PGC-1α controls mitochondrial biogenesis and dynamics in lead-induced neurotoxicity

Exposure to lead induces mitochondrial dysfunction, energy depletion and subsequent apoptosis of dopaminergic neurons

N27 cells, derived from rat mesencephalon, were used to evaluate the effect of lead on mitochondrial function. Cells were treated with a range of lead (II) acetate concentrations over a period of 48 hours. Cells stained with MitoSOX and analyzed by flow cytometry revealed a 25% increase in mitochondrial superoxide at 5 μM and 100 μM concentrations and almost a 100% increase at 500 μM (Fig. 1A). DiOC₆(3), a selective mitochondrial dye, and propidium iodide (PI) staining revealed that lead causes mitochondrial membrane depolarization and neuronal cell death, which was much more marked at 500 μM than at 5 or 100 μM (Fig. 1B, C). PARP cleavage analysis (Fig. 1D) confirmed this, with the effect being far more pronounced at the highest lead concentration. At 5μM and 100μM, effects on mitochondrial dysfunction and cell death were similar (Fig. 1A-D). Apoptotic cell death was confirmed by an increased activation of caspases3/7 and caspase 8 in N27 cells exposed to 100μM and 500μM lead (Supplementary Figure1-S-1, A-a and b). Morphological characteristics of apoptotic cells, including condensed nuclei, were also evident in cells treated with 500μM lead (Supplementary Figure1-S-1, B and C). Morphological changes in the cells were apparent only at higher concentrations than changes in molecular markers of apoptosis, most likely reflecting the temporal engagement of the respective hallmarks of apoptosis.

Figure 1. Exposure to lead induces mitochondrial dysfunction and subsequent apoptosis of N27 dopaminergic neurons. Cells were incubated for 48-hours with lead acetate, at the concentrations shown. (A) Mitochondrial superoxide levels were measured using MitoSOX staining followed by FACS analysis. (B) The effect of lead treatment on mitochondrial membrane potential (MMP) loss was detected by DiOC₆(3). (C) Analysis of cell death measured by propidium iodide (PI). (D) Western blot analysis of PARP cleavage (PARP to PARPc) following lead exposure. Profiles of different parameters of mitochondrial function (E-I) were determined using Seahorse XF24 Analyser. *P<0.05, **P<0.01, ***P<0.001, n=3; mean ± SE.

Mitochondrial function was assessed by Seahorse XF24 analysis allowing for the measurement of a number of parameters including mitochondrial basal respiration, ATP production, oxygen consumption rate (OCR), maximum respiratory capacity and mitochondrial spare respiratory capacity. While a reduction in basal respiration was observed in cells treated with 100 and 500 μM lead (Fig. 1E), this was not statistically significant. There was a concentration-dependent decrease in ATP production (Fig. 1F), with the highest lead concentration causing a statistically significant (P<0.01) decrease of more than 50%. Proton leak was not significantly affected by lead treatment (Fig. 1G). Maximal respiration decreased in a concentration-dependent manner (Fig. 1H). Interestingly, mitochondrial reserve capacity levels were very low across all of the lead-treated cells (P<0.01) including 5 μM, a concentration which did not have a significant impact on other aspects of mitochondrial function investigated with the Seahorse XF24 compared with control (Fig. 1I).

PGC1α protects DA neurons from neurotoxic insults from lead

Real-time QPCR and protein expression analyses were performed to investigate the expression of Pgc1α and its downstream targets, which regulate mitochondrial biogenesis, in lead-treated N27 cells. Interestingly, only the 100 μM treatment with lead for 48 h caused a significant increase of Pgc1α expression, at both the RNA and protein levels (Fig. 2A, 2B). Downstream targets of PGC1α, Tfam and Nrf1 showed a similar pattern of expression, with changes apparent only at 100 μM lead (Fig. 2C). In order to determine whether the highest lead concentration affects Pgc1α expression at an early exposure time point, Pgc1α mRNA levels were investigated after a 3-hour lead exposure. Surprisingly, the expression of Pgc1α mRNA levels was markedly upregulated 3-hour post-treatment, with a ten-fold increase observed at 500 μM (Fig. 2D).

Figure 2. PGC1α protects N27 dopaminergic neuronal cells from Pb²⁺-induced neurotoxicity. Cells were incubated with lead acetate at the concentrations shown. (A,B) qRT-PCR and western blotting measurement of mRNA and protein expression levels, respectively of Pgc1α in N27 cells treated with lead for 48-hours. (C) mRNA levels of Pgc1α target genes (Tfam, Nrf1)inN27 cells treated with lead for 48-hours.(D)N27 Pgc1αmRNA levels after 3-hours lead treatment.(E) Pgc1α mRNA levels and (F) colony formation in N27 cells with stable downregulation of PGC1α. (G) Mitochondrial superoxide levels were measured in shPGC1α cells using MitoSOX staining and (H) the effect of lead treatment on mitochondrial membrane potential loss in shPGC1α was detected by DiOC₆(3) in N27 cells treated with lead for 48-hours. (I,K) Confocal imaging analysis of mitochondrial morphology in wild type PGC1α and in overexpressed PGC1α N27 cells ((Pgc1α) was investigated using Mito Tracker and DAPI staining recorded by fluorescence microscopy. (J) qRT-PCR analysis of Pgc1α in N27 cells expressing exogenous Pgc1α versus(Pgc1α cells, treated with lead for 48-hours.*P<0.05, **P<0.01, n=3; mean ± SE.

Since no significant difference was observed between cells treated with 5 μM and 100 μM in terms of disruption of mitochondrial function, ROS generation or cell death(Fig. 1A-D), but there was a significant difference in Pgc1α mRNA levels between the two groups, we sought to investigate whether Pgc1α plays a protective role against lead insult at 100 μM. To interrogate this hypothesis, we produced a stable knock-down of Pgc1α using a lentiviral shRNA vector. Successful knock-down was confirmed by Pgc1α mRNA levels(Fig. 2E) and clear downregulation of PGC-1α protein levels, as shown by immunocyto-chemistry (Supplementary Figure3-S3, A). sh-PGC1α transduction did not affect colony formation (Fig. 2E and F). FACS analysis of cells stained independently with MitoSOX and DiOC₆(3) showed that reduction of Pgc1α expression renders N27 cells more susceptible to lead-induced mitochondrial stress (Fig. 2G, H), with morphological characteristics of apoptosis apparent in cells treated with 100μM or 500μM lead (Supplementary Figure1-S1, B).

The influence of lead treatment on mitochondrial organisation was investigated by immunofluorescence (Fig. 2I). Exposure to 100 μM lead caused mitochondrial filament elongation, while mitochondria in cells exposed to 500 μM Pb²⁺ were very fragmented and localized to the nuclear perimeter (Fig. 2I).

An equivocal role of PGC1α in neuroprotection and toxicity

To determine whether up-regulation of PGC1α was responsible for the changes in mitochondrial morphology and organization and subsequent protection of DA neurons from lead insult, we examined the effects of PGC1α overexpression on the structure of mitochondria (Fig. 2J and K) and (Fig. 3A and B). Intriguingly, overexpression of PGC1α actually increased susceptibility of N27 cells to lead toxicity at both 100 and 500 μM, as manifested by increased fragmentation of mitochondria, measured as a decrease in mitochondrial surface area (Fig. 3A and B). Whereas a two-fold increase in PGC1α expression protected cells against lead-induced neurotoxicity (endogenous levels of the protein doubled in response to 100 μM lead), higher expression levels (>50 fold) of the protein had the opposite effect and exacerbated the toxicity of lead (Fig. 2J and K). Furthermore, cells overexpressing PGC1α treated with 100μM and 500μM leaded showed condensed nuclei, an apoptotic hallmark (Supplementary Figure1-S1, C).

Figure 3. Lead-induced changes in mitochondrial morphology and dynamics. Immunocytochemistry and confocal imaging analysis of N27 cells co-transfected with an empty vector and mitoYFP (A, a-c) or PGC1α-encoding vector and mitoYFP (A, d-f). The extent of the mitochondrial network both under basal conditions (a), Pb²⁺ 100μM (b), Pb²⁺ 500μM (c) and PGC1α overexpression alone (d) or combined with Pb²⁺ 100μM (e) or 500 μM (f) was measured by Spectrum. (A, a’-f’) mitochondrial area surface determined using ImageJ->threshold->Intermodes,(B,W). (B) Mitochondrial surface area in cells was determined using ImageJ software.(C) N27 cells treated with 100μM lead for 48-hours were analysed for the expression of mitochondrial fusion (Mfn2) and fission (Drp1, Fis1) genes by qRT-PCR. (D) qRT-PCR analysis of the expression of Drp1 and Mfn1 in PGC1α overexpressing cells.(E) Chromatin immunoprecipitation (ChIP) with a PGC1α antibody was performed to investigate PGC1α protein binding to its promoter, Nrf1, Drp1 and SOD2 gene promoters,*P<0.05, **P<0.01, ***P<0.001, n=3; mean ± SE.

Since exposure to lead induced changes in mitochondrial organization, the expression profiles of genes encoding proteins regulating mitochondrial dynamics were investigated. RT-QPCR analysis revealed that 100 μM lead shifts the mitochondrial dynamic profile towards fusion (Fig. 3C). While Mfn2 mRNA levels were increased, the transcript levels of Drp1, a fission promoting factor, were significantly decreased upon treatment. The Fis1 mRNA expression profile remained unchanged. To investigate the role of PGC1α alone in regulation of mitochondrial dynamics, Drp1 and Mfn1expression levels were investigated in PGC1α-overexpressing cells. Levels of Mfn1 expression mRNA) were significantly increased whilst those of Drp1 were significantly decreased (mRNA and protein) (Fig. 3D and Supplementary Figure 3-S3, B).

This suggested that overexpression of PGC1α was accompanied by a shift of the mitochondrial dynamics profile towards fusion (Fig. 3D). ChIP assay was performed to identify regulatory targets of PGC1α. This revealed significantly increased binding of PGC1α to its own promoter and unexpected binding to the Drp1 promoter (Fig. 3E). Moreover, PGC1α bound to Nrf1 and SOD2 promoters.

In order to compare the effect of lead on neuronal cells in vitro and in vivo, tissue samples were obtained from substantia nigra of rats given drinking water containing 500 ppm lead acetate for a period of 14 weeks as well as from control animals given water with acetate only. Real time QPCR analysis showed that Pgc1α mRNA levels, as well as levels of its downstream targets (Nrf1, Nrf2, Tfam) were significantly increased in the substantia nigra of animals exposed to lead (Fig. 4A). Expression of both mitofusins was also significantly induced by lead (Fig. 4A). Interestingly, unlike N27 dopaminergic neurons exposed to lead for 48-hours in vitro, levels of theDrp1 transcript were increased together with levels of Mfn1 and Mfn2 transcipts, after lead exposure in vivo, with no decrease in the number of tyrosine hydroxylase-expressing, dopaminergic SN neurons cells (Fig. 4B and C). Since SN tissue used in the experiment was a heterogeneous mixture of different cell types, including neurons, we performed dual fluorescence immunohistochemistry of DRP1 and NeuN (a neuronal specific marker) in the ventral mesencephalon to determine whether the previously observed increase in Drp1 expression was specific to DA neurons (Fig. 4D). Further analysis confirmed that the expression of DRP1 protein was significantly induced by lead in SN pars compact a neurons only (Fig. 4E). As adult rats are less susceptible to lead toxicity than young rats, and a significant increase of Drp1 expression was observed in vivo at 48-hours, we looked at Drp1 expression in vitro after short-term exposure (Fig. 4F). After 3-hours of lead exposure in N27 cells, an elevation of Drp1 mRNA levels was observed (P<0.01).

Figure 4. In vivo sub-lethallead concentration in drinking water increases the expression of genes controlling mitochondrial biogenesis and dynamics. Adult rats received drinking water containing 500 ppm lead acetate for 14 weeks and control animals received water with acetate only. Substantia nigra tissue was isolated and mRNA was extracted and processed for Real time QPCR analysis. (A) mRNA levels of Pgc1α and its target genes Nrf1, Nrf2 and Tfam as well asMfn1, Mfn2and Drp1 were analysed by qRT-PCR; (B) Immunohistochemistry was performed to detect tyrosine hydroxylase (TH) positive neurons and (C) stereological counting of TH-positive neurons in the substantia nigra of rats given drinking water containing 500 ppm lead acetate for 14 weeks. (D) Immunohistochemistry of DRP1 and NeuN co-localisation in the ventral mesencephalon neuronal cells. (E) Drp1 signal intensity was analysed by confocal laser scanning microscope (Zeiss LSM 7 DUO), using the associated software package (ZEN 2010) and quantified using an Image Analysis Program from Soft Imaging System (analySIS®, Germany). (F) MfnI and Drp1 mRNA levels in N27 cells after 3-hours of lead exposure. *P<0.05, **P<0.01, ***P<0.001, n=6 (F, n=3); mean ± SE.

Lead treatment causes a disruption of cellular calcium balance

The possible role of ROS in lead-induced cell death in N27 cells was investigated using N-acetylcysteine (NAC), a well-known ROS scavenger. However, NAC pre-treatment of cells had no effect on lead-induced changes in mitochondrial ROS or membrane potential (Fig.5A and B). Therefore, we explored the possibility that changes in intracellular Ca²⁺ levels might mediate the effects of lead, since lead is known to mimic this cation inside the cell and compete with it for binding to Ca²⁺-dependent proteins. Moreover, calcium-regulated proteins have been shown to play a role in control of PGC1α expression [19].

Figure 5. Lead treatment causes a disruption of cellular calcium balance. Pre-treatment of N27 cells with 1 mM N-acetylcysteine (NAC) for 1 hour followed by a 48-hours lead-exposure was used to investigate the impact on mitochondrial ROS (A) and mitochondrial depolarisation (MMP) (B) N27 cells treated with lead for 48-hours were tested for cytoplasmic (C) and ER calcium levels (D) using thapsigargin (Tg) and Rhod-2 staining followed by flow cytometry analysis (D, a and b). Calcineurin transcript levels 48-hours post-lead exposure were investigated using qRT-PCR (E). Pre-treatment of N27 cells with 5μM BAPTA-AM for 1 hour followed by a 48-hours lead-exposure was used to investigate the impact of calcium on mitochondrial depolarisation (F) and cell death (G) following lead treatment. (H) Western blot of BAP31 protein levels after 48-hours lead treatment. (I) Western blot analysis of BAP31 levels in shBAP31 N27 cells. N27 cells with stable downregulation of BAP31 (shBAP31) were used to investigate the involvement of BAP31 in lead-induced mitochondrial superoxide generation (J), mitochondrial depolarisation (MMP) (K) and activation of caspases 3/7 (L) and 8 (M). *P<0.05, **P<0.01, ***P<0.001, n=3; mean ± SE.

Staining of cells with Fluo4-AM, a cytoplasmic calcium dye, revealed that lead causes an increase of cytosolic Ca²⁺ with depletion within the ER (Fig. 5C and D). Pre-treatment of cells with BAPTA-AM, a cytoplasmic calcium chelator, conferred protection against lead insult (Fig. 5F and G). This finding, together with a lack of response to NAC on cellular susceptibility to lead, implicates a primary role for calcium in the response of dopaminergic neurons to lead. mRNA levels of calcineurin, a modulator of a number of key Ca²⁺ signalling pathways in neurons, were significantly increased on exposure to higher concentrations of lead, with peak expression occurring at 100 μM (Fig. 5E). Calcineurin mRNA levels at 100 μM and 500 μM mirrored those of Pgc1α and its target genes Nrf1 and Tfam (Fig 1A-C).

Caspase-8 cleavage of BAP31 at the ER stimulates Ca2+-dependent mitochondrial fission and subsequently enhances pro-apoptotic signals between the ER and mitochondria [6, 20]. Since the protein level of BAP31 increased after exposure to 100μM and 500μM lead (Fig. 5H), we investigated whether BAP31 down-regulation had any neuroprotective effect against lead insult. shRNA delivered by a lentiviral vector was used to knock down Bap31 expression in N27 cells (Fig. 5I). shRNA-Bap31 conferred significant protection against lead toxicity (Fig. 5J-M) implying an important role of BAP31 in the response to lead and in calcium pathway regulation.

Cell culture and treatments

Rat mesencephalon-derived cell line (1RB₃AN₂₇; hereafter referred to as N27 cells)represent a homogenous population of tyrosine hydroxylase-expressing cells with functional characteristics resembling dopaminergic neurons. N27 cells were maintained in RPMI-1640 medium supplemented with 10% FBS, 100 U/ml penicillin and 100 μg/ml streptomycin (Sigma) at 37°C in a 5% CO₂ humidified atmosphere. Cells were exposed to lead (II) acetate (Sigma) dissolved in distilled water added to the growth medium at 5, 100 and 500 μM for 48-hoursor 3- hours, as indicated.

Stable cell line production

The pLKO.1-TRC cloning vector was obtained from AddGene. Oligonucleotides containing shRNA were cloned into the vector according to the manufacturer's protocol. For shRNA primers used for PGC1α and BAP31 see Supplementary Material Table 1.

Virions were produced in HEK293T cells, by combining three packaging plasmids (pVSV-G), pRev, pGag.Pol (Invitrogen) with pAdVantage (Promega), and pLKO vector. Water and 2 M CaCl₂ were added and complexes were mixed with 2× HBS (500 μl) and added to cells drop wise. After an overnight incubation, fresh DMEM medium containing 1 mM sodium butyrate was added. Post-transfection the supernatant was collected, spun down and filter-sterilized. N27 cells were transduced for 8-hours in the presence of polybrene (Sigma). Cells were selected with puromycin 3 days after transduction.

Successful knockdown was confirmed by the reduction in specific mRNA (Fig. 2E and protein, as determined immunocytochemically (Supplementary figure3-S3, A).

Caspase activity measurement

Caspase activity assays were performed using CaspaTag™ caspase-3/7 or 8 assay kit (Millipore) according to the manufacturer's protocol followed by data collection with FACSCalibur™ (BD Biosciences) and analysis with Cyflogic (CyfLo Ltd.)

Mitochondrial superoxide measurement

Mitochondrial superoxide levels were measured using MitoSOX™ Red (Invitrogen) according to the manufacturer's guidelines. FACSCalibur™ (BD Biosciences) was used for data collection and Cyflogic (CyfLo Ltd.) for the analysis.

Mitochondrial membrane depolarisation

Cells were harvested as described above and resuspended in 150 μl of PBS containing 40 nM 3,3-dihexaoxacarbocyanine iodide (DiOC6) and PI (50 μM) for 40 minutes at 37°C. Cells were then analysed using FACSCalibur™.

RNA extraction from cells

Total RNA extraction was performed using QiaShredder columns and the RNeasy extraction kit (Qiagen).

Animals and surgery

Twenty male albino Wistar rats (250-270 g) were kept at constant room temperature of 22 ± 1 °C and relative humidity (60%) with a 12-h light-dark cycle (8:00 a.m.-8:00 p.m.), with free access to food and water containing either sodium lead acetate (500 ppm) or sodium acetate for 14 weeks. Then, half of the animals in each group were killed by decapitation (RT-PCR experiments) or by perfusion (immunohisto-chemistry experiments).

Fluorescence immunohistochemistry

Thaw-mounted 20-μm coronal sections were cut on a cryostat at −15 °C and mounted in gelatin-coated slides. For double-labelling of DRP1 with NeuN, sections were blocked with PBS containing 1% normal goat serum and horse serum (Vector) for 1 h. The slides were washed three times in PBS and then incubated overnight at 4°C with either rabbit-derived anti-DRP1 (1:300; Cell Signalling) or mouse-derived anti-NeuN (1:1000; Millipore) diluted in PBS containing 1% normal goat/horse serum and 0.25% Triton X-100. Sections were incubated with goat anti-rabbit secondary antibody conjugated to fluorescein (1:200, for DRP1; Vector) and horse anti-mouse secondary antibody conjugated to Texas Red (1: 200, for NeuN; Invitrogen) for 1 h. Nuclei were counterstained with Hoechst (1 μg ml⁻¹, Molecular Probes). Fluorescence images were acquired using a confocal laser scanning microscope (Zeiss LSM 7 DUO) and processed using the associated software package (ZEN 2010). Signal intensity was quantified using an Image Analysis Program from Soft Imaging System (analySIS®, Germany).

Stereological counting of TH-positive neurons in SN

The number of TH-positive neurons in the SN was estimated using a fractionator sampling design. Counts were made at regular predetermined intervals (x = 150 μm and y = 200 μm) within each section. An unbiased counting frame of known area (40 × 25 μm = 1000 μm²) was superimposed on the tissue section image under a 100× oil immersion objective. Therefore, the area sampling fraction is 1000/(150 × 200) = 0.033. The entire z-dimension of each section was sampled; hence, the section thickness sampling fraction was 1. In all animals, 20-μm sections, each 100 μm apart, were analyzed; thus, the fraction of sections sampled was 20/100 = 0.20. The number of neurons in the analyzed region was estimated by multiplying the number of neurons counted within the sample regions by the reciprocals of the area sampling fraction and the fraction of section sampled[39].

RNA extraction from tissue

Rat substantia nigra were dissected with reference to a standard rat brain atlas (Paxinos and Wilson, 6^th Edition, Elsevier), immediately snap-frozen in liquid nitrogen and stored at −80C. Total RNA was extracted using the phenol-chloroform method. Ice-cold Trizol reagent (Life Technologies) was added to 25-50mg of sample according to the manufacturer's instructions. Brain tissue kept on ice was homogenized using a tissue homogenizer (OMNI international).

Real-time PCR

Five μg RNA was used to synthesize cDNA using the ThermoScript™ RT-PCR first strand synthesis system (random hexamer method) according to manufacturer's instructions (Invitrogen). Platinum SYBR Green qPCR Super Mix-UDG (Invitrogen) was used for qPCR analysis. Primers were designed using Primer Blast (supplementary data, table 1). Real time PCR reactions were performed using Stratagene Mx3000P instrument. MxPro Software (Stratagene) was used to obtain Ct values. Transcript levels relative to GAPDH were determined using the standard curve method.

Mitochondrial function assessment

The Seahorse XF analyzer was used according to manufacturer's guidelines (Seahorse Bioscience). OCR values were measured by XF24 extracellular flux analyzer.

Immunocytochemistry

Cells were stained with MitoTracker Red (Molecular probes) according to the manufacturer's protocol, fixed with 4% paraformaldehyde and blocked in 10mM HEPES, 0.3% TX-100, 3% BSA, pH 7.4 for 1 hour at RT. Coverslips were subsequently incubated with PGC1α antibody (overnight; Everest) and AlexaFluor 488 anti-goat IgG secondary antibody (RT, 1 h; Molecular Probes). Nuclei were counterstained with Hoechst (Molecular Probes).

Western blot analysis

Cells were homogenized on ice in lysis buffer (1 M Tris-HCL, 2.5 M NaCl, glycerol, 0.5 M β-glycerophosphate, Tween 20 and Nonidet P40). The lysate was cleared by centrifugation at 900 g for 10 min at 4 °C. 50-120 μg of proteins were run on a 10% SDS-PAGE gel and transferred to nitrocellulose membranes (GE healthcare). Membranes were blocked with 5% non-fat dry milk dissolved in PBS with 0.01% Tween 20 (Sigma). Incubation with primary antibodies was performed overnight at 4°C (see supplementary materials for the antibody list). Peroxidase activity was developed using enhanced chemiluminescence (ECL).

Chromatin immunoprecipitation

N27 cells were treated and fixed with formaldehyde and quenched with glycine. Chromatin immunoprecipitation of samples was performed as described previously [40] using PGC1α antibody, followed by Q-RT-PCR analysis. An irrelevant goat antibody (IgG) was used as a negative control, to control for the possibility of non-specific clumping of chromatin. C_t values were normalised according to the following equation: ∆C_t[normalised ChIP] = (C_t[ChIP] – (C_t[input] – log₂ (0.1))). The % input was calculated with the following equation: % input = 2^{(−∆Ct[normalised ChIP])}. For the list of target promoter primer pairs see supplementary materials. PGC-1α binding sites were all within promoter regions, defined as ±3 kb from a transcriptional start site (TSS). (Please see supplementary figure 3-S3, C).