Detection of a Novel Intraneuronal Pool of Insoluble Amyloid beta  Protein that Accumulates with Time in Culture

Daniel M. Skovronsky, Robert W. Doms, and Virginia M.-Y. Lee

Center for Neurodegenerative Disease Research, Department of Pathology and Laboratory Medicine, University of Pennsylvania School of Medicine, Philadelphia, Pennsylvania 19104

    Abstract
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

The amyloid-beta peptide (Abeta ) is produced at several sites within cultured human NT2N neurons with Abeta 1-42 specifically generated in the endoplasmic reticulum/intermediate compartment. Since Abeta is found as insoluble deposits in senile plaques of the AD brain, and the Abeta peptide can polymerize into insoluble fibrils in vitro, we examined the possibility that Abeta 1-40, and particularly the more highly amyloidogenic Abeta 1-42, accumulate in an insoluble pool within NT2N neurons. Remarkably, we found that formic acid extraction of the NT2N cells solubilized a pool of previously undetectable Abeta that accounted for over half of the total intracellular Abeta . Abeta 1-42 was more abundant than Abeta 1-40 in this pool, and most of the insoluble Abeta 1-42 was generated in the endoplasmic reticulum/intermediate compartment pathway. High levels of insoluble Abeta were also detected in several nonneuronal cell lines engineered to overexpress the amyloid-beta precursor protein. This insoluble intracellular pool of Abeta was exceptionally stable, and accumulated in NT2N neurons in a time-dependent manner, increasing 12-fold over a 7-wk period in culture. These novel findings suggest that Abeta amyloidogenesis may be initiated within living neurons rather than in the extracellular space. Thus, the data presented here require a reexamination of the prevailing view about the pathogenesis of Abeta deposition in the AD brain.

    Introduction
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References

ALZHEIMER'S disease (AD)1 is characterized by accumulation of fibrillar amyloid-beta peptides (Abeta ) in senile plaques. That the accumulation of Abeta is essential for the pathogenesis of AD is supported by genetic studies showing that mutations in the amyloid-beta precursor protein (APP) (which gives rise to Abeta through proteolytic processing) are linked to a subset of familial AD (FAD) cases with autosomal penetrance, and alter Abeta production (reviewed in Selkoe, 1997). For example, the double mutation found in a Swedish FAD kindred leads to overproduction of Abeta , while other mutations alter the relative levels of the two major forms of Abeta , resulting in an increased Abeta 1-42/1-40 ratio (Citron et al., 1992; Scheuner et al., 1996). Previous studies have shown that Abeta 1-42 is more insoluble than the more abundant Abeta 1-40, and that it is the most prevalent Abeta species found in senile plaques (Iwatsubo et al., 1994). Other FAD mutations that account for the majority of early-onset FAD cases have been linked to the Presenilin 1 (PS1) and Presenilin 2 (PS2) genes (Levy-Lahad et al., 1995; Sherrington et al., 1995). Mutations in these genes, like some of those in the APP gene, also increase the Abeta 1-42/1-40 ratio (Borchelt et al., 1996; Duff et al., 1996; Scheuner et al., 1996).

Since genetic studies have established a role for Abeta in the pathogenesis of AD, it is essential to understand how Abeta is produced from APP. For example, it has been shown that APP is cleaved by beta -secretase(s) to generate the NH2 terminus of Abeta , and by gamma -secretase(s) to generate the COOH terminus of Abeta (Haass et al., 1992; Shoji et al., 1992). These cleavages may occur in a variety of subcellular locations, including the endoplasmic reticulum/intermediate compartment (ER/IC; Chyung et al., 1997; Cook et al., 1997; Hartmann et al., 1997; Xu et al., 1997), the trans-Golgi network (TGN; Xu et al., 1997), and the endosomal/lysosomal system (Koo and Squazzo, 1994). Whereas Abeta produced by these pathways may be secreted (as has been shown for TGN-generated Abeta ) or may remain intracellular (as has been shown for Abeta generated by the ER/ IC pathway), the relative roles of intracellular and secreted Abeta in the pathogenesis of AD remain to be determined.

While numerous studies have documented that nonneuronal cells engineered to express APP secrete both Abeta 1-40 and Abeta 1-42, intracellular Abeta is not commonly seen in these cells (Forman et al., 1997; Xu et al., 1997). However, intracellular Abeta can be detected readily in human NT2N neurons after metabolic labeling, and its production precedes that of secreted Abeta (Wertkin et al., 1993; Turner et al., 1996). Analysis of intracellular Abeta by ELISA indicates that intracellular and secreted Abeta are composed of different ratios of Abeta 1-42/1-40, with Abeta 1-40 being more prevalent in secreted material (Turner et al., 1996). In addition to being produced by mechanisms with different time courses, and being composed of different proportions of Abeta 1-40 and Abeta 1-42, intracellular and secreted Abeta can be produced by different pathways in NT2N neurons. Recent studies have shown that Abeta 1-42, but not Abeta 1-40, is produced by an ER/IC pathway, and that this pathway does not contribute to the secreted pool of Abeta (Cook et al., 1997). Finally, secretion of Abeta by NT2N neurons increases with time in culture (Turner et al., 1996). An age-dependent increase in Abeta secretion by neurons in vivo may play a role in the deposition of Abeta into senile plaques in the extracellular space of the brain during normal aging and in AD, as well as in the cortex and hippocampus of transgenic mice that overexpress mutant forms of APP (Games et al., 1995; Hsiao et al., 1996).

In addition to forming insoluble extracellular plaques, Abeta may also accumulate intracellularly in an aggregated insoluble pool. For example, exogenous Abeta 1-42 added to culture medium can be taken up by cells, after which it can be solubilized only by formic acid extraction (Knauer et al., 1992; Yang et al., 1995). Thus, these findings raise the possibility that endogenously produced intracellular Abeta may aggregate within neurons as well. Because formic acid is required to solubilize Abeta from senile plaques, we sought to detect the presence of insoluble Abeta within NT2N neurons and other cell lines by formic acid extraction, and found that a significant fraction of the total intracellular Abeta , particularly Abeta 1-42, was retained as an insoluble pool within these cells. Further, this insoluble pool of Abeta increased 12-fold in postmitotic NT2N neurons over a period of 7 wk in culture. Since the prevailing view of amyloidogenesis in AD is that plaque formation is initiated in the extracellular space by secreted Abeta , our findings challenge this assumption by implicating the intracellular compartment as a site where Abeta may accumulate in an insoluble form.

    Materials and Methods
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

Cell Culture

NT2 cells derived from a human embryonal carcinoma cell line (Ntera2/ cl.D1) were grown and passaged as described previously (Pleasure et al., 1992; Pleasure and Lee, 1993). Cells were differentiated by two weekly retinoic acid treatments (10 µM) for 5 wk, and were replated (replate 2 cells) in the presence of mitotic inhibitors to yield nearly pure NT2N neurons (Pleasure et al., 1992). To obtain 99% pure neurons (replate 3 cells), replate 2 cells were removed enzymatically and mechanically, and were replated in 10-cm dishes (Pleasure et al., 1992). Cultures of Replate 2 or Replate 3 NT2N cells were used for experiments when they were 3-4 wk old unless otherwise indicated. CHO Pro5 cells were grown and passaged three times per week in Alpha-MEM (Life Technologies, Inc., Gaithersburg, MD) containing 10% FBS and penicillin/streptomycin. Baby hamster kidney (BHK-21) cells were grown and passaged three times per week in Glascow MEM (Life Technologies, Inc.) supplemented with 10% tryptose phosphate, 5% FBS, and 0.02 M Hepes. CHO-695 cells were obtained from Dr. S.S. Sisodia, and were grown and passaged as described above for CHO Pro5 cells with the addition of 0.2 mg/ml of G418 to the culture medium.

Preparation of Semliki Forest Virus and Infection of Cultured Cells

Semliki Forest Virus (SFV) expressing wild-type APP695 (SFV-APPwt) or an APP mutant in which the third and fourth amino acids from the carboxyl terminus of APP have been changed to lysines (SFV-APPDelta KK) were prepared and titered as previously described (Chyung et al., 1997; Cook et al., 1997). CHO-Pro5, BHK-21, NT2, and NT2N cells were infected in serum-free medium at a multiplicity of infection of ~10. After 1 h, complete growth medium was replaced and infection was allowed to proceed for 12 h.

Metabolic Labeling, Immunoprecipitation, and Gel Electrophoresis

Cultured NT2N cells were methionine-deprived by incubation in methionine-free DMEM (Life Technologies, Inc.) for 30 min before adding [S35]methionine (500 µCi/ml in methione-free DMEM + 5% dialyzed FBS; DuPont-NEN, Boston, MA) for a 12-h labeling period. Cells were washed twice in PBS and lysed in 600 µl RIPA buffer (0.5% sodium deoxycholate, 0.1% SDS, 1% NP40, 5 mM EDTA in TBS, pH 8.0) with a cocktail of protease inhibitors (1 µg/ml each of Pepstatin A, Leupeptin, TPCK, TLCK, STI, and 0.5 mM PMSF). After brief sonication, cell lysates were centrifuged at 40,000 g for 20 min at 4°C, and the supernatant was subjected to immunoprecipitation with 6E10 (a monoclonal antibody specific for Abeta 1-17; Kim et al., 1988) as previously described (Turner et al., 1996). The remaining pellets were resuspended in 100 µl 70% formic acid and sonicated until clear. For direct extraction into formic acid, cells were scraped in 1 ml PBS, pelleted by centrifugation, and lysed in 100 µl of 70% formic acid with sonication. Formic acid from both directly extracted and sequentially extracted samples was removed by vacuum centrifugation for 40 min, and the resulting dry pellet was resuspended in 100 µl of 60% acetonitrile. RIPA buffer (1.9 ml) was added to each of the samples before they were subjected to immunoprecipitation with 6E10. Immunoprecipitated Abeta was resolved on a 10/16.5% step gradient Tris-Tricine gel, fixed in 60% methanol, dried, and placed on PhosphorImager (Molecular Dynamics, Inc., Sunnyvale, CA) plates for 72 h.

Trypsin Treatment of CHO Cells

CHO Pro5 cells were infected with SFV-APPwt for 12 h, rinsed twice in PBS, and incubated on ice for 20 min in either PBS alone, 10 µg/ml of trypsin (Life Technologies, Inc.) in PBS, or 10 µg/ml trypsin plus 0.1% Triton X-100 in an adaptation of a previously described technique (Turner et al., 1996; Chyung et al., 1997). Trypsin was then inactivated by adding 100 µg/ml soybean trypsin inhibitor. The treated cells were then washed with ice-cold PBS, scraped into PBS buffer, centrifuged at 2,000 g for 2 min, resuspended in 100 µl formic acid, sonicated, and centrifuged at 40,000 g for 20 min at 4°C. The supernatant was neutralized with 1.9 ml of 1 M Tris base and diluted 1:3 in H2O for quantification of Abeta 1-40 and Abeta 1-42 by sandwich-ELISA.

Lysis of Cells and Sandwich ELISA

For serial extraction in RIPA and formic acid, cells were washed twice in PBS and then lysed in 600 µl RIPA buffer and centrifuged for 20 min at 40,000 g at 4°C. Supernatant was subjected directly to sandwich ELISA, and the pellet was resuspended in 100 µl 70% formic acid with sonication until clear. Formic acid samples were then neutralized by adding 1.9 ml 1 M Tris base and diluted 1:3 in H2O before quantifying Abeta by sandwich-ELISA.

For direct extraction into formic acid, cells were scraped in PBS after washing twice with PBS. Cells were pelleted by centrifugation at 2,000 g for 2 min, and were then lysed in 100 µl formic acid. Insoluble material was pelleted by centrifugation at 40,000 g at 4°C for 20 min, and the supernatant was neutralized by adding 1.9 ml 1 M Tris base and diluted 1:3 in H2O before quantification of Abeta by sandwich-ELISA.

For extraction into PBS, cells were scraped in PBS after washing twice with PBS. Cells were lysed by sonication, and insoluble material was pelleted by centrifugation at 40,000 g at 4°C for 20 min, and Abeta in the soluble fraction was quantitated by sandwich-ELISA.

Sandwich-ELISA was performed as described previously using mAbs specific for different species of Abeta (Suzuki et al., 1994; Turner et al., 1996). BAN-50 (a mAb specific for the first 10 amino acids of Abeta ) was used as a capturing antibody, and horseradish peroxidase-conjugated BA-27 (a mAb specific for Abeta 1-40) and horseradish peroxidase-conjugated BC-05 (a mAb specific for Abeta 1-42) were used as secondary antibodies. To calibrate the sensitivity of the ELISA for detecting Abeta after formic acid extraction and neutralization, synthetic Abeta 1-40 and Abeta 1-42 peptides (Bachem Bioscience Inc., King of Prussia, PA) used to generate the standard curves were treated with formic acid and neutralized in the same manner as the cell lysates. Under these conditions, the sandwich ELISA had a detection limit of <1 femtomole of synthetic Abeta per sample. The BAN50, BA-27, and BC-05 mAbs were prepared and characterized as described previously (Suzuki et al., 1994).

Cycloheximide Treatment

For experiments involving cycloheximide treatments, NT2N cells were incubated in media containing 150 µg/ml cycloheximide for various time points up to 24 h. Cells were harvested and extracted sequentially in RIPA and formic acid as described above. Samples were then subjected to sandwich ELISA analysis.

Western Blot Analysis of APP Levels

RIPA-extracted cell lysates (15 µg as determined by BCA assay) were resolved on a 7.5% Tris-glycine acrylamide gel and transferred to nitrocellulose for immunoblotting with Karen (a goat anti-APP antibody) at a 1:1,000 dilution (Turner et al., 1996; Chyung et al., 1997). After application of a rabbit anti-goat IgG linker, [125I]Protein A was applied, and radiolabeled APP was quantitated by PhosphorImager analysis.

    Results
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

Neurons Contain Insoluble Amyloid beta  Peptide

To evaluate the possibility that Abeta exists in multiple intracellular pools with different solubility characteristics, NT2N neurons were sequentially extracted in aqueous buffer (PBS), detergent buffer (RIPA), and then 70% formic acid. The levels of Abeta 1-40 and Abeta 1-42 present in each fraction were quantified by sandwich-ELISA. Previous studies have shown that nonionic detergents liberate intracellular Abeta , but not Abeta deposited in senile plaques or fibrillar Abeta formed in vitro (Selkoe et al., 1986; Burdick et al., 1992; Harigaya et al., 1995; Turner et al., 1996). However, more rigorous solubilization methods using 70% formic acid liberate Abeta from these insoluble aggregates. Sonication of cells in PBS in the absence of detergent failed to release any soluble Abeta (data not shown). By contrast, significant levels of Abeta 1-40 and Abeta 1-42 were solubilized by RIPA buffer. Nonetheless, RIPA buffer released only a fraction of the total intracellular Abeta since subsequent extraction of the detergent-insoluble material with 70% formic acid revealed a much larger pool of both Abeta species (Fig. 1 A). Since increased production of Abeta 1-42 relative to Abeta 1-40 has been associated with AD (Borchelt et al., 1996; Duff et al., 1996; Scheuner et al., 1996), we examined the ratios of these Abeta species in the detergent-soluble and -insoluble pools in NT2N neurons. The ratio of Abeta 1-42/1-40 in the RIPA soluble pool was 1.0 ± 0.1 (Fig. 1 B), consistent with previous studies in a variety of experimental systems (Cook et al., 1997; Forman et al., 1997). However, Abeta 1-42 was more abundant in the detergent-insoluble pool, with an Abeta 1-42/1-40 ratio of 2.7 ± 0.3 (Fig. 1 B). This finding is consistent with the reduced solubility of Abeta 1-42 relative to Abeta 1-40 in vitro, and the predominance of Abeta 1-42 in insoluble deposits in the AD brain (Jarrett et al., 1993a; Iwatsubo et al., 1994).


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Fig. 1.   FA extraction of NT2N neurons reveals a large pool of insoluble intracellular Abeta . (A) 10-cm dishes of NT2N cells (replate 2, 4 wk old) were either sequentially extracted in RIPA followed by FA, or lysed directly into FA. Abeta 1-40 and Abeta 1-42 levels in the RIPA and FA samples were quantified by sandwich ELISA. Mean results and standard errors are shown (six separate experiments, each done with duplicate samples). (B) 1-42/1-40 ratios were calculated for the RIPA soluble pool of Abeta , the RIPA insoluble (FA soluble) pool of Abeta , and the total intracellular pool of Abeta for these NT2N neurons, demonstrating that the insoluble intracellular pool of Abeta consists mainly of Abeta 1-42, while the soluble pool contains similar amounts of Abeta 1-40 and Abeta 1-42.

The identification of a large and previously undetected pool of insoluble Abeta in NT2N neurons prompted us to establish precise conditions for reproducible recovery of the maximum amount of formic acid-extractable Abeta . Sonication was found to be necessary for efficient Abeta extraction, and a volume of 100 µl formic acid was found to extract Abeta optimally from cell lysates containing ~1 mg of total protein. However, longer incubation times in formic acid (up to 24 h) or high incubation temperatures (up to 37°C) did not increase Abeta recovery (data not shown). To confirm that formic acid-extracted Abeta was present in intracellular compartments and not attached to the cells or culture dish, cells were treated with trypsin in the presence or absence of 0.1% Triton X-100. We found that formic acid- solubilized intracellular Abeta was resistant to trypsin digestion in the absence of detergent, but sensitive to trypsin digestion after solubilization by Triton X-100 (data not shown). This finding indicates that the formic acid-soluble pool of Abeta is located intracellularly, and is accessible to trypsin only when cell membranes are first permeabilized by detergent. Finally, we found that cells extracted directly into formic acid yielded amounts of Abeta similar to the sum of RIPA-soluble and RIPA-insoluble Abeta (Fig. 1 A). From these studies, we concluded that neurons contain at least two major pools of intracellular Abeta : a detergent soluble pool, and a larger formic acid soluble pool that is enriched in Abeta 1-42.

Insoluble Abeta is Present in a Range of APP-Expressing Cell Types

To determine if insoluble intracellular Abeta is present in cell types other than neurons, NT2, CHO Pro5, and BHK-21 cells were sequentially extracted with RIPA followed by formic acid, and Abeta levels were measured by sandwich-ELISA (Fig. 2). To evaluate the consequences of increased APP production on the generation of soluble and insoluble intracellular Abeta , each cell type was also infected with a recombinant SFV vector that led to the expression of high levels of APP695. Additionally, Abeta levels in stably transfected CHO cells expressing APP695 (CHO-695) were examined (Fig. 2 A). Steady-state APP levels present in each cell type were determined by Western blotting in order to correlate the levels of intracellular Abeta with APP (Fig. 2 B).


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Fig. 2.   FA extraction of a variety of cell lines reveals the presence of varying levels of insoluble intracellular Abeta . (A) Uninfected and SFV-APPwt-infected NT2 cells, NT2N cells (replate 2, 4 wk old), CHO Pro5 cells, CHO-695 cells (uninfected only), and BHK-21 cells were sequentially extracted in RIPA followed by FA. Abeta 1-40 and 1-42 levels in the RIPA and FA samples were quantified by sandwich ELISA. Means and standard errors (four separate experiments done in triplicate) of Abeta levels are shown. (B) Samples from RIPA cell lysates of each of these cell lines (both uninfected and SFV-APPwt infected) were resolved on a 7.5% Tris-glycine acrylamide gel, and immunoblotted with Karen antibody; bands were detected by PhosphorImager after using an I125-labeled secondary antibody.

In contrast to NT2N neurons, retinoic acid naïve NT2 cells did not produce significant amounts of Abeta , despite expressing nearly equivalent levels of APP (Fig. 2, A and B). This observation is consistent with previous experiments that have demonstrated that NT2 cells do not efficiently process APP by the beta -secretase pathway, and thus generate only low levels of Abeta (Wertkin et al., 1993; Forman et al., 1997). Furthermore, the engineered expression of APP695 in NT2 cells at levels similar to those found in NT2N neurons resulted in only a modest increase in intracellular Abeta levels (Fig. 2, A and B), indicating that the lack of intracellular Abeta in NT2 cells relative to NT2N neurons was not due to differential expression of APP isoforms in the two cell types (APP751/770 in NT2 cells vs. APP 695 in NT2N cells), but to differential processing of APP. In addition, the fact that only low levels of Abeta were detected by sandwich-ELISA in this cell line further confirms that this assay is highly specific for Abeta , and does not significantly cross-react with other cellular proteins, including full-length APP, other Abeta -containing carboxy-terminal fragments, or non-Abeta APP-derived fragments.

CHO Pro5 and BHK-21 cells expressed barely detectable levels of APP, and they did not produce detectable levels of soluble or insoluble Abeta , further confirming the specificity of the Abeta ELISA. CHO-695 cells, however, did produce intracellular Abeta , 22 ± 3% of which was insoluble (Fig. 2 A). Likewise, infection of CHO Pro5 cells and BHK-21 cells with SFV-APPwt led to a markedly increased production of APP as well as intracellular Abeta , of which up to 74 ± 5% was insoluble. This dramatic increase in Abeta production over a relatively short period of time could favor Abeta aggregation, resulting in a decrease in Abeta solubility. Indeed, CHO cells stably expressing APP contained a much lower proportion of insoluble Abeta than did SFV-APP-infected CHO cells (Fig. 2 A).

These findings indicate that in addition to cell type-specific factors, the level of APP expression also governs deposition of insoluble Abeta . In cells that efficiently use the beta -secretase pathway to generate Abeta , increased APP expression generally resulted in increased levels of both soluble and insoluble Abeta . However, while CHO-695 cells and NT2N neurons both expressed similar levels of APP and produced similar levels of soluble Abeta , NT2N neurons accumulated significantly higher levels of insoluble Abeta (Fig. 2; compare tracks labeled NT2N vs. CHO-695). This difference may be due to the higher metabolic rate of CHO-695 cells, which may result in increased turnover of Abeta , thus hindering aggregation. Alternatively, Abeta aggregation in CHO-695 cells may be impeded by continual dilution due to cell division. In postmitotic neurons, Abeta may accumulate intracellularly over time, and thus favor the formation of insoluble aggregates.

Taken together, these results indicate that while NT2N neurons accumulate intracellular insoluble Abeta as a consequence of endogenous APP production, other cell types also exhibit this property when they overexpress APP. It is interesting to note that increased expression of APP in NT2N neurons as a consequence of SFV-APPwt infection did not result in increased levels of intracellular Abeta 1-42, consistent with some of our previous work indicating that gamma -secretase cleavage in the ER/IC pathway is rate-limiting (Cook et al., 1997). By contrast, increased expression of APP in NT2N neurons resulted in increased levels of intracellular Abeta 1-40. That this increase was due solely to increased levels of soluble Abeta 1-40 is consistent with this form of Abeta being produced late in the secretory pathway, and being recovered from cells before secretion.

Abeta Can be Immunoprecipitated from an Insoluble Pool

To further confirm that the material recovered by extraction with formic acid and measured by sandwich ELISA was indeed Abeta , SFV-APPwt-infected NT2N and CHO cells were metabolically labeled with [35S]methionine for 12 h, and Abeta was immunoprecipitated using 6E10. As shown in Fig. 3, a band of ~4 kD was immunoprecipitated by an Abeta -specific antibody in the RIPA-soluble cell lysate. Additional Abeta was immunoprecipitated from the RIPA-insoluble (formic acid-extracted) cellular fraction, thus confirming that a pool of Abeta remained insoluble in RIPA buffer, and could be extracted by formic acid (Fig. 3). However, the yield of Abeta after formic acid extraction was lower than that predicted by Abeta sandwich ELISA. To determine if formic acid extraction compromised the recovery of Abeta by immunoprecipitation, [35S]methionine-labeled SFV-APPwt-infected cells were extracted directly into formic acid. Direct extraction of cells into formic acid would be expected to yield amounts of Abeta equal to the sum of Abeta extracted in the RIPA-soluble and -insoluble pools. However, lower levels of Abeta than expected were recovered by this method (Fig. 3; compare lane 3 with lanes 1 and 2, and lane 6 with lanes 4 and 5). Thus, immunoprecipitation of formic acid-extracted cells was not quantitative, and resulted in only partial recovery of Abeta . This low recovery of Abeta may have been due to incomplete resolubilization of Abeta in acetonitrile after lyophilization, or reaggregation of Abeta during immunoprecipitation. To evaluate the contribution of each of these factors to the incomplete recovery of Abeta by immunoprecipitation, we measured Abeta levels in the formic acid-extracted cell lysate before and after lyophilization and immunoprecipitation by sandwich-ELISA. We found that ~43% of formic acid-extracted Abeta could be resolubilized in acetonitrile after lyophilization, and ~45% of this resolubilized Abeta could be captured by immunoprecipitation with the antibody 6E10 (data not shown). Nevertheless, despite the shortcomings of the immunoprecipitation protocol as compared with the Abeta sandwich-ELISA, these data confirm that the formic acid-extracted pool does indeed contain Abeta .


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Fig. 3.   Abeta can be recovered from RIPA-soluble and RIPA- insoluble (FA-solubilized) fractions of cell lysate. SFV-APPwt- infected NT2N cells and CHO Pro5 cells were metabolically labeled for 12 h before being lysed in either FA or RIPA, followed by extraction of insoluble material by FA. RIPA and FA samples were subjected to immunoprecipitation with 6E10, and were resolved on a 10/16.5% step gradient Tris-tricine gel. Molecular weight standards and Abeta bands are labeled.

Differential Production of Insoluble Abeta 1-40 and Abeta 1-42 in Subcellular Compartments

While it has been shown that secreted Abeta is mainly produced in the TGN, intracellular Abeta 1-42, but not Abeta 1-40, is produced in the ER/IC (Cook et al., 1997). To determine if Abeta 1-42 produced in the ER/IC enters the insoluble pool, NT2N neurons and CHO Pro5 cells were infected with SFV-APPwt or SFV-APPDelta KK (an APP mutant containing the dilysine ER retrieval sequence). Infection of both cell types with SFV-APPDelta KK gave similar results: almost a complete abrogation of Abeta 1-40 production, with no diminution of Abeta 1-42 production relative to SFV-APPwt infected cells (Fig. 4, A and B). Importantly, the levels of insoluble Abeta 1-42 were the same in SFV-APPwt and SFV-APPDelta KK-infected cells. These results demonstrate that Abeta 1-42 produced in the ER/IC pathway represents the bulk of the insoluble Abeta 1-42 inside cells. By contrast, insoluble Abeta 1-40 is produced by a post-ER/IC pathway. Finally, these results also prove that insoluble Abeta can accumulate in the absence of secretion, and they provide additional evidence that the Abeta solubilized by formic acid is intracellular.


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Fig. 4.   Insoluble Abeta 1-42 can be produced by APPDelta KK expressing NT2N and CHO cells. (A) N2TN cells and (B) CHO Pro5 cells were infected with SFV-APPwt or SFV-APPDelta KK for 12 h before sequential extraction in RIPA, followed by FA. Abeta in each sample was quantified by sandwich-ELISA. Means and standard errors (two separate experiments each done in triplicate) are shown.

Time-dependent Accumulation of Insoluble Abeta

Our previous studies have shown that secretion of Abeta 1-40 and Abeta 1-42 by the NT2N neurons increases with time in culture without an increase in APP synthesis (Turner et al., 1996). However, a time-dependent increase in intracellular Abeta was not detected. Conversely, we found that retention of APP in the ER/IC resulted in continued production of Abeta 1-42, but without either secretion or intracellular accumulation (Cook et al., 1997). Our observation here that intracellular Abeta (particularly the Abeta 1-42 species produced in the ER/IC) forms an insoluble pool provided a possible explanation for both of these earlier findings. To test the hypothesis that insoluble Abeta can accumulate intracellularly over time, NT2N neurons were analyzed at various time points after replating by sequential extraction in RIPA and formic acid, followed by sandwich-ELISA for Abeta quantitation. We found a dramatic increase (12-fold over 7 wk in culture) in the levels of formic acid- extractable intracellular Abeta 1-40 and Abeta 1-42 in NT2N cells concomitant with increased time in culture (Fig. 5, A and B). In addition to an increase in the absolute amount of insoluble Abeta with longer times in culture, an increase in the fraction of insoluble Abeta was also observed. For example, at 4 wk, ~58% of Abeta was insoluble, while at 7 wk ~78% of Abeta was insoluble (Fig. 5, A and B). This result suggests that the equilibrium of soluble to insoluble Abeta may be shifted to favor insoluble Abeta in NT2N cells that were cultured longer (i.e., older neurons).


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Fig. 5.   Insoluble intracellular Abeta accumulates over time within NT2N cells. 10-cm dishes of Replate 2 NT2N cells were harvested each week 3-7 wk after replating. Cells were lysed in RIPA buffer, and insoluble material was resuspended in FA. Abeta 1-40 (A) and 1-42 (B) in the soluble and insoluble intracellular pools were quantified by sandwich-ELISA. Data were normalized for total protein present (in mg). The experiment was repeated three times, and means and standard errors (three to five samples per time point) are shown for a representative experiment.

The Intracellular Accumulation of Abeta Over Time in Culture is Due to the Slow Turnover of Insoluble Abeta

The time-dependent accumulation of insoluble intracellular Abeta in neurons could be due to several factors, including the slow turnover of insoluble Abeta . To examine this possibility, we treated NT2N cells with cycloheximide to prevent protein synthesis, and measured endogenous levels of Abeta in the soluble and insoluble pools over time in culture. This approach was needed (rather than a standard pulse-chase analysis) because immunoprecipitation of Abeta after formic acid extraction was not quantitative (Fig. 3). Fig. 6 shows that over the 24-h cycloheximide treatment, soluble Abeta 1-40 and Abeta 1-42 decreased by ~63% and ~ 77%, respectively. Assuming a constant rate of degradation, we calculated half-lives of ~18 h and ~12 h for the decay of intracellular soluble Abeta 1-40 and Abeta 1-42, respectively. By contrast, insoluble Abeta 1-40 and Abeta 1-42 levels did not decrease significantly over 24 h. The slow turnover of the insoluble pool of Abeta precluded an accurate estimate of the half-life of this pool. In addition, this analysis is complicated by two factors. First, although no new APP will be synthesized in the presence of cycloheximide, existing pools of APP continue to be processed to generate Abeta . However, the half-life of APP in NT2N neurons is ~3 h. Thus, de novo production of Abeta from existing pools of APP is unlikely to contribute significantly to intracellular Abeta pools, especially at later time points. Second, soluble Abeta 1-40 and Abeta 1-42 may enter the insoluble pool over time, again making accurate estimates of turnover rates difficult. Nevertheless, our results show that intracellular insoluble Abeta is very long-lived, and that this long life is likely to play an important role in the time-dependent accumulation of insoluble Abeta we observed in NT2N cells over weeks in culture.


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Fig. 6.   The insoluble pool of intracellular Abeta is stable over 24 h, while the soluble pool of cellular Abeta turns over more rapidly. 10-cm dishes of Replate 2 NT2N cells were treated with 150 µg/ml cyclohexamide for the times indicated before being sequentially extracted in RIPA and FA. Abeta 1-40 and 1-42 in the soluble (A) and insoluble intracellular pools (B) were quantified by sandwich-ELISA. The experiment was repeated three times, and means and standard errors (three samples per time point) are shown for a representative experiment.

    Discussion
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

The presence of insoluble aggregates of Abeta in senile plaques is a well-characterized feature of AD (Selkoe, 1997). Abeta is composed of two major species that terminate at residues 40 and 42 of the intact Abeta sequence. Both species can be recovered from the CSF of normal and AD individuals, with Abeta 1-40 being approximately 10-fold more abundant than Abeta 1-42 (Citron et al., 1992). However, Abeta 1-42 is the major Abeta species present in senile plaques, with Abeta 1-40 being only a minor constituent (Iwatsubo et al., 1994). That alterations in APP processing can lead to development of AD has been shown by several FAD-associated APP mutations that, when expressed in vitro or in transgenic animals, lead to either an overall increase in Abeta production or an increase in the amount of Abeta 1-42 relative to Abeta 1-40 (Borchelt et al., 1996; Duff et al., 1996). The differential production of Abeta 1-40 and Abeta 1-42 as a consequence of AD-associated APP mutations as well as the preferential deposition of Abeta 1-42 in senile plaques raises important questions as to the intracellular sites of Abeta 1-42 generation, the origin of Abeta that is recovered from senile plaques, and the factors that control its deposition.

Both Abeta 1-40 and Abeta 1-42 are constitutively produced and secreted from cells in vitro and in vivo as judged by their recovery from conditioned medium and CSF (Shoji et al., 1992; Tamaoka et al., 1996). Since FAD-associated APP mutations lead to increased secretion of Abeta , and senile plaques are extracellular lesions, it is possible that secreted Abeta is ultimately deposited in senile plaques, even though the factors controlling its deposition are obscure. However, we have recently discovered that retention of APP in the ER/IC induced by a variety of methods leads to continued production of intracellular Abeta 1-42, but not Abeta 1-40 (Cook et al., 1997). While Abeta 1-42 is constitutively produced by this novel pathway in NT2N neurons, other cell types can also process APP to generate Abeta in the ER/ IC after overexpression of APP (Wild-Bode et al., 1997). Abeta 1-42 has also been shown to be localized to the ER/IC by immunoelectron microscopy and by cell fractionation (Hartmann et al., 1997; Wild-Bode et al., 1997). Interestingly, this compartment also is the site where PS1 and PS2 are localized (Cook et al., 1996; Kovacs et al., 1996). Since mutations in PS1 and PS2 account for the majority of early-onset FAD cases, and FAD-associated PS1 and PS2 mutations have been shown to result in an increased ratio of Abeta 1-42/1-40 (Borchelt et al., 1996; Duff et al., 1996; Scheuner et al., 1996), colocalization of the presenilins with a major site of constitutive Abeta 1-42 production raises the possibility that alterations in Abeta production by the ER/IC pathway may play an important role in AD pathogenesis.

While retention of APP in the ER/IC resulted in continued and selective production of Abeta 1-42, we were unable to document either secretion of this material or its intracellular accumulation (Cook et al., 1997). Taken at face value, this result indicates that the production and turnover of Abeta 1-42 by the ER/IC pathway are in equilibrium. However, given the propensity of Abeta 1-42 to aggregate in vivo and in vitro, we asked whether Abeta 1-42 also aggregated intracellularly. Since formic acid has been shown to effectively solubilize aggregated Abeta present in senile plaques, we solubilized cell lysates in formic acid. Using this approach, we found that a considerable fraction of total intracellular Abeta 1-42, and to a lesser extent Abeta 1-40, could be solubilized by formic acid, but not by a variety of detergents. Currently, we do not know whether or not the formic acid-extractable Abeta self-aggregates or coaggregates with other proteins. Our observation that none of the cell-associated Abeta (including that targeted for secretion) can be extracted with aqueous buffer suggests that it may be bound to other cellular proteins. On the other hand, in vitro studies of Abeta aggregation suggest that Abeta is prone to self-aggregation. Future ultrastructural studies on the accumulated intracelullar Abeta will help to resolve this issue. Intracellular insoluble Abeta was recovered in a number of different APP-expressing cell lines. Overexpression of APP generally resulted in increased production of insoluble Abeta . However, insoluble Abeta was produced most efficiently in NT2N neurons. Thus, while aggregation of intracellular Abeta is not cell type-specific, the subcellular environment in neurons appears to favor this process.

Identification of a novel form of intracellular Abeta that has previously escaped detection could explain our failure to detect secretion or intracellular accumulation of Abeta 1-42 produced by the ER/IC pathway (Cook et al., 1997). To test this possibility, we expressed APP bearing an ER/IC retrieval signal in the cytoplasmic domain in NT2N neurons and in CHO cells. Production of intracellular soluble and insoluble Abeta 1-40 was almost completely inhibited after expression of this construct, while levels of soluble and insoluble intracellular Abeta 1-42 were unchanged by ER retention. Thus, almost all of the formic acid-soluble Abeta 1-42 can be derived from the ER/IC pathway. By extension, insoluble Abeta 1-40 must be produced by a post-ER/IC compartment. Production of insoluble Abeta 1-40 and Abeta 1-42 in different subcellular compartments may help explain the predominance of Abeta 1-42 in the intracellular pool. Abeta 1-40 is produced late in the biosynthetic pathway, and may spend relatively little time in the cell before secretion, thereby minimizing the opportunity for aggregation. By contrast, the bulk of the intracellular Abeta 1-42 is produced by the ER/IC pathway. This fact represents an environment distinct from that in which Abeta 1-40 is produced, and one that does not result in Abeta 1-42 secretion. The long-lived nature of Abeta 1-42, its continued production at an intracellular site from which it cannot be secreted, and the fact that it is intrinsically less soluble than Abeta 1-40 all may contribute to its propensity to enter a stable, intracellular pool of insoluble material. It will be important to define further the factors that govern Abeta deposition in this insoluble pool, and to more carefully study its physical state.

During the course of our experiments, we found that recovery of insoluble Abeta from NT2N neurons was somewhat variable. However, we found that this result was due to a time-dependent accumulation of Abeta . Specifically, we found that Abeta levels increased by 12-fold as the NT2N neurons aged over 7 wk in culture. While insoluble Abeta 1-40 and Abeta 1-42 accumulated at similar rates, more detailed kinetic studies are needed to determine if production of insoluble Abeta 1-40 and Abeta 1-42 is contemporaneous, or if generation of insoluble Abeta 1-42 seeds subsequent polymerization of Abeta 1-40, as has been reported in vitro. In any event, time-dependent accumulation of insoluble Abeta could be due to increased production, decreased turnover, or stable accumulation of Abeta at a relatively constant rate. We found that intracellular insoluble Abeta was exceptionally stable. Thus, even slow addition of Abeta to the insoluble pool over weeks in culture could result in steady accumulation of Abeta seen in the insoluble pool over time. This observation may have implications for AD pathogenesis, where it is thought that accumulation of Abeta occurs slowly over decades. Since AD is an age-dependent disease, the data presented here suggest that gradual accumulation of intracellular Abeta may be a factor in the slow onset and progression of AD. It will be important to determine if accumulation of intracellular insoluble Abeta is simply the result of the stability of this form of Abeta , or if other time-dependent factors (such as altered APP processing or neurotoxic insults) contribute to this process.

Although intracellular beta -amyloid fibrils have been observed in the AD brain (Kim et al., 1988) as well as in a transgenic mouse model of AD (Masliah et al., 1996), it is unclear whether Abeta fibrils can form within neurons from endogenously produced Abeta . The experiments presented here demonstrate that significant levels of Abeta are insoluble within neurons. The observation that Abeta can accumulate with time in a relatively stable insoluble pool may explain how Abeta deposition in senile plaques can begin despite relatively low levels of secreted and CSF-soluble Abeta 1-42. Concentrated intracellular Abeta 1-42 could rapidly nucleate fibril formation, and intracellularly produced Abeta 1-42 and Abeta 1-40 could add to these fibrils over time, thus serving as a nidus for a developing senile plaque.

    Footnotes

Received for publication 20 January 1998 and in revised form 31 March 1998.

   We gratefully thank Dr. N. Suzuki and Tekeda Pharmaceutical for providing the monoclonal antibodies for the Abeta sandwich-ELISA. We also thank Dr. J. Wang for her help with neuronal aging experiments, and Dr. J.Q. Trojanowski for critically reading the manuscript.
   Address all correspondence to Dr. Virginia M.-Y. Lee, Center for Neurodegenerative Disease Research, Department of Pathology and Laboratory Medicine, Maloney 3, HUP, Philadelphia, PA 19104-4283. Tel.: 215-662-6427; Fax: 215-349-5909; E-mail: vmylee{at}mail.med.upenn.edu

This work was supported by grants from the National Institute on Aging. D.M. Skovronsky is the recipient of a Medical Scientist Training Program Predoctoral Fellowship from the National Institutes of Health, and R.W. Doms is the recipient of a Paul Beeson Faculty Scholar Award.

    Abbreviations used in this paper

Abeta , amyloid-beta peptide; AD, Alzheimer's disease; APP, amyloid-beta precursor protein; BHK, baby hamster kidney; ER/IC, endoplasmic reticulum/intermediate compartment; FAD, Familial Alzheimer's disease; SFV, Semliki Forest Virus; SFV-APPwt, SFV expressing wild-type APP695; TGN, trans-Golgi network.

    References
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References

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