The neuronal ceroid lipofuscinoses (NCLs) are a class of individually rare, primarily autosomal recessive, neurodegenerative diseases occurring in an estimated 2 to 4 of 100,000 live births (Nita et al., 2016). Collectively, NCLs represent the most prevalent class of hereditary pediatric neurodegenerative disease (Haltia, 2006). The NCLs are characterized by progressive neurodegeneration, blindness, cognitive and motor deterioration, seizures, and premature death. The cardinal feature of all NCLs is the intracellular accumulation of proteolipid material, termed lipofuscin (Jalanko and Braulke, 2009; Nita et al., 2016). While lipofuscin accumulates in all cells of affected individuals, it deposits most robustly in neurons. This accumulation is concurrent with rapid and progressive neurodegeneration, particularly of thalamic and primary sensory cortical areas (Bible et al., 2004; Kielar et al., 2007). The NCLs are categorized into CLN1-14 based on the age of onset and the causative gene mutated. The products of CLN genes are lysosomal and endosomal proteins, therefore NCLs are also classified as lysosomal storage disorders (LSDs) (Bennett and Hofmann, 1999; Jalanko and Braulke, 2009). The infantile form of disease, CLN1, presents as early as 6 months of age with progressive psychomotor deterioration, seizure, and death at approximately 5 years of age (Haltia, 2006; Jalanko and Braulke, 2009; Nita et al., 2016). CLN1 disease is caused by mutations in the gene CLN1, which encodes the enzyme palmitoyl-protein thioesterase 1 (PPT1)(Camp and Hofmann, 1993; Camp et al., 1994; Vesa et al., 1995; Jalanko and Braulke, 2009). PPT1 is a depalmitoylating enzyme responsible for the removal of palmitic acid from modified proteins (Camp and Hofmann, 1993; Lu and Hofmann, 2006).

Protein palmitoylation, the addition of a 16-carbon fatty acid (palmitic acid) to cysteine residues, is a crucial regulator of protein trafficking and function, particularly in neurons (Hayashi et al., 2005; Hayashi et al., 2009; Fukata et al., 2006; Kang et al., 2008; Fukata and Fukata, 2010; Han et al., 2015). This post-translational modification is mediated by palmitoyl acyltransferases (PATs) of the DHHC enzyme family (Fukata et al., 2006; Fukata and Fukata, 2010). In contrast to other types of protein acylation, palmitoylation occurs via a reversible thioester bond (s-palmitoylation), permitting dynamic control over target protein interactions and function. Further, palmitoylated proteins require depalmitoylation prior to lysosomal degradation (Lu et al., 1996; Lu and Hofmann, 2006). Consequently, protein palmitoylation and depalmitoylation contribute significantly to mechanisms underlying synaptic plasticity and endosomal-lysosomal trafficking of proteins (Hayashi et al., 2005; Hayashi et al., 2009; Kang et al., 2008; Lin et al., 2009; Noritake et al., 2009; Fukata and Fukata, 2010; Mattison et al., 2012; Thomas et al., 2012; Thomas et al., 2013; Fukata et al., 2013; Han et al., 2015). Indeed, PPT1 is a lysosomal depalmitoylating enzyme that localizes to the axonal and synaptic compartments (Verkruyse and Hofmann, 1996; Ahtiainen et al., 2003; Kim et al., 2008). The synaptic association of PPT1 and prominence of palmitoylated synaptic proteins suggests that PPT1 influences synaptic functions through, at least, protein turnover. Many synaptic proteins undergo palmitoylation, including, but not limited to postsynaptic density protein 95 (PSD-95), all GluA subunits of AMPARs, and the GluN2A/2B subunits of NMDARs (Kang et al., 2008). However, the role of depalmitoylation in regulating synaptic protein function remains less clear.

N-methyl-D-aspartate receptors (NMDARs) are voltage-dependent, glutamate-gated ion channels consisting of two obligatory GluN1 subunits and two GluN2 subunits that undergo a developmental change (Cull-Candy et al., 2001; van Zundert et al., 2004; Lau and Zukin, 2007; Paoletti et al., 2013). NMDARs play a crucial role in synaptic transmission, postsynaptic signal integration, synaptic plasticity, and have been implicated in various neurodevelopmental and psychiatric disorders (Lau and Zukin, 2007; Lakhan et al., 2013; Paoletti et al., 2013; Yamamoto et al., 2015). NMDAR subunit composition, receptor localization, and downstream signaling mechanism undergo developmental regulation (Watanabe et al., 1992; Monyer et al., 1994; Sheng et al., 1994; Li et al., 1998; Stocca and Vicini, 1998; Tovar and Westbrook, 1999; Losi et al., 2003; van Zundert et al., 2004; Paoletti et al., 2013; Wyllie et al., 2013). Specifically, GluN2B-containing NMDARs are expressed neonatally and display prolonged decay kinetics, which allows comparatively increased calcium influx thought to facilitate forms of synaptic plasticity critical for neurodevelopment (Sobczyk et al., 2005; Zhao et al., 2005; Zhao et al., 2013; Zhang et al., 2008; Evans et al., 2012; Shipton and Paulsen, 2014). These GluN2B-containing receptors are supplanted at the synapse by diheteromeric GluN1/GluN2A NMDARs or triheteromeric (GluN1/GluN2A/GluN2B) receptors in response to experience-dependent neuronal activity (Quinlan et al., 1999b; Quinlan et al., 1999a; Tovar and Westbrook, 1999; Philpot et al., 2001; Liu et al., 2004; Paoletti et al., 2013; Tovar et al., 2013). This developmental switch of GluN2B- to GluN2A-containing NMDARs during brain maturation is mediated by the postsynaptic scaffolding receptors, SAP102 and PSD-95, respectively; SAP102-GluN2B-NMDAR complexes are replaced by PSD-95-GluN2A-NMDAR complexes in response to developmental, experience-dependent activity (Sans et al., 2000; van Zundert et al., 2004; Elias et al., 2008). While PSD-95, GluN2B, and GluN2A all undergo palmitoylation, how depalmitoylation regulates the turnover of these proteins, let alone during the GluN2B to GluN2A subunit switch, is unclear.

In the current study, we investigated the cellular and synaptic effects of PPT1-deficiency using the Ppt1^-/- mouse model of CLN1 disease. We focused on the visual system in Ppt1^-/- animals for two reasons. First, cortical blindness is a characteristic feature of CLN1 disease. Second, the rodent visual system is a well-studied model of cortical development and synaptic plasticity/maturation and it therefore serves as an optimal experimental model to examine the role of PPT1-mediated depalmitoylation during development. We found that lipofuscin accumulated in the Ppt1^-/- visual cortex shortly after eye-opening at postnatal day (P) 14, a timing earlier than previously documented (Gupta et al., 2001). Using biochemistry and electrophysiology, we found impeded developmental NMDAR subunit switch from GluN2B to GluN2A in Ppt1^-/- mice compared to wild-type (WT). This NMDAR disruption is associated with disrupted dendritic spine morphology in vivo. To gain further mechanistic insight into neurodegeneration in CLN1, we used cultured cortical neurons and found that Ppt1^-/- cells recapitulate the disrupted dendritic spine phenotype and GluN2B to GluN2A switch, leading to excessive extrasynaptic calcium transients and enhanced vulnerability to NMDA-mediated excitotoxicity. We directly examined protein palmitoylation state and found hyperpalmitoylation of GluN2B as well as Fyn kinase, which facilitates GluN2B surface retention, in Ppt1^-/- neurons. Finally, we demonstrate that chronic treatment of Ppt1^-/- neurons with palmitoylation inhibitors normalized GluN2B and Fyn kinase hyperpalmitoylation and rescued the enhanced susceptibility to excitotoxicity. Our results indicate that PPT1 plays a critical role in the developmental GluN2B to GluN2A subunit switch and synaptic maturation. Further, our results indicate that these dysregulated mechanisms contribute to CLN1 pathophysiology and may be shared features of common adult-onset neurodegenerative diseases.

To understand synaptic dysregulation in CLN1 disease, we utilized the visual cortex of Ppt1^-/- animals as a model system. The rodent visual cortex undergoes timed, experience-dependent plasticity, which has been well-characterized at the systemic, cellular, and molecular levels (Bear et al., 1990; Gordon and Stryker, 1996; Hensch et al., 1998; Quinlan et al., 1999a; Fagiolini and Hensch, 2000; Mataga et al., 2001; Mataga et al., 2004; Philpot et al., 2001; Desai et al., 2002; Yoshii et al., 2003; Hensch, 2005; Cooke and Bear, 2010). We examined WT and Ppt1^-/- littermates at the following ages: P11, P14, P28, P33, P42, P60, P78, P120, which correspond to particular developmental events in visual cortex. In mice, P11 and P14 are prior to and just after eye opening (EO), respectively. Further, the critical period in the visual cortex peaks at P28 and closes from P33 to P42. We chose postnatal day 60, P78, and P120 were selected as adult time points. We determined whether experience-dependent synaptic maturation is altered during the progression of CLN1 pathology.

Lipofuscin deposits immediately following eye opening in visual cortex of Ppt1^-/- mice

Although it remains controversial whether lipofuscin is toxic to neurons or an adaptive, neuroprotective mechanism, its accumulation correlates with disease progression. Therefore, we examined lipofuscin deposition in the visual cortex as a marker of pathology onset and progression. Lipofuscin aggregates are readily detectable as autofluorescent lipopigments (ALs) without staining under a confocal microscope. To examine the temporal and spatial accumulation of ALs in Ppt1^-/- mice, we performed quantitative histology on the visual cortex (area V1) of WT and Ppt1^-/- mice during early development. Visual cortical sections were imaged at the above-mentioned developmental time points, and ALs were quantified in a laminar-specific manner. We found that ALs are detectable first at P14 in Ppt1^-/- visual cortex, earlier than previously reported at 3 or 6 months (Figure 1A–C)(Gupta et al., 2001; Blom et al., 2013). Further, ALs accumulated rapidly through the critical period (Berardi et al., 2000; Hensch, 2005; Maffei and Turrigiano, 2008) and plateaued by adulthood (P60) (Figure 1A–C). This result suggests that neuronal AL load is saturable, and that this saturation occurs early on in disease, as Ppt1^-/- animals do not perish until around 10 months of age.

Figure 1

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Whether lipofuscin accumulation is directly neurotoxic or not, profiling the temporospatial and sub-regional pattern of AL deposition will be valuable for assessing therapeutic interventions in future studies. The pattern of deposition revealed herein suggests a correlation between systemic neuronal activation and AL accumulation, as we found that AL deposition started immediately following EO, the onset of patterned visual activity, and accumulated rapidly during development (Figure 1A,C, Supplementary file 1). These findings suggest that neuronal activity or experience-dependent plasticity may be linked to lipofuscin deposition.

NMDAR subunit composition is biased toward immaturity in Ppt1^-/- visual cortex

To examine the role of PPT1 in excitatory synapse function, we focused on the NMDAR subunits, GluN2B and GluN2A, which are both palmitoylated (Hayashi et al., 2009). The developmental GluN2B to GluN2A subunit change (Paoletti et al., 2013) is critical for NMDAR function and maturation, which facilitates refinement of neural circuits and a higher tolerance to glutamate-mediated excitotoxicity (Hardingham and Bading, 2002; Hardingham and Bading, 2010; Hardingham et al., 2002). Furthermore, previous work shows evidence for NMDA-induced excitotoxicity in the pathogenesis of CLN1 (Finn et al., 2012). We biochemically analyzed WT and Ppt1^-/- visual cortices from P11 to P60 and measured levels of GluN2B and GluN2A subunits in synaptosomes and whole lysates of WT and Ppt1^-/- visual cortices. Although GluN2B levels were comparable between WT and Ppt1^-/- at all ages, GluN2A levels in synaptosomes were significantly lower in Ppt1^-/- than WT (Figure 2A). This decrease was present at time points during, and just following, the critical period in visual cortical development (P33, P42, and P60). When analyzed as a ratio of GluN2A/GluN2B, a robust and persistent decrease is observed in Ppt1^-/- visual cortex (Figure 2B). GluN1 levels were unchanged between WT and Ppt1^-/- in synaptosomes (Figure 2C), indicating the selective obstruction of GluN2A incorporation into NMDARs.

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The developmental shift from GluN2B-containing NMDARs to synaptic GluN2A-containing NMDARs is mediated by the postsynaptic scaffolding proteins, SAP102 and PSD-95 (Townsend et al., 2003; van Zundert et al., 2004; Elias et al., 2008). SAP102 preferentially interacts with GluN2B-containing NMDARs, which are enriched neonatally (Sans et al., 2000; van Zundert et al., 2004; Zheng et al., 2010; Chen et al., 2011). In contrast, PSD-95 has greater affinity to GluN2A-containing NMDARs, particularly in the mature brain (Sans et al., 2000; van Zundert et al., 2004; Dongen, 2009; Yan et al., 2014). Thus, we examined the expression of these scaffolding proteins in WT and Ppt1^-/- visual cortex. While SAP102 levels remained unchanged, PSD-95 levels reduced at P33-P60, the same developmental time points where GluN2A expression also decreased (Figure 2D). Together, these results suggest reduced incorporation and scaffolding of GluN2A-containing NMDARs in Ppt1^-/- synapses, indicating immature or dysfunctional synaptic composition.

Next, we measured PPT1 protein level across the same time points in WT animals to examine whether the expression profile of PPT1, and presumably its cellular activity, temporally correlated with the observed reductions in mature synaptic components in Ppt1^-/- animals. Indeed, PPT1 expression in synaptosomes is low at P11 and P14 and increases with age, reaching peak levels between P33 and P60 (Figure 2E). This expression profile correlates with the time course of AL accumulation (Figure 1B and C) and fits with the notion that PPT1 activity at the synapse plays a role in neurodevelopmental processes.

To examine whether the reduction in GluN2A is due to selective exclusion from the postsynaptic site or alterations in the total protein amount, we also measured NMDAR subunit levels in whole lysates. These findings closely match our findings in synaptosomes. Namely, GluN2A levels showed reductions in Ppt1^-/- lysates beginning at the same time point (P33) (Figure 2—figure supplement 1A), while GluN2B levels were stable (Figure 2—figure supplement 1B). The GluN2A/2B ratios in Ppt1^-/- whole lysates were also lower than those in WT lysates and the reduction was comparable to that observed in synaptosomes (Figure 2—figure supplement 1C). GluN1 levels, however, were unaltered between genotypes (Figure 2—figure supplement 1D), again indicating a selective reduction in the expression of GluN2A. Interestingly, while PPT1 levels in whole lysates were, similarly to synaptosomes, low at P11 and P14, expression subsequently peaked at P28 and P33 before declining at P60 (Figure 2—figure supplement 1E). Together, these results indicate a selective decrease in the total amount of mature synaptic components in Ppt1^-/- brains that temporally correlates with the cellular PPT1 expression profile in developing WT neurons. Furthermore, our findings in whole lysates suggest that synaptosomal reductions in GluN2A and PSD-95 may result from altered transcription or translation of these proteins instead of direct depalmitoylation by PPT1.

NMDAR-mediated EPSCs are altered in Ppt1^-/- visual cortex

Next, we sought to correlate our biochemical findings with electrophysiological changes in NMDAR functionality (Figure 2). While human CLN1 patients present with retinal degeneration and the Ppt1^-/- mouse model of CLN1 phenocopies the human disease, the electroretinogram (ERG) is effectively unaltered at 4 months in the mouse model (Lei et al., 2006), allowing for detailed study of the electrophysiological changes in the visual cortex associated with early disease states. We recorded evoked, NMDAR-mediated excitatory postsynaptic currents (EPSCs) in layer II/III cortical neurons in visual cortical slices of WT and Ppt1^-/- mice at P42. The NMDAR-EPSCs were pharmacologically isolated (see Materials and methods section) and were recorded in whole cell patch mode clamped at +50 mV. As GluN2A- and GluN2B-containing NMDARs exhibit differential receptor kinetics, with GluN2A displaying fast (~50 ms) and GluN2B displaying slow decay kinetics (~300 ms), their relative contribution is reliably interpolated by fitting the EPSC decay phase with a double exponential function (Stocca and Vicini, 1998; Vicini et al., 1998). From the fitting of absolute amplitude-normalized, WT and Ppt1^-/- NMDAR-EPSCs (Figure 3A), we measured the following parameters: the ratios of the amplitudes (A) of the fast, A_f/A_f + A_s, and slow, A_s/A_f + A_s components, and the weighted decay time constants (τ_w). The fast component (A_f/A_f +A_s) of NMDAR-ESPC amplitudes decreased in Ppt1^-/- mice as compared to WT, while the slow component (A_s/A_f +A_s) increased (Figure 3B). Further, Ppt1^-/- neurons showed a significant increase in weighted decay time τ_w as compared to WT (Figure 3C). Remarkably, the rise time (time to peak amplitude) of Ppt1^-/- NMDAR-EPSCs was slightly but significantly longer than WT (Figure 3D), suggesting that the response involves the receptors more distant from the presynaptic release site. Indeed, previous studies documented similar observations and postulated that a longer rise time is characteristic of GluN2B-containing NMDARs that are preferentially localized on the extrasynaptic membrane (Townsend et al., 2003; van Zundert et al., 2004; Sanz-Clemente et al., 2013).

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Next, we treated cortical slices with Ro 25–6981, a potent and selective inhibitor of GluN2B-containing NMDARs (Fischer et al., 1997) and asked if these receptors are overrepresented in Ppt1^-/- neurons. We recorded NMDAR-EPSCs at baseline and during bath infusion of Ro 25–6981 (30 min, 3μM), then compared the percent inhibition (τ_w percent of baseline) between WT and Ppt1^-/- groups. Ro 25–6981 treatment significantly decreased the τ_w of NMDAR-EPSCs from the baseline in both WT and Ppt1^-/- cells at P42 (Figure 3E). To our surprise, no significant effects were present between the two genotypes after Ro 25–6981 treatment by two-way ANOVA, suggesting that NMDARs are inhibited to the same degree in WT and Ppt1^-/- cortices.

At first glance, the above result did not fulfill the anticipation that NMDA-EPSCs in Ppt1^-/- neurons would respond to Ro 25–6981 treatment to a greater degree than in WT, due to an overrepresentation of GluN2B-containing receptors. However, cortical neurons predominantly express NMDARs consisting of two GluN1, one GluN2A and one GluN2B subunits (Sheng et al., 1994; Luo et al., 1997; Tovar and Westbrook, 1999). These triheteromeric NMDARs display prolonged decay kinetics compared to GluN2A-diheteromeric NMDARs, while being largely insensitive to GluN2B-specific antagonists (Stroebel et al., 2018). Indeed, the fast component of the amplitude is reduced in Ppt1^-/- neurons (Figure 3B), indicating a functional decrease in the contribution of GluN2A to NMDAR-mediated EPSCs in these cells. Moreover, analysis of the weighted decay time constant (Figure 3C) suggests a larger contribution of GluN2B to the overall NMDAR-EPSC in Ppt1^-/- cells (Stocca and Vicini, 1998; Vicini et al., 1998). Thus, our findings suggest an enhanced incorporation of triheteromeric NMDARs at Ppt1^-/- synapse, and corroborate our biochemical findings (see Discussion). Collectively, our data indicate a functionally immature NMDAR phenotype in Ppt1^-/- layer II/III visual cortical neurons.

Dendritic spine morphology is immature in Ppt1^-/- visual cortex

The morphology of dendritic spines is dynamic and modified by synaptic plasticity (Engert and Bonhoeffer, 1999; Parnass et al., 2000; Yuste and Bonhoeffer, 2001; Matsuzaki et al., 2004). During visual cortical development that is concomitant with the GluN2B to GluN2A switch, dendritic spine morphology undergoes robust structural plasticity at excitatory synapses. Dendritic spines contribute to experience-dependent synaptic plasticity via the generation, maturation, and long-term stabilization of spines, ultimately giving rise to established synaptic circuits. Typically, by P33, dendritic spines begin to demonstrate a reduction in turnover and an increase in mushroom-type spines, indicating synaptic maturity. Importantly, dendritic spines are morphologically disrupted in many neurodevelopmental disorders, typically skewing toward an immature phenotype (Purpura, 1979; Irwin et al., 2001; Penzes et al., 2011).

We hypothesized that dendritic spine morphology is immature or disrupted in Ppt1^-/- neurons, particularly given that GluN2A subunit incorporation is disrupted in vivo. Thus, we used in utero electroporation to sparsely label layer II/III cortical neurons in the visual cortex using a GFP construct (Matsuda and Cepko, 2004). GFP-expressing cells from WT and Ppt1^-/- animals were imaged for detailed analysis of dendritic spine morphology (spine length, spine volume, and spine head volume) at P33, a time point when dendritic spine morphology is typically considered mature and GluN2A is reduced at Ppt1^-/- synapses.

We analyzed dendritic spine characteristics of GFP-expressing cells (procedure schematized in Figure 4A) from WT and Ppt1^-/- visual cortex (Figure 4B) using the Imaris software (Bitplane). While WT neurons exhibited mushroom-type spine morphology with high-volume spine heads (Figure 4C, arrows), Ppt1^-/- neurons showed longer, filopodial protrusions or stubby spines (Figure 4C, arrowheads). Quantification of spine length and spine volume demonstrated that Ppt1^-/- spines were longer and less voluminous compared to WT (Figure 4D–E). Further, the volume of dendritic spine heads was reduced in Ppt1^-/- neurons (Figure 4E, inset). Interestingly, dendritic spine density was significantly increased in Ppt1^-/- neurons, signifying dysregulated synapse formation or refinement in the Ppt1^-/- brain (Figure 4F). These data indicate that dendritic spine morphology is disrupted in the developing CLN1 visual cortex, corresponding with the finding that NMDAR composition is immature at P33 and suggesting a reduced ability to compartmentalize calcium and other localized biochemical signals in CLN1.

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NMDAR subunit composition and dendritic spine morphology are also immature in Ppt1^-/- primary cortical neurons

The GluN2B to GluN2A switch and maturation of dendritic spine characteristics in WT primary neurons has been previously demonstrated (Williams et al., 1993; Zhong et al., 1994; Papa et al., 1995). We established that the developmental switch from GluN2B- to GluN2A-containing NMDARs and dendritic spine morphology are impaired in the Ppt1^-/- mouse brain. To understand these mechanisms more comprehensively and examine protein palmitoylation more directly, we used dissociated neuronal cultures. First, we analyzed these developmental events in WT and Ppt1^-/- primary cortical neurons to determine whether the biochemical and structural features of disease are recapitulated in vitro.

We collected lysates from cultured cortical neurons for 7, 10, or 18 days in vitro (DIV 7, 10, or 18) harvested and performed immunoblot analyses for markers of immature (GluN2B) or mature (GluN2A, PSD-95) excitatory synapses. Expression of GluN2B clearly preceded that of mature synaptic markers, peaking in both WT and Ppt1^-/- neurons at DIV10 and decreasing slightly thereafter (Figure 5A). In contrast, levels of both GluN2A and PSD-95 remained low until DIV18, at which point expression was robust (Figure 5B,C). Importantly, GluN2A, PSD-95, and GluN2A/GluN2B ratio levels showed reductions in Ppt1^-/- neurons compared to WT at DIV18, indicating that the biochemical phenotype is recapitulated to an extent in vitro (Figure 5B–D).

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To analyze dendritic spine morphology, we transfected primary cortical neurons from fetal WT and Ppt1^-/- mice with the GFP construct as above (Matsuda and Cepko, 2004) and cultured until DIV 15 or 20, then performed live cell imaging (Figure 6A). We measured dendritic spine length and volume in transfected cells using the Imaris software (Bitplane). At both DIV 15 and 20, we observed a significant alterations in the dendritic spine length (Figure 6B–E). The distribution of spine length in Ppt1^-/- neurons at DIV15 significantly shifted toward longer protrusions as compared to WT cells (Figure 6B). The averaged spine length was also robustly increased in Ppt1^-/- neurons (Figure 6C). Similar changes were also present at DIV20 (Figure 6D–E). Next, we analyzed differences in dendritic spine volume (Figure 6F–I). Ppt1^-/- neurons exhibit a significant reduction in the percentage of spines with volumes greater than ~0.2 μm³ at both DIV15 (Figure 6F) and DIV20 (Figure 6H). The averaged spine volume was also reduced in Ppt1^-/- neurons at both DIV15 (Figure 6G) and DIV20 (Figure 6I). As observed in vivo, Ppt1^-/- neurons showed an increase in the dendritic spine density at DIV15 (Figure 6J) and DIV20 (Figure 6K), again suggesting aberrant synapse formation, a failure of synaptic pruning, or both in Ppt1^-/- neurons. Together, these data demonstrate that Ppt1^-/- neurons in culture give rise to morphologically immature dendritic spines and corroborate our in vivo findings.

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Calcium imaging reveals extrasynaptic calcium dynamics in Ppt1^-/- neurons

Intracellular calcium dynamics, compartmentalization, and signaling play a critical role in synaptic transmission and plasticity. These properties are altered by glutamate receptor composition and location (Lau and Zukin, 2007; Hardingham and Bading, 2010; Paoletti et al., 2013). GluN2B-containing NMDARs maintain a prolonged open conformation compared to GluN2A-containing receptors, allowing increased calcium entry per synaptic event (Sobczyk et al., 2005). Moreover, previous studies indicate that GluN2A-containing NMDARs are generally inserted in the PSD, whereas GluN2B-containing NMDARs are localized extrasynaptically and associated with SAP102 (Tovar and Westbrook, 1999; Townsend et al., 2003; van Zundert et al., 2004; Washbourne et al., 2004; Groc et al., 2007; Elias et al., 2008; Martel et al., 2009). To determine more directly the effects of our biochemical and electrophysiological findings on calcium dynamics, we analyzed calcium signals in WT and Ppt1^-/- neurons transfected with the genetically encoded calcium sensor, GCaMP3 (Tian et al., 2009).

While WT neurons exhibited primarily compartmentalized calcium signals that were restricted to individual spines (Figure 7A–C, left, see Video 1), Ppt1^-/- neurons demonstrated diffuse calcium influxes that spread through the dendritic shaft (Figure 7A–C, right, see Video 2). These extrasynaptic transients appear rarely in WT cells (Figure 7, see Videos). To analyze the calcium dynamics in more detail, measurements of ΔF/F₀ were made for each dendritic segment, from each cell over the course of the captured videos (see Materials and methods). Multiple transients from the same synaptic site are shown as a heat map of ΔF/F₀ measurements and they are largely consistent across time in both WT and Ppt1^-/- neurons (Figure 7B). Further, plotting of the averaged ΔF/F₀ transients at an individual synaptic site demonstrates that local fluorescence increases in WT cells are confined to a short distance from the peak ΔF/F₀ at synaptic sites (Figure 7B and C, left), while those of Ppt1^-/- neurons diffuse longer distances within the dendrite (Figure 7B and C, right). To quantitatively compare these properties, we performed measurements of area under the curve (AUC) and calcium diffusion distance (see shaded region in Figure 7C) for each synaptic site from WT and Ppt1^-/- neurons. These analyses revealed a robust increase in both the AUC (Figure 7D) and the calcium diffusion distance (Figure 7E) in Ppt1^-/- neurons compared to WT. Furthermore, performing correlation analysis of calcium events across time (see Materials and methods) within a given neuron demonstrates that calcium influxes are more synchronous (increased correlation coefficient) in Ppt1^-/- neurons compared to WT (Figure 7F). This result may involve mechanisms underlying synaptic cluster plasticity, including synaptic integration via translational activation influenced by excessive Ca²⁺ entry (Govindarajan et al., 2006), enhanced biochemical crosstalk between synapses by, for example, small GTPases (which are generally palmitoylated proteins) (Harvey et al., 2008), or direct cooperative multi-synaptic Ca²⁺ signaling (Weber et al., 2016) in Ppt1^-/- neurons. Together, these data indicate that calcium entry and dispersion are enhanced at Ppt1^-/- synapses in vitro.

Figure 7

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These data are in line with our biochemical and electrophysiological findings and suggest that GluN2B-containing NMDARs mediate the observed calcium signals. To further test this possibility, we next treated WT and Ppt1^-/- neurons with Ro 25–6981 (1 μM, added in imaging medium following 2.5 min imaging at baseline) and performed calcium imaging. Ro 25–6981 had virtually no effect on calcium signals recorded from WT cells (Figure 7G–I, see Video 3). In contrast, Ppt1^-/- neurons treated with Ro 25–6981 showed a reduction in dendritic calcium influxes within shafts, while few residual, compartmentalized transients persisted (Figure 7G–I, see Video 4). Quantitatively, both AUC (Figure 7H) and calcium diffusion (Figure 7I) distance were rescued to WT levels following Ro 25–6981 treatment of Ppt1^-/- neurons. Together, these data suggest that Ppt1^-/- neurons have extrasynaptic calcium signaling compared to WT that is sensitive to GluN2B-NMDAR blockade.

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Ppt1^-/- cultured neurons show enhanced vulnerability to NMDA-mediated excitotoxicity

GluN2B-predominant NMDARs are implicated in enhanced neuronal susceptibility to NMDA-mediated neuronal death (Martel et al., 2009; Martel et al., 2012). Our results from biochemical, electrophysiological, and live-imaging analyses indicate decreased GluN2A/2B ratio suggesting an intriguing possibility that Ppt1^-/- neurons are more vulnerable to excitotoxicity (Finn et al., 2012). Therefore, we treated WT and Ppt1^-/- cultured neurons with NMDA (varying doses, 10–300 μm) and glycine (1–30 μm, always in 1:10 ratio with NMDA) for 2 hr and assayed cell viability 24 hr later using the PrestoBlue reagent (ThermoFisher Scientific) (Figure 8A). As expected, both WT and Ppt1^-/- neurons demonstrated dose-dependent reductions in cell viability in response to increasing concentrations of NMDA/glycine (Figure 8B). Importantly, Ppt1^-/- neurons were more vulnerable to NMDA insult, as exposure to 10 μM NMDA was sufficient to reduce cell viability significantly in Ppt1^-/- neurons but not WT cells (WT = 93 ± 4.1%; Ppt1^-/- = 76 ± 3.5%; *p=0.046; Figure 8B). Further, at 100 μM NMDA, WT neuron viability decreased by 35%, while Ppt1^-/- neuron viability was reduced significantly further, by 58% (WT = 65 ± 1.8%; Ppt1^-/- = 42 ± 4.5%; **p=0.0043; Figure 8B). At 300 μM NMDA treatment this effect plateaued, as cell viability between WT and Ppt1^-/- neurons was comparable (Figure 8B). These results indicate Ppt1^-/- neurons are more vulnerable to excitotoxicity and are consistent with our calcium imaging data that demonstrated the predominance of extrasynaptic, GluN2B-mediated NMDAR activity.

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Palmitoylation inhibitors rescue enhanced vulnerability to NMDA-mediated excitotoxicity in Ppt1^-/- cultured neurons

We next asked whether this enhanced vulnerability to excitotoxicity results from hyperpalmitoylation of neuronal substrates, and if it can be corrected by balancing the level of synaptic protein palmitoylation/depalmitoylation. First, we found that 77% of cultured Ppt1^-/- neurons accumulate ALs spontaneously at DIV18-20 (Figure 9A and B). In agreement, an immunostaining for lysosomal-associated membrane protein-2 (LAMP-2) showed colocalization of the lysosomal marker with ALs in Ppt1^-/- but not WT neurons (vehicle treatment in Figure 9B, Figure 9—figure supplement 1). Further, lysosomes appeared swollen in vehicle-treated Ppt1^-/- neurons (see arrows in Figure 9B, Figure 9—figure supplement 1D–F). Treatment with the palmitoylation inhibitors, 2-bromopalmitate (2 BP, 1 μM, 7 day treatment) and cerulenin (1 μM, 7 day treatment) reduced the percentage of AL-positive neurons (Figure 9C) and the area occupied with ALs per neuron (Figure 9D). Further, the mean lysosomal size also normalized in Ppt1^-/- neurons when these cells were treated with 2 BP or cerulenin (Figure 9E).

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To examine the efficacy of these compounds in preventing NMDA-mediated toxicity, we pretreated a subset of neurons with the same palmitoylation inhibitors, 2 BP (1 μM, DIV12-18) and cerulenin (1 μM, DIV12-18) prior to treatment with NMDA and glycine. Notably, pretreatment with both 2 BP and cerulenin improved cell viability of Ppt1^-/- neurons to that of WT following excitotoxicity induction, while the chronic low-dose treatment alone had no effect on neuronal viability (Figure 9F). These results indicate at least two features of Ppt1^-/- neurons, accumulation of proteolipid materials and a higher vulnerability to excitotoxicity, can be mitigated by correcting a balance between palmitoylation and depalmitoylation.

Palmitoylation inhibitor treatment improves pathological calcium dynamics in Ppt1^-/- neurons

To determine whether palmitoylation inhibitor treatment had a functional effect on the calcium dynamics in Ppt1^-/- neurons, we treated a subset of Ppt1^-/- cells from DIV12-18 with 2 BP (1 μM) or cerulenin (1 μM) before imaging under the same conditions described for Figure 7 (see Videos 5 and 6). Notably, treatment with both 2 BP and cerulenin decreased the AUCs of specified synapses compared to untreated Ppt1^-/- cells (see Video 7), nearly to WT levels (Figure 10A), indicating more compartmentalized calcium influx. In fact, the morphology of treated Ppt1^-/- neurites appeared more mature (see Videos 5–7). However, the AUC of cerulenin-treated Ppt1^-/- neurons was still significantly increased compared to WT, indicating a partial rescue of phenotype (Figure 10A). We also observed similar changes for the calcium diffusion distance at synaptic sites, as groups followed the order: Ppt1^-/- > Ppt1^-/- + cerulenin ≥ Ppt1^-/- + 2 BP=WT. These results further confirm the efficacy of palmitoylation inhibitor treatment (Figure 10B). Further, calcium transient frequency was higher in Ppt1^-/- cells than in WT but was lowest in Ppt1^-/- neurons treated with 2 BP or cerulenin (Figure 10C and D). It is plausible that chronic palmitoylation inhibitor treatment caused dissipation of synaptic proteins, including NMDARs (El-Husseini et al., 2002; Li et al., 2003), thereby reducing transient frequency.

Figure 10

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Video 5

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Video 6

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Video 7

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Palmitoylation inhibitors rescue Fyn kinase and GluN2B hyperpalmitoylation in Ppt1^-/- neurons

Finally, we directly examined the palmitoylation state of neuronal proteins to gain insight into the mechanisms by which hyperpalmitoylation of neuronal substrates may lead to NMDA-mediated excitotoxicity in Ppt1^-/- neurons. We also asked whether palmitoylation inhibitors can correct these abnormalities. We employed a modified acyl-biotin exchange procedure (Drisdel and Green, 2004), termed the APEGS assay (acyl-PEGyl exchange gel-shift) (Yokoi et al., 2016). The APEGS assay effectively tags the palmitoylation sites of neuronal substrates with a 5 kDa polyethylene glycol (PEG) polymer, causing a molecular weight-dependent gel shift in immunoblot analyses. Thus, we quantitatively analyzed the palmitoylated fraction of synaptic proteins and palmitoylated signaling molecules that may influence NMDAR function.

To test the feasibility of the APEGS assay in our primary cortical neuronal cultures, we collected lysates at DIV18 from WT, Ppt1^-/-, and palmitoylation inhibitor-treated (2 BP or cerulenin, DIV12-18, 1 μm) neurons and examined two palmitoylated proteins, PSD-95 and Fyn, which have been successfully quantified using this method (Yokoi et al., 2016).

We examined the palmitoylation state of PSD-95 at baseline and in response to palmitoylation inhibitor treatment (Figure 11A). As we found in our initial immunoblotting analyses of the homogenates derived from cortical tissues (Figure 2) and neuronal cultures (Figure 5), Ppt1^-/- neurons had lower amounts of total PSD-95 protein than WT, as evidenced by decreased overall band density (Figure 11B). Remarkably, the palmitoylation states of PSD-95 were comparable between Ppt1^-/- and WT neurons (Figure 11C), suggesting that PSD-95 may not be a PPT1 substrate. This finding is consistent with previous results showing that PSD-95 is not depalmitoylated by PPT1 (Yokoi et al., 2016). Further, in line with previous data (El-Husseini et al., 2002; Fukata et al., 2013), 2 BP treatment decreased the relative palmitoylation level (ratio of palm/non-palm) of PSD-95 by nearly 50% in WT neurons (Figure 11C). However, we did not observe this effect in Ppt1^-/- neurons. Also, cerulenin treatment had no consistent effects on PSD-95 levels or palmitoylation state in WT or Ppt1^-/- cells (Figure 11B and C). However, the specificity of palmitoylation inhibitors is incompletely understood and compensatory mechanisms may restore PSD-95 palmitoylation due to chronic low-dose inhibitor treatment.

Figure 11

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We performed the same analysis on another well-studied palmitoylated protein, Fyn kinase (Figure 11D). Fyn is a prominent member of the Src family kinases that phosphorylates and thereby stabilizes GluN2B at the synaptic surface (Prybylowski et al., 2005; Trepanier et al., 2012). Further, Fyn palmitoylation is important for its localization to the plasma membrane, where it may interact with GluN2B (Sato et al., 2009). Hence, Fyn hyperpalmitoylation can be a mechanism by which GluN2B retention may be enhanced in Ppt1^-/- neurons. Indeed, total levels of Fyn were increased in Ppt1^-/- neurons compared to WT, and were significantly suppressed by 2 BP and cerulenin treatment in both WT and Ppt1^-/- neurons (Figure 11E). 2 BP and cerulenin treatments also significantly reduced the ratio of palmitoylated/non-palmitoylated Fyn in WT and Ppt1^-/- neurons (Figure 11F). These findings imply that palmitoylation of Fyn regulates its stability and that Fyn hyperpalmitoylation may play a vital role in the stagnation of GluN2B to GluN2A subunit switch (Figure 11E and F).

Next, we examined the palmitoylation state of GluN2B (Figure 11G). First, total GluN2B levels were comparable between WT and Ppt1^-/-, vehicle-treated neurons at DIV18 (Figure 11G and H). While 2 BP had no effect on total GluN2B levels in WT neurons, 2 BP treatment decreased total GluN2B in Ppt1^-/- neurons compared to vehicle-treated cells (Figure 11H). Cerulenin had the same effect (Figure 11H). Importantly, both 2 BP and cerulenin corrected the ratio of palmitoylated/non-palmitoylated GluN2B in Ppt1^-/- neurons, with values approaching those of vehicle-treated WT cells (Figure 11I). No effect was observed in WT neurons treated with cerulenin. The latter results indicate that GluN2B palmitoylation state is less sensitive to chronic, low-dose palmitoylation inhibitor treatment than Fyn palmitoylation state. One possibility, therefore, is that enhanced surface retention of GluN2B-containing NMDARs in Ppt1^-/- neurons results from Fyn hyperpalmitoylation. Alternatively, hyperpalmitoylation of GluN2B may directly lead to enhanced surface retention of GluN2B-containing NMDARs in Ppt1^-/- neurons (Mattison et al., 2012). Nevertheless, these data are in line with Figure 10C, which shows the frequency of calcium influxes in Ppt1^-/- neurons is robustly decreased by treatment with 2 BP or cerulenin.

Finally, we examined the palmitoylation state of GluN2A at baseline and in response to palmitoylation inhibitor treatment (Figure 11J). As we found in our biochemical analyses of the cortical homogenates (Figures 2 and 5), Ppt1^-/- neurons showed decreases in the band intensity of total GluN2A level as compared to WT (Figure 11J and K). The palmitoylation state (ratio of palm/non-palm) of GluN2A, however, was unchanged between WT and Ppt1^-/- at baseline (Figure 11L), suggesting that PPT1 is not directly involved in the palmitoylation state of GluN2A. Interestingly, chronic treatment with 2 BP and cerulenin had dissimilar effects on GluN2A levels and palmitoylation state in WT and Ppt1^-/- neurons. In WT cells, 2 BP had no effect on GluN2A levels or palmitoylation state (Figure 11K and L), indicating that the chronic low-dose treatment does not intervene the GluN2A depalmitoylation in WT neurons. In contrast, 2 BP treatment in Ppt1^-/- neurons robustly increased both the total level and palmitoylation state of GluN2A, resulting in the nearly equal representation of two distinct GluN2A palmitoylated species (Figure 11K and L). Differing from 2 BP, cerulenin treatment modestly decreased total GluN2A levels and GluN2A palmitoylation state in WT cells (Figure 11K and L). In contrast, cerulenin treatment of Ppt1^-/- cells increased the total GluN2A protein level (Figure 11K and L), albeit not nearly as robustly as 2 BP. Overall, these results illustrate complex and potentially indirect effects of palmitoylation inhibitor treatment on the palmitoylation state of GluN2A in Ppt1^-/- neurons. While there may be other possibilities, the data suggest that 2 BP treatment may have corrected a defect in palmitoylation in Ppt1^-/- neurons (e.g. Fyn hyperpalmitoylation), thereby initiating or facilitating the GluN2B to GluN2A switch. Nevertheless, the increase in GluN2A levels and palmitoylation in Ppt1^-/- cells may ultimately account for the beneficial effects of inhibitor treatment in our complementary analyses (Figures 9 and 10).

In aggregate, these data indicate hyperpalmitoylation of Fyn and GluN2B, and point to mechanisms by which chronic low-dose palmitoylation inhibitor treatment may decrease the synaptic stabilization of GluN2B, thereby reducing calcium load in Ppt1^-/- neurons and mitigating the enhanced susceptibility to excitotoxicity. Further, these data imply that the progression of CLN1 may be mediated by the palmitoylation of Fyn kinase, which is also being targeted for the treatment of Alzheimer’s disease (Kaufman et al., 2015; Nygaard et al., 2015).

Since the development of the first Ppt1^-/- mouse and knock-in mouse model of CLN1, much progress has been made in understanding the temporal, regional, and cell-type specific effects of lipofuscin accumulation and neuronal degeneration, particularly in late stage disease (Gupta et al., 2001; Bible et al., 2004; Kielar et al., 2007; Bouchelion et al., 2014). In addition, comprehensive data characterizing the behavioral dysfunction of the Ppt1^-/- mouse has recapitulated clinical symptoms of the disease (Dearborn et al., 2015). Recent data demonstrate that PPT1 localizes to synaptic compartments and influences presynaptic localization and mobility of prominent presynaptic proteins, including SNAP25 (Kim et al., 2008). These findings correlate with histological and electrophysiological findings in cultured Ppt1^-/- neurons, demonstrating a depletion of presynaptic vesicle pool size (Virmani et al., 2005). Moreover, presynaptic protein localization and function are altered in CLN1 models and in human tissue (Kanaani et al., 2004; Kim et al., 2008; Aby et al., 2013).

In the current study, we have identified a role for PPT1 in postsynaptic maturation in the Ppt1^-/- mouse model of CLN1. Our principal finding demonstrates a role for PPT1 in the regulation of NMDAR composition and function that leaves Ppt1^-/- neurons vulnerable to excitotoxic insult, which is alleviated by chronic low-dose palmitoylation inhibitor treatment. Together, these data implicate dysregulated GluN2 subunit switch as a major pathogenic mechanism in CLN1.

All data generated or analysed during this study are included in the manuscript.

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Author details

1. Kevin P Koster

   Department of Anatomy and Cell Biology, University of Illinois at Chicago, Chicago, United States

   Contribution

   Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Writing—original draft, Writing—review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-2935-3427

2. Walter Francesconi

   Department of Anatomy and Cell Biology, University of Illinois at Chicago, Chicago, United States

   Contribution

   Formal analysis, Investigation, Methodology, Writing—review and editing

   Competing interests

   No competing interests declared

3. Fulvia Berton

   Department of Anatomy and Cell Biology, University of Illinois at Chicago, Chicago, United States

   Contribution

   Formal analysis, Investigation, Methodology, Writing—review and editing

   Competing interests

   No competing interests declared

4. Sami Alahmadi

   Department of Anatomy and Cell Biology, University of Illinois at Chicago, Chicago, United States

   Contribution

   Data curation, Formal analysis, Validation

   Competing interests

   No competing interests declared

5. Roshan Srinivas

   Department of Anatomy and Cell Biology, University of Illinois at Chicago, Chicago, United States

   Contribution

   Data curation, Software, Formal analysis

   Competing interests

   No competing interests declared

6. Akira Yoshii

   1. Department of Pediatrics, University of Illinois at Chicago, Chicago, United States
   2. Department of Neurology, University of Illinois at Chicago, Chicago, United States

   Contribution

   Conceptualization, Resources, Supervision, Funding acquisition, Investigation, Writing—original draft, Writing—review and editing

   For correspondence

   ayoshii@uic.edu

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0001-8305-006X

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