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Neurochem Int. Author manuscript; available in PMC Feb 1, 2012.
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PMCID: PMC3040984
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Effects of fatty acid unsaturation numbers on membrane fluidity and α-secretase-dependent amyloid precursor protein processing

Abstract

Fatty acids may integrate into cell membranes to change physical properties of cell membranes, and subsequently alter cell functions in an unsaturation number-dependent manner. To address the roles of fatty acid unsaturation numbers in cellular pathways of Alzheimer's disease (AD), we systematically investigated the effects of fatty acids on cell membrane fluidity and α-secretase-cleaved soluble amyloid precursor protein (sAPPα) secretion in relations to unsaturation numbers using stearic acid (SA, 18:0), oleic acid (OA, 18:1), linoleic acid (LA, 18:2), α-linolenic acid (ALA, 18:3), arachidonic acid (AA, 20:4), eicosapentaenoic acid (EPA, 20:5), and docosahexaenoic acid (DHA, 22:6). Treatments of differentiated human neuroblastoma (SH-SY5Y cells) with AA, EPA and DHA for 24 h increased sAPPα secretion and membrane fluidity, whereas those treatments with SA, OA, LA and ALA did not. Treatments with AA and DHA did not alter the total expressions of amyloid precursor protein (APP) and α-secretases in SH-SY5Y cells. These results suggested that not all unsaturated fatty acids but only those with 4 or more double bonds, such as AA, EPA and DHA, are able to increase membrane fluidity and lead to increase in sAPPα secretion. This study provides insights into dietary strategies for the prevention of AD.

Keywords: Fatty acids, membrane fluidity, sAPPα, Alzheimer's disease

1. Introduction

The accumulation of neurotoxic amyloid-β peptide (Aβ) is a pathologically profound characteristic of Alzheimer's disease (AD) (McGeer et al., 1987; Perlmutter et al., 1990; Frautschy et al., 1998; Dickson, 1999; Stalder et al., 1999; Selkoe, 2000). Aβ is derived from amyloid precursor protein (APP) processing through the amyloidogenic pathway, in which APP is cleaved sequentially by β- and γ-secretases (Vassar, 2004). BACE1 is the major β-secretase for generation of Aβ by neurons (Cai et al., 2001). Alternatively, in the non-amyloidogenic pathway, APP is cleaved by α-secretases between amino acids 16 and 17 within the Aβ domain to produce sAPPα. α-secretases are members of ADAMs (a disintegrin and metalloprotease), including ADAM9, 10, 17 and 19. sAPPα is neurotrophic and neuroprotective (Thornton et al., 2006) and enhancing APP processing by α-secretases has been suggested as a potential therapeutic strategy for AD (Cheng et al., 2007). Since APP, α-, β- and γ-secretases are membrane protein molecules, APP processing should be governed by the local membrane environment. For example, the activity of β-secretase takes place preferentially in highly molecularly-ordered lipid rafts which are cholesterol, saturated phospholipids and sphingolipids-enriched microdomains (Tun et al., 2002; Cordy et al., 2003; Ehehalt et al., 2003; Marlow et al., 2003; Kaether and Haass, 2004; Vetrivel et al., 2004), while the activity of α-secretase is favorable in non-raft domains (Reid et al., 2007). Therefore, APP processing can be manipulated by changing the contents of membrane components, such as cholesterol and sphingolipids (Simons et al., 1998; Kojro et al., 2001; Sawamura et al., 2004; von Arnim et al., 2008).

Since fatty acids are capable of modulating membrane organization and functions (Yehuda et al., 2002; Stillwell et al., 2005; Shaikh and Edidin, 2006; Pepe, 2007; Shaikh and Edidin, 2008), we hypothesized that the effects of fatty acids (e.g. AA) on membrane fluidity and sAPPα secretion are dependent on their unsaturation numbers (i.e. the number of double bonds in the hydrocarbon chains). In this study, we systematically examined the effects of fatty acids with unsaturation numbers ranging from 0 to 6 double bonds including stearic acid (SA, 18:0), oleic acid (OA, 18:1), linoleic acid (LA, 18:2), α-linolenic acid (ALA, 18:3), arachidonic acid (AA, 20:4), eicosapentaenoic acid (EPA, 20:5) and docosahexaenoic acid (DHA, 22:6) on membrane fluidity and sAPPα secretion in differentiated SH-SY5Y cells. Since these fatty acids are ingredients in daily food, information derived from this study should provide potential dietary strategy for prevention of AD.

2. Materials and Methods

2.1. Chemicals and Reagents

Dulbecco's modified Eagle's medium (DMEM)/F12 medium (1:1) and fetal bovine serum (FBS) and 5-hexadecanoylaminofluorescein were from Invitrogen (Carlsbad, CA). Stearic acid (SA, 18:0), oleic acid (OA, 18:1), linoleic acid (LA, 18:2), α-linolenic acid (ALA, 18:3), arachidonic acid (AA, 20:4), eicosapentaenoic acid (EPA, 20:5), docosahexaenoic acid (DHA, 22:6), cis-parinaric acid, Albumin from bovine serum (BSA), phorbol 12-myristate 13-acetate (PMA), dimethyl sulfoxide (DMSO) and all-trans retinoic acid (RA) were from Sigma-Aldrich (St. Louis, MO). Cis-2-eicosenoic acid and cis-5, 8, 11-eicosatrienoic acid were from Cayman (Ann Arbor, MI). Farnesyl-(2-carboxy-2-cyanovinyl)-julolidine (FCVJ) was from Dr. Haidekker's Laboratory (Univerisity of Georgia) (Nipper et al., 2008).

2.2. Cell culture

Human neuroblastoma SH-SY5Y cells (1.0×106 cells/dish) were seeded into 60 mm dishes and were cultured in DMEM/F12 medium (1:1) containing 10% FBS. For differentiation, SH-SY5Y cells were exposed to 10 μM RA for 6 days. Culture medium was replaced by fresh culture medium every other day. Treatments of cells with different fatty acids including SA, OA, LA, ALA, AA, EPA, DHA, cis-2-eicosenoic acid and cis-5, 8, 11-eicosatrienoic acid were in DMEM/F12 medium (1:1) containing 1% BSA for 24 h. All cells were maintained at 37°C in a 5% CO2 humidified incubator.

2.3. Fluorescent fatty acids labeled differentiated SH-SY5Y cells

Cells were incubated with 1μM 5-hexadecanoylaminofluorescein or cis-parinaric acid for 40 min and excess fluorescent fatty acids were then removed by washing cells with PBS three times. Fluorescent images were obtained at room temperature using a Nikon TE-2000 U fluorescence microscope with an oil immersion 60× objective lens. Images were acquired using a CCD camera controlled by a computer running a MetaVue imaging software (Universal Imaging, PA).

2.4. Characterization of membrane fluidity by fluorescence microscopy of FCVJ-labeled cells

A fluorescent molecular rotor, farnesyl-(2-carboxy-2-cyanovinyl)-julolidine (FCVJ), was used to measure the relative membrane fluidity in differentiated SH-SY5Y cells. FCVJ was designed to be a more membrane-compatible fluorescent molecular rotor (Haidekker et al., 2001) with the quantum yield strongly dependent on the local free volume. A higher fluorescent intensity of FCVJ reflects the intramolecular-rotational motions being restricted by a smaller local free volume, indicating a more viscous membrane. The hydrocarbon chain of FCVJ has longer to help improve plasma membrane localization and reduce migration into the inner compartments of the cell, cytosolic staining and background fluorescence (Haidekker et al., 2001). Previously, we have verified the application of FCVJ for measuring membrane viscosity by comparing the results obtained using FCVJ with those from the technique of fluorescence recovery after photobleaching (FRAP) (Nipper et al., 2008). In this study, we adapted the protocol from Haidekker et al. (2001) to fluorescently label cells with FCVJ. Briefly, after undergoing different treatments, e.g. SA, OA, LA, ALA, AA, EPA and DHA, SH-SY5Y cells were washed with PBS and incubated in DMEM containing 20% FBS and 1μM FCVJ for 20 minutes. Excess FCVJ was removed by washing cells with PBS three times. Fluorescent intensity measurements were performed at room temperature using a Nikon TE-2000 U fluorescence microscope with an oil immersion 60× objesctive lens. Images were acquired using a CCD camera controlled by a computer running a MetaVue imaging software (Universal Imaging, PA). The fluorescent intensities of FCVJ per cell were measured. Background subtraction was done for all images prior to data analysis.

2.5. Western blot analysis of sAPPα released from differentiated SH-SY5Y cells

After different treatments, e.g. SA, OA, LA, ALA, AA, EPA and DHA, culture medium was collected and the same volume of the cell lysate from each sample was used for Western blot analysis using β-actin as internal standard. Medium was centrifuged at 12,000 × g for 5 min to remove cell debris, and the same volume of medium from each sample (e.g., 40 μl) was diluted with Laemmli buffer, boiled for 5 min, subjected to electrophoresis in 7.5% SDS-polyacrylamide gels, and transferred to nitrocellulose membranes. Membranes were blocked for 1 h with 5% (w/v) nonfat dry milk in Tris-buffered saline containing 0.1% (v/v) Tween 20 (TBST) and were incubated overnight at 4°C in 3% (w/v) bovine serum albumin (BSA) with 0.02% (w/v) sodium azide in TBST with a 6E10 monoclonal antibody (1:1000 dilution; Covance, Princeton, NJ) that recognizes residues 1–16 of the Aβ domain of sAPPα. Membranes were washed three times during a 15-min period with TBST and incubated with horseradish peroxidase-conjugated goat anti-mouse IgG antibody (1:2000 dilution; Santa Cruz Biotechnology, Santa Cruz, CA) in 5% (w/v) nonfat dry milk in TBST at room temperature for 1 h. After washing with TBST for three times, the membrane was subjected to SuperSignal West Pico Chemiluminescent detection reagents from Pierce (Rockford, IL) to visualize bands. The protein bands detected on x-ray film were quantified using a computer-driven scanner and Quantity One software (Bio-Rad).

2.6. Western blot analysis of APP, ADAM9, ADAM10, ADAM17, ADAM19 and BACE1 in differentiated SH-SY5Y cells

After treatments, the protein concentration of the cell lysate was determined by BCA protein assay kit (Pierce Biotechnology, Rockford, IL) according to manufacture's instruction. Equivalent amounts of protein from each sample (e.g., 30 μg) was diluted with Laemmli buffer, boiled for 5 min, subjected to electrophoresis in 7.5% SDS-polyacrylamide gels, and transferred to nitrocellulose membranes. Membranes were blocked for 1 h with 5% (w/v) nonfat dry milk in Tris-buffered saline containing 0.1% (v/v) Tween 20 (TBST) and were incubated overnight at 4°C in 3% (w/v) BSA with 0.02% (w/v) sodium azide in TBST with 6E10 monoclonal antibody, anti-ADAM9 antibody or anti-ADAM19 antibody (1:1000 dilution; Abcam, Cambridge, MA), anti-ADAM10 antibody (1:1000 dilution; Millipore, Billerica, MA) or anti-ADAM17 antibody (1:1000 dilution; Santa Cruz Biotechnology, Santa Cruz, CA), anti-BACE1 antibody (1:1000 dilution; Sigma-Aldrich, St. Louis, MO). Membranes were washed three times during a 15-min period with TBST and incubated with horseradish peroxidase-conjugated goat anti-mouse or anti-rabbit IgG antibody (1:2000 dilution; Santa Cruz Biotechnology, Santa Cruz, CA) in 5% (w/v) nonfat dry milk in TBST at room temperature for 1 h. After washing with TBST for three times, the membrane was subjected to SuperSignal West Pico Chemiluminescent detection reagents from Pierce (Rockford, IL) to visualize bands. The protein bands detected on x-ray film were quantified using a computer-driven scanner and Quantity One software (Bio-Rad).

2.7. Quantification of Aβ1-42

After treatments, culture medium and cell lysates were collected, supplemented with protease inhibitor cocktail and centrifuged at 12,000 × g for 5 min at 4°C to remove cell debris. An aliquot (100 μl) of supernatant was used for Aβ1-42 quantification using an ELISA kit (Invitrogen, Carlsbad, CA) following manufacturer's recommendation. According to the instruction manual, substances including Aβ1–12, Aβ1–20, Aβ12–28, Aβ22–35, Aβ1–40, Aβ1–43, Aβ42–1 and APP have no cross-reactivity. The minimum detectable dose of Aβ1-42 is <1.0 pg/ml which is similar to a previous study (Prasanthi et al., 2009). The level of Aβ1-42 in each sample was measured in duplicates and expressed in pg/ml.

2.8. Statistical analysis

Data are presented as mean ± SD from at least three independent experiments. There are three trials in each experiment. Comparison between two groups was made with student's t test. Comparisons of more than two groups were made with one-way ANOVA, followed by Bonferroni's post hoc tests. Values of p< 0.05 are considered to be statistically significant.

3. Results

3.1. Exogenous fatty acids integrated into cellular membranes

In order to study the effects of fatty acids with different unsaturations on cellular membrane fluidity and sAPPα secretion, we first confirmed if fatty acids were able to integrate into membranes of SH-SY5Y cells. Therefore, we incubated differentiated SH-SY5Y cells for 40 min with 5-hexadecanoylaminofluorescein and cis-parinaric acid. Both 5-hexadecanoylaminofluorescein and cis-parinaric acid are fluorescent fatty acids representing saturated and unsaturated fatty acids, respectively. Fig. 1A and Fig. 1B showed that 5-hexadecanoylaminofluorescein and cis-parinaric acid were incorporated into cellular membranes of SH-SY5Y cells, suggesting that exogenous fatty acids used in this study are capable of integrating into cellular membranes in SH-SY5Y cells.

Figure 1
Fluorescently labeled fatty acids incorporated into membranes in differentiated SH-SY5Y cells. 5-hexadecanoylaminofluorescein (A) and cis-parinaric acid (B) represent saturated and unsaturated fatty acids, respectively. After incubation for 40 min, fluorescent ...

3.2. Fatty acids with 3 or less double bonds did not alter membrane fluidity and sAPPα secretion

To study the effects of fatty acids on membrane fluidity, we applied a fluorescent molecular rotor, FCVJ. As explained in the Materials and Methods section, FCVJ integrated into a highly fluidized membrane exhibits a lower quantum yield, as reflected by a lower fluorescent intensity. The application of this technique for the measurement of membrane fluidity has been validated (Nipper et al., 2008; Yang et al., 2010). FCVJ-labeled differentiated SH-SY5Y cells were exposed to SA, OA, LA and ALA for 24 h. Interestingly, cells did not exhibit a significant change in membrane fluidity as indicated by the unchanged fluorescent intensity of FCVJ as compared to control (Fig. 2A, B, C, D, right). These results suggest that SA, OA, LA and ALA do not alter membrane fluidity in SH-SY5Y cells.

Figure 2
SA, OA, LA and ALA on sAPPα secretion and membrane fluidity in differentiated SH-SY5Y cells

To study the effects of SA, OA, LA and ALA on sAPPα secretion, cell culture medium was subjected to western blot analysis of sAPPα after the treatments of cells with SA, OA, LA and ALA. SA, OA, LA and ALA did not alter sAPPα secretion in SH-SY5Y cells (Fig. 2A, B, C, D, left). Since it has been reported that PMA, a PKC agonist, increases sAPPα secretion (Caporaso et al., 1992; Slack et al., 1993; Camden et al., 2005), treatment with PMA (10nM) was used as a positive control.

To test whether positions of double bonds plays a role in membrane fluidity and sAPPα secretion, differentiated SH-SY5Y cells were exposed to cis-2-eicosenoic acid and cis-5, 8, 11-eicosatrienoic acid, which have the same numbers of double bonds as OA and ALA respectively, but the double bonds are closer to the carboxylic groups of fatty acids. Results showed that these two fatty acids also had no effect on membrane fluidity and sAPPα secretion in SH-SY5Y cells, like OA and ALA (Fig. 3), suggesting that the positions of double bonds in fatty acids do not play a role in their effects on membrane fluidity and APP processing.

Figure 3
cis-2-Eicosenoic acid (20:1) and cis-5, 8, 11-Eicosatrienoic acid (20:3) on sAPPα secretion and membrane fluidity in differentiated SH-SY5Y cells

3.3. Fatty acids with 4 or more double bonds increased membrane fluidity and sAPPα secretion

We then tested the effects of fatty acids with 4 or more double bonds including AA, EPA and DHA on membrane fluidity and sAPPα secretion. After treatment with AA, EPA and DHA, cells exhibited more fluidized membranes as indicated by lower fluorescent intensities of FCVJ compared to control (Fig. 4A, B, C, right). Western blot analysis showed that AA, EPA and DHA increased sAPPα secretion in differentiated SH-SY5Y cells (Fig. 4A, B, C, left).

Figure 4
AA, EPA and DHA increased the sAPPα secretion and membrane fluidity in differentiated SH-SY5Y cells

Together with results from SA, OA, LA and ALA treatments, these data showed that unsaturation numbers of fatty acids, but not the positions of double bonds, determined their effects on membrane fluidity and sAPPα secretion, as summarized in Fig. 5. These results suggested that fatty acids with 4 or more double bonds increased sAPPα secretion through increasing membrane fluidity in differentiated SH-SY5Y cells.

Figure 5
The effects of the number of double bonds in fatty acids on sAPPα secretion and membrane fluidity in differentiated SH-SY5Y cells

3.4. AA and DHA did not alter the expressions of total APP and α-secretases in SH-SY5Y cells

To rule out the changes of sAPPα secretion possibly due to the changes in the expressions of total APP and α-secretases in cells, western blot analyses of APP and α-secretases were performed after the treatments of cells with AA and DHA for 24 h. Fig. 6 shows that AA and DHA did not alter the expressions of total APP and different isoforms of α-secretases including ADAM9, ADAM10, ADAM17 and ADAM19. Mature APP blots were shown in this study. These results suggested that AA, DHA and EPA do not alter α-secretases and APP expressions to contribute to the increase in sAPPα secretion.

Figure 6
Effects of AA and DHA on total APP and α-secretases expression in differentiated SH-SY5Y cells

3.5. AA, EPA and DHA did not alter BACE1 expression, Aβ1-42 secretion and expression in SH-SY5Y cells

Since AA, EPA and DHA increase sAPPα secretion, they may alter expression of BACE1, the major β-secretase, and Aβ1-42 in SH-SY5Y cells. Western blot analysis showed that AA, EPA and DHA did not change BACE1 expression (Fig. 7A). ELISA measurement was used to quantify Aβ1-42 in culture medium. Fig. 7B showed that AA, EPA and DHA did not change the Aβ1-42 secretion and expression level in SH-SY5Y cells.

Figure 7
Effects of AA, EPA and DHA on BACE1 expression and Aβ1-42 from differentiated SH-SY5Y cells

4. Discussion

In this study, we have systematically examined how unsaturation numbers of fatty acids affected membrane fluidity and sAPPα secretion in differentiated SH-SY5Y cells. We found that not all unsaturated fatty acids, but only fatty acids with at least 4 or more double bonds increased membrane fluidity and sAPPα secretion (Fig. 5). The effects on membrane fluidity and sAPPα secretion were not dependent on the positions of double bonds in fatty acids. In addition, treatments of cells with fatty acids did not alter the expressions of APP and α-secretases in differentiated SH-SY5Y cells, which ruled out the effects of fatty acids on sAPPα secretion possibly due to the changes in APP and α-secretases expressions.

Studies have shown the roles of fatty acids on health and diseases (Bazan, 2007; Das, 2008; Hooijmans and Kiliaan, 2008). Fatty acids are important ingredients in various dietary sources (Connor, 2000; Chow, 2007). For example, ALA, an essential fatty acid, can be found in many vegetable oils (Mozaffarian, 2005), which has been reported to be neuroprotective in in vivo model of global ischemia (Lauritzen et al., 2000). SA is common in animal fats, and cocoa butter (Beare-Rogers et al., 2001). OA is a mono-unsaturated ω-9 fatty acid, which can be found in peanut oil and poppy seed oil (Untoro et al., 2006). DHA, mainly found in fishes, is also abundant in neuronal cell membranes, especially in synaptic membranes (Bazan and Scott, 1990) and myelin sheaths (Ansari and Shoeman, 1990). DHA is also essential for prenatal brain development and normal brain functions. Its levels in serum and brains are lower in AD patients compared with those in age-matched controls (Tully et al., 2003). Furthermore, greater consumption of DHA significantly reduced the likelihood of developing AD (Schaefer et al., 2006). AA is another abundant fatty acid in the brain. It is a second messenger (Khan et al., 1995) and a precursor for synthesis of eicosanoids (Zhou and Nilsson, 2001). It also helps maintain cell membrane fluidity (Fukaya et al., 2007). The disturbed metabolism of AA is associated with neurological disorder such as AD (Rapoport, 2008).

APP processing depends on the local membrane environment (Tun et al., 2002; Cordy et al., 2003; Ehehalt et al., 2003; Marlow et al., 2003; Kaether and Haass, 2004; Vetrivel et al., 2004; Reid et al., 2007) and can be altered by manipulating the membrane lipid composition (Simons et al., 1998; Kojro et al., 2001; Sawamura et al., 2004; von Arnim et al., 2008). Since fatty acids modulate membrane organization and functions (Yehuda et al., 2002; Stillwell et al., 2005; Shaikh and Edidin, 2006; Pepe, 2007; Shaikh and Edidin, 2008), they may affect APP processing. Therefore, we tested the hypothesis that the effects of fatty acids on membrane fluidity and sAPPα secretion depend on the number of double bonds (i.e. the unsaturation number) in the hydrocarbon chains of fatty acids.

To test the hypothesis, we investigated the effects of fatty acids with different unsaturation (from 0 to 6 double bonds) on membrane fluidity and sAPPα secretion in differentiated SH-SY5Y cells. Our data showed that fatty acids with 3 or less double bonds including SA, OA, LA and ALA had no effects on membrane fluidity and sAPPα secretion (Fig. 2). We also exposed SH-SY5Y cells to cis-2-eicosenoic acid and cis-5, 8, 11-eicosatrienoic acid, which has 1 or 3 double bonds as OA and ALA, respectively, but their double bonds are closer to the carboxylic group of fatty acids. Results showed that these two acids had no effect on membrane fluidity and sAPPα secretion in differentiated SH-SY5Y cells either, suggesting that the position of the double bonds may not play a role in altering membrane fluidity and APP processing (Fig. 3). On the other hand, fatty acids with 4 or more double bonds including AA, EPA and DHA increased membrane fluidity and sAPPα secretion (Fig. 4). Our data are consistent with other studies that AA increased membrane fluidity in hippocampal neurons in vivo (Fukaya et al., 2007) and cultured human umbilical vein, cerebral endothelial cells (Villacara et al., 1989; Beck et al., 1998). In addition, DHA has been shown to increase membrane fluidity and sAPPα secretion in HEK cells and SH-SY5Y cells overexpressing APP (Kogel et al., 2008). Our study demonstrated that not all the unsaturated fatty acids, but only fatty acids of unsaturation number ≥4, are able to increase membrane fluidity, and subsequently promote sAPPα secretion.

To explore if AA, EPA, DHA effects on membrane fluidity and secretion of sAPPα have dose dependency, we exposed SH-SY5Y cells to AA, EPA and DHA with lower doses at 10nM and 100nM. At lower doses, AA, EPA and DHA have no effect on membrane fluidity and sAPPα secretion (data not shown). Taken together, our data show that polyunsaturated fatty acids (PUFAs) only begin to have effects on membrane fluidity and APP processing at certain threshold value between 100nM and 1μM.

Membrane fluidity plays important roles in pathogenesis of AD. Hippocampal membranes of AD patients showed a significant lower fluidity compared with membranes from elderly non-demented controls (Eckert et al., 1999). Membrane fluidity in AD patients was correlated with abnormal APP processing and cognitive decline (Zainaghi et al., 2007). The clinical study by Croisile et al. (1993) indicated that long-term and high-dose treatment with piracetam may slow down the progression of AD and increased membrane fluidity (Eckert et al., 1999). DHA protection effect in Aβ-infused rats was associated with increased membrane fluidity (Hashimoto et al., 2006) which also provided oxidative stress resistance in hippocampal cells (Clement et al., 2010). In vivo, increased membrane fluidity in rat hippocampus improved memory formation (Muller et al., 1997; Scheuer et al., 1999), whereas reduced fluidity impaired memory (Schaeffer et al., 2005; Schaeffer and Gattaz, 2007). Furthermore, membrane fluidity affected not only APP processing (Kojro et al., 2001), but also Aβ aggregation size and hydrophobicity (Kremer et al., 2000), which is critical for the formation of Aβ plaques and downstream cellular pathways.

PUFAs play a central role in the normal development and functioning of brain (Schuchardt et al., 2010). Diets enriched in ω-3 PUFA increased membrane fluidity, affect signal transduction and modulate gene expression for brain function (Horrocks and Farooqui, 2004). Preclinical studies suggested that DHA maintains membrane fluidity, improved synaptic and neurotransmitter functioning, enhanced learning and memory performances and displayed neuroprotective properties (Carrie et al., 2009). Meanwhile, DHA decreased the amount of vascular Aβ deposition (Hooijmans et al., 2007) and reduced Aβ burden (Lim et al., 2005) in aged Alzheimer mouse model. In AD mouse model, DHA modulated APP processing by decreasing both α- and β-APP C-terminal fragment products and full-length APP (Lim et al., 2005), which is contradictory to the idea that higher membrane fluidity increased sAPPα production. More works are warrant to resolve this discrepancy. Despite favorable effects of DHA, it should be noted that DHA has a potential to increase oxidative stress (Meydani et al., 1991), resulting in lipid peroxidation (Song and Miyazawa, 2001), especially with long-term and high dose treatments (Grundt et al., 2003; Wang et al., 2003). Peroxidation of lipids and lipoproteins, especially low-density lipoprotein (LDL), was thought to lead to the development of atherosclerotic plaques (Wiklund et al., 1991). As such, although the impact of PUFAs on lipid peroxidation is controversial (Nenseter and Drevon, 1996), considerations should be taken when PUFAs are used for dietary supplementation.

Although we report that AA promoted sAPPα secretion, it is important to note that AA exerts both neurotrophic and neurotoxic effects (Farooqui and Horrocks, 2006). The concentration of AA (<10 μmol/kg) in the brain tissue is very low under normal conditions. AA performs a variety of functions, such as regulating activity of protein kinase A, protein kinase C, NADPH oxidase, choline acetyltransferase, and caspase-3 (Farooqui and Horrocks, 2006). 1 μM of AA also significantly potentiates the elongation of neurites in hippocampal cultures (Katsuki and Okuda, 1995). Here we report that 1 μM of AA increased membrane fluidity and promoted the secretion of sAPPα which is neurotrophic and neuroprotective. However, under pathological conditions, higher concentrations (~ 0.5 mmol/kg) of AA are present in the brains (Farooqui and Horrocks, 2006). Higher concentrations of AA result in mitochondrial dysfunction (Schapira, 1996). AA also triggers nuclear factor-κB (NF-κB) and increases neuronal death (Toborek et al., 1999). AA can be converted to inflammatory mediators, prostaglandins and leukotrienes, by prostaglandin synthetases and lipoxygenases, respectively (Farooqui and Horrocks, 2006). Therefore, whether AA is beneficial to brains is highly concentration-dependent.

Other compounds capable of altering membrane fluidity can also modulate sAPPα secretion. For example, treatment with methyl-β-cyclodextrin (MβCD) or lovastatin to reduce cellular cholesterol resulted in increase in membrane fluidity and sAPPα secretion (Kojro et al., 2001). In addition, decreased sAPPα secretion by Pluronic F68 (PF68) is associated with decreased membrane fluidity, while increased sAPPα by benzyl alcohol (C6H5OH) is associated with increased membrane fluidity (Peters et al., 2009). These data suggested that sAPPα secretion can be altered by manipulating membrane fluidity (Wolozin, 2001). Taken together, our data and those from Kojro et al (2001) and Peter et al (2009) provide the explanation that fatty acids with 4 or more double bonds promote sAPPα secretion through increasing membrane fluidity in differentiated SH-SY5Y cells.

sAPPα has been shown to be neurotrophic. sAPPα enhanced neuronal survival and neurite extension in a dose-dependent manner with a maximum effect at approximately 100 nM in rat primary cerebral cortical neurons (Araki et al., 1991) and significantly increased neurite length and branching in pheochromocytoma PC12 cells (Milward et al., 1992) and stimulated neurite outgrowth of hippocampal neurons in an isoform-dependent manner (Qiu et al., 1995). ADAM10 over-expression, which led to increased sAPPα production, increased cortical synaptogenesis (Bell et al., 2008). sAPPα is also neuroprotective, although mechanisms are yet to be fully understood. For example, sAPPα enhanced basal glucose and glutamate transport and protected against oxidative impairment of glucose and glutamate transport in synaptosomes by a cyclic GMP-mediated mechanism (Mattson et al., 1999); sAPPα exerted neuroprotective effects through regulating intraneuronal calcium (Mattson et al., 1993); sAPPα antagonized dendritic degeneration and neuron death triggered by proteasomal stress (Copanaki et al., 2010), and this anti-apoptotic effect of sAPPα was associated with inhibition of the stress-triggered c-Jun N-terminal kinase (JNK)-signalling pathway; Statins, widely used cholesterol-lowering drugs, exerted neuroprotective effects through stimulating α-secretase cleavage of APP (Ma et al., 2009). Obviously, more efforts are needed to further understand the neurotrophic and neuroprotective mechanisms of sAPPα.

Information derived from this study should provide potential dietary strategies for prevention of AD.

Acknowledgments

This work was supported by NIH Grants 1P01 AG18357 and 1R21 NS052385.

Abbreviations used

AA
arachidonic acid
AD
Alzheimer's disease
ADAM
a disintegrin and metalloprotease
ALA
α-linolenic acid
APP
amyloid precursor protein
amyloid-β peptide
BCA
bicinchoninic acid
BSA
bovine serum albumin
DHA
docosahexaenoic acid
DMEM
dulbecco's modified Eagle's medium
DMSO
dimethyl sulfoxide
EPA
eicosapentaenoic acid
FBS
fetal bovine serum
FCVJ
farnesyl-(2-carboxy-2-cyanovinyl)-julolidine
FRAP
fluorescence recovery after photobleaching
LA
linoleic acid
MβCD
methyl-β-cyclodextrin
NF-κB
nuclear factor-κB
OA
oleic acid
PF68
Pluronic F68
PMA
phorbol 12-myristate 13-acetate
PUFA
polyunsaturated fatty acid
RA
all-trans retinoic acid
SA
stearic acid
sAPPα
α-secretase-cleaved soluble APP
sPLA2-III
secretory phospholipase A2 type III

Footnotes

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