Acid sphingomyelinase modulates the autophagic process by controlling lysosomal biogenesis in Alzheimer’s disease

We first sought to confirm whether sphingolipid metabolism is altered in AD patient samples. We examined ASM and acid ceramidase (AC) activities, and the levels of several sphingolipids, including sphingomyelin and ceramide, in samples from normal individuals and AD patients. Consistent with previous results (He et al., 2010), ASM was significantly increased in plasma and fibroblasts from individuals with AD compared with normal aged individuals (Fig. 1, A and B). To assess whether increased ASM activity was an AD-specific signature, we analyzed ASM activity in samples from individuals with Parkinson’s disease (PD). The activity of ASM was not elevated in PD-derived samples compared with normal (Fig. 1, A and B). ASM activity also did not show passage differences between AD and normal fibroblasts (Fig. 1 C). Sphingomyelin levels were decreased in the AD plasma compared with normal (Fig. 1 D). No significant differences in the ceramide and AC levels were found between the two groups (Fig. 1 D). These results confirmed that elevation of ASM, an important sphingolipid-modulating factor, is AD specific and may influence disease progression and/or pathogenesis.

To investigate the influence of ASM on AD pathology, we first generated amyloid precursor protein (APP)/presenilin 1 (PS1) double mutant and APP/PS1/ASM^−/− triple mutant mice (Fig. 1 E). AD-related pathologies in APP/PS1 mice normally begin at ∼6–7 mo of age; however, our APP/PS1/ASM^−/− mice died young (Fig. 1 F). We presume that the early death of the APP/PS1/ASM^−/− animals was due to their ASM^−/− phenotype because these animals (originally developed as a model of the neurodegenerative type A NPD) usually die by ∼6–8 mo of age. The APP/PS1/ASM^−/− mice showed significantly decreased body weight compared with APP/PS1 mice (Fig. 1 G), and indicators of brain injury, such as cell death and inflammation, were significantly increased (Fig. 1, H–J). These data demonstrated that complete deletion of ASM in APP/PS1 mice exacerbated brain pathology, and that APP/PS1/ASM^−/− mice were not suitable to examine the correlation of ASM and AD pathology.

To overcome these obstacles, we generated APP/PS1/ASM^+/− triple mutant mice (with partial genetic deletion of the ASM gene; Fig. 2, A and B). Similar to AD patients, ASM activity was elevated in plasma, brain, and fibroblasts of 9-mo-old APP/PS1 mice (Fig. 2 C), likely due to the stress response related to the progression of AD-like disease in these animals. Next, to further investigate the cell contribution of increased ASM activity in AD mouse brain, we isolated neurons and microglia from the brain. Although ASM activity was slightly increased in APP/PS1 microglia compared with WT microglia, the degree of ASM increase was greater in neurons than microglia (Fig. 2 D), indicating that neurons were the main contributor of elevated ASM activity in AD mouse brain.

Importantly, ASM activity in age-matched APP/PS1/ASM^+/− mice was significantly decreased compared with the APP/PS1 mice to levels within the normal range or lower (Fig. 2 C). Other sphingolipid factors were unaltered in the APP/PS1/ASM^+/− mice except sphingomyelin, which was modestly reduced in APP/PS1 mice and elevated in the triple mutant animals (Fig. 2 E).

To determine whether the reduced ASM activity in the APP/PS1/ASM^+/− mice affected AD pathology, we first determined the Aβ profile. Thioflavin S staining, immunofluorescence, and ELISA results of Aβ40 and Aβ42 showed significantly lower Aβ levels in the 9-mo-old APP/PS1/ASM^+/− mice compared with age-matched APP/PS1 mice (Fig. 2, F–I). In APP/PS1/ASM^+/− mice, cerebral amyloid angiopathy and C-terminal fragment of APP were also reduced (Fig. 2 J; and see Fig. 4, D and F). There were no significant differences of tau hyperphosphorylation between the two groups (Fig. 2 K).

Next, to assess the potential effect of partial genetic ASM inhibition on learning and memory in APP/PS1 mice, we performed the Morris water maze task and fear conditioning. Aged APP/PS1 mice showed severe deficits in memory formation and APP/PS1/ASM^+/− mice were largely protected from this defect (Fig. 3). Collectively, these results suggested that restored ASM activities to the normal range in APP/PS1 mice decreased Aβ load and improved learning and memory.

A reduction in APP/PS1/ASM^+/− mouse cerebral amyloidosis could be due to a decreased inflammatory response, attenuated APP expression, or activation of proteases involved in Aβ degradation. We first assessed the apoptotic and inflammatory responses in brain samples derived from APP/PS1 and APP/PS1/ASM^+/− mice but did not detect differences between the two strains (Fig. 4, A–C). To determine whether reduction of ASM activity affected APP expression, we compared the levels of APP in the two strains. We found that partial genetic inhibition of ASM did not influence the overall expression levels of APP (Fig. 4, D and E). We also examined the Aβ generating enzyme Bace-1 using brain homogenates. Bace-1 was slightly decreased in APP/PS1/ASM^+/− mice compared with APP/PS1 mice, but the reduction did not reach statistical significance (Fig. 4 G). To address Aβ clearance by microglia, we analyzed microglia activation and Aβ degrading enzyme release by microglia but again did not detect differences (Fig. 4, H and I). Similarly, these changes did not show any differences between APP/PS1 and APP/PS1/ASM^+/− mice in 5-mo-old young mice (Fig. 4, J–M). There were also no significant differences of brain pathology, such as apoptosis, inflammation, and Aβ deposition, between the WT and ASM^+/− mice (Fig. 4, A–I). Overall, these data suggested that the partial inhibition of ASM in APP/PS1/ASM^+/− did not alter major inflammatory pathways or expression of APP.

Dysfunction of the normal proteolytic degradation system also could affect AD pathogenesis and lead to enhanced Aβ deposition (Lee et al., 2010b). Autophagy, a major degradative pathway of the lysosomal system, is known to be markedly impaired in AD (Boland et al., 2008). We found that the microtubule-associated protein 1 light chain 3 (LC3)–II levels were significantly increased in human AD-derived fibroblasts compared with control fibroblasts (Fig. 5, A–C). Increased LC3-II levels could stem from overinduction of autophagy or may be a product of reduced autophagic turnover and defects in the latter stages of autophagic degradation. We therefore measured the level of beclin-1 expression in the AD cells, which is part of a kinase complex responsible for autophagy induction (Zeng et al., 2006), and found that it did not vary between the groups (Fig. 5, A and B).

To examine autophagic turnover of protein, we then analyzed proteolysis of long-lived proteins (Lee et al., 2010b) in control and AD fibroblasts. When autophagic/lysosomal degradation was induced through serum withdrawal, proteolysis was increased in control fibroblasts but not significantly changed in PS1 and apolipoprotein E4 (ApoE4)–derived AD patient fibroblasts (Fig. 5 D). Lysosome stability relating to autophagy also could be affected by lysobisphosphatidic acid (LBPA) binding with ASM (Kirkegaard et al., 2010). We therefore measured LBPA immunofluorescence intensity in control and AD fibroblasts, and found that it did not vary between the groups (Fig. 5 E). Collectively, these data indicated that the autophagosome accumulation in AD is due to dysregulation of autophagic protein degradation, similar to previous results (Lee et al., 2010b).

To examine how genetic inhibition of ASM affected the autophagic pathway in AD, we also analyzed fibroblasts and brain samples derived from 9-mo-old WT, APP/PS1, APP/PS1/ASM^+/−, and ASM^+/− mice. Compared with WT, APP/PS1 mice showed increased LC3-II, similar to human AD fibroblasts. This enhanced LC3-II level was reduced in APP/PS1/ASM^+/− mice. Beclin-1 expression did not vary between the groups (Fig. 5, F, G, K, and L). Metabolic analysis of protein turnover (Cuervo et al., 2004) was assessed using fibroblasts from WT, APP/PS1, and APP/PS1/ASM^+/− mice. Under culture conditions that induced autophagy (absence of serum), degradation of long-lived proteins was significantly lower in cells from APP/PS1 mice compared with WT mice but was increased in cells derived from APP/PS1/ASM^+/− mice (Fig. 5 I). The differences of cell senescence levels in cultured mice fibroblasts were not found between the groups, indicating that these changes were not related to the cell senescence (Fig. 5 J). The levels of cathepsin D, a lysosomal hydrolase, were elevated in APP/PS1 mice compared with WT. Enhanced cathepsin D level was ameliorated in APP/PS1/ASM^+/− mice (Fig. 5, F, G, K and L). However, the activity of cathepsin D was not changed between the groups (Fig. 5, H and M). This result indicates that the elevated levels of cathepsin D in APP/PS1 mice did not ultimately translate into a significant increase of enzyme activity. We also analyzed the expression of p62, indicator of autophagic turnover. Increased p62 levels in APP/PS1 mice were reduced in APP/PS1/ASM^+/− mice (Fig. 5, F, G, K and L).

To corroborate the immunoblotting results, we performed electron microscope (EM) analysis using mouse brain samples. As previously reported (Yu et al., 2005), APP/PS1 mouse brain regions showed an increased number of autophagic vacuoles (AVs), whereas brains of APP/PS1/ASM^+/− mice showed a reduced number of these vesicles, albeit still higher than WT mice (Fig. 5 N).

The endocytic pathway is also considered a major contributor to Aβ deposition in AD (Ginsberg et al., 2010; Li et al., 2012). To determine whether the endocytic pathway was affected by partial ASM inhibition, we examined Rab5 and Rab7 expression in our animals. The expression pattern of these proteins showed no difference between the groups (Fig. 5 O). Although additional studies of endocytic pathway are required to identify the exact mechanism, our results showed that endocytic pathway was not a main mechanism by ASM inhibition. Collectively, these results revealed dysfunctional changes in the turnover of AVs in the APP/PS1 mice, and that partial genetic ASM inhibition could reverse this abnormality and improve autophagic degradation of proteins.

To gain more direct insights into the relationship of ASM and autophagic dysfunction, we treated human fibroblasts and neurons with recombinant 1–10 µM ASM and determined the LC3-II and p62 levels. ASM strongly accelerated LC3-II and p62 levels in human fibroblasts and neurons in a concentration-dependent manner (Fig. 6, A–C). The level of beclin-1 expression was not affected by ASM (Fig. 6, A and C), indicating that the accumulation of autophagosomes was not due to the biogenesis pathway. ASM is found in a secretory and a lysosomal form (Jenkins et al., 2009), and the mannose-6-phosphate (M6P) receptor system is involved in trafficking of ASM to the lysosome (Dhami and Schuchman, 2004). To elucidate which ASM form affected lysosomal/autophagic dysfunction, cells were incubated with ASM alone, or in the presence of M6P. As expected, the activity of ASM was significantly increased in ASM-treated cells compared with nontreated cells. This enhanced ASM activity was reduced in the presence of M6P (Fig. 6 D). To confirm whether the ASM treatment reached the lysosomes, we also examined the colocalization of ASM and lysosomes using immunocytochemistry. Double immunostaining of ASM and lysosomal-associated membrane protein 1 (Lamp1) showed that most ASM-positive vesicles were colocalized with Lamp1-positive vesicles, indicating that treated ASM was located in lysosome (Fig. 6 E). The ASM-induced autophagosome accumulation was significantly decreased by inhibition of lysosomal ASM uptake using M6P or M6P receptor siRNA (Fig. 6, F and G). These results suggested that elevation of lysosomal ASM may lead to autophagic dysfunction in AD. ASM treatment did not affect the cell survival (Fig. 6 H). The levels of LC3 and cell senescence in the fibroblast also did not show the differences with passage number, indicating that ASM caused the abnormal autophagy (Fig. 6, I and J).

As discussed above, the accumulation of AVs in cells can result from either autophagy induction or the blockade of autophagic degradation. To further distinguish between these possibilities, we performed an autophagic flux assay (Rubinsztein et al., 2009) in the presence or absence of NH₄Cl that blocks autophagic degradation but does not affect autophagosome formation. It was hypothesized that if ASM treatment enhanced autophagy induction, in the presence of NH₄Cl (which inhibits degradation) a considerable increase in LC3-II would be expected due to the combined effects of blocking degradation and enhancing induction. However, compared with NH₄Cl treatment alone, dual treatment of fibroblasts with ASM and NH₄Cl did not show any significant changes of LC3-II (Fig. 7 A, left). In contrast, the addition of NH₄Cl (Fig. 7 A, middle) or ASM (Fig. 7 A, right) in serum starvation culture resulted in a significant but similar increase of LC3-II levels. Furthermore, we measured the autophagic flux by detecting the abundance of p62. The levels of p62 were markedly increased in the cells treated with ASM or NH₄Cl (Fig. 7 B). We also performed LC3 flux assay in human AD fibroblasts and 9-mo-old WT, APP/PS1, and APP/PS1/ASM^+/− mice fibroblasts. Autophagic flux was measured by assessing the changes of LC3-II in the presence and absence of NH₄Cl–mediated lysosomal inhibition. Under basal condition, human AD and APP/PS1 fibroblasts showed significantly increased LC3-II levels compared with normal cells. NH₄Cl–induced lysosome inhibition led to marked increase of LC3-II levels in the normal fibroblasts, but this increase was significantly less in the AD cells (Fig. 7, C and D). APP/PS1/ASM^+/− fibroblast showed similar pattern in LC3-II increase compared with normal cell (Fig. 7 D). Collectively, these results indicated that enhanced lysosomal ASM in AD caused a defect of autophagic degradation but not induction.

To further investigate the relationship of elevated ASM and defective lysosomal/autophagic degradation, we evaluated alteration in lysosomal pH using the acidotropic dye LysoTracker red. H₂O₂- and NH₄Cl-treated cells were used as positive and negative controls, respectively. Flow cytometry and fluorescent microscopic analysis did not show any differences of lysosomal pH between ASM- and vehicle-treated fibroblast (Fig. 7 E). Recently, the transcription factor EB (TFEB) was identified as a master regulator of the autophagy–lysosome pathway (ALP) and lysosome biogenesis (Settembre et al., 2011). Enhancement of TFEB function is able to stimulate ALP function and promote protein clearance. To examine whether ASM could affect the ALP and lysosome biogenesis, we tested endogenous levels of TFEB. ASM-treated fibroblasts and neurons showed significantly decreased TFEB levels (Fig. 7, F and G). Also, the levels of the lysosomal structural protein Lamp1 were decreased in ASM-treated cells (Fig. 7, F–H). To further validate our observation, we investigated TFEB subcellular localization after ASM treatment. Interestingly, ASM-treated cells showed a reduced TFEB expression in the nuclear compartment (Fig. 7 I). Similarly, the expression levels of TFEB target genes related to lysosome were significantly decreased in ASM-treated fibroblasts (Fig. 7 J). Conversely, to determine whether autophagic degradation affected ASM, we evaluated ASM activity in fibroblasts after NH₄Cl treatment. Blocking of autophagic degradation via NH₄Cl did not show any significant changes of ASM activity (Fig. 7 K). These results further suggested that lysosomal ASM acts not as an inducer but rather as an inhibitor of autophagic protein degradation by reducing ALP function and lysosome biogenesis.

To examine the in vivo effect of ASM activation on autophagic dysfunction, we introduced conditioned medium (CM) from cultured ASM-overexpressing cells into C57BL/6 mice via intracerebral (i.c.) and intravenous (i.v.) injections. ASM-CM–treated mice showed elevated ASM activity in the brain and plasma (Fig. 8, A and B), as well as increased LC3-II without changes of beclin-1 expression in the brains (Fig. 8, C and D). Cathepsin D level was increased in the ASM-CM (i.c.)–treated mice compared with control mice, but actual activity was not changed (Fig. 8, C and E). p62 also was increased in the ASM-CM (i.c.)–treated mice compared with control mice (Fig. 8 C). ASM-CM (i.c.)–treated mice further exhibited abnormal ALP function, indicative of decreased TFEB and Lamp1 levels (Fig. 8 F). Although ASM-CM (i.v.)–treated mice showed slightly increased cathepsin D and p62 levels and decreased ALP function proteins, this did not reach statistical significance (Fig. 8, D and G). The activity of cathepsin D was also not changed (Fig. 8 E). These relatively modest effects of ASM-CM (i.v.) treatment on autophagy dysfunction might be due to presence of the blood–brain barrier because only a slight increase of ASM activity in the brain was achieved by these treatments, and the activity of ASM did not reach those of APP/PS1 mice.

To examine whether partial genetic inhibition of ASM affected the ALP in APP/PS1 mice, we also analyzed TFEB and Lamp1 levels in the brain samples derived from WT, APP/PS1, ASM^+/−, and APP/PS1/ASM^+/− mice. Compared with WT, APP/PS1 mice showed significantly decreased TFEB and Lamp1 expression, which were increased in APP/PS1/ASM^+/− mice (Fig. 8 H). Together, these findings show for the first time a direct correlation of lysosomal ASM and the function of the ALP, and suggest that abnormal autophagic degradation in AD may be due to the effects of elevated ASM expression on this pathway.

The ASM-mediated lysosomal/autophagic dysfunction in AD prompted us to examine possible therapeutic implications of this pathway. To decrease ASM in APP/PS1 mice, we undertook pharmacological inhibition using amitriptyline-hydrochloride (AMI) for 4 mo (Fig. 9 A). AMI is a known inhibitor of ASM that can cross the blood–brain barrier. At 9 mo of age, AMI-treated APP/PS1 mice exhibited decreased ASM activity compared with vehicle-treated mice (Fig. 9 B). Other sphingolipid metabolites were not changed (Fig. 9 C). Aβ levels were decreased in the AMI-treated APP/PS1 mice compared with the nontreated littermates (Fig. 9, D–G). The levels of LC3-II, p62, and cathepsin D were decreased in the AMI-treated APP/PS1 mice (Fig. 9, H and I). Actual activity of cathepsin D was not changed by AMI treatment (Fig. 9 J). AMI treatment significantly increased TFEB and Lamp1 protein levels in APP/PS1 mice (Fig. 9, H and I). Similarly, APP/PS1 mice treated with AMI showed recovery of memory function (Fig. 9, K–P). Overall, these positive but relatively moderate results (e.g., Aβ levels) in AMI-treated APP/PS1 mice might be due to under dosing of the animals. We speculate that this may be improved in the future by adjusting the dose or using modified, more potent drugs of a similar class.

To further validate our observation made by partial ASM inhibition in AD mice, we studied possible changes in autophagy dysfunction in human AD fibroblast after ASM inhibition. Elevated ASM levels in human AD fibroblasts (PS1–familial AD [FAD] and ApoE4) were restored to normal range by ASM siRNA treatment (Fig. 10 A). ASM siRNA-treated human AD fibroblasts (PS1-FAD and ApoE4) showed decreased LC3-II and p62 accumulation compared with control siRNA-treated cells (Fig. 10 B). Also, ASM siRNA was able to increase lysosome levels (as judged by Lamp1 expression) by activating TFEB in the human AD fibroblasts (Fig. 10 C).

Many insights into the pathogenesis in neurodegenerative disease have come from investigating postmortem brain tissues due to the difficulty of invasive access to living human CNS. The recent developments in induced pluripotent stem cells (iPSCs) and induced neurons have allowed investigation of pathogenesis of neurological diseases in vitro (Kondo et al., 2013). To explore whether the observed effects of ASM in previous results are paralleled by similar alterations in AD human neurons, we first established iPSCs with PS1 mutation (PS1 iPSC-2, -4, and -21) by transduction of human fibroblast with retroviruses encoding OCT4, SOX2, KLF4, and c-Myc. The PS1-iPSC cell line was shown to be fully reprogrammed to pluripotency, as judged by colony morphology, alkaline phosphatase (AP) staining, expression of pluripotency-associated transcription factors and surface markers, karyotype stability, and generation of teratomas (Fig. 10, D–G). To establish whether the PS1 mutation may affect neuronal differentiation, PS1 iPSC and control iPSC lines were induced to differentiate into neurons for 10 d. Consistent with previous results (Kondo et al., 2013), no obvious differences in the ability to generate neurons were observed between control and PS1-iPSCs (Fig. 10 H). Aβ42 secretion level was increased in PS1 iPSC-derived neurons compared with control iPSC-derived neuron (Fig. 10 I).

Next, we investigated whether elevated ASM in fibroblasts was also evident increased in PS1 iPSC and iPSC-derived neurons. The ASM activity in PS1 iPSC was not changed except for PS1-4 iPSC in comparison to those in control iPSC, but the activity of ASM was significantly higher in PS1 iPSC-derived neurons compared with control iPSC-derived neuron (Fig. 10 J). Elevated ASM levels in PS1 iPSC-derived neurons were restored to normal range by ASM siRNA treatment (Fig. 10 J). Neurons from PS1-4 iPSCs also had significantly higher abnormal autophagic markers than neurons from control iPSC (Fig. 10 K). ASM siRNA treatment significantly decreased the protein level of abnormal autophagic markers in PS1 iPSC-derived neurons (Fig. 10 K). To corroborate the immunoblotting results, we performed EM analysis using control and PS1 iPSC-derived neurons. As expected, PS1 iPSC-derived neurons exhibited increased AV accumulation, whereas ASM siRNA-treated PS1 iPSC-derived neurons showed a reduced number of these vesicles (Fig. 10 L). Decreased TFEB target genes in PS1 iPSC-derived neurons were also significantly increased by ASM siRNA treatment (Fig. 10 M). These results confirm that abnormal autophagy observed in AD mice and human fibroblasts by ASM also occur in AD patient neurons, and restoration of ASM back to normal levels is able to ameliorate autophagic dysfunction by restoring lysosomal biogenesis in AD patient cells.

Although the exact causes of AD are unknown, the complex interactions of genetic and environmental factors are likely play important roles in the pathogenesis. ASM activity is known to be increased by environmental stress and in various diseases, and is elevated in AD patients (He and Schuchman, 2012). One downstream consequence of increased ASM is elevated ceramide, contributing to cell death, inflammation, and other common disease findings. Although elevated ASM is known to occur in AD, the cellular mechanisms that link ASM and AD have not been fully characterized. The data presented here suggest a previously unknown role of ASM in the down-regulation of lysosomal biogenesis and inhibition of lysosome-dependent autophagic proteolysis. The findings also establish proof of concept for ASM inhibitor therapy in AD.

Our previous study showed that sphingolipid metabolism was severely impaired in the human AD brain, and that ASM activity was positively correlated with the Aβ levels (He et al., 2010). Consistent with our previous study, we found that ASM was significantly increased in fibroblasts, brain, and/or plasma from patients with AD and in AD mice, although other sphingolipid factors were unaltered. There are some differences between the previous and this study. For example, the previous results showed increased ceramide level in AD, but we could not found significant changes of sphingolipid factors including ceramide in AD compared with normal samples. These differences might be related to the fact that once formed, ceramide can rapidly enter several metabolic pathways. It may be used for either the biosynthesis of complex lipids or broken down into sphingosine, which itself is rapidly converted to sphingosine 1 phosphate.

Accumulation of abnormal AVs has been observed in AD (Lee et al., 2010b; Nixon and Yang, 2011), and the autophagy pathway is increasingly regarded as an important contributor to Aβ-mediated pathogenesis in AD (Yu et al., 2005; Nixon, 2007). Moreover, it has been shown that the abnormal autophagic flux in AD may be due to dysfunction at the late autophagy stage associated with the lysosome (Lai and McLaurin, 2012; Zhou et al., 2012). Similar to previous results (Lee et al., 2010b; Lai and McLaurin, 2012), we found that the impaired autophagic flux in AD was associated with reduced autophagic degradation due to decreased ALP function. In addition, we show for the first time that this is directly linked to elevated ASM activity. Suppression of ASM expression or inhibition of its uptake and delivery to lysosomes using M6P reversed abnormal autophagic degradation. These findings indicate that increased lysosomal ASM plays a negative role in AD by causing autophagic dysfunction, suggesting that therapeutic strategies to restoring ASM activity to the normal range may be beneficial for AD pathology.

This was further studied in the AD mice by finding that partial genetic or systemic inhibition of ASM activities in these animals largely reversed autophagic pathology by restoring ALP function, as well as reducing the accumulation of incompletely digested substrates within the autophagic-lysosomal compartments (e.g., LC3-II and p62). Aβ accumulation also was reduced in response to ASM inhibition, as was cathepsin D expression. There are several challenges associated with interpretation of cathepsin D levels in AD. Although some reports have shown that cathepsin D activities were decreased in AD (Lee et al., 2010b), many studies indicate that cathepsin D is elevated in AD and contributes to the pathogenesis, such as Aβ formation (Cataldo et al., 2004; Lai and McLaurin, 2012; Zhou et al., 2012). A recent paper suggested that COP9 signalosome deficiency increased cathepsin D levels but reduced the autophagic degradation. They suggested that these results were associated with a failure of lysosomal assembly of cathepsin D because only a lysosomal cathepsin D could affect autophagic degradation (Su et al., 2011). In this study, we have found that maturation of cathepsin D was increased in AD mice, but the actual enzyme activity was not changed between the groups. This result indicated that the elevated levels of cathepsin D did not ultimately translate into a significant increase of enzyme activity. Based on these papers and our data, a plausible interpretation of increased cathepsin D in our AD mice is that AD microenvironment attempts to increase cathepsin D synthesis, but this does not have a direct impact on lysosomal function because the activity of the enzyme is unchanged. Therefore, enhanced cathepsin D level in our AD mice induced by increased ASM is more likely a compensatory response to an impaired lysosome system. The present study also provides the first evidence of increased ASM activity and autophagic dysfunction in living human (iPSC-derived) neurons derived from AD patients and that restoring normal levels of ASM in AD neurons effectively blocks abnormal autophagy.

Overall, the data presented here show that increased ASM activity in AD contributes to the abnormal lysosomal/autophagic process by leading to dysfunction of ALP. This results in an inability to break down appropriate substrates during the autophagy process. Restoration of ASM effectively blocks AD progression by increasing autophagic degradation. Although the involvement of other ASM-related mechanisms in AD remains to be explored, the data in this study demonstrate that inhibition of ASM improves Aβ clearance and rescues impaired memory in a validated mouse model of AD, suggesting this as a potential therapy for AD patients in the future.

Transgenic mouse lines overexpressing the hAPP695swe (APP) and presenilin-1M146V (PS1) mutations, respectively, were generated at GlaxoSmithKline by standard techniques as previously described (Howlett et al., 2004). In brief, a Thy-1–APP transgene was generated by inserting the 695 aa isoform of human cDNA (APP695) harboring the Swedish double familial mutation (K670N; M671L) into a vector containing the murine Thy-1 gene. The Thy-1–PS-1 transgene was generated by inserting the coding sequence of human PS-1 cDNA harboring the M146V familial mutation into a vector containing the murine Thy-1 gene. Transgenic lines were generated by pronuclear microinjection into fertilized oocytes from either C57BL/6xC3H mice in the case of Thy-1-APP transgene, or into fertilized oocytes from pure C57BL/6 mice in the case of Thy-1–PS-1 transgene. Thy-1 APPswe mice were generated and backcrossed onto a pure C57BL/6 background before crossing with TPM (PS-1 M146V) mice to produce heterozygote double mutant mice. ASM^+/− mice (C57BL/6 background; Horinouchi et al., 1995) were bred with APP/PS1 mice to generate APP/PS1/ASM^+/− mice. Because APP/PS1 mice show sex difference in disease progression, we used only male mice. Data analysis of APP/PS1 mice was done at 5 or 9 mo old. Block randomization method was used to allocate the animals to experimental groups. To eliminate the bias, we were blinded in experimental progress such as data collection and data analysis. SCID Beige mice (Charles River) were used for teratoma formation assay. Mice were housed at a 12 h day/12 h night cycle with free access to tap water and food pellets. Mouse studies were approved by the Kyungpook National University Institutional Animal Care and Use Committee (IACUC).

Human plasma samples were obtained from individuals with AD, PD, and age-matched, non-AD controls from Yonsei University Severance Hospital (Table S1). Informed consent was obtained from all subjects according to the ethics committee guidelines at the Yonsei University Severance Hospital.

Human fibroblast lines (normal, PS1, ApoE4, and PD) acquired from the Coriell Institute were maintained in DMEM with 15% FBS. The human cortical neuronal cell line HCN-2 was acquired from ATCC. Cells with a passage number 10–15 were used in this study. To obtain CM containing ASM, 5 × 10⁵ Chinese hamster ovary cells overexpressing human ASM (He et al., 1999) were cultured in DMEM for 2 d. Cells were washed with PBS and changed with new media. 24 h later, the CM was collected, centrifuged, and filtered using a 0.22 µm filter. We isolated WT, APP/PS1, APP/PS1/ASM^+/−, and ASM^+/− mouse tail fibroblasts as previously described (Takahashi et al., 2007a) from 9-mo-old mice. For some experiments, cells were treated with purified, recombinant ASM or ASM siRNA to measure autophagy regulation. NH₄Cl was used to inhibit the autophagic flux. For the inhibition of lysosomal ASM uptake, 10 mM M6P or M6P receptor siRNA were added to the fibroblast culture media at the same time as ASM.

4-mo-old APP/PS1 mice received 100 µg/g body weight AMI (Sigma-Aldrich) per os in their drinking water for 4 mo, and a control group received water without drug. 3-mo-old C57BL/6 mice were treated with ASM-CM via i.v. (100 µl) or i.c. (3 µl) injections on 10 consecutive days.

Thioflavin S staining was done according to previously described procedures (Lee et al., 2012). We used anti-20G10 (mouse, 1:1,000, provided by D.R. Howlett, GlaxoSmithKline, Harlow, Essex, UK) for Aβ 42, anti-G30 (rabbit, 1:1,000, provided by D.R. Howlett) for Aβ40, rabbit anti–Iba-1 (1:500; Wako), rabbit anti-GFAP (1:500, Dako), mouse anti–α-SMA (1:400; Sigma-Aldrich), rabbit anti-AT8 (1:500; Thermo Fisher Scientific), and rabbit anti–active caspase3 (1:50; EMD Millipore). The sections were analyzed with a laser-scanning confocal microscope (FV1000; Olympus) or with a BX51 microscope (Olympus). MetaMorph software (Molecular Devices) was used to quantification.

For measurement of Aβ40 and Aβ42, we used commercially available ELISA kits (BioSource). Hemispheres of mice were homogenized in buffer containing 0.02M guanidine. ELISA was then performed for Aβ40 and Aβ42 according to the manufacturer’s instructions.

We performed behavioral studies to assess spatial learning and memory in the Morris water maze as previously described (Lee et al., 2012). Animals were given four trials per day for 10 d to learn the task. At 11 d, animals were given a probe trial in which the platform was removed. Fear conditioning was conducted as previously described techniques (Kojima et al., 2005). On the conditioning day, mice were individually placed into the conditioning chamber. After a 60-s exploratory period, a tone (10 kHz, 70 dB) was delivered for 10 s; this served as the conditioned stimulus (CS). The CS co-terminated with the unconditioned stimulus (US), a scrambled electrical footshock (0.3 mA, 1 s). The CS-US pairing was delivered twice at a 20-s intertrial interval. On day 2, each mouse was placed in the fear-conditioning chamber containing the same exact context, but with no administration of a CS or foot shock. Freezing was analyzed for 5 min. On day 3, a mouse was placed in a test chamber that was different from the conditioning chamber. After a 60-s exploratory period, the tone was presented for 60 s without the footshock. The rate of freezing response of mice was used to measure the fear memory.

RNA was extracted from the brain homogenates and cell lysates using the RNeasy Lipid Tissue Mini kit and RNeasy Plus Mini kit (QIAGEN) according to the manufacturer’s instructions. cDNA was synthesized from 5 µg of total RNA using a commercially available kit (Takara Bio Inc.). Quantitative real-time PCR was performed using a Corbett research RG-6000 real-time PCR instrument. Used primers are described in Table S2.

Brain tissues and cells were fixed in 3% glutaraldehyde/0.1 M phosphate buffer, pH 7.4, and postfixed in 1% osmium tetroxide in Sorensen’s phosphate buffer. After dehydration in ethyl alcohol, the tissue and cells were embedded in epon (Electron Microscopy Sciences). Samples were cut serially and placed on copper grids and analyzed using a transmission EM (Tecnai). Images were captured on a digital camera and Xplore3D tomography software.

Total protein degradation in cultured cells was measured by pulse-chase experiments with 48 h pulse with 2 µCi/ml [³H]-leucine for 48 h to preferentially label long-lived proteins (Lee et al., 2010b).

Samples were immunoblotted as previously described (Settembre et al., 2011; Lee et al., 2012). Primary antibodies to the following proteins were used: BACE-1 (mouse, 1:1,000; Millipore), LC3 (rabbit, 1:1,000; Cell Signaling Technology), Beclin-1 (rabbit, 1:1,000; Cell Signaling Technology), p62 (rabbit, 1:1,000; Cell Signaling Technology), rab5 (rabbit, 1:1,000; Cell Signaling Technology), rab7 (rabbit, 1:1,000; Cell Signaling Technology), TFEB (rabbit, 1:1,000; Cell Signaling Technology), Lamp1 (rabbit, 1:1,000; Abcam), TFEB (rabbit, 1:500; Novus Biologicals), cathepsin D (goat, 1:500; R&D Systems), 6E10 (mouse, 1:500; Signet), Histon H3 (rabbit, 1:1,000; Cell Signaling Technology), and β-actin (1:1,000; Santa Cruz Biotechnology, Inc.). We performed densitometric quantification using the ImageJ software (National Institutes of Health).

We performed the measurements as previously described methods using a UPLC system (Waters; He et al., 2010).

We prepared samples for lipid extraction as previously described (Lee et al., 2010a). To quantify the sphingomyelin and ceramide levels, the dried lipid extract was resuspended in 0.2% Igepal CA-630 (Sigma-Aldrich) and the levels of each lipid were determined using the UPLC system.

We isolated neuron and microglia from the mouse brain as previously described (Brewer and Torricelli, 2007). In brief, 9-mo-old mice cerebrum were minced in HibernateA (Brainbits LLC)/B27 (Invitrogen) medium and dissociated using papain solution. After tissue trituration, cells were separated by density gradient centrifugation. Fractionated cells were used for ASM activity assay.

LysoTracker red (Invitrogen) was used at a final concentration of 75 nM. The cells were trypsinized, resuspended in PBS, and analyzed on a FACSCalibur using FACSDiva software (BD).

Cell viability was quantified by using MTT (3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazoliumbromide; Sigma-Aldrich) reagent. In brief, human fibroblast was seeded onto 96-well plates at a density of 1.5 × 10³ cells per well. 10 µM ASM was added to the culture media, and the cells were incubated for 24 h. MTT stock solution (5 mg/ml) was prepared in PBS (Gibco) and added to the culture media at a final concentration of 1 mg/ml. After 90 min incubation, the media was removed, and the chromogen in the cells was dissolved in DMSO containing 0.01 N NaOH. The absorbance was measured at 570 nm using a 96-well microplate spectrophotometer.

Enzyme activity of cathepsin D was determined with cathepsin D activity fluorometric assay kit according to the manufacturer’s protocol (Abcam).

PS1-iPSCs were established from the PS1 patient’s skin fibroblasts (Coriell Institute) as previously described (Okita et al., 2007), with slight modifications. In brief, PS1 fibroblasts were seeded at 3 × 10⁵ cells in 60 mm² dishes coated with gelatin. On day 1, the VSV-G pseudotyped retroviral vector system carrying OCT4, SOX2, KLF4, and c-Myc was added to fibroblast cultures. On day 2, cells were subjected to the same transduction procedures and harvested 24 h later. Transduced cells were replated on mouse embryonic fibroblast (MEF) layers in 100 mm² dishes containing the fibroblast medium. On the next day, the medium was changed to complete ES medium with 0.5 mM valproic acid (Sigma-Aldrich), and thereafter changed every other day. After 20 d, ES-like colonies appeared and were picked up to be reseeded on new MEF feeder cells. Cloned ES-like colonies were subjected to further analysis. Normal iPSC line (HPS0063) was obtained from the RIKEN Bioresource Center (Takahashi et al., 2007b).

Neural differentiation of iPSCs was performed as described previously (Okada et al., 2008). iPSC colonies were detached from feeder layers and cultured in suspension as embryonic body for ∼30 d in bacteriological dishes. Embryonic bodies were then enzymatically dissociated into single cells and the dissociated cells cultured in suspension in serum-free media hormone mix media (Okada et al., 2008) for 10–14 d to allow the formation of neurospheres. Neurospheres were passaged repeatedly by dissociation into single cells followed by culture in the same manner. Typically, neurospheres between passages 3 and 8 were used for analysis. For terminal differentiation, dissociated neurospheres were allowed to adhere to poly-l-ornithine– and laminin-coated coverslips and cultured for 10 d.

AP staining was performed using an ES-AP detection kit (EMD Millipore) according to manufacturer’s recommendations. SA-β-gal activity was detected using SA-β-gal staining kit (Cell Signaling Technology) according to manufacturer’s protocol. For immunocytochemical analysis, we used anti-SSEA4, TRA-1-60, TRA-1-81 (mouse, 1:100; EMD Millipore), anti–β-III-tubulin (mouse, 1:400; EMD Millipore), rabbit anti–LC-3B (1:200; Cell Signaling Technology), rabbit anti-ASM (1:1,000, Abcam), mouse anti-LAMP1 (1:100; Abcam), and mouse anti-LBPA (1:500; Echelon).

Established iPSCs were prepared at 10⁷ cells/ml in PBS. Suspended cells (1–3 × 10⁶) were injected into testes of anesthetized male SCID Beige mice. 8 wk after transplantation, mice were sacrificed and tumors were dissected. Tumor samples were fixed in 10% formalin and embedded in paraffin. Sections were stained with hematoxylin and eosin.

Comparisons between two groups were performed with Student’s t test. In cases where more than two groups were compared with each other, a one-way analysis of variance (ANOVA) was used, followed by Tukey’s HSD test. All statistical analysis was performed using SPSS statistical software. P < 0.05 was considered to be significant.

Table S1 shows subjects’ characteristics. Table S2 shows sequences of primer pairs.

This work was supported by the Bio & Medical Technology Development Program (2010-0020234, 2011-0019356, 2012M3A9C6049913, and 2012M3A9C6050107) of the National Research Foundation (NRF) of Korea funded by the Ministry of Science, ICT & Future Planning, Republic of Korea.

The authors declare no competing financial interests.

Author contributions: J.K. Lee, H.K. Jin, M.H. Park, B.R. Kim, P.H. Lee, H. Nakauchi, J.E. Carter, and X. He performed experiments and analyzed data, J.K. Lee, H.K. Jin, and J.S. Bae designed the study and wrote the paper. E.H. Schuchman and J.S. Bae interpreted the data and reviewed the paper. All authors discussed results and commented on the manuscript.