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Keywords:

  • apoptosis;
  • cadmium;
  • c-Jun N-terminal kinase;
  • extracellular signal-regulated kinase 1/2;
  • mammalian target of rapamycin;
  • rapamycin

Abstract

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

Cadmium (Cd) may be accumulated in human body through long-term exposure to Cd-polluted environment, resulting in neurodegeneration and other diseases. To study the mechanism of Cd-induced neurodegeneration, PC12 and SH-SY5Y cells were exposed to Cd. We observed that Cd-induced apoptosis in the cells in a time- and concentration-dependent manner. Cd rapidly activated the mitogen-activated protein kinases (MAPK) including extracellular signal-regulated kinase 1/2 (Erk1/2), c-Jun N-terminal kinase (JNK) and p38. Inhibition of Erk1/2 and JNK, but not p38, partially protected the cells from Cd-induced apoptosis. Consistently, over-expression of dominant negative c-Jun or down-regulation of Erk1/2, but not p38 MAPK, partially prevented Cd-induced apoptosis. To our surprise, Cd also activated mammalian target of rapamycin (mTOR)-mediated signaling pathways. Treatment with rapamycin, an mTOR inhibitor, blocked Cd-induced phosphorylation of S6K1 and eukaryotic initiation factor 4E binding protein 1, and markedly inhibited Cd-induced apoptosis. Down-regulation of mTOR by RNA interference also in part, rescued cells from Cd-induced death. These findings indicate that activation of the signaling network of MAPK and mTOR is associated with Cd-induced neuronal apoptosis. Our results strongly suggest that inhibitors of MAPK and mTOR may have a potential for prevention of Cd-induced neurodegeneration.

Abbreviations used
4E-BP1

eukaryotic initiation factor 4E binding protein 1

Akt

protein kinase B (PKB)

ASK1

apoptosis signal-regulating kinase 1

Cd

cadmium

DAPI

4′, 6-diamidino-2-phenylindole

DMEM

Dulbecco’s Modified Eagle’s Medium

Erk1/2

extracellular signal-regulated kinase 1/2

FBS

fetal bovine serum

FDA

fluorescein diacetate

JNK

c-Jun N-terminal kinase

MAPK

mitogen-activated protein kinase

MEK

mitogen extracellular kinase

MKK

MAPK kinase

mTOR

mammalian target of rapamycin

PARP

poly (ADP-ribose) polymerase

PBS

phosphate buffered saline

PDL

poly-d-lysine

PI

propidium iodide

PI3K

phosphatidylinositol 3′-kinase

ROS

reactive oxygen species

S6K1

S6 kinase 1

SDS

sodium dodecyl sulfate

Cadmium (Cd), one of the toxic heavy metals is mainly released from smelting and refining of metals and cigarette smoking, resulting in the pollution of water, air, and soil. Cd can be absorbed and accumulated in plants and animals, and thereby can be accumulated in human body either through direct exposure to Cd-contaminated environment or by food chain. Such accumulation contributes to carcinogenesis, immunodepression and neurodegeneration (Chuang et al., 2000; Lopez et al., 2003; Kim et al., 2005). Clinically, exposure to Cd not only causes pulmonary edema, respiratory tract irritation, renal dysfunction, anemia, osteoporosis, and cancer in humans (Satarug et al., 2000; Kim et al., 2005; Lau et al., 2006), but also severely affects the function of the nervous system (Lopez et al., 2003). For example, Cd induces neurological disorders, such as learning disabilities and hyperactivity in children (Pihl and Parkes, 1977; Marlowe et al., 1985). Workers exposed to Cd show olfactory dysfunction and neurobehavioral defects in attention, psychomotor speed, and memory (Kim et al., 2005). Cd-induced oxidative stress is closely associated with Parkinson’s disease and Alzheimer’s disease (Okuda et al., 1997; Johnson, 2001; Panayi et al., 2002).

Recent studies have shown that the sustained activation of mitogen-activated protein kinases (MAPKs) signaling pathways are involved in Cd-induced cell death (Chuang et al., 2000; Kim and Sharma, 2004; Kim et al., 2005; Lag et al., 2005; Miguel et al., 2005; Martin et al., 2006). To date, three MAPK family members, including extracellular signal-regulated protein kinase 1/2 (Erk1/2), c-Jun N-terminal kinase (JNK), and p38 MAPK have been well characterized. These MAPKs are regulated by the distinct stimuli (Kyriakis and Avruch, 2001; Pearson et al., 2001). Erk1/2 is predominantly activated by growth factors or mitogens leading to cell differentiation, growth, and survival, whereas JNK and p38 are preferentially activated by oxidative stress and cytokines resulting in inflammation and apoptosis (Fan and Chambers, 2001). However, in response to some stress stimuli, e.g., cisplatin, all three MAPK members can be activated (Fan and Chambers, 2001). Cd activates Erk1/2, JNK and/or p38, depending on cell types and the concentration of Cd tested (Chuang et al., 2000; Kim and Sharma, 2004; Kim et al., 2005; Lag et al., 2005; Miguel et al., 2005; Martin et al., 2006). Cd activates Erk1/2 in human embryonic kidney 293 cells, human liver cells, and human cervical cancer cells, rat primary embryo fibroblasts, new born rat osteoblasts and mouse bone-derived macrophages (Martin et al., 2006), Erk/JNK in murine macrophages (Chuang et al., 2000; Kim and Sharma, 2004; Kim et al., 2005), p38 in human lymphoma (U937) cells (Miguel et al., 2005), Erk/JNK/p38 in rat lung cells (Lag et al., 2005) and human non-small cell lung carcinoma cell line (CL3) (Chuang et al., 2000; Kim and Sharma, 2004; Kim et al., 2005). However, how Cd activates MAPK family members in neuronal cells remains to be defined. Also, whether Cd targets other signaling pathways responsible for neuronal cell survival is largely unknown.

The mammalian target of rapamycin (mTOR), a 289 kDa Ser/Thr kinase, lies downstream of phosphatidylinositol 3′-kinase (PI3K), and senses mitogenic stimuli, nutrient conditions (Fang et al., 2001; Kim et al., 2002) and ATP (Dennis et al., 2001). Activated PI3K or protein kinase B (PKB) (Akt) may positively regulate mTOR, leading to increased phosphorylation of ribosomal p70 S6 kinase (S6K1) and eukaryotic initiation factor 4E binding protein 1 (4E-BP1), the two best characterized downstream effector molecules of mTOR (Bjornsti and Houghton, 2004). mTOR has been widely recognized as a central controller for cell proliferation, growth and survival (Bjornsti and Houghton, 2004). Studies have shown that Cd potently inhibits cell proliferation/growth and survival of neuronal cells (Lopez et al., 2003; Kim et al., 2005; Lau et al., 2006). Of interest, among the proteins regulated by mTOR, a number of them, such as cyclin D1, eukaryotic initiation factor 4E, c-myc, nuclear factor κB, Akt, and protein kinase C are also deregulated by Cd (Othumpangat et al., 2005; Qu et al., 2005; Washington et al., 2006; Brama et al., 2007), suggesting that Cd may execute its cytotoxic effect by targeting mTOR signaling.

Here we show that Cd activates Erk1/2, JNK and p38 as well as the upstream kinases, such as apoptosis signal-regulating kinase 1 (ASK1), MAPK kinase (MKK4), MKK3/6, in neuronal (PC12 and SH-SY5Y) cells. Inhibition of Erk1/2 and JNK, but not p38, partially protected the cells from Cd-induced apoptosis. Unexpectedly, we found that in PC12 and SH-SY5Y cells, Cd also activates mTOR signaling, a critical pathway for survival in many other cell types. Of importance, pre-treatment with mTOR inhibitor, rapamycin, markedly prevented Cd-induced apoptosis. Over-expression of dominant negative c-Jun, down-regulation of Erk1/2 or mTOR by RNA interference also in part, rescued cells from Cd-induced death. These findings indicate that Cd-induced neuronal apoptosis is associated with the activation of the signaling network of MAPK and mTOR pathways.

Materials and methods

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

Materials

Cadmium chloride (Sigma, St Louis, MO, USA) was dissolved in distilled water to prepare the stock solutions (0–120 mmol/L), filtered through a 0.22 μm pore size membrane, aliquoted and stored at 22°C. Dulbecco’s modified Eagle’s medium (DMEM) was purchased from Mediatech (Herndon, VA, USA). Horse serum and fetal bovine serum (FBS) were supplied by Hyclone (Logan, UT, USA), whereas 0.05% Trypsin–EDTA was from Invitrogen (Grand Island, NY, USA). Enhanced chemiluminescence solution was from Perkin-Elmer Life Science (Boston, MA, USA). CellTiter 96® AQueous One Solution Cell Proliferation Assay kit was from Promega (Madison, WI, USA). The MAPK inhibitors, SB203580, U0126, and SP600125 were obtained from LC Laboratories (Woburn, MA, USA). Fluorescein diacetate (FDA) and propidium iodide (PI) were from Alfa Aesar (Ward Hill, MA, USA) and MP Biomedicals Inc. (Solon, OH, USA), respectively. The following antibodies were used: ASK1, phospho-ASK1 (Thr845), MKK4, phospho-MKK4 (Ser257/Thr261), phospho-mTOR (Ser2448), phospho-mTOR (Ser2481) (all from Cell Signaling Technology, Beverly, MA, USA), JNK1, phospho-JNK (Thr183/Tyr185), c-Jun, phospho-c-Jun (Ser63), Erk2, phospho-Erk1/2 (Thr202/Tyr204), p38, phospho-p38 (Thr180/Tyr182), mitogen extracellular kinase (MEK3/6), phospho-MEK3/6, phospho-S6K1 (Thr389), S6K1, phospho-Akt (Thr308), phospho-Akt (Ser473), Akt, FLAG (all from Santa Cruz Biotechnology, Santa Cruz, CA, USA), 4E-BP1 (Zymed Laboratories, South San Francisco, CA, USA), β-tubulin (Sigma), and mTOR (a gift from Dr Peter J. Houghton, St Jude Children’s Research Hospital, Memphis, TN, USA) (Zhang et al., 2002). Annexin V-FITC Apoptosis Detection Kit I was purchased from BD Biosciences (San Jose, CA, USA). Poly-d-lysine (PDL), 4′, 6-diamidino-2-phenylindole (DAPI) and all the other chemicals were purchased from Sigma.

Cell culture

Rat pheochromocytoma (PC12) and human neuroblastoma (SH-SY5Y) cell lines were from ATCC (Manassas, VA, USA). PC12 cells were grown in antibiotic-free DMEM supplemented with 10% horse serum and 5% FBS, whereas SH-SY5Y cells were grown in antibiotic-free DMEM supplemented with 10% FBS. Cells were maintained in a humid incubator (37°C, 5% CO2).

One solution assay

Cells were seeded at a density of 1 × 10cells/well in a flat-bottomed 96-well plate, pre-coated with (for PC12) or without (for SH-SY5Y) PDL (0.2 μg/mL). Next day, cells were treated with various concentrations of Cd (0–120 μmol/L) for 24 h, or with 20 μmol/L Cd for different time (0–24 h) with 6–12 replicates of each treatment. After incubation, each well was added 20 μL of one solution reagent (Promega) and incubated for 4 h. Cell viability was determined by measuring the optical density at 490 nm using a Wallac 1420 Multilabel Counter (Perkin-Elmer Life Sciences, Wellesley, MA, USA).

Cell morphological analysis

Cells were seeded at a density of 1 × 106 cells/well in a six-well plate, pre-coated with (for PC12) or without (for SH-SY5Y) PDL (0.2 μg/mL). Next day, Cd (0–40 μmol/L) was added. After incubation for 24 h, images were taken with an Olympus inverted phase-contrast microscope (Olympus Optical Co., Melville, NY, USA) (200×) equipped with the Quick Imaging system.

Fluorescein diacetate–propidium iodide staining

Cell viability was also evaluated using FDA–PI staining (Jones and Senft, 1985). Briefly, a stock solution of FDA was made by dissolving 10 mg of FDA in 1 mL of acetone. A working solution was prepared by adding 0.04 mL of stock solution to 10 mL of phosphate buffered saline (PBS) at pH 7.0. A stock solution of PI was made by dissolving 0.5 mg of PI in 50 mL of PBS. After treatment, cells were trypsinized, centrifuged at 250 g for 5 min, resuspended in 200 μL of PBS (at a final concentration of 1 × 107 cells/mL), and stained with 0.1 mL of FDA working solution and 0.03 mL of PI for 5 min. A drop of cell suspension was applied to a slide and allowed to air-dry. Photographs were taken with a Nikon Eclipse TE300 fluorescence microscope (Nikon Instruments Inc., Melville, NY, USA) equipped with a digital camera at excitation wavelengths of 490 nm for FDA and 545 nm for PI. At least 500 viable (green)/non-viable (red) cells were counted for each treatment.

DAPI staining

Cells were seeded at a density of 5 × 105 cells/well in a six-well plate containing a PDL-coated glass coverslip per well. Next day, Cd was added. After treatment with CdCl2 (0–40 μmol/L) for 24 h, cells were fixed with 4% paraformaldehyde prepared in PBS for 2 h at 4°C. The cells were washed three times with PBS, and then stained with DAPI (4 μg/mL in deionized water) for 30 min at 22°C in the dark. Following a brief washing with PBS, slides were mounted in glycerol/PBS (1 : 1, v/v) containing 2.5% 1,4-diazabiclo-(2,2,2)octane. Photographs were taken with a Nikon Eclipse TE300 fluorescence microscope equipped with a digital camera. Cells with condensed nuclei were scored to be apoptotic.

Flow cytometry

Cells were seeded in 100-mm dishes, pre-coated with (for PC12) or without (for SH-SY5Y) PDL, at a density of 2 × 106 cells/dish in completed growth medium. Next day, cells were exposed to CdCl2 (0–80 μmol/L) for 24 h, followed by apoptosis assay using the Annexin V-FITC Apoptosis Detection Kit I (BD Biosciences), as described (Beevers et al., 2006).

Adenoviral infection of cells

Recombinant adenovirus encoding FLAG epitope-tagged dominant negative c-Jun (FLAG-Δ169) (Ad-dn-c-Jun) was a gift from Dr Jonathan Whitfield (Eisai London Research Laboratories, University College London, London, UK) (Whitfield et al., 2001). The virus was amplified and titrated as described previously (Huang et al., 2003). For experiments, PC12 and SH-SY5Y cells were grown in six-well plates in the corresponding medium, and infected with the Ad-dn-c-Jun for 24 h at the multiplicity of infection of 5. Subsequently, cells were treated with Cd (0–20 μmol/L) for ca. 24 h. Ad-GFP encoding the green fluorescence protein (GFP) (Liu et al., 2006) served as a control. Expression of FLAG-tagged dn-c-Jun was confirmed by western blot with antibodies to FLAG.

Lentiviral shRNA cloning, production, and infection

To generate lentiviral shRNAs to Erk1/2 and p38 MAPK, oligonucleotides containing the target sequences were synthesized, annealed and inserted into FSIPPW lentiviral vector (Kanellopoulou et al., 2005) via the EcoR1/BamH1 restriction enzyme site. Oligonucleiotides used were: Erk1 sense: 5′-AATTCCCGACCTGCTGGACCGGATGTTATGCAAGAGATAACATCCGGTCCAGCAGGTC TTTTTG-3′, anti-sense: 5′-GATCCAAAAAGACCTGCTGGACCGGATGTTATCTCTTGCATAACATCCGGTCCAGCAGGTCGGG-3′; Erk2 sense: 5′-AATTCCCGATTCCAGCCAGGATACAGATTGCAAGAGAATCTGTATCCTGGCTGGAATCTTTTTG-3′, anti-sense: 5′-GATCCAAAAAGATTCCAGCCAGGATACAGATTCTCTTGCAATCTGTATCCTGGCTGGAATCGGG-3′; p38 sense: 5′-AATTCCCGACTGTGAGCTGAAGATTCTGCAAGAGAGAATCTTCAGCTCACAGTCTTTTTG-3′, antisense: 5′-GATCCAAAAAGACTGTGAGCTGAAGATTCTCTCTTGCAGAATCTTCAGCTCACAGTCGGG-3′. Lentiviral shRNA constructs targeting mTOR and green fluorescence protein (control) were described (Liu et al., 2006). To produce lentiviral shRNAs, above constructs were co-transfected together with pMD2G and psPAX2 (Addgene, Cambridge, MA, USA) to 293TD cells using LipfectamineTM 2000 reagent (Invitrogen, Carlsbad, CA, USA). Each virus-containing medium was collected 36 and 60 h post-transfection, respectively. For use, monolayer cells, when grown to about 70% confluence were infected with above lentivirus-containing supernatant in the presence of 8 μg/mL polybrene for 12 h twice at an interval of 6 h. Uninfected cells were eliminated by exposure to 2 μg/mL puromycin for 48 h before use.

Western blot analysis

After treatment, cells were briefly washed with cold PBS. On ice, cells were lysed in the radioimmunoprecipitation assay buffer [50 mmol/L Tris, pH 7.2; 150 mmol/L NaCl; 1% sodium deoxycholate; 0.1% sodium dodecyl sulfate (SDS); 1% Triton X-100; 10 mmol/L NaF; 1 mmol/L Na3VO4; protease inhibitor cocktail (1 : 1000, Sigma)]. Lysates were sonicated for 10 s and centrifuged at 16 000 g for 10 min at 4°C. Protein concentration was determined by bicinchoninic acid assay with bovine serum albumin as standard (Pierce, Rockford, IL, USA). Equivalent amounts of protein were separated on 7.5–12% SDS–polyacrylamide gel and transferred to polyvinylidene difluoride membranes (Millipore, Bedford, MA, USA). Membranes were incubated with PBS containing 0.05% Tween 20 and 5% non-fat dry milk to block non-specific binding and were incubated with primary antibodies, then with appropriate secondary antibodies conjugated to horseradish peroxidase. Immunoreactive bands were visualized by using Renaissance chemiluminescence reagent (Perkin-Elmer Life Science). To check the amount of protein loaded, the immunoblots were treated with stripping solution (62.5 mmol/L Tris buffer, pH 6.7, containing 2% SDS and 100 mmol/L β-mercaptoethanol) for 30 min at 50°C and incubated with mouse monoclonal anti-β-tubulin antibody (Sigma) followed by horseradish peroxidase-coupled goat anti-mouse IgG (Pierce).

Statistical analysis

Results were expressed as mean values ± SE. Statistical analysis was performed by Student’s t-test (statistica, Statsoft Inc., Tulsa, OK, USA). A level of < 0.05 was considered to be significant.

Results

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

Cd reduced viability and altered morphology of neuronal cells

PC12 and SH-SY5Y, the two neuronal cell lines were chosen as a model to study the mechanism by which Cd-induced apoptosis of neuronal cells. To find an appropriate concentration and treatment time of Cd for the mechanism studies, we first performed cell viability assays. As shown in Fig. 1a, treatment with Cd for 24 h resulted in a concentration-dependent decrease of PC12 cell viability. At 20 μmol/L, Cd reduced the cell viability by ca. 50%, compared to the vehicle control. A time-dependent decline in cell viability occurred during the 24 h period (Fig. 1b). Starting from 1 h treatment, Cd (20 μmol/L) significantly decreased cell viability. By phase-contrast microscopic observation, more cells became round or shrunken, when exposed to increasing concentrations of Cd (2.5–40 μmol/L) (Fig. 1c). In comparison with PC12, SH-SY5Y cells appeared to be more sensitive to Cd. When exposed to 10 μmol/L of Cd for 12 h, ca. 60% of PC12 cells remained viable, whereas only about 10% of SH-SY5Y cells were viable (Fig. 6b). The data suggest that Cd may induce remarkable apoptosis of the neuronal cells at concentrations of > 10 μmol/L within 24 h.

image

Figure 1.  Cadmium (Cd) reduced viability and altered morphology of neuronal cells. Cell viability of PC12 cells treated with different concentrations of Cd for 24 h (a), or treated with 20 μmol/L Cd for various time (b) was evaluated using one solution assay. Morphology of PC12 cells treated with different concentrations of Cd for 24 h (c) was assessed using an Olympus inverted phase-contrast microscope (200×) equipped with Quick Imaging system. Scale bar: 100 μm. Results are presented as mean ± SE, = 6–12. *< 0.05, **< 0.01, difference with control group.

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image

Figure 6.  Inhibition of mammalian target of rapamycin (mTOR) markedly reduced Cadmium (Cd)-induced neuronal cell apoptosis. (a) Following the fluorescein diacetate (FDA)–propidium iodide (PI) staining, viable (green)/non-viable (red) PC12 cells treated with/without Cd for 12 h following pre-incubation with rapamycin (Rap) for 48 h were visualized under a fluorescence microscope. (i) Control, (ii) 0.2 μg/mL Rap, (iii) and (iv) 10 and 20 μmol/L Cd, respectively, (v) and (vi) 0.2 μg/mL Rap plus 10 and 20 μmol/L Cd, respectively. Scale bar: 50 μm. (b) Percentage of viable SH-SY5Y and PC12 cells was obtained, following FDA–PI staining. Results are presented as mean ± SE, = 3–6. a< 0.05, b< 0.01, difference with control group; c< 0.01, difference with 10 μmol/L Cd group; d< 0.01, difference with 20 μmol/L Cd group. (c) PC12 cells were co-treated with Cd and Rap for an indicated concentration and time, followed by western blot analysis with antibodies against S6K1 (Thr389), S6K1, eukaryotic initiation factor 4E binding protein 1, and β-tubulin (loading control). (d) Down-regulation of mTOR (by ca. 90%) by lentiviral shRNA to mTOR in SH-SY5Y cells, as detected by western blot with antibodies to mTOR (upper panel), partially prevented Cd-induced cell death (bottom panel).

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Cd-induced apoptosis of neuronal cells

Nuclear fragmentation and condensation are one of major events, when cells are undergoing apoptosis. To test whether the loss of cell viability was because of Cd-induced apoptosis, nuclear staining with DAPI was used. We found that treatment with Cd for 24 h increased nuclear fragmentation and condensation (arrows) in a concentration-dependent manner (Fig. 2a and b). Consistently, significant reduction of cell number was observed at higher concentrations (> 10 μmol/L) of Cd (Fig. 2a).

image

Figure 2.  Cadmium (Cd)-induced apoptosis of neuronal cells. (a) PC12 cells exposed to 0–40 μmol/L Cd for 24 h, displayed nuclear condensation (arrows) and reduction in cell number via 4′,6-diamidino-2-phenylindole staining. (b) Cd increased percentage of cells with fragmented nuclei in a concentration-dependent manner. Scale bar: 20 μm. (c) Cd-induced apoptosis, as determined by annexin-V-FITC and propidium iodide staining followed by the fluorescence-activated cell sorting using flow cytometry. Results are presented as mean ± SE, = 3–5. *< 0.05, **< 0.01, difference with control group.

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To further quantify the extent of apoptosis in a larger cell population, we performed annexin-V-FITC and PI staining, followed by the fluorescence-activated cell sorting using flow cytometry. As shown in Fig. 2c, exposure of PC12 to Cd for 24 h induced apoptosis in a concentration-dependent manner, which is in agreement with the DAPI staining results. Significant increase in apoptosis occurred at concentrations of > 10 μmol/L (Fig. 2c). At 20 μmol/L, Cd increased apoptosis by ca. 2.5-fold. It appears that apoptosis was the major cause for Cd-reduced-cell viability.

Cd-induced neuronal cell apoptosis was associated with activation of MAPK pathways

Previous studies have demonstrated that Cd may trigger cell death by activation of Erk1/2, JNK and/or p38, depending on the cell types and the concentration of Cd tested (Chuang et al., 2000; Kim and Sharma, 2004; Kim et al., 2005; Lag et al., 2005; Miguel et al., 2005; Martin et al., 2006). Recently, Kim et al. (2005) found that Cd activates JNK and p38, but not Erk1/2, in SH-SY5Y cells, and identified that Cd-induced apoptosis of the neuronal cells occurs through activation of the ASK1/MKK4/JNK/c-Jun signaling pathway (Kim et al., 2005). To examine whether this is a cell-type context, we determined phosphorylation of Erk1/2, JNK, and p38 in PC12, as well as in SH-SY5Y cells (for control). Our western blot analysis shows that treatment of PC12 cells with Cd for 24 h resulted in robust phosphorylation of Erk1/2, JNK, and p38 starting at 5 μmol/L (Fig. 3a and b). This is further supported by the findings that Cd also activated their upstream kinases, such as ASK1, MKK4, and MKK3/6, as detected by phospho-specific antibodies (Fig. 3a and b). We noticed that Cd-activation of JNK resulted in 10-fold increase in protein expression and phosphorylation of c-Jun (Fig. 3a). Similar results were also seen in SH-SY5Y cells (Fig. 3e–g).

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Figure 3.  Cadmium (Cd) rapidly activated mitogen-activated protein kinase (MAPK) cascade. PC12 cells treated with 0–20 μmol/L Cd for 24 h, or with 20 μmol/L Cd for 0–12 h were harvested and total lysates were subjected to western blot analysis using antibodies against the indicated proteins. The blots were probed for β-tubulin as a loading control. Similar results were observed in at least three independent experiments. Cd activated c-Jun N-terminal kinase (JNK), extracellular signal-regulated kinase 1/2 (Erk1/2), and p38 as well as the upstream apoptosis signal-regulating kinase 1, MAPK kinase 4 (MKK4), and MKK3/6 in a concentration-dependent (a and b) and time-dependent manner (c and d). Inhibitors suppressed Cd-activated phosphorylation of MAPKs (e–g). SH-SY5Y and PC12 cells were treated for 4 h with/without Cd (10, 20 μmol/L) following pre-incubation with 10–40 μmol/L JNK inhibitor SP600125, 5 μmol/L mitogen extracellular kinase 1/2 (upstream of Erk1/2) inhibitor U0126, or 10 μmol/L p38 inhibitor SB203580, for 30 min, respectively. Western blot analysis was performed using indicated antibodies.

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As Cd dramatically induced apoptosis of the PC12 and SH-SY5Y cells at 20 μmol/L within 24 h (Fig. 2), we next tested whether Cd also activates the MAPKs under this condition. As shown in Fig. 3c and d, within 4–6 h, Cd obviously increased phosphorylation of Erk1/2, JNK and p38, and this increased phosphorylation was sustained for over 12 h. Consistently, high level of c-Jun and phospho-c-Jun was induced.

To dissect the role of activation of Erk1/2, JNK, and p38 in Cd-induced neuronal apoptosis, PC12 and SH-SY5Y cells were exposed to Cd (10, 20 μmol/L) for 4 h after pre-treatment with MEK1/2 (upstream of Erk1/2) inhibitor U0126, JNK inhibitor SP600125, or p38 inhibitor SB203580 for 30 min, respectively. As shown in Fig. 3e, Cd-induced phosphorylation of c-Jun, as readout of JNK activity was obviously attenuated by SP600125 (40 μmol/L). Similarly, U0126 (5 μmol/L) and SP203580 (10 μmol/L) blocked Cd-induced phosphorylation of Erk/12 and p38, respectively (Fig. 3f and g). As these inhibitors could inhibit MAPKs phosphorylation induced by Cd at 10 μmol/L, we next studied whether the individual inhibitor could prevent Cd-induced apoptosis of the neuronal cells. To this end, PC12 cells were pre-treated with each inhibitor for 30 min, followed by exposure to Cd (10 μmol/L) for 24 h. Morphological analysis reveals that SP600125, SB203580 or U0126 alone did not obviously alter cell viability. However, SP600125 or U0126 partially rescued cells from Cd-induced apoptosis (Fig. 4a). The protective effect of SB203580 was undetectable (data not shown). The results suggest that Cd-induced apoptosis of the neuronal cells at least partially by activation of JNK and Erk1/2. The findings were confirmed by gene silencing or gene over-expression experiments. Infection of PC12 (right panel, Fig. 4b) or SH-SY5Y cells (data not shown) with Ad-dn-c-Jun resulted in expression of a dominant negative c-Jun, as detected by western blotting. Expression of the dominant negative c-Jun partially protected PC12 (left panel, Fig. 4b) or SH-SY5Y cells (data not shown) from apoptosis induced by Cd. Infection of PC12 (upper panel, Fig. 4c) or SH-SY5Y cells (data not shown) with lentiviral shRNAs to Erk1/2 and p38 MAPK, respectively, down-regulated expression of these proteins by ca. 90%. Silencing of expression of Erk1/2, but not p38 MAPK, in part reduced Cd-induced cell death in PC12 (Fig. 4c) or SH-SY5Y cells (data not shown).

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Figure 4.  Inhibition of mitogen-activated protein kinases (MAPKs) partially prevented Cadmium (Cd)-induced apoptosis of neuronal cells. (a) PC12 cells were treated with Cd (10 μmol/L) for 24 h following pre-incubation with each inhibitor for 30 min. Representative pictures show that SP600125 and U0126 partially rescued cells from Cd-induced apoptosis. (b) Over-expression of FLAG-tagged dominant negative c-Jun by infection of PC12 cells with Ad-dn-c-Jun, as detected by western blot with antibodies to FLAG (right panel), partially rescued cells from death induced by Cd (0–20 μmol/L, 24 h) (left panel). (c) Down-regulation of extracellular signal-regulated kinase 1/2 (Erk1/2; by ca. 90%), but not p38 MAPK (by ca. 90%) in PC12 cells, by lentiviral shRNAs to Erk1/2 and p38 MAPK, respectively, as detected by western blot with antibodies to Erk1/2 and p38 MAPK (upper panel), in part, protected the cells from Cd-induced death (bottom panel). Cell viability in (a), (b), and (c) was evaluated via morphological observation using an Olympus inverted phase-contrast microscope (200×) equipped with Quick Imaging system. Scale bar: 100 μm.

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Cd-induced neuronal cell apoptosis was associated with activation of Akt/mTOR signaling

Akt/mTOR signaling is crucial not only for cell proliferation/growth, but also for cell survival (Bjornsti and Houghton, 2004). We therefore hypothesized that Cd may inhibit Akt/mTOR signaling, leading to apoptosis of the neuronal cells. However, to our surprise, when PC12 cells were treated with different concentrations of CdCl2 (0–20 μmol/L) for 24 h, or with 20 μmol/L CdCl2 for 0–12 h, a concentration- and time-dependent increase of phosphorylation of Akt was detected by western blot analysis (Fig. 5a and b). Cd-induced activation of Akt started at 2 h and sustained for > 12 h (Fig. 5b). Similarly, we found that Cd also enhanced phosphorylation of mTOR and its downstream effector molecules, S6K1 and 4E-BP1 (Fig. 5a and b). Cd did not obviously alter total protein levels of those proteins (Fig. 5a and b). Similar data were observed in SH-SY5Y cells (data not shown). It should be mentioned that a considerable basal level of phosphorylation of 4E-BP1 was detected (Figs 5a, b and 6c) in PC12 or SH-SY5Y cells. Phosphorylation state of 4E-BP1 was detected with an antibody to 4E-BP1. Phosphorylation of 4E-BP1 decreases its electrophoretic mobility during SDS–polyacrylamide gel electrophoresis (Brunn et al., 1997). As shown in Fig. 5a, Cd increased phosphorylation of 4E-BP1 in a concentration-dependent manner, as indicated by the increase in the intensity of the uppermost band γ and by the decrease in the higher mobility band α and β that corresponds to a less phosphorylated form of 4E-BP1. The findings clearly indicate that Cd activated Akt/mTOR signaling pathways in the neuronal cells.

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Figure 5.  Cadmium (Cd) activated mammalian target of rapamycin-mediated signaling pathways. PC12 cells treated with 0–20 μmol/L Cd for 24 h (a), or with 20 μmol/L Cd for 0–12 h (b) were harvested and total lysates were subjected to western blot analysis using antibodies against the indicated proteins.

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To determine whether Cd-activation of mTOR signaling is related to neuronal apoptosis, PC12 and SH-SY5Y cells were pre-treated with rapamycin (0.2 μg/mL), an inhibitor of mTOR, for 48 h, and then exposed to Cd (10, 20 μmol/L) for 12 h, followed by FDA–PI staining. We observed that rapamycin alone did not apparently alter cell viability. However, rapamycin markedly prevented Cd-induced apoptosis of the cells (Fig. 6a and b). Furthermore, using western blotting, we confirmed that the Cd-activated phosphorylation of 4E-BP1 and S6K1 was severely blocked by rapamycin (Fig. 6c). The role of mTOR in Cd-induced cell death was also confirmed by RNA interference. As shown in Fig. 6d, down-regulation of mTOR partially prevented Cd-induced apoptosis of SH-SY5Y cells as well. Therefore, our data clearly indicate that Cd may induce apoptosis of the neuronal cells at least in part, through activation of Akt/mTOR signaling pathway. Inhibition of mTOR by rapamycin may exert a beneficial effect against Cd-induced neuronal cell death.

Discussion

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

Cadmium pollution in the environment may be accumulated in human body through direct exposure or food chain, resulting in neurodegeneration as well as many other diseases. Cd may induce Parkinson’s disease, Alzheimer’s disease, and other neurodegenerative disorders primarily by triggering neuronal cell death. However, the underlying mechanism remains to be determined. Here, we show for the first time that Cd-induced apoptosis is associated not only with activation of MAPK cascade, but also with activation of mTOR-mediated signaling pathway. Previous studies have demonstrated that sustained activation of JNK, Erk1/2 and/or p38 is responsible for Cd-induced apoptosis in various cells, including neuronal (SH-SY5Y) cells (Chuang et al., 2000; Hung et al., 1998; Rockwell et al., 2004; Kim et al., 2005). Therefore, it is not surprising that Cd activated JNK, Erk1/2 and p38 in PC12 and SH-SY5Y cells, and the inhibitors of those MAPKs partially prevented Cd-induced apoptosis of the cells. mTOR has been widely recognized as a central and positive controller for cell proliferation/growth and survival (Bjornsti and Houghton, 2004). Originally, we hypothesized that Cd might suppress mTOR signaling, causing neuronal cell death. However, to our surprise, exposure of the neuronal cells to Cd increased phosphorylation of Akt, mTOR and its downstream effector molecules, such as S6K1 and 4E-BP1. Of importance is that pre-treatment with an mTOR inhibitor, rapamycin, blocked Cd-increased phosphorylation of S6K1 and 4E-BP1, and markedly prevented Cd-induced apoptosis of the neuronal cells. Our findings clearly indicate that activation of mTOR signaling also contributes to Cd-induced apoptosis.

Here we show that Cd activated JNK, Erk1/2, and p38, and both JNK and Erk1/2, but not p38, participated in Cd-induced apoptosis in PC12 and SH-SY5Y cells. This is evidenced by our findings: (i) SP600125 (JNK inhibitor) and U0126 (Erk1/2 inhibitor, actually MEK1/2 inhibitor), but not SB203580 (p38 MAPK inhibitor), partially prevented Cd-induced apoptosis of the neuronal cells, (ii) expression of dominant negative c-Jun in part rescued cells from Cd-induced death, (iii) down-regulation of Erk1/2, but not p38 MAPK, partly protected the cells from apoptosis induced by Cd. The data are consistent with the recent findings that p38 is not involved in Cd-induced apoptosis in SH-SY5Y cells (Kim et al., 2005). However, Kim et al. (2005) failed to see Cd-activation of Erk1/2 in SH-SY5Y cells, which is in contrast to our results. Probably this is because of the difference in experimental conditions or cell phenotypes used. Comparing with PC12 cells, SH-SY5Y cells exhibited a higher level of phosphorylation of Erk1/2 under normal growth medium (Fig. 3f). However, this phosphorylation was considerably weak (Fig. 3f). Kim et al. (2005) claimed that there existed a high level of phosphorylation of Erk1/2 in untreated SH-SY5Y cells, and this phosphorylation remained unchanged in response to Cd treatment. If the phosphorylation of Erk1/2 reaches a maximal level, Cd may not induce more phosphorylation in those SH-SY5Y cells.

We also noticed that Cd-induced expression of c-Jun protein, but the total levels of other proteins remained unchanged in PC12 and SH-SY5Y cells (Fig. 3a, b). This is in agreement with the previous findings in CL3 (Chuang et al., 2000). Though Cd induced c-Jun mRNA expression, it did not alter protein levels of JNK, Erk and p38 MAPK in CL3 cells (Chuang et al., 2000). It appears that Cd can specifically induce expression of some genes, at least including c-Jun, c-fos and c-myc (Chuang et al., 2000; Yu et al., 2007), though the underlying mechanism is not clear.

In the studies, we also observed that Cd increased proteolytic cleavages of caspase 3 and poly (ADP-ribose) polymerase (PARP) in PC12 cells in a concentration- and time-dependent manner (data not shown). Pre-treatment of the cells with a broad spectrum caspase inhibitor, Z-VAD-FMK did prevent cleavage of caspase 3 or PARP, but only partially prevented Cd-induced apoptosis (data not shown), suggesting that Cd-induced apoptosis in the neuronal cells in a caspase-dependent and -independent manner. Similar findings have been reported in other types of cells, such as lymphoblastoid cells (Coutant et al., 2006) and human embyronic kidney 293 cells (Mao et al., 2007). We also found that U0126 (5 μmol/L), SP600125 (20 μmol/L) and rapamycin (100 ng/mL), but not SP203580 (10 μmol/L) (data not shown), partially inhibited cleavage of caspase 3 or PARP. This may be related to their partial protection against Cd-induced neuronal cell death.

It has been reported that several heavy metals, such as arsenic, nickel, and zinc, can activate PI3K/Akt or mTOR signaling (Kim et al., 2000; An et al., 2005; Yoon et al., 2006). This is consistent with our findings that Cd activates Akt/mTOR pathway. However, so far, little is known about how those heavy metals activate Akt or mTOR signaling. Cd is a well-known inducer of reactive oxygen species (ROS) generation in cells (see reviews by Filipic et al., 2006; Valko et al., 2006). In our studies, we also detected ROS generation induced by Cd in a time- and dose-dependent manner in PC12 and SH-SY5Y cells (data not shown). Activation of mTOR signaling by ROS has been described (Bae et al., 1999; Huang et al., 2002; Jung et al., 2003). Treatment of cells with hydrogen peroxide (H2O2) or its inducers, such as arsenite and ultraviolet irradiation, increased phosphorylation of S6K1, which could be blocked by the anti-oxidant N-acetyl-L-cystein or catalase (a specific H2O2 scavenger) (Bae et al., 1999; Huang et al., 2002; Jung et al., 2003). In this study, we observed that treatment with N-acetyl-L-cystein also prevented Cd-induced activation of MAPKs and mTOR pathways (data not shown). Thus, we tentatively conclude that Cd may activate mTOR (including MAPKs) pathway through induction of ROS generation in the neuronal cells. Undoubtedly, more studies are needed to address this issue. Our data indicate that Cd-induced apoptosis can be in part prevented by SP600125, U0126, or rapamycin, which suggests that the inhibitors of JNK, Erk1/2 and mTOR may have preventive effect against the neurodegeneration induced by Cd.

In conclusion, we have identified that Cd induces apoptosis of neuronal cells by activation of JNK, Erk1/2, and mTOR signaling network. Our findings support the notion that inhibitors of these pathways may be exploited for prevention of Cd-induced Parkinson’s disease, Alzheimer’s disease, and other neurodegenerative disorders.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

We thank Drs Peter J. Houghton and Jonathan Whitfield for generously providing antibodies and recombinant adenoviral constructs. This work was supported in part by Stiles Award (SH), Feist-Weiller Cancer Research Award (SH) and Start-up Fund (SH) jointly from Louisiana State University Health Sciences Center in Shreveport, LA.

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  6. Acknowledgements
  7. References
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