BioMed
Research International. (Annual 2014):
Copyright: COPYRIGHT 2014 Hindawi Publishing Corp.
Abstract:
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.
1. Introduction
Both acute and chronic alcohol consumption
have well-documented effects on the immune system leading to increased
susceptibility to community acquired pneumonia and tuberculosis [1]. When
subjects with an alcohol use disorder get pneumonia, they are more likely to be
infected with serious Gram-negative bacteria [2] and these increased risks
occur even in those who do not meet the diagnostic criteria for an alcohol use
disorder [3]. This results in a higher rate of intensive care use, longer
inpatient stays, higher healthcare costs, and a 2-4 times greater mortality
rate [4]. There is also an increased risk of ventilator-associated pneumonia
which worsens the morbidity and mortality rates [5]. 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 3].
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 [3].
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 [8]. 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 [11]. 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 [12]. 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 ([O.sub.2.sup.*-]), hydrogen peroxide
([H.sub.2][O.sub.2]), and hydroxyl radicals ([sup.*]OH). 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 [17].
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 [18]. During respiration, the NADH is oxidized
to [NAD.sup.+] and the [NAD.sup.+]/NADH ratio has been recognized as a key
regulator in energy metabolism, aging, and immunological functions [19]. For
example, decreases in [NAD.sup.+] or in the [NAD.sup.+]/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) [23] which uses
[NAD.sup.+] as a cofactor. Thus, acetaldehyde metabolism decreases
mitochondrial [NAD.sup.+] pools and increases NADH. The resulting decreases in
the [NAD.sup.+]/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 (n = 5/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% [+ or -] 0.03, as published by our group [26] 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 xg 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 [degrees]C in 5%
C[O.sub.2] 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[degrees]C in a 5% C[O.sub.2] 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 24hr of
the 5d EtOH treatment, some cells were also treated with the
mitochondria-targeted antioxidant mitoTEMPOL (mitoT, 100 [micro]M) [31].
2.3. Measurement of Intracellular ROS Generation.
After EtOH exposure, the cellular ROS
sensitive probe CM-[H.sub.2]DCFDA (Invitrogen; Carlsbad, CA) and the
mitochondrial superoxide probe mitoSOX (Invitrogen; Carlsbad, CA) were added to
the medium (10 [micro]M, 30 min, 37 [degrees]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-[H.sub.2]DCFDA
and MitoSOX were excited at 488 nm and detected at 530 [+ or -] 15 nm or 585 [+
or -] 42 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 degrees]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 [micro]L of a cell resuspension ([10.sup.3] - [10.sup.4]
cells) was mixed with 100 [micro]L 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.sup.+]/NADH assay kit (Bioassay
Systems; Hayward, CA) was used to determine the mitochondrial ratio of
[NAD.sup.+]/NADH. In brief, mitochondria were isolated from MH-S cells or mAMs
using a Mitochondria Isolation Kit (Thermo Fisher Scientific, Rockford, IL).
[NAD.sup.+] 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 x [10.sup.6] 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 [micro]M mitoT
for 24hrs. ROS probes CM-[H.sub.2]DCFDA (10 [micro]M) or mitoSOX (10 [micro]M)
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 x 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 (4[pi] x Area/([perimeter.sup.2])), 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. Results
3.1. Chronic EtOH Exposure Induced Mitochondrial ROS Generation.
CM-[H.sub.2]DCFDA is oxidized to DCF
(dichlorofluorescein) by cellular ROS [33] 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-[H.sub.2]DCFDA 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.
[FIGURE 1 OMITTED]
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.sup.+]/NADH (Figures 4(c) and
3(c)). Since NADH is oxidized to [NAD.sup.+]+ in the process of transferring
electrons in the mitochondrial electron transfer chain, decreases in the
[NAD.sup.+]/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.sup.+]/NADH ratio were
accompanied by a ~25% decrease in ATP production (Figure 4(b)). Ethanol-induced
decreases in ATP and [NAD.sup.+]/NADH levels were both normalized through the
addition of mitoT (Figures 4(b) and 4(c)).
[FIGURE 2 OMITTED]
[FIGURE 3 OMITTED]
[FIGURE 4 OMITTED]
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 [micro][m.sup.2] 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.
[FIGURE 5 OMITTED]
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)).
[FIGURE 6 OMITTED]
4. Discussion
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
[7]. 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 [35]. 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 [35]. For AMs, the GSH/GSSG redox state was oxidized by -30 mV
after chronic ethanol ingestion [36]. Across intracellular and extracellular
GSH pools in alveolar cells, the GSH/GSSG redox state was consistently oxidized
by 30-60mV. AMs are the only intra-alveolar phagocyte that responds to
inflammation [37] 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 [36].
Previous studies in our laboratory
demonstrated that EtOH promotes oxidative stress in AMs through increased ROS
production by NADPH oxidases (Nox) [10]. In that mouse model, chronic EtOH
ingestion increased the level of mRNA and protein expression of Nox1, Nox2, and
Nox4. Since mitochondria prodduce 90% of cellular ROS compared to a 10%
cytosolic ROS contribution under baseline conditions [38], we examined the
potential contribution of mitochondria to the ROS. Like the alveolar type II
cells [35], 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 triphenylphosphonium 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 super-oxide dismutase activity that promotes the detoxification of
ferrous iron and prevents toxic hydroxyl radicals formation in the reaction of
[H.sub.2][O.sub.2] 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 [41] 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.
[FIGURE 7 OMITTED]
[FIGURE 8 OMITTED]
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.sup.+]/NADH was also decreased in
both the in vitro and in vivo models of ethanol exposure. Although the
isolation protocol may have increased [NAD.sup.+] 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, NAD+
was 0.33 [+ or -] 0.03 a.u. (absolute unit) and NADH was 0.20 [+ or -] 0.01
a.u. for the control group. For the ethanol group, NAD+ was 0.25 [+ or -] 0.03
a.u. and NADH was 0.76 [+ or -] 0.20 a.u. Therefore, the decrease in
[NAD.sup.+] and increase in NADH resulted in a decrease in the mitochondrial
[NAD.sup.+]/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 [35]. As with
ethanol-induced mitochondrial ROS, treatment with mitoT normalized
mitochondrial [NAD.sup.+]/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 [36], 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-[beta] and IL-13 in AMs which subsequently promotes
alternative activation (M2 activation) [7]. In that study, TGF-[beta] and IL-13
activated an autocrine loop that was central to AM alternative activation.
EtOH-induced upregulation of TGF-[beta] expression also promoted another
self-activating autocrine loop with the constitutively active NOX 4 resulting
in chronic cytosolic ROS generation [10]. Additional studies are needed to
determine whether the ethanol-induced activation of TGF-[beta]/IL-13 and the
TGF[beta]/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-[beta]/NOX4 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 [35].
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-[H.sub.2]DCFDA 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.sup.+]/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.
http://dx.doi.org/10.1155/2014/371593
Conflict of Interests
The authors declare that there is no conflict
of interests regarding the publication of this paper.
Acknowledgments
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.
References
[1]
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.
[2]
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.
[3]
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.
[4]
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.
[5]
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.

[6]
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.
[7]
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.
[8]
A. Aderem and D. M. Underhill, "Mechanisms of phagocytosis in
macrophages," Annual Review of Immunology, vol. 17, pp. 593-623, 1999.
[9]
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.
[10]
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.
[11]
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.
[12]
L. L. Ji, "Antioxidants and oxidative stress in exercise,"
Proceedings of the Society for Experimental Biology and Medicine, vol. 222, pp.
283-292, 1999.

[13]
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.
[14]
D. C. Wallace, W. Fan, and V. Procaccio, "Mitochondrial energetics and
therapeutics," Annual Review of Pathology, vol. 5, pp. 297-348, 2010.
[15]
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.
[16]
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.
[17]
D. E. Handy and J. Loscalzo, "Redox regulation of mitochondrial
function," Antioxidants and Redox Signaling, vol. 16, no. 11, pp.
1323-1367, 2012.
[18]
V. Sosa, T. Moline, 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.
[19]
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.

[20]
D. Esterhazy, 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.
[21]
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.
[22]
N. Braidy, G. J. Guillemin, H. Mansour, T. Chan-Ling, A. Poljak, and R. Grant,
"Age related changes in [NAD.sub.+] metabolism oxidative stress and sirt1
activity in wistar rats," PLoS ONE, vol. 6, no. 4, Article ID e19194,
2011.
[23]
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
[24]
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.
[25]
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.
[26]
M. C. Wagner, S. M. Yeligar, L. A. Brown, and C. Michael Hart,
"PPAR[gamma] ligands regulate NADPH oxidase, eNOS, and barrier function in
the lung following chronic alcohol ingestion," Alcoholism, vol. 36, no. 2,
pp. 197-206, 2012.

[27]
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.
[28]
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.
[29]
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.
[30]
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.
[31]
K. Nakah










