Mitochondrial depolarization following hydrogen sulfide exposure in erythrocytes from a sulfide-tolerant marine invertebrate
1 Department of Zoology, University of Florida, Gainesville, FL 32611-8525,
USA
2 Mount Desert Island Biological Laboratory, Bar Harbor, ME 24533,
USA
* Author for correspondence (e-mail: djulian{at}ufl.edu)
Accepted 25 August 2005
![]() |
Summary |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Key words: hydrogen sulfide, Glycera dibranchiata, mitochondria, coelomocyte, free radical
![]() |
Introduction |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
In normally functioning mitochondria, oxidative phosphorylation maintains
an electrochemical proton (H+) gradient, and therefore a
mitochondrial electrical potential (m), which drives the ATP
synthase (F1FO-ATP synthase). In mammalian cells,
reduced
m can result from increased cellular demand for ATP,
mitochondrial outer membrane permeabilization (MOMP) and uncouplers of
oxidative phosphorylation (Bernardi et
al., 2001
; Green and Kroemer,
2004
; Ly et al.,
2003
). Although it might be assumed that inhibition of oxidative
phosphorylation would directly decrease
m, this is not necessarily
true. For example, azide and cyanide in vitro decrease
m
in some mammalian cells (Feeney et al.,
2003
; Jensen et al.,
2002
; Prabhakaran et al.,
2002
), but other mammalian cells show only a small reduction in
m during cyanide exposure unless inhibitors of glycolysis are
added simultaneously (Lawrence et al.,
2001
). With regard to sulfide exposure, one study has reported
that isolated rat hepatocytes exposed to 0.5 mmol l-1 sulfide
in vitro lost over 50% of
m within 1 h, and that this
effect was reduced when the cells were supplemented with glycolytic substrate
(Eghbal et al., 2004
). In
contrast, another study on the same cell type reported that supplementation
with glycolytic substrate had no effect on cell survival during sulfide
exposure, although it did increase cell survival during cyanide exposure
(Thompson et al., 2003
).
Therefore, whether sulfide exposure causes cellular toxicity via
decreased
m is unclear. Furthermore, short-term sulfide exposure
in mice under controlled conditions greatly decreased metabolic rate but had
no apparent long-term effect on the animal's health, indicating that
inhibition of oxidative phosphorylation by sulfide does not necessarily lead
to cell death (Blackstone et al.,
2005
). Whether
m was affected in the mice was not
determined.
One possible mechanism of sulfide-induced mitochondrial depolarization is
MOMP, which in mammalian cells can result from exposure to heavy metals,
reactive oxygen species (ROS) and a variety of other toxins
(Green and Kroemer, 2004). Two
separate mechanisms appear to induce MOMP: (1) opening of the mitochondrial
permeability transition (PT) pore, which permeabilizes the mitochondrial inner
membrane, leading to dissipation of
m, influx of water into the
matrix and eventual rupturing of the outer membrane, and (2) formation of a
pore that permeabilizes the mitochondrial outer membrane
(Green and Kroemer, 2004
). In
either case, MOMP leads inexorably to cell death through the release of
pro-apoptotic factors from the mitochondrial inter-membrane space into the
cytosol (He and Lemasters,
2002
; Hunter et al.,
1976
; Kim et al.,
2003a
; Ly et al.,
2003
). In rat hepatocytes in vitro, the PT pore
inhibitors cyclosporine A (CsA) and trifluoperazine (TFP) decreased
sulfide-induced cell death by 50% compared to sulfide alone, but had no effect
on cyanide-induced cell death (Thompson et
al., 2003
), suggesting that sulfide and cyanide cause cell death
via different mechanisms. Therefore, in contrast to cyanide, which in
mammalian cells does not necessarily dissipate
m
(Lawrence et al., 2001
) and
may not induce PT pore opening (Thompson
et al., 2003
), sulfide exposure of mammalian cells in
vitro causes loss of
m that appears to be at least partially
via PT pore opening (Eghbal et
al., 2004
; Thompson et al.,
2003
).
It is not known whether sulfide exposure affects m in cells of
sulfide-adapted invertebrates, and if so, whether the action is similar to
that of cyanide and therefore likely occurs primarily via COX
inhibition. This is particularly relevant, since recent evidence suggests that
the mitochondria of at least some invertebrates do not respond as expected to
classical PT pore inducers. Menze et al.
(2005
) have shown that
mitochondria from the brine shrimp Artemia franciscana, unlike
mitochondria from mammalian cells, do not swell or release the mitochondrial
pro-apoptotic factor cytochrome c when exposed to high
Ca2+, although the expected molecular components of the PT pore are
present (as indicated by western blot). Furthermore, Sokolova et al.
(2004
) found that in
vitro Cd2+ exposure of hemocytes from the oyster
Crassostrea virginica caused an apparent increase in
m,
whereas Cd2+ exposure of isolated oyster mitochondria caused
inhibition of respiration, a slight decrease in
m but no
significant mitochondrial swelling (in contrast, PT pore opening and
m loss are characteristic of Cd2+ toxicity in mammalian
cells in vitro).
In the present study, we investigated whether sulfide exposure in
vitro causes loss of m in erythrocytes from the bloodworm
Glycera dibranchiata Ehlers 1868 (Annelida: Polychaeta: Phyllodocida:
Glyceridae). To determine whether the action of sulfide was likely
via COX inhibition, we also tested the effect of cyanide and several
other mitochondrial electron transport chain inhibitors on
m. To
test whether the toxicity of sulfide was associated with free radical
production, we assessed intracellular oxidative stress and mitochondrial
superoxide generation during sulfide exposure. G. dibranchiata is
abundant in fine mud with high organic content in the North Atlantic region of
the USA (Wilson and Ruff,
1988
), and is therefore likely to be exposed to sulfide in its
natural habitat. It lacks a vascular system and its hemoglobin-containing
erythrocytes, which are nucleated and contain functional mitochondria, are
circulated by cilia into the thin-walled parapodia for gas exchange
(Mangum, 1994
), suggesting
that these cells are at risk of sulfide exposure.
![]() |
Materials and methods |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Fluorescent dyes
Live-cell fluorescent dyes were used to measure m, ROS and
superoxide in G. dibranchiata erythrocytes in vitro.
Erythrocytes (diluted up to 1:100 in incubation buffer) were loaded with dye
prior to toxin exposure, except as noted below in the sulfide recovery
experiment. The dye was diluted into incubation buffer from a stock solution
(stored at -20°C and prepared as described below), and the erythrocytes
were incubated with the dye in the dark at 15°C for 20-30 min. For each
dye, loading conditions were established in preliminary experiments by
confirming the appropriate labeling of cells or subcellular structures (i.e.
mitochondria) using fluorescence microscopy. All dyes were from Molecular
Probes, Inc. (now Invitrogen Corporation, Carlsbad, CA, USA).
JC-1 and TMRM
The cell-permeant, cationic, lipophilic fluorophores tetramethylrhodamine
methyl ester (TMRM) and
5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolylcarbocyanine
iodide (JC-1) are selective for polarized mitochondria membranes under
appropriate loading conditions, where they fluoresce with an intensity
proportional to m (Bernardi et
al., 1999
).
Erythrocytes were loaded with JC-1 at 16 µmol l-1 from a 10 mmol l-1 stock (in dimethylsulfoxide). To reduce the presence of dye particulates prior to incubation, the JC-1 solution was sonicated for 10 min followed by centrifugation (10 min at 14 000 g), with the supernatant being used for dye loading.
Erythrocytes were loaded with TMRM at 0.03-0.3 µmol l-1 from a 10 mmol l-1 stock (in ethanol). After loading, the cells were pelleted by a brief centrifugation (2 s pulse to 10 000 g) and resuspended in fresh incubation buffer prior to measuring fluorescence.
H2DCFDA and MitoSOXTM Red
Intracellular oxidative stress and free radical production were estimated
using the oxidation-sensitive dyes
2',7'-dichlorodihydrofluorescein diacetate (H2DCFDA)
and MitoSOXTM Red mitochondrial superoxide indicator (Invitrogen Corp.),
respectively. Both are nonfluorescent, cell-permeant dyes that form highly
fluorescent products upon oxidization.
Oxidation of H2DCFDA to the fluorescent product
2',7'-dichlorofluorescein (DCF) serves as an indicator of the
overall degree of intracellular oxidative stress
(Barja, 2002). Erythrocytes
were loaded with H2DCFDA at 1 µg ml-1 from a 3 mg
ml-1 stock solution (in dimethlysulfoxide). The cells were not
rinsed by centrifugation prior to measurement of fluorescence.
Intracellular accumulation of superoxide was estimated using MitoSOXTM Red, which is selectively targeted to the mitochondria, where it is oxidized by superoxide and exhibits red fluorescence upon binding to nucleic acids. Cells were loaded with MitoSOXTM at 5 µmol l-1, and were rinsed by centrifugation (as described above for TMRM) three times prior to measuring fluorescence.
Toxins
In vitro sulfide exposures were carried out in 20 ml glass vials.
After loading with dye, 100-400 µl of diluted erythrocytes were added to a
vial. The vial was then sealed with a rubber stopper and a volume of
H2S gas (pure or diluted 1:10 or 1:100 with room air) was added to
the vial air space to bring the final H2S composition in the gas
phase to 0.1%, 0.32%, 1.0%, 3.2% or 10.0%. Pure H2S gas was
prepared fresh daily in a fume hood by reacting 0.5 g Na2S
9H2O with 10 ml 6 mol l-1 HCl in a 50 ml syringe. After
injection of H2S, the vials were placed inside a dark, 15°C
incubator. After 1 h exposure, the erythrocytes were removed from the vials
and placed in microcentrifuge tubes, quickly pelleted by centrifugation (a 2 s
pulse to 10 000 g) and resuspended in fresh incubation medium.
In preliminary experiments, incubation buffer both with and without
erythrocytes was assayed for total sulfide after 1 h exposure to the range of
H2S gas compositions using the Methylene Blue method
(Cline, 1969). Total sulfide
concentration in buffer with erythrocytes was 0.77±0.14 the sulfide
concentration of buffer alone. In all experiments measuring
m, the
mitochondrial uncoupler carbonyl cyanide m-chlorophenylhydrazone
(CCCP; 0.10 mmol l-1 from a 20 mmol l-1 stock made in
ethanol and maintained at -20°C) was used as a positive control for
mitochondrial membrane depolarization.
A modification of the sulfide exposure technique was conducted to determine whether the effect of sulfide was reversible after several hours. Since mitochondrial dye would tend to leak back out of the cells over a long recovery period, the erythrocytes in this experiment were loaded with dye at the end of the recovery period rather than before sulfide exposure. Erythrocytes were first exposed to sulfide for 1 h as above, and then rinsed twice by centrifugation and allowed to `recover' at 15°C for an additional 2 or 5 h. The erythrocytes were then loaded with TMRM 30 min prior to the end of the recovery period and the fluorescence assayed as described above.
Incubations of erythrocytes with toxins other than sulfide were carried out in glass vials, as described above, or in 96-well microplates. To inhibit specific complexes of the mitochondrial electron transport chain, erythrocytes were exposed to the following toxins, for each of which the stock solutions were prepared immediately prior to the experiment: rotenone (complex I inhibitor; 10-100 µmol l-1 from a 20 mmol l-1 stock in dimethyl formamide), antimycin A (complex III inhibitor; 1-100 µmol l-1 from a 20 mmol l-1 stock in ethanol), azide (COX inhibitor; 1-10 mmol l-1 from a 2 mol l-1 N3Na stock in incubation buffer), and cyanide (COX inhibitor; 0.1-10 mmol l-1 from 20 mmol l-1 NaCN stock in incubation buffer).
To determine whether the change in m from sulfide exposure is
dependent on opening of the mitochondrial PT pore, G. dibranchiata
erythrocytes were exposed to sulfide for 1 h, as described above, except that
the PT pore inhibitor pair cyclosporine A (CsA; 0.5 µmol l-1)
and trifluoperazine (TFP; 5 µmol l-1) were added to the
erythrocytes 30 min prior to initiation of sulfide exposure
(Thompson et al., 2003
).
Control erythrocytes at each sulfide concentration were exposed to sulfide
alone or sulfide with CCCP (0.10 mmol l-1).
Measurement of fluorescence emission intensity
For toxin dose-response measurements using JC-1 and TMRM fluorescence
emission intensity, erythrocytes that were pre-loaded with dye and had already
been exposed to the toxin of interest were placed in a multi-well chamber
having a #1 glass coverslip bottom (this chamber was custom-manufactured, but
suitable plates are also available from Greiner Bio-One, Inc., Longwood, FL,
USA). The chamber was then placed on the stage of an inverted, reflected
fluorescence microscope (Olympus IX-70) and the erythrocytes were excited with
light from a 100 W mercury bulb that had passed through a 90% or 99% neutral
density filter (Thorlabs, Inc., Newton, NJ, USA) and a 484/15 nm (JC-1) or
555/15 nm (TMRM) excitation filter (Chroma Technology Corp, Rockingham, VT,
USA). The excitation filters were positioned and shuttered using a
computer-controlled filter wheel (TOFRA Inc., Palo Alto, CA, USA). Emitted
light from the erythrocytes was filtered with a triple bandpass polychroic
filter set (DAPI/FITC/TRITC; Chroma Technology Corp.) mounted in the standard
microscope filter carousel. For quantitation of emission intensity, light from
the microscope side-port was passed through a collimating lens (Thorlabs,
Inc.) coupled with an SMA connector (ThorLabs, Inc.) to a 600 µm diameter
fiber optic cable, and then to a linear diode-array fluorometer (SF2000, Ocean
Optics Inc., Dunedin, FL, USA). Each excitation typically lasted 1 s or less.
JC-1 fluorescence was quantified ratiometrically by simultaneously measuring
emission intensity at 590/20 nm and 530/20 nm (i.e. red:green emission ratio),
whereas TMRM fluorescence emission was measured only at 590/20 nm (i.e. red
emission intensity). All fluorescence measurements were performed at
18-20°C. Digital images were acquired with a monochrome, cooled CCD camera
equipped with a color LCD filter system (Retiga 2000R, QImaging Corp.,
Burnaby, BC, Canada).
For measurement of sulfide exposure effects over time, erythrocytes preloaded with TMRM were placed in a gas-tight Leiden chamber (Harvard Apparatus, Holliston, MA, USA) to which 10% H2S gas was added by syringe. Emission intensity was then measured, as otherwise described above, at 0, 5, 10, 20, 30, 45 and 60 min at the same location in the well, and therefore from the same erythrocyte population.
The JC-1 frequently loaded unevenly, because even after sonicating and centrifuging the JC-1 incubation solution, minute particles of dye frequently persisted and adhered to the coverslip or the erythrocytes. Therefore, to prevent these particles from biasing the measurements, JC-1 emission intensity was measured from single erythrocytes that were pre-selected as being appropriately labeled. In this technique, single erythrocytes were placed in the center of the microscope's field of view and imaged using a 100x oil-immersion objective with additional 1.5x magnification provided by the microscope's intermediate magnification changer. The excitation light was then constrained to just outside the perimeter of the cell using the Hg lamp field iris diaphragm, and the fluorescence emission was ported to the diode array fluorometer, as described above. Despite measuring from only a single cell, integration times of 64-256 ms were typically sufficient for counts reaching between 20% and 80% of the fluorometer's dynamic range. To avoid experimenter bias in selecting appropriate erythrocytes for measurement, the order of toxin treatments on the multi-well chamber was randomized by another person; i.e. the measurements were made `single-blind'. In contrast to JC-1, TMRM generally loaded evenly, so fluorescence emission was measured with a larger field of view (20x or 40x objective, thereby imaging up to hundreds of cells simultaneously, depending on the extent of dilution).
To confirm that effects of the sulfide on JC-1 and TMRM fluorescence were
due to a change in erythrocyte m rather than to direct chemical
interaction of sulfide with the dye molecules, JC-1 and TMRM were each diluted
in incubation buffer and exposed to control conditions (air) or 10%
H2S gas for 30 min in sealed gas vials. Each solution was then
placed in a 1 cm quartz fluorescence cuvette and excited at 484/15 nm (JC-1)
or 555/15 nm (TMRM) from the microscope Hg lamp via a fiber optic
cable. The emission light was then collected normal to the excitation light
with a collimating lens attached to a second fiber optic cable in a 4-way
cuvette holder (Ocean Optics, Inc.), and the emission spectrum was scanned
from 500-750 nm using the diode-array fluorometer. The sulfide concentration
of each dye solution was tested as above to confirm that H2S
exposures achieved sulfide concentrations of at least 1 mmol
l-1.
Oxidation of nonfluorescent H2DCFDA to fluorescent DCF and fluorescence of MitoSOXTM in erythrocytes exposed to sulfide were measured in black-wall, clear-bottom, fluorescence 96-well plates (Corning Inc. Life Sciences, Acton, MA, USA) with a multimode microplate reader (Synergy SIAFRT, Bio-Tek Instruments, Inc., Winooski, VT, USA) in bottom-reading mode using 485/20 nm excitation with 530/25 nm emission for H2DCFDA, and 528/20 nm excitation with 590/35 nm emission for MitoSOXTM.
Measurement of O2 consumption
To confirm that cyanide and azide were inhibiting COX, erythrocytes were
obtained from G. dibranchiata as described above, but the cells were
diluted 5x in pre-aerated incubation buffer. The diluted erythrocytes
were then placed in a 0.6 ml volume respiration chamber (Instech Laboratories,
Inc., Plymouth Meeting, PA, USA) that was held at 15°C with a
refrigerating, circulating water bath. The rate of decline in oxygen pressure
(PO2) was then measured using a polarographic electrode
until the PO2 reached ca. 80% of air-saturation (i.e. ca.
17 kPa), at which point 2 µl cyanide or azide were added from stock
solutions (prepared daily) using a 10 µl syringe (Hamilton Co., Reno, NV,
USA) to achieve 1 mmol l-1. The rate of PO2
decline was subsequently measured after toxin addition until the
PO2 was stable or reached 50% of air-saturation (ca. 10
kPa). Because the O2 P50 of G.
dibranchiata hemolysate is 0.60 kPa
(Harrington et al., 1978),
unloading of O2 at 50% air-saturation was minimal. A change in
O2 consumption rate before and after toxin addition was therefore
interpreted as a toxin effect. Any apparent increase in
PO2 immediately after addition of toxin was attributed to
a solvent artifact. Calibration of the O2 electrode was performed
daily following the manufacturer's instructions.
|
For sulfide dose-response measurements, the change in fluorescence emission
(or emission ratio) with increasing sulfide concentration was fit to the
standard sigmoidal dose-response curve
,
where a is the minimum relative fluorescence emission,
EC50 is the concentration of toxin at which TMRM fluorescence is
decreased by 50% relative to the control, x is log10 of
the toxin concentration, and n is the Hill coefficient. Data for the
curve fitting were normalized to the control value for each erythrocyte
population, and were fit using the non-linear curve-fitting function of
SigmaPlot 9.01 (Systat Software, Inc., Point Richmond, CA, USA).
![]() |
Results |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
Although we found that both JC-1 and TMRM labeled mitochondria in G.
dibranchiata erythrocytes, each dye had specific advantages and
disadvantages. JC-1 staining solution was more difficult to prepare, primarily
owing to its low solubility in aqueous medium and tendency to form
particulates that were difficult to remove even with prolonged centrifugation.
This necessitated measuring the fluorescence of erythrocytes individually in
the modified fluorescence microscope, which proved quite time-consuming (note
that a flow cytometer would likely be a useful alternative in this case).
However, an advantage of JC-1 was its specificity; it had little tendency to
label cellular structures other than mitochondria. Nonetheless, for reasons
that were not clear, JC-1 sometimes failed to label any structures. TMRM, in
contrast, while simple and inexpensive to prepare, had a tendency to label
non-mitochondrial structures (presumably endoplasmic reticulum or nuclear
membranes), especially at higher dye concentrations, necessitating careful
attention to loading conditions and confirmation of specificity by
fluorescence microscopy before each experiment. Another difference is that
while JC-1 fluorescence is measured at two emission wavelengths
ratiometrically, TMRM fluorescence is typically measured at a single
wavelength (Bernardi et al.,
1999), although note that TMRM emission can also be measured at
two excitation wavelengths, which has been reported to provide increased
sensitivity (Scaduto and Grotyohann,
1999
) and has been used for measuring
m in oyster
hemocytes (Sokolova et al.,
2004
). In our hands, JC-1 and TMRM had similar sensitivity and
signal-to-noise ratio (data not shown). On balance, TMRM proved the most
practical, and therefore after validating the effect of sulfide on
m in erythrocytes (below), the data for the remaining experiments
were with TMRM.
Sulfide-m dose-response
Exposure of G. dibranchiata erythrocytes loaded with JC-1 to
sulfide concentrations of 110 µmol l-1 to 1.9 mmol
l-1 for 1 h produced a significant decrease in the JC-1
fluorescence ratio (Fig. 3A;
N=8, RM ANOVA F8,64=11, P<0.0001),
consistent with a dose-dependent decrease in m, with a significant
change from control fluorescence first evident at 0.73 mmol l-1
(P=0.00052). The data fitted well with a sigmoidal dose-response
curve (r2=0.49, P<0.0001), with an apparent
50% maximal effect (EC50) at 0.50 mmol l-1 sulfide.
Exposure to CCCP (0.10 mmol l-1) for 1 h also produced a decrease
in the JC-1 fluorescence that was similar in magnitude to the decrease
recorded with the maximum sulfide concentration ratio
(Fig. 3B; N=8,
one-tail paired t-test t7=5.4,
P=0.00049). Erythrocytes loaded with TMRM showed a similar
dose-dependent decrease in fluorescence that was characteristic of a decrease
in
m (Fig. 3C;
N=4, RM ANOVA F7,21=6.7, P<0.00030),
with a significant change from control fluorescence first evident at 1.1 mmol
l-1 (P=0.0050). As with JC-1, the TMRM response fitted
well with a sigmoidal dose-response curve (r2=0.63,
P<0.0001), with the apparent EC50 at 0.83 mmol
l-1 sulfide. Exposure to CCCP for 1 h produced a decrease in
fluorescence that was similar to that of the highest sulfide concentration
(Fig. 3D; N=4,
one-tail paired t-test t3=3.8,
P=0.016).
|
|
Recovery from sulfide exposure
To determine whether the effect of sulfide on m is transient
and reversible, G. dibranchiata erythrocytes were exposed to a range
of sulfide concentrations for 1 h, rinsed twice in sulfide-free incubation
buffer and then allowed to recover for 2 or 5 h (3 and 6 h experiment
duration, respectively). Unlike the dose-response experiments (described in
the previous section), erythrocytes in this recovery experiment were not
loaded with TMRM until the last 30 min of the recovery period. Therefore,
given that H2S volatilization and sulfide oxidation should have
rapidly eliminated any free sulfide, it is reasonable to assume that
mitochondrial TMRM incorporation at 2 and 5 h recovery occurred under
sulfide-free conditions. After exposure to 0.3, 0.5, 0.8 and 1.2 mmol
l-1 sulfide for 1 h, there was a significant decrease in TMRM
fluorescence due to sulfide, but no effect of time and no interaction between
these factors (Fig. 5;
N=5, two-factor RM ANOVA; sulfide main effect
F4,8=33, P<0.0001; time main effect
F2,4=0.20, P=0.83; sulfidextime effect
F8,16=0.44, P=0.88). Immediately following 1 h
sulfide exposure, TMRM fluorescence was significantly decreased at all sulfide
concentrations compared to the control erythrocytes, as expected. After 2 h
recovery, only the erythrocytes that had been exposed to the highest sulfide
concentration were still significantly different from the control, whereas
after 5 hrecovery, erythrocytes exposed to all but the lowest sulfide
concentration were significantly below the control erythrocytes.
|
|
|
PT pore inhibitors
The PT pore inhibitors CsA (0.5 µmol l-1) and TFP (5 µmol
l-1) were added to G. dibranchiata erythrocytes prior to
sulfide exposure to test whether the decrease in m during sulfide
exposure was at least partially dependent on PT pore opening. As expected,
sulfide alone caused a decrease in TMRM fluorescence
(Fig. 8, circles, N=5,
F3,9=8.0, P=0.0067), but the addition of CsA and
TFP with sulfide exposure did not affect TMRM fluorescence compared to sulfide
alone (Fig. 8, triangles,
N=4-5, P=0.087). In fact, addition of CsA and TFP caused
decreased fluorescence even in the absence of sulfide (P=0.010). CCCP
caused a decrease in fluorescence in the absence of sulfide
(P=0.00016), as expected, but TMRM fluorescence in erythrocytes
exposed to the combination of 0.73 mmol l-1 sulfide with CCCP was
significantly lower than fluorescence in erythrocytes exposed to CCCP alone
(i.e. there was an additive effect of sulfide on CCCP;
Fig. 8, squares,
F3,12=13, P=0.00041).
|
|
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
When G. dibranchiata erythrocytes loaded with JC-1 or TMRM were
exposed to 1.9 mmol l-1 sulfide for 1 h, the fluorescence emission
of both dyes changed in a manner similar to that seen after exposure of
erythrocytes to CCCP (i.e. decreased red:green ratio with JC-1 and decreased
red intensity with TMRM). This suggested that sulfide caused a loss of
m. However, an alternative explanation for the change in
fluorescence would be that sulfide (whether as H2S, HS-
or S2-) interacted directly with each dye to cause decreased red
fluorescence. However, two lines of evidence suggest that this is not likely.
First, when JC-1 and TMRM were dissolved in incubation buffer (without
erythrocytes) and exposed to 1 mmol l-1 sulfide for 30 min, the
fluorescence emission spectra of both dyes were indistinguishable from their
spectra obtained prior to sulfide exposure
(Fig. 2). Second, JC-1 and TMRM
were specifically selected for this study because they differ substantially in
chemical structure and action, even though they are both lipophilic cations
that accumulate in mitochondria with high
m. One consequence is
that JC-1 exists as a green-fluorescent monomer in cells having mitochondria
with low
m, but forms red-fluorescent `J-aggregates' upon
concentration into mitochondrial membranes with high
m
(Smiley et al., 1991
), whereas
TMRM remains as a monomer in membranes with high
m but exhibits a
red-shift in the fluorescence emission spectrum
(Scaduto and Grotyohann,
1999
). Therefore, we consider it unlikely that both dyes interact
directly with sulfide to cause fluorescence changes mimicking loss of
m.
After exposure to a range of sulfide concentrations for 1 h, G.
dibranchiata erythrocytes loaded with JC-1 or TMRM exhibited a
concentration-dependent change in fluorescence consistent with decreased
m (Fig. 3).
Furthermore, the magnitude of the fluorescence change at sulfide
concentrations of 1.2 mmol l-1 and higher was similar to that of
CCCP, suggesting that sulfide induced mitochondrial depolarization to an
extent equivalent to pharmacological uncoupling. Similarly, Eghbal et al.
(2004
) used rhodamine 123
incorporation (rather than fluorescence change) to show that isolated rat
hepatocytes exposed to 0.5 mmol l-1 sulfide for 1 h decreased
m by 50%. Using CCCP-induced depolarization as a reference point,
we found a comparable sensitivity in G. dibranchiata erythrocytes, in
which the sulfide EC50 for
m was 0.50-0.83 mmol
l-1 (using JC-1 and TMRM, respectively), suggesting a surprisingly
similar sulfide cytotoxicity for rats and G. dibranchiata.
Although inhibition of COX by sulfide is reversible in vitro
(Nicholls and Kim, 1982) and
probably in vivo (Blackstone et
al., 2005
), the loss of
m in G. dibranchiata
erythrocytes following 1 h sulfide exposure in vitro was irreversible
over a subsequent 5 h recovery period (Fig.
5). This implied that the toxic effect of sulfide was not simply
due to inhibition of COX. We further tested this by exposing erythrocytes to
four other mitochondrial electron transport chain inhibitors: rotenone,
antimycin, azide and cyanide. Of these, only rotenone and cyanide caused a
significant change in
m, but for neither of these inhibitors was
the degree of depolarization comparable to that achieved with CCCP or sulfide
(Fig. 6). That 10 mmol
l-1 azide and cyanide had either no effect or caused only a 20%
decrease in
m (in comparison to CCCP) was surprising since, like
sulfide, both are COX inhibitors, and both caused an immediate, substantial
reduction in O2 consumption even when applied at only 1 mmol
l-1 concentration (Fig.
7). Taken together, the irreversibility of sulfide and the absence
of a similar effect from other COX inhibitors provide strong evidence that the
mechanism by which sulfide causes mitochondrial depolarization is not
via COX inhibition. A similar conclusion was reached by Thompson et
al. (2003
) who, like Eghbal et
al. (2004
), studied the effect
of sulfide on primary rat hepatocytes in culture. Thompson et al.
(2003
) found that hepatocytes
exposed to sulfide or cyanide were killed in a dose-dependent manner, as
expected, but differed in their response to supplementation by glycolytic
substrate. Specifically, addition of fructose to the hepatocytes, which
greatly enhances glycolytic ATP production in these cells
(Nieminen et al., 1994
),
substantially decreased the toxicity of cyanide, but had no effect on the
toxicity of sulfide, and the PT pore inhibitors CsA and TFP decreased cell
death during sulfide exposure but not during cyanide exposure
(Thompson et al., 2003
). It
should noted, however, that since the response of hepatocytes to fructose
supplementation is not a characteristic of other mammalian cells, the reduced
toxicity of cyanide, but not sulfide, with added fructose may be unique to
liver.
Mitochondrial membrane permeability
Sulfide-induced, irreversible opening of the PT pore would be consistent
with the loss of m seen in G. dibranchiata erythrocytes.
In support of this, the PT pore inhibitors CsA and TFP decreased cell death by
up to 50% in rat hepatocytes exposed to sulfide
(Thompson et al., 2003
).
However, in the present study, addition of CsA and TFP did not prevent loss of
m during sulfide exposure (Fig.
8). Thus, CsA and TFP together reduce sulfide cytotoxicity but not
sulfide-induced loss of
m. Reconciling these findings requires
that (1) sulfide-induced loss of
m does not necessarily lead to
erythrocyte death and (2) at least some sulfide-induced cell death is
via a mechanism that is inhibited by CsA and TFP but does not result
in maintenance of
m. Although this has not been tested with
sulfide, loss of
m can result from many events other than PT pore
opening in mammalian cells, including increased ATP demand and uncoupling of
oxidative phosphorylation (Bernardi et
al., 2001
; Green and Kroemer,
2004
; Ly et al.,
2003
), and cell death is not necessarily a consequence if
m loss is transient (Ly et
al., 2003
). With regard to an alternative protective role of CsA
or TFP, in addition to its effect on the PT pore, CsA inhibits all
cyclophilins and also has recently been shown to inhibit Ca2+
uptake by mitochondria through the Ca2+ uniporter
(Montero et al., 2003
).
Furthermore, release of pro-apoptotic proteins from mitochondria requires
increased permeability of the mitochondrial outer membrane, whereas the PT
pore spans the inner membrane. The apparent mechanistic link is that PT pore
opening with the subsequent influx of ions into the mitochondrial matrix
promotes the osmosis of sufficient water into the mitochondrial matrix to
rupture the outer membrane (Green and
Kroemer, 2004
; Halestrap et
al., 2002
). Therefore, it is worth noting that neither sulfide nor
cyanide caused swelling in mitochondria from rat hepatocytes
(Thompson et al., 2003
),
suggesting that neither caused PT pore opening.
Nonetheless, this does not rule out PT pore opening in sulfide-exposed
erythrocytes. The structure of the PT pore is controversial, but it appears to
be a complex with three major components: a voltage-dependent anion channel
(VDAC) from the outer membrane, an adenine nucleotide translocator (ANT) from
the inner membrane, and cyclophilin D (CyP-D), which binds to ANT and promotes
PT pore opening (Green and Kroemer,
2004; Halestrap et al.,
2002
; Kim et al.,
2003a
; Ly et al.,
2003
). CsA normally inhibits PT pore opening by binding to CyP-D,
but pore opening is insensitive to CsA at high concentrations of various
inducers (Halestrap et al.,
2002
; He and Lemasters,
2002
), suggesting that sulfide could initiate PT pore opening even
in the presence of CsA. Furthermore, TFP is not a potent inhibitor of PT pore
opening under de-energized conditions
(Halestrap et al., 1997
), and
therefore may not protect cells from sulfide exposure. Finally, the ANT has
three cysteine residues that appear to regulate CyP-D activity and that can be
modified by thiol reagents and reactive oxygen species
(Halestrap et al., 1997
).
Unlike cyanide or azide, sulfide is a highly reactive thiol compound that may
be capable of directly modifying one or more of these residues, potentially
inducing pore opening. Finally, an additional possibility is that sulfide
induces opening of an unregulated PT pore in G. dibranchiata
erythrocytes. The presence of an unregulated pore, which is not inhibited by
CsA (He and Lemasters, 2002
;
Menze et al., 2005
), has been
proposed in the crustacean A. franciscana
(Menze et al., 2005
).
ROS production and oxidative stress
Increased oxidative stress is an additional mechanism by which sulfide
exposure could cause toxicity
(Abele-Oeschger, 1996;
Abele-Oeschger et al., 1994
;
Morrill et al., 1988
), and
therefore mitochondrial depolarization. Sulfide oxidizes spontaneously in the
presence of divalent metals (both dissolved and in metalloenzymes), generating
oxygen-centered and sulfur-centered radicals in aqueous solutions
(Chen and Morris, 1972
;
Tapley et al., 1999
) and in
animal tissues (Tapley, 1993
).
Recently, Eghbal et al. (2004
)
used the ROS indicator H2DCFDA in rat hepatocytes and showed that
free radical-induced oxidation of this dye to DCF was 2-3 times faster when
the cells were exposed to 0.5 mmol l-1 sulfide than when they were
exposed to cyanide or control conditions. Furthermore, the addition of ROS
scavengers decreased cell death by up to 40% in hepatocytes exposed to 0.5
mmol l-1 sulfide for 3 h, although this protective effect was not
seen after 1 h (Eghbal et al.,
2004
). In this study, we found that G. dibranchiata
erythrocytes loaded with H2DCFDA and exposed to sulfide for 1 h
showed a 60% increase in DCF fluorescence at the highest sulfide concentration
(1.9 mmol l-1), indicating oxidation of H2DCFDA
(Fig. 9A). This effect was not
seen at 0.73 mmol l-1 and lower sulfide concentrations. Increased
H2DCFDA oxidation was not due to spontaneous sulfide oxidation,
since H2DCFDA in cell-free incubation medium oxidized to DCF at
less than 2% of the rate seen in medium with erythrocytes. Therefore, the
effect of sulfide on H2DCFDA in G. dibranchiata
erythrocytes is comparable to its effect in rat hepatocytes, although in
G. dibranchiata the effect is right-shifted with respect to sulfide
concentration. Because superoxide may be a reaction product in the first
sulfide oxidation step (Chen and Morris,
1972
; Tapley et al.,
1999
), we specifically investigated whether sulfide exposure
increases superoxide generation. We utilized the dye MitoSOXTM Red, which
is taken up by mitochondria and is readily oxidized by superoxide but not by
other ROS or reactive nitrogen species (data from Invitrogen Corp.). Similar
to the results with oxidative stress indicator H2DCFDA, we found
that MitoSOXTM oxidation was increased threefold in G.
dibranchiata erythrocytes exposed to 1.9 mmol l-1 sulfide for
1 h, but not by exposure to 0.73 mmol l-1 or lower sulfide
concentrations (Fig. 9B). In
addition to sulfide oxidation as a source, superoxide is also generated within
mitochondria as a byproduct of oxidative phosphorylation
(Halliwell and Gutteridge,
1999
), and the present study does not distinguish between these
sources. Theoretically, increased mitochondrial ROS production may also occur
specifically from periodic exposure to sulfide (such as with tidal cycles for
mudflat animals), because the resultant periodic COX inhibition might increase
mitochondrial free-radical production by a process essentially identical to
that of hypoxia/reoxygenation injury (Kim
et al., 2003b
). However, whether this occurs following sulfide
exposure has not been investigated.
While increased oxidation of H2DCFDA will result from augmented
ROS production, DCF production may also be enhanced by depletion of
intracellular antioxidants. For this reason, the rate of DCF production is
best interpreted as an estimator of the overall degree of oxidative stress
within cells (Barja, 2002),
with the same presumably also applying to MitoSOXTM. Therefore, since
significant H2DCFDA oxidation occurred at lower sulfide
concentrations in rat hepatocytes than in G. dibranchiata
erythrocytes, this suggests that either G. dibranchiata erythrocytes
have increased antioxidant defenses compared to rat hepatocytes, or that rat
hepatocytes produce more free radicals than G. dibranchiata
erythrocytes at a given sulfide concentration. In fact, both G.
dibranchiata erythrocytes and rat hepatocytes may be particularly
susceptible to oxidative damage from sulfide exposure, since both contain high
concentrations of heme-containing proteins, which can act as pro-oxidants
(Halliwell and Gutteridge,
1999
). Specifically, G. dibranchiata erythrocytes contain
hemoglobin (Mangum et al.,
1989
), while liver cells have high concentrations of CYP450.
Consistent with this, Eghbal et al.
(2004
) showed that addition of
the CYP450 inhibitor benzylimidazole decreased sulfide cytotoxicity by up to
50% in rat hepatocytes. Antioxidant defenses of sulfide-adapted animals
against sulfide exposure could include increased activities or concentrations
of antioxidant enzymes such as superoxide dismutase and catalase, but also
increased concentrations of low molecular mass free radical scavengers.
Interestingly, glutathione and hypotaurine, which are present in high
concentrations in tissues of sulfide-adapted invertebrates
(Pruski et al., 1997
;
Yancey et al., 2002
;
Yin et al., 2000
), are both
potent free radical scavengers (Halliwell
and Gutteridge, 1999
).
Significance
Many sulfidic environments have communities of invertebrates adapted to
sulfide concentrations from 0.05 mmol l-1 to more than 12 mmol
l-1 (Childress and Fisher,
1992; Grieshaber and
Völkel, 1998
; Urcuyo et
al., 2003
; Van Dover and Lutz,
2004
). Although intact sulfide detoxification mechanisms may
reduce sulfide toxicity for the majority of cells in these animals,
integumentary tissues, epithelial tissues at respiratory surfaces and
circulating respiratory cells are still exposed to potentially toxic sulfide
concentrations. If the irreversible mitochondrial toxicity observed in the
present study occurs in epithelial cells of sulfide-adapted animals in their
natural environment, then it may be an important factor affecting sulfide
tolerance by necessitating substantially upregulated mitochondrial repair or
degradation and biogenesis. In support of this, some histological studies of
epithelial tissues from sulfide-adapted, sulfide-exposed invertebrates have
identified mitochondrial swelling and the presence of electron-dense
mitochondrial matrices and granules (Duffy
and Tyler, 1984
; Janssen and
Oeschger, 1992
; Jouin and
Gaill, 1990
; Menon and Arp,
1993
,
1998
;
Menon et al., 2003
), which may
be evidence of mitochondrial injury, but such changes have not been seen in
all studies (Dubilier et al.,
1997
). Irreversible mitochondrial depolarization from sulfide
would be likely to result in MOMP, which would be followed by release of
mitochondrial pro-apoptotic factors into the cytoplasm and cell death soon
thereafter (Green and Kroemer,
2004
). Indeed, the smooth appearance of G. dibranchiata
erythrocytes exposed to sulfide (Fig.
1) suggests cellular swelling, which is a characteristic of cell
death following PT pore opening in mammalian cells
(Nieminen, 2003
). Once
mitochondrial injury and depolarization begin, the only mechanism available to
the organism to prevent cell death would be autophagic ingestion of the
injured mitochondria (Bauvy et al.,
2001
; Shintani and Klionsky,
2004
). Indeed, it has been proposed that electron-dense
organelles, which are characteristic of epidermal tissues in sulfide-adapted
annelids (e.g. Giere et al.,
1988
; Hourdez and
Jouin-Toulmond, 1998
; Jouin
and Gaill, 1990
;
Jouin-Toulmond et al., 1996
;
Menon et al., 2003
), represent
`secondary lysosomes' containing autophagocytosed, sulfide-damaged and
degenerating mitochondria (Arp et al.,
1995
). Whether mitochondrial injury, represented by
depolarization, results in increased mitochondrial autophagy remains to be
determined.
Conclusions
In this study, we demonstrate that mitochondria in G. dibranchiata
erythrocytes can be successfully labeled with the m indicator dyes
JC-1 and TMRM and that these dyes show changes in fluorescence emission
characteristic of mitochondrial depolarization when the erythrocytes are
exposed to the mitochondrial uncoupler CCCP. When erythrocytes were exposed to
various concentrations of sulfide for 1 h, JC-1 and TMRM showed a similar,
dose-dependent change in fluorescence, suggesting that sulfide caused loss of
m. This change was irreversible over 5 h and was not seen to the
same extent with azide or cyanide, which both inhibit COX, or with other
mitochondrial electron transport chain inhibitors, with the possible exception
of rotenone. JC-1 and TMRM in cell-free incubation buffer showed no change in
fluorescence spectra when exposed to sulfide, which increases our confidence
that the changes in fluorescence emission observed in these dyes when loaded
into G. dibranchiata erythrocytes exposed to sulfide are not due to a
chemical interaction between each dye and sulfide. Furthermore, erythrocytes
that had been exposed to sulfide for 1 h, then rinsed in sulfide-free buffer,
incubated in sulfide-free conditions for up to 5 h and only then loaded with
TMRM, showed fluorescence similar to that of cells immediately following
exposure to sulfide or CCCP (Fig.
5). Since it is highly improbable that any free sulfide would have
remained after 5 h, it is likely the fluorescence change represents loss of
m.
The mechanism by which sulfide irreversibly depolarizes mitochondria is not
clear. Although the PT pore inhibitors CsA and TFP have been shown to improve
cell survival during sulfide exposure, we found that they did not prevent loss
of m. One potential mechanism of mitochondrial injury is free
radical damage from increased ROS production. This was supported by the
observation that H2DCFDA oxidation and MitoSOXTM oxidation
were increased two- to threefold in erythrocytes exposed to sulfide,
suggesting increased oxidative stress and superoxide production, respectively.
However, it remains to be determined whether ROS production is a contributing
factor to, or a consequence of,
m loss during sulfide
exposure.
![]() |
Acknowledgments |
---|
![]() |
References |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Abele-Oeschger, D. (1996). A comparative study of superoxide dismutase activity in marine benthic invertebrates with respect to environmental sulphide exposure. J. Exp. Mar. Biol. Ecol. 197,39 -49.[CrossRef]
Abele-Oeschger, D., Oeschger, R. and Theede, H. (1994). Biochemical adaptations of Nereis diversicolor (Polychaeta) to temporarily increased hydrogen peroxide levels in intertidal sandflats. Mar. Ecol. Prog. Ser. 106,101 -110.
Akiyama, T. and Okada, M. (1992). Spatial and
developmental changes in the respiratory activity of mitochondria in early
Drosophila embryos. Development
115,1175
-1182.
Arp, A. J., Menon, J. G. and Julian, D. (1995). Multiple mechanisms provide tolerance to environmental sulfide in Urechis caupo. Am. Zool. 35,132 -144.
Bagarinao, T. (1992). Sulfide as an environmental factor and toxicant: tolerance and adaptations in aquatic organisms. Aquatic Toxicol. 24, 21-62.[CrossRef]
Bagarinao, T. and Vetter, R. D. (1992). Sulfide-hemoglobin interactions in the sulfide-tolerant salt-marsh resident, the California killifish Fundulus parvipinnis. J. Comp. Physiol. B 162,614 -624.
Barja, G. (2002). The quantitative measurement of H2O2 generation in isolated mitochondria. J. Bioenerg. Biomem. 34,227 -233.[CrossRef][Medline]
Bauvy, C., Gane, P., Arico, S., Codogno, P. and Ogier-Denis, E. (2001). Autophagy delays sulindac sulfide-induced apoptosis in the human intestinal colon cancer cell line HT-29. Exp. Cell Res. 268,139 -149.[CrossRef][Medline]
Beauchamp, R. O., Bus, J. S., Popp, J. S., Boreiko, C. J. and Andjelkovich, D. A. (1984). A critical review of the literature on hydrogen sulfide toxicity. CRC Crit. Rev. Toxicol. 13,25 -97.
Bernardi, P., Scorrano, L., Colonna, R., Petronilli, V. and Di
Lisa, F. (1999). Mitochondria and cell death: mechanistic
aspects and methodological issues. Eur. J. Biochem.
264,687
-701.
Bernardi, P., Petronilli, V., Di Lisa, F. and Forte, M. (2001). A mitochondrial perspective on cell death. Trends Biochem. Sci. 26,112 -117.[CrossRef][Medline]
Blackstone, E., Morrison, M. and Roth, M. B.
(2005). H2S induces a suspended animation-like state
in mice. Science 308,518
.
Chen, K. Y. and Morris, J. C. (1972). Oxidation of sulfide by O2: catalysis and inhibition. J. San. Eng. Div. Proc., Am. Soc. Civ. Eng. 98,215 -227.
Childress, J. J. and Fisher, C. R. (1992). The biology of hydrothermal vent animals: physiology, biochemistry, and autotrophic symbioses. Oceanog. Mar. Biol. Annu. Rev. 30,337 -441.
Cline, J. D. (1969). Spectrophotometric determination of hydrogen sulfide in natural waters. Limnol. Oceanog. 14,454 -458.
Cossarizza, A., Cooper, E. L., Quaglino, D., Salvioli, S., Kalachnikova, G. and Franceschi, C. (1995). Mitochondrial mass and membrane potential in celomocytes from the earthworm Eisenia foetida: studies with fluorescent probes in single intact cells. Biochem. Biophys. Res. Commun. 214,503 -510.[CrossRef][Medline]
Dorman, D. C., Moulin, F. J.-M., McManus, B. E., Mahle, K. C.,
Arden James, R. and Struve, M. F. (2002). Cytochrome
oxidation inhibition induced by acute hydrogen sulfide inhalation: correlation
with tissue sulfide concentrations in the rat brain, liver, lung, and nasal
epithelium. Toxicol. Sci.
65, 18-25.
Dubilier, N., Windoffer, R., Grieshaber, M. K. and Giere, O. (1997). Ultrastructure and anaerobic metabolism of mitochondria in the marine oligochaete Tubificoides benedii - effects of hypoxia and sulfide. Mar. Biol. 127,637 -645.[CrossRef]
Duffy, J. and Tyler, S. (1984). Quantitative differences in mitochondrial ultrastructure of a thiobiotic and an oxybiotic turbellarian. Mar. Biol. 83, 95-102.[CrossRef]
Eghbal, M. A., Pennefather, P. S. and O'Brien, P. J. (2004). H2S cytotoxicity mechanism involves reactive oxygen species formation and mitochondrial depolarisation. Toxicol. 203,69 -76.[CrossRef][Medline]
Feeney, C. J., Pennefather, P. S. and Gyulkhandanyan, A. V. (2003). A cuvette-based fluorometric analysis of mitochondrial membrane potential measured in cultured astrocyte monolayers. J. Neurosci. Methods 125,13 -25.[CrossRef][Medline]
Gainey, L. F. and Greenberg, M. J. (2005).
Hydrogen sulfide synthesized in the gills of the clam Mercenaria
mercenaria acts seasonally as a modulator of branchial muscle
contraction. Biol. Bull.
209, 11-20.
Giere, O., Rhode, B. and Dubilier, N. (1988). Structural peculiarities of the body wall of Tubificoides benedii (Oligochaeta) and possible relations to its life in sulphidic sediments. Zoomorphol. 108,29 -39.[CrossRef]
Green, D. R. and Kroemer, G. (2004). The
pathophysiology of mitochondrial cell death. Science
305,626
-629.
Grieshaber, M. K. and Völkel, S. (1998). Animal adaptations for tolerance and exploitation of poisonous sulfide. Annu. Rev. Physiol. 60,33 -53.[CrossRef][Medline]
Halestrap, A. P., Woodfield, K. Y. and Connern, C. P.
(1997). Oxidative stress, thiol reagents, and membrane potential
modulate the mitochondrial permeability transition by affecting nucleotide
binding to the adenine nucleotide translocase. J. Biol.
Chem. 272,3346
-3354.
Halestrap, A. P., McStay, G. P. and Clarke, S. J. (2002). The permeability transition pore complex: another view. Biochimie 84,153 -166.[CrossRef][Medline]
Halliwell, B. and Gutteridge, J. (1999). Free Radicals in Biology and Medicine. Oxford: Oxford University Press.
Hand, S. C. and Somero, G. N. (1983). Energy-metabolism pathways of hydrothermal vent animals - adaptations to a food-rich and sulfide-rich deep-sea environment. Biol. Bull. 165,167 -181.
Harrington, J. P., Suarez, G., Borgese, T. A. and Nagel, R. L. (1978). Subunit interactions of Glycera dibranchiata hemoglobin. J. Biol. Chem. 253,6820 -6825.[Abstract]
He, L. and Lemasters, J. J. (2002). Regulated and unregulated mitochondrial permeability transition pores: a new paradigm of pore structure and function? FEBS Lett. 512, 1-7.[CrossRef][Medline]
Hourdez, S. and Jouin-Toulmond, C. (1998). Functional anatomy of the respiratory system of Branchipolynoe species (Polychaeta, Polynoidae), commensal with Bathymodiolus species (Bivalvia, Mytilidae) from deep sea hydrothermal vents. Zoomorphol. 118,225 -233.[CrossRef]
Hunter, D. R., Haworth, R. A. and Southard, J. H. (1976). Relationship between configuration, function, and permeability in calcium-treated mitochondria. J. Biol. Chem. 251,5069 -5077.[Abstract]
Janssen, H. and Oeschger, R. (1992). The body wall of Halicryptus spinulosus (priapulida) - ultrastructure and changes induced by hydrogen sulfide. Hydrobiologia 230,219 -230.[CrossRef]
Jensen, M. S., Ahlemeyer, B., Ravati, A., Thakur, P., Mennel, H. D. and Krieglstein, J. (2002). Preconditioning-induced protection against cyanide-induced neurotoxicity is mediated by preserving mitochondrial function. Neurochem. Int. 40,285 -293.[CrossRef][Medline]
Jouin, C. and Gaill, F. (1990). Gills of hydrothermal vent annelids: structure, ultrastructure and functional implications in two alvinellid species. Prog. Oceanog. 24, 59-69.[CrossRef]
Jouin-Toulmond, C., Augustin, D., Desbruyeres, D. and Toulmond, A. (1996). The gas transfer system in alvinellids (annelida polychaeta, terebellida). Anatomy and ultrastructure of the anterior circulatory system and characterization of a coelomic, intracellular haemoglobin. Cahiers Biol. Mar. 37,135 -151.
Julian, D., Statile, J., Roepke, T. and Arp, A.
(2005). Sodium nitroprusside potentiates H2S-induced
contractions in body wall muscle from a marine worm. Biol.
Bull. 209,6
-10.
Khan, A. A., Schuler, M. M., Prior, M. G., Yong, S., Coppock, R. W., Florence, L. Z. and Lillie, L. E. (1990). Effects of hydrogen sulfide exposure on lung mitochondrial respiratory chain enzymes in rats. Toxicol. Appl. Pharmacol. 103,482 -490.[CrossRef][Medline]
Kim, J. S., He, L. and Lemasters, J. J. (2003a). Mitochondrial permeability transition: a common pathway to necrosis and apoptosis. Biochem. Biophys. Res. Commun. 304,463 -470.[CrossRef][Medline]
Kim, J. S., He, L., Qian, T. and Lemasters, J. J. (2003b). Role of the mitochondrial permeability transition in apoptotic and necrotic death after ischemia/reperfusion injury to hepatocytes. Curr. Mol. Med. 3,527 -535.[CrossRef][Medline]
Kimura, H. (2002). Hydrogen sulfide as a neuromodulator. Mol. Neurobiol. 26, 13-19.[CrossRef][Medline]
Kraus, D. W., Doeller, J. E. and Powell, C. S.
(1996). Sulfide may directly modify cytoplasmic hemoglobin
deoxygenation in Solemya reidi gills. J. Exp.
Biol. 199,1343
-1352.
Lawrence, C. L., Billups, B., Rodrigo, G. C. and Standen, N. B. (2001). The KATP channel opener diazoxide protects cardiac myocytes during metabolic inhibition without causing mitochondrial depolarization or flavoprotein oxidation. Br. J. Pharmacol. 134,535 -542.[CrossRef][Medline]
Lloyd, D., Kristensen, B. and Degn, H. (1982). The effects of cyanide, azide, carbon monoxide and salicylhydroxamic acid on whole-cell respiration of Acanthamoeba castellanii. J. Gen. Microbiol. 128,185 -188.[Medline]
Ly, J. D., Grubb, D. R. and Lawen, A. (2003). The mitochondrial membrane potential (deltapsi(m)) in apoptosis; an update. Apoptosis 8,115 -128.[CrossRef][Medline]
Mangum, C. P. (1994). Multiple sites of gas-exchange. Am. Zool. 34,184 -193.
Mangum, C. P., Colacino, J. M. and Vandergon, T. L. (1989). Oxygen binding of single red blood-cells of the annelid bloodworm Glycera dibranchiata. J. Exp. Zool. 249,144 -149.[CrossRef]
Menon, J. and Arp, A. (1993). The integument of
the marine echiuran worm Urechis caupo. Biol.
Bull. 185,440
-454.
Menon, J. G. and Arp, A. J. (1998). Ultrastructural evidence of detoxification in the alimentary canal of Urechis caupo. Invert. Biol. 117,307 -317.
Menon, J., Willsie, J. K., Tauscher, A. and Arp, A. J. (2003). Epidermal ultrastructure and implications for sulfide tolerance in six species of deep-sea polychaetes. Invert. Biol. 122,334 -346.
Menze, M. A., Hutchinson, K., Laborde, S. M. and Hand, S. C. (2005). Mitochondrial permeability transition in the crustacean Artemia franciscana: Absence of a Ca2+-regulated pore in the face of profound calcium storage. Am. J. Physiol. 289,R68 -R76.[CrossRef]
Montero, M., Lobaton, C. D., Gutierrez-Fernandez, S., Moreno, A. and Alvarez, J. (2003). Calcineurin-independent inhibition of mitochondrial Ca2+ uptake by cyclosporin A. Br. J. Pharmacol. 141,263 -268.[Medline]
Morrill, A. C., Powell, E. N., Bidigare, R. R. and Shick, J. M. (1988). Adaptations to life in the sulfide system: a comparison of oxygen detoxifying enzymes in thiobiotic and oxybiotic meiofauna (and freshwater planarians). J. Comp. Physiol. 158B,335 -344.
Nguyen, P. V., Marin, L. and Atwood, H. L.
(1997). Synaptic physiology and mitochondrial function in
crayfish tonic and phasic motor neurons. J.
Neurophysiol. 78,281
-294.
Nicholls, P. (1975). The effect of sulphide on cytochrome aa3. Isosteric and allosteric shifts of the reduced alpha-peak. Biochim. Biophys. Acta 396, 24-35.[Medline]
Nicholls, P. and Kim, J. K. (1982). Sulphide as an inhibitor and electron donor for the cytochrome c oxidase system. Can. J. Biochem. 60,613 -623.[Medline]
Nieminen, A. L. (2003). Apoptosis and necrosis in health and disease: role of mitochondria. Int. Rev. Cytol. 224,29 -55.[Medline]
Nieminen, A. L., Saylor, A. K., Herman, B. and Lemasters, J. J. (1994). ATP depletion rather than mitochondrial depolarization mediates hepatocyte killing after metabolic inhibition. Am. J. Physiol. 267,C67 -C74.[Medline]
Powell, M. A. and Arp, A. J. (1989). Hydrogen sulfide oxidation by abundant nonhemoglobin heme compounds in marine invertebrates from sulfide-rich habitats. J. Exp. Zool. 249,121 -132.[CrossRef]
Prabhakaran, K., Li, L., Borowitz, J. L. and Isom, G. E.
(2002). Cyanide induces different modes of death in cortical and
mesencephalon cells. J. Pharmacol. Exp. Ther.
303,510
-519.
Pruski, A. M., Fiala-Médioni, A. and Colomines, J.-C. (1997). High amounts of sulphur-amino acids in three symbiotic mytilid bivalves from deep benthic communities. Compt. Rend. 320,791 -796.
Rosenegger, D., Roth, S. and Lukowiak, K.
(2004). Learning and memory in Lymnaea are negatively altered by
acute low-level concentrations of hydrogen sulphide. J. Exp.
Biol. 207,2621
-2630.
Scaduto, R. C., Jr and Grotyohann, L. W.
(1999). Measurement of mitochondrial membrane potential using
fluorescent rhodamine derivatives. Biophys. J.
76,469
-477.
Shintani, T. and Klionsky, D. J. (2004).
Autophagy in health and disease: A double-edged sword.
Science 306,990
-995.
Smiley, S., Reers, M., Mottola-Hartshorn, C., Lin, M., Chen, A.,
Smith, T., Steele, G., Jr and Chen, L. (1991).
Intracellular heterogeneity in mitochondrial membrane potentials revealed by a
J-aggregate-forming lipophilic cation JC-1. Proc. Natl. Acad. Sci.
USA 88,3671
-3675.
Smith, L., Kruszyna, H. and Smith, R. P. (1977). The effect of methemoglobin on the inhibition of cytochrome c oxidase by cyanide, sulfide or azide. Biochem. Pharmacol. 26,2247 .[CrossRef][Medline]
Sokolova, I. M., Evans, S. and Hughes, F. M.
(2004). Cadmium-induced apoptosis in oyster hemocytes involves
disturbance of cellular energy balance but no mitochondrial permeability
transition. J. Exp. Biol.
207,3369
-3380.
Tapley, D. (1993). Sulfide-Dependent Oxidative Stress in Marine Invertebrates, especially Thiotrophic Symbioses, 159pp. Maine: University of Maine Press.
Tapley, D. W., Beuttner, G. R. and Shick, J. M.
(1999). Free radicals and chemiluminescence as products of the
spontaneous oxidation of sulfide in seawater, and their biological
implications. Biol. Bull.
196, 52-56.
Thompson, R. W., Valentine, H. L. and Valentine, W. M. (2003). Cytotoxic mechanisms of hydrosulfide anion and cyanide anion in primary rat hepatocyte cultures. Toxicol. 188,149 -159.[Medline]
Urcuyo, I. A., Massoth, G. J., Julian, D. and Fisher, C. R. (2003). Habitat, growth and physilogical ecology of a basaltic community of Ridgeia piscesae from the Juan de Fuca Ridge. Deep-Sea Res. (I, Ocean. Res. Papers) 50,763 -780.[CrossRef]
Van Dover, C. L. and Lutz, R. A. (2004). Experimental ecology at deep-sea hydrothermal vents: a perspective. J. Exp. Mar. Biol. Ecol. 300,273 -307.[CrossRef]
Völkel, S. and Berenbrink, M. (2000).
Sulphaemoglobin formation in fish: A comparison between the haemoglobin of the
sulphide-sensitive rainbow trout (Oncorhynchus mykiss) and of the
sulphide-tolerant common carp (Cyprinus carpio). J. Exp.
Biol. 203,1047
-1058.
Völkel, S. and Grieshaber, M. K. (1994). Oxygen dependent sulfide detoxification in the lugworm Arenicola marina. Mar. Biol. 118,137 -147.[CrossRef]
Wilson, W. and Ruff, R. (1988). Species profiles: life histories and environmental requirements of coastal fishes and invertebrates (North Atlantic) - sandworms and bloodworms. In US Fish Wildlife Service Biological Report, p.23 . US Army Corps of Engineers.
Wohlgemuth, S. E., Taylor, A. C. and Grieshaber, M. K. (2000). Ventilatory and metabolic responses to hypoxia and sulphide in the lugworm Arenicola marina (L.). J. Exp. Biol. 203,3177 -3188.[Abstract]
Yancey, P. H., Blake, W. R. and Conley, J. (2002). Unusual organic osmolytes in deep-sea animals: adaptations to hydrostatic pressure and other perturbants. Comp. Biochem. Physiol. 133A,667 -676.[CrossRef]
Yin, M., Palmer, H. R., Fyfe-Johnson, A. L., Bedford, J. J., Smith, R. A. J. and Yancey, P. H. (2000). Hypotaurine, N-methyltaurine, taurine, and glycine betaine as dominant osmolytes of vestimentiferan tubeworms from hydrothermal vents and cold seeps. Physiol. Biochem. Zool. 73,629 -637.[CrossRef][Medline]