthesis. Intron retention in the 3’ UTRs of endogenously expressed transcripts also targets them to NMD [13]. For instance, the mRNA of the synaptic plasticity protein Arc is a known target of NMD due to intron retention in its 3’ UTR [13]. Arc is highly expressed in dendrites [14–17] and is required for synaptic plasticity and memory consolidation [18–21]. Similar to Arc mRNA, various mRNAs with intron retentions are localized to dendrites [22] and have been implicated in synaptic plasticity and memory [19, 23–26]. However, the extent to which NMD is physiologically relevant to synaptic plasticity is not clear.
Despite its putative link to many mental disorders including autism and schizophrenia [27–32], whether NMD is relevant to the synaptic pathology and behavioral deficits of these diseases remains unclear. A recent study showed that UPF3B-null mice exhibit deficits in fear-conditioned learning, supporting a role for NMD in regulating synaptic plasticity and cognition [33]. However, because NMD is ubiquitous and required for neuronal differentiation [34], disruption of this pathway during development hinders determining the direct role of NMD in these processes. Therefore, understanding the precise, cell-type specific role of NMD in developing and adult brain is likely to provide novel insights into key mechanisms critical for synaptic plasticity and behavior.
Synaptic plasticity underlies changes in neuronal network dynamics and is therefore considered to be the foundation of learning and memory. It takes several forms, including the modification of synaptic strength and spine structure. Synaptic transmission can be influenced by activity, becoming either enhanced, through long-term potentiation (LTP), or depressed, through long-term depression (LTD) [35]. Many higher cognitive disorders are associated with alterations in spine density and synaptic plasticity [36, 37]. Mechanisms that regulate synaptic plasticity are therefore fundamental to understanding higher brain functions, including learning and memory, and have been implicated in the etiology of neuropsychiatric diseases. Both LTP and LTD rely on local protein synthesis in dendrites [21, 38]. Locally synthesized proteins are thought to either strengthen a pre-existing synaptic connection, by inducing structural remodeling, or to promote the formation of new synaptic connections. Deficits in mRNA regulation can result in ectopic protein synthesis and lead to cognitive disabilities [39–45], having downstream effects on synaptic plasticity and higher-order cognition. For example, exaggerated mRNA translation leads to alterations in spine density, LTP and LTD [42, 45–47] suggesting that a tight control of protein synthesis is required at many levels within dendrites [40]. A great amount of prior research has addressed the pathways that regulate translational derepression in dendrites. However, the mechanisms that control mRNA quantity during synaptic function remain unknown. Importantly, it is not known whether NMD functions locally to regulate mRNA levels in dendrites, and if mRNA degradation contributes to the regulation of synaptic plasticity.
Here we show that both synaptic plasticity and cognitive function is physiologically regulated by neuronal NMD. Conditional deletion of the NMD component UPF2 in adult glutamatergic neurons in vivo disrupts learning and memory and potentiates perseverative/repetitive behavior. Deletion of UPF2 also results in altered LTP, but not LTD, in the adult hippocampus. This phenotype is accompanied by a significant reduction in total GluR1 levels in conditional knockout mice. We show that NMD regulates GluR1 surface levels through mechanisms that influences GluR1 internalization and nascent protein synthesis. Selective depletion of UPF2 from cultured hippocampal neurons reduces both total and surface expression of GluR1 without affecting GluR1 mRNA levels. This phenotype is recapitulated when NMD is disrupted in dendrites via the targeted-inhibition of UP3B local synthesis; the only dendritically synthesized component of the NMD machinery. Disruption of NMD increases internalization of GluR1 in dendrites, which may account for the reduced surface levels of this receptor. This is consistent with previous findings that Arc mediates GluR1 internalization and thereby reduces surface levels of GluR1. However, elevated levels of Arc are, at most, only partially responsible for the reduced surface levels of GluR1 in UPF2-deficient dendrites. In addition to its altered internalization, the local synthesis of GluR1 is also reduced in UPF2-deficient dendrites due to increased levels of AMPK mRNA. Normalizing the levels of AMPK and Arc mRNAs, together, completely corrects GluR1 levels in UPF2-deficient dendrites. These data demonstrate a role for NMD in regulating cognitive function, the dendritic proteome, and synaptic plasticity, as well as potentially other essential dendritic functions.
Results
NMD is required for learning and memory and suppresses perseverative behavior
We first sought to determine the requirement for neuronal NMD in cognitive function and synaptic plasticity. To do this, we genetically disrupted NMD in postmitotic glutamatergic neurons using a conditional knockout mouse of UPF2 (UPF2fl/fl) [48]. The function of UPF2 is exclusive to the NMD pathway, and this line has been successfully used to target NMD in various tissues [11, 48, 49]. Because NMD is ubiquitous and required for neuronal differentiation [34], deleting the UPF2 gene during development would preclude determining an explicit role of NMD in adult plasticity and cognition. Therefore, we temporally controlled NMD disruption by crossing the UPF2fl/fl line with a mouse line that expresses tamoxifen-inducible Cre under the αCaMKII promoter (αCaMKII:CreERT2) [50]. In all behavioral experiments, we induced Cre in 2-month old conditional-knockout (UPF2fl/fl;αCaMKII:CreERT2; CKO) mice as well as in their control littermates (UPF2wt/wt;αCaMKII:CreERT2; CTRL) by tamoxifen. This experimental design ensures that phenotypes are not caused by an acute effect of tamoxifen, or a non-specific effect of Cre at cryptic LoxP sites. Loss of UPF2 protein in excitatory neurons was successfully achieved by 200mg/kg of tamoxifen administered every other day for 5 doses (Figure S1a). A 2-week washout period followed tamoxifen treatment before plasticity and behavioral analyses began. We first evaluated the general locomotor activity of conditional UPF2 CKO mice in locomotor photocells [51]. No difference in locomotor activity was detected between UPF2 CKO mice and controls, indicating that NMD does not induce hypo-or hyperactivity (Figure S1b). Likewise, no evidence of anxiety-related behavior was observed in UPF2 CKO mice in the elevated-plus maze or open field assays (Figure S1c-d), nor was there a difference in sociability between groups (Figure S1e).
We next examined hippocampus-dependent learning and memory in UPF2 CKO mice. To do this, we used multiple behavioral paradigms, including the Y-maze, contextual fear conditioning and the Morris water maze. On the Y-maze, UPF2 CKO mice showed no preference for the novel arm relative to the other arms, and displayed significantly worse novel arm preference than the control group (Figure 1a), suggesting that UPF2 CKO mice have disrupted short-term spatial memory. As an additional measure of hippocampal function, we tested UPF2 CKO mice for contextual fear memory. In this test, UPF2 CKO mice froze for significantly less time than controls by the last tone-shock pairing during conditioning, and this conditioning deficit resulted in worse contextual fear memory in UPF2 CKO mice relative to control mice 24 hr later (Figure 1b-c). To further disambiguate whether our observed phenotypes on the Y-maze and contextual fear conditioning represented an acquisition failure as opposed to a memory deficit, we subjected mice to the Morris water maze. In the Morris water maze, UPF2 CKO mice displayed a robust attenuation of learning behavior across training and correspondingly decreased exploration of the target quadrant and increased latencies to enter the (now removed) hidden platform zone on the probe trial (Figure 1d-f). UPF2 CKO mice failed in most trials to even attempt to locate the hidden platform despite being guided to its location after each trial. Indeed, the final task performance of these mice did not surpass the performance of control mice after just two days of training. These data collectively suggest that UPF2 CKO mice have attenuated learning behavior, which produces worse performance on common assays of hippocampus-dependent memory function.
Because the NMD pathway has been previously linked to autism and schizophrenia [27–32], where stereotypies are common to both disorders, we also tested for perseverative/repetitive behaviors [45, 52–55]. This primarily included an assessment of marble burying and self-grooming. In our marble burying assay, UPF2 CKO mice buried significantly more marbles than control mice (Figure 1g; 81.14% vs 58.86% of marbles). In an activity-sampling test session, where grooming behavior was continuously monitored for 20 min, CKO mice groomed for longer and displayed a greater frequency of grooming bouts compared to the control group (Figure 1h-i). Other stereotypies such as digging, circling, and backflipping were not significantly different between the genotypes (Figure 1i). Together, these data suggest that NMD is required for proper learning and memory, and its disruption potentiates perseverative/repetitive behaviors.
NMD ensures proper spine density and L-LTP in hippocampus
We next examined spine density in UPF2 CKO mice. During LTP, activation of glutamate receptors increases connectivity of specific neurons through the enlargement of pre-existing spines or the formation of new spines, which supports learning and memory. To determine whether NMD is required for proper spine density, we quantified spine density in hippocampal dendrites of both UPF2 CKO and control brains cut at 200 μm. To visualize dendrites and spines, we injected AAV-αCaMKII-eGFP virus into the dorsal CA1 field 7d prior to perfusion. We observed significantly reduced CA1 spine density in UPF2 CKO hippocampal dendrites compared to control dendrites in vivo (Figure 1j). We next sought to investigate the role of NMD in well-studied forms of synaptic plasticity in the adult brain by assessing both LTD and LTP in these mice. To do this, we prepared hippocampal slices from 3-month old UPF2 CKO mice and their control littermates following induction of Cre expression by tamoxifen at 2-month of age. We incubated 400 μm slices for at least 3 hr in artificial cerebral spinal fluid (ACSF) before measuring LTP. LTP can be divided into two phases: early phase LTP (E-LTP) and late phase LTP (L-LTP). E-LTP does not require new gene transcription or mRNA translation, whereas L-LTP requires new gene transcription and mRNA translation and results in an increase in the number of AMPA receptors accompanied by an increase in the size of the synaptic connection. Briefly, we recorded field excitatory postsynaptic potentials (fEPSPs) from the stratum radiatum and activated Schaffer collateral/commissural afferents at 0.05 Hz. We used a single 100 Hz high-frequency stimulation (HFS) to induce E-LTP and three trains of HFS with a 60 sec intertrain interval to induce L-LTP. We found that L-LTP, but not E-LTP, was significantly decreased in UPF2-deficient slices compared to control slices (Figure 1k-l). On the other hand, LTD, which was induced by application of 100 μM DHPG for 10 min, was not different in UPF2-deficient slices compared to control slices (Figure S1f-g). These data suggest that NMD is required for proper LTP and spine density but not for LTD in hippocampus.
NMD positively regulates GluR1 surface expression
We next addressed the potential mechanisms through which NMD regulates synaptic plasticity and cognitive function. It was previously shown that disruption of the EJC, an essential component for targeting mRNAs to the NMD pathway, causes an increase in the levels of AMPA receptor GluR1 in hippocampal dendrites in vitro [13]. Functionally, AMPA receptors mediate the vast majority of excitatory synaptic transmission by ensuring rapid responses to glutamate, the principal neurotransmitter in the mammalian central nervous system. Modulation of AMPA-receptor integrity plays a crucial role in synaptic strength. Strengthening of synapses by LTP or weakening of synaptic strength by LTD can be represented by the synaptic insertion or removal of AMPA receptors, respectively [56]. For example, low surface expression of AMPA receptor GluR1 increases the number of dendritic spines [57]. Similarly, reductions in GluR1 expression cause working-memory deficits in mice [58]. If EJC function were specific to the NMD pathway, the EJC disruption study would suggest that NMD negatively regulates GluR1 levels; but EJC proteins have multiple functions, including transportation and translation of mRNAs [59]. Intriguingly, contrary to what is observed in studies of disrupted EJC proteins, we found that the loss of UPF2 led to a decrease in GluR1 levels in vivo (Figure 1m). To explore this phenotype in vitro, we infected hippocampal neurons with UPF2-shRNA:GFP lentivirus to interrupt endogenous NMD activity at 7 days in vitro (DIV7) and examined GluR1 expression on DIV21. We tested 2 UPF2-shRNA lentiviruses. Infection of hippocampal neurons with either UPF2-shRNA 1 or UPF2-shRNA 2 led to a nearly complete loss of UPF2 protein in these neurons (Figure S2). We measured the total levels of GluR1 using an anti-Cterminus-GluR1 antibody in permeabilized cells and found that total levels of GluR1 were decreased both in cell bodies and dendrites of hippocampal neurons upon knockdown of UPF2 (Figure S3a-c). Thus, the reduction in GluR1 observed in UPF2 CKO mouse hippocampus was recapitulated in neurons and their dendrites in vitro.
We next asked whether NMD regulates GluR1 surface expression in neurons. An essential component of GluR1 signaling involves the dynamic insertion and removal of the GluR1 receptor to the synaptic surface [60]. To label surface GluR1, we incubated fixed, but not permeabilized, neurons with a monoclonal anti-Nterminus-GluR1 antibody at 4°C as described previously [61, 62]. We found that, similar to the total levels, the loss of UPF2 led to a decrease in the surface levels of GluR1 (Figure 2; see also Figure S3c and S4 for UPF2 shRNA 2 knockdown and surface GluR1/PSD-95 quantifications, respectively). This suggests that NMD positively regulates GluR1 surface expression. GluR1 mRNA is not a canonical target of NMD. However, it is known that some mRNAs that do not carry any NMD-inducing features can also be degraded by NMD [63]. In addition, it has been suggested that GluR1 transcription is regulated by Arc [64]. Consistent with Arc mRNA being an established target of NMD [11, 13], the levels of both Arc mRNA and Arc protein are increased in hippocampal neurons upon knockdown of UPF2 (Figure S5). We, therefore, examined cytoplasmic GluR1 mRNA levels in UPF2-deficient cell bodies. Overall GluR1 mRNA levels were unaltered following knockdown of UPF2 in hippocampal neurons, indicating that the reduction in total and surface GluR1 levels is not due to low levels of its mRNA (see Figure S7 for viral infection efficiency in hippocampal cultures). These data suggest that NMD modulates GluR1 signaling by restraining a mechanism that negatively regulates the synthesis, surface localization or surface turnover of GluR1 in dendrites.
NMD negatively regulates GluR1 internalization in dendrites
We next sought to determine how NMD regulates surface expression of GluR1 in dendrites. It is possible that the reduction in total GluR1 levels (Figure 1m and Figure S3a-b) might account for decreased surface levels of this receptor. However, locally synthesized Arc is also known to cause GluR1 endocytosis, leading to a decrease in the surface levels of this receptor [21, 65]. This suggests that increased levels of Arc might also contribute to downregulated surface-GluR1 levels upon loss of UPF2. To determine if GluR1 endocytosis is altered after disruption of NMD, we measured the basal internalization rate of GluR1 upon knockdown of UPF2 in hippocampal dendrites using a previously established protocol [66]. We observed significantly increased GluR1 basal internalization in UPF2-deficient hippocampal dendrites (Figure 3a-b). This is consistent with the increased levels of Arc in these neurons, namely because Arc-mediated internalization of GluR1 only influences surface but not total levels of GluR1 [67]. This suggests that UPF2 regulates total GluR1 levels via yet defined NMD targets.
NMD positively regulates local synthesis of GluR1 in dendrites
An overall reduction in GluR1 protein levels likely contributes to the decreased dendritic GluR1 levels upon loss of UPF2 in hippocampal neurons (Figure S3). However, the total levels of dendritic GluR1 are determined by the amount of GluR1 protein trafficked to dendrites from cell bodies as well as a balance of local protein synthesis [68] and local protein degradation [69, 70] in dendrites. Thus, we also investigated the possibility of enhanced GluR1 proteolysis or decreased GluR1 synthesis in UPF2-deficient dendrites. To do this, we used a custom-fabricated tripartite microfluidic device, which, unlike traditional microfluidic chambers with two channels [71], contains three channels (Figure 3c and Figure S6-7). As each channel is fluidically isolated, tripartite chambers permit the selective treatment or harvesting of discrete neuronal populations or compartments. In order to access a high number of dendritic synapses, a ‘synaptic’ channel was placed 200 μm from the edge of one channel and 500 μm away from the other. Because the average length of dendrites extending into the microgrooves is less than 300 μm at DIV21, the closer “postsynaptic cell channel” allows a large number of dendrites to project from this channel to the “synaptic” channel (Figure 3c). This enabled us to selectively isolate, treat and harvest dendrites in subsequent experiments.
When AMPA receptors are internalized, endocytosed receptors undergo lysosome- or proteasome-mediated degradation [70, 72–75]. To study local degradation of GluR1, we cultured E16 hippocampal neurons in tripartite chambers and applied control- or UPF2-shRNA viruses only to postsynaptic cell channels at DIV7. This resulted in disruption of NMD only in postsynaptic cell bodies and dendrites but not in presynaptic cells (Figure S7). At DIV21, we inhibited both proteosomal and lysosomal degradation by treating the synaptic channels with MG132 and leupeptin. Following 6 hr of treatment, we measured the surface levels of GluR1 in control and UPF2-deficient dendrites in the absence and presence of degradation inhibitors. Co-inhibition of lysosome- and proteasome-mediated degradation did not restore GluR1 levels in UPF2-deficient dendrites (Figure S8). Thus, degradation of GluR1 receptors at the protein-level does not contribute to the reduction in surface GluR1 observed in UPF2-deficient dendrites.
We next asked whether local synthesis of the GluR1 receptor is altered in NMD-deficient dendrites. To measure the dendritic synthesis of GluR1, we used a CLICK-chemistry based approach and labeled locally synthesized nascent proteins at synapses (Figure 3c-e). We cultured E16 hippocampal neurons in tripartite chambers and targeted NMD only in postsynaptic cells by applying the UPF2-shRNA virus into the postsynaptic cell channel at DIV7. At DIV21, we exchanged the growth medium in the synaptic channels to methionine-free medium containing the methionine analog azidohomoalanine (AHA) [76] to label all newly synthesized proteins at synapses (Figure 3d). After harvesting the material from the synaptic channels, we incubated lysates with DBCO-biotin [77] to label newly synthesized proteins with biotin. We purified biotin-containing proteins over streptavidin columns, separated by SDS-PAGE, and detected protein by immunoblotting with an anti-Cterminus-GluR1 antibody. The amount of GluR1 protein that bound the streptavidin in the UPF2-deficient synapses was significantly lower compared to the amount of GluR1 that bound the streptavidin in the control synapses (Figure 3e). This data indicates that dendritic synthesis of GluR1 is reduced upon disruption of UPF2.
Local degradation of Arc and AMPK mRNAs by NMD in dendrites
We next sought to determine whether local NMD contributes to the regulation of GluR1 protein synthesis in dendrites. Similar to Arc mRNA, the mRNA of AMP-activated protein kinase (AMPK) also contains an intron retention in its 3’UTR [13], making this mRNA a potential substrate for NMD. Activated AMPK interrupts translation by either inhibiting the mammalian target of rapamycin (mTOR) kinase or the translation elongation factor 2 [78–83]. Interestingly, AMPK has been suggested to negatively regulate GluR1 protein levels in dendrites [25]. The AMPK mRNA that carries NMD-inducing feature is transcribed from the PRKAG3 gene and encodes the non-catalytic subunit of the AMPK heterodimer protein. Intriguingly, both Arc and AMPK mRNAs are localized to dendrites, suggesting that they might be degraded by NMD in these compartments.
To explore this possibility, we first asked whether NMD functions locally in dendrites. NMD occurs in axonal growth cones and influences the local proteome by regulating mRNA dynamics [11]. Similar to growth cones, we found that the core proteins of the NMD machinery are localized to dendrites (Figure 4a), suggesting that local NMD might contribute to regulation of GluR1 levels in these compartments. We first examined Arc and AMPK mRNA levels in UPF2-deficient synaptic regions. To do this, at DIV7, we selectively infected postsynaptic neurons with UPF2-shRNA lentivirus and isolated RNA from synaptic channels at DIV21. qRT-PCR showed an increase in the levels of both Arc and AMPK mRNAs in UPF2-deficient synaptic channels compared to control synaptic channels, suggesting that these targets are subjected to NMD in dendrites (Figure 4b).
To determine whether NMD-mediated mRNA degradation occurs in dendrites, we examined the AMPK and Arc mRNAs upon selective inhibition of translation in these compartments. NMD can only occur when ribosomes recognize NMD-inducing features in mRNAs during the first round of translation [84, 85]. Inhibition of translation hinders NMD and leads to the accumulation of NMD-target mRNAs [7]. Therefore, a widely used approach to determine whether an mRNA is subjected to NMD is to determine if it accumulates after the treatment of cells with cycloheximide (CHX), a translation inhibitor. To confirm whether these targets undergo translation-dependent degradation in synaptic regions, we cultured E16 neurons in tripartite chambers and, at DIV21, inhibited translation selectively in synaptic channels. qRT-PCR on material harvested from the synaptic channels showed that both AMPK and Arc mRNAs accumulated in these compartments upon inhibition of translation (Figure 4c).
We next asked whether NMD contributes to the regulation of GluR1 surface levels locally in dendrites. To address this, we targeted NMD selectively in synaptic channels by inhibiting the local synthesis of UPF3B, a protein specific to the NMD pathway [86, 87]. From the NMD machinery, only the UPF3B mRNA is localized to dendrites [22] (Figure S9a). We confirmed that the UPF3B mRNA is locally translated in dendrites, and that its local translation is the only source for UPF3B protein in synaptic channels (Figure S9b). To disrupt local NMD in dendrites, we targeted the UPF3B mRNA by siRNA transfection of synaptic channels. The siRNA-mediated depletion of targets has been successfully used in microfluidic devices previously, allowing exclusive knockdown in neurites without influencing expression in cell bodies or even microgrooves [88–93]. We cultured E16 neurons in cell-body channels and, starting at DIV14, repeatedly treated synaptic channels with siRNAs against UPF3B mRNA for seven days. This resulted in complete loss of UPF3B protein in synaptic channels (Figure S9d). To determine whether the expression of GluR1 is affected upon a strictly local disruption of NMD in synaptic channels, we measured the frequency of surface GluR1 upon knockdown of local UPF3B in these channels. Similar to the knockdown of UPF2 in postsynaptic cells, the surface frequency of GluR1 was reduced upon selective loss of UPF3B protein in synaptic channels (Figure 4d).
Taken together, these experiments establish the local requirement for NMD in synaptic regions and suggest that the local regulation of GluR1 by this pathway might involve AMPK degradation in addition to Arc.
AMPK negatively regulates GluR1 expression in dendrites
We next examined whether AMPK locally influences GluR1 expression in synaptic channels. To do this, we cultured E16 neurons in cell-body channels and, starting at DIV14, repeatedly treated synaptic channels with siRNAs against the AMPK mRNA for seven days. This resulted in an almost complete loss of AMPK protein in synaptic channels (Figure S9e). We then evaluated the total levels of GluR1 in synaptic channels upon loss of AMPK. Selective inhibition of AMPK in synaptic channels significantly increased the total levels of GluR1 (Figure 4e).
Because the synaptic channels of tripartite chambers contain both dendrites and axons, it is possible that GluR1 phenotypes observed after knockdown of local UPF3B or local AMPK may arise also from axonal changes and therefore not selectively from dendrites. To clarify this, we examined the proteins of the NMD machinery and targets of NMD in mature axons. Neither the NMD-specific components of the NMD machinery, nor AMPK or Arc mRNAs, were present in isolated hippocampal axons at DIV21 (Figure S10). In fact, the axonal transcriptome significantly shrinks as axons mature [94] and does not include any known NMD targets [95, 96]. These data indicate that alterations in GluR1 expression upon selective disruption of either UPF3B or AMPK proteins in synaptic channels solely originate from a local effect within dendrites.
Normalization of AMPK and Arc mRNA levels rescues GluR1 surface expression in UPF2-deficient dendrites
Lastly, we functionally tested a mechanistic role of locally synthesized AMPK and Arc in the NMD-mediated regulation of surface GluR1 expression. To do this, we systematically modulated the elevated levels of Arc and AMPK mRNAs in UPF2-disrupted dendrites by siRNA transfection of synaptic channels. As siRNA treatment affects mRNA expression in a concentration-dependent manner [97–100], we first determined the amount of siRNA cocktail (see Methods) required to correct levels of Arc and AMPK mRNAs in UPF2-deficient dendrites. First, we applied the UPF2-shRNA virus to postsynaptic cell channels at DIV7. Starting at DIV14, we treated the synaptic channels of UPF2-deficient dendrites with non-overlapping siRNA cocktails at different concentrations for each target mRNA (Figure S11). After determining the amount of siRNA needed to normalize the exaggerated levels of Arc and AMPK mRNAs, we modulated the levels of these targets in synaptic channels containing UPF2-deficient dendrites (Figure 5a). After a week of siRNA treatment, we evaluated the surface expression of GluR1. Compared to synaptic channels with intact NMD (i.e., those treated with control-shRNA lentivirus), cultures infected with UPF2-shRNA lentivirus had significantly lower GluR1 density as expected. Restoring expression levels of Arc or AMPK mRNAs in synaptic channels led to a significant but incomplete recovery of surface GluR1 in these compartments for each target (Figure 5b-c). We next considered whether the surface GluR1 phenotype in UPF2-deficient dendrites might be a combinatorial effect of both elevated Arc and AMPK, with altered GluR1 surface levels arising from an amalgamation of both increased internalization and reduced protein synthesis. Normalization of both Arc and AMPK mRNAs together led to a complete recovery of GluR1 density in UPF2-deficient dendrites. This data indicates that local NMD contributes to the regulation of GluR1 surface expression through physiological degradation of both Arc and AMPK mRNAs in dendrites.
Discussion
Our study identifies neuronal NMD as a mechanism that contributes to the regulation of synaptic plasticity and cognitive function. Specifically, we report that NMD influences synaptic plasticity as well as learning and memory. In addition to the enrichment of NMD-machinery components in dendrites, which suggests that NMD functions locally in these compartments, our data indicates that local NMD participates in GluR1 regulation by limiting the amount of Arc and AMPK proteins in dendrites.
Role of NMD in regulating synaptic plasticity and cognitive function
Our data indicate that proper synaptic plasticity and cognitive function requires NMD. Our data show that UPF2-deficient hippocampal slices show decreased L-LTP. In addition, spine density in the hippocampus of conditional UPF2 mutant mice is reduced relative to control mice. Mice also show attenuated learning and memory upon conditional ablation of UPF2 in excitatory neurons. Memory deficits were readily detectable in UPF2-deficient mice on three independent hippocampus-dependent behavioral assays. In addition to attenuated learning and memory, loss of UPF2 in excitatory neurons resulted in perseverative behavior in mice, including increased marble burying and self-grooming behavior. Together, these data implicate a role for NMD in both synaptic plasticity and behavioral endophenotypes common to a spectrum of neurodevelopmental diseases including autism and schizophrenia.
Our data also provide comprehensive insight into the neuron-specific and spatiotemporal requirement for NMD in plasticity and behavior. Recently, a UPF3B null mutant mouse line was shown to have fear-conditioning defects but, unlike our observations in UPF2 conditional knockout mice, no spatial learning deficit [33]. There may be several reasons for this. For instance, NMD is ubiquitous and required for neuronal differentiation [34, 101]. Consistent with this, global interruption of NMD by ablation of UPF3B during brain development alters neuronal maturation and is thereby likely to contribute to the behavioral phenotypes observed in the UPF3B null mouse line. In addition, while UPF2 and UPF3 proteins are known to be specific to the NMD machinery, it has been suggested that NMD might have different branches in which NMD exerts its function independent of UPF2 or UPF3B [87, 102, 103]. Therefore, studies of conditional UPF2 deletion provide a platform to dissect not only the temporal and cell-type specific functions of the NMD machinery, but also branch-specific functions of NMD in brain development and behavior.
NMD-mediated GluR1 regulation and its contribution to behavior
Our data also demonstrate that NMD positively regulates GluR1 signaling in hippocampus. The dynamic insertion and removal of GluR1 receptors to the synaptic surface directly modulates GluR1 signaling [49]. We report that loss of UPF2 in adult hippocampal neurons caused a concomitant decrease in total and surface levels of this receptor. GluR1 mRNA expression was not significantly changed in UPF2-deficient neurons, indicating that GluR1 mRNA is not degraded upon disruption of UPF2. This suggests that synthesis or degradation of the GluR1 receptor must be regulated via proteins affected by NMD-mediated degradation. Our CLICK-chemistry experiments suggest that NMD positively regulates GluR1 protein synthesis. GluR1 synthesis is repressed in the absence of UPF2 protein, causing a drop in the total levels of GluR1 protein. In addition, loss of UPF2 results in a higher GluR1 internalization rate. These mechanisms together account for the surface GluR1 phenotype caused by the disruption of NMD. Therefore, NMD-dependent GluR1 regulation involves restraining synthesis and internalization of the GluR1 receptor.
Modulation of AMPA-receptor integrity plays a crucial role in synaptic strength. Strengthening of synapses by LTP or weakening of synaptic strength by LTD can be represented by the synaptic insertion or removal of AMPA receptors, respectively [56]. Consistent with this, alterations in GluR1 levels are linked to impairments in LTP as well as abnormalities in memory [52–54]. Our data show that hippocampal L-LTP, which is thought to underlie memory consolidation, is decreased in conditional UPF2 mutant mice. Activation of glutamate receptors facilitates LTP either by through the enlargement of pre-existing spines or promoting the formation of new spines. Therefore, a reduction in spine density is consistent with a decrease in L-LTP in conditional UPF2 mutants. This phenotype is accompanied by behavioral defects, including a lack of short-term spatial memory in the Y-maze, defective associative fear learning and memory, and attenuated spatial learning and memory on the Morris water maze. Consistent with these data, reductions in GluR1 expression were shown to cause spatial memory dissociations in mice [58]. Similarly, contextual conditioning requires increased levels and surface localization of GluR1 [52, 104, 105]. We show that UPF2-deficient neurons display reduced GluR1 synthesis, which results in an overall decrease in GluR1 levels. NMD-dependent control of GluR1 signaling is therefore likely to contribute to the plasticity and behavioral phenotypes conditional UPF2 mutant mice exhibit. However, we cannot exclude the possibility that NMD regulates other receptors or intracellular signaling pathways that might also contribute to LTP, learning and memory. Although the ionotropic glutamate receptors, N-methyl-D-aspartate receptors (NMDARs), are more static than AMPA receptors in mature neurons, subunits of NMDARs are also locally translated in dendrites [106], suggesting a potential function for NMD in regulation of NMDAR dynamics.
Role of local AMPK in GluR1 regulation and synaptic plasticity
Our data also show that the NMD-target AMPK, whose levels are elevated in UPF2-deficient dendrites, negatively regulates GluR1 levels. The AMP kinase consists of one α, one β, and one γ subunit. We found that the mRNA of the γ subunit, which is required for AMPK activation, is a substrate of NMD (Figure 4b-c). Our data shows that local disruption of AMPK within dendrites led to an increase in GluR1 levels, suggesting that local AMPK functions to negatively regulate GluR1 levels. Activated AMPK is known to interrupt translation by either inhibiting the mammalian target of rapamycin (mTOR) kinase or the translation elongation factor 2 [78–83]. Our data indicates that the local synthesis of GluR1 protein in dendrites is reduced upon disruption of UPF2, which coincides with an elevation of AMPK levels in UPF2-deficient dendrites. We also show that modulation of AMPK levels within these dendrites is required to restore GluR1 surface levels (Figure 5b-c). These suggest that while AMPK represses GluR1 synthesis, NMD limits the amount of AMPK thereby serving as a balance in regulating total levels of the GluR1 receptor. Whether AMPK regulates the synthesis of other synaptic proteins and how this specificity is achieved is yet to be determined. Intriguingly, inhibition of AMPK was shown to alleviate impairments in hippocampal synaptic plasticity in a mouse model of Alzheimer’s disease [107]. Inhibition of AMPK corrected the Amyloid β-induced inhibition of LTP suggesting that AMPK has negative effects on LTP induction. This finding is consistent with our data that L-LTP was decreased in UPF2-deficient hippocampal slices.
Activity-induced local NMD as a regulator of the dendritic proteome
We found that NMD can occur locally in dendrites. Arc mRNA was shown to be locally degraded in hippocampus [108]. However, the involvement of NMD in this, as well as more broadly hippocampal function, has not been demonstrated. Our data show that the NMD machinery is localized to dendrites, and the levels of the NMD targets Arc and AMPK are elevated upon selective inhibition of translation in these compartments (Figure 4c). The mRNA of UPF3B is trafficked to [22] and locally translated in dendrites (Figure S9). Local inhibition of dendritic UPF3B synthesis results in dysregulated surface localization of GluR1, indicating that NMD locally influences GluR1 levels within not just neuronal cell bodies but also their dendrites. This is consistent with the dynamic modulation of GluR1 surface levels at synapses.
Our data demonstrate that local translation is regulated by mechanisms that control mRNA degradation in dendrites. Synaptic plasticity relies on local protein synthesis at synaptic sites [21, 38]. Locally synthesized proteins are thought to either strengthen a pre-existing synaptic connection, by inducing structural remodeling, or to promote the formation of new synaptic connections. NMD is likely to have more targets than Arc and AMPK in dendrites. In addition to its canonical targets, NMD can also degrade mRNAs that do not carry any NMD-inducing features [63]. The mechanisms by which NMD recognizes its atypical targets remain unclear. This raises the possibility that NMD might be regulating various physiological mRNAs in dendrites that cannot be predicted based on sequence information. Similarly, RNA sequencing cannot distinguish between 1) true NMD targets, 2) secondary effects mediated by NMD-targets or 3) compensatory mechanisms that may occur upon disruption of NMD. Therefore, we cannot exclude the possibility that atypical targets of NMD may also contribute to the NMD-mediated GluR1 regulation at synapses as well as other synaptic events.
Our data also suggest that NMD is positively regulated by synaptic activity. mRNA degradation is regulated during development of the nervous system. microRNAs negatively regulate NMD activity in neural progenitors and impact the expression of a broad range of NMD targets that are important for appropriate differentiation [109]. UPF3B mRNA is locally translated in dendrites, where local translation of specific mRNAs is induced by synaptic stimuli [110]. This suggests that NMD activity might also be regulated in the context of synaptic function. Our data show that dendritic UPF3B protein is upregulated when synaptic channels are selectively treated with the group I mGluR agonist DHPG, a well-known synaptic stimulus (Figure S12). Activation of mGluRs induces the local translation of Arc [21], directing Arc mRNA to NMD in dendrites. NMD induction is thus likely to modulate the impact of synaptic activity on GluR1 internalization by limiting the amount of Arc protein. It is not known whether synaptic activity induces local synthesis of AMPK to suppress GluR1 synthesis in the long term. If that were the case, NMD would also limit the amount of AMPK to positively regulate total GluR1 levels upon synaptic activity.
STAU1-mediated mRNA decay (SMD) has been suggested as a competitive pathway to NMD in mammalian cells [111]. When bound to the 3’UTR, STAU1 can elicit translation-dependent mRNA degradation. STAU1 is known to associate with dendritic mRNA transport granules. Like NMD, numerous neuronal mRNAs may be natural SMD targets. Thus, mRNA decay in dendrites might be a broader phenomenon that serves as a natural brake to influence local translation, which is in turn required for synaptic plasticity.
Conclusions
Synaptic plasticity requires the selective translation of specific mRNAs targeted to dendrites. If this process is disrupted neurodevelopmental disorders may arise. Our experiments provide fundamental support for the idea that NMD-mediated mRNA decay modulates dendritic mRNA availability and therefore synaptic plasticity, learning, and memory. Through a number of locally-confined experiments in microfluidic devices and behavioral/LTP assays in neuron-restricted UPF2 conditional knockout mice, we define a specific role for the NMD pathway in dendrites and synaptic function (see schematic model in Figure S13). Therefore, our paper provides novel insights into basic mechanisms that regulate synaptic function that are also likely to play a role in NMD-implicated diseases such as autism and schizophrenia.
Author contributions
M.N. performed and/or analyzed most of the experiments including behavioral assays, and manuscript preparation/writing. M.A. prepared all neuronal cultures. F.L. performed electrophysiology experiments. N.V. provided technical assistance. M.T. provided infrastructure support. N.L.J. designed and manufactured microfluidic devices. E.K. supervised electrophysiology experiments. D.C. initiated and conceived the project; designed, analyzed, and supervised experiments; and wrote the manuscript. All authors contributed to the experimental design and interpretation and commented on the manuscript.
Methods
Mice and constructs
Hippocampal neuronal cultures were prepared from C57BL/6J mouse embryos (Charles River Laboratories). UPF2-shRNA 1 virus is on a piLenti-shRNA-GFP backbone carrying one shRNA against the UPF2 mRNA (AGGCGTATTCTGCACTCTAAAGGCGAGCT). UPF2-shRNA 2 virus is on a piLenti-shRNA-GFP backbone carrying four shRNAs against the UPF2 mRNA (TGAAAGACTATGTTATTTGTTGTATGATA). In experiments with UPF2 deletion in excitatory neurons, a UPF2 conditional knockout mouse line, which carries flanked loxP sites in the second exon of this gene, [1] was crossed with αCaMKII:CreERT2 line [2], and UPF2wt/wt;αCaMKII:CreERT2 (CTRL) or UPF2fl/fl;αCaMKII:CreERT2 (CKO) adults were used. To induce Cre, mice were treated with 200 mg tamoxifen/kg body weight every other day for five days. To outline dendrites and spines, an AAV-CaMKIIa-eGFP virus (Addgene; 50469-AAV5) was injected into the adult hippocampus of CTRL and CKO mice.
Hippocampal neuronal cultures
For culturing hippocampal neurons, we followed a previously described protocol with some modifications [3]. We typically isolated hippocampal neurons from embryos at embryonic day 16 (E16) and plated 1×105 neurons per coverslip (diameter: 12 mm) pre-coated with poly D-lysine (PDL) in Neurobasal medium containing 2% B27, 1 mM sodium pyruvate, 2 mM Glutamax, 30% D-glucose and penicillin (100 units/ml) /streptomycin (100 μg/ml).
Compartmentalized culture in microfluidic devices
Tri-partite microfluidic device was designed and manufactured by the Jeon lab. Microfluidic chambers were prepared in house as described previously described [4]. To isolate synaptic regions or to selectively treat the synaptic regions as well as post- or presynaptic cells with either pharmacological agents or sh/siRNAs, hippocampal neurons were cultured in tripartite microfluidic devices. The polymethyldimethylsiloxane (PDMS) devices were fabricated as previously described [4]. These devices have different channels separated by a physical barrier with embedded microgrooves (10 μm wide, 3 μm high). Unlike traditional microfluidic chambers with two channels [5], the tripartite PDMS chamber contains three channels (Figure 3). To access a high number of dendritic synapses, a ‘synaptic’ channel was placed 200 μm from the edge of one channel and 500 μm away from the other. Since the average length of dendrites extending into the microgrooves is less than 300 μm at 21 days in culture (DIV21), the closer “postsynaptic cell channel” allows a large number of dendrites to project from this channel to the new “synaptic” channel. On the other hand, the “presynaptic cell channel” that is located 500 μm away from the new channel ensures that only axons from this channel can extend into the synaptic channel. Fluidic isolation is established across the microgrooves, making it possible to isolate synaptic regions or to selectively treat the synaptic regions as well as post- or presynaptic cells. Reagents applied to synaptic channels were as follows: Cycloheximide (10 μM, Calbiochem), MG132 (10 μM, Sigma), and Leupeptin (10 μM, Sigma). siRNAs were applied to synaptic channels at different concentrations using 10% NeuroPORTER (Sigma) following manufacturer’s instructions.
For microfluidic cultures, we reversibly affixed the tripartite chambers to PDL-coated coverslips (Assistant; 50×24mm; two per coverslip) and then place them in 10 cm dishes. For culturing hippocampal neurons, we followed a previously described protocol with some modifications [3]. We typically isolated hippocampal neurons from embryos at embryonic day 16 and plate 1×105 neurons per coverslip (diameter: 12mm) pre-coated with poly D-lysine (PDL) in Neurobasal medium containing 2% B27, 1 mM sodium pyruvate, 2 mM Glutamax, 30% D-glucose and penicillin (100 units/mL) /streptomycin (100 μg/mL). We plated 8×104 neurons in cell-body channels and maintain these cultures in neuronal medium for up to 21 days. For immunostaining, we fixed neurons inside microfluidic chambers and continued with the staining procedure upon careful removal of chambers.
Immunohistochemistry
Hippocampal neuronal cultures were fixed in 4% paraformaldehyde (PFA)/PBS for 15 min, which was followed by 3 × 10 min washes in PBS. Standard immunohistochemistry was performed as previously described [4]. Briefly, samples were incubated overnight in PBS containing 1% Triton X-100, 10% normal goat serum and the relevant dilution of primary antibodies. The following morning, samples were washed with PBS (3 × 10 min) to remove any residual primary antibody. Secondary antibodies were incubated in PBS containing 1% Triton X-100 and 10% normal goat serum for 2 hr. Samples were subsequently washed with PBS (3 × 20 min) and mounted (Vectashield, Vector Laboratories). Primary commercial antibodies included AMPK (rabbit; Thermofisher, PA5-13797, PRKAG3 subunit), Arc (rabbit; Synaptic Systems, 156002), MAP2 (mouse; Abcam, ab11276), synapsin (rabbit; Abcam, ab64581) and UPF3B (rabbit; Sigma, HPA001882). Two commercial GluR1 antibodies were used: mouse GluR1 antibody (Abcam, ab174785) for stainings with synapsin and rabbit GluR1 antibody (Calbiochem, pc246) for the internalization assay. Anti-UPF1 and anti-UPF2 antibodies (both rabbit) were a gift from Jens Lykke-Andersen. Secondary antibodies were Alexa 488, 546, and 647 conjugated (Molecular Probes, 1:2000).
qRT-PCR
RNA was isolated from the synaptic channels of microfluidic devices by perfusion with TRIzol reagent (Invitrogen), with RNA being subsequently extracted from lysates following the manufacturer’s instructions. Purified RNA was inbcubated with DNase I (RNase-Free DNase Set, Qiagen) over a mini kit column (RNAeasy, Qiagen) to ensure DNA digestion. SuperScript III First-Strand Synthesis SuperMix (Invitrogen) was used to amplify cDNAs in a 20 μl reaction using 1 μg of RNA. Quantitative Real-Time PCR (qRT-PCR) reactions were performed as previously described [4] using the iQ SYBR Green Supermix (Bio-Rad) and an Eppendorf Mastercycler EP Realplex Thermocycler. For each reaction 10 ng cDNA was used. Relative expression levels were either normalized to GAPDH or Beta-Actin.
Primers used for qRT-PCR:
Arc Primers:
(FW) 5’CTGAGATGCTGGAGCACGTA3’
(REV) 5’GCCTTGATGGACTTCTTCCA3’
AMPK primers:
(FW) 5’ CAGGTCTACATGCACTTCATGC 3’
(REV) 5’ AAAGCTCTGCTTCTTGCTGTCC 3’
GluR1 primers:
(FW) 5’AGAGAAGAGGAGGAGAGCAG 3’
(REV) 5’ CTATCTGGATATTGTTGGGGA 3’
GAPDH primers:
(FW) 5’ ACATGGTCTACATGTTCC 3’
(REV) 5’ CAGATCCACAACGGAATAC 3’
Beta-Actin primers:
(FW) 5’ AGTGTGACGTTGACATCCGT 3’
(REV) 5’ TGCTAGGAGCCAGAGCAGTA 3’
Dicer-substrate siRNAs were acquired from Integrated DNA Technologies, and were predesigned using their proprietary algorithm which integrates design principles from 21mer siRNA theory with updated 27mer criteria. This design process ensures siRNA target sites do not match known single nucleotide polymorphisms or exons that are the product of alternative splicing, with all sequences having been pre-screened using Smith-Waterman analysis to reduce off-target binding.
Arc siRNA Cocktail
Duplex 1:
5’ CUGAUGGCUAUGACUAUACCGUUAG 3’
3’ UCGACUACCGAUACUGAUAUGGCAAUC 5’
Duplex 2:
5’ CUACAUGGACUGAACAUCAAGAAGC 3’
3’ ACGAUGUACCUGACUUGUAGUUCUUCG5’
AMPK siRNA Cocktail
Duplex 1:
5’ GACCUUGUUUCAAAUUGAAACAATA 3’
3’ UUCUGGAACAAAGUUUAACUUUGUUAU 5’
Duplex 2:
5’ CGAGAAAUUCAAAAUCUUAAACUCT 3’
3’ UUGCUCUUUAAGUUUUAGAAUUUGAGA 5’
UP3B siRNA Cocktail
Duplex 1:
5’ AAAUUGAAGCCAAAAAUCGAGAATT 3’
3’ CCUUUAACUUCGGUUUUUAGCUCUUAA 5’
Duplex 2:
5’ GGGUCAAGAAUAUCAUGCUAUAGTA 3’
3’ UUCCCAGUUCUUAUAGUACGAUAUCAU 5’
Fluorescent in-situ hybridization (FISH) for UPF3B mRNA
For the detection of UP3B mRNA, a mix of five non-overlapping antisense probes were used. Each oligonucleotide was 49 base pairs in length. FISH was visualized using the Tyramide In Situ System (Perkin Elmer) as previously described [4]. Briefly, samples were pretreated with hybridization buffer (50% formaldehyde, 5 × SSC, 0.1% tween, 100 ug/ml tRNA [Roche], 50 ug/ml heparin [Sigma]) for 2 hr at 65°C. Riboprobes were (100 ng/μl) were next heated to 97°C for 7 min and left to cool on ice. Hybridization was performed overnight at 65°C, and was followed by a series of washes at room temperature in hybridization buffer (no tRNA or heparin, 2 × 30 min), hybridization buffer/PBT (0.1% tween in PBS, 2 × 30 min), and lastly just PBT (4 × 30 min). Samples were next blocked for 1 hr (Roche Blocking Solution) and incubated with anti-DIG-POD (Roche) 1:800 in PBT overnight at 4°C. The following morning, samples were washed in PBT (3 × 20 min). Detection was visualized with tyramide-488 1:50 in supplied amplification buffer from the TSA-Plus in situ kit (Perkin Elmer).
Oligonucleotides used for antisense UPF3B probes
1-GGAAGAAAAAGATCATAGGCCTAAGGAGAAACGAGTGACCCTGTTTACG
2-GTCAAGAATATCATGCTATAGTAGAATTTGCACCATTTCAAAAAGCTGC
3-AAGAAAAGAGAGAAGAAAGGAGGAGACGGGAAATAGAGAGGAAAAGGCA
4-AAGAAAGAGCCAGTGGGCACAGTTATACTCTGCCCAGGCGTTCTGATGT
5-GGCACTTCGAGATAAAGGAAAGAAGAGTGAGAATACAGAATCAATATGC
GluR1 internalization assay
GluR1 internalization experiments were carried-out and quantified as previously described [6, 7]. Briefly, we incubated live DIV21 hippocampal cultures infected with control or UPF2 shRNAs at 37°C for 5 min with GluR1 antibody (Calbiochem, PC246) to allow labeling of surface GluR1. After washing, we incubated neurons at 37°C for 10 min to allow for basal internalization. We then fixed neurons with 4% PFA in PBS and blocked in a detergent-free blocking solution for 1 hr. Following blocking, we incubated neurons with AlexaFluor546 secondary antibody for 1 hr to label the surface population. To label the pre-labeled internalized fraction, we post-fixed neurons with 100% methanol at −20°C for 1 min and stained with Alexa-647 secondary antibody. We calculated the percent internalization by dividing the integrated far-red intensity (internalized GluR1) by the total (red + far red) intensity between control and NMD-deficient neurons.
Click chemistry and SDS-page
To measure the dendritic translation of GluR1, we used a click chemistry based approach and labeled locally synthesized GluR1 at synaptic regions. At DIV21, we exchanged the growth media in the synaptic channels to methionine-free medium containing the methionine-analog azidohomoalanine (AHA; 25 μM) for 6 hr. AHA is incorporated into all newly synthesized proteins. After harvesting the material from the synaptic channels in RIPA, we incubated the lysate with DBCO-biotin [8] (30 μM; Click Chemistry Tools LLC) for 1 hr at ambient temperature. DBCO reacts with AHA [9], functionalizing newly synthesized proteins with biotin. We purified biotin-containing proteins with Dynabeads (150 μl per reaction; Dynabeads M-280 Streptavidin-Thermo Fisher Scientific). Dynabeads were separated over a magnetic rack and subsequently washed 2 times in PBS containing 0.1% Bovine Serum Albumin (BSA). Proteins were dissociated and denatured from beads by boiling Dynabeads for 5 min in 0.1% SDS. Lysates were subsequently analyzed for protein concentration via BCA Protein Assay (Thermofisher). Proteins were separated via SDS-PAGE, as previously described [10]. Briefly, protein aliquots were mixed with an equal volume of loading buffer, denatured at 95°C and loaded onto 8% gels before being subject to SDS-PAGE. Proteins were subsequently transferred to nitrocellulose membrane, incubated in 5% BSA diluted in TBST for 1hr and then incubated with primary anti-C-terminus GluR1 antibody overnight (Abcam, ab31232). Membranes were washed in TBST (3 × 5 min), incubated for 1 hr in IRDye 800CW goat anti-rabbit secondary (Licor) diluted in 5% low-fat milk diluted in TBST, and bands resolved on a Licor Odyssey CLx imager.
Electrophysiology
The electrophysiology experiments were performed as previously described (114). All experiments were performed on transvers hippocampal slices (400 μm) from 3-month old (LTP studies) or 6-week-old (LTD studies) UPF2fl/fl;αCaMKII:CreERT2 mice and their control littermates (UPF2wt/wt;αCaMKII:CreERT2) following induction of Cre expression by tamoxifen (Sigma) at 2-months (for LTP) and 1-month (for LTD) of age. Given that Tamoxifen is fat soluble, we prepared our tamoxifen solution by dissolving the relevant quantity of drug into sunflower oil with 1:10 ETOH added. Tamoxifen solutions were incubated at 37°C with rocking for 6-8 hr. Tamoxifen was administered via oral gavage at 200 mg tamoxifen/kg body weight every other day for five total doses. This tamoxifen regimen led to successful ablation of UPF2 protein in the adult brain (Figure S1a). Slice preparation and aCSF composition were performed as described previously [11, 12]. Briefly, slices were isolated and transferred to recording chambers (preheated to 32°C), where they were superfused with oxygenated aCSF. We recorded field excitatory postsynaptic potentials (fEPSPs) from the stratum radiatum of CA1 using microelectrodes filled with artificial cerebrospinal fluid (ACSF) (resistance 1-4 MΩ). A bipolar Teflon-coated platinum electrode was placed in the stratum radiatum of CA3 to activate Schaffer collateral/commissural afferents at 0.05 Hz. Slices recovered in the recording chamber at least 1 hr before recordings began. In all experiments, basal field excitatory postsynaptic potentials (fEPSPs) were stable for at least 20 min before the start of each experiment. LTP was induced with either one (E-LTP) or three 1 sec 100-Hz high-frequency stimulation (HFS) trains, with an intertrain interval of 60 sec. To induce mGluR-LTD, slices were incubated with DHPG (100 μM) for 10 min. After induction of either E-LTP or mGluR-LTD, we collected fEPSPs for an additional 60 min (180 min for L-LTP). Slope values were compared from the conditional NMD mutant mice and their control littermates.
Stereotaxic injection of virus and CA1 spine density analysis
To measure hippocampal spine density, the spine density of CA1 hippocampal neurons in control (CTRL: UPF2wt/wt;αCaMKII:CreERT2) and Conditional Knockout (CKO: UPF2fl/fl;αCaMKII:CreERT2) mice was examined post-labelling with GFP-expressing virus. To target the dorsal CA1 field, the stereotaxic coordinates AP: −2, ML: 1.6, DV: 1.5-.25 were used. 1 μl of AAV-CaMKIIa-eGFP (Addgene, #50469-AAV5) virus was injected bilaterally using a 10 μl nanofil syringe (World Precision Instruments) fitted with a 33-gauge beveled needle. Post-surgery, wounds were closed with tissue adhesive (Vetbond, 3M). Mice were transcardially perfused 7 days later with 4% PFA (without PBS flush) brains removed, sectioned at 200 μm and apical CA1 dendrites imaged on an Olympus FluoView-FV1000 confocal microscope. All spine quantifications were completed in ImageJ.
Behavioral assays
To study the role of NMD in cognitive function, we induced Cre in 2-month old UPF2 CKO mice as well as in their control littermates by tamoxifen. This experimental design ensured that phenotypes are not caused by an acute effect of tamoxifen, or a non-specific effect of Cre at cryptic LoxP sites. A 2-week washout period followed tamoxifen dosing before behavior began, at approximately 3-months of age. The Y-maze [13, 14], fear conditioning [14, 15], Morris water maze [16, 17], marble burying [18], grooming (12), locomotor hyperactivity/open field [19], elevated-plus maze [19, 20] and three-chamber social interaction assay [21] were completed and analyzed as previously described, except where outlined below. Briefly, the Y-Maze comprised exposing mice to 2 of 3 arms of the maze for 10 min as a training trial. The third, blocked, arm of the maze was designated as the novel arm, and was pseudorandomized between mice. Following a 1 hr delay, mice were returned to the maze but were free to explore all 3 arms for 5 min. The time spent exploring the novel arm, relative to the other familiar arms, was quantified as our index of short-term spatial memory. To examine associative memory, we utilized a simple 2-day fear conditioned memory protocol. On day 1, mice were exposed to a 6 min conditioning trial that involved 3 tone-shock pairings. A 30-sec tone duration and 30 sec ISI was utilized, while all shock stimuli comprised a 0.7mA scrambled foot-shock that co-terminated with tone presentation. Learning was evaluated by plotting % freezing to each successive tone presentation as a time-course. On day 2, mice were returned to their conditioning context for 6 min, with % freezing over the test session being quantified as contextual fear memory. All experiments were completed in Colbourn fear chambers. The Morris water maze was used to test more sophisticated learning and longer-term spatial memory. Learning was measured by training mice to find a hidden platform (four trials per day, 75 sec/trial) over 4 consecutive days of testing. Trials on each day of testing were averaged, and plotted as a time-course, with gradually decreasing escape latencies being used as our index of spatial learning behavior. To assess spatial memory, we conducted a probe trial 24 hr after the last day of training on day 5. In this trial, the hidden escape platform was removed from the pool. Latency to enter the hidden platform zone, and time spent exploring the target quadrant, were quantified as measurements of spatial memory. Reversal learning was not evaluated as UPF2 CKO mice did not display adequate learning behavior, thus making it impossible to distinguish a general learning deficit from a reversal learning deficit even with addition of extra training trials. Assays were performed in order of least stressful (locomotor hyperactivity/open field) to most stressful (Morris water maze). Mice were bred and tested across 4-5 small cohorts, ensuring that all phenotypes were independent of idiosyncratic effects or nuisance variability that may contaminate any single day of testing.
Data analysis
Where total protein levels were measured via immunostaining, Image J was used to measure total fluorescence. In these analyses, a defined area containing dendrites were identified and the integrated fluorescent density was measured in each region. Background fluorescence was measured by selecting a region adjacent to dendrites, and was subtracted from positive fluorescent signal-measurements to derive corrected total fluorescence. Depending on the test, behavior was analyzed using a between-groups or mixed-model ANOVA with genotype and sex added as main effects. For tests that involved multiple observations, data were corrected using Tukey’s method. Consistent with prior research [10, 14, 22], as no significant interaction involving the main effect of sex was observed on any given behavioral assay - explicating no modulatory effect of sex on genotype-dependent effects - data from mice were pooled to increase power. No specific exclusion criteria were applied to the datasets, excepting the removal of outliers (defined as values falling outside of ± 2 S.D.). Significance was set at p=0.05 per Fisher’s tables. Data analysis was completed using the Graphpad Prism software package.
Acknowledgements
We thank J. Lykke-Andersen for generously providing the anti-UPF1 and anti-UPF2 antibodies, M.E. Ross for helpful comments and suggestions and M. Toth for helpful suggestions regarding behavioral assays. We also thank Nicole Volk, the research assistant of Colak lab, for technical help. This work was supported by a NHMRC CJ Martin Biomedical Fellowship awarded to M.N., KoreaNRF-2015R1A2A1A09005662 to N.L.J., NIH grants NS034007 and NS047384 to E.K, and NIH R01 MH114888 and Leon Levy Foundation Grants to D.C.
References
- 1.↵
- 2.↵
- 3.↵
- 4.↵
- 5.↵
- 6.↵
- 7.↵
- 8.↵
- 9.↵
- 10.↵
- 11.↵
- 12.↵
- 13.↵
- 14.↵
- 15.↵
- 16.↵
- 17.↵
- 18.↵
- 19.↵
- 20.↵
- 21.↵
- 22.↵
- 23.↵
- 24.↵
- 25.↵
- 26.↵
- 27.↵
- 28.
- 29.
- 30.↵
- 31.↵
- 32.↵
- 33.↵
- 34.↵
- 35.↵
- 36.↵
- 37.↵
- 38.↵
- 39.↵
- 40.↵
- 41.
- 42.↵
- 43.
- 44.
- 45.↵
- 46.
- 47.↵
- 48.↵
- 49.↵
- 50.↵
- 51.↵
- 52.↵
- 53.
- 54.↵
- 55.↵
- 56.↵
- 57.↵
- 58.↵
- 59.↵
- 60.↵
- 61.↵
- 62.↵
- 63.↵
- 64.↵
- 65.↵
- 66.↵
- 67.↵
- 68.↵
- 69.↵
- 70.↵
- 71.↵
- 72.↵
- 73.
- 74.
- 75.↵
- 76.↵
- 77.↵
- 78.↵
- 79.
- 80.
- 81.
- 82.
- 83.↵
- 84.↵
- 85.↵
- 86.↵
- 87.↵
- 88.↵
- 89.
- 90.
- 91.
- 92.
- 93.↵
- 94.↵
- 95.↵
- 96.↵
- 97.↵
- 98.
- 99.
- 100.↵
- 101.↵
- 102.↵
- 103.↵
- 104.↵
- 105.↵
- 106.↵
- 107.↵
- 108.↵
- 109.↵
- 110.↵
- 111.↵
References for Methods
- 1.
- 2.
- 3.
- 4.
- 5.
- 6.
- 7.
- 8.
- 9.
- 10.
- 11.
- 12.
- 13.
- 14.
- 15.
- 16.
- 17.
- 18.
- 19.
- 20.
- 21.
- 22.
Supplementary References
- 1.
- 2.
- 3.
- 4.
- 5.
- 6.
- 7.
- 8.
- 9.
- 10.
- 11.
- 12.
- 13.
- 14.
- 15.
- 16.
- 17.
- 18.
- 19.
- 20.
- 21.
- 22.
- 23.
- 24.
- 25.
- 26.
- 27.
- 28.
- 29.
- 30.
- 31.