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The brain depends on redox electrons from NADH to produce ATP and oxyradicals (ROS). Since ROS damage and mitochondrial dysregulation are prominent in aging and Alzheimer’s disease (AD) and their relationship to redox state is unclear, we wanted to know whether an oxidative redox shift precedes these markers and leads to macromolecular damage in a mouse model of AD. We used the 3xTg-AD mouse model that displays cognitive deficits beginning at 4 months. Hippocampal/cortical neurons were isolated across the age-span and cultured in common nutrients to control for possible hormonal and vascular differences. We found an increase of NAD(P)H levels and redox state in non-transgenic neurons until middle age, followed by a decline in old age. The 3xTg-AD neurons maintained much lower resting NAD(P)H and redox state after 4 months, but the NADH regenerating capacity continuously declined with age beginning at 2 months. These redox characteristics were partially reversible with nicotinamide, a biosynthetic precursor of NAD+. Nicotinamide also protected against glutamate excitotoxicity. Compared to non-transgenic neurons, 3xTg-AD neurons possessed more mitochondria/neuron and lower glutathione levels which preceeded age-related increases in ROS levels. These glutathione deficits were again reversible with nicotinamide in 3xTg-AD neurons. Surprisingly, low macromolecular ROS damage was only elevated after 4 months in the 3xTg-AD neurons if anti-oxidants were removed. The present data suggest that a more oxidized redox state and a lower antioxidant glutathione defense can be dissociated from neuronal ROS damage, changes that precede the onset of cognitive deficits in the 3xTg-AD model.
The brain depends on the redox reaction of NADH to transfer electrons to oxygen in the production of ATP by oxidative phosphorylation. Some of the electrons escape tight controls to reduce oxygen (O2−•) that is further reduced to OH•, ONOO−3, OH−, collectively called reactive oxygen species (ROS) that can damage macromolecules. With aging and in Alzheimer’s disease (AD), ROS damages brain proteins (Butterfield et al., 1998; Hensley et al., 1998; Smith et al., 1998), lipids (Montine et al., 1998) and nucleic acids (Nunomura et al., 1999) but determining whether these are a consequence of metabolic aging or causal to AD has been elusive. The major electron transfer currencies for oxidation and reduction reactions in cells are the redox couples NADH/NAD+ and NADPH/NADP+. A reductive shift in redox is associated with cell proliferation; an oxidative shift with morbidity and apoptosis (Jones, 2006). These principles led us to formulate the epigenetic oxidized redox shift (EORS) theory of aging (Brewer, 2010) and propose that an oxidized redox state is upstream of ROS generation. Here, we test the hypothesis that an age-related oxidized redox shift is upstream of elevated ROS levels and reported cognitive deficits. We determine the redox shift as intrinsic fluorescence of intracellular NADH (Chance and Williams, 1956; Chance et al., 1979) in isolated neurons with reference to the oxidized flavin nucleotide, FAD (Parihar et al., 2008). Since 0.1 mM NADH is the major intracellular redox currency with 8 times more free energy than ATP (Klaidman et al., 1995; Brewer, 2010) and millimolar glutathione (GSH) (Kudo et al., 1990) is the major redox buffer in cells (Das and White, 2002), we further hypothesized that aging is associated with an oxidized redox state evident as an oxidative shift in the NADH redox ratio and a decline in glutathione levels that precede elevations in ROS.
Nicotinamide, a biosynthetic precursor of NAD+, was previously shown to increase NADH and NADPH levels in striatum and thalamus of whole brain tissue of adult mouse (Klaidman et al., 2001). However, the AD and age-dependent effects of nicotinamide on neurons remains unclear. Nicotinamide also improved the cognitive deficits, which begin at 4 months, in a hippocampal and amygdala dependent task in a triple transgenic mouse model of AD (3xTg-AD) (Oddo et al., 2003a; Oddo et al., 2003b) by selectively decreasing Thr231 phosphotau, increasing stabilizing acetylated α-tubulin and increasing synaptogenic p25, all associated with improved learning and memory (Green et al., 2008). These findings led us to investigate if nicotinamide could increase intracellular NAD(P)H levels and change the redox state in 3xTg-AD mice. We measured other possible events upstream of AD-like pathology including cytochrome C release, glutathione antioxidant defenses and macromolecular damage in aging and in male 3xTg-AD mouse model to determine the relationship of ROS damage to the onset of cognitive changes in these mice. We used cultured adult neurons (Brewer, 1997; Brewer and Torricelli, 2007) that enable age-related comparisons of neurons under common culture conditions that control or eliminate hormonal, vascular and inflammatory changes present with aging in vivo.
We used LaFerla’s triple transgenic mouse model of Alzheimer’s disease (3xTg-AD) with human transgenes APP (SWE), PS1 (M146V) and Tau (P301L) under control of thy 1.2 promoters (Oddo et al., 2003a). Non-transgenic (non-Tg) mice on the same mixed C57BL6/129 background from LaFerla were used as controls. For all our experiments, we used male 3xTg-AD and non-Tg animals housed 1–4 per cage, fed rodent diet 5001 (LabDiet, Purina, distributed by El Mel, St Louis, MO, with 28.5% calories from protein, 13.5% from fat and 58% from carbohydrates) ad libidum at controlled temperature, humidity and a 12 hour light, 12 hour dark cycle. All animals were genotyped before experiments. DNA was extracted from tail snips using Roche High Pure PCR Template Preparation Kit (Indianapolis, IN). We used two primer sets for identifying the 3xTg-AD mutant. The following primers identified the PS1 gene F5′-CAC ACG CAA CTC TGA CAT GCA CAG GC – 3′, R5′ – AGG CAG GAA GAT CAC GTG TTC AAG TAC – 3′ (Invitrogen, Carlsbad, CA). The PCR product was digested with the restriction enzyme BSTEII (New England Biolabs, Ipswich, MA). Non-Tg mice showed a single band (~ 550 bp) and 3xTg-AD mouse showed two bands (~ 300 and 250 bp). The following primers identified the hAPP gene F5′ - GCT TGC ACC AGT TCT GGA TGG – 3′, R5′ – GAG GTA TTC AGT CAT GTG CT -3′. Non-Tg mice showed no amplification and 3xTg mouse showed one band (~ 300 bp).
Adult hippocampal and cortical neurons were isolated from Non-Tg and 3xTg-AD age matched male mice at 2, 4, 8, 11, and 21 month time points (Brewer and Torricelli, 2007). The combined hippocampus and frontal-cortex of each hemisphere (~ 140 mg tissue) were sliced at 0.5 mm and combined in Hibernate A (BrainBitsLLC.com), 2% B27 supplement (Invitrogen), 0.5 mM Glutamax (Invitrogen). The tissue was digested with 2 mg/mL papain (Worthington, Lakewood, NJ) for 30 min at 30°C. Slices were triturated and the neurons separated from debris and microglia on an Optiprep density gradient. In order to limit the toxicity caused by excess debris, each hippocampus and hemi-cortex was digested, triturated, and spun through a density gradient in separate tubes. After the gradient, the neuron-enriched fractions of each hippocampus and frontal-cortex were combined. Viable cells were counted by exclusion of trypan blue. Neurons were plated at 32,000 cells/cm2 on 12 mm and 15 mm glass coverslips (Assistent brand, Carolina Biologicals, Burlington, NC) that were coated overnight with poly-D-lysine, 100 μg/mL in water. The cells were plated and cultured in Neurobasal A/B27/Glutamax with 10 ng/mL FGF2 and 10 ng/mL PGDFbb (Invitrogen, Carlsbad, CA) for trophic support. Cells were cultured for 7–12 days at 37°C in 5% CO2, 9% O2 at saturated humidity (Forma, Marietta, OH).
For neuron survival, live cells on glass coverslips were stained with fluorescien diacetate (15μg/ml; Sigma–Aldrich, St. Louis, MO) and dead cells with propidium iodide (4.6μg/ml; Sigma–Aldrich) (Brewer et al., 1993). After washing the slips with HBSS (Invitrogen), cells were observed by blue and green fluorescence excitation through a 20x objective (Olympus) for green and red fluorescence. Survival was calculated as the average percent live divided by the total cells (live+dead) in 8–12 adjacent fields.
Single live cells were imaged for simultaneous NAD(P)H and FAD measurements as before (Parihar et al, 2008) with slight modification. Since our system cannot distinguish between NADH and NADPH fluorescence, we use the more general term NAD(P)H for our experiments. However, 80% of autofluorescence originates from NADH (Eng et al., 1989), so we can assume that NADH contributes maximally to the NAD(P)H autofluorescence pool. Cells cultured for 8 days on 15 mm glass coverslips were mounted on a slip holder (Warner Instruments, Hamden, CT) in 500 μl Hibernate A Low Fluorescence (BrainBits LLC, Springfield, IL) and 0.5 mM Glutamax. Hibernate A clamps the pH at 7.3 and contains the same amino acids, salts and vitamins as the culture medium. 20 μm polystyrene beads (Polysciences, Warrington, PA) were added to approximate the neuron diameter and create a fixed space between two coverslips to reduce evaporation and provide a constant optical thickness. They also served as a focusing aid. 350 nm light was used to excite NADH using a 75 W Xenon arc lamp (Deltaram V, slits open 2 turns each; (Photon Technology International (PTI), Birmingham, NJ) attached to a dual Photometer model D 104 controlled by Felix GX software (PTI) and observed under Nikon 40x oil objective. The dichroic used for simultaneously exciting NADH and FAD was a 450 dcxru (Nikon, Japan) in a Nikon Diaphot 300 microscope with a beam splitter having a 490 nm DCLP dichroic with 460 nm bandpass emitter filter (#S460/36 Chroma, Brattleboro, VT) for NADH and a 490 nm longpass filter for FAD. Since emission spectra for NADH and NADPH overlap, we use the general term NAD(P)H to indicate that fluorescence was generated from either or both nucleotides. The photomultiplier voltage for NAD(P)H was set at 1000 V and FAD at 850 V. Each cell was isolated with a constant 20 μm square slit and measured for one sec every 6 seconds over 60 sec. 5–6 cells were imaged in a programmed order with the help of an automated stage (Ludl electronic products Ltd., Hawthorne, NY). In order to convert fluorescent intensities (counts/second) to μM concentrations, the system response was calibrated to a concentration curve (0 –200 μM) of NAD(P)H and FAD (Parihar et al, 2008). Background fluorescence was subtracted before analyzing the results for NAD(P)H, FAD concentration and redox ratio (NAD(P)H/FAD).
To measure NADH regenerating capacity, we determined the difference of NADH in neurons with 5 μM rotenone to block consumption at complex I (maximum) from NADH after treatment with 1 μM oligomycin to block ATP synthase activity followed by 1 μM FCCP to uncouple the respiratory chain to drive maximum NADH consumption (minimum NADH).
Nicotinamide (Sigma, St. Louis, MO) was prepared in Neurobasal A, 0.5 mM Glutamax. Neurons were treated for 15–16 hours with or without nicotinamide.
Extraction, derivitization, and analysis of brain NAD/NADH was adapted from Klaidman et al. (1995). Anesthetized animals were decapitated into liquid nitrogen to rapidly quench all metabolism. Approximately 0.4 g of combined hippocampus and cortex was dissected in Hibernate A at 4°C. The tissue was weighed and homogenized in a 1.5 mL tube of 0.2 M KCN, 0.06 M KOH, 1 mM bathophenanthroline disulfonic acid (BPDS, Sigma 146617) with a plastic pestle (K749520-0000, Fisher Scientific) on ice and reacted for 5 min to produce stable cyanide derivatives of the adenine nucleotides. The derivitized homogenate was extracted with chloroform and centrifuged at 17,000 g for 5 min at 4°C. The upper layer was again extracted with chloroform and centrifuged. The aqueous portion was then filtered through a 0.45 um nylon filter (8170, CoStar Spin-X, Corning, NY) by centrifuging at 14,000 g for 10 min at 4°C. The filter tube was then rinsed with 30 μL of 0.2 M KCN, 0.06 M KOH, 1 mM BPDS and spun for an additional 5 min to recover residual sample. Samples were stored at −70°C until analysis. Before injection on to the column, samples were diluted with 1/16th part 1.6 M ammonium acetate, pH 6.0 (8x mobile phase). An injection volume of 20 μL was applied to a 250 × 4.6 mm, 5 μm ProteCol-GP C18–125 reverse-phase column (250210, SGE, Austin, TX). Analytes were detected by fluorescence (Ex 330, Em 460 nm) on a Waters Breeze 2 HPLC System (Milford, MA). The mobile phase consisted of two solutions. Solvent A was 0.2 M ammonium acetate (Fisher Scientific), pH 6.0. Solvent B was 50% 0.2 M ammonium acetate, pH 6.0 and 50% high-performance liquid chromatography (HPLC) grade methanol. A gradient of mobile phase began as 92% solvent A, 8% solvent B. After 1 min, solvent B was increased at a rate of 0.4% for 25 min. The flow rate was 1 mL/min. Freshly made 1.5 mM standards of NAD and NADH were diluted to 5–30 μM with 0.2 M KCN, 0.06 M KOH, 1 mM BPDS and reacted for 5 min. Derivitized standards were diluted with 1/16th part 1.6 M ammonium acetate, pH 6.0. We used the Nernst equation to calculate the redox state of the tissue, Eh = E0 − 2.3(RT/nF)log([NADH]/[NAD]), with the standard potential relative to hydrogen, E0 = −370 mV, the gas constant R=1.987 cal/(degree × mol), T= 310° K, n=1 electron transfer, and the Faraday constant, F = 23.062 cal/(mol × mV).
To determine ROS levels and antioxidant glutathione defenses, we followed the protocol as used before (Brewer et al., 2010) with few changes. At 8 days in culture, 100% of the media containing anti-oxidants in the B27 was replaced with Neurobasal A/Glutamax without B27 for 15 hours at 37°C, 5 % CO2 and 9% O2. Prior to experiments, cells attached to 15 mm glass cover slips were mounted in a slip holder in 800 μL of Neurobasal A, Glutamax, 20 μM 2′, 7′-dichlorofluorescein diacetate (DCFDA, # 0399, Invitrogen) for 20 min. During the last 5 minutes of DCF incubation, 100 μM monochlorobimane (MCB, # M1381, Invitrogen) was added to measure glutathione (Hulbert and Yakubu, 1983). After incubation, cells were rinsed twice with Hibernate A low fluorescence, Glutamax containing 4.6 μg/ml propidium iodide (PI) to stain any dead cells. Dead cells were not included in the analysis due to their high ROS and low glutathione signal. Cells were imaged through a 40x objective/NA 0.60 using Olympus DAPI (MCB), FITC (DCF), and TRITC (PI) optics. Image-pro plus software (version 7.0, Media Cybernetics, Bethesda, MD) was used for analysis of fluorescence intensity.
Live adult neurons were imaged for intracellular calcium as described (Brewer et al., 2006). Briefly, neurons were loaded with 5 μM Fura 2-AM (Molecular Probes, Eugene, OR) for 45 min in Krebs buffer (100 mM NaCl, 20 mM KCl, 1 mM MgCl2, 1 mM CaCl2, 1 mM NaH2PO4, 4.2 mM NaHCO3, 10 mM glucose, 12.5 mM MOPS, pH 7.3) in 5% CO2, 9% O2, followed by de-esterification for 30 min in Kreb’s buffer. 20–30 neurons were imaged using easy ratio pro software (PTI) and CoolSnap HQ 25 CCD camera (Photometrics, Tuscon). Fluorescence was excited alternatively between 340 nm and 380 nm for 400 ms using a Fura2/rhodamine filter (Chroma) and Nikon 40x oil objective.
Neurons cultured for 8 days attached to 15mm glass coverslips were fixed with 4% paraformaldehyde in PBS (minus Mg and Ca) for 10 minutes. Then, a blocking and permeabilization solution of 1% Triton X-100, 1% BSA in PBS was applied to the slips for 10 min. Primary antibodies, rabbit anti-OH-nonenal IgG (#ALX-210-767, Enzo Life Sciences, Plymouth Meeting, PA) and mouse anti-8-OH-guanosine (#12501, QED Biosciences, San Diego, CA), were diluted with 0.02% Triton X-100, 1% BSA in PBS 1:500 and 1:82.5, respectively. The primary antibodies were allowed to bind overnight at 4°C. Secondary antibodies, Alexafluor 488 goat anti-rabbit (Invitrogen) diluted 1:300 and Alexafluor 568 goat anti-mouse (Invitrogen) diluted 1:2000, were allowed to bind for 1 hr at room temperature. Bisbenzamide, final concentration 0.33 μg/mL, was applied to slips for 2 min to label nuclei. Neurons used as a positive control were treated with 20 μM hydrogen peroxide (H2O2) for 10 minutes. Coverslips were mounted on a slide and imaged at 60X with an oil immersion objective using Olympus dichroic filters for DAPI (bisbenzamide), FITC (OH-nonenal) and TRITC (OH-guanosine). Image-pro plus software was used for analysis of fluorescence intensity with constant thresholds for each age and genotype. For experiments without antioxidants, neurons were cultured with B27 without antioxidants (Invitrogen # 10889-038). Total fluorescence per cell was calculated as fluorescence intensity x area of cell (μm2)/1000.
Neurons were rinsed with PBS and fixed with 4% paraformaldehyde in PBS (minus Mg and Ca). After blocking with 3% BSA, 0.05% saponin in PBS, cytochrome C levels were detected using sheep anti-rabbit cytochrome C (#C5723, Sigma; diluted 1:200) and Alexafluor 488 donkey anti-rabbit antibodies (#A11015, Invitrogen; diluted 1:2000). Images were acquired at 100x and analyzed using Image Pro Plus software. Images were segmented into bright and dim objects. The bright intensity objects measured cytochrome C concentrated in mitochondria while the dim intensity objects measured the cytoplasmic cytochrome C.
Data are presented as means and standard errors. Student’s T test was used to assess the difference of means using Prostat (Poly Software, Pearl River, NY). We used p< 0.05 to reject the null hypothesis.
In order to evaluate neuron function with age, independent of an aging hormonal, immune and vascular system, we isolated neurons from the frontal cortex and hippocampus of mice across their age-span. Figure 1 shows that non-Tg (Figure 1A) and 3xTg-AD (Figure 1B) neurons in uniform culture medium have similar healthy morphology. There is also no difference between viability in non-Tg and 3xTg-AD neuronal cultures over the entire age-span of the mouse. Neuronal viability also does not decrease with age in uniform cultures (Figure 1C).
Since brain deficits in the primary electron donor (NADH) to the electron transport chain would severely limit the capacity to generate ATP, we examined the NAD(P)H levels in isolated neurons as a function of age or AD-like genotype. Since autofluorescence of NADH cannot be distinguished from NADPH in our system, we used the term NAD(P)H for our measurements. In order to determine the NAD(P)H concentration of non-Tg and 3xTg-AD cultured neurons, intrinsic fluorescence of NAD(P)H was measured microscopically from individual neurons with detection by split-beam photomultipliers to simultaneously measure the reduced NAD(P)H and the oxidized FAD. Figure 2A shows the NAD(P)H concentration in live neurons from both non-Tg and 3xTg-AD mice at different ages starting from sexual maturity at 2 months, through middle-age at 11 months and into old-age at 21 months, near the median life-span. In non-Tg mouse neurons (Figure 2A), the NAD(P)H started at a low concentration (60 μM) that rose 85% with development to 111 μM at 11 mo. middle-age, after which the NAD(P)H concentration dropped 33% to 77 μM in old-age. Although 3xTg-AD mouse neurons started at the same NAD(P)H concentration as non-Tg (58 μM), NAD(P)H concentration increased toward middle-age by only 21%, then began to decline earlier after 8 months by 56% to 30 μM in old-age. The final NAD(P)H concentration in 3xTg-AD neurons was only 39% that of non-Tg neurons.
The ratio of the reduced [NAD(P)H] divided by the oxidized [FAD] is a useful proxy for the redox state of the cell, more precisely indicated by the ratio of the [NADH]/[NAD+] (Chance and Williams, 1956; Chance et al., 1979). The use of the FAD measure is necessitated by the non-fluorescent properties of NAD+ while FAD and NAD(P)H exhibit intrinsic fluorescence microscopically measurable in single cells. Figure 2B shows the ratio of reduced [NAD(P)H]/oxidized [FAD] (redox ratio) in neurons of both genotypes across the age-span. In non-Tg neurons, the redox ratio mimics the NAD(P)H profile with a shift toward a reduced redox peak at middle-age and an oxidized shift in old-age. In contrast to non-Tg neurons, the redox ratio in the 3xTg-AD neurons declined continuously with age toward a 62% more oxidized redox state. These results suggest that the redox state of live single neurons varies with age and genotype in large reproducible ways with potential impact on numerous metabolic functions.
Since aging and the 3xTg-AD genotype caused a lower NAD(P)H concentration and a more oxidized redox state, we wanted to investigate whether this lower NAD(P)H in 3xTg-AD neurons was due to higher NADH consumption by complex I of the electron transport chain or lower NADH regenerating capacity by lactate and pyruvate dehydrogenases and Kreb’s cycle dehydrogenases compared to non-Tg neurons. Either way, the imbalance would produce a more oxidized redox state insufficient to withstand a challenge of oxidative stress. We compared NADH levels under two conditions. First, to minimize NADH levels and maximize consumption, we added oligomycin to block reversal of ATPase and FCCP to dissipate the proton gradient and to maximally consume NADH by stimulation of complex I, (NADH dehydrogenase), the major consumer of NADH for electron transport down the chain to oxygen. Secondly, we blocked complex I with 5 μM rotenone to minimize NADH consumption and maximize NADH levels. At 2 months for both genotypes, the resting NADH levels were near those with FCCP added, indicating that the youngest neurons were operating near their maximum capacity to consume NADH. At all subsequent ages, the resting levels of NADH were near the midpoint between uncoupled FCCP levels and consumption-inhibited rotenone levels (Figure 2C).
The difference between the maximum and minimum NADH concentrations evoked by rotenone and FCCP provides a measure of the NADH regenerating capacity. Figure 2D shows that both genotypes showed an increase in NADH regenerating capacity until middle-age after which they dramatically declined to similar low levels in old-age. Compared to non-Tg neurons, 3xTg-AD neurons showed significant deficits in NADH regenerating capacity from the earliest age of 2 months. However, this early deficit was amplified at 4, 8 and 11 months, suggesting that NADH regenerating capacity is further impaired with age. Since FCCP lowers NADH levels and rotenone raises NADH levels in all ages and both genotypes, complex I utilization is not impaired in either the 3xTg-AD or the non-Tg neurons.
Nicotinamide, a precursor of NAD+ (Ying, 2008), reverses learning deficits in the 3xTg-AD mouse (Green et al., 2008) with a mechanism attributed to epigenetic changes based on acetylation of phospho-tau Thr231. Here, we consider an alternative mechanism in which nicotinamide is first an NAD+ and then an NADH precursor to supply more energy to neurons. Treatment of 21 month old neurons with increasing doses of nicotinamide for 15 h. produced an inverted U-response in neuron NAD(P)H levels and redox ratios (Figure 3A, B). 2 mM nicotinamide produced a maximum response. In 21 month non-Tg neurons, nicotinamide raised the [NAD(P)H] from 76 to 95 μM and produced a reductive shift in the redox ratio from 4.3 to 5.4, increasing both concentration and redox ratio by 25 % (Figure 3A, B). Nicotinamide treatment of 21 month 3xTg-AD neurons led to an increase in [NAD(P)H] from 30 to 54 μM and in redox ratio from 1.8 to 2.5, leading to larger increases of 80% in NAD(P)H and 39% in redox ratio respectively, but failed to restore these neurons to levels present in non-Tg neurons. Treatment of 11 month neurons with 2 mM nicotinamide produced non-Tg shifts from 110 to 129 μM NAD(P)H and 3xTg-AD shifts from 68 to 109 μM NAD(P)H. Treatment of 2 month neurons with 2 mM nicotinamide produced non-Tg shifts from 61 to 86 μM NAD(P)H and 3xTg-AD shifts from 58 to 80 μM NAD(P)H.
To determine the effect of nicotinamide on the regenerating capacity, we treated cells with FCCP and rotenone as before. We observed that in 21 month animals of both genotypes, nicotinamide treatment increased the maximal NADH (rotenone added), but had no effect on the minimal NADH levels (FCCP added) (Figure 3C). Similarly, treatment of 2 and 11 month neurons of either genotype with nicotinamide did not change the minimum NADH levels generated by FCCP (data not shown), suggesting complex I was not impaired in the 3xTg-AD neurons. However, as in Figure 3C for 21 month neurons and for 2 and 11 month neurons (data not shown), the maximum levels of NADH in the presence of rotenone were increased. The net result in Figure 3D, E show the effect of 2 mM nicotinamide on the NADH regenerating capacity. At 2, 11 and 21 months, treatment with nicotinamide increased the regenerating capacity similarly for both 3xTg-AD and non-Tg neurons. These results suggest that NADH regenerating capacity is impaired in 3xTg-AD neurons, but can be fully restored to non-Tg levels by 15 hr. treatment with nicotinamide at either 2 or 21 months.
Increased ROS levels is one of the major co-pathologies in aging and AD (Emerit et al., 2004; Carney et al., 1994). Since an NAD(P)H oxidized redox state in 3xTg-AD neurons and in old-age non-Tg neurons also decreases its anti-oxidant ability, we wanted to determine the relationship of impaired NAD(P)H levels to ROS levels across the age-span as well as simultaneous measures of the antioxidant buffer glutathione. The oxidized redox state could result from increased ROS levels and/or decreased glutathione defenses. We used fluorescent probes dichlorofluorescein diacetate (fluorescent product DCF) to detect ROS levels (Parihar and Brewer, 2007) and monoclorobimane (MCB) to label glutathione production (Hulbert and Yakubu, 1983). Although DCF does not react directly with singlet oxygen, it can react with several reactive species like OH•, NO2•, peroxynitrites etc (Kalyanaraman et al., 2012) and damage macromolecules. ROS and glutathione levels from live non-Tg and 3xTg-AD neurons from 8 month brains are shown in Figure 4(A, B). Qualitatitively, non-Tg neurons (Figure 4A) show stronger intensity of glutathione label than 3xTg-AD neurons (Figure 4B) with a merge as light blue neurons. The reverse was observed for ROS with non-Tg neuron levels low (Figure 4A) and higher intensity levels in 3xTg-AD neurons (Figure 4B) with the merged image showing the stronger ROS green stain.
Compared quantitatively, the ROS levels of 3xTg-AD and non-Tg neurons began at low and equal levels at 2 and 4 months of age (Figure 4C). However, after 4 months, ROS levels increase sharply for both genotypes, with a significantly faster increase with age for 3xTg-AD than non-Tg neurons. The old-age level of ROS levels for non-Tg neurons was already attained at 8 months in the 3xTg-AD neurons. In the same cells, we simultaneously determined the levels of glutathione (GSH) as the primary cellular anti-oxidant buffer. We hypothesized that increased ROS would be buffered by increased glutathione, but in old age, either production of GSH would decline or more GSH would be consumed by even higher ROS. Further, we predicted that the 3xTg-AD neurons with their increased energy demands from birth due to higher resting calcium levels would reach this critical point of insufficient ROS buffering at an earlier age. Figure 4D shows a significant deficit in GSH levels in 3xTg-AD neurons relative to non-Tg neurons starting at 2 months of age. While both genotypes increased their GSH levels as ROS increased through middle-age, the increase for 3xTg-AD neurons was significantly less. Both genotypes produced lower levels of glutathione from middle to old age, reaching levels below those at the start of life. Compared to non-Tg neurons, GSH levels in 3xTg-AD neurons remained significantly lower at all ages.
These simultaneous measures allow comparisons of the relative aging kinetics of increases in ROS and declines in glutathione buffering of ROS. Focusing first on non-Tg neurons, Figure 4E shows that glutathione levels generally increase at the same rate as ROS levels until middle age. In old age, glutathione levels decline much faster, creating an arbitrary crossover point at 17 months. In 3xTg-AD neurons, a similar crossover point occurs at only 7 months of age (Figure 4F). The larger rise in ROS in the 3xTg-AD neurons from 4–11 month, compared to non-Tg may contribute to the failure of GSH to rise over this period. The further rise in ROS from 11 to 21 month in both genotypes, which correlates with a greater decline in GSH over this period, highlights the need for further studies regarding a threshold level of ROS, above which GSH is catastrophically consumed. This experiment suggests that 3xTg-AD deficits in glutathione occur before excess ROS in 3xTg-AD neurons that become more acute as ROS levels increase and glutathione buffers are consumed into middle and old ages.
Since nicotinamide partially reversed the oxidative redox shift toward a reduced redox state and increased NAD(P)H concentration, we hypothesized that nicotinamide treatment would lower ROS levels by first increasing GSH regeneration from its oxidized form GSSG via NADPH-dependent glutathione reductase followed by GSH reduction of H2O2 by glutathione peroxidase and other reactions. The first reaction may not be detectable by measuring GSH regeneration if a cell is at steady state equilibrium of GSH/GSSG. To test our hypothesis, we chose the 8-month time point since GSH levels in 3xTg-AD neurons declined after 8 months. Figure 5A shows that pre-treatment with nicotinamide for non-Tg neurons did not alter GSH levels, while in 3xTg-AD neurons nicotinamide increased GSH levels by 30%. Simultaneous measures of ROS levels (Figure 5B) revealed that non-Tg neurons did not alter their ROS levels in response to nicotinamide, treated 3xTg-AD neurons showed a 20% decrease in the ROS level. However, despite this significant decrease in ROS levels in 3xTg-AD neurons, the overall ROS levels remained significantly higher than the non-Tg neurons. One possible explanation why ROS is not restored to the same levels as in non-Tg neurons is that other ROS sources remain unaffected by nicotinamide-enhanced NAD(P)H. Also, the unaltered GSH levels in non-Tg neurons treated with nicotinamide suggests that non-Tg neurons are already regenerating enough GSH from GSSG to maintain a normal redox state.
Since glutamate excitoxicity is implicated with aging and AD, we wanted to determine further if nicotinamide can rescue neurons from glutamate induced excitotoxicity. We chose our middle-aged (11-month) and old-aged (21-month) animals to compare neuron viability. Figure 6 shows percentage dead in 11-month (Figure 6A) and 21-month (Figure 6B) neurons of both genotypes with 100 μM glutamate and 2 mM nicotinamide treatment. Compared to untreated neurons, glutamate treatment induced a 2-fold increase in dead neurons in both non-Tg and 3xTg-AD neurons. However, co-treatment with nicotinamide and glutamate, largely protected neurons from glutamate excitotoxicity (p < 0.001 for 11 month; p < 0.02 for 21 month). Although nicotinamide reduced the glutamate toxicity in non-Tg neurons to that of untreated neurons, the treatment only reduced by 50% the toxicity in 21 month 3xTg-AD neurons. This increased susceptibility of 3xTg-AD neurons to glutamate toxicity suggests that factors other than NAD(P)H and GSH are involved in AD-like toxicity since these neuroprotectants appear to be fully restored by nicotinamide.
3xTg-AD neurons have 3–5 fold higher intracellular calcium concentration (Smith et al., 2005a; Stutzmann et al., 2006) at birth. To confirm calcium dysregulation in adult neurons, we performed intracellular calcium imaging of 11 month non-Tg and 3xTg-AD neurons. We found that while the [Ca]i in non-Tg neurons was 109 nM, 3xTg-AD neurons had more than 2 fold higher [Ca]i at 253 nM (data not shown), which confirmed the previous study in primary neurons, cultured from 8–12 month 3xTg-AD mice (Lopez et al., 2008).
Mitochondrial function is impaired in both aging and Alzheimer’s disease. However it remains unanswered if mitochondrial dysfunction causes the rise in ROS and oxidative shift in redox or whether ROS causes the mitochondrial dysfunction and oxidative shift? One line of evidence for causation is precedence: if event A precedes event B, then B cannot cause A. In our model system, to hypothesize that mitochondrial dysfunction causes a redox shift, early changes in mitochondrial function such as NADH regenerating capacity would precede redox changes and/or ROS. Since healthy mitochondria retain and stressed mitochondria release cytochrome C, we determined the effect of age and genotype on neuronal and mitochondrial health.
According to hypothesis A, the increased genotypic stress on mitochondria causes increased release of cytochrome C from mitochondria into the cytoplasm with age. Examination of the mean cytoplasmic cytochrome C in Figure 7A shows low levels at 2 months followed by increased cytoplasmic levels in non-Tg and even higher levels in 3xTg-AD neurons at middle-age that remained high at old-age. This increase in cytoplasmic cytochrome C could lead to apoptosome activation. Hypothesis B was that 3xTg-AD neurons have fewer healthy mitochondria because they are poisoned by the excess beta-amyloid and calcium so that those remaining have to work harder. A corollary is that harder working mitochondria produce more ROS and/or consume more NADH. In contradiction to this hypothesis, Figure 7B shows a higher mitochondrial count/cell in 3xTg-AD neurons than non-Tg, in early and middle-age. These results suggest that the 3xTg-AD neurons compensate for the increased genotypic stress by upregulating mitochondria in each cell. Higher measures of mitochondria/cell in 3xTg-AD compared to non-Tg neurons supports a compensatory bioenergetic mechanism in 3xTg-AD neurons. This higher mitochondrial count/cell in 3xTg-AD neurons is closely paralleled in Figure 7C by a higher early and middle-age intensity of bright-level cytochrome C immunoreactivity in the 3xTg-AD neurons than non-Tg, likely associated with mitochondria.
If ROS levels increase with age or genotype beyond the detoxification ability of the cell, then nucleic acid and lipid macromolecular damage will occur (Reed et al., 2009). In order to assess ROS-induced macromolecular damage in these cultured neurons as a function of mouse age and genotype, Figure 8A shows 11 month neurons double stained for oxidative damage to DNA and RNA using anti-8-OH-(deoxy)guanosine and for lipid peroxidation using anti-4-hydroxynonenal in the same neurons. To demonstrate the reliability of these probes and cellular responses, we also treated 4 month neurons with 20 μM H2O2 for 10 min to induce excess oxidative damage to the cells (positive control). Probes for both nucleic acid damage and lipid peroxidation dramatically increased following the H2O2 treatment. Despite an increase in ROS and oxidized redox state shown in Figure 4C and Figure 2B, we see in Figure 8(B,C) no indication of ROS-induced macromolecular damage in cultured neurons with age or genotype. This suggests that the anti-oxidants in the cells and in the culture medium were sufficient to protect against significant ROS damage to macromolecules. As an additional control, we removed the SOD, catalase, glutathione, vitamin E acetate and vitamin E from the culture medium. This external manipulation was sufficient to elicit both nucleic acid and lipid damage to 3xTg-AD neurons above those of non-Tg neurons from 4 month old brains, while the 2 month neurons did not show any genotypic difference even with antioxidant removal (Figure 8E).
In order to observe the redox state of brain tissue in vivo, which includes neurons with the support of other surrounding cell types, we analyzed hippocampal and cortical brain lysates by HPLC. Figure 9A shows a 22% decrease in NADH concentration in 2-month 3xTg-AD brain. The redox state of 3xTg-AD brain from 2 month animals (Figure 9B) was significantly more oxidized (−372 mV) than non-Tg brain (−387 mV). These results indicate that the redox state in the whole 3xTg-AD brain reflects an oxidative shift as seen in the cultured neuron model.
The ability of neurons from old animals to regenerate in culture and their lack of abundant ROS damage argue against the theory that aging results from an irreversible lifelong buildup of ROS damage. Nevertheless, neurons from old non-Tg and 3xTg-AD mice showed deficits in NAD(P)H levels, lower capacity to generate NAD(P)H, lower glutathione levels, late elevated ROS and fail to increase mitochondria to meet these energy demands. These aging effects indicate an early oxidized redox shift before ROS damage to macromolecules. Further, because of the rapid reversibility by nicotinamide of the oxidized redox shift and glutamate toxicity in neurons from old animals, we argue that processes associated with aging cause an important metabolic shift in reductive supply of energy as NADH to the electron transport chain. We (Brewer, 2010) and others (Martin et al., 2010; Anderson and Weindruch, 2010; Safdar et al., 2010) have hypothesized that a sedentary lifestyle causes this metabolic shift.
Regarding the AD model, we demonstrated that the redox ratio in the 3xTg-AD neurons never increased beyond 2 month levels and that NAD(P)H levels never increased above those of 4 month non-Tg neurons. Therefore, we predicted that restoration of NAD(P)H levels would reverse the low energetic capacity. Since these genotype deficits could be reversed in less than 24 hr. with the NAD+ precursor, nicotinamide, we argue that the age-related shift in redox state is a critical target that the AD-like transgenes exacerbate beyond normal aging. Further, the 3xTg-AD model shows impairment in GSH regeneration from 2 months on, never reaching levels of middle-age non-Tg neurons. Therefore, the supply of key metabolites NADH and GSH may restore sufficient reducing energy and anti-oxidant capacities to counter the impact of increased transgene expression.
These results support the novel proposition that an oxidative redox shift precedes increased ROS damage, cell damage and cognitive decline with age and in a mouse model of AD. In the absence of ROS damage, we found a robust oxidative shift in NAD(P)H in non-Tg old age that began earlier in the 3xTg-AD neurons. The capacity to generate NADH under stress showed substantial deficits in 3xTg-AD neurons beginning at 2 months of age. Compared to the non-Tg from the earliest 2 month age, we also found 3xTg-AD neurons deficient in GSH which continued throughout the lifespan. Both NAD(P)H and glutathione levels, declined in old age in both genotypes. We also observed that in 2 month old brain extracts, 3xTg-AD had a significantly lower NADH concentration and a more oxidized redox state than the non-Tg. ROS levels were low before 8 months for both genotypes then increased until middle age, but increases were again reversible with nicotinamide. Surprisingly, macromolecular ROS damage was not detected with age or genotype in our in vitro model, unless environmental anti-oxidants were removed (reduced glutathione, vitamin E, vitamin E acetate, catalase and superoxide dismutase). However, in brain extracts, malondialdehyde, hydroxynonenal and TBARS levels were elevated in the 3xTg-AD relative to non-Tg mouse (Resende et al., 2008). This observation, together with an elevated SOD and glutathione peroxidase suggests an insufficient antioxidant response in the 3xTg-AD brain in vivo. Interestingly, oxidized glutathione was increased and the enzyme to recycle to GSH, glutathione reductase, was not increased in the 3xTg-AD brain. These observations could be related to our studies since NADPH is a cofactor for glutathione reductase in the regeneration of GSH from GSSG. Thus, our measures of lower NAD(P)H in the 3xTg-AD neurons would deny sufficient reducing power to the enzyme needed to recycle the GSSG to GSH and maintain redox balance. If the extracellular anti-oxidants were removed, then 3xTg-AD neurons have significantly higher oxidative damage than non-Tg neurons, as in whole brain.
Several factors could contribute to a decreased NAD(P)H and an oxidized redox state with age and AD-like genotype including downregulation of TCA cycle enzymes that generate NADH (Bubber et al., 2005), lower rates of glycolysis with age (Patel and Brewer, 2003) and AD (Hoyer et al., 1988; Hoyer et al., 2005) and high energy demands with age and genotype. In addition to glyceraldehyde-3-phosphate dehydrogenase in glycolysis, important sources of NADH are pyruvate dehydrogenase at the entry to the TCA cycle and within the TCA cycle, isocitrate dehydrogenase, alpha-ketoglutarate dehydrogenase and malate dehydrogenase. Down-regulation of any of these enzymes could be responsible for the decreased NADH levels in the 3xTg-AD neuron (Bubber et al., 2005). Yao et al. (2009) observed a significant decline in pyruvate dehydrogenase (PDH) and cytochrome c oxidase (COX) activity in mitochondria isolated from 9 and 12 month 3xTg-AD brain compared to non-Tg. The levels of PDH protein were also significantly lower in 3xTg-AD from the young age of 3 month to middle age of 12 month, while COX levels were significantly lower than non-Tg at 9 and 12 months. In younger embryonic neurons from 3xTg-AD mice, maximum respiration stimulated by FCCP was impaired in the 3xTg-AD compared to the non-Tg (Yao et al., 2009). Thus, the availability of NADH for electron transfer is restricted in 3xTg-AD mouse. In neurons from aged rats, our lab has also shown by flux control analysis of the electron transport chain that respiration was limited at the substrate level (NADH) for the NADH-ubiquinone oxidoreductase (Jones and Brewer, 2010). Consequently, a more oxidized environment will contribute to higher rates of ROS generation. This corresponds to our result of decreased NAD(P)H capacity and excessive ROS after middle age.
Nicotinamide, a biosynthetic precursor of NAD+, was previously shown to increase NADH and NADPH levels in striatum and thalamus of whole brain tissue of adult mouse (Klaidman et al., 2001). Nicotinamide also improved cognitive deficits in hippocampal and amygdala dependent tasks in 3xTg-AD mice associated with decreased Thr231 phospho-tau, increased levels of the stabilizing form of acetylated α-tubulin and increased synaptogenic p25, with improved learning and memory (Green et al., 2008). In our study, we found that nicotinamide can in fact, improve earlier age-related deficits in NAD(P)H concentration, improve redox ratio to a more normal reduced level, as well as increase NADH regenerating capacity. 3xTg-AD mice have an elevated intracellular Ca2+ concentration from birth due to the mutant presenilin (Stutzmann et al., 2006; Smith et al., 2005b), supporting the hypothesis that elevated calcium contributes to AD (Khachaturian, 1994; Mattson et al., 2000). Calcium can disrupt the mitochondrial membrane potential and increase opening of the mitochondrial transition pore precipitating cell death. Physiologically elevated calcium also upregulates the NADH-generating TCA cycle enzyme, α-ketogluterate dehydrogenase (KGDH) in a concentration-related manner (Lai and Cooper, 1986). However, at higher concentrations, calcium inhibits KGDH activity. Thus, age-related elevated calcium in 3xTg-AD neurons could downregulate KGDH activity with less NADH generation.
While NAD(P)H acts as redox energy currency, GSH acts as a dynamic redox energy buffer (Das and White, 2002) with millimolar concentrations in rat brain (Kudo et al., 1990; Das and White, 2002) compared to micromolar NADH (Gupte et al., 2005). An age-related decrease in GSH is associated with neurodegenerative diseases like Parkinson’s disease and AD (Schulz et al., 2000) which could contribute to increased ROS levels. It is possible that 3xTg-AD neurons are unable to generate more GSH due to the decline in NADPH and eventually cannot keep up with higher ROS levels. The reversal of low NAD(P)H, GSH and high ROS in 8 month 3xTg-AD neurons by nicotinamide suggests that these redox agents are interlinked.
Cytochrome C is an essential electron carrier in the mitochondrial respiratory chain. We observed that at 2 months, 3xTg-AD neurons already had a significantly higher number of mitochondrial cytochrome C than the non-Tg neurons. Interestingly, though the mitochondrial count per neuron was high in 3xTg-AD, the NAD(P)H concentration was the same for both non-Tg and 3xTg-AD neurons. The higher mitochondrial count in 3xTg-AD neurons may be needed to compensate for inefficiency in ATP generation due to elevated calcium (Nicholls, 2009) or A-beta exposure (Alberdi et al., 2010), either of which would depolarize the mitochondrial membrane potential to lower energy to drive ATP synthesis. However, with age, 3xTg-AD mitochondria have lower NAD(P)H concentrations suggesting that a depolarized membrane potential drives higher rates of NADH-ubiquinone oxidoreductase that lower NADH without sufficient regeneration.
Here, we were able to show that an oxidized redox state (NAD(P)H/FAD), lower NADH regenerating capacity and lower GSH levels precede excessive ROS in 3xTg-AD compared to non-Tg, a clear effect of redox changes with age on the intrinsic physiology of neurons from the AD model. We also showed that macromolecular damage in neurons was higher in 3xTg-AD neurons when antioxidants were removed from the culture media, but not until 4 months of age. Nicotinamide, which was previously shown to increase NAD(P)H in mouse brain, could also reverse the oxidized redox state to a more reduced state, increase GSH levels and decrease ROS in the neurons. In elegant studies in rats virally transfected with either SOD1 or catalase, age-related learning deficits were also dissociated from accumulated damage (Lee et al., 2011). Since increased ROS and macromolecular damage along with decreased GSH are hallmarks of AD, any reversal and possible prevention of this damage with nicotinamide may have clinical significance.
This work was supported by NIH grants R01 AG032431, AG13435 and the Stark Endowed Chair for Alzheimer’s Research. We thank Salvatore Oddo and Frank LaFerla for contributing the founder mice used in this study.
No conflict of interest
Author contributions: Ghosh designed and performed research, analyzed data and wrote the paper. LeVault performed research, analyzed data and wrote the paper. Barnett performed research, analyzed data and wrote the paper. Brewer directed the project, designed experiments and wrote the paper.