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Introduction

Prion diseases or transmissible spongiform encephalopathies (TSEs) are neurodegenerative disorders that are characterized by loss of motor control, dementia, central nervous system (CNS) spongiosis, and microglial activation (1,2).

TSEs are caused by an infectious agent, prion, whose major component is a pathological form of the prion protein termed the scrapie isoform (PrPSc) (3). PrPsc acts as a template for the conversion of normal form of the prion protein (the cellular isoform, PrPc) to PrPsc (4). In many cases this is also accompanied by the accumulation of the PrPSc that leads to neuronal apoptosis, extensive neuronal loss, and mitochondrial disruption (5). Many pathogenic characteristics of PrPSc have been confirmed in a peptide corresponding to residues 106-126 of PrP [PrP (106-126)] (6). Moreover, PrP (106-126) was reported to induce apoptotic cell death via dysregulation of mitochondrial homeostasis in neuronal cells (7). Thus, PrP (106-126) has been used as a model to study prion-induced neuronal cell death and has been postulated to induce mitochondrial dysfunction (8).

Mitochondria are essential organelles found in various cell types that play a principal role in cell survival and apoptotic cell death (9). Mitochondrial oxidative damage contributes to a range of degenerative diseases (10). Mitochondrial dysfunction caused by unnatural regulation of mitochondrial dynamic proteins may lead to neuropathological changes in prion disorders (11). In addition, PrP (106-126)-induced neuronal cell damage that occurs in neurodegenerative disorders causes mitochondrial disruption (12). Furthermore, oxidative stress is key in mitochondrial-mediated apoptotic cell death (13).

Oxidative stress is a baneful condition caused by reactive oxygen species (ROS) and/or a decrease in antioxidant levels (14). In neurodegenerative disorders, oxidative stress-induced neurodegeneration is mediated by ROS production (15). In addition, mitochondrial dysfunction is associated with ROS (16). PrP (106-126)-induced neuronal cell damage occurs in neurodegenerative disorders via regulation of cellular oxidation pathways (17).

Lactoferrin (LF) is an 80 kDa protein found in colostrum, milk, and mucosal secretions such as blood, saliva, and tears (18). It is a multifunctional protein of the transferrin family, which is involved in the regulation of immune responses, regulation of neutrophil apoptosis, antioxidation, iron binding ability, and antimicrobial activity (19). The antioxidation capability of LF is due to the scavenging of ROS (20). For example, LF inhibits the subsequent production of ROS by neutrophils (21). However, the molecular mechanism of LF-mediated neuronal survival is only beginning to be understood.

We hypothesized that LF can prevent PrP (106-126)-induced oxidative stress and neuronal cell death by regulating ROS generation. To test this hypothesis, we investigated the antioxidant effect of LF in PrP (106-126)-induced neuronal cell death. In particular, we tested whether LF protects from neuronal cell death by PrP (106-126) and assessed the therapeutic value of LF in the treatment of neurodegenerative disorders.

Materials and methods

Cell culture

The SH-SY5Y human neuroblastoma cell line was obtained from the American Type Culture Collection (ATCC, Rockville, MD, USA). Cells were cultured in Minimum Essential Medium (MEM; Invitrogen-Gibco, Grand Island, NY, USA) that contained 10% fetal bovine serum (FBS; Invitrogen-Gibco) and penicillin-streptomycin (both 100 U/ml) in a humidified incubator maintained at 37°C and 5% CO2.

Reagents

LF from bovine colostrums was purchased from Sigma-Aldrich (St. Louis, MO, USA). The antioxidant agents glutathione (GSH) and N-acetylcysteine (NAC) were purchased from Sigma-Aldrich.

PrP (106-126) treatment

Synthetic PrP (106-126) (sequence, Lys-Thr-Asn-Met-Lys-His-Met-Ala-Gly-Ala-Ala-Ala-Ala-Gly-Ala-Val-Val-Gly-Gly-Leu-Gly) was synthesized by Peptron (Seoul, Korea). The peptide was dissolved in sterile dimethylsulfoxide (DMSO) at a concentration of 10 mM and stored at −80°C.

Western blot analysis

SH-SY5Y was lysed in a buffer containing 25 mM HEPES; pH 7.4, 100 mM NaCl, 1 mM EDTA, 5 mM MgCl2, 0.1 mM dithiothreitol (DTT), and protease inhibitor mixture. Proteins were electrophoretically resolved by 10–15% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), and immunoblotting was performed as previously described. Equal amounts of lysate protein were similarly electrophoretically resolved and electrophoretically transferred to a nitrocellulose membrane. Immunoreactivity was detected through sequential incubation with horseradish peroxidase-conjugated secondary antibody and enhanced chemiluminescence reagents. The antibodies used for immunoblotting were phospho-c-Jun, N-terminal kinase (p-JNK; Santa Cruz Biotechnology, Inc., Santa Cruz, CA, USA), cleaved caspase-3 (Cell Signaling Technology, Danvers, MA, USA), and β-actin (Santa Cruz Biotechnology, Inc.).

Cellular fractionation

SH-SY5Y cells were resuspended in mitochondrial buffer (210 mM sucrose, 70 mM mannitol, 1 mM EDTA, 10 mM HEPES), broken by a 26-gauge needle, and centrifuged at 700 × g for 10 min. The postnuclear supernatant was centrifuged at 10,000 × g for 30 min. The pellet was used as the mitochondrial fraction and the supernatant was used as the cytosolic fraction. Total proteins were obtained and subjected to western blotting.

Annexin V assay

Apoptosis was assessed by a commercial Annexin V assay (Santa Cruz Biotechnology, Inc.) according to the manufacturer’s protocol. Annexin V content was determined by measuring fluorescence at an excitation wavelength of 488 nm and emission wavelengths of 525 and 530 using a Guava easyCyte HT System (Millipore, Billerica, MA, USA).

Immunofluorescence

SH-SY5Y cells cultured on glass cover-slips were treated with PrP (106-126). Cells were washed with phosphate-buffered saline (PBS) and fixed with cold acetone for 90 sec. Cells were washed with PBS, blocked with 5% FBS in Tris buffer saline containing Tween-20, and incubated with anti-caspase-3 (2 μg/ml) and anti-p-JNK (2 μg/ml) monoclonal antibodies for 48 h at 20°C. Unbound antibody was removed by an additional PBS wash, and cells were incubated with labeled anti-rabbit Alexa Fluor 546 (for anti-caspase-3) IgG antibody (4 μg/ml) and Alexa Fluor 488 (for anti-p-JNK) IgG antibody (4 μg/ml) for 2 h at 20°C. Finally, cells were mounted with DakoCytomation fluorescent medium and visualized via fluorescence microscopy.

Terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) assay

TUNEL analysis was performed to measure the degree of cellular apoptosis using an in situ ApoBrdU DNA fragmentation assay kit (BioVision, San Francisco, CA, USA) following the manufacturer’s instructions.

DCFH-DA assay

SH-SY5Y cells were incubated in minimum essential medium (Hyclone Laboratories, Logan, UT, USA) containing 10 μM 2′,7′-dichlorodihydrofluorescein diacetate (H2-DCFDA) at 37°C for 30 min. Cells were washed with PBS and lysed in the aforementioned lysis buffer. Cells were transferred to a clear 96-well plate and fluorescent emission from the bottom of the plate was measured at 515 nm with an excitation wavelength of 488 nm using a SpectraMax M2 instrument (Molecular Devices, Sunnyvale, CA, USA). SH-SY5Y cells were cultured on coverslips positioned in a 24-well plate. Cells were incubated in MEM (Hyclone Laboratories) containing 10 μM H2-DCFDA) at 37°C for 30 min. Cells were washed with PBS.

Mitochondrial transmembrane potential (MTP) assay

The change in MTP was evaluated by the cationic fluorescent indicator JC-1 (Molecular Probes, Eugene, OR, USA), which aggregates in intact mitochondria (red fluorescence) indicating high or normal MTP and low MTP when it remains in monomeric form in the cytoplasm (green fluorescence). SH-SY5Y cells were incubated in MEM containing 10 μM JC-1 at 37°C for 30 min, washed with PBS, and then transferred to a clear 96-well plate. JC-1 aggregate fluorescent emission was measured at 583 nm with an excitation wavelength of 526 nm, and JC-1 monomer fluorescence intensity was measured with an excitation and emission wavelength of 525 and 530 nm, respectively, using a Guava easyCyte HT System (Millipore). SH-SY5Y cells were cultured on coverslips in a 24-well plate, incubated in MEM containing 10 μm JC-1 at 37°C for 30 min, and then washed with PBS. Finally, cells were mounted with DakoCytomation fluorescent medium and visualized via fluorescence microscopy.

Statistical analysis

All data are expressed as the means ± standard deviation (SD), and the data were compared using the Student’s t-test and the ANOVA Duncan test with the SAS statistical package (SAS, Cary, NC, USA). The results were considered to indicate statistically significant differences at *P<0.05 or **P<0.01.

Results

PrP (106-126)-induced neuronal cell death is decreased by LF treatment in SH-SY5Y neuroblastoma cells

In a previous study, it was shown that LF inhibits prion accumulation (22). Thus, we presently examined whether LF protects against PrP (106-126)-mediated neurotoxicity. To study the influence of LF on PrP (106-126)-induced neuronal cell death, SH-SY5Y cells were pretreated with various concentrations of LF (12 h) and then exposed to 100 μM PrP (106-126) for 8 h (Fig. 1B). The preventative effect of LF was evaluated using the Annexin V assay of cell viability. As shown in Fig. 1A, LF treatment prevented PrP (106-126)-induced neuronal cell death. SH-SY5Y cells were responsive to PrP (106-126) treatment (46.94% increase in Annexin V-positive cells) and PrP (106-126)-induced neuronal cell death was decreased by LF pretreatment (Fig. 1A). TUNEL assay revealed the protective effect of LF on PrP (106-126)-induced apoptosis of SH-SY5Y cells (Fig. 1C). These results suggest that LF prevents PrP (106-126)-induced neuronal cell death.

Figure 1. (A) LF protects against PrP (106-126)-induced neuronal cell damage. SH-SY5Y neuronal cells were pretreated with LF (12 h) and then exposed to 100 μM PrP (106-126) for 12 h. Cell viability was measured by an Annexin V assay using flow cytometry. (B) Bar graph indicates the averages of Annexin V-positive cells. *P<0.05 vs. control; **P<0.01 vs. PrP (106-126). (C) Representative immunofluorescence images of TUNEL-positive (white) SH-SY5Y cells at 12 h after exposure to PrP (106-126) (100 μM) in the absence or presence of LF (12 h). The cells were counterstained with propidium iodide (dark gray) to show all cell nuclei. Magnification ×400; scale bar, 100 μM.

LF treatment suppresses PrP (106-126)-mediated protein activation

We examined the effects of LF treatment on the JNK and caspase-3 activation. Western blot analyses revealed that activation of JNK and caspase-3 increased expression in the 100 μM PrP (106-126)-treated group compared to the LF (200 μg/ml)-pretreated group and the control group (Fig. 2A). PrP (106-126) treatment induced the activation of JNK and caspase-3 in SH-SY5Y cells. However, LF treatment inhibited the activation of JNK and caspase-3 (Fig. 2A and B). Consistent with these results, immunofluorescence monitoring also showed that LF treatment completely inhibited PrP (106-126)-mediated protein activation (Fig. 2C). These results suggest that LF treatment suppresses PrP (106-126)-induced protein activation.

Figure 2. PrP (106-126)-induced JNK, Cas3 activation is attenuated by LF treatment. SH-SY5Y neuronal cells were pretreated with (A) LF (12 h) or (B) LF (12 h) in a dose-dependent manner, and then exposed to 100 μM PrP (106-126) for 12 h. (C) The treated cells were assessed for phospho-JNK, cleaved Cas3 production by western blot analysis. Results were normalized with β-actin. The treated cells were also assessed for phospho-JNK, cleaved Cas3 production by immunofluorescence.

LF treatment decreases PrP (106-126)-induced oxidative stress via ROS scavenging

In a previous study, it was shown that LF is a scavenger of ROS (20), and that this protects against ROS-mediated cell death. PrP (106-126)-induced neuronal cell death is mediated by ROS generation (23). Thus, we next assessed whether the protective effect of LF on PrP (106-126)-induced neuronal cell death was related to ROS generation. SH-SY5Y cells were preincubated 12 h with 200 μg/ml LF and then exposed to 100 μM PrP (106-126) for 12 h. LF treatment reduced PrP (106-126)-induced ROS generation (Fig. 1A). How LF treatment might induce PrP (106-126) resistance was studied by assessing the antioxidative properties and generation of ROS after treatment. Intracellular ROS production was spectrophotometrically measured by the DCFH-DA assay (Fig. 3A). After exposure to 100 μM PrP (106-126), DCF fluorescence intensity in SH-SY5Y cells increased significantly to 175% of the control value, whereas LF (200 μg/ml) or anti-oxidants (800 μM GSH or 4 mM NAC) led to a decrease in DCF fluorescence intensity (Fig. 3B). These results suggest that LF protects PrP (106-126)-induced neuronal cell death via the prevention of PrP (106-126)-induced ROS generation (Fig. 3C).

Figure 3. PrP (106-126)-induced ROS generation is suppressed by LF treatment. SH-SY5Y neuronal cells were pretreated with (A) LF or (B) anti-oxidant agents (800 μM GSN and 4 mM NAC), and then exposed to 100 μM PrP (106-126) for 12 h. The treated cells were used for measuring ROS release with the (A) DCFH-DA assay spectrophotometrically or (B) by flow cytometry *P<0.05 vs. control; **P<0.01 vs. PrP (106-126). (C) The treated cells or antioxidant agents (800 μM GSN and 4 mM NAC) were then exposed to 100 μM PrP (106-126) for 12 h. Cell viability was measured by the Annexin V assay via flow cytometry.

PrP (106-126)-induced mitochondrial dysfunction is suppressed by LF treatment

PrP (106-126)-induced apoptosis is mediated by mitochondrial disruption (12). Mitochondrial dysfunction occurs after apoptotic signals, including loss of MTP and release of apoptotic factors into the cytosol (24). We examined the effects of LF or antioxidants on PrP (106-126)-induced mitochondrial dysfunction. MTP was measured by flow cytometry. PrP (106-126)-treated cells showed increased JC-1 monomers, while LF pretreatment reduced PrP (106-126)-induced JC-1 monomers (Fig. 4A). Furthermore, pretreatment of antioxidants also reduced PrP (106-126)-induced JC-1 monomers. These results were confirmed by fluorescence microscopy images of JC-1 stained cells (Fig. 4B). Consistent with these results, LF-treatment cells prevented PrP (106-126)-induced cytochrome c release and Bax translocation (Fig. 4C).

Figure 4. LF prevents against PrP (106-126)-induced mitochondrial dysfunction via attenuation of Bax translocation. (A) SH-SY5Y cells were pretreated with LF or antioxidant agents (800 μM GSN and 4 mM NAC), and then exposed to 100 μM PrP (106-126) for 12 h. The treated cells were measured using the MTP assay, *P<0.05 vs. control; **P<0.01 vs. PrP (106-126). (B) The treated cells also were photographed using a fluoroscope. The JC-1 monomer fluorescence intensity (dark gray) was measured at 530 nm (emission of JC-1 monomeric form) when excited at 490 nm. (C) Cells were homogenized in a mitochondrial buffer, as analyzed by western blotting using antibodies against cytochrome c and Bax protein.

Discussion

Prion diseases are fatal neurodegenerative disorders (25). The main component of prion disease is the abnormal isoform of prion protein (PrPsc) (26). PrP (106-126) maintains the neurotoxic characteristics of the entire pathological PrPSc and is commonly used as a suitable model to study the mechanism of prion disorders (5). However, this peptide mechanism is not fully understood. In previous studies, it has been shown that PrP (106-126) induces neurotoxicity via mitochondrial disruption and ROS generation. LF is an 80 kDa protein. It is a multifunctional protein of the transferrin family and its functions include antimicrobial activity, antibacterial activity, cell proliferation, and antioxidant ability (27). LF protects from programmed cell death via antioxidant activity that is due to the scavenging of ROS (20). Moreover, LF inhibits PrPsc accumulation in scrapie-infected cells (22). However, the affirmative effect of LF on PrP (106-126)-induced neuronal cell death is not completely understood. In this study, LF treatment protected against PrP (106-126)-induced neuronal cell death (Fig. 1). In addition, PrPc-deficient mice were more sensitive to oxidative stress (28). Oxidative stress plays an important role in neurodegenerative disorders (13). Thus, we considered whether LF treatment could mediate ROS scavenger ability. Our results demonstrate that LF protects against PrP (106-126)-induced ROS generation in SH-SY5Y cells (Fig. 3A and B). These results suggest that PrP (106-126) mediates apoptotic cell death and ROS generation, and that these consequences are decreased by LF treatment. ROS can activate JNK protein. Indeed, PrP (106-126) induces neuronal cell damage by activating JNK and caspase-3 proteins (Fig. 2). JNK activation has been documented in neurodegenerative diseases (29). By contrast, LF treatment inhibits PrP (106-126)-mediated protein activation including JNK and caspase-3 (Fig. 2). These results indicate that LF treatment inhibits PrP (106-126)-mediated JNK and caspase-3 activation, and support the view that LF-mediated ROS scavenging downregulates PrP (106-126)-mediated protein activation. NAC protects cells against mitochondrial dysfunction (30). Furthermore, PrP (106-126)-induced apoptotic cell death occurs through mitochondrial disruption in neuronal cells (12). Our findings additionally show that LF or antioxidants (GSH and NAC) prevent neuronal cell death due to PrP (106-126)-mediated mitochondrial dysfunction (Fig. 4). Collectively, these results indicate that LF treatment protects from PrP (106-126)-induced neuronal cell death by ROS scavenging associated antioxidant activity. Moreover, LF possesses antioxidant activity and prevents PrP (106-126)-mediated mitochondrial disruption. In addition, these findings also suggest that LF may have clinical benefits when used for neurodegenerative chemotherapy such as in patients with prion disorders.

Acknowledgements

This study was supported by the Cooperative Research Program for Agriculture Science and Technology Development (PJ907116) in Rural Development Administration and by the National Research Foundation of the Korea Grant funded by the Korean Government (2010-E00019).

References