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BioMed Research International
Volume 2014 (2014), Article ID 371593, 13 pages
Alcohol Induced Mitochondrial Oxidative Stress and Alveolar Macrophage Dysfunction
1Department of Pediatrics, Emory University, Atlanta, GA 30322, USA
2Children's Healthcare of Atlanta Center for Developmental Lung Biology, Atlanta, GA 30322, USA
3Division of Pulmonary, Allergy & Critical Care Medicine, Emory University School of Medicine, Atlanta, GA 30322, USA
4Division of Neonatal-Perinatal Medicine, Department of Pediatrics, Emory University, 2015 Uppergate Drive, Atlanta, GA 30322, USA
Received 30 April 2013; Revised 27 December 2013; Accepted 28 December 2013; Published 19 February 2014
Academic Editor: Victor J. Thannickal
Copyright © 2014 Yan Liang et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.
An alcohol use disorder increases the risk of invasive and antimicrobial resistant community-acquired pneumonia and tuberculosis. Since the alveolar macrophage (AM) orchestrates the immune response in the alveolar space, understanding the underlying mechanisms by which alcohol suppresses AM phagocytosis is critical to improving clinical outcomes. In the alveolar space, chronic alcohol ingestion causes severe oxidative stress and depletes antioxidants which are critical for AM function. The mitochondrion is important in maintaining cellular redox balance and providing the ATP critical for phagocytosis. The focus of this study was to understand how alcohol triggers mitochondrial reactive oxygen species (ROS), stimulates cellular oxidative stress, and induces AM dysfunction. The current study also investigated the capacity of the mitochondrial targeted antioxidant, mitoTEMPOL (mitoT), in modulating mitochondrial oxidative stress, and AM dysfunction. Using in vitro ethanol exposure and AMs from ethanol-fed mice, ethanol promoted mitochondrial dysfunction including increased mitochondrial ROS, decreased mitochondrial membrane potential, and decreased ATP. Treatment with mitoT reversed these effects. Ethanol-induced decreases in phagocytosis and cell viability were also attenuated with mitoT. Therefore, antioxidants targeted to the mitochondria have the potential to ameliorate ethanol-induced mitochondrial oxidative stress and subsequent decreases in AM phagocytosis and cell viability.
Both acute and chronic alcohol consumption have well-documented effects on the immune system leading to increased susceptibility to community acquired pneumonia and tuberculosis . When subjects with an alcohol use disorder get pneumonia, they are more likely to be infected with serious Gram-negative bacteria  and these increased risks occur even in those who do not meet the diagnostic criteria for an alcohol use disorder . This results in a higher rate of intensive care use, longer inpatient stays, higher healthcare costs, and a 2–4 times greater mortality rate . There is also an increased risk of ventilator-associated pneumonia which worsens the morbidity and mortality rates . Alcohol abuse is also associated with a 2-3-fold increased risk of the acute respiratory syndrome (ARDS), representing ~50% of all ARDS cases with an average age of 30–35 . For subjects without a history of alcohol abuse, pneumonia will lead to sepsis in ~35% of the cases and ~30% will progress to ARDS. In contrast, pneumonia will lead to sepsis in ~60% of the cases if the subject has a history of alcohol abuse and 70% will progress to ARDS .
A seminal feature is that chronic alcohol abuse causes severe oxidative stress in the fluid lining the alveolar space, which includes the depletion of the critical antioxidant glutathione (GSH) and oxidation of the GSH/GSSG redox state by ~40 mV in subjects with an alcohol use disorder [6, 7]. GSH depletion and oxidation within the alveolar space are particularly critical for alveolar macrophages (AM) since they are constantly bathed by this fluid and depend on this GSH pool for cellular uptake and protection against the oxidative stress generated during immune responses. Residing at the inner epithelial surfaces of airway and alveoli, AMs are the only lung phagocytes exposed directly to the environment. Therefore, AMs represent the first line of cellular defense in the lower respiratory tract . However, oxidative stress can impair AM phagocytosis [9, 10]. In addition to impaired clearance of microbes, impaired phagocytosis can cause insufficient clearance of dying or dead cells and lead to pathological inflammation. Therefore, alcohol-induced oxidative stress can be a critical contributor to pulmonary pathophysiology, risk of infection, and contribute to the increased risk of tissue injury associated with ARDS.
There are multiple cellular sources of reactive oxygen species (ROS) including the mitochondria, the cytochrome P450 family, xanthine oxidoreductase, peroxisomes, cyclooxygenases, lipoxygenases, and the family of NADPH oxidases . The consequences of the ROS depend on the type of the ROS generated, the amount of ROS, and where it is generated. Under resting conditions, the majority of the cellular ROS generated is derived from the mitochondria where ~90% of the oxygen used by a cell is consumed during energy metabolism . In this mitochondrial process, nicotinamide adenine dinucleotide (NADH) is oxidized to support electrochemical coupling of oxidative phosphorylation and ATP synthesis [13–16]. However, respiration also generates ROS such as superoxide anions (), hydrogen peroxide (H2O2), and hydroxyl radicals (). To protect against the ROS generated during respiration, mitochondria also maintain redox balance through numerous ROS defense systems including mitochondrial manganese superoxide dismutase (MnSOD), GSH, thioredoxin 2 (Trx2), and catalase . Neutralization of mitochondrial ROS is critical for mitochondrial function and, ultimately, cellular functions but low-level concentrations of ROS are also required for signal transduction . During respiration, the NADH is oxidized to NAD+ and the NAD+/NADH ratio has been recognized as a key regulator in energy metabolism, aging, and immunological functions . For example, decreases in or in the NAD+/NADH are associated with increased production of superoxide by the mitochondria and subsequent alteration of the mitochondrial redox system [20–22].
Alcohol metabolism can interrupt this complex integrated redox system within the mitochondria. Whether it is metabolized by alcohol dehydrogenase or cytochrome P450, the primary metabolite produced during alcohol metabolism is acetaldehyde. Within the mitochondria, acetaldehyde is metabolized by mitochondrial aldehyde dehydrogenase (ALDH2)  which uses NAD+ as a cofactor. Thus, acetaldehyde metabolism decreases mitochondrial NAD+ pools and increases NADH. The resulting decreases in the NAD+/NADH ratio and subsequent increases in mitochondrial ROS can change the mitochondrial redox balance leading to cellular oxidative stress and damage. Our research team has previously demonstrated that chronic alcohol ingestion resulted in impaired phagocytosis by AMs and chronic oxidative stress was central to the impaired immune functions [9, 10], while alcohol-induced upregulation of ROS through NADPH oxidases is linked to impaired phagocytosis; we speculated that there was also a role for alcohol-induced mitochondrial ROS generation in impaired AM phagocytosis. The results presented in this paper demonstrate that chronic alcohol ingestion in a mouse model induced mitochondrial ROS generation and mitochondrial dysfunction which contribute to impaired AM phagocytosis. Treatment with mitochondrial specific antioxidants reversed mitochondrial dysfunction and restored phagocytosis.
2. Materials and Methods
2.1. Mouse Model of Chronic Ethanol Ingestion
All animal studies were performed in accordance with the National Institutes of Health guideline outlined in the Guide for the Care and Use of Laboratory Animals. All described protocols were reviewed and approved by the Emory University Institutional Animal Care and Use Committee. Mice (C57BL/6; age 6–8 weeks) were fed standard laboratory chow ad libitum with incremental increases of ethanol in the drinking water over 3 weeks (5%/week) to a final concentration of 20%. Mice were maintained at 20% ethanol (EtOH) in the drinking water for 10–12 weeks (/group) [24, 25]. The controls were pair-fed in order to control for the calories due to EtOH as well as any differences in food intake. The weight of the chow consumed by the mice with ethanol in the drinking water is routinely determined and this historical data was then used to establish a pair-feeding model for the controls. This regimen produced clinically relevant elevations in blood alcohol concentrations of 0.12% ± 0.03, as published by our group  and others [27, 28]. After euthanasia, tracheas were cannulated and a bronchoalveolar lavage (BAL; three 1 mL of saline) performed. Mouse AMs (mAMs) were then isolated from the fluid by centrifugation at 1000 ×g for 10 min. After differential staining with Diff-Quik (Dade Behring, Newark, DE) and counting with a hemocytometer, the cell population was determined to be ~95% alveolar macrophages. The cell pellet was resuspended in RPMI 1640 medium containing 2% FBS and 1% penicillin/streptomycin and cells were incubated at 37°C in 5% CO2 atmosphere before the experiments outlined below were performed.
2.2. MH-S Cell Culture and EtOH Exposure
The mouse AM cell line, MH-S (American Type Culture Collection, Manassas, VA), was used as a model system for studying the direct effects of EtOH exposure in vitro. Cells were cultured in RPMI 1640 medium containing 10% FBS and 1% penicillin/streptomycin and incubated at 37°C in a 5% CO2 atmosphere. MH-S cells were treated with 0.2% EtOH for 5 consecutive days with the media changed daily. This EtOH concentration (0.2%) is representative of the blood alcohol content (BAC) when a 120 lb person consumes 5 drinks at a single sitting [28–30]. During the last 24 hr of the 5d EtOH treatment, some cells were also treated with the mitochondria-targeted antioxidant mitoTEMPOL (mitoT, 100 μM) .
2.3. Measurement of Intracellular ROS Generation
After EtOH exposure, the cellular ROS sensitive probe CM-H2DCFDA (Invitrogen; Carlsbad, CA) and the mitochondrial superoxide probe mitoSOX (Invitrogen; Carlsbad, CA) were added to the medium (10 μM, 30 min, 37°C). Cells were then harvested, washed, and resuspended in phosphate buffered saline (PBS) for FACS analysis by BD Canto II Flow Cytometer (Becton Dickinson, Franklin Lakes, NJ). CM-H2DCFDA and MitoSOX were excited at 488 nm and detected at nm or nm, respectively. Data analysis was performed using Flowjo (http://www.flowjo.com/).
2.4. Measurement of Mitochondrial Membrane Potential
After the different treatments, the mitochondrial membrane potential was determined by incubating the cells with tetra-methylrhodamine, ethyl ester (10 nM, 30 min, 37°C; TMRE; Sigma, St. Louis, MO). This cell-permeable, positively charged, red-orange fluorescent dye is readily sequestered by active mitochondria due to the relative negative charge of the fluorophore. However, depolarization of the mitochondrial membrane results in a failure to sequester TMRE. Cells were then harvested, washed, resuspended in PBS, and analyzed by FACS analysis. Data analysis was performed using Flowjo.
2.5. Measurement of Mitochondrial ATP Production
ATP production was measured by a plate reader bioluminescence assay following the manufacturer’s instructions (abcam, Boston, MA). In brief, MH-S cells were harvested after the appropriate exposures, stained with Diff-Quik (Dade Behring; Newark, DE), and counted using a hemocytometer. 10 μL of a cell resuspension ( cells) was mixed with 100 μL of the reaction mix for 5–10 min and then read in a luminometer. The ATP values were normalized to the cell count for each sample.
2.6. Colorimetric Assay for Measuring the Mitochondrial Ratio of NAD/NADH
EnzyChrom NAD+/NADH assay kit (Bioassay Systems; Hayward, CA) was used to determine the mitochondrial ratio of NAD+/NADH. In brief, mitochondria were isolated from MH-S cells or mAMs using a Mitochondria Isolation Kit (Thermo Fisher Scientific, Rockford, IL). NAD+ and NADH were then extracted with the extraction buffer provided in the assay kit, mixed with assay buffer, and absorbance-read at 565 nm.
2.7. Measurement of Phagocytosis and Cell Viability
To determine the phagocytic capacity of macrophages, pHrodo Red S. aureus bioparticles conjugate (Invitrogen, Carlsbad, CA) was added to the culture media according to the manufacturer’s recommendations with ~2 × 106 cells per 2 mg vial of pHrodo-labeled bioparticles. Cells were incubated with the pHrodo labeled bioparticles for 2 hrs and then collected for FACS analysis and data analysis by Flowjo. This phagocytosis assay is based on the fact that there is a minimal fluorescence signal when the pHrodo Red S. aureus bioparticle conjugate is adherent to the outer surface of the phagocyte. Once the S. aureus is internalized and incorporated into the acidic environment of the phagosome, the bioparticle conjugates emit a strong red fluorescence. Internalization was verified by live cell confocal imaging (Olympus FluoView FV1000, Center Valley, PA). To assess changes in viability due to ethanol, MH-S cells were stained with the Dead Cell Apoptosis Kit with Annexin V Alexa Fluor 488 and Propidium Iodide (PI) (Invitrogen, Carlsbad, CA) before analysis by flow cytometry.
2.8. Fluorescence Microscopy and Image Analysis
mAMs isolated from EtOH-fed and control mice were cultured overnight in 8-well cover glass bottom chambers (Lab-Tek; Scotts Valley, CA) with RPMI 1640 medium containing 2% FBS and 1% penicillin/streptomycin. Some mAMs were also treated with 500 μM mitoT for 24 hrs. ROS probes CM-H2DCFDA (10 μM) or mitoSOX (10 μM) was added to the media and images were taken after a 30 min incubation. Images were acquired with Olympus FluoView FV1000 Confocal Microscope using a 63 × oil objective. Images were viewed and analyzed by FV10-ASW 2.0 (Olympus, Center Valley, PA). For mitochondrial morphology analysis, acquired images were subjected to particle analysis using ImageJ Particle Analyzer (National Institutes of Health (http://rsbweb.nih.gov/ij/)). After thresholding, individual particles (mitochondria) were analyzed for area, perimeter, circularity ( Area/(perimeter2)), and the lengths of major and minor axes of fit ellipse. From these values, form factor (FF; the reciprocal of circularity value) and aspect ratio (AR; major/minor) were calculated. Both FF and AR have a minimal value of 1 when a particle is a perfect circle and the values increase as the noncircle features of the particle increase. Specifically, AR is a measure of mitochondrial length and the increase of FF represents the increase of mitochondrial length and branching. This procedure is similar to mitochondrial morphology analysis as previously described [30, 32].
3.1. Chronic EtOH Exposure Induced Mitochondrial ROS Generation
CM-H2DCFDA is oxidized to DCF (dichloro-fluorescein) by cellular ROS  and ethanol increased DCF fluorescence (images in Figure 1(a)). MitoSOX Red, which selectively targets mitochondria and is rapidly oxidized by superoxide, was used to monitor mitochondrial superoxide production (images in Figure 1(a)). In MH-S cells, five days of EtOH exposure increased both cellular ROS and mitochondrial superoxide production by ~100% and 50%, respectively (Figures 1(b) and 1(c)). However, EtOH-induced upregulation of cellular ROS and mitochondrial superoxide in MH-S cells were reversed by treatment with the mitochondrial targeted antioxidant, mitoT. To determine whether chronic EtOH ingestion induced ROS generation in vivo, mAMs were isolated from control or EtOH-fed mice and stained with MitoSOX and CM-H2DCFDA before flow cytometry or confocal imaging. Chronic ethanol ingestion upregulated cellular ROS in mAMs by ~2-fold and mitochondrial superoxide by ~3-fold (Figures 2(a) and 2(b)). Similar to that observed with MH-S cells, 24 h in vitro treatments of the mAMs with mitoT reversed EtOH-induced cellular and mitochondrial ROS production.
3.2. Chronic EtOH Exposure Resulted in Mitochondrial Dysfunction
Mitochondrial membrane potential is a key indicator of mitochondrial function and integrity. It can be determined with TMRE, a cell permeable cationic dye that readily accumulates in active mitochondria because of the relative negative charge of the mitochondrial membrane potential. EtOH exposure resulted in two mitochondrial populations with TMRE staining. The population with higher TMRE intensity represents those with polarized mitochondria and greater capacity to transport the fluorophore. The population with lower TMRE intensity represents the depolarized mitochondria and decreased capacity to transport the fluorophore. In mAMs, chronic EtOH ingestion decreased the population of cells with higher TMRE staining by 10% (Figures 3(a) and 3(b)) suggesting loss of mitochondrial membrane potential. Similarly, EtOH exposure of MH-S cells incrementally decreased the percentage of cells with higher TMRE staining relative to the period of EtOH exposure (Figure 4(a)). In addition, in vitro and in vivo alcohol exposure also decreased the ratio of NAD+/NADH (Figures 4(c) and 3(c)). Since NADH is oxidized to NAD+ in the process of transferring electrons in the mitochondrial electron transfer chain, decreases in the NAD+/NADH ratio indicate a mitochondrial redox imbalance and loss of mitochondrial function. Indeed, this EtOH-induced loss of mitochondrial membrane integrity and perturbations in the NAD+/NADH ratio were accompanied by a ~25% decrease in ATP production (Figure 4(b)). Ethanol-induced decreases in ATP and NAD+/NADH levels were both normalized through the addition of mitoT (Figures 4(b) and 4(c)).
3.3. Chronic EtOH Exposure Induced Mitochondria Condensation and Perinuclear Clustering
Mitochondria are dynamic organelles which constantly change their size and shape by fusion and fission and their morphological dynamics are linked to the regulation of normal cell physiology and disease. We next examined whether the EtOH-induced mitochondrial ROS generation and mitochondrial depolarization were linked to changes in mitochondrial morphology. In control MH-S, the mitochondrial network was spread throughout the cell (Figure 5(a)). With 5 consecutive days of EtOH exposure (0.2%), the majority of MH-S cells had mitochondria that were condensed and located in the perinuclear region (Figure 5(a)). To quantitatively address mitochondrial morphology changes, we analyzed mitochondrial morphology using a computer-assisted morphometric analysis, which calculates form factor (FF) and aspect ratio (AR) as discussed above. With a minimal value of 1 representing a perfect circle (major axis = minor axis), the mitochondria within control MH-S cells had AR values distributed above 4 suggesting that the mitochondria were elongated. With EtOH exposure, the majority of the AR values were below 4, suggesting a transition from an elongated shape to a more spherical shape. In addition, mitochondrial areas exceeded 15 μm2 in the EtOH treatment group suggesting mitochondrial clustering. We next investigated the mitochondrial morphology after chronic EtOH ingestion. Figure 5(b) is comprised of representative confocal microscopic images of AMs isolated from control and EtOH-fed mice. Because primary mAM cells were taken from their original environment in mouse lungs, they are more fragile, and their morphology was not well retained like that for the MH-S cell line. However, the mitochondria in the AMs from the EtOH-fed mice were more fragmented and clustered at the perinuclear area when compared to the AMs from the control mice.
3.4. EtOH-Induced Impairment of Macrophage Phagocytosis Was Reversed by mitoT
As demonstrated previously [9, 10], EtOH exposure decreased the phagocytic capacity of mAMs and MH-S cells. Since mitoT attenuated cellular and mitochondrial ROS, we next examined whether mitoT would reverse the effects of ethanol on phagocytosis. As demonstrated in our previous studies, EtOH exposure suppressed phagocytosis of the S. aureus bioparticle conjugates by 30% (Figure 6). Treatment with mitoT during the last 24 h of EtOH exposure restored the phagocytic ability of MH-S cells suggesting that mitochondrial-derived oxidative stress was central to EtOH-induced disruptions in phagocytosis. We also examined whether mitoT could restore phagocytosis to mAMs from ethanol-fed mice. Similar to that observed with MH-S cells, in vitro treatments with mitoT restored phagocytosis to the mAMs suppressed by chronic ethanol ingestion (Figure 7). These results further confirmed the association between EtOH-induced mitochondrial oxidative stress and impaired mAM phagocytosis.
3.5. EtOH Induced Early Apoptosis but Was Prevented by mitoT Treatment
As demonstrated previously [9, 10], chronic ethanol ingestion increases AM apoptosis 3-fold with ~30% of the cells expressing markers of apoptosis. In MH-S cells with 5 days of ethanol exposure, the percentage of cells with an early marker of apoptosis, Annexin V positive staining, increased 5-fold (Figures 8(a)–8(c)) when compared to the control group. For cells positive for Annexin V plus loss of cytoplasm, ethanol increased the percentage of cells positive for late apoptosis but statistical significance was not achieved. There also was no statistically significant increase in the percentage of cells positive for a marker of necrosis, propidium iodide staining of DNA (Figure 8(c)). When MH-S cells were pretreated with mitoT, ethanol-induced early apoptosis, Annexin V positive staining, was attenuated (Figure 8(d)).
Chronic alcohol abuse is associated with an increased risk of respiratory infections, pneumonia, and tuberculosis, even in those without a clinical diagnosis of an alcohol use disorder [1, 3]. However, the underlying mechanisms by which alcohol abuse increases the risk of respiratory infections are unclear. One central effect of chronic alcohol ingestion is severe oxidative stress and depletion of critical antioxidants . Within the alveolar space, GSH levels in the alcoholic subjects were significantly decreased when compared with those of nonalcoholic subjects. In the alveolar epithelial lining fluid, alcohol abuse caused an ~40 mV change in the glutathione and glutathione disulfide (GSH/GSSG) redox potential (Eh) [6, 7, 34]. Changes in the extracellular GSH/GSSG redox status were echoed in the intracellular GSH/GSSG redox balance of alveolar type II cells. Indeed, chronic alcohol ingestion caused a 60% decrease in GSH and induced GSH/GSSG oxidation by 40 mV in alveolar type II cells from ethanol-fed adult male rats . In the mitochondria of type II cells, chronic alcohol ingestion also induced a 60 mV oxidation of the GSH/GSSG redox potential when compared to the cells from control rats . For AMs, the GSH/GSSG redox state was oxidized by ~30 mV after chronic ethanol ingestion . Across intracellular and extracellular GSH pools in alveolar cells, the GSH/GSSG redox state was consistently oxidized by 30–60 mV. AMs are the only intra-alveolar phagocyte that responds to inflammation  and their function is dependent on the oxidation/reduction balance in the alveolar lining fluid. Indeed, ex vivo GSH antioxidant supplementation can reverse the EtOH-induced suppression of phagocytosis in rodent models of chronic alcohol abuse .
Previous studies in our laboratory demonstrated that EtOH promotes oxidative stress in AMs through increased ROS production by NADPH oxidases (Nox) . In that mouse model, chronic EtOH ingestion increased the level of mRNA and protein expression of Nox1, Nox2, and Nox4. Since mitochondria produce 90% of cellular ROS compared to a 10% cytosolic ROS contribution under baseline conditions , we examined the potential contribution of mitochondria to the ROS. Like the alveolar type II cells , the current study demonstrated that EtOH exposure (in vitro or in vivo) also upregulated mitochondrial ROS generation. Under baseline conditions, increased mitochondrial superoxide production can be enzymatically dismutated to hydrogen peroxide which is subsequently removed by catalase or glutathione peroxidase. However, chronic increases in mitochondrial ROS generation can overwhelm normal scavenging mechanisms, promote the uncoupling of the respiratory chain, and result in more ROS generation, loss of mitochondrial membrane potential, and decreased ATP. In the current study, a role for ethanol-induced mitochondrial ROS was further demonstrated by treatment of AM cells with mitoTEMPOL (mitoT) which reversed EtOH-induced mitochondrial ROS. MitoT contains a lipophilic triphenyl-phosphonium cation added to the TEMPOL antioxidant moiety which promotes its accumulation in the mitochondria. The TEMPOL moiety is a piperidine nitroxide, which has been widely used as a mitochondrial specific antioxidant for in vivo and in vitro studies. The TEMPOL moiety has superoxide dismutase activity that promotes the detoxification of ferrous iron and prevents toxic hydroxyl radicals formation in the reaction of H2O2 with ferrous iron [39, 40]. Although our analysis of ROS by redox sensitive fluorophores has its limitations, treatment with mitoT blocked or reversed EtOH-induced ROS generation in the mitochondria further supporting that EtOH promoted mitochondrial ROS. Whether EtOH-induced mitochondrial ROS is due to interference with mitochondrial redox balances  or other mechanisms remains to be determined. Furthermore, mitoT also blocked the ethanol-induced increases in cytosolic ROS suggesting that mitochondrial ROS contributes to the generation of cytosolic ROS. Additional studies are needed to determine whether ROS generation through NADPH oxidases, CYP2E1, or other ROS generators are dependent on mitochondrial ROS.
EtOH-induced mitochondrial ROS was also associated with mitochondrial dysfunction. Mitochondrial integrity and function were interrupted by EtOH as evidenced by decreased mitochondrial membrane potential in MH-S cells. More importantly, mAMs from EtOH-fed mice displayed decreased mitochondrial function as evidenced by decreased mitochondrial membrane potential and ATP. The ratio of NAD+/NADH was also decreased in both the in vitro and in vivo models of ethanol exposure. Although the isolation protocol may have increased and NADH leak from the mitochondria, it is unlikely that the leak of one component would be preferential over the other. Since the actual leak may differ between mitochondrial preparations, we decided that expressing the concentrations of the two components as a ratio would be more accurate. In these studies, was a.u. (absolute unit) and NADH was a.u. for the control group. For the ethanol group, was a.u. and NADH was a.u. Therefore, the decrease in and increase in NADH resulted in a decrease in the mitochondrial NAD+/NADH ratio suggesting its oxidation in the mitochondria.
Mitochondria are organelles which supply energy for normal cellular functions making it a key regulator of cell function. For the AM, the energy intensive cellular process such as phagocytosis is particularly dependent on the capacity of the mitochondria to generate ATP. In the current studies, EtOH exposure promoted significant mitochondrial morphological changes, a central indicator of the organelle’s integrity and function. In the control group, there was a network of mitochondria with an elongated shape. With EtOH exposure, the mitochondria became more spherical in shape and were present in condensed perinuclear clusters. These EtOH-induced mitochondrial morphological changes are generally associated with cellular oxidative stress and are associated with cell death . As with ethanol-induced mitochondrial ROS, treatment with mitoT normalized mitochondrial NAD+/NADH as well as the ATP pool after ethanol exposure. This further supports a causative role for ethanol-induced mitochondrial ROS in the corresponding mitochondrial dysfunction. As observed in previous studies with ethanol-fed animals , there was impaired phagocytosis, a key immune function of AMs. However, in vitro mitoT treatments reversed the injurious effects of EtOH on the mitochondria and restored the phagocytic capacity even in mAMs from mice fed ethanol for 12 weeks. In addition to impaired bacterial clearance, ethanol decreased cell viability as evidenced by increased early markers of apoptosis. The capacity of mitoT treatment to block apoptosis suggested a central role for ethanol-induced mitochondrial dysfunction in the apoptotic process.
Since mitoT decreased ethanol-induced mitochondrial- and cytosolic-derived ROS, one potential mechanism for the beneficial effects of mitoT could be through its positive cytosolic effects. In previous studies, we demonstrated that chronic EtOH ingestion increases the production of TGF-β and IL-13 in AMs which subsequently promotes alternative activation (M2 activation) . In that study, TGF-β and IL-13 activated an autocrine loop that was central to AM alternative activation. EtOH-induced upregulation of TGF-β expression also promoted another self-activating autocrine loop with the constitutively active NOX 4 resulting in chronic cytosolic ROS generation . Additional studies are needed to determine whether the ethanol-induced activation of TGF-β/IL-13 and the TGFβ/NOX 4 autocrine loops are causative or secondary to ethanol-induced mitochondrial dysfunction. Alternatively, the primary driver could be through ethanol-induced mitochondrial ROS that overwhelm the large antioxidant capacity of the mitochondria, leak into the cytosol, and activate various mechanisms for cytosolic ROS such as the TGF-β/NOX 4 autocrine loop. In the current study, mitoT attenuated ethanol-induced mitochondrial dysfunction such as ROS, decreased ATP levels, and decreased NAD+/NADH. The mechanisms by which mitoT maintains these events that are critical for the highly energy-dependent processes of phagocytosis and maintenance of cell viability are unclear but may be through maintenance of mitochondrial GSH/GSSG, a critical event for type II cells .
Although each piece of data is not singularly definitive, the collective data from diverse measures indicated that ethanol increased mitochondrial ROS: (1) increased cellular ROS as indicated by CM-H2DCFDA oxidation which was blocked by the mitochondrial specific antioxidant mitoT; (2) increased mitochondrial ROS as indicated by MitoSOX fluorescence which was also blocked by mitoT; (3) increased cellular and mitochondrial ROS with in vitro and in vivo EtOH exposure; (4) the ability of mitoT to attenuate cellular and mitochondrial ROS in the AMs even after chronic EtOH ingestion; and (5) increased oxidation of the mitochondrial NAD+/NADH ratio. This mitochondrial oxidation was also associated with mitochondrial dysfunction as evidenced by loss of mitochondrial morphology, depolarization of the mitochondrial membrane potential, and decreased ATP generation. The mitochondria-targeted antioxidant mitoT not only reversed EtOH-induced mitochondrial and cytosolic ROS generation, it also reversed EtOH-induced mitochondrial dysfunction, restored AM phagocytosis, and maintained cell viability. Phagocytosis and cell viability are both complex cellular processes that are controlled at multiple points; additional studies are needed to determine the actual roles of mitochondrial ROS in EtOH-induced disruption of these cellular events. Chronically, alcohol can dampen the inflammatory responses of alveolar macrophages and the chronic suppression of phagocytosis decreases the capacity of alveolar macrophages to clear microbes. Therefore, EtOH-induced mitochondrial ROS and dysfunction in AMs may be pivotal in the increased risk of respiratory infections and ARDS in subjects with an alcohol use disorder.
Conflict of Interests
The authors declare that there is no conflict of interests regarding the publication of this paper.
This work was supported by a NIAAA T32 Training Grant (5T32AA013528-08), the Emory Alcohol and Lung Biology Center (1P50AA135757), and NIAAA (R01 AA12197). The authors acknowledge the facilities and the scientific and technical assistance of the Flow Cytometry Core Facility and Imaging Core Facility at Emory + Children’s Pediatrics Research Center.
- M. Bhatty, S. B. Pruett, E. Swiatlo, and B. Nanduri, “Alcohol abuse and Streptococcus pneumoniae infections: consideration of virulence factors and impaired immune responses,” Alcohol, vol. 45, no. 6, pp. 523–539, 2011.
- C. W. Chen, G. M. Jong, J. J. Shiau et al., “Adult bacteremic pneumonia: bacteriology and prognostic factors,” Journal of the Formosan Medical Association, vol. 91, no. 8, pp. 754–759, 1992.
- A. J. Mehta and D. M. Guidot, “Alcohol abuse, the alveolar macrophage and pneumonia,” American Journal of the Medical Sciences, vol. 343, no. 3, pp. 244–247, 2012.
- M. J. Fine, M. A. Smith, C. A. Carson et al., “Prognosis and outcomes of patients with community-acquired pneumonia: a meta-analysis,” Journal of the American Medical Association, vol. 275, no. 2, pp. 134–141, 1996.
- G. V. Bochicchio, M. Joshi, K. Bochicchio, K. Tracy, and T. M. Scalea, “A time-dependent analysis of intensive care unit pneumonia in trauma patients,” Journal of Trauma, vol. 56, no. 2, pp. 296–303, 2004.
- M. Y. Yeh, E. L. Burnham, M. Moss, and L. A. S. Brown, “Chronic alcoholism alters systemic and pulmonary glutathione redox status,” American Journal of Respiratory and Critical Care Medicine, vol. 176, no. 3, pp. 270–276, 2007.
- M. Moss, D. M. Guidot, M. Wong-Lambertina, T. ten Hoor, R. L. Perez, and L. A. S. Brown, “The effects of chronic alcohol abuse on pulmonary glutathione homeostasis,” American Journal of Respiratory and Critical Care Medicine, vol. 161, no. 2 I, pp. 414–419, 2000.
- A. Aderem and D. M. Underhill, “Mechanisms of phagocytosis in macrophages,” Annual Review of Immunology, vol. 17, pp. 593–623, 1999.
- S. D. Brown, T. W. Gauthier, and L. A. S. Brown, “Impaired terminal differentiation of pulmonary macrophages in a guinea pig model of chronic ethanol ingestion,” Alcoholism, vol. 33, no. 10, pp. 1782–1793, 2009.
- S. M. Yeligar, F. L. Harris, C. M. Hart, and L. A. S. Brown, “Ethanol induces oxidative stress in alveolar macrophages via upregulation of NADPH oxidases,” Journal of Immunology, vol. 188, no. 8, pp. 3648–3657, 2012.
- I. Al Ghouleh, N. K. H. Khoo, U. G. Knaus et al., “Oxidases and peroxidases in cardiovascular and lung disease: new concepts in reactive oxygen species signaling,” Free Radical Biology and Medicine, vol. 51, no. 7, pp. 1271–1288, 2011.
- L. L. Ji, “Antioxidants and oxidative stress in exercise,” Proceedings of the Society for Experimental Biology and Medicine, vol. 222, pp. 283–292, 1999.
- A. Y. Andreyev, Y. E. Kushnareva, and A. A. Starkov, “Mitochondrial metabolism of reactive oxygen species,” Biochemistry, vol. 70, no. 2, pp. 200–214, 2005.
- D. C. Wallace, W. Fan, and V. Procaccio, “Mitochondrial energetics and therapeutics,” Annual Review of Pathology, vol. 5, pp. 297–348, 2010.
- P. S. Brookes, Y. Yoon, J. L. Robotham, M. W. Anders, and S. Sheu, “Calcium, ATP, and ROS: a mitochondrial love-hate triangle,” American Journal of Physiology—Cell Physiology, vol. 287, no. 4, pp. C817–C833, 2004.
- A. Patenaude, M. R. ven Murthy, and M. Mirault, “Mitochondrial thioredoxin system: effects of TrxR2 overexpression on redox balance, cell growth, and apoptosis,” The Journal of Biological Chemistry, vol. 279, no. 26, pp. 27302–27314, 2004.
- D. E. Handy and J. Loscalzo, “Redox regulation of mitochondrial function,” Antioxidants and Redox Signaling, vol. 16, no. 11, pp. 1323–1367, 2012.
- V. Sosa, T. Moliné, R. Somoza, R. Paciucci, H. Kondoh, and M. E. LLeonart, “Oxidative stress and cancer: an overview,” Ageing Research Reviews, vol. 12, pp. 376–390, 2013.
- W. Ying, “NAD(+)/NADH and NADP(+)/NADPH in cellular functions and cell death: regulation and biological consequences,” Antioxidants and Redox Signaling, vol. 10, no. 2, pp. 179–206, 2008.
- D. Esterházy, M. S. King, G. Yakovlev, and J. Hirst, “Production of reactive oxygen species by complex I (NADH:ubiquinone oxidoreductase) from Escherichia coli and comparison to the enzyme from mitochondria,” Biochemistry, vol. 47, no. 12, pp. 3964–3971, 2008.
- L. Kussmaul and J. Hirst, “The mechanism of superoxide production by NADH:ubiquinone oxidoreductase (complex I) from bovine heart mitochondria,” Proceedings of the National Academy of Sciences of the United States of America, vol. 103, no. 20, pp. 7607–7612, 2006.
- N. Braidy, G. J. Guillemin, H. Mansour, T. Chan-Ling, A. Poljak, and R. Grant, “Age related changes in NAD+ metabolism oxidative stress and sirt1 activity in wistar rats,” PLoS ONE, vol. 6, no. 4, Article ID e19194, 2011.
- H. J. Edenberg, “The genetics of alcohol metabolism: role of alcohol dehydrogenase and aldehyde dehydrogenase variants,” Alcohol Research and Health, vol. 30, no. 1, pp. 5–13, 2007.
- R. T. Cook, A. J. Schlueter, R. A. Coleman et al., “Thymocytes, pre-B Cells, and organ changes in a mouse model of chronic ethanol ingestion: absence of subset-specific glucocorticoid-induced immune cell loss,” Alcoholism, vol. 31, no. 10, pp. 1746–1758, 2007.
- R. T. Cook, X. Zhu, R. A. Coleman et al., “T-cell activation after chronic ethanol ingestion in mice,” Alcohol, vol. 33, no. 3, pp. 175–181, 2004.
- M. C. Wagner, S. M. Yeligar, L. A. Brown, and C. Michael Hart, “PPARγ ligands regulate NADPH oxidase, eNOS, and barrier function in the lung following chronic alcohol ingestion,” Alcoholism, vol. 36, no. 2, pp. 197–206, 2012.
- T. R. Jerrells, J. A. Pavlik, J. DeVasure et al., “Association of chronic alcohol consumption and increased susceptibility to and pathogenic effects of pulmonary infection with respiratory syncytial virus in mice,” Alcohol, vol. 41, no. 5, pp. 357–369, 2007.
- K. Song, R. A. Coleman, X. Zhu et al., “Chronic ethanol consumption by mice results in activated splenic T cells,” Journal of Leukocyte Biology, vol. 72, no. 6, pp. 1109–1116, 2002.
- N. Khalil, R. N. O'Connor, K. C. Flanders, S. W. Shing Wade, and C. I. Whitman, “Regulation of type II alveolar epithelial cell proliferation by TGF-beta during bleomycin-induced lung injury in rats,” American Journal of Physiology, vol. 267, no. 5, pp. L498–L507, 1994.
- T. Yu, J. L. Robotham, and Y. Yoon, “Increased production of reactive oxygen species in hyperglycemic conditions requires dynamic change of mitochondrial morphology,” Proceedings of the National Academy of Sciences of the United States of America, vol. 103, no. 8, pp. 2653–2658, 2006.
- K. Nakahira, J. A. Haspel, V. A. K. Rathinam et al., “Autophagy proteins regulate innate immune responses by inhibiting the release of mitochondrial DNA mediated by the NALP3 inflammasome,” Nature Immunology, vol. 12, no. 3, pp. 222–230, 2011.
- J. Irobi, A. Holmgren, V. D. Winter, et al., “Mutant HSPB8 causes protein aggregates and a reduced mitochondrial membrane potential in dermal fibroblasts from distal hereditary motor neuropathy patients,” Neuromuscular Disorders, vol. 22, pp. 699–711, 2012.
- Y. Chen, M. B. Azad, and S. B. Gibson, “Superoxide is the major reactive oxygen species regulating autophagy,” Cell Death and Differentiation, vol. 16, no. 7, pp. 1040–1052, 2009.
- M. Y. Yeh, E. L. Burnham, M. Moss, and L. A. S. Brown, “Non-invasive evaluation of pulmonary glutathione in the exhaled breath condensate of otherwise healthy alcoholics,” Respiratory Medicine, vol. 102, no. 2, pp. 248–255, 2008.
- L. A. S. Brown, F. L. Harris, R. Bechara, and D. M. Guidot, “Effect of chronic ethanol ingestion on alveolar type II cell: gutathione and inflammatory mediator-induced apoptosis,” Alcoholism, vol. 25, no. 7, pp. 1078–1085, 2001.
- L. A. S. Brown, X. Ping, F. L. Harris, and T. W. Gauthier, “Glutathione availability modulates alveolar macrophage function in the chronic ethanol-fed rat,” American Journal of Physiology—Lung Cellular and Molecular Physiology, vol. 292, no. 4, pp. L824–L832, 2007.
- K. Tsushima, L. S. King, N. R. Aggarwal, A. De Gorordo, F. R. D'Alessio, and K. Kubo, “Acute lung injury review,” Internal Medicine, vol. 48, no. 9, pp. 621–630, 2009.
- R. S. Balaban, S. Nemoto, and T. Finkel, “Mitochondria, oxidants, and aging,” Cell, vol. 120, no. 4, pp. 483–495, 2005.
- J. Trnka, F. H. Blaikie, A. Logan, R. A. J. Smith, and M. P. Murphy, “Antioxidant properties of MitoTEMPOL and its hydroxylamine,” Free Radical Research, vol. 43, no. 1, pp. 4–12, 2009.
- M. P. Murphy and R. A. J. Smith, “Targeting antioxidants to mitochondria by conjugation to lipophilic cations,” Annual Review of Pharmacology and Toxicology, vol. 47, pp. 629–656, 2007.
- S. Manzo-Avalos and A. Saavedra-Molina, “Cellular and mitochondrial effects of alcohol consumption,” International Journal of Environmental Research and Public Health, vol. 7, no. 12, pp. 4281–4304, 2010.