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J. Biol. Chem., Vol. 278, Issue 34, 31584-31592, August 22, 2003
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From the Department of Molecular Biology and Biotechnology, The University of Sheffield, Firth Court, Western Bank, Sheffield S10 2TN, United Kingdom
Received for publication, March 31, 2003 , and in revised form, June 3, 2003.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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More recent evidence for NO production by E. coli has come from expression of the Paracoccus denitrificans transcription factor NNR in E. coli. This protein is activated by NO, and transcription of a target melR-lacZ promoter in E. coli was attributed to formation of NO (or related species) from nitrate by molybdenum-dependent nitrite reductase (12). NO production from nitrite, however, was not dependent on molybdenum cofactor biosynthesis.
Since the initial reports of NO production by E. coli (6, 13), advances have been made that prompt a reinvestigation. First, sensitive NO electrodes with markedly improved selectivity have been developed (14). Second, several proteins have functions firmly linked to the stresses imposed by NO and nitrosation (15–17), yet the potential physiological stresses imposed by endogenously generated NO have not been assessed. The best understood is the flavohemoglobin, Hmp, of E. coli, encoded by a gene that is inducible under both aerobic and anaerobic conditions by nitrate, but especially nitrite and NO (18). NO consumption by Hmp aerobically generates nitrate (19–21), but, anaerobically, Hmp detoxifies NO by converting it to NO– with N2O appearing as a product (20, 22). The global aerobic-anaerobic regulators Fnr and MetR are involved in E. coli hmp regulation (15, 23). Third, certain bacteria (e.g. Nocardia (24)) have been shown to possess nitric-oxide synthases similar to those reported in certain plants (25) and mammalian systems, suggesting a role for NO in bacterial physiology. These findings suggest an interplay between NO-producing and NO-consuming activities of physiological relevance in non-denitrifying bacteria.
In this report, we provide direct and sensitive measurements of NO production by E. coli cells from nitrite and explore the roles of Hmp and of cytochrome bd (previously shown to react with and reduce nitrate (26, 27)) in this process. We demonstrate the involvement of the nrfA-encoded cytochrome c nitrite reductase in generating NO and the lack of NO generation in mutants lacking the flavohemoglobin Hmp, the narG-encoded nitrate reductase, or the global regulator Fnr.
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EXPERIMENTAL PROCEDURES |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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Media and Culture Conditions—Cells were grown at 37 °C
in Luria Broth (LB), pH 7.0
(32), supplemented as
appropriate with kanamycin (100 µg/ml) or ampicillin (200 µg/ml).
KNO3 was added as a filter-sterilized solution to LB at a final
concentration of 40 or 100 mM, as indicated. Aerobic cultures were
grown with shaking (200 rpm) in side-arm flasks containing 1/20th of their own
volume of medium. Anaerobic cultures (for NO evolution experiments) were grown
in screw-cap glass tubes filled to the brim and containing a glass bead
(
1-mm diameter) to aid resuspension of cells that had sedimented during
static culture. Anaerobic cultures (for cytochrome c552
assays) were grown in 500-ml Duran bottles, filled to the brim (total volume
620 ml), for 24 h to stationary phase of growth. Cells were harvested by
centrifugation and washed twice at 4 °C with phosphate-buffered saline
(PBS), pH 7.3, then suspended in PBS to a final volume of 1.0 ml. Strains
JCB352 and JCB387 were washed and resuspended in LB. A Klett-Summerson
photoelectric colorimeter (Klett Manufacturing Co., New York, NY) equipped
with a no. 66 (red) filter was used to monitor turbidity of cultures grown in
screw-cap glass tubes or side-arm flasks. Culture optical density at 420 nm
(A420) was measured using a Jenway 6100 spectrophotometer
in cells of 1-cm pathlength.
Preparation of Periplasmic and Cytoplasmic
Extracts—Periplasmic and cytoplasmic extracts were made using a
modified version of earlier protocols
(33,
34). Aerobic cultures (200 ml
in a 1-liter conical flask, 200 rpm) were grown to an A420
of 0.5–0.6 (4 h after inoculation) and then conditioned for osmotic
shock by the addition of NaCl and Tris-HCl buffer (pH 7.3) to a final
concentration of 30 mM each. Cells from 400 ml of culture were
resuspended in 6 ml of a buffer containing (final concentrations) 33
mM Tris-HCl (pH 7.3), 20% (w/v) sucrose, and 1 mM sodium
EDTA. After incubation for 20 min at room temperature, cells were collected by
centrifugation at room temperature and then rapidly resuspended in 6 ml of
ice-cold water. After mixing for 45 s on ice, Mg2+ was
added to a final concentration of 1 mM, and the suspension was
mixed on ice for 45 s, then left on ice for a further 10 min. This suspension
was centrifuged at 10,000 x g for 5 min, and the resulting
supernatant fraction (periplasm) was stored on ice prior to use. The remaining
pellet was resuspended in a buffer that contained (final concentrations) 20%
sucrose, 1 mM sodium EDTA, and 200 mM Tris-HCl (pH 7.5).
Sonication (5 x 15-s bursts with 30-s intervals) was followed by high
speed centrifugation (1.57 x 105 g for 70 min),
yielding a cytoplasmic fraction, which was also stored on ice. Purified Hmp
was obtained as described before
(21).
Enzyme and Protein Assays—Assays of 
-galactosidase and
alkaline phosphatase activities were used to determine the purity of
periplasmic and cytoplasmic samples produced. Assays were carried out at
ambient room temperature, 20 °C. -Galactosidase
(32) and alkaline phosphatase
(35,
36) activities were measured
by monitoring the hydrolysis of
o-nitrophenyl--D-galactopyranoside and
p-nitrophenyl phosphate, respectively, and measuring
A420. -Galactosidase activities are expressed in
Miller units (32). Alkaline
phosphatase activities are expressed as (
A420
x 1000)/min/ml of sample. The protein content of all samples was
measured using the protocol of Markwell et al.
(37). Each sample was assayed
a minimum of six times.
Nitrite Reductase Assay (Cytochrome
c552)—Cytochrome
c552 concentrations in periplasmic and cytoplasmic
bacterial extracts were determined spectroscopically
(38) except that an SDB-4
dual-wavelength scanning spectrophotometer was used (University of
Pennsylvania Biomedical Instrumentation Group, and Current Designs Inc.,
Philadelphia, PA) (39).
Dithionite-reduced minus persulfate-oxidized difference spectra were
computed. Spectral data were analyzed and plotted using Soft SDB (Current
Designs Inc.) and CA-Cricket Graph III software. The absorbance coefficient
(
422–410) for cytochrome c552
(reduced minus oxidized) was taken to be 61.4
mM–1 cm–1, deduced
from the data of Fujita
(40).
NO Evolution and O2 Measurements—Concentrations of dissolved oxygen and NO were measured in a Clark-type polarographic oxygen electrode system (Rank Bros., Bottisham, Cambridge, UK) modified to accommodate a World Precision Instruments ISO NOP sensor (2-mm diameter) (21). Cell suspension was diluted in the chamber with PBS buffer. Sodium formate was added, giving a working volume of 2.0 ml. A close-fitting lid, with a fine hole for injections using a Hamilton syringe, was inserted. After oxygen had reached undetectable levels, the suspension was further supplemented with NaNO2 (final concentrations stated in text) and NO evolution was measured. The O2 electrode was calibrated using air-equilibrated PBS and the addition of sodium dithionite (a few grains) to achieve anoxia. The NO electrode was calibrated using S-nitroso-N-acetyl penicillamine as described by the manufacturer. Additions of anoxic, NO-saturated solutions, NaNO2 and 10 mM carboxy-PTIO were made using Hamilton syringes. NO was generated exactly as in Poole et al. (18). A stock solution of 0.5 M NaNO2 was freshly made each day. A suba-seal (VWR International) was added to the mouth of the bottle, making an airtight seal. Nitrogen gas was bubbled through the solution for 20–30 min, making it anoxic. Carboxy-PTIO was purchased from Calbiochem. Aliquots (0.1 ml) of a 10 mM aqueous solution were stored at –20 °C for up to 1 month.
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RESULTS |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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generated 10 nmol of
NO/mg of protein and 5 mM
elicited 31 nmol NO/mg of protein.
Even at high concentrations (25
mM), NO evolution did not exceed 44 nmol of NO/mg of protein. To
verify the selectivity of the electrode response, we used the imidazolineoxyl
N-oxide, carboxy-PTIO, a stable radical compound that reacts
stoichiometrically with NO to form either nitrate or nitrite and
imidazolineoxyls in neutral solution
(41). Addition of carboxy-PTIO
to the electrode chamber rapidly decreased the electrode response in a
dose-dependent manner (Fig.
1B), confirming that cells were producing NO. However,
higher concentrations of carboxy-PTIO were required for diminution of the NO
signal than would have been anticipated from the equistoichiometric reaction
of carboxy-PTIO and the NO concentration determined by calibration of the
electrode (see, for example, trace a,
Fig. 1B). This might
be due to loss of carboxy-PTIO owing to the lipophilicity and incorporation
into bacterial membranes. Addition to the reaction chamber of deoxymyoglobin
also quenched the NO signal (not shown). Cells cultured aerobically in the
presence or absence of 100 mM KNO3 failed to produce NO
(results not shown).
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Evolution of NO from Nitrite Reductase Mutants—Previous work on NO evolution by E. coli has suggested that a nitrate reductase is responsible for the production of NO. Ji and Hollocher (6, 11, 13) concluded that NO production by E. coli is due to the reduction of nitrite to NO as a secondary activity of the membrane-bound respiratory nitrate reductase. Nitrosation reactions have also been genetically and biochemically linked to nitrate reductase genes (42–44). Smith (45) and Ralt et al. (44) correlated nitrous oxide production with nitrate reductase activity in E. coli. However, Hutchings et al. (12) showed that the nitrate reductases of E. coli (all of which are molybdoenzymes) are not directly responsible for NO production from nitrite. Activation of NNR (an NO-responsive transcription factor of P. denitrificans) was abolished in an E. coli mobAB::Km mutant, defective in Mo cofactor biosynthesis, when nitrate, but not nitrite, was provided. The ability of certain nitrite reductases to reduce nitrite to NO (3) makes them strong candidates for the NO evolution observed here and by Hutchings et al. (12), particularly because E. coli lacks NO synthase.
E. coli possesses two nitrite reductases, namely a periplasmic
cytochrome c enzyme (Nrf) that generates predominantly ammonium ion,
and a cytoplasmic siroheme-dependent reductase (Nir)
(46,
47). Therefore, we compared
the NO evolution capacity of nir, nir nrf, and wild-type strains.
Both the nir mutant JCB387 and the wild-type AN2342 produced NO, and
use of carboxy-PTIO confirmed the presence of NO in both cases
(Fig. 2A). The level
of NO accumulated by the nir mutant (6.6 nmol/mg of cell protein,
mean of two measurements) was approximately a third of that produced (17
nmol/mg of protein) by the wild-type strain. However, the nir nrf
double mutant produced no detectable NO
(Fig. 2A). The
nrfA gene, implicated in NO production by this result, encodes a
52-kDa periplasmic pentaheme cytochrome c552, which acts
as a nitrite reductase in association with the NrfB cytochrome c as
redox partner. Therefore, we tested for the presence of cytochrome c
in periplasmic and cytoplasmic extracts of the above strains. Dithionite
reduced minus persulfate-oxidized spectra
(Fig. 2B) revealed the
presence of cytochrome c552 with a 
-peak at 422 nm
in both the wild-type strain AN2342 and the nir mutant JCB387 (0.10
and 0.11 nmol of cytochrome c552 per mg of protein,
respectively). Weaker signals at 552 nm are due to the 
-band but were
not used for quantification. The NrfA protein has absorbance maxima in the
reduced state (absolute spectrum) at 420.5 (), 523.5 (), and 552
nm () (48). Purified
NrfB protein, in its reduced state, has a sharp absorbance peak at 551 nm.
Unlike NrfA, however, NrfB has no charge transfer band at 630
nm.2 However,
cytochrome c552 was undetectable in the nir nrf
double mutant. As expected, cytoplasmic fractions (not shown) from all three
strains showed no detectable levels of cytochrome c552 but
the presence of b-type cytochrome(s) (not shown).
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Effects of an hmp Mutation on NO Production—Hmp plays a key
role in protecting cells against nitrosative stress by catalysis of either the
oxygen-dependent formation of nitrate ion or the anaerobic reduction of NO to
nitrous oxide (15). We
hypothesized that an hmp mutant would produce significantly higher
levels of NO compared with an isogenic wild-type strain, due to an inability
to detoxify any NO formed. However, the results obtained were quite the
opposite, as shown in Fig.
2C. The wild-type strain VJS676 produced NO (35 nmol of
NO/mg of protein) at levels higher than wild-type strain AN2342 (23 nmol of
NO/mg of protein; both means of two values). The hmp mutant strain
RKP4545 produced no detectable NO, regardless of growth conditions
(0–100 mM 
in the
medium) and the concentration of NaNO2 (in the range 2.5–25
mM) added to the electrode chamber.
Given the demonstration (Fig.
2B) that nrf mutants lacking periplasmic
cytochrome c552 were unable to generate NO from nitrite,
we assayed cytochrome c552 in periplasmic and cytoplasmic
extracts of wild-type and hmp mutant strains. Dithionite reduced
minus persulfate-oxidized spectra
(Fig. 2D) revealed the
presence of cytochrome c552 with peaks at 422 and 552 nm
in the periplasm of the wild-type strain AN2342 (0.15 nmol of cytochrome
c552 per mg of protein). The somewhat higher levels of
cytochrome c in this strain correlate with higher levels of NO
evolution (see above). However, cytochrome c552 was
undetectable in the -region of reduced minus oxidized spectra
of the periplasm of the hmp mutant strain RKP4545, but a weak
-spectral signal at 422 nm was observed. As expected, cytoplasmic
fractions (not shown) from both strains showed no detectable levels of
cytochrome c552, but one or more b-type
cytochromes were present. An Hmp-overproducing strain (RKP4717) also produced
NO under these experimental conditions, but quadruplicate assays failed to
show any significant differences from wild-type levels.
One explanation of the failure of an hmp mutant to generate NO
might be that the flavohemoglobin Hmp itself is able to reduce nitrite ion to
NO; indeed, Hmp possesses a broad spectrum of reductase activities
(49). However, purified Hmp
protein was unable to produce NO anoxically from nitrite. Briefly, 730 ng of
purified Hmp protein was added to the electrode chamber, with 1.5 ml of PBS
(pH 7.0). Oxygen levels were reduced to zero following addition of NADH, then
25 mM was added. In
duplicate experiments, no NO evolution was observed (not shown).
Effects of an fnr Mutation on NO Production—The absence of NrfA or NrfB cytochromes in the hmp mutant prompted study of the link between Hmp, NO metabolism, and nitrite reductase. Recently we have demonstrated that the O2-responsive regulator Fnr, which represses hmp gene transcription, also senses NO (23). The [4Fe-4S]2+ cluster of Fnr reacts anaerobically with, and is inactivated by, NO. Active Fnr is a positive activator of the nrf operon, nirB, narG, and other genes involved in anaerobic nitrite and nitrate dissimilation (46). We therefore hypothesized that, because a mutant lacking Hmp is unable to detoxify NO, NO will inactivate Fnr, leading to a failure to up-regulate nrf (and other Fnr-regulated genes). If NrfA/NrfB are largely responsible for NO generation from nitrite, an fnr mutant should fail to produce NO. This hypothesis is supported by the results in Fig. 3A. Both wild-type strains used here showed similar levels of NO evolution (17 and 14 nmol of NO/mg of protein for strains AN2342 and RKP2178, respectively), whereas the fnr mutant (VJS5369) did not produce NO. The use of carboxy-PTIO again confirmed the presence of NO.
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Mutational and NO Effects on Fnr Are Reflected in Anaerobic Growth
Patterns—Anaerobic growth curves of the fnr mutant strain
(VJS5369) and its corresponding wild-type strain (RKP2178) are shown in
Fig. 3B. In the
absence of added nitrate, growth of the fnr+ strain
(RKP2178) proceeded rapidly after inoculation but after 10 h reached a
saturation density of about 35 Klett units. Inclusion of nitrate substantially
stimulated the growth rate in the first 10 h and then allowed growth to a
significantly higher final cell yield (58 Klett units). In contrast, the
fnr mutant grew more slowly than the wild-type in the first 10 h
irrespective of the presence of nitrate. The inclusion of nitrate allowed a
slower subsequent phase of growth, allowing the culture after 50 h to reach a
final population density of about 56 Klett units.
To test further the hypothesis that an hmp mutant is defective in fnr-regulated gene function, anaerobic growth curves of wild-type (VJS676) and hmp mutant (RKP4545) strains were compared. Fig. 3C shows that the hmp mutant grew anaerobically in the absence of nitrate in a similar fashion to the wild-type. However, nitrate markedly stimulated growth of the wild-type strain but not of the hmp mutant. Wild-type strains VJS676 and AN2342 grew anaerobically in a similar fashion (results not shown). These growth data demonstrate that the fnr mutation adversely affects anaerobic growth and particularly anaerobic growth on nitrate. The data also reveal a previously undescribed phenotype of an hmp mutant, namely the inability of nitrate to stimulate anaerobic growth.
Fnr* Restores NO Evolution in an hmp Mutant—To further investigate the relationship between the E. coli flavohemoglobin and Fnr, an hmp strain carrying fnr* on a plasmid (50, 51) was constructed (RKP2825). It was hypothesized that this hmp fnr* strain would evolve NO, because the dimeric Fnr* protein is insensitive to NO inactivation even though the strain lacks a functional flavohemoglobin with which to detoxify NO. Under anoxic conditions, nitrate enhanced growth of the hmp fnr* strain to levels similar to the wild-type strain in the absence of nitrate (Fig. 4A). As observed previously, growth of the wild-type strain was significantly enhanced in the presence of nitrate, and the hmp mutant strain grew poorly regardless of the presence or absence of nitrate. The hmp fnr* strain did produce NO (Fig. 4B), although at lower levels than a wild-type strain. The hmp mutant strain failed to produce NO, as observed previously.
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Is NO Produced in a narG Mutant Strain?—In an attempt to
clarify the role of the membrane-bound nitrate reductase in the production of
NO, we tested the NO-producing ability of a narG mutant. The
narG gene encodes the -subunit of the membrane-associated
respiratory nitrate reductase. NO was not produced by the narG mutant
strain VJS789 when grown anaerobically in the presence of nitrate but was
produced by wild-type strain AN2342 (results not shown). In the absence of
nitrate in the growth medium, both the wild-type and the narG mutant
strain failed to produce NO (results not shown). In view of the recent
demonstration that nitrate reductase is not required for NNR activation
(12), it is possible that the
failure to detect NO evolution from the narG mutant is due to the
lack of nitrite formed from Nar activity during growth.
NO Production by a cydD Mutant—Cytochrome bd is a
terminal quinol oxidase (46),
which appears to have additional physiological roles that are not readily
explicable by its oxidase function
(52). Hubbard et al.
(26) observed a decrease in
the absorbance maximum (630 nm) of reduced cytochrome d in membrane
particles upon the addition of nitrate ions. Nitrite, trioxodinitrate, and NO
also caused qualitatively similar, but faster, changes in the spectrum of
cytochrome d. Nitrate gave a slower reaction rate, possibly due to a
rate-determining reduction of nitrate to nitrite catalyzed by a nitrate
reductase. It was concluded
(26,
27) that this spectral shift
was due to the presence of a cytochrome d-nitrosyl complex. Haddock
et al. (53) also
observed a shift in the spectrum of cytochrome d following anaerobic
growth on nitrate. The above evidence led us to investigate the role of
cytochrome bd in NO evolution in E. coli.
Fig. 5 shows that NO is evolved
by a cydD mutant at levels (11 and 28 nmol of NO/mg of protein at
12.5 and 25 mM ,
respectively) approximately one-third less than its wild-type. Thus cytochrome
bd is not essential for NO production.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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-dependent NO evolution in E.
coli although at lower levels than those measured before
(13). These differences could
be due to the different strains used or techniques employed in the measurement
of NO.
The mechanisms of NO formation during denitrification are reasonably well
understood as a result of high resolution x-ray structures of both classes of
nitrite reductases in which the substrate
() or product (NO) are bound at the
active site (see (5)). In
E. coli however, both nitrite reductases were thought to be
assimilatory, producing ammonium ion, not NO, until now
(47).
In our work, NO generation was not detectable in an nrf mutant, nor in nrf+ strains in which the nrf-encoded nitrite reductase is not expressed or synthesized at spectrally undetectable levels, as a consequence of hmp or fnr mutations. Expression of the NrfA and NirB operons is elevated during anaerobic growth by Fnr (29, 54). Wang and Gunsalus (55) concluded that nrfA operon expression is induced only when nitrite concentrations are low, whereas NirB seems to be optimally synthesized only when nitrate or nitrite are in excess of the cell's capacity to consume them.
The physiological significance of NO generation by Nrf remains to be
determined. However, the present mutant-based analysis suggests that other
NO-generating mechanisms that have been proposed are insignificant when cells
are grown anaerobically with . These
include the chemical reduction of nitrite by ferrous ion
(56) or by formate
(57). Zumft
(58) suggested that
non-enzymatic transformations could be responsible for NO formation in
non-denitrifiers, as demonstrated in humans; inorganic nitrite is chemically
reduced to NO under acidic, reducing conditions
(59).
"Chemodenitrification" has also been suggested as an NO-producing
mechanism (60), involving the
decomposition of hydroxylamine into NO and N2O
(58). However, the reducing,
acidic conditions required were not met in our electrode chamber
experiments.
An hmp mutation prevented NO formation, presumably a consequence
of the failure of Hmp to remove the NO that accumulates during growth with
nitrate. However, under the anoxic growth conditions employed here, it is
improbable that the "oxygenase"
(19) or denitrosylase
(61) activity of Hmp in which
NO is converted to nitrate could operate, particularly because the apparent
Km of this reaction for O2 is
relatively high (50–90 µM)
(16,
21). Recent work by Gardner
and Gardner (62) suggests
that, although Hmp is highly efficient as an oxygen-dependent NO-detoxifying
enzyme, its role in anaerobic NO metabolism and detoxification is minimal and
a flavorubredoxin is proposed to possess NO-scavenging activity during
anaerobic growth. However, it should be noted that, although hmp
transcription is not significantly up-regulated anaerobically compared with
aerobically, the presence of nitrate, and especially nitrite and NO, which
will be formed under the growth conditions used here, dramatically up-regulate
hmp transcription
(18). This increase may
provide sufficient NO–-detoxifying activity to protect Fnr
from inactivation. Nrf is periplasmically located, so that NO damage to Fnr
and intracellular targets will require facile NO diffusion to the cytoplasm.
Thus, membrane-associated NO-detoxifying activity would be particularly
effective in NO removal, and there is evidence for the presence of Hmp
apoprotein in the E. coli periplasm
(63).
A model of the fates of nitric oxide in E. coli, is shown in Fig. 6. Fnr senses pathophysiological levels of NO (5–10 µM) in vivo, and NO inactivates Fnr (23). Thus, one consequence of NO accumulation within E. coli would be loss of a major mechanism (via Fnr) for up-regulation of nitrate and nitrite reductases. During anaerobic growth, the potentially harmful accumulation of NO from the combined activities of nitrate and nitrite reductases is prevented by the combined NO-detoxifying activities of Hmp and flavorubredoxin, both of which are up-regulated under such conditions (18, 62). However, mutation of hmp alone appears sufficient for NO accumulation, inactivation of Fnr, and consequent down-regulation of nrf, nar, and other genes required for anaerobic growth. Failure to form nitrate reductase is shown in the present work by the lack of growth stimulation of an hmp mutant by nitrate (Fig. 3C), whereas loss of periplasmic nitrite reductase in an hmp mutant is reflected in the spectral analysis. Although lacking a functional flavohemoglobin, the hmp fnr* strain still generated NO (Fig. 4B), demonstrating the role of Fnr inactivation. Strain RKP4545 (hmp) displayed poorer anaerobic growth in the presence of nitrate (Fig. 3C) than did the fnr mutant (Fig. 3B). Although Fnr inactivation has global effects on anaerobic growth capability, the consequences of mutation of hmp appear more severe, because of failure to detoxify NO that arises from Fnr-independent routes and multiple sites of NO toxicity.
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Although the reaction of nitrite with flavohemoglobin might be anticipated to be similar to the reaction of nitrite/nitrous acid with human deoxyhemoglobin (64), we could find no evidence that purified Hmp forms NO from nitrite under anoxic conditions. Nitrate reductase-dependent mutagenesis in E. coli was observed in LB medium containing no added nitrate or nitrite (65). The requirement for hypoxia for maximum mutagenesis by nitrite suggests that hypoxia may induce an enzyme that generates NO from nitrite as proposed here for nitrite reductase(s).
Finally, the formation of free NO in cells growing under conditions of
respiration has major implications
for energy metabolism in facultative bacteria. Because NO is a potent
inhibitor of both terminal oxidases in E. coli
(66), formation by anaerobic
respiratory enzymes may inhibit oxidase activity, thereby augmenting the
shut-down of aerobic metabolism activated primarily by regulation of gene
expression.
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FOOTNOTES |
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To whom correspondence should be addressed. Tel.: 44-114-222-4447; Fax:
44-114-272-8697; E-mail:
r.poole{at}sheffield.ac.uk.
1 The abbreviations used are: NO, nitric oxide; carboxy-PTIO,
2-(4-carboxyphenyl)-4,4,5,5-tetramethylimidazoline-1-oxyl-3-oxide; DAN,
2,3-diaminonaphthalene; PBS, phosphate-buffered saline; Nrf, periplasmic
cytochrome c nitrite reductase; Nir, cytoplasmic siroheme-dependent
nitrite reductase; NNR, nitrite and nitric-oxide reductase regulator.
2 D. Richardson, personal communication.
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ACKNOWLEDGMENTS |
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REFERENCES |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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ven, R., Vollmer, M., and Bock, E. (1992)
Arch. Microbiol. 158,
439–443
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