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Department of Surgery, Duke University Medical Center, Durham, NC 27710 3 Abbreviations used in this paper: SSH, suppression subtractive hybridization; CcO,cytochrome c oxidase; CcO I, CcO subunit 1; iNOS, inducible NO synthase; L-NAME, NG-nitro-L-arginine methyl ester; L-NIL, L-N-(1-iminoethyl)lysine hydrochloride; mtNOS, mitochondrial NO synthase; SNAP, S-nitroso-N-acetyl-penicillamine.
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Introduction |
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In LPS-mediated states of sepsis, inducible NO synthase (iNOS) expression and NO production regulate molecular functions that determine the host inflammatory response. NO inhibits mitochondrial respiration by nitrosylation of the iron-sulfur centers of aconitase, complex I (NADH-ubiquinone oxidoreductase), complex II (succinate-ubiquinone oxidoreductase), and complex IV (cytochrome c oxidase) (4, 5). Short term exposure to low concentrations of NO specifically and reversibly inhibits CcO activity (4). A mitochondrial heavy strand gene product, CcO, is the terminal complex of the mitochondrial respiratory chain, responsible for 90% of cellular oxygen consumption and essential for cellular energy production (6, 7, 8, 9). CcO I is considered to be the most critical of the various CcO component subunits. In this regard although the effect of NO on mitochondrial respiratory physiology has been extensively characterized, the role of NO has not been examined with respect to the transcriptional program for mitochondrial expression of proteins critical to the cellular respiration. In this study using LPS-stimulated ANA-1 murine macrophages, we demonstrate that the expression and activity of the mitochondrial protein, CcO I, are significantly decreased as the result of an NO-dependent post-transcriptional regulatory mechanism.
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Materials and Methods |
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ANA-1 macrophages (gift from Dr. G. Cox, U.S. Uniformed Health Services, Bethesda, MD) were maintained in DMEM with 10% heat-inactivated FCS, 100 U/ml penicillin, and 100 µg/ml streptomycin. LPS (0–10,000 ng/ml) was added in the absence of FCS (10%) to induce NO synthesis. In selected instances the competitive substrate inhibitor of NO synthase, L-NAME (250 ng/ml); the selective inhibitor of iNOS, L-N-(1-iminoethyl)lysine hydrochloride (L-NIL; 100 µM); the NO donor, S-nitroso-N-acetyl-penicillamine (SNAP; 100 µM); or a combination of these compounds was added. After incubation for 12 h at 37°C in 5% CO2, the supernatants and cells were harvested for assays.
Assay of NO production
NO released from cells in culture was quantified by measurement of the NO metabolite, nitrite. Cell culture medium (50 µl) was removed from culture dish and centrifuged; the supernatants were mixed with 50 µl sulfanilamide (1%) in 0.5 N HCl. After a 5-min incubation at room temperature, an equal volume of 0.02% N-(1-naphthyl)ethylenediamine was added. Following incubation for 10 min at room temperature the absorbance of samples at 540 nm was compared with that of an NaNO2 standard on a MAXLINE microplate reader (Molecular Devices, Sunnyvale, CA).
Suppression subtractive hybridization
SSH was performed using the Clontech PCR-Select cDNA Subtraction and PCR-Select Differential Screening kits. Total RNA was isolated from near-confluent cells treated with LPS or LPS plus L-NAME for a period of 12 h; poly(A)+ RNA was prepared from total RNA using a Dynabeads mRNA purification kit (Dynal Biotech, Oslo, Norway). A reverse transcribed reaction was performed with 2 µg poly(A)+ RNA, 50 mM Tris-HCl (pH 8.5), 8 mM MgCl2, 30 mM KCl,1 mM DTT, 10 µM dNTP, 10 µM primer (5-TTTTGTACAAGCTT30N1N-3), and 2 U AMV reverse transcriptase for 1.5 h at 37°C. Second-strand cDNA synthesis was performed using DNA polymerase I and T4 DNA polymerase. Double-strand cDNA was treated with a restriction enzyme, RsaI, to generate a blunt-ended, double-stranded cDNA fragment. It was then phenol-extracted and ethanol-precipitated. Adaptors (adaptor 1 or adaptor 2R; Clontech Laboratories, Palo Alto, CA) were attached in 1x ligation buffer containing 1 U/µl T4 DNA ligase (Clontech Laboratories) and incubated at 16°C for 18 h. Subtraction hybridization of cDNA was conducted by hybridizing adaptor 1-ligated cDNA and the adaptor 2R-ligated cDNA at 68°C for 8 h after denaturation at 98°C for 1.5 min. A second hybridization was performed at 68°C overnight after mixing the two primary hybridization samples and adding an excess of freshly denatured adaptor-ligated cDNA. Two hundred microliters of 20 mM HEPES-HCl (pH 8.3), 50 mM NaCl, and 0.2 mM EDTA (pH 8.0) were then added, and the resulting solution was incubated at 68°C for 7 min. Differentially expressed sequences in subtracted cDNA were subjected to PCR to amplify only cDNA with different adaptors at both ends, which were further enriched by a second round of PCR amplification with the nested primer. The desired differentially expressed sequences amplified by PCR were inserted into pT-Adv cloning vector (Clontech Laboratories). After a simple blue/white visual assay, PCR was used to rapidly amplify the cDNA insert using the following PCR primers: forward, AAA CAG CTA TGA CCA TGA; and reverse, TAA TAC GAC TCA CTA TAG GG. Each PCR product was blotted on a nylon membrane (Hybond N+; Amersham Pharmacia Biotech, Piscataway, NJ). 32P-labeled cDNA probes were synthesized as first-strand cDNA from tester and driver. Clones corresponding to differentially expressed mRNAs hybridize only with the tester probe. Following hybridization and washing, the membrane was exposed to x-ray film. Positive clones were sequenced with the sequencing primer AAA CAG CTA TGA CCA TGA, using the ABI PRISM 377 Genetic Analyzer (PE Applied Biosystems, Foster City, CA). Resulting sequences were compared with the GenBank database using the National Center for Biotechnology Information BLAST server.
Isolation of mitochondria
Mitochondria were isolated using the ApoAlert Cell Fractionation Kit (Clontech Laboratories). The purity of mitochondrial yield was determined by immunoblot analysis for the CcO IV mitochondrial protein marker.
CcO activity
CcO activity was determined using a spectrophotometric microtiter plate assay based upon the reduction of cytochrome c by CcO and the subsequent rereduction by 3,3'-diaminobenzidine-tetrachloride resulting in an oxidized 3,3'-diaminobenzidine-tetrachloride polymer detectable at 450 nm (10). The rate of formation was directly proportional to CcO activity. Results were normalized to succinate dehydrogenase activity and total cell protein.
RNA preparation and Northern blot analysis
Total RNA was isolated from ANA-1 macrophages using TRIzol reagent (Life Technologies BRL, Gaithersburg, MD). The RNA samples (10 µg/lane) were fractionated by electrophoresis on a 1% agarose formaldehyde gel and transferred to Hybond-C nylon membrane (Amersham Pharmacia Biotech). 32P-dATP-labeled probes were constructed based upon the murine CcO I mitochondrial cDNA sequence (GenBank number AF259518; nt +616 to +1216), murine CcO II sequence (GenBank number AF378830; nt +26 to +725), and the murine NADH I sequence (GenBank number NC001569; nt +2760 to +3459). cDNA probes were prepared by random primer labeling, followed by purification using a Sephadex G-50 minicolumn (BioMax, Odenton, MD). Hybridization was performed at 42°C for 18 h in ULTRAhyb hybridization buffer (Ambion, Austin, TX). Following hybridization, filters were washed twice and subjected to autoradiography using Fuji film (Fuji, Tokyo, Japan) for a period of 14 h. The mitochondrial H-strand gene for 16S rRNA was used as the housekeeping gene. Quantification was performed using a PhosphorImager (Storm 840; Molecular Dynamics, Sunnyvale, CA).
Immunoprecipitation and immunoblot analysis
Cell culture medium was removed, and plates were rinsed with PBS at room temperature. All the following steps were performed using ice-cold buffers. RIPA buffer (0.6 ml; 1x PBS, 1% Nonidet P-40, 0.5% sodium deoxycholate, 0.1% SDS, 100 µg/ml PMSF, and 60 µg/ml aprotinin) was added to a 65-mm cell culture plate. Plates were scraped, and the cells were lysed. Ten microliters of 10 mg/ml PMSF stock was added, followed by incubation for 30–60 min on ice. Whole cell lysate was precleared by adding 0.25 µg normal mouse control IgG together with protein A-agarose conjugate, and incubation was performed at 4°C for 30 min. The beads were pelleted, and the supernatant was incubated with primary Ab (monoclonal mouse CcO I, CcO II, or NADH I Ab; Molecular Probes, Eugene, OR). Resuspended protein A-agarose was added, and the tubes were incubated at 4°C on a rocker platform overnight. The pellet was collected by centrifugation at 1000 x g for 5 min at 4°C, and the supernatant was discarded. The pellet was washed with RIPA buffer multiple times and resuspended in electrophoresis sample buffer. The protein concentration was determined by absorbance at 650 nm using protein assay reagent (Bio-Rad, Hercules, CA). Cell lysate (50 µg/lane) was separated by SDS-12% PAGE, and the products were electrotransferred to polyvinylidene difluoride membrane (Amersham Pharmacia Biotech). The membrane was blocked with 5% skim milk PBS/0.05% Tween for 1 h at room temperature. After being washed three times, blocked membranes were incubated with primary mouse CcO I, CcO II, or NADH I mAb (Molecular Probes) for 1 h at room temperature, washed three times in PBS/0.05% Tween, and incubated with HRP-conjugated secondary Ab for 1 h at room temperature. After an additional three washes, bound peroxidase activity was detected by the ECL detection system (Amersham Pharmacia Biotech). Quantification was performed using densitometric analysis.
Mitochondria run-on assays
Mitochondria run-on assays were performed as previously
described (11, 12). Macrophage mitochondria were prepared
in lysis buffer (10 mM Tris-Cl (pH 7.4), 10 mM NaCl, 3 mM
MgCl2, and 0.5% Nonidet P-40) and pelleted at
500 x g. The mitochondria (2 x
107) were resuspended in 100 µl glycerol
buffer, then 150 µCi [
-32P]UTP (800
Ci/mmol) in 100 µl 10 mM Tris-Cl (pH 8.0), 5 mM DTT, 5 mM
MgCl2, 300 mM KCl, and 1 mM each of ATP, CTP, and
GTP for 30 min at 30°C were added. Labeled RNA was treated with 10 U
RNase-free DNase I (Life Technologies) for 5 min at 30°C and
extracted with phenol/chloroform (24/1) and chloroform alone. Before
ethanol precipitation, 10 µg yeast tRNA was added, and labeled RNA
was treated with 0.2 M NaOH for 10 min on ice. The solution was
neutralized by the addition of HEPES (acid free) to a final
concentration of 0.24 M. After ethanol precipitation, the RNA pellet
was resuspended in 10 mM N-Tris (hydroxymethyl)
methyl-2-aminoethanesulfonic acid (pH 7.4), 0.2% SDS, and 10 mM EDTA.
CcO I cDNA was spotted onto nylon membranes with a slot-blot apparatus
(Bio-Rad). Mitochondrial H-strand DNA and pT-Adv vector served as
positive and negative controls, respectively. Hybridization was
performed at 42°C for 48 h with 5 x
106 cpm labeled RNA in hybridization buffer (50%
formamide, 4x SSC, 0.1% SDS, 5x Denhardt’s solution, 0.1 M sodium
phosphate (pH 7.2), and 100 µg/ml salmon sperm DNA). After
hybridization, the membranes were washed twice at room temperature in
2x SSC and 0.1% SDS and three times at 56°C in 0.1x SSC and 0.1%
SDS. The membranes were exposed with an intensifying screen to Fuji
x-ray film for a period of 36 h. Quantification was performed
using a PhosphorImager (Molecular Dynamics Storm 840).
Statistical analysis
Data are expressed as the mean ± SD of three or four assays. Statistical analysis was performed using Student’s t test. A value of p < 0.05 was considered significant.
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Results |
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ANA-1 macrophage production of NO in response to a 12-h incubation with LPS (0–10 µg/ml) was determined in the presence and the absence of the competitive substrate inhibitor, L-NAME (250 ng/ml), or the specific inhibitor of iNOS, L-NIL (100 µM). The nitrite level in unstimulated control cells was 23.1 ± 6.6 nmol/mg. There was a significant concentration-dependent increase in medium levels of nitrite, the NO metabolite, in response to LPS stimulation (by ANOVA, p = 0.0001). In the presence of an LPS concentration of 100 ng/ml, nitrite production was 74 ± 11.2 nmol/mg. LPS- plus L-NAME-treated and LPS- plus L-NIL-treated cells exhibited levels of NO production that were not significantly from controls for all concentrations of LPS used. Nitrite levels of cells treated with L-NAME alone or L-NIL alone did not differ from those of unstimulated control cells (19.3 ± 5.8 and 18.4 ± 4. vs 23.1 ± 6.6 nmol/mg). In subsequent assays an LPS concentration of 50 ng/ml was used unless otherwise stated.
Ceullar CcO activity
Cellular CcO enzyme activities were normalized to succinate
dehydrogenase activity and total cell protein and determined after 1,
6, and 12 h of treatment (Table I
).
One hour following LPS stimulation, CcO activity was decreased by
approximately 50% compared with that in unstimulated control cells.
Ablation of NO synthesis in LPS- plus L-NAME-treated cells
resulted in restoration of CcO activity. Repletion of NO by addition of
SNAP to LPS- plus L-NAME-treated cells again significantly
decreased CcO activity. L-NAME or SNAP alone did not alter
CcO activity. Following 6 and 12 h of stimulation there was a
serial decline in CcO activity in LPS-treated and LPS-,
L-NAME-, plus SNAP-treated cells. At 12 h of
stimulation CcO activity was 20 and 25% of that noted in controls in
LPS-treated and LPS-, L-NAME-, plus SNAP-treated cells,
respectively. Cell viability, as measured by trypan blue exclusion, was
not significantly different among the various treatment groups
following 1, 6, and 12 h of incubation. Subsequently, 1- and 12-h
treatment groups of LPS-stimulated cells were washed, medium containing
L-NAME was added to inhibit NO production, and CcO activity
was measured after an additional 6 h. In the 1-h treatment group
CcO activity (1.6 ± 0.2/mg protein) was restored to levels noted
in unstimulated control cells. In contrast, in the 12-h treatment group
CcO activity (0.4 ± 0.1/mg protein) remained significantly
depressed and was not significantly different from that noted before
L-NAME addition. These data suggest that CcO enzyme
activity is significantly decreased by LPS-mediated NO synthesis. In
addition, depression of CcO activity is irreversible following 12-h
incubation in the setting of LPS-mediated NO synthesis.
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Cellular CcO I protein expression was determined using immunoblot
analysis (Fig. 1). CcO IV protein,
encoded by the nuclear genome, was used as a marker to normalize for
levels of total mitochondria. Under all treatment conditions, CcO IV
expression was unaltered, indicating that total mitochondrial mass and
subunits of CcO encoded by the nuclear genome did not change with the
various experimental conditions. LPS treatment decreased normalized CcO
I protein by >12-fold compared with control cells
(p < 0.01 vs control). Inhibition of NO
synthesis by addition of L-NAME with LPS restored
CcO I protein to levels not significantly different from those in
control cells. Repletion of NO in the form of SNAP to LPS- plus
L-NAME-treated cells again significantly
decreased CcO I protein. In this instance, CcO I protein was >10-fold
less than the control value (p < 0.01 vs
control). Treatment of ANA-1 cells with L-NAME or
SNAP alone did not significantly alter CcO I expression compared with
that in controls. These results indicate that LPS-mediated NO
production inhibits CcO I protein expression. NO alone is necessary,
but insufficient.
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Steady state mRNA levels of cellular CcO I were determined by
Northern blot analysis (Fig. 2).
-Actin and 16S rRNA mRNA were used as measures of constitutive
nuclear and H-strand mitochondrial gene expression. Of note, both
-actin and 16S rRNA mRNA were unaltered by treatment conditions. LPS
treatment decreased CcO I mRNA by >20-fold compared with control cells
(p < 0.01 vs control). Inhibition of NO
synthesis by addition of L-NAME with LPS restored
CcO I mRNA to levels not significantly different from those in control
cells. Repletion of NO in the form of SNAP to LPS- plus
L-NAME-treated cells again significantly
decreased CcO I mRNA. In this instance CcO I mRNA was largely
undetectable (p < 0.01 vs control). Treatment
of ANA-1 cells with L-NAME or SNAP alone did not
significantly alter CcO I mRNA expression relative to the control
value. These results indicate that LPS-mediated NO production inhibits
steady state CcO I mRNA expression. Again, NO alone is necessary, but
insufficient.
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Mitochondrial RNA polymerase is resistant to normal inhibitors of
nuclear RNA polymerase (12, 13). Eads and Hand
(12) have demonstrated that actinomycin D (100 µg/ml)
inhibits mitochondrial gene transcription by 85%, while 0°C
incubation during the assay is associated with 96% inhibition.
Therefore, CcO I mRNA half-life was determined with both actinomycin D
(100 µg/ml) and incubation at 0°C to completely inhibit
mitochondrial RNA polymerase (Fig. 3).
One hour following stimulation, actinomycin D was added. Expression of
mRNA was normalized to that of the housekeeping gene, 28S rRNA, and
that of CcO I at time zero.
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). In control cells the CcO I mRNA
half-life was 2.9 ± 0.3 h. LPS stimulation decreased the CcO
I mRNA half-life to 1.6 ± 0.4 h (p
< 0.01 vs control). Inhibition of NO synthesis by addition of
L-NAME with LPS restored the CcO I mRNA half-life
(3.2 ± 0.4 h) to a value not significantly different from
that in control cells. Repletion of NO in the form of SNAP to LPS- plus
L-NAME-treated cells again significantly
decreased the CcO I mRNA half-life to 2.0 ± 0.1 h
(p = NS vs LPS; p < 0.01 vs
control). Treatment of ANA-1 cells with L-NAME or
SNAP alone did not significantly alter the CcO I mRNA half-life
relative to that in controls. These results indicate that LPS-mediated
NO production significantly decreases the CcO I mRNA half-life,
suggesting enhanced post-transcriptional degradation. Of interest, at
1 h inhibition of NO synthesis in LPS- plus
L-NAME-treated cells resulted in a 30% increase
in steady state CcO I mRNA levels. This finding supports the converse
idea that LPS-mediated NO synthesis inhibits CcO I mRNA expression.
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In most instances the rate of transcription determines relative
differences in mRNA expression. To determine the effect of LPS-mediated
NO synthesis on mitochondrial gene transcription, mitochondrial run-on
assays were performed (Fig. 5). Whole
mitochondrial H-strand transcription was used as a positive control,
while pT-Adv vector served as a negative control. The mitochondrial
H-strand contains genes for two rRNAs, 14 tRNAs, and 12 polypeptides,
including CcO I. Because transcription for the entire H-strand is
initiated from a single promoter, the entire polycistronic H strand
transcript was chosen as the positive control. There was no statistical
difference between H-strand DNA and CcO I transcription for all
treatment groups. These results indicate that NO-induced differences in
steady state CcO I mRNA levels are not the result of altered rates of
transcription.
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The effect of NO on other proteins encoded by the mitochondrial H
strand genes is largely unknown. Certainly this study indicates that
16S rRNA expression is not changed. In the setting of LPS-mediated NO
synthesis in ANA-1 macrophages, we have previously demonstrated that
cytochrome b protein expression is inhibited in the presence
of unaltered transcription (14). Our SSH data suggest that
CcO II and NADH I expression may also be down-regulated in the presence
of NO. Therefore, immunoblot and Northern blot analyses were performed
(Fig. 6). CcO II and NADH I protein and
steady state mRNA levels were not altered under the various treatment
conditions. Additional mitochondrial run-on experiments examining CcO
II and NADH dehydrogenase I transcription demonstrated no significant
changes in transcription under the various treatment conditions (data
not shown). These data indicate that LPS-induced NO synthesis does not
uniformly decrease levels of all transcripts encoded by the
mitochondrial H-strand.
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Discussion |
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A mitochondrial heavy strand gene product, CcO, is the terminal complex of the mitochondrial respiratory chain, responsible for 90% of cellular oxygen consumption and essential for cellular energy production. The primary three subunits, CcO I, II, and III, are encoded by mitochondrial DNA and perform the catalytic functions of the holoenzyme. CcO I binds three of the enzyme’s redox centers (heme a, heme a3, and CuB) and is the most highly conserved member of the entire CcO complex. As such, it is considered to be the most critical of the 13 CcO subunits (4, 6, 7, 9). In theory, inhibition of CcO I protein expression by NO should dramatically decrease CcO holoenzyme function and inhibit mitochondrial respiration. In this regard it is known that exposure to NO for prolonged time periods or at higher concentrations results in irreversible inhibition of mitochondrial respiration. Induction of iNOS in our studies fulfills these criteria. Previous studies have found that NO damages the mitochondrial iron-sulfur centers and nitrosylates essential thiols in complex I. In addition, the NO metabolic product, peroxynitrite, may damage mitochondrial complexes I and II, ATP synthase, creatine kinase, and mitochondrial DNA and induce mitochondrial swelling and uncoupling (4, 5). The effect of NO on expression of essential protein components of the electron transport chain has not been extensively characterized in this regard. Our results suggest that iNOS-mediated irreversible inhibition of mitochondrial respiration may be the result of NO-dependent post-transcriptional augmentation of CcO I mRNA.
NO-dependent post-transcriptional acceleration of CcO I mRNA
degradation has not been previously described in the setting of LPS
stimulation. In a model of RAW 264.7 macrophages Lehrer-Graiwer and
colleagues (15) found that exposure to an exogenous source
of NO for <1 h significantly increased steady state CcO I mRNA and
protein expression. Consistent with our observations after
administration of SNAP alone, these authors found that an NO donor
alone did not significantly alter CcO I protein expression. However,
there was no change found in CcO enzyme activity, and addition of the
NO donor to the enzyme assay mixture resulted in complete inhibition of
CcO activity (15). The difference in results between our
two groups is explained by the difference in experimental models. We
used a model of LPS-induced NO production with an emphasis on prolonged
exposure, and as a consequence, our data indicate that NO is necessary,
but insufficient, by itself to inhibit CcO I protein expression.
However, in settings using only the NO donor SNAP we found no change in
CcO I mRNA or protein expression after an incubation period of 12
h. Janssen et al. (16) exposed rat epithelial cells to
spermine 2-(N, N-diethylamino)-diazenolate
2-oxide for 4 h and found that mRNA for the mitochondrial
genes NADH dehydrogenase subunits 5 and 6 was significantly decreased.
Corresponding protein levels and enzyme activity were not measured.
Again, the role of LPS-induced NO production is not addressed in this
study. Finally, in LPS- and IFN-
-stimulated astroglial and mixed
cortical cell cultures, Nicoletti et al. (17) found that
NO induced increased CcO I mRNA levels and transcription following an
incubation period of 18 h. However, cytochrome oxidase enzyme
activity was decreased. These data contrast with ours and may simply
result from differences in cell models. These results notwithstanding,
our findings suggest a unique mechanism by which LPS-mediated NO
synthesis inhibits CcO activity by enhancing mRNA degradation of CcO I,
a mitochondrial gene product critical for electron transport.
The mitochondrial genome is a closed circular dsDNA molecule of approximately 16.6 kb that is highly conserved among mammals (9). The H-strand encodes two rRNAs, 14 tRNAs, and 12 polypeptides, including CcO I. The L strand codes for eight tRNAs and a single polypeptide. All 13 polypeptide products are constituents of enzyme complexes of the oxidative phosphorylation system. Transcription and replication depend upon trans-acting nuclear-encoded factors. An H-strand promoter with two potential initiation sites controls transcription of the H strand. Once initiated, the strands are transcribed as single polycistronic precursor RNAs. Increased mitochondrial transcription has been found in rapidly dividing immortal cells requiring elevated respiratory function and anoxia-induced quiescence in Artemia franciscana embryos and in early Xenopus embryogenesis (11, 12, 13, 18). However, differential rates of transcription have not been identified among individual mitochondrial polypeptide mRNAs, indicating that differential mRNA stability accounts for varying levels of mitochondrial mRNA levels. Our run-on results with the various mitochondrial genes corroborate this finding. The mechanisms that control mitochondrial mRNA degradation are less well defined. Polyadenylation of mRNAs is thought to create functional translation stop codons that are encoded in the DNA while also enhancing degradation (9, 18). In Trypanosoma brucei, Militello and Read (18) found that mitochondrial mRNA is degraded by two biochemically distinct turnover pathways. The first pathway is dependent upon an mRNA poly(A) tail and requires exogenous UTP, while the second requires neither a poly(A) tail nor UTP. The first pathway results in rapid turnover with a mRNA t1/2 of approximately 10–20 min, while the second results in slower turnover with a t1/2 of approximately 3 h. Although the applicability of these observations to our system is unknown, the second slower pathway may be responsible. The details of the interplay of NO with this second mitochondrial mRNA degradation pathway are as yet unknown. Alternatively, exclusive of these pathways, NO may interact with the mitochondrial enzymes, poly(A) polymerase or RNase P, to enhance activity, resulting in increased mitochondrial mRNA degradation (9).
The source of the NO in our studies requires consideration. The two potential sources of NO in our system are inducible NOS and mitochondrial NO synthase (mtNOS) (19, 20). Characterization of the mtNOS isoform indicates that is similar to iNOS, but is constitutively expressed and localized to the mitochondrial membrane. It is unknown whether mtNOS expression or activity is enhanced in the presence of LPS or proinflammatory cytokines. Our attempts to induce NO production and demonstrate decreased CcO I protein expression or activity in isolated mitochondria stimulated with LPS were unsuccessful. However, these preliminary results do not rule out involvement of mtNOS in the inhibition of CcO I activity.
In summary, in a system of LPS-stimulated ANA-1 murine macrophages, CcO I protein and mRNA levels, CcO activity, and CcO I mRNA half-life were significantly decreased in the presence of an NOS inhibitor. NO is necessary, but insufficient, to alter CcO I expression. LPS-dependent signal transduction pathways are also required. This effect does not uniformly affect all mitochondrial H-strand genes. The role of NO in mitochondrial respiratory physiology has been extensively characterized in states of inflammation and sepsis. However, the role of NO has not been previously examined with respect to the mitochondrial transcriptional programs that regulate proteins critical for cellular respiration and oxidative phosphorylation. In this regard our observations suggest a novel mechanism by which NO may inhibit mitochondrial function and cellular respiration by enhancing degradation of CcO I mRNA, a critical electron transport protein.
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Footnotes |
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2 Address correspondence and reprint requests to Dr. Paul C. Kuo, Duke University Medical Center, 110 Bell Building, Box 3522, Durham, NC 27710. E-mail address: kuo00004{at}mc.duke.edu
Received for publication October 16, 2001. Accepted for publication February 26, 2002.
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