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The Journal of Neuroscience, August 8, 2007, 27(32):8475-8485; doi:10.1523/JNEUROSCI.0792-07.2007
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Cellular/Molecular
Neurite Degeneration Induced by Heme Deficiency Mediated via Inhibition of NMDA Receptor-Dependent Extracellular Signal-Regulated Kinase 1/2 Activation
Tatyana Chernova, Joern R. Steinert, Christopher J. Guerin, Pierluigi Nicotera, Ian D. Forsythe, andAndrew G. Smith Medical Research Council Toxicology Unit, Hodgkin Building, University of Leicester, Leicester LE1 9HN, United Kingdom
Abstract
Top
Abstract
Introduction
The early stages of many neurodegenerative diseases and age-related degeneration are characterized by neurite damage and compromised synaptic function that precede neuronal cell death. We investigated the signaling mechanisms underlying neurite degeneration using cortical neuron cultures. Inhibition of heme synthesis caused neurite damage, without neuronal death, and was mediated by reduced NMDA receptor (NMDAR) expression and phosphorylation. The signaling toward the degenerative phenotype involved suppression of the extracellular signal-regulated kinase 1/2 (ERK1/2) pathway, and electrophysiological recording showed that the neurodegeneration is accompanied by reduced NMDAR current and Ca2+ influx, as well as reduced voltage-gated sodium currents, consistent with compromised neurite integrity. Rescue from the degenerative phenotype by heme replacement was dependent on restoration of NR2B subunit phosphorylation and expression of NMDAR currents with higher Ca2+ permeability, consistent with triggering prosurvival ERK1/2 signaling to maintain and extend neurites. This study demonstrated a new mechanism of neurodegeneration in which impaired heme synthesis led to NMDAR signaling dysfunction, suppression of the prosurvival ERK1/2 pathway, and progressive fragmentation of neuronal projections.
Key words: neurodegeneration; heme; ERK pathway; NMDA receptor; patch clamp; neuron
Introduction
The early pathological finding of neurodegenerative diseases is a selective loss of synaptic connections in the absence of cell soma demise (Fiala et al., 2002). Many neurodegenerative disorders are age-related and characterized by progressive cognitive or motor impairment. Mitochondrial dysfunction leading to oxidative damage is thought to be a major contributor to brain aging and injury and occurs at an early stage of all major age-related neurodegenerative diseases (Liu and Ames, 2005
The importance of heme in mammalian neurobiology, however, extends beyond its functions as a direct constituent of mitochondrial cytochromes to being a prosthetic group in a large number of proteins in oxygen sensing and metabolism, detoxification, control of oxidative damage, growth and differentiation, and production of CO and nitric oxide (Padmanaban et al., 1989
Here we show that heme deficiency in cultured cortical neurons leads to a neurodegenerative phenotype with neurite fragmentation, combined with reduced NMDAR-mediated and voltage-gated sodium currents. The neuronal damage and the reversal of neurodegeneration by heme replacement are mediated via the NMDAR-dependent ERK1/2 pathway. These findings suggest that heme may act as a signaling molecule to maintain neurite integrity.
Materials and Methods
Primary cell culture. Primary cortical neurons were isolated from 14-d-old fetuses of the BALB/c mouse strain bred in house. The BALB/c Fechm1Pas mouse strain was obtained from the The Jackson Laboratory (Bar Harbor, ME). The Fechm1Pas mutant contains a point mutation in the ferrochelatase gene (Tutois et al., 1991Inhibition and measurement of heme synthesis. To inhibit heme synthesis, cells were cultured in serum-free medium with 0.5 mM succinyl acetone (SA) (Sigma, Dorset, UK) continuously for the duration of the experiments. For measurement of heme synthesis, cells were incubated with 0.4 µCi of [3,5-3H]aminolevulinic acid hydrochloride (ALA; 2.6 Ci/mmol; PerkinElmer, Boston, MA) for 24 h. Heme was extracted from the cells by acetone-HCl and diethyl ether. The amount of radioactivity in extracted heme was measured by scintillation counting as described previously (Shedlofsky et al., 1987
Viability assay. Cell viability in heme depletion experiments was estimated by using the LIVE/DEAD Reduced Biohazard Viability/Cytotoxicity kit (Invitrogen) according to the instructions of the manufacturer. The amounts of live and dead cells were scored with a fluorescence microscope.
RNA extraction and quantitative real-time PCR analysis. Treated and untreated cells at different time points were collected, and total RNA was isolated by using TRI-reagent (Sigma). cDNA synthesis was performed using random primers and Superscript II (Invitrogen). PCR primers were selected using the Primer Express version 2.0 Software program (Applied Biosystems, Foster City, CA). Primers sequences were as follows: ß-actin forward primer, 5'-GATTACTGCTCTGGCTCCTAGCA-3'; ß-actin reverse primer, 5'-GTGGACAGTGAGGCCAGGAT-3';
-aminolevulinate synthase 1 (ALAS1) forward primer, 5'-TCTTC-CGCAAGGCCAGTCT-3', ALAS1 reverse primer, 5'-TGGGCTTGAGCAGCCTCTT-3'; HMOX1 forward primer, 5'-CACTTCGTCAGAGGCCTGCTA-3'; HMOX1 reverse primer, 5'-GTCTGGGATGAGCTAG-TGCTGAT-3'; NMDA NR2B forward primer, 5'-CTTAATCTGTCC-GCCTAGAGCTTT-3'; and NMDA NR2B reverse 5'-TGCGCTGGG-CTTCATCTT-3'.
Primers were designed to cross exon–exon boundaries, and the concentration was optimized (300–900 nM) to ensure that the efficiency of the target amplification and the efficiency of the endogenous reference amplification are approximately equal. PCR was performed using SYBR Green PCR Master Mix, primers, and 10 ng of reverse-transcribed cDNA in the ABI PRISM 7700 Sequence Detection System (Applied Biosystems); the thermal cycler protocol was as follows: stage one, 50°C for 2 min; stage two, 95°C for 10 min; stage three, 40 cycles at 95°C for 15 s and 60°C for 1 min. Each sample was run in triplicate. Quantification was performed using the comparative CT method (
CT). Data are presented as the mean ± SD (n = 3–8 for each group). Statistical significance was assessed as p < 0.05 using one-way ANOVA.
Immunoblotting. Proteins were extracted from primary neurons after 14 and 21 d of culturing using lysis buffer (7 M urea, 50 mM Tris-HCl, pH7.5, and 5 mM DTT) followed by brief sonication. SDS electrophoresis and immunoblotting were performed (Davies et al., 2005
-tubulin, B-Raf, and phospho-B-Raf from Santa Cruz Biotechnology (Santa Cruz, CA); and NMDA NR2B and phospho-NMDA NR2B from PhosphoSolutions (Aurora, CO). Results were quantified using densitometry and Image Quant 5.2 software. Statistical significance of data were estimated using two-tailed Student's t test.
Immunocytochemistry. After treatment, cells were fixed with 4% paraformaldehyde at room temperature for 20 min and permeabilized with 0.2% Triton X-100 (Sigma) in PBS for 5 min. Cells were then incubated with rabbit anti-ßIII-tubulin (1:3000) antibody (Cell Signaling Technology) at room temperature for 1 h. Secondary antibody (goat anti-rabbit Alexa Fluor 488 at 1:500) were then added for 1 h, followed by nuclear staining with 300 nM 4'-6-diamidino-2-phenylindole for 10 min. Secondary antibodies alone were used as specificity controls and uniformly resulted in very low background levels of fluorescence.
Treatment with inhibitors. Kinase inhibitors were applied to the neurons undergoing various treatments as follows: 10 µM U0126 [1,4-diamino-2,3-dicyano-1,4-bis(o-aminophenylmercapto)butadiene] (Calbiochem, La Jolla, CA), 50 µM LY294002 [2-(4-morpholinyl)-8-phenyl-1(4H)-benzopyran-4-one] (Cell Signaling Technology), 10 µM GF109203X (LC Laboratories, Woburn, MA), 100 nM Ro 31-8220 (2-[1-(3-(amidinothio)propyl)-1H-indol-3-yl]-3-(1-methylindol-3-yl)maleimide) (Sigma), and 50 nM PP2 [4-amino-5-(4-chlorophenyl)-7-(t-butyl)pyrazolo[3,4-d]pyrimidine] (Calbiochem) for 30 min. NMDAR competitive antagonist 50 µM
Imaging and quantification of immunostaining. Images of cultured cells were obtained with a Zeiss (Jena, Germany) LSM 510 META confocal microscope equipped with a 25x, 0.8 numerical aperture water immersion lens. Fluorochromes were excited with a pulsed 780 nm titanium/sapphire laser (Newport Spectra Physics, Didcot, Oxfordshire, UK). Images were collected as Z series of 12 sections with voxel dimensions of x,y,z = 0.32 x 0.32 x 0.32 µm (4 µm total thickness). Staining for ßIII-tubulin was quantified using Velocity (Improvision, Coventry, UK) on three-dimensional reconstructions of datasets using an intensity classifier and size exclusion criteria to identify neurites. Images were collected from at least three different randomly selected areas from each of two duplicate cell cultures. Average volume data and total volume per nuclei from each time point and each treatment was compared using a one-way ANOVA, with a Tukey's multiple comparison test (Prism software; GraphPad Software, San Diego, CA).
NMDA puffing. Agonist application experiments on primary cortical cultures between 6 and 14 d in vitro (DIV) were performed using a Picospritzer III (Intracel, Frederick, MD) connected to a patch electrode filled with artificial CSF (aCSF) containing 100 µM NMDA. The tip of the electrode was placed near a patch-clamped cortical neuron, and the maximal response to the agonist was optimized by adjusting the pipette position. Puffs were applied for 30 ms every 8 s (20 psi) in the presence of DNQX (10 µM), D-serine (10 µM), and bicuculline (10 µM). No extracellular Mg2+ was added to these solutions. To create an current–voltage (I–V) relationship for NMDA-evoked currents, we applied NMDA while holding at different potentials ranging from –70 to +40 mV.
Calcium imaging. For [Ca2+]i determinations, subconfluent cortical neurons were cultured on glass coverslips and loaded for 30 min at room temperature with 5 µM fura 2-AM (Invitrogen) aCSF. After loading, cells were kept in aCSF for up to 1 h. Fluorescence was measured as described previously (Grynkiewiecz et al., 1985
Electrophysiology. Whole-cell patch-clamp recordings were made from primary cultures of cortical neurons [visualized at 60x with a Nikon (Tokyo, Japan) E600FN fitted with differential interference contrast optics] using a multiclamp 700B amplifier (Molecular Devices) and pClamp9 software (Molecular Devices). Data were sampled at 50 kHz and filtered at 10 kHz. Patch pipettes were pulled from borosilicate glass capillaries (GC150F-7.5; Harvard Apparatus, Edenbridge, UK) using a two-stage vertical puller (PC-10; Narishige, Tokyo, Japan). Tips were polished (Micro Forge; Narishige), and pipettes had a final open-tip resistance of
5 M
when filled with a patch solution containing the following (in mM): 130 CsCl, 10 HEPES, 5 EGTA, and 1 MgCl2, pH adjusted to 7.2 with CsOH. Whole-cell access resistances were <20 M
Results
To explore the role of heme signaling in neuronal survival, we developed a model of heme deficiency using cortical neuron cultures. Treatment with SA, a specific inhibitor of 5-aminolaevulinic acid dehydratase (Tschudy et al., 1981Inhibition of heme synthesis causes neurodegeneration
To examine morphological changes, heme-deficient cortical neuron cultures were monitored by staining for the cytoskeletal protein ßIII-tubulin. Immunostaining on 8, 10, and 15 DIV revealed that heme-deficient cultures exhibit neurodegeneration (Fig. 1). In SA-treated cells (Fig. 1B,E,H), the ßIII-tubulin staining was discontinuous, showing increasing neurite fragmentation and disruption in the axonal/dendritic microtubule structure. Fragmentation increased progressively throughout the treatment. By 15 DIV, only a limited number of neurites in the heme-deficient culture were intact (Fig. 1H). Neurites in the control cultures remained intact through 10 DIV and by 15 DIV had a few bright spots along some processes, consistent with slight degeneration as a result of aging (Fig. 1A,D,G). Strikingly, cultures treated with SA and heme maintained a well established network of projections for the entire duration of the experiments, showing only minor fragmentation of neuronal processes with age in culture and far less than in the SA-treated samples (Fig. 1C,F,I).
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Figure 1. Neurodegeneration of cultured cortical neurons caused by heme deficiency. Confocal micrographs of maximum projection Z series of cultured neurons stained for ßIII-tubulin on 8, 10, and 15 DIV. A, D, G, Control cultures (C); B, E, H, cells treated with SA; C, F, I, cells treated with SA and heme (H). Quantification of immunostaining for ßIII-tubulin reflecting the degree of neurodegeneration. K, Average neurite volume in control and treated cultures at each time point. J, Total volume per nuclei in image fields examined. *p < 0.05; **p < 0.01; ***p < 0.001.
To quantify neurite fragmentation, staining for ßIII tubulin was assessed on three-dimensional reconstructions of datasets. Total volume per nuclei was used as a measure of equality between cell numbers in image fields examined (Fig. 1J). No significant differences were found between control and treatments, indicating that the sampling was free from cell density bias. Average neurite volume per cell was used as a measure of neurite fragmentation and was compared between all treatments at each time point (Fig. 1K). Significant differences were found between control and heme-deficient neurons at 10 and 15 DIV. Heme replacement essentially prevented fragmentation of neurites at all ages. To determine whether this observation reflects cell death-independent degeneration, we applied a viability/cytotoxicity assay. The proportion of live and dead cells was assessed using fluorescent microscopy at 5, 9, and 14 DIV. No significant difference was observed at these time points between controls and any of the treatments.The results of a cell viability assay conducted on 16, 19, and 22 DIV showed that long-term treatment with SA eventually results in death of the cell body after 3–4 weeks (supplemental Fig. 1, available at www.jneurosci.org as supplemental material). The proportion of dead cells after 16 d of treatment was 34 ± 4% (compared with 29 ± 5% in control culture), but the loss of the projections at this stage was dramatic as reflected in Figure 1H. By 19 DIV, the number of dead cells increased to 60 ± 5%, whereas virtually all neurites were fragmented. On 22 DIV, the culture had no visible neurite network, and 95 ± 4% of the cells were dead. Control cultures on 22 DIV still had a well established neurite network, and the number of dead cells was 48 ± 4%.
Heme depletion and restoration are reflected in Alas1 and Hmox1 expression
Our model was based on sustained 70% reduction of heme synthesis by treatment with 0.5 mM SA. Inhibition of heme synthesis by SA caused upregulation of Alas1, the first and rate controlling enzyme (Fig. 2B) as a result of end-product negative feedback regulation (Sassa and Nagai, 1996
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Figure 2. Inhibition of heme synthesis in primary cortical neurons. Neurons were cultured in the presence of 1, 0.5, or 0.25 mM SA for 14 d. A, Treated with SA and control (C) cells were incubated with [3H]ALA for 24 h on day 14, and then labeled heme was extracted and measured (n = 5). B, C, Expression of ALAS1 and HMOX1 in control and 0.5 mM SA-treated on day 14 with or without 100 nM heme cultures, estimated by real-time reverse transcription-PCR. **p < 0.01, statistically different from control group.
Heme deficiency impairs basal activation of ERK1/2 pathway
Previously, we have shown that heme deficiency suppresses NMDAR subunit expression and causes premature neuronal aging (Chernova et al., 2006
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Figure 3. Rescue effect of heme on basal activation of ERK1/2 pathway in the neurons with inhibited heme synthesis and aged cells. A, BALB/c neurons were treated with SA with or without heme for 14 d. Additional treatments with 10 µM MEK inhibitor U0126, with 50 µM LY294002 (PI3K inhibitor), with 1 µM GF109203X (PKA inhibitor at 2 µM and PKC inhibitor at 10 nM), and with 10 µM Ro 31-88220 (PKC inhibitor) were applied for 30 min before harvesting the cells. B, Neurons were isolated from BALB/c or Fechm1Pas mice and cultured for 14 or 21 d, respectively. Treatment with 0.1 µM hemin (H) was conducted for the entire length of the experiments with Fechm1Pas (Fe) neurons and for 7 d (15–21 DIV) before harvesting of BALB/c neurons. Total cell lysates were analyzed by Western blot, probing for phospho-ERK and total ERK (A, B), phospho-p38, phospho-JNK, and
Rescue of ERK1/2 activation by heme requires b-Raf but not c-Raf
These above results indicate that the survival signaling of heme is specifically mediated by the ERK1/2 pathway, upstream of MEK1/2 and independent of PI3K, PKC, or PKA. To test for upstream affectors of ERK1/2, we measured c-Raf and b-Raf kinases in the heme-depletion model. In the highly conserved Ras/Raf/MEK/ERK signaling pathway, different Raf isoforms activate MEK in a range of cell types (Mercer et al., 2002
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Figure 4. Expression and phosphorylation of b- and c-Raf in heme-deficient and heme-sufficient cortical neurons. Western blots were performed on cultured BALB/c neurons treated with SA and hemin for 14 d. Neurons from Fechm1Pas mouse were treated with hemin for 24 h before harvesting. A, Decreased expression of b-Raf in SA-treated neurons. Twenty micrograms of proteins in total cell lysates were analyzed by Western blot, probing for total b-Raf, and the same blots were reprobed for
Exogenous heme has a differential effect on phosphorylation of NMDAR subunits
The observed loss of neurite integrity and consequent loss of connectivity in heme-deficient cultures could severely impair synaptic signaling and impact on expression of synaptic receptors. Activation of the ERK1/2 signaling pathway is dependent on calcium influx through synaptic NMDARs (Sweatt, 2004
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Figure 5. Differential effect of heme on total expression and phosphorylation of NMDAR subunits in heme-deficient neurons. Total expression of NMDAR subunits NR1 and NR2B detected by Western blot (A) and by real-time PCR (B). Neurons were treated with SA and hemin for 14 d. Additional treatments with 10 µM MEK1/2 inhibitor U0126, with 50 µM LY294002 (PI3K inhibitor), with 1 µM GF109203X (PKA inhibitor at 2 µM and PKC inhibitor at 10 nM), and with 10 µM Ro 31-8220 (PKC inhibitor at nanomolar concentration and PKA inhibitor at micromolar concentration) were applied for 30 min before harvesting the cells. Twenty micrograms of proteins were loaded for each sample, probing for total NR1 (A), phospho-NR1 (Ser890) (C), phospho-NR1 (Ser896) (D), phospho-NR1 (Ser897) (E), phospho-NR2B (Tyr1336) (F), and
NMDAR phosphorylation also regulates channel kinetics and membrane localization/trafficking (Cull-Candy et al., 2001NR2B is the main tyrosine-phosphorylated protein regulated by Src family kinases (SFKs) in the postsynaptic density (Moon et al., 1994
Neuronal rescue by heme replacement is abolished by NMDAR block
NMDAR activity is regulated by numerous endogenous modulators, including activation by serine–threonine and tyrosine phosphorylation. To elucidate the mechanism by which heme is involved in the activation of the prosurvival ERK1/2 pathway, we blocked NMDAR activity in SA- and heme-treated cells, using the competitive inhibitor AP-5 (50 µM) and abolished the stimulatory effect of heme on ERK1/2 in heme-depleted culture (Fig. 6A). Another way of suppressing NMDAR gating is by inhibiting the members of the SFKs, which enhance NMDAR function. Inhibition of SFKs by PP2 eliminated the activation of ERK1/2 by added heme (Fig. 6B). Inhibition of NR2B-containing NMDARs using the selective antagonist ifenprodil (10 µM) (Williams, 1993
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Figure 6. Effects of NMDAR inhibition on the rescue of ERK1/2 phosphorylation by heme. A, Cortical neurons treated for 14 d with SA and heme were incubated for 30 min with the competitive NMDAR antagonist 50 µM APV. Total cell lysates were analyzed by immunoblotting to detect activated ERK1/2. The bottom shows the same blot reprobed with an antibody recognizing total ERK1/2. B, Cortical neurons treated for 14 d with SA and heme (H) were incubated for 30 min with SFK inhibitor PP2 (1 µM). Equal amounts of protein extracts were analyzed by Western blot with phospho-ERK1/2 antibody. The bottom shows the same blot reprobed with anti-total ERK1/2 antibody. C, Cortical neurons treated for 14 d with SA and heme were incubated for 30 min before harvesting with 10 µM ifenprodil (IFE) and with 1 µM PP2. Each column is the mean ± SD of phospho-ERK1/2/ERK1/2 ratio from three independent experiments. *p < 0.05. C, Control.
Heme depletion reduces NMDA-evoked and voltage-gated sodium currents
Although we can clearly demonstrate molecular changes in NMDARs on heme depletion, it was important to establish whether these changes were directly translated into differences in NMDAR expression in the neuronal membrane and/or to determine whether other ionic channels may be involved. Whole-cell patch-clamp recordings were made from single cortical neurons maintained in tissue culture under identical conditions to the above experiments. The voltage-gated and leak potassium currents were similar in control and heme-depleted cultures, as shown in the I–V relationship in Figure 7A. However, there was a strong reduction of voltage-gated Na+ currents in response to SA treatment, which recovered with heme replacement (Fig. 7B). NMDA currents were measured in the absence of extracellular magnesium ions so as to avoid voltage-dependent block at negative holding potentials. Pressure application of NMDA (100 µM, 30 ms) was used over a range of holding potential to generate an I–V relationship in control cultures and compared with I–Vs recorded from SA-treated neurons at similar ages (>12 DIV). Heme depletion resulted in significantly reduced NMDA-mediated whole-cell current (Fig. 7C, raw traces). The half-duration for these NMDA currents (t(1/2), measured at –70 mV in the absence of [Mg2+]o) increased after SA treatment and did not completely recover with heme replacement (Fig. 7C, insets) (t(1/2)control = 1.45 ± 0.28 s, n = 8, t(1/2)SA = 3.1 ± 1.32 s, n = 6; t(1/2)SA+heme = 2.64 ± 0.19 s **, n = 6; **p < 0.01). A summary of data averaged across the age range of 6–13 DIV is shown in Figure 7D, with data from SA-treated cells normalized to current magnitudes in control cells at 12 DIV. Leak currents were measured at –110 mV, voltage-dependent outward K+ currents (high-voltage-activated IK) were measured at +30 mV, voltage-gated Na+ currents (Nav) were measured at –20 mV, and NMDA-evoked whole-cell currents (INMDA) were measured at +40 mV. Both Nav and INMDA were significantly reduced after 12 d of heme depletion. NMDA and Na+ currents exhibited age-dependent changes in magnitude, showing a developmental increase under control conditions over 6–11 DIV. NMDA currents were significantly reduced in SA-treated cells at all ages, but surprisingly, heme replacement did not restore NMDA current amplitudes to control levels at 13 DIV (Fig. 7C,E).
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Figure 7. Heme depletion reduces NMDA and voltage-gated sodium currents but not potassium currents. A, I–V relationship of primary cultured cortical neurons at 12–13 DIV (control vs SA treatment) shows no change in leak or voltage-activated outward currents (insets shows superimposed data traces). B, Representative traces for whole-cell sodium currents evoked by stepping the holding potential from –70 to –20 mV for cells treated with SA and SA in the presence of heme. C, Whole-cell currents evoked by pressure application of NMDA (100 µM) recorded at holding potentials of –40, 0, and +40 mV in control cells (top), cells treated with SA (middle), and SA in the presence of heme (bottom). Insets at each condition show averaged traces at –70 mV to illustrate the change in decay kinetics. D, Summary of ionic current changes caused by heme depletion. Mean data are normalized and expressed relative to control values in 12 DIV cells. Both INa and INMDA were significantly reduced. E, NMDA currents increased with days in culture. Mean data for untreated, SA-treated, and SA plus heme-treated cells are normalized to control values at 12 DIV cells. F, Voltage-activated Na+ currents increased with days in culture. Mean data for untreated, SA-treated, and SA plus heme-treated cells are normalized to control values at 12 DIV cells. SA treatment reduced Na+ currents at 12–13 DIV after an initial augmentation at 6–8 DIV. Heme replacement reestablished these current at 13 DIV. Data denote mean ± SEM of at least six different cells per data point. Significant differences are shown using Student's unpaired t test; *p < 0.05, **p < 0.01 compared with control; #p < 0.05 relative to SA treatment.
Voltage-gated Na+ currents showed an initial augmentation in SA-treated cultures, consistent with accelerated senescence and then strongly declined to <20% control levels by 13 DIV (Fig. 7F). Heme replacement dramatically enhanced Nav currents.Heme replacement favors NMDAR channels that exhibit higher calcium permeability
The above electrophysiological data on NMDA currents appears in conflict with previous data showing that heme replacement restores NMDAR expression to control levels. Because NMDARs are highly Ca2+ permeable (Schneggenburger et al., 1993
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Figure 8. Heme depletion reduces NMDA-evoked Ca2+ influx. A, Representative traces of intracellular Ca2+ ([Ca2+]i) recordings (expressed as 340/380 nm ratio) in control cells (Ctrl), cells treated with SA, and cells treated with SA plus heme at 12 DIV. Top, False color images taken at indicated time points. B, NMDA-evoked I–V relationships of two control cells at 12 DIV. Note that the arrows indicate the Er. C, I–V relationships of NMDA-evoked currents in SA-treated cells at 12 DIV. Note the negative shift in reversal potential compared with controls in B. D, NMDA-evoked
Discussion
Using heme depletion of cortical neurons in primary culture, we have shown that heme deficiency leads to a morphological phenotype similar to that seen in slow neurodegenerative diseases. We demonstrate that inhibition of heme synthesis causes neurodegeneration via NMDAR-dependent suppression of the ERK1/2 pathway. Reduced NMDAR expression is combined with changes in the relative Ca2+ permeability of functional NMDARs and reduced calcium influx. Associated reductions in voltage-gated sodium currents with heme depletion would further compromise neuronal signaling. These results provide important new insights into how compromised heme synthesis and availability influence key aspects of neuronal signaling associated with neurodegeneration and aging.In a previous study, we characterized this heme-depletion model as exhibiting key features of senescence (Chernova et al., 2006
The progressive fragmentation of neurons without an increase in cell soma loss caused by heme depletion resembles the early progress of many neurodegenerative diseases and age-related dementias. Our purpose here was to explore the molecular signaling pathways leading from compromised heme metabolism to neurodegeneration. A recent study has demonstrated that activation of ERK1/2 in primary neurons delays Wallerian degeneration of synaptic terminals induced by proteasome inhibitors (MacInnis and Campenot, 2005
Neuronal processes may be more sensitive to heme-related mitochondrial dysfunction attributable to their decreased size compared with the cell soma and therefore the inability to buffer or compensate for even small functional changes. The rescue of neurons from neurodegenerative changes by exogenous heme indicates the specificity of the phenomenon and is consistent with promotion of neuroblastoma neurite outgrowth by exogenous heme but not biliverdin (Ishii and Maniatis, 1978
Functional heme deficiency could be a contributing factor to Alzheimer's disease (Atamna and Boyle, 2006
We found that the damage induced by heme deficiency and the rescue by heme replacement were mediated via the ERK1/2 pathway, which is a key regulatory cascade of neuronal survival. Similar results were observed in the Fechm1Pas transgenic mouse and in SA-treated neurons, directly linking the phenomenon with heme synthesis. The phosphorylation of ERK2, which is the isoform of greater importance (English and Sweatt, 1996
Direct or indirect upstream regulators of ERK1/2 include Ras, Src, glutamate (via NMDAR increased gating), Ca2+, Zn2+, calcium/calmodulin-dependent kinase II, and some growth factors (Subramaniam and Unsicker, 2006
NMDA-initiated ERK1/2 activation is calcium-dependent because the activation of Ras by RasGRF1 occurs only if it binds calcium/calmodulin (Farnsworth et al., 1995
The recent evidence for spontaneous formation of the stable hemin-Slo1 BK channel (Tang et al., 2003
Together, these findings lead to the conclusion that the major neuronal injury caused by heme deficiency is focused around NMDAR dysfunction. Compromised heme metabolism may be the starting point for a vicious circle: dysfunctional NMDARs lead to decreased Ca2+influx, triggering diminished ERK1/2 activation, followed by loss of connectivity of neuronal processes and causing loss of synaptic NMDARs, further decreased Ca2+influx, etc. This study suggests a new scenario of neurodegeneration, in which compromised heme availability produces a subtle deficit in NMDAR-dependent ERK1/2 activation leading to neurite loss rather than somatic death. Additional exploration of the molecular and cellular mechanisms of heme biology may provide broad insights into the early stages of neurodegenerative disease and age-related neurodegeneration.
Footnotes
Received Feb. 21, 2007; revised June 8, 2007; accepted June 11, 2007.We thank the members of the Biomedical Services staff for their assistance.
Correspondence should be addressed to Tatyana Chernova, Medical Research Council Toxicology Unit, Hodgkin Building, University of Leicester, Lancaster Road, Leicester LE1 9HN, UK. Email: tc28{at}le.ac.uk
Copyright © 2007 Society for Neuroscience 0270-6474/07/278475-11$15.00/0
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