Axonal generation of amyloid-β from palmitoylated APP in mitochondria-associated endoplasmic reticulum membranes

Raja Bhattacharyya; Sophia E Black; Madhura S Lotlikar; Rebecca H Fenn; Mehdi Jorfi; Dora M Kovacs; Rudolph E Tanzi

doi:10.1016/j.celrep.2021.109134

. Author manuscript; available in PMC: 2021 Jul 19.

Published in final edited form as: Cell Rep. 2021 May 18;35(7):109134. doi: 10.1016/j.celrep.2021.109134

Axonal generation of amyloid-β from palmitoylated APP in mitochondria-associated endoplasmic reticulum membranes

Raja Bhattacharyya ^1,^*, Sophia E Black ¹, Madhura S Lotlikar ¹, Rebecca H Fenn ¹, Mehdi Jorfi ¹, Dora M Kovacs ¹, Rudolph E Tanzi ^1,^2,^*

PMCID: PMC8287518 NIHMSID: NIHMS1706347 PMID: 34010653

SUMMARY

Axonal generation of Alzheimer’s disease (AD)-associated amyloid-β (Aβ) plays a key role in AD neuropathology, but the cellular mechanisms involved in its release have remained elusive. We previously reported that palmitoylated APP (palAPP) partitions to lipid rafts where it serves as a preferred substrate for β-secretase. Mitochondria-associated endoplasmic reticulum (ER) membranes (MAMs) are cholesterol-rich lipid rafts that are upregulated in AD. Here, we show that downregulating MAM assembly by either RNA silencing or pharmacological modulation of the MAM-resident sigma1 receptor (S1R) leads to attenuated β-secretase cleavage of palAPP. Upregulation of MAMs promotes trafficking of palAPP to the cell surface, β-secretase cleavage, and Aβ generation. We develop a microfluidic device and use it to show that MAM levels alter Aβ generation specifically in neuronal processes and axons, but not in cell bodies. These data suggest therapeutic strategies for reducing axonal release of Aβ and attenuating β-amyloid pathology in AD.

In brief

Bhattacharyya et al. show that the modulation of mitochondrial-associated endoplasmic reticulum membranes (MAMs) via sigma-1 receptor regulates Aβ generation from axons via cell surface trafficking and β-secretase cleavage of MAM-resident palmitoylated APP (palAPP).

Graphical Abstract

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INTRODUCTION

The subcellular localization of amyloid-β (Aβ) generation and downstream effects on neurons are key events in Alzheimer’s disease (AD) neuropathogenesis (Simons 1995; Yamazaki et al., 1995; Koo et al., 1996; Muresan et al., 2009). Synaptic and neuritic Aβ can be generated via processing of APP in neuronal processes (Kamal et al., 2001; Cirrito et al., 2008; Tampellini et al., 2009; Das et al., 2013), ~40% of neuronal Aβ production takes place in axons (Niederst et al., 2015; Das et al., 2016). However, the molecular mechanisms underlying axonal generation of Aβ remain elusive.

To transport proteins to the cell surface, axons use endoplasmic reticulum (ER)-rafts or mitochondria-associated ER membranes (MAMs), found on the ER juxtaposed with mitochondria (Merianda et al., 2009). 5%–20% of the mitochondrial surface associates with the ER to form MAMs (Hajnóczky et al., 2006; Rowland and Voeltz, 2012). MAMs move bidirectionally as isolated small ER (sER) vesicles in neuronal processes (Pizzo and Pozzan, 2007; Giorgi et al., 2009). Mitochondria are essential for the formation and maintenance of synapses and move along axonal microtubules, neurofilaments, and actin tracks (Kamal et al., 2001; Cirrito et al., 2008; Tampellini et al., 2009; Das et al., 2013).

MAM function has been shown to be significantly increased in fibroblasts from AD patients (Area-Gomez et al., 2012). Smaller MAMs (<10 nm) are found in hippocampal neurons in APP transgenic rats (Martino Adami et al., 2019). APP, BACE1, and γ-secretase components are all found in MAMs (Schon and Area-Gomez, 2013; Erpapazoglou et al., 2017), and MAMs bear remarkable similarity to lipid rafts, preferred subcellular microdomains for β-secretase cleavage of APP and Aβ generation (Urano et al., 2005; Vetrivel et al., 2005, 2009; Kosicek et al., 2010). Other proteins enriched in MAMs (e.g., inositol-1,4,5-trisphosphate receptor subunit 3 [IP3R3], sigma-1 receptor [S1R], and acyl-coenzyme A:cholesterol acyltransferase [ACAT]) (Schon and Area-Gomez, 2013) have emerged as AD drug targets (Bhattacharyya and Kovacs, 2010).

We recently reported APP is palmitoylated in the ER prior to localizing to lipid rafts, where it serves as a preferential substrate for cleavage by β-secretase (Bhattacharyya et al., 2013). Reduced palmitoylation leads to ER-retention and slower maturation of APP (Bhattacharyya et al., 2013). We previously showed that cortical extracts from mice exhibit age-dependent increases in palmitoylated APP (palAPP). Treatment with palmitoylation inhibitors (e.g., cerulenin [Cer] and 2-bromopalmitate [2-BP]) prevents APP palmitoylation and lowers Aβ production. Recently, we reported palAPP forms strong cis-dimers that undergo β-secretase cleavage in detergent resistant membranes (DRMs) or lipid rafts (Bhattacharyya et al., 2016). Loss of the ER/MAM-resident enzyme, acyl-coenzyme A:ACAT (Rusiñol et al., 1994; Lewin et al., 2002), reduces secreted Aβ by up to 92% (Puglielli et al., 2001; Hutter-Paier et al., 2004; Huttunen et al., 2007, 2010; Bhattacharyya and Kovacs, 2010; Bryleva et al., 2010; Murphy et al., 2013; Shibuya et al., 2015). ACAT inhibition decreases levels of lipid raft palAPP and Aβ generation by up to 76% (Bhattacharyya et al., 2013). Thus, palmitoylation plays an important role in APP metabolism and Aβ generation (Bhattacharyya et al., 2013).

Here, we investigated the role of MAMs in APP processing and trafficking and axonal generation of Aβ. We employed differentiated neurons from human neuronal progenitor cells expressing APP with familial AD (FAD) mutations (FAD hNPCs) in a three-dimensional (3D) neural cell culture model of AD (Choi et al., 2014). We also employed a biochemical assay to separate palAPP from non-palAPP and total APP (APP_tot) and developed a microfluidic system to study MAMs in axonal versus somal microenvironments in neurons. First, we show that association of palAPP with MAMs regulates the processing and cell surface localization of palAPP and Aβ generation. Second, we show that modulating MAM levels directly influences β-secretase cleavage of palAPP and Aβ generation. Third, we show that MAM-based generation of Aβ occurs specifically in neuronal processes and axons. These findings carry major implications for developing AD therapies based on targeting MAM-associated palAPP in axons to alleviate Aβ pathology.

RESULTS

palAPP is predominantly localized to MAMs in human neuronal cells and in mouse brains

To quantify the distribution of palAPP in lipid rafts versus MAMs in AD neuronal cells, we compared palAPP levels in flotillin-positive raft fractions (lipid rafts) versus inositol-1,4,5-trisphosphate receptor subunit 3 (IP3R3)-positive raft fractions (MAMs) in ReN-VM neural progenitor stem cells (hNPCs) constitutively overexpressing human APP containing K670N/M671L (Swedish) and V717I (London) FAD mutations (APP_Swed/Lon) (Choi et al., 2014). FAD-APP (APP_Swe/lon) expressing hNPCs (FAD hNPCs) were generated by transfecting hNPCs with IRES-mediated polycistronic lentiviral vectors encoding human APP_Swe/lon with GFP as a reporter. Fluorescence-activated cell sorting (FACS) was employed to enrich the population. FAD hNPCs differentiated in a 3D matrix recapitulate β-amyloid and Tau pathology (D’Avanzo et al., 2015; Kim et al., 2015; Kwak et al., 2020). We metabolically labeled FAD hNPCs with chemically reactive alkylene-palmitic acid (Alkyl-C16) for 18 h, as described previously (Bhattacharyya et al., 2013). Labeled cells were subjected to sucrose gradient centrifugation (Figure 1A), and fractions were probed with the raft marker, flotillin, and MAM-marker, IP3R3. Most flotillin distributed to fraction 3, whereas most IP3R3 partitioned to fraction 4 with some overlap, indicating separation of post-ER rafts and MAMs. Although we detected equal amounts of APP_tot in raft fractions positive for flotillin (PM-rafts) or IP3R3 (MAMs), most APP_tot partitioned to the non-raft fractions (fractions 9 and 10) (Figure 1A), as expected. ABE palmitoylation assay revealed that ~10% of APP_tot is palmitoylated (palAPP) in FAD hNPCs (Figure S1), consistent with our previous findings (Bhattacharyya et al., 2013, 2016). To detect palAPP distribution in FAD hNPCs, we subjected each fraction to click-iT addition of TAMRA (tetramethylrhodamine) label, as previously described (Bhattacharyya et al., 2013) (Figure 1A, click-iT palmitoylation). In flotillin-positive lipid rafts, palAPP constituted only 20.7% ± 3.1% of APP_tot, whereas in IP3R3-positive MAMs, palAPP was enriched to 68.12% ± 8.8% of APP_tot (Figure 1B). Meanwhile, <2% of APP_tot was palmitoylated in non-rafts fractions 9 and 10 (Figure 1B).

Figure 1. — (A) FAD hNPCs were plated on a thin-layer 3D matrix prior to differentiation. Lipid rafts and non-raft fractionation of 10-day differentiated FAD hNPCs pre-labeled with reactive alkylene-palmitic acid (alkyl-C16) prior to western blot analysis. First, second, and third blots: detection of total APP (APP_tot) by probing with anti-APP C-terminal antibody (C66), detection of rafts by probing with anti-flotillin, or anti-IP3R3 antibodies, respectively. Fourth blot: detection of palmitic acid labeled APP (palAPP) after TAMRA (tetramethylrhodamine)-labeling via click-iT reaction and probing with anti-TAMRA antibody.

(B) Quantitation of palAPP in flotillin- and IP3R3-positive fractions shown in (A) normalized to APP_tot. Data represent Mean ± SEM (n = 3, p < 0.004).

(C) Fractionations of MAMs and non-MAMs from crude ER-mitochondria membranes isolated from FAD-NPCs. Fractions were probed for indicated proteins.

(D) Badrilla palmitoylation assay to isolate palmitoylated and non-palmitoylated proteins from MAM and non-MAM fractions isolated in (C).

(E) Quantitation of palAPP and non-palAPP in MAMs and non-MAMs. Data represent Mean ± SEM (n = 3, p < 0.01).

(F) MAM-fractionation of total membrane extract (TM) from mouse brain via sequential extraction and Percoll gradient centrifugation to isolate crude ER- mitochondria (ER/mito), pure mitochondria (mito), and pure MAMs. Fractions were subjected to ABE palmitoylation assay followed by probing total APP (input) and ABE isolated palAPP (ABE) in each fraction with anti-APP(C-term) antibody (IB: C66). Fractions were probed with indicated antibodies. IP3R3 and ACAT1 partitioned in the MAMs. VDAC1 partitioned in both mito and MAMs. Cyto C was detected exclusively in the mito fractions. Representative image of two separate and independent experiments.

To confirm the localization of palAPP in MAMs, we extracted crude ER-mitochondria (ER-mito) from FAD hNPCs and used sucrose density gradient centrifugation to isolate MAMs (IP3R3 and ACAT-rich fractions) and non-MAMs (fractions 8–10; Figure 1C). APP_tot distributed into both MAM and non-MAM fractions (Figure 1C). To assess palAPP distribution, the pooled MAM and non-MAM fractions were subjected to the recently developed Badrilla palmitoylation assay (Figure 1D). This assay captures palmitoylated proteins by exchanging palmitic acid with a thiol-bound CAPTUREome resin following thioester cleavage and isolates non-palmitoylated proteins in the CAPTUREome-unbound fractions (Figure S2). We observed strong staining for palAPP in MAM fractions, with little or no evidence of non-palmitoylated APP (non-palAPP) (Figure 1D). Conversely, non-palAPP was mainly distributed in the non-MAM fractions (Figure 1D). Quantitative analysis revealed ~9.8-fold more palAPP was associated with MAMs as compared to non-palAPP. In contrast, ~10.5-fold more non-palAPP distributed in non-MAMs compared to palAPP (Figure 1E). Thus, MAMs almost exclusively contain palAPP.

We next asked whether palAPP localizes to MAMs in mouse brain. Mouse brain homogenates were subjected to Percolgradient centrifugation as previously described (Wieckowski et al., 2009). Total membrane (TM), crude mixture of ER-mitochondria (ER/mito), ER, mitochondria (mito), and MAMs were isolated (Figure 1F). Equal amounts of proteins were subjected to the acyl-biotin exchange (ABE) palmitoylation assay and probed with an anti-APP antibody (C66) to detect palAPP in the indicated membranes (Figure 2, ABE, IB:C66). In addition to detecting APP, BACE1, IP3R3, and ACAT1 in MAMs, as previously described (Area-Gomez et al., 2009, 2012), we detected palAPP in MAMs (Figure 2). Thus, palAPP mainly resides in MAMs.

Figure 2. — (A) Representative confocal microscopic images of FAD hNPCs after silencing of sigma1 receptor (S1R) expression. Cells were either introduced with a non-targeting siRNA (si-non) or with a SMART pool of siRNAs against human sigma1 receptor gene Sigmar1 (si-S1R) for 48 h. Left: GFP-expressing (green) FAD NPCs containing si-non or si-S1R. Right: endogenous S1R (red) expression in cells containing si-non or si-S1R.

(B) Representative immunoblots of cell lysates (TCL) from FAD NPCs containing si-non or si-S1R probed with indicated antibodies.

(C) Quantitative analysis of S1R and IP3R3 levels from 3B. Data were analyzed using ImageJ and expressed as fold ± SEM relative to control (si-non) (n = 3, *p < 0.005).

(D) Representative electron micrographs (EM) showing ER (ER)-mitochondria (M) contact sites or MAMs (arrowheads) in FAD NPCs containing si-non or si-S1R for 48 h.

(E) Quantitation of MAMs in control (si-non) and S1R-silenced (si-S1R) cells per mitochondria per frame (MAM per M). More than 8 frames were used for each analysis (n = 3, *p < 0.01).

(F) Immunoblot of FAD NPCs treated with increasing amounts of S1R agonist PRE-084 or with S1R antagonist NE-100. MAM levels were identified by probing with antibody against MAM-protein IP3R3 (IB:anti-IP3R3). Levels of mitochondrial protein GRP74 or housekeeping protein GAPDH were detected by probing with anti-GRP74 or anti-GAPDH antibodies, respectively.

(G) Quantitative analyses of IP3R3 levels from untreated (0 μM), PRE- (5 and 10 μM), or NE- (5 and 10 μM) treated cells. IP3R3 levels were presented as fold ± SEM (n = 3, *p < 0.01).

(H) Immunoblots of pure MAM fractions (MAMs) isolated from crude ER/mitochondria (ER/mito) of FAD hNPCs after treatment with control vehicle (veh) or with 10 μM PRE or NE for 16 h. Crude ER-mitochondria extracts (ER/mito), pure ER (ER), pure mitochondria (mito), and pure MAM (MAMs) fractions were probed with antibodies against MAM-resident IP3R3 or with mitochondrial marker Cyto C. Representative image of a triplicate experiments was presented.

(I) Quantitation of the IP3R3 levels in the MAMs from control (veh), PRE- or NE-treated cells. The levels were represented as fold ± SEM.

Silencing of MAM-resident sigma 1 receptor (S1R) reduces MAM levels in FAD NPCs

S1R is a chaperone protein residing in MAMs, where it interacts with proteins such as IP3R3, which anchors the outer mitochondrial membrane protein voltage-dependent anion channel isoform 1 (VDAC1) to the ER-associated molecular chaperone glucose-regulated protein 74 (GRP74) (Hayashi and Su, 2007). Loss or inhibition of MAM-resident S1R disrupts MAM-assembly by destabilizing the MAM-resident IP3R3 (Hayashi and Su, 2007). Genetic ablation of the S1R gene in mouse brain (sigmar1^—/—) results in a robust decrease in IP3R3 levels and significant reduction of ER-mito contact sites (~30% of mitochondria forming MAMs in wild-type neuron versus ~18% in sigmar1^—/— neurons) compared to non-transgenic control mice (Bernard-Marissal et al., 2015).

To study the effect of disrupted MAMs on the processing and β-secretase cleavage of palAPP, we silenced the expression of S1R in FAD hNPCs, using a SMART pool small interfering RNA (siRNA) against sigmar1 (si-S1R) (Amer et al., 2013). We performed the CytoTox-ONE assay to measure lactate dehydrogenase (LDH) levels in the culture media to assess cell viability after introducing si-S1R for 0, 48, and 72 h. We observed ~77% reduction of S1R expression after 48 h, and >90% reduction after 72 h transfection with si-S1R. The LDH-assay revealed that 72 h transfection with si-S1R led to ~53% loss of cell viability, whereas 48 h transfection exhibited little or no reduction in cell viability (Figure S2). Cells transfected with si-S1R for 48 h were fixed and subjected to confocal and EM analyses and immune-labeling with anti-S1R antibody. Confocal microscopy revealed significant reduction of S1R expression in S1R-silenced (si-S1R) cells, compared to control cells (si-non) (Figure 2A). Equal amounts of proteins from si-non and si-S1R cells were probed for IP3R3 and ACAT1. si-S1R cells exhibited dramatic reductions of IP3R3 and ACAT1 levels compared to si-non cells (Figure 2B), indicating that silencing of S1R reduced MAM levels in FAD hNPCs. Meanwhile, S1R-knockdown had little or no effect on VDAC1, a primarily mitochondrial protein, or GRP74, a primarily ER-protein (Figure 2B), suggesting that S1R-silencing specifically affected MAMs, not bulk mitochondria or ER. Quantitation revealed ~85% reduction of S1R expression in si-S1R cells compared to si-non cells, whereas IP3R3 levels decreased by ~67% in the same cells (Figure 2C). Thus, silencing S1R expression effectively downregulates MAMs.

Next, S1R-silenced and control cells were subjected to transmission electron microscopy (TEM) to identify ER-mito contact sites or MAMs. TEM revealed robust reduction of ER-mito contact sites in si-S1R cells compared to si-non cells (Figure 2D). MAMs were quantified directly by counting the number of ER-mito contact sites (5–20 nm size) per mitochondria per field of the electron micrographs (Bernard-Marissal et al., 2015). Quantitation revealed ~67% reduction in ER-mito contact sites or MAMs per mitochondria (MAMs per mito) in si-S1R cells versus si-non cells (Figure 2E). Thus, silencing S1R expression in FAD hNPCs significantly reduces MAM levels.

S1R-agonist PRE-084 and S1R-antagonist NE-100 regulate MAM levels in FAD hNPCs

S1R function can be regulated using the S1R agonist PRE-084 (PRE) and antagonist NE-100 (NE) (Bernard-Marissal et al., 2015). Total IP3R3 levels were increased in the presence of 5.0 or 10.0 μM PRE and decreased following treatment with 5.0 or 10.0 μM in FAD hNPCs (Figure 2F). 10 μM PRE increased total IP3R3 levels by ~43% (1.43- ± 0.2-fold; p < 0.05, n = 3), whereas treatment with 10 μM NE decreased total-IP3R3 levels by ~20% (0.79- ± 0.08-fold; p < 0.05, n = 3) versus vehicle control (veh) cells (Figure 2G).

To confirm regulation of S1R-activity modulated MAM-associated IP3R3 levels, we isolated MAM, ER, mixed ER/mitochondria, and mitochondria fractions from FAD hNPCs following pre-treatment with 10 μM PRE or NE. The fractions were probed with antibodies against IP3R3 and cytochrome C (cyto C) to determine the purity of MAMs and of mitochondria (mito), respectively (Figure 2H). MAM-IP3R3 levels were increased (3.73- ± 0.43- fold) following treatment with PRE and significantly reduced (~55%; 0.45- ± 0.13-fold) following treatment with NE compared to veh cells (Figures 2H and 2I). The significant effects of S1R-activation or -inactivation on MAM-IP3R3 levels as opposed to the more modest effects on total-IP3R3 suggest S1R preferentially modulates MAMs in FAD hNPCs.

Modulation of MAMs via S1R stabilizes palAPP in FAD-NPC neurons

Palmitoylation localizes and stabilizes transmembrane proteins in lipid-rich membrane compartments (Linder and Deschenes, 2007). We previously reported palmitoylation extends the half-life of APP, from t_1/2 = 2 h for total APP to t_1/2 >4 h for palAPP, indicating palAPP is twice as stable as total APP (Bhattacharyya et al., 2016). To test whether association with MAMs stabilizes palAPP, we measured the half-life of palAPP in differentiated FAD-NPC neurons following upregulation of MAMs with 10 μE or downregulation with 10 μM NE. MAM-dependent palAPP stability was assessed by cycloheximide (CHX)-chase experiments (Bhattacharyya et al., 2016). To measure palAPP levels, ABE palmitoylation assays were performed after 0, 1, 2, 4, and 6 h of chase with cycloheximide (Figure 3A). ~40% of palAPP levels remained after 4 h chase, confirming our previous studies (Figure 3B) (Bhattacharyya et al., 2016). Treatment with PRE stabilized palAPP: ~90% of palAPP remained after the 4-h chase (Figure 3B). Conversely, treatment with NE dramatically reduced palAPP stability: ~20% of palAPP remained after 4 h of chase (Figure 3B). Meanwhile, neither PRE- nor NE-treatment altered the stability of APP_tot.

Figure 3. — (A–E) FAD-NPCs were subjected to cycloheximide chase analysis followed by ABE (A and B) or Badrilla (C–E) palmitoylation assays to determine half-life of *pal*APP in absence or presence of S1R agonist/antagonist.

(A) ABE palmitoylation assay of FAD NPCs pre-treated without (veh) or with 10 μM PRE-084 (PRE) or NE-100 (NE) prior to cycloheximide (CHX) chase for indicated time intervals (0–18 h) revealed an increased half-life of *pal*APP in the presence of PRE and a decreased half-life of *pal*APP in the presence of NE in comparison to untreated (veh) cells (ABE palmitoylation assay). Half-life of total APP (APP_tot) remained unaltered in presence of PRE or NE.

(B) Quantitative analysis of *pal*APP/APP_tot after cycloheximide chase in absence (veh) or presence of PRE or NE (n = 3).

(C) Representative western blots of Badrilla palmitoylation assay detecting palmitoylated APP (*pal*APP) (CAPTUREome, IP:C66) and non-palmitoylated APP (non-*pal*APP) (unbound, IB:C66) from cells chased with CHX for 0, 4, and 6 h in absence (veh) or presence of PRE and NE.

(D) Quantitation of *pal*APP/APP_tot in absence (veh) or presence of PRE or NE after CHX-chase.

(E) Quantitation of non-*pal*APP/APP_tot. Error bars represent percent (%) ± SEM (n = 3).

Next, we compared the half-life of palAPP versus non-palAPP using the Badrilla palmitoylation assay. We observed a marked decrease in palAPP levels after 4 h of chase following CHX-treatment in the control (veh) experiment (Figure 3C). A similar reduction of palAPP levels was observed only after 6 h of chase in cells pre-treated with PRE (Figure 3C), suggesting stabilization. Conversely, palAPP levels were significantly decreased in NE-treated cells (Figure 3C). 41.1% ± 2.2% of palAPP remained after 6 h of chase in control cells. 68.9% ± 1.7% versus 17.0% ± 1.3% of palAPP remained after 4 h of chase in cells pre-treated with PRE or NE, respectively (Figure 3D), whereas the stability of non-palAPP remained unchanged by either PRE- or NE- treatment after 4 h of chase: ~40% of non-palAPP remained in control, PRE-, or NE-treated cells (Figure 3E). Thus, S1R-activation specifically stabilizes palAPP but not APP_tot or non-palAPP. Conversely, S1R-inactivation specifically destabilizes palAPP.

MAM assembly regulates levels of cell surface-associated palAPP but not total APP

We next asked whether modulation of MAM assembly affects levels of cell surface-associated palAPP. We used a two-step pull-down assay to simultaneously detect cell surface palmitoylated and non-palmitoylated APP in FAD hNPCs, which were first surface biotinylated with sulfo-NHS-SS-biotin. Surface biotinylated-palmitoylated proteins were isolated from Neutravidin beads using tris(2-carboxyethyl)phosphine (TCEP) prior to carrying out the Badrilla palmitoylation assay (Figure S4). To test the effect of MAM formation on cell surface-associated palAPP, we performed the two-step pull-down assay in untreated FAD hNPCs and those treated with 10 μM PRE or NE (Figure 4A). We detected a robust (~1.5-fold) increase of surface labeled palAPP on PRE-treatment, whereas NE-treatment resulted in ~30% loss of cell surface palAPP (Figure 4B). In contrast, treatment with neither PRE nor NE affected cell surface biotinylation of APP_tot or non-palAPP (Figures 4A and 4B). Cell surface association of the control protein transferrin receptor (Tfr) also remained unaffected following PRE- or NE-treatment (Figures 4A and 4B), consistent with Tfr being nearly absent in MAMs (Sala-Vila et al., 2016). Thus, MAM-assembly specifically promotes cell surface association of palAPP without affecting cell surface association of APP_tot.

Figure 4. — (A) Badrilla palmitoylation assay of biotinylated cell surface proteins of FAD NPCs pre-treated without (+veh) or with PRE-084 (PRE) or NE-100 (NE). Left panels (a–c): western blot detecting increased or decreased cell surface *pal*APP in presence of PRE or NE, respectively, in comparison to untreated (veh) cells (a). Cell surface total APP (APP_tot) and flotillin are represented in b and c. Right panel: representative western blot images of *pal*APP, APP_tot, or flotillin in total cell lysates from untreated (veh), PRE- or NE-treated cells.

(B) Quantitation showed ~1.5-fold increase of cell surface *pal*APP after activation of S1R by PRE-treatment, whereas inactivation of S1R by NE-treatment significantly decreased cell surface *pal*APP level. Neither PRE- nor NE-treatment had any significant effect on cell surface association of either non-*pal*APP or Tfr. Error bars are fold ± SEM (n = 2, **p < 0.05).

Regulation of MAM-assembly in FAD hNPCs via S1R modulates β-secretase cleavage of palAPP

Next, we asked whether silencing the expression of S1R in FAD hNPCs attenuates cleavage of APP by β-secretase. si-S1R decreased ER-mito contact sites by ~70% (Figure 2). The ABE palmitoylation assay revealed that silencing S1R significantly reduced palAPP levels in FAD hNPC cell lysates (Figure 5A, ABE assay, palAPP). In contrast, si-S1R led to no significant differences in levels of palmitoylated flotillin versus control cells (not shown) and only a small reduction in levels of APP_tot (Figure 5A, ABE assay, APP_tot). Quantification of the reduction of palAPP in si-S1R cells in comparison to APP_tot (palAPP/APP_tot) revealed 48.61% ± 14.60% (p < 0.01, n = 3) palAPP/APP_tot in si-S1R cells as compared to si-non (Figure 5B), indicating an ~51% reduction of palAPP levels following S1R-silencing. We detected a smaller (14.72% ± 4.77%; p < 0.03, n = 3) decrease in APP_tot in si-S1R cells versus si-non (data not shown), in agreement with 10%–20% of APP_tot consisting of palAPP.

Figure 5. — (A) Upper panel: ABE palmitoylation assay of control (si-non) and S1R-silenced (si-S1R) FAD NPCs detected lower level of *pal*APP on S1R silencing (ABE assay). Lower panel: conditioned media (CM) from control (si-non) and S1R-silenced (si-S1R) cells showed reduction of soluble sAPPβ (sAPPβ) level but not that of total sAPP (sAPPtot).

(B) Quantitative analyses of *pal*APP or sAPPβ levels from si-non and si-S1R cells. *pal*APP levels were normalized with total APP level (*pal*APP/APP_tot) and sAPPβ levels were normalized with total sAPP level (sAPPβ/sAPP_tot). We used ImageJ analysis and expressed the data as fold ± SEM (n = 3, *p < 0.01).

(C) ABE palmitoylation assay from FAD NPCs demonstrated increase or decrease of palmitoylated sAPPβ (*pal*-sAPPβ) release from cells treated with PRE-084 (PRE) or NE-100 (NE), respectively, as compared to untreated (veh) cells (ABE IB:anti-sAPPβ). Total sAPPβ (sAPPβ), sAPP (sAPP_tot,) and sAPPα (sAPPα) were detected in the conditioned media (CM, IB:anti-sAPPβ, IB:22C11 and IB: anti-sAPPα, respectively).

(D) Quantitation of *pal*-sAPPβ release compared to total sAPPβ release from untreated (veh), PRE-, or NE-treated cells. The data were represented as fold ± SEM (n = 4, **p < 0.01).

We next asked whether decreased palAPP levels following S1R-silencing affects β-secretase cleavage of APP. Conditioned media from si-non and si-S1R cells probed revealed a robust reduction in sAPPβ release relative to total sAPP following si-S1R (Figure 5A, CM, sAPPβ, or sAPP_tot, respectively). The ratio of sAPPβ/sAPP_tot in conditioned media from si-S1R cells was 44.62% ± 16.84% (p < 0.01, n = 3) of that observed in si-non cells (Figure 5B), with an ~58% reduction in sAPPβ release following si-S1R. Thus, ~70% downregulation of MAM-assembly following silencing of S1R leads to significant reduction in β-secretase cleavage of APP, most likely by reducing palAPP levels.

Next, we measured pal-sAPPβ release from FAD hNPCs treated with 10 μM PRE or NE. Figure 4D shows that treatment with PRE increased MAM-IP3R3 levels by ~3.7-fold, whereas NE decreased MAM-IP3R3 levels by ~45%. Next, pal-sAPPβ levels were assessed by the ABE palmitoylation assay as done before (Bhattacharyya et al., 2013). Treatment with PRE led to a significant increase in pal-sAPPβ release compared to veh cells (Figure 7C, ABE, IB:anti-sAPPβ). In contrast, pal-sAPPβ release was dramatically decreased following treatment with NE (Figure 5C, ABE, IB:anti-sAPPβ). Neither PRE nor NE-treatment led to any significant effect on total sAPP (sAPP_tot) or sAPPβ release (Figure 5C), nor on APP-C-terminal fragment (CTF) levels (Figure S6). In comparison to sAPPβ_tot-release, pal-sAPPβ (pal-sAPPβ/sAPPβ_tot) was increased by 2.14- ± 0.39-fold (p < 0.001, n = 3) following treatment with PRE (Figure 5D). Conversely, NE-treatment reduced pal-sAPPβ release by ~50% (0.508- ± 0.073-fold as compared to control, p < 0.001, n = 3) (Figure 5D). Thus, modulation of MAM levels via S1R specifically affects β-secretase but not α-secretase cleavage of palAPP and does not affect β- and α-secretase cleavage of APP_tot.

Figure 7. — (A) Confocal microscopy of FAD hNPCs seeded in a Matrigel 3-D matrix on the somal chambers of microfluidic devices to develop axons along the capillaries (arrows). Hoechst (blue) labeled nuclei in the somal chamber, indicating cell bodies. GFP signals (green) showed cell morphology. Antibodies against axonal markers neurofilament H (NFH)-labeled (red) or tau (tau)-labeled (white) axons in axonal chambers.

(B) Aβ ELISA of conditioned media isolated from axonal chambers of the microfluidic devices undergoing a fluidic separation confirmed increased or decreased number of Aβ₄₀ molecules per axon after treatment with PRE-084 (PRE) or NE-100 (NE), respectively. Number of Aβ calculated by measuring Aβ₄₀ (in pM) levels in the conditioned media. Although PRE-treatment increased number of Aβ₄₀-release from axons by ~3-fold, NE-treatment decreased axonal Aβ₄₀-release to undetectable level. No significant changes in the number of Aβ₄₀ molecules released per cell body were detected after PRE- or NE-treatment. Treatment with BACE-inhibitor IV (BI-IV) reduced both somal and axonal Aβ₄₀ levels, as expected. Error bars indicated SEM from three independent experiments (**p < 0.01).

Regulation of S1R-activity specifically modulates MAMs in neuronal processes

To test for S1R-dependent changes in MAM-assembly in neuronal processes versus cell bodies, we performed confocal microscopy to visualize the ER-mito contact sites in cells pre-labeled with cell-permeable GFP-tagged ER (CellLight ER-GFP) and RFP-tagged mitochondrial (CellLight mito-RFP) probes. A MAM-resident Ca²⁺-sensor protein Miro has been shown to be co-transported with mitochondria down axons (Misko et al., 2010). Elevated calcium levels halt axonal transport of Miro, indicating MAM-transport along axons could be regulated by physiological stimuli (Macaskill et al., 2009; Wang and Schwarz, 2009).

We first confirmed that the GFP- and RFP-tagged probes could correctly identify ER-mito contact sites in mouse cortical neurons from C57BL/6 embryonic day 16 (E-16) mice. Confocal microscopy revealed that ER-GFP (green fluorescence) and mito-RFP (red fluorescence) distinctly labeled ER and mitochondria, respectively, in cell bodies as well as in processes (Figure 6A, a–c). The overlapping areas between ER and mitochondria (ER-mito) were identified as MAMs (Figure 6A, d, white arrow). Employing a modified MAM-imaging technique (Filadi et al., 2015) that eliminated all non-overlapping fluorescence (representing ER and mitochondria), we were able to highlight the contact areas (white) (Figure 6A, e–g) and demonstrated MAMs in the processes (Figure 6A, h–j) as well as in cell bodies (Figure 6A, k–m).

ER-mito contact sites >10 nm in both processes (Figure 6A, n–p) and in cell bodies (not shown) were counted as MAMs, using the ImageJ particle measurement tool. Quantitation of MAMs (ER-mito contact sites) revealed that treatment with 10 μM PRE increased MAMs in axons and neuronal processes by ~2.5-fold, without affecting MAMs in the cell bodies or soma of cultured primary neurons (Figure 6B). Conversely, treatment with 10 μM NE decreased ER-mito contact sites by ~50%, only in axons and neuronal processes (Figure 6B). Similar results were obtained when we counted ER-mito contact sites in neuronal processes of FAD hNPCs pre-labeled with the ER and mitochondria probes (Figure 6C). NE-treatment decreased ER-mito contact sites, whereas treatment with PRE increased ER-mito contact sites, primarily in neuronal processes, without altering ER-mito contact sites in cell bodies (Figure 6D). We next used a bidirectional fluorescent compensation (BiFC) assay to assess the effect of S1R-inactivation on axonal MAMs in cells co-expressing split GFP ER and mitochondria probes (ER-GFP(1–10) and mito-GFP11, respectively) (Yang et al., 2018). NE treatment reduced the number of MAMs per axon by ~35% (from 27.87 ± 1.52 MAMs/axon in control to 3.125 ± 1.23 MAMs/axons in NE-treated) (Figure S6). Thus, modulation of S1R activity regulates MAM assembly specifically in axons and neuronal processes, not in the cell body or soma.

To further validate that pharmacological regulation of S1R specifically affects axonal MAMs, we developed a microfluidic device (Figure 6E) that separates bulk neurons from axons following physical axotomy (Figure 6F). Although we accumulated a sufficient number of axons from 10 devices to obtain detectable levels of MAM-resident proteins including IP3R3, ACAT, and VDAC1 (Hedskog et al., 2013; Liu et al., 2019), this number of axons was insufficient for MAM purification. Thus, we assessed MAM levels in axons by measuring total levels of MAM-resident proteins, IP3R3, ACAT, and VDAC1, following axotomy (Figure 6G). We confirmed that the axonal fractions were not contaminated with bulk neurons following axotomy, by probing the fractions with an antibody against the nuclear envelope protein, lamin B1 (Figure 6G, IB:anti-lamin B1). We observed significant increases in all levels of all three axonal MAM-proteins measured following treatment with 10 μM PRE versus veh cells (Figure 6G). Conversely, the levels of all three MAM-proteins were dramatically reduced in axons following treatment with 10 μM NE (Figure 6G). Meanwhile, treatment with either PRE or NE had little or no effect on levels of IP3R3, ACAT, and VDAC1 in bulk neurons (Figure 6G). Quantitative analysis revealed that treatment with PRE increased levels of MAM-resident IP3R3, ACAT, and VDAC1 in the axons by 3.16- ± 0.41-fold, 1.4- ± 0.18-fold, and 1.79- ± 0.11-fold, respectively, as compared to axons from veh cells (Figure 6H). Conversely, axons from NE-treated cells contained 25% (0.25- ± 0.09-fold) IP3R3, 21% (0.21- ± 0.03-fold) ACAT, and only 3% (0.03- ± 0.008-fold) VDAC1 compared to veh cells (Figure 6H).

Modulation of MAMs in neuronal processes/axons regulates axonal Aβ generation from FAD hNPCs

APP and amyloidogenic processing enzymes, β- and γ-secretases, are localized in axons and dendrites (Kaether et al., 2000), and anterograde axonal transport delivers Aβ and sAPPβ to neurites around amyloid plaques. Axons have been heavily implicated in AD-related Aβ-generation and tau phosphorylation (Kamal et al., 2001; Cirrito et al., 2008; Tampellini et al., 2009; Das et al., 2013). Thus, we next investigated whether MAM-assembly in neuronal processes and axons affects Aβ generation. To measure axonal Aβ release, we employed a microfluidic chamber system (Xona Microfluidics) to perform fluidic separation of cell somas from axons in differentiated FAD hNPCs (Niederst et al., 2015). In this system, capillary channels draw axons from the somal chamber into the axonal chamber. To prevent cells from migrating into the axonal chamber, we plated FAD hNPCs in a 3D matrix on the somal side prior to differentiation. Both somal and axonal chambers were probed either with Hoechst dye to label the nuclei or with antibody against tau and/or neurofilament heavy (NFH) antibodies to label axons (Figure 7A, tau and NFH). We then carried out microfluidic isolation of the axonal microenvironments from bulk neuronal environments by establishing a small (30 μL) volume difference between axonal and somal compartments (Taylor et al., 2005). After 10–15 days in differentiating media, conditioned media from each chamber was subjected to Aβ ELISA to detect Aβ₄₀ levels (Aβ₄₂ levels were below detection limit). Levels of Aβ₄₀ in the somal chamber reached 372.4 ± 50.7 pM per chamber in 24 h. To account for axonal Aβ₄₀ in the somal chambers, we determined axonal numbers by subtracting the number of tau-positive axons in the axonal chambers from the number of cells (~30,000 DAPI-stained nuclei in the somal chambers) in the somal chambers, assuming each cell produced only one axon. Aβ₄₀-release in the axonal chambers was then calculated as the number of Aβ₄₀ molecules per tau-positive axon, using a previously described method of measurement (Niederst et al., 2015). We found differentiated FAD hNPC neurons generated 10.69 × 10⁸ ± 4.2 × 10⁸ Aβ₄₀ per cell bodies and 2.97 × 10⁸ ± 5.3 × 10⁶ Aβ₄₀ per axon in 24 h (Figure 7B).

Next, we pharmacologically modulated MAM assembly and tested for effects on Aβ generation in neuronal processes/axons versus neuronal cell somas. We measured Aβ₄₀ levels in the somal and axonal chambers of 10- to 15-day differentiated FAD hNPCs in microfluidic systems that received either vehicle (veh), 10 μM PRE, or 10 μM NE for 24 h. Neither PRE- nor NE-treatment altered Aβ₄₀ levels in the somal chamber. However, PRE treatment increased the number of Aβ₄₀ molecules released per axon by an additional 11.63 × 10⁸ ± 9.2 × 10⁷. In contrast, NE treatment reduced axonal Aβ₄₀ release to nearly undetectable levels (Figure 7B). Thus, modulation of MAM levels governs Aβ generation, specifically in axons and neuronal processes, but not in the cell soma.

DISCUSSION

We have previously shown that palmitoylation of APP targets palAPP to lipid rafts where it becomes a preferred substrate for β-secretase. Here, we show that palAPP is primarily localized to specialized lipid rafts known as MAMs, in both FAD hNPCs and in mouse brain. We also show that genetically silencing or pharmacologically inactivating the MAM-resident receptor, S1R (with the S1R-antagonist NE100) to downregulate MAM assembly (by destabilizing MAM-resident IP3R3), leads to greatly decreased levels of palAPP, reduced stability and cell surface trafficking of palAPP, decreased β-secretase cleavage of palAPP, and reduced generation of sAPPβ and Aβ. In contrast, the S1R-agonist PRE-084 upregulated MAM levels and increased palAPP stability and cell surface trafficking, β-secretase cleavage of palAPP, and Aβ generation. Furthermore, we show that modulation of S1R regulates MAM assembly, stability, and cell surface trafficking of palAPP, β-secretase cleavage of palAPP, and Aβ generation specifically in axons. Thus, we demonstrate a clear connection between MAM-assembly and Aβ generation from palAPP, specifically in axons and neuronal processes.

Previous studies have shown that SigR1 knock out (Sigmar1^—/—) decreases MAM-assemblies by nearly 50% (Bernard-Marissal et al., 2015). We demonstrated that silencing of S1R expression not only reduces MAMs (Figure 2) but also reduces the levels of palAPP with minimal to no effects on APP_tot levels (Figure 5A). We have previously shown that palAPP and/ or palAPP-dimers are better substrates for β-secretase cleavage in lipid-rich microdomains as compared to APP_tot (Bhattacharyya et al., 2013, 2016). Here, we show that levels of sAPPβ were reduced with silencing of MAM-resident S1R expression, and regulation of S1R activity by an agonist (PRE-084) or an antagonist (NE-100) not only modulates MAMs (Figure 2), but also specifically promotes or reduces β-secretase cleavage of palAPP, respectively (Figures 5C and 5D). Thus, MAMs and palmitoylation of APP directly modulate β-secretase cleavage of palAPP and subsequent Aβ generation.

The observation that S1R activation and inactivation oppositely modulate the stability of palAPP with no effects on APP_tot (Figure 3) is consistent with previous studies reporting evidence for palmitoylation-dependent stability of transmembrane proteins in lipid-rich membranes (Linder and Deschenes, 2007). Here, we show that modulation of S1R-activity affects cell-surface association of palAPP without affecting that of APP_tot (Figure 4). Although it remains unclear how association with MAMs promotes the cell surface association of palAPP, it is worth noting that ~11% of MAM-proteins are found in the plasma membrane (Poston et al., 2013). Additionally, MAM-like ER-rafts fusing with plasma membrane, called PAMs, have been detected in several tissues involved in regulating Ca²⁺ homeostasis, lipid trafficking, and mitochondrial structures (Szymański et al., 2017).

We also showed S1R activity mainly modulates MAM levels in axons and neuronal processes as opposed to neuronal cell bodies and soma (Figure 6). Thus, S1R-activity specifically regulates the assembly of a novel subset of MAMs enriched in axons and neuronal processes, while the assembly of MAMs in cell bodies is independent of S1R-activity. A previous study reported that S1R knockout only reduced MAM-assemblies in neurons by >50%, suggesting that more than half of MAM-assemblies were independent of S1R activity (Bernard-Marissal et al., 2015). Moreover, MAMs also differ in size and motility (Volpe et al., 1991; Bannai et al., 2004). We discovered that modulation of MAMs in neuronal processes via alteration of S1R-activity specifically affects β-secretase cleavage of palAPP without affecting cleavage of total APP_tot (Figures 5 and S5). Small molecule agonists and antagonists of S1R are currently being considered as therapeutics for numerous neurodegenerative diseases (Ryskamp et al., 2019).

Using a system incorporating FAD gene-expressing hNPCs grown in a 3D matrix inside microfluidic chambers, we were able to reveal that S1R regulates MAM-dependent Aβ generation exclusively in axons and neuronal processes, but not in soma or from bulk neurons (Figure 7). Axons and synapses are critical sites for APP-processing and Aβ production. (Lazarov et al., 2002; Sheng et al., 2002; Brendza et al., 2003; Spires et al., 2005). Previous studies have shown that intra-axonal Aβ is generated ahead of extracellular Aβ especially when axons are under stress (Suo et al., 2004). Moreover, amyloid deposition has been reported to decrease when β-secretase processing of APP and Aβ generation is shifted away from axonal to somal compartments (Lee et al., 2005). A recent study reported that ER-mitochondria tethering is increased in axons following injury (Lee et al., 2019). Here, we show that axons specifically use MAMs, containing ~70% of preferred β-secretase substrate, palAPP, to generate Aβ. Other studies have reported that Aβ and/or the β-cleaved C-terminal fragment of APP (C99 or APP-βCTF) modulate MAM function in AD (Schreiner et al., 2015; Pera et al., 2017). Knocking down the expression of MAM-resident mitofusin 2 (Mfn 2) resulted in increased ER-mito contact and decreased levels of Aβ (Leal et al., 2016). Our data reveal an additional converse mechanism whereby modulation of MAMs regulates Aβ generation in axons and neuronal processes. Collectively, these findings suggest a possible feed-forward mechanism for MAM-dependent AD pathogenesis.

The subcellular location in which APP primarily undergoes β-secretase cleavage in neurons has remained a topic of intensive investigation. We previously reported that APP undergoes palmitoylation targeting it to lipid rafts where it serves as a preferential substrate for β-secretase. In addition to lipid rafts in plasma membranes, intracellular MAMs have been increasingly investigated in neurodegenerative diseases such as AD and amyotrophic lateral sclerosis. We previously reported that reduced palmitoylation leads to ER-retention of APP. Here, we expand on these findings by showing that most palAPP is partitioned to MAMs. We and others have previously shown that the loss or inhibition of the MAM-resident enzyme acyl-co-enzyme A:ACAT reduces cell surface localization of APP and Aβ generation (Huttunen et al., 2009; Bhattacharyya and Kovacs, 2010; Murphy et al., 2013). More recently, we showed that inhibition of ACAT reduces palAPP in lipid-rafts by ~76%, in vitro (Bhattacharyya et al., 2013). Palmitoylation inhibitors 2-bromopalmitate and cerulenin also reduced palAPP level and Aβ generation in vitro (Bhattacharyya et al., 2013, 2016). Confocal imaging revealed that cerulenin (Cer) treatment reduced APP distribution in axonal MAMs (IP3R3-labeled), but not in somal MAMs in differentiated PC12 and NPCs (Figure S7). Based on these data, we propose that palAPP partitions into axonal MAMs, whereas total or de-palmitoylated APP resides in somal MAMs. Future studies will be needed to explore whether palmitoylation inhibition as well as ACAT inhibition ameliorates β-amyloid pathology by reducing palAPP levels specifically in axonal MAMs.

In this study, we used FAD hNPCs expressing the FAD mutant of APP (APP_Swe/Lon) to examine the role of MAM-association of palAPP in axonal Aβ generation. Lipid raft fractionation, cell surface biotinylation, and ABE palmitoylation assays (Figures 1A, 4A, and S1) have confirmed that the overall cellular localization and palmitoylation of total APP in FAD hNPCs remain similar to those observed in vitro and in vivo (Bhattacharyya et al., 2013, 2016). These observations suggested that the Swedish and the London mutations in APP do not affect APP’s overall cellular localization or palmitoylation. Several reports have shown that Swedish and/or London mutations in APP in non-neuronal (MDCK and COS) and neuronal (SN56) cells altered the endocytosis of APP without altering its cell surface association. The APP_swe mutant has been shown to undergo β-secretase cleavage in secretory vesicles en route to the cell surface, whereas APP_wt undergoes β-secretase cleavage in post-Golgi compartments after endocytosis (Haass et al., 1995). The Swedish and London mutations in APP slowed the rapid transport from cell surface to lysosomes required for APP-degradation (Lorenzen et al., 2010). In a recent study using the iPSC neurons, Kwart et al. (2019) have shown that CRISPR/Cas9-edited cells expressing APP_Swe have promoted enlargement of Rab5-positive endosomes, which is an early hallmark of AD (for review, see Peric and Annaert, 2015). Therefore, it would be interesting in future studies to investigate whether the Swedish/London mutations in APP (APP_Swe/Lon) affect endocytosis of palAPP to lysosomes or alter endosome size in AD.

The degree of dementia in AD is primarily correlated with loss of synapses. Synaptic dysfunction, preceded by reduced synaptic transmission and loss of dendritic spines, is largely driven by neurotoxic Aβ₄₂-oligomers (Cleary et al., 2005; Haass and Selkoe, 2007). Physiological levels of Aβ₄₂-oligomers have also been shown to suppress long-term potentiation (LTP) in hippocampal slices (Mango et al., 2019). In contrast, Aβ in the picomolar range has also been shown to be required for LTP induction (Koppensteiner et al., 2016). Thus, a key question regarding AD pathology is how Aβ is generated in axons and neuronal processes. Previous studies have shown that APP can be transported anterogradely in axons (Koo et al., 1990; Sisodia et al., 1993; Buxbaum et al., 1998), and Aβ can be made in axonal terminals (Cirrito et al., 2005). To date, the cellular and molecular mechanisms by which this critical pool of axonal Aβ is generated have remained unclear. Our microfluidic chamber system showed that Aβ may be released from axons even in the absence of synapses. This observation is in line with several reports showing focally increased toxic Aβ species in axonal swellings of dystrophic neurons in AD (Stokin et al., 2005; Chevalier-Larsen and Holzbaur, 2006; Kanaan et al., 2013). We reason that MAM-upregulation in axons promotes focally increased secretion of Aβ species from regions along the axons. Axonal swelling along axons has been reported at the early stage of AD pathology leading to axonal dysfunction and Aβ deposits surrounding dystrophic neurons in later stage of the disease (Teipel et al., 2007; Cross et al., 2008; Serrano-Pozo et al., 2011). The mechanisms in the early phase leading to axonal damage in AD pathology remain unclear. MAM dysregulation is an early event in AD pathogenesis (for review, see Yu et al., 2021). Axonal BACE1 levels were increased in axonal swellings in an AD mouse model by genetic deletion of an adaptor protein GGA3, which regulates lysosomal degradation of BACE1, leading to increased production of Aβ (Lomoio et al., 2020). Thus, Aβ released from axons (even in the absence of synapses) is likely to impact AD pathogenesis. In summary, we have demonstrated that axonal Aβ generation is specifically modulated by MAMs via stabilization and cell surface trafficking of palAPP, followed by β-secretase cleavage. These data strongly suggest that modulation of MAM-associated palAPP, specifically in axons (e.g., by regulation of S1R), may be considered a therapeutic strategy for ameliorating Aβ-induced neurodegeneration in AD.

STAR★METHODS

Detailed methods are provided in the online version of this paper and include the following:

RESOURCE AVAILABILITY

Lead contact

Further information and requests for resources and reagents should be directed to and will be fulfilled by the Lead Contact, Rudolph E. Tanzi (email: tanzi@helix.mgh.harvard.edu).

Materials availability

All expression vectors used in this study are available upon request.

Data and code availability

All data are available in the manuscript or in the supplementary materials.

EXPERIMENTAL MODEL AND SUBJECT DETAILS

We have used ReN-VM neural progenitor stem cells (hNPCs) constitutively overexpressing human APP containing K670N/M671L (Swedish) and V717I (London) FAD mutations (APP_Swed/Lon) (Choi et al., 2014). FAD-APP (APP_Swe/lon) expressing hNPCs (FAD hNPCs) were generated by transfecting naive hNPCs with IRES-mediated polycistronic lentiviral vectors encoding human APP_Swe/lon with GFP as a reporter for viral infection. Fluorescence-activated cell sorting (FACS) was employed to enrich the population. FAD hNPCs differentiated in 3D matrix were thoroughly characterized as one of the most effective cellular models tha mimicked amyloid pathology of human AD (D’Avanzo et al., 2015; Kim et al., 2015; Kwak et al., 2020). In addition, mouse cortical neurons from C57BL/6 embryonic day 16 (E-16) mice (female) were used to confirm effect of S1R-modulation on axonal and somal MAMs. All experiments involving animals were performed accordance to the guidelines for animal welfare approved by Institutional Animal Care and Use Committee (IACUC) of Harvard University.

METHOD DETAILS

Isolation of lipid rafts

For lipid raft fractionation from FAD hNPCs, cells were plated on thin-layer 3D matrix (1:10 Matrigel containing culture media), routinely used as a cellular model for AD (Kim et al., 2015). Lipid rafts were isolated from 0.5% Lubrol WX lysates of 10-day differentiated FAD hNPCs following published protocol (Bhattacharyya et al., 2013). To isolate ER-lipid rafts or MAMs, we first prepared crude Mitochondria fractions (Annunziata et al., 2013, 2018) prior to sucrose gradient centrifugation to isolate MAMs and non-MAMs. Briefly, 90% confluent cultured differentiated neuronal cells were resuspended in 5-volume homogenization buffer (10mM HEPES, pH 7.4, 0.25 M sucrose, protease inhibitors). The suspension was homogenized by 20-strokes in a glass homogenizer twice. Removing cell debris and nuclei after centrifugation (600 X g for 5min), the supernatant was subjected to sequential centrifugation steps as described before (Wieckowski et al., 2009) to isolate crude mitochondria (containing ER-rafts or MAMs) by discarding plasma membrane, lysosomes, microsomes and the cytosol. The crude mitochondria pellet was suspended in 0.5% Lubrol in 50 mM HEPES pH 7.4 containing 0.15 M NaCl, 5mM EDTA and protease inhibitor prior to sucrose gradient centrifugation as described before (Bhattacharyya et al., 2013). 1ml fractions were gently collected from the top of the gradient before immunoblotting with antibodies against MAM-proteins IP3R3 (anti-IP3R3) or ACAT (anti-ACAT) to determine MAM and non-MAM fractions.

Isolation of MAMs (mitochondria-associated ER membrane)

Purification of ER, mitochondria and MAMs from cultured cells were performed and analyzed as described (Area-Gomez et al., 2009). Briefly, FAD hNPCs were plated over a thin-layer 3-D matrigel matrix as described before (D’Avanzo et al., 2015), prior to differentiation. 10-day differentiated FAD hNPCs were harvested in resuspension buffer (5 mM HEPES, pH 7.4, 250 mM Manitol and 0.5 mM EGTA). Crude ER membranes were isolated after sequential centrifugation. Crude mitochondria containing MAM and Mito were layered on 30% Percol gradient prior to centrifugation at 100,000 g for 30 min. The upper layer containing MAMs were diluted and further centrifuged at 100,000 g for 1 hr to isolate pure MAMs. Purity of MAMs in the MAM-fractions was measured by immunoblotting the fractions with antibody against MAM-resident IP3R3 antibody (anti-IP3R3) as was done before (Area-Gomez et al., 2009). Probing the fractions with antibody against the mitochondrial marker cytochrome C (anti-Cyto C) showed little or no distribution of cytochrome C (Cyto C) in the MAM fractions, confirming purity of the isolated MAMs. Anti-Cyto C, however, detected Cyto C in both crude ER and mitochondria fractions.

We isolated MAMs from mouse brains following published methods (Wieckowski et al., 2009). We collected brains from non-transgenic (non-Tg) mice (C57BL/6J background). All animal handling and tissue harvesting were performed in accordance with the guidelines of Massachusetts General Hospital Animal Care Use Committee. Mice used in this study were of either sex. Three-month old mice were sacrificed, and brains were immediately removed and stored at −80°C. Briefly, mouse brains were homogenized in five volumes of homogenization buffer prior to fractionation and Percol density gradient centrifugation to isolate pure MAMs. Isolation of pure MAMs were confirmed by determining the presence of MAM-proteins such as IP3R3 and ACAT1, and absence of mitochondrial Cyto C.

Metabolic labeling with Alkyl-C16

Briefly, cells were metabolically labeled with a chemical palmitic acid probe, alkylene palmitic acid (Alkyl-C16; Invitrogen) as described previously (Bhattacharyya et al., 2013). Six hours after labeling, cells were subjected to sucrose-density gradient centrifugation to collect lipid rafts and non-rafts. APP was pulled-down from the fractions using anti-APP C-terminal antibody (C66) prior to addition of a biorthogonal alkyne-labeled fluorescent chromophore, tetramethylrhodamine (alkyne-TAMRA; Invitrogen) via “click chemistry.” The samples were then probed with an anti-TAMRA antibody (Invitrogen).

ABE palmitoylation assay

This assay is based on previously described three-step chemical method where free unmodified cysteine thiols are blocked with NEM (N-ethyl maleimide) followed by palmitoylation thioesters cleavage by hydroxylamine (NH₂OH); finally, loading thiol-specific biotinylating reagent (HPDP-biotin in our experiments) to the newly exposed cysteinyl thiols. Biotinylated proteins are then affinity purified with streptavidin–agarose beads and probed for the protein of interest (Bhattacharyya et al., 2013). The assay was performed on total cell lysates extracted from differentiated FAD NPCs. In some cases, cells were treated with sigma1 receptor (S1R) agonist PRE-084 or antagonist NE-100 before ABE assay. ABE assays were also performed on cells undergoing cycloheximide (CHX) chase for 0–18h. For lipid raft, MAMs and non-raft palmitoylation studies, protein extracts from lipid rafts, MAMs and non-rafts were subjected to ABE assay according to a method reported previously (Bhattacharyya et al., 2013). Briefly, lipid raft and MAM fractions were extracted with 60 mM n-Octylglucoside (Sigma), and non-raft fractions were extracted with 1% Triton X-100. The protein extracts were precipitated using chloroform/methanol before ABE assay.

Badrilla palmitoylation assay

Palmitoylation assay was performed using CAPTUREome S-Palmitoylated Protein assay kit from Badrilla, UK, following the manufacturer’s protocol. This assay used the technique employed in ABE assay with some exceptions. It replaced the biotinylating step of ABE assay with direct conjugation of thioester-cleaved palmitoylated proteins with CAPTUREome resin containing thiol-reactive groups. This resulted in the covalent capture of proteins on a resin, permitting more stringent treatment to harvest S-acylated proteins with ease and high purity. Most importantly, due to the absence of the biotinylation step in ABE assay, this assay was able to not only capture palmitoylated proteins but also was able to capture cell surface biotinylated palmitoylated proteins. In addition, we were also able to isolate non-palmitoylated proteins in the unbound fractions from CAPTUREome precipitates. Briefly, cells were lysed in presence of a blocking reagent before extraction. S-palmitate groups were removed via cleavage of thioester bonds, by incubating the cell extracts with the thioester cleavage reagent. This resulted in the creation of free thiol groups on proteins where the palmitic acid group used to be. As a control, the cell extracts were incubated with acyl prevention reagent. After incubation, CAPTUREome captures resin containing reactive thiol group was added to both control and samples containing thioester cleavage reagent. After washing three times, pulled-down samples were immunoblotted with appropriate antibodies to test palmitoylation. Proteins captured by CATUREome from samples treated with thioester reagent represented palmitoylated proteins. Unbound fractions were subjected to two sequential CAPTUREome precipitation to remove all palmitoylated protein, while the proteins in the final unbound fractions contained non-palmitoylated proteins. In some cases, the assay was performed on MAMs and non-MAMs isolated from sucrose gradient centrifugation of the ER-mitochondria extract of FAD hNPCs.

Cell surface biotinylation assay of palmitoylated proteins

This assay was a combination of cell surface biotinylation and Badrilla palmitoylation assays. Covalent attachment of cell surface proteins with cell impermeable thiol-cleavable amine-reactive biotinylation reagent (EZ-link sulfo-NHS-S-S-Biotin, Thermo Scientific) was a technique to monitor trafficking of proteins to the cell surface. Badrilla assay of cell surface biotinylated proteins were designed to detect palmitoylated proteins at the cell surface. The combined assay was performed in two steps. First, cultured cells were subjected to cell surface biotinylation assay following published protocol (Sannerud et al., 2011). Briefly, cultured cells were placed on ice to restrict endocytosis prior to labeling with 0.5 mg/ml biotin reagent for 60 min. Cells were then washed and the unbound biotin reagent was quenched with 100 μM lysine for 5 min. After three-washes with PBS, cells were lysed with extraction buffer containing 10mM Tris (pH 7.4), 2mM EDTA, 150mM NaCl, 1% Triton X-100, 0.5% sodium deoxycholate, 0.2% SDS, and protease inhibitors. 750 μg of cell extract was subjected to pull-down with Neutravidin beads to purify biotinylated cell surface proteins. Proteins were extracted from the beads either by incubating with SDS sample buffer containing 0.1% β-mecrcaptoethanol (βME) or by extracting with 25mM TCEP solution in 50mM Tris-buffer, pH 8.5. βME-extracted samples were probed with appropriate antibodies to detect cell surface association of APP (C66 antibody), transferrin receptor (anti-Tfr antibody) or β₂ adrenergic receptor (anti-β₂AR antibody). Next, the TCEP purified cell surface proteins were subjected to Badrilla palmitoylation assay to detect palmitoylated proteins at the cell surface. CAPTUREome-bound proteins constituted palmitoylated cell surface proteins. APP and β₂AR were detected in the CAPTUREome bound fractions showing cell surface palmitoylated APP and β₂AR at the cell surface. We used transferrin receptor as a positive control for cell surface biotinylation assay, and a negative control for cell surface Badrilla assay because Tfr was not palmitoylated. CAPTUREome beads did not pull-down Tfr confirming the efficiency of the combined assay to detect palmitoylated proteins at the cell surface. The combined assay was performed on cells untreated (veh) or pre-treated with S1R-agonist (PRE- 084) or antagonist (NE-100).

Cycloheximide chase assay

Cultured cells were incubated with 100 μM cycloheximide (CHX, Sigma) for various times, as described before (Dai et al., 2013). Cell lysates were prepared for ABE or Badrilla Palmitoylation assays prior to western blot analysis. In some cases, CHX-chase were performed in cells treated with 10 μM PRE or NE.

Confocal microscopy

Differentiated hNPCs, FAD hNPCs and primary neurons were grown on coverslips or inside microfluidic chamber slides prior to labeling with antibodies following methods described before (Bhattacharyya et al., 2016). Briefly, cells were fixed in 4% paraformaldehyde (PFA) prior to labeling with antibodies against S1R, tau or pNFH followed by appropriate fluorescence conjugated secondary antibodies. Fluorescence microscopy was performed under Nikon confocal microscope using 40X objective. Images were processed by ImageJ software.

To determine MAM-assemblies, cells were incubated with membrane-permeable ER and mitochondria probes (Cell Light ER-GFP and Cell Light Mito-RFP, respectively) for 16h prior to confocal microscopy. Cell Light reagents (Thermo Fisher) containing 1 X 10⁸ particles/ml were used. We used 25 particles per cell (PPC) of each probe. Precisely, we differentiated 40,000 cells and probed with 10 μL of Cell Light ER-GFP or Cell Light Mito-RFP. Confocal images were processed by Photoshop to eliminate GFP and RFP signals by minimizing the saturations to zero without altering the GFP and RFP overlapped regions. We highlighted the overlapped regions white that represented the ER-mitochondria contact sites or MAMs. Next, we measured the number of ER-Mito sites or MAMs using ImageJ software. Specifically, we used “analyze particle and image calculator” option to measure the number of areas > 10 nm because the sizes of MAMs ranged between 5 and 20 nm (Szymański et al., 2017). We counted the number of MAMs in cell bodies, as well as in the neuronal processes. The mean of the MAM-counts was used for quantification of MAMs in cell bodies and in neuronal processes.

Bio-immunofluorescence (BiFC) assay

Bio-immunofluorescence of split GFP constructs (BiFC) assay was performed as done before (Bhattacharyya et al., 2016). We have used expression plasmids encoding split-GFP (spGFP) BiFC probes for ER and mitochondria. Specifically, we transiently transfected (Lipofectamine 2000, Invitrogen) plasmids encoding spGFP(1–10)-tagged ER protein Ubc6 [ER-GFP(1–10)] and a spGFP11-tagged mitochondrial protein Tomm70 (Mito-GFP11) (Yang et al., 2018). The plasmids are generous gifts from Dr. Chao Tong, Professor, Life Sciences Institute, Zhejiang University, China. Co-expression of these probes hNPCs resulted in punctate GFP-signals confirming close contacts between these 2 organelles, suggesting detection of MAMs. Cells were then treated with S1R-antagonist NE-100 for 16 h before fixing in 4% paraformaldehyde in 1x PBS at room temperature (RT) for 30 min. Fluorescence microscopy was performed under Nikon confocal microscope using 60X objective. The fluorescent puncta in axons and in cell bodies were counted and plotted to determine the effect of NE-100 on the number of MAMs per axon or per cell body. We assessed the expression of ER- and Mito-probes by labeling with anti-GFP antibody. BiFC puncta in the cell bodies or in axons of 20 untreated and 20 treated cells were counted for analysis.

Transmission electron microscopy

Transmission Electron Microscopy of cells containing control non-targeting siRNA (si-non) or siRNA against S1R (si-S1R) were performed at the Harvard Medical School Electron Microscopy Core facility. Briefly, cells were fixed in routine fixative containing 2.5% Glutaraldehyde 2.5% Paraformaldehyde in 0.1 M sodium cacodylate buffer (pH 7.4) overnight at 4°C. Cells were then washed in 0.1M Sodium cacodylate buffer pH 7.4, post fixed for 30 min in 1% Osmium tetroxide (OsO4)/1.5% Potassium ferrocyanide (KFeCN6), washed in water 3x and incubated in 1% aqueous uranyl acetate for 30min followed by 2 washes in water and subsequent dehydration in grades of alcohol (5min each; 50%, 70%, 95%, 2× 100%). Cells were removed from the dish in propylene oxide, pelleted at 3000 rpm for 3min and infiltrated for 2hrs to ON in a 1:1 mixture of propylene oxide and TAAB Epon (Marivac Canada Inc. St. Laurent, Canada). The samples subsequently embedded in TAAB Epon and polymerized at 60°C for 48 hr. Ultrathin sections (about 60nm) were cut on a Reichert Ultra cut-S microtome, picked up on to copper grids stained with lead citrate and examined in a JEOL 1200EX Transmission electron microscope or a TecnaiG² Spirit BioTWIN and images were recorded with an AMT 2k CCD camera. We manually counted the ER-mitochondria (ER-Mito) contact sites > 10 nm per mitochondria per frame as described before (Bernard-Marissal et al., 2015). We counted more than 25 frames. Average of the counts were used to quantitate the fold decrease of ER-Mito contact sites or MAMs in S1R-silenced (si-S1R) cells compared to control (si-non) cells.

Microfluidic two-chamber device design and fabrication

The microfluidic chips composed of two distinct chambers connected through microgroove arrays (5 × 8 × 450 mm in height, width and length) were fabricated using standard soft photolithography techniques (Qin et al., 2010). Briefly, the master for preparing the PDMS array was microfabricated with SU-8 photoresist (MicroChem Co.) on a silicon wafer. The first layer (5 μm depth) contained 195 microgrooves and the second layer (100 μm depth) of SU-8 contained the compartments were patterned by photolithography using high-resolution chromium masks. The PDMS array was then molded by casting the liquid prepolymer composed of a mixture of 10:1 silicon elastomer and a curing agent (Sylgard 184). The mixture was cured at 75 °C for 4 h, and the PDMS mold was peeled from the silicon wafer. The inlet and outlet wells were punched using 3 mm diameter puncher. The molded PDMS block was then bonded to 6 well glass bottom plates (−1.5H-N, Cellvis) using oxygen-plasma treatment. The microfluidic devices were then oxygen-plasma treated and immediately coated with 1:100 (Matrigel: Expansion medium) and incubated at 37 °C and 5% CO₂ atmosphere for at least two hours before cell culture.

Axotomy

To compare MAM levels between bulk neurons and axons, we performed axotomy by physically severing the axonal chambers from the somal chambers of the Microfluidic Two-Chamber Devices containing FAD hNPCs. The severed chambers were carefully collected in two separate conical tubes labeled as bulk neurons and axons. Protein extraction buffer was added to each tube to extract proteins from bulk neurons or axons. Equal amounts of the extracts were subjected to Western Blot analysis and probed with anti-IP3R3, anti-ACAT1 and anti-VDAC1 antibodies to determine MAM levels. To assess the purity of the axons, the extracts were also probed with antibody against nuclear envelope protein, Lamin B1 (anti-Lamin B1), present only in the nucleus of bulk neurons but not in axons. Axonal extracts showing no detectable level of Lamin B1 were considered to be pure axons.

Compartmentalized microfluidic chambers

To measure somal and axonal Aβ₄₀ we exclusively used the Xona microfluidic devices because these were able to generate fluidic separation between somal and axonal chambers. We created a thin-layer 3D culture of FAD hNPCs in these microfluidic devices to minimize contamination of soma in axonal chambers. Briefly, ~30,000 FAD hNPCs were mixed with ice-cold cell differentiation media containing BD Matrigel stock solution (1:10 dilution ratio). The mixture was immediately added to the somal chamber following the manufacturer’s instruction. The chamber was transferred to 37°C for 10 min to form thin-layer (100–300 μm) 3D gels. The axonal chambers were loaded with differentiation media. Cells were allowed to differentiate for 10-days with changes of media every 3-days. We maintained a fluidic separation of 60 μL between the somal and the axonal chambers. Somal chambers contained 180 μL while the axonal chamber contained 120 μL conditioned media (CM). On the tenth day, media was exchanged with fresh media (veh) or with media containing 10 μM PRE-084 (PRE) or 10 μM NE-100 (NE). CM were collected after 16h prior to immunoblotting or Aβ-ELISA assay to determine Aβ (Aβ₄₀ and Aβ₄₂)-release. The neurons were subjected to indirect immunofluorescence analysis using anti-p-NFH or anti-tau antibodies to label axons in both axonal and somal chambers followed by confocal microscopy. Staining the nucleus with Hoechst dye determined the total number of cell bodies in somal and axonal chambers.

ELISA and western blot

Aβ (Aβ₄₀ and Aβ₄₂) species were measured from conditioned media of untreated or treated (PRE, CE, or BACEi-IV) Ren-GA neurons using commercially available ELISA kits from WAKO, as described before (Bhattacharyya et al., 2013, 2016). Conditioned media from the somal and axonal chambers from cells differentiated in microfluidic chambers were subjected to WAKO ELISA to measure Aβ (Aβ₄₀ and Aβ₄₂) from cell bodies and axons, respectively. Aβ₄₂ levels in the axonal chambers were below detectable limits, thus only axonal-Aβ₄₀ levels were determined. We determined the number of Aβ₄₀ molecules per axons or per cell body following methods described by Niederst et al. (2015). Briefly, we first calculated the amounts of total Aβ₄₀ release from each axonal and each somal chambers by multiplying the concentration (in μM) of Aβ₄₀ with total volume of conditioned media of the chambers (120 μL for axonal chamber and 150 μL for somal chamber) divided by the number of tau-positive axons per axonal chamber and Hoechst-positive soma per somal chamber. Next, we calculated the number of Aβ molecule per soma by multiplying the Aβ₄₀ concentrations with Avogadro’s number. To calculate the number of Aβ molecules per axon, first, we multiplied Aβ₄₀ concentrations times Avogadro’s number. Next we counted the small number of Hoechst-positive cells in the axonal chambers and calculated the number of Aβ molecules generated from these cells as somal Aβ. This number was then subtracted from the number of Aβ molecules per axon calculated before to obtain the number of Aβ molecule per axon.

Protein measurement

Protein concentrations were determined by Bio Rad BCA assay. For western blot analysis samples from palmitoylation assays or from sucrose or Percol fractionation were prepared in sample buffers containing β-mercaptoethanol (βME) for denaturation. Samples were resolved on a 4%–12% gradient Bis-Tris gels (Invitrogen) prior to immunoblotting analysis using appropriate antibodies. The blots were visualized by a Li-Cor Odyssey imaging system. Band intensities were measured by ImageStudio software.

siRNA mediated knock down

A SMART pool siRNA (Dharmacon, Inc.) targeting four sequences within the human sigma1 receptor (S1R) transcript (CUAUUAAUAAAGAUUUGUU, CGAGUAGUGCUGCUCUUC, GGGAUAUCCAUGCUUAUGU and GUUCUAGAGUUAAGGAUGG), based on a previous report (Amer, McKeown et al., 2013). A non-targeting (si-non) siRNA was obtained as control siRNA. To knock down S1R, cells were introduced with 100nM SMART pool siRNA against S1R (si-S1R) or with the control (si-non) via electroporation technique using Amexa Mouse Neuron Nucleofector kit (Lonza) according to the manufacturer protocol. For FAD hNPCs, we used 5 X 10⁶ cells to introduce si-S1R and si-non.

Cell viability assay

We used CytoTox-ONE Assay (Promega) to measure cell viability. The CytoTox-ONE Assay is a rapid, fluorescent measure of the release of lactate dehydrogenase (LDH) from cells with a damaged membrane. We measured cell viability by assessing the LDH release from cells undergoing siRNA-mediated silencing, following the manufacturer’s instruction. Briefly, fresh conditioned media (CM) were collected and mixed with CytoTox-ONE Reagent in a 96 well plate. The mixtures were incubated at 22°C for 10 minutes prior to adding Cyto-Tox-ONE Stop solution. LDH-release was measured by recording fluorescence (Ex = 560 nm and Em = 590 nm). Experiments were performed in triplicates. The percent cell viability was calculated by using the average fluorescence values from experimental, maximum LDH release, and culture medium background for each experiment.

QUANTIFICATION AND STATISTICAL ANALYSIS

All statistical analyses were performed using Microsoft Excel or GraphPad Prism v.6 software (Graphpad). Exact values for experimental numbers and p values are reported in the figures and corresponding figure legends. Bars and error bars on the graphs represent mean values and SEM for multiple independent experiments as specified in each legend. Unless otherwise mentioned, statistical significances were determined by unpaired Student’s t test for two groups or one-way ANOVA with Tukey’s multiple comparisons test for multiple groups. All statistical analyses were performed using a two-tailed Student’s t test. Data in graphs are expressed as mean values SEM p < 0.01 were considered significant.

Supplementary Material

NIHMS1706347-supplement-1.pdf^{(4.2MB, pdf)}

KEY RESOURCES TABLE

REAGENT or RESOURCE	SOURCE	IDENTIFIER
Antibodies

22C11 (anti-APP N terminus)	Sigma-Millipore	Cat#MAB348, RRID: AB_2056583
anti-s APP beta	IBL International	Cat#10321, RRID: AB_1630822
anti-Sigma Receptor Antibody (B-5) (Mouse)	Santa Cruz	Cat#sc-137075, RRID: AB_2285870
Purified Mouse anti-IP3R3	BD Transduction Laboratories	Cat#610312, RRID: AB_397704
anti-SOAT-1/ACAT-1 Polyclonal (Rabbit)	Cayman Chemical	Cat#100028, RRID: AB_327803
Purified Mouse Anti-Flotillin-1	BD Transduction Laboratories	Cat#610820, RRID: AB_398140
anti-MAP2 (Rabbit)	Cell Signaling Technologies	Cat#4542, RRID: AB_10693782
Anti-Grp75 antibody	Abcam	Cat# ab129201, RRID:AB_11141817
Anti-Neurofilament heavy polypeptide antibody	Abcam	Cat#ab8135, RRID: AB_304560
anti-Tau	Cell Signaling Technologies	Cat#A0024, RRID: AB_10013724
anti-GAPDH	Life Technologies	Cat#MA5-15738-D680, RRID: AB_2537657
anti- beta2 Adrenergic Receptor (Rabbit)	Abcam	Cat#ab176490, RRID: AB_2861265
anti-Transferrin Receptor (Rabbit)	Abcam	Cat#ab84036, RRID: AB_10673794
Anti-GRP94 (C-terminal) (Rabbit)	Sigma	Cat#G4420, RRID: AB_477017
Anti-VDAC1 / Porin antibody [EPR10852(B)] (Rabbit)	Abcam	Cat#ab154856, RRID: AB_2687466
anti-Laminb1	Abcam	Cat#ab65986, RRID: AB_1140888
Alexa Fluor 488 and 568 secondary antibody	Life Technologies	Cat#A32723, RRID: AB_2633275; A32731, RRID: AB_2633280; A-11011, RRID: AB_143157; A-11004, RRID: AB_2534072; A-11041, RRID: AB_2534098
HRP conjugate secondary antibody	Life Technologies	Cat#G-21040, RRID: AB_2536527; G-21234, RRID: AB_2536530
Anti-Cytochrome C antibody [7H8.2C12] (mouse)	Abcam	Cat#ab13575, RRID: AB_300470
Anti-Tamara	Invitrogen, Thermo Fisher	Cat#MA1-041, RRID: AB_2536728
Anti-GFP (chicken)	Abcam	Cat#ab13970, RRID: AB_300798
C66 (Anti-APP C terminus)	In-house	https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3718372/
DMEM/F12 with L-glutamine	GIBCO/Thermo fisher	Cat#11320-033
B-27 Supplement (50X), serum free	GIBCO/Thermo fisher	Cat#17504044
bFGF	R&D System	Cat#233-FB
EGF	Sigma Aldrich	Cat#92090408
Penicillin/Streptomycin/Amphotericin B	Lonza	Cat#17-745E
StemPro Accutase	GIBCO	Cat#2023-01-30
DMEM 4.5 g/L Glucose w/o L-Gln w/Phenol Red	Lonza	Cat#BE12-614F
Alkylene Palmitic acid (Alkyl C-16)	Invitrogen	Cat#C10265
Click-it reagents, alkyne-Tetramethylrhodamine	Invitrogen	Cat#C10269, T10183
PRE-+B34:C48084 hydrochoride	Tocris Bioscience	Cas#138847-85-5 Batch# 2A/222270
NE-100	Sigma Aldrich	Cat#SML0631
Cerulenin	Sigma Aldrich	Cat#P2607
EZ-Link Sulfo-NHS-SS-Biotin	Thermo Scientific	Cat#21335
BACE1 inhibitor IV	(BACE1-IV) Calbiochem	Cat#565788
CellLight ER-GFP BacMam 2.0	Thermo fisher	Cat#C10590
CellLigh Mitochondria-RFP BacMam 2.0	Thermo fisher	Cat#C10601
Hoechst 3342	Thermo fisher	Cat#H3570
Lubrol	Calbiochem	Cat#205528
Percol	Sigma Aldrich	Cat#P4937
Triton X-100	Sigma Aldrich	Cat#T8787
n-Octylglucoside	Sigma Aldrich	Cat#10634425001
Cyclohexamide	Sigma Aldrich	Cat#C7698
Lipofectamine 2000 Transfection Reagent	Invitrogen	Cat#11668019
NuPAGE 4-12% Bis-Tris gel	Invitrogen	Cat#NP0321BOX
Matrigel Basement Membrane Matrix	Corning	Cat#356234

Critical commercial assays

CAPTUREome S-Palmitoylation Protein mini kit	Badrilla	Cat#K010-310
Human/Rat beta amyloid (40) ELISA Kit	Wako	Cat#294-62501
Human/Rat beta amyloid (42) ELISA Kit	Wako	Cat#290-62601
Amexa mouse Neuron Ncleofactor kit	Lonza	Cat#VPG-1001
Cyto Tox-One assay	Promega	Cat#G3582

Software

ImageJ Software	ImageJ 1.53a	N/A
Photoshop	Adobe Photoshop CC 20.0.10	N/A
Graphpad Prism	Prisom 9, version 9.0.2	N/A
MS Excel	Microsoft Excel, version 16.30	N/A

Oligonucleotides

split GFP constructs (provide sequence for both ER and mito)	Dr. Chao Tong, Professor, Life Sciences Institute, Zhejiang University, China	N/A
SMART pool siRNA against sigma-1 receprot (S1R)	Dharmacon, Inc.	N/A

Other/devices

XonaChip 450um	Xona Microfluidics	Cat#XC450
XonaChip 150um	Xona Microfluidics	Cat#XC150
Microfluidic Chips (5 × 8 × 450 um) (H x W x L)	In-house (designed by M.J.)	N/A

Open in a new tab

Highlights.

Palmitoylated APP (palAPP) is enriched in mitochondrial-associated ER membranes (MAMs)
Upregulation of MAM increases cell surface palAPP levels and axonal Aβ generation
Downregulation of MAM attenuates cell surface palAPP levels and axonal Aβ generation

ACKNOWLEDGMENTS

We thank Dr. Doo Y. Kim, Associate Professor, Massachusetts General Hospital for providing us with the 3D neural cell model of AD (FAD hNPCs); Dr. Chen Meng of Massachusetts General Hospital for help in the preparation of primary neurons; Maria Ericsson of Harvard Medical School Electron Microscopy Core facility and Diane E. Capen from MGH Center for System Biology for their help with electron microscopy; Dr. Chao Tong, Professor, Zhejiang University, China, for providing the split-GFP BiFC plasmids; and Nick Waterhouse, summer intern, for initial testing of CellLight probes in non-neuronal cells. This study was funded by grants from the NIH (5R01NS045860-19) and the Cure Alzheimer’s Fund.

Footnotes

DECLARATION OF INTERESTS

R.E.T. and D.M.K. hold intellectual property and patents on the topic of acyl-CoA:cholesterol acyltransferase inhibitors (patent no. US20050118226A1) and methods and compositions relating to modulating amyloid precursor protein cleavage (patent no. US20050170437A1).

INCLUSION AND DIVERSITY

We worked to ensure sex balance in the selection of non-human subjects. We worked to ensure diversity in experimental samples through the selection of the cell lines. We worked to ensure diversity in experimental samples through the selection of the genomic datasets.

SUPPLEMENTAL INFORMATION

Supplemental information can be found online at https://doi.org/10.1016/j.celrep.2021.109134.

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Supplementary Materials

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Data Availability Statement

All data are available in the manuscript or in the supplementary materials.

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PERMALINK

Axonal generation of amyloid-β from palmitoylated APP in mitochondria-associated endoplasmic reticulum membranes

Raja Bhattacharyya

Sophia E Black

Madhura S Lotlikar

Rebecca H Fenn

Mehdi Jorfi

Dora M Kovacs

Rudolph E Tanzi

SUMMARY

In brief

Graphical Abstract

INTRODUCTION

RESULTS

palAPP is predominantly localized to MAMs in human neuronal cells and in mouse brains

Figure 1. palAPP is predominantly distributed in the ER-rafts or MAMs in differentiated human stem cell-derived neural progenitor FAD-NPC neurons and in mouse brain.

Figure 2. Silencing or inactivation of MAM-resident sigma-1 receptor (S1R) in FAD hNPCs decreases MAM levels and reduces palAPP level and sAPPβ release.

Silencing of MAM-resident sigma 1 receptor (S1R) reduces MAM levels in FAD NPCs

S1R-agonist PRE-084 and S1R-antagonist NE-100 regulate MAM levels in FAD hNPCs

Modulation of MAMs via S1R stabilizes palAPP in FAD-NPC neurons

Figure 3. Activation or inactivation of S1R specifically increases or decreases palAPP stability, respectively.

MAM assembly regulates levels of cell surface-associated palAPP but not total APP

Figure 4. Activation or inactivation of S1R specifically increases or decreases cell surface association of palAPP, respectively.

Regulation of MAM-assembly in FAD hNPCs via S1R modulates β-secretase cleavage of palAPP

Figure 5. Silencing of S1R and modulation of S1R-activity regulates β-secretase cleavage of APP and palAPP, respectively.

Figure 7. Activation of S1R increases axonal Aβ40 release from FAD hNPCs in 3-D neural cell cultures.

Regulation of S1R-activity specifically modulates MAMs in neuronal processes

Figure 6. Regulation of S1R-activity modulates MAMs in neuronal processes or axons without affecting MAMs in cell bodies or bulk neurons.

Modulation of MAMs in neuronal processes/axons regulates axonal Aβ generation from FAD hNPCs

DISCUSSION

STAR★METHODS

RESOURCE AVAILABILITY

Lead contact

Materials availability

Data and code availability

EXPERIMENTAL MODEL AND SUBJECT DETAILS

METHOD DETAILS

Isolation of lipid rafts

Isolation of MAMs (mitochondria-associated ER membrane)

Metabolic labeling with Alkyl-C16

ABE palmitoylation assay

Badrilla palmitoylation assay

Cell surface biotinylation assay of palmitoylated proteins

Cycloheximide chase assay

Confocal microscopy

Bio-immunofluorescence (BiFC) assay

Transmission electron microscopy

Microfluidic two-chamber device design and fabrication

Axotomy

Compartmentalized microfluidic chambers

ELISA and western blot

Protein measurement

siRNA mediated knock down

Cell viability assay

QUANTIFICATION AND STATISTICAL ANALYSIS

Supplementary Material

Highlights.

ACKNOWLEDGMENTS

Footnotes

REFERENCES

Associated Data

Supplementary Materials

Data Availability Statement

ACTIONS

PERMALINK

RESOURCES

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Links to NCBI Databases

Figure 7. Activation of S1R increases axonal Aβ₄₀ release from FAD hNPCs in 3-D neural cell cultures.