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Applied and Environmental Microbiology, October 2001, p. 4694-4700, Vol. 67, No. 10
Department of Plant Science, University of
Manitoba, Winnipeg, Manitoba R3T 2N2, Canada
Received 5 June 2001/Accepted 31 July 2001
Gluconacetobacter diazotrophicus is an
N2-fixing endophyte isolated from sugarcane. G.
diazotrophicus was grown on solid medium at atmospheric partial
O2 pressures (pO2) of 10, 20, and 30 kPa for 5 to 6 days. Using a flowthrough gas exchange system, nitrogenase activity and respiration rate were then measured at a range of atmospheric pO2 (5 to 60 kPa). Nitrogenase activity was
measured by H2 evolution in N2-O2
and in Ar-O2, and respiration rate was measured by
CO2 evolution in N2-O2. To validate
the use of H2 production as an assay for nitrogenase
activity, a non-N2-fixing (Nif Gluconacetobacter
diazotrophicus (47) (previously known as
Acetobacter diazotrophicus [15]) is a strict
aerobe and an N2-fixing endophyte originally
isolated from sugarcane roots and stems (6). It has been
estimated that G. diazotrophicus can fix up to 150 kg of N
ha Aerobic endophytic diazotrophs require a high flux of
O2 to their respiratory systems to enable an
adequate supply of reductant and ATP to support
N2 fixation (e.g., see reference
13), yet paradoxically, an excessive flux of
O2 to the bacterium can result in an inhibition
of nitrogenase activity (14, 21, 26). The inhibition of
nitrogenase activity by O2 in aerobic diazotrophs can be reversible or irreversible, depending on the organism and the
nature (i.e., duration and severity) of the increase in
O2 flux (33, 37, 39). Reversible
inhibition of nitrogenase activity (i.e., a temporary "switch-off"
of the nitrogenase activity while O2 flux is
excessive) can be due to a conformational change in nitrogenase, as
seen in Azotobacter (11, 32), to an
ADP-ribosylation of dinitrogenase reductase, as seen in the purple
nonsulfur bacteria (46) and Azospirillum
(49), or to a diversion of electrons from nitrogenase to
other reduction pathways, as proposed for Azotobacter
(16, 29).
G. diazotrophicus has the ability to fix
N2 at ambient atmospheric partial
O2 pressures (pO2) (i.e.,
approximately 20 kPa of O2) in semisolid medium
(6) and as colonies on solid medium (10). The
ability to fix N2 in colonies on solid medium is
especially interesting, as there is evidence that G. diazotrophicus exists in situ in the intercellular spaces of
sugarcane vascular tissue as mucoid microcolonies (9).
Dong (8) also reported that colony morphology on solid
medium and the relative distribution of the bacteria within these
highly mucilaginous colony changed with changes in the partial pressure
of O2 surrounding the colonies.
Reis and Döbereiner (40) measured nitrogenase
activity in liquid cultures of G. diazotrophicus by
acetylene reduction in closed batch assays and found that activity was
maximal when the culture was at equilibrium with 0.2 kPa of
O2 in the gas phase. However, nitrogenase
activity of G. diazotrophicus grown in colonies on solid
medium in response to changes in atmospheric pO2
has not yet been well characterized. Given that G. diazotrophicus exists in situ as microcolonies adhering to plant
cell walls (9), characterization of the response of the
bacterium on solid medium to changes in atmospheric
pO2 is particularly relevant.
The objective of our study was to characterize the effect of
atmospheric pO2 on nitrogenase activity of
G. diazotrophicus grown on solid medium using flowthrough
gas exchange measurements. Treatments included long-term growth of the
bacterium on a range of atmospheric pO2 (10 to 30 kPa) and subsequent rapid changes in atmospheric
pO2 in small (5- to 10-kPa) and large (40-kPa) steps. We found that nitrogenase activity by G. diazotrophicus is adaptive to both short-term and long-term
changes in atmospheric pO2 and that the bacterium
has a switch-off/switch-on mechanism for protection of nitrogenase from
rapid changes in atmospheric pO2.
Organism and culture.
G. diazotrophicus PAL-5
(ATCC 49037; obtained from the American Type Culture Collection,
Manassas, Va.) was cultured for 2 days at 30°C, shaken at 150 rpm in
LGI-P liquid medium (M. McCulley [Carleton University], personal
communication), a modified version of LGI medium (6).
LGI-P medium differs from the original LGI medium in containing
0.02 g of Na2MoO4
· 2H2O liter
0099-2240/01/$04.00+0 DOI: 10.1128/AEM.67.10.4694-4700.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
Response of the Endophytic Diazotroph
Gluconacetobacter diazotrophicus on Solid Media to
Changes in Atmospheric Partial O2 Pressure
ABSTRACT
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Abstract
Introduction
Materials and Methods
Results
Discussion
References
) mutant of
G. diazotrophicus was tested and found to have a low rate of uptake hydrogenase (Hup+) activity
(0.016 ± 0.009 µmol of H2 1010
cells1 h1) when incubated in an atmosphere
enriched in H2. However, Hup+ activity was not
detectable under the normal assay conditions used in our experiments.
G. diazotrophicus fixed nitrogen at all atmospheric
pO2 tested. However, when the assay atmospheric
pO2 was below the level at which the colonies had been
grown, nitrogenase activity was decreased. Optimal atmospheric
pO2 for nitrogenase activity was 0 to 20 kPa above the
pO2 at which the bacteria had been grown. As atmospheric
pO2 was increased in 10-kPa steps to the highest levels (40 to 60 kPa), nitrogenase activity decreased in a stepwise manner.
Despite the decrease in nitrogenase activity as atmospheric
pO2 was increased, respiration rate increased marginally. A
large single-step increase in atmospheric pO2 from 20 to 60 kPa caused a rapid 84% decrease in nitrogenase activity. However, upon
returning to 20 kPa of O2, 80% of nitrogenase activity was recovered within 10 min, indicating a "switch-off/switch-on"
O2 protection mechanism of nitrogenase activity. Our study
demonstrates that colonies of G. diazotrophicus can fix
N2 at a wide range of atmospheric pO2 and can
adapt to maintain nitrogenase activity in response to both long-term
and short-term changes in atmospheric pO2.
INTRODUCTION
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Abstract
Introduction
Materials and Methods
Results
Discussion
References
1 year1 in sugarcane
(2). Such high levels of N2 fixation
have not been reported in any other system outside legume-rhizobium
symbioses. The bacterium has subsequently been isolated from sweet
potato (38), coffee (23), pineapple
(44), sorghum (22), finger millet
(31), and several other tropical grass species
(24).
MATERIALS AND METHODS
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Abstract
Introduction
Materials and Methods
Results
Discussion
References
1, 0.1 mg
of biotin liter1, 0.2 mg of pyridaxol HCl
liter1, and 5 ml of sugarcane juice (pressed
from fresh sugarcane stem) liter1, and the
final pH was adjusted to 5.5 using 1% acetic acid. Diluted cells were
spread on solid LGI-P agar medium (15 g of agar
liter1 plus 50 mg of yeast extract
liter1). Before serial dilution, 5 ml of
culture was vortexed with glass beads to prevent clumping of the
colonies and to obtain an even distribution of individual separate
colonies on the petri plates.
Cell enumeration. The numbers of viable cells per colony of all petri plate cultures used in gas exchange measurements were determined by plate counting. Five colonies per plate were cut out of the agar, and as much agar subtending the colonies as possible was removed without disturbing the integrity of the colonies. These five colonies were then vortexed together in a small test tube containing 5 ml of 0.85% NaCl solution and glass beads. This suspension was then serially diluted in 0.85% NaCl solution and plated. Cell number per plate was calculated by multiplying the colony number per plate by the cell number per colony. On average, colonies contained 107 to 108 viable cells each and 8.5-cm-diameter petri plates contained 100 to 150 colonies each. No differences were noted in these growth parameters for cultures grown at 10, 20, or 30 kPa of O2.
Gas exchange measurements of respiration rate and nitrogenase and hydrogenase activities. A flowthrough gas exchange system (45) was used to measure the effects of changing atmospheric pO2 on respiration rate and nitrogenase activity of G. diazotrophicus colonies. The system includes computer-controlled mass flow controllers (MKS Instruments Inc., Nepean, Canada) for gas mixing and delivery, an infrared gas CO2 analyzer (ADC-225MKS; Analytical Development Co. Ltd., Hoddesdon, United Kingdom) for measurement of respiration rate, and an H2 analyzer (27) for measurement of nitrogenase activity. A chamber with inner dimensions of 50 by 20 by 5 cm with four shelves for holding up to 40 petri plates was constructed from 0.9-cm-thick acrylic sheeting. The void volume when the chamber was fully loaded with cultures was 1.86 liters. Gas was introduced at one end of the chamber, flowed horizontally across the petri plates, and exited at the opposite end of the chamber.
Measurements of nitrogenase activity and respiration rate by G. diazotrophicus colonies were taken at various atmospheric pO2. Gas mixtures fed into the assay chamber were composed of various partial pressures of O2 in N2 (N2-O2) or in Ar (Ar-O2). Respiration rate was measured as the rate of CO2 evolved from the colonies with N2-O2 as the input gas. Nitrogenase activity was measured as H2 evolution in N2-O2 and in Ar-O2 (21, 26). Gas exchange measurements were made in the following manner. Twenty to 40 petri plates containing 5- or 6-day-old cultures of G. diazotrophicus on solid LGI-P medium were sealed in the chamber and then connected to the gas exchange system. Gas mixtures were passed through the chamber at a rate of 500 ml min1.
For testing the response of G. diazotrophicus to small (5- or 10-kPa) changes in atmospheric pO2, gas
exchange measurements were initiated at the pO2
under which the cultures had been grown (i.e., 10, 20, or 30 kPa of
O2). Initially, H2 and
CO2 evolution was quantified at this
pO2 in a gas mixture of
N2-O2. Once these readings
had been made, the input gas was switched to
Ar-O2 at the same pO2.
Normally, 30 min to 1 h was required before the H2 evolution came to steady state, and the
H2 evolution rate was always measured 1 h
after the switch from N2-O2
to Ar-O2. After the H2
evolution rate had been measured in Ar-O2, the
input stream was change back to
N2-O2, but at a new
atmospheric pO2 (±5 or 10 kPa from the previous
reading). Upon returning to an atmosphere of
N2-O2, 10 to 30 min was
required for H2 and CO2
evolution rates to come to steady state. This cycle was then repeated
until nitrogenase activity and respiration rate had been measured at
all atmospheric pO2 (stepping down from initial
level of atmospheric pO2 to the lowest levels
tested and then stepping up to the highest levels tested). To observe
the temporal response of G. diazotrophicus to large
(40-kPa), single-step increases and decreases in atmospheric pO2, gas exchange measurements were initiated at
the atmospheric pO2 under which the bacteria had
been grown (approximately 20 kPa), and then atmospheric
pO2 was increased in a single step to 60 kPa,
where it remained for approximately 15 min before being returned to 20 kPa in a single step. All gas exchange measurements were made at room
temperature (22 ± 1°C). Preliminary experiments showed that
once steady-state rates of H2 and
CO2 evolution were reached, they remained steady
for many hours (i.e., up to 12 h).
Production of H2 is an obligate reaction of the
nitrogenase enzyme complex during the fixation of
N2 (4). The rate of
H2 evolution in
N2-O2 is a measure of
partial or "apparent" nitrogenase activity (i.e., proton reduction
to H2 by nitrogenase in the presence of
N2 fixation) (21, 26). The rate of
H2 evolution in Ar-O2 is a
measure of total nitrogenase activity (i.e., in the absence of
N2 as a substrate, total electron flow through
nitrogenase is used to reduce protons to H2)
(20, 21, 26). In
N2-O2, the proportion of
total electron flow through nitrogenase being directed to
N2 fixation is known as the electron allocation
coefficient (EAC) (12) and is calculated as 1 
(H2 evolution in
N2-O2 
H2
evolution in Ar-O2). EAC can be viewed as
a measure of an aspect of nitrogenase "efficiency" (i.e., the
higher the EAC, the greater the proportion of electrons going to fix
N2 and the lower the proportion of electrons
going to the "wasteful" process of proton reduction).
The accuracy of measuring nitrogenase activity by
H2 evolution is dependent upon a lack of
hydrogenase activity leading to either H2
production or consumption by the test organism under the assay
conditions. Experiments with a non-N2-fixing
(Nif) mutant of G. diazotrophicus
(strain MAD3A) (42) were performed to determine if
H2 evolution from G. diazotrophicus
was associated only with nitrogenase activity and if the bacterium had
hydrogenase uptake (Hup+) activity. The
Nif mutant was designed by insertional
mutagenesis of the nifD gene of wild-type G. diazotrophicus. The resulting mutant strain (MAD3A) was generously
donated by C. Kennedy, University of Arizona. G. diazotrophicus MAD3A was grown and handled as described above for
wild-type G. diazotrophicus PAL5 except that 200 µg of
kanamycin ml1 was added to the medium. The
growth rate of the Nif mutant was not
significantly different from that of wild-type G. diazotrophicus for the first 5 days of culture.
Three replicates of 40 plates each of G. diazotrophicus
MAD3A were tested for H2 production in air and
Ar-O2 (80:20) in preliminary experiments in our
gas exchange system. MAD3A did not produce H2
production under any conditions (data not shown).
Hydrogenase uptake activity by G. diazotrophicus was
assessed in flowthrough and closed-assay systems. For the flowthrough assay, MAD3A was grown for 4 days at ambient pO2
(approximately 20 kPa) as described above. Three replicates of 20 petri
plate cultures were then placed in the gas exchange chamber and flushed continuously with air containing 2 ppm (vol/vol) of
H2 at a flow rate of 20 ml
min1 for approximately 24 h. This
concentration of H2 was used because it is the
typical level of H2 evolution from wild-type
G. diazotrophicus in air under our normal assay conditions.
After exposure to 2 ppm of H2 for 24 h, the
gas flow rate was increased to 500 ml min1 (the
normal flow rate for our assays) and the concentration of H2 exiting the chamber was measured. For the
closed assay, wild-type and Nif mutant strains
of G. diazotrophicus were grown on solid medium for 4 days.
On the fifth day, 10 petri plate cultures were placed in the gas
exchange chamber and the chamber was flushed with 50 ppm of
H2 in air at flow rate of 20 ml
min1. After 24 h at this flow rate, the
chamber was sealed, and evolution (wild-type strain) and consumption
(Nif strain) were monitored immediately after
sealing and then every 30 to 60 min for the next 6 to 8 h. Gas
samples (1 ml) were taken from the chamber and injected into an air
stream entering the H2 analyzer at a flow rate of
300 ml min1 for analysis as described by
Layzell et al. (27). Hydrogen consumption and evolution
rates were calculated by linear regression. The tests were replicated
four times each for the wild-type and Nif
strains of G. diazotrophicus.
The aerobic, facultative chemoautotroph Ralstonia eutropha
(ATCC 17699) was used as a positive control in the assessment of Hup+ activity. Early-log-phase cells grown in
Difco 0003 liquid medium (Becton Dickinson, Franklin Lakes, N.J.) were
plated onto Difco 0001 solid medium and grown for 4 days before being
assayed. H2 consumption by these colonies was
assay as described above for G. diazotrophicus MAD3A (i.e.,
closed assays at 50 ppm of H2 for 8 h).
Four experiments were conducted to test the response of G. diazotrophicus to changes in atmospheric
pO2. The experiments consisted of (i) testing
responses of G. diazotrophicus grown at 20 kPa of
O2 to small (5- or 10-kPa) stepped changes in
atmospheric pO2; (ii) testing responses of
G. diazotrophicus grown at 20 kPa of O2 to a large (40-kPa) stepped change in
atmospheric pO2; (iii) testing responses of
G. diazotrophicus grown at 10 kPa of
O2 to small (5- or 10-kPa) stepped changes in
atmospheric pO2; and (iv) testing responses of
G. diazotrophicus grown at 30 kPa of
O2 to small (10-kPa) stepped changes in
atmospheric pO2. Gas exchange measurements in a
single chamber containing 20 to 40 petri plates of G. diazotrophicus cultures was considered a single replicate. For
each experiment, gas exchange measurements were replicated four times.
All data were normalized by calculating gas evolution per cell (cell
number was determined for each replication of each experiment; see
enumeration method above). Data were analyzed using the general
linear model of the SAS statistical package (SAS Institute,
Cary, N.C.), assuming a completely randomized design, and mean
separation was tested using the least-significant-difference procedure (P = 0.95).
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RESULTS |
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Abstract
Introduction
Materials and Methods
Results
Discussion
References
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Effects of small stepped changes in atmospheric pO2 on
G. diazotrophicus grown at 20 kPa of
O2.
For G. diazotrophicus grown at 20 kPa
of O2, 10-kPa stepped increases in atmospheric
pO2 above 30 kPa of O2
resulted in a decrease in total nitrogenase activity
(H2 evolution in Ar-O2) (Fig. 1). However, nitrogenase was still
active even at atmospheric pO2 of 60 kPa (29% of
the rate at 20 kPa of O2). Stepped decreases in
atmospheric pO2 from 20 to 10 to 5 kPa also
resulted in decreases in total nitrogenase activity. The optimal
atmospheric pO2 for G. diazotrophicus
grown at 20 kPa of O2 were 20 and 30 kPa of O2.
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0.05.
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Effects of a large (40-kPa) stepped change in atmospheric
pO2 on G. diazotrophicus grown at 20 kPa of
O2.
When G. diazotrophicus colonies grown
at atmospheric pO2 of 20 kPa were exposed to a
40-kPa single-step increase in atmospheric pO2,
nitrogenase activity decreased rapidly and severely (Fig. 4). After this decrease, nitrogenase
activity at 60 kPa of O2 was steady at
approximately 26% of the activity at 20 kPa of
O2. After 15 min at 60 kPa of
O2, oxygen concentration was then switched back
to 20 kPa, and nitrogenase activity increased almost immediately. Within 10 min of returning to 20 kPa of O2,
nitrogenase activity by G. diazotrophicus had recovered to
approximately 80% of the original activity. Changes in nitrogenase
activity (Fig. 4) and respiration rate (data not shown) in response to
the single-step change from 20 to 60 kPa of O2
were similar in magnitude (i.e., a 74% decrease in nitrogenase
activity and an approximate 10% increase for respiration) to those
observed when the increase in from 20 and 60 kPa of
O2 took place in several 10-kPa steps (Fig. 1 and
3).
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Effects of small stepped changes in atmospheric pO2 on
G. diazotrophicus grown at 10 or 30 kPa of
O2.
G. diazotrophicus colonies were grown
under 10 or 30 kPa of atmospheric O2 for 5 to 6 days and then assayed for total nitrogenase activity
at a range of atmospheric
pO2 (5 to 60 kPa) (Fig. 5 and 6). In both cases, maximal nitrogenase
activity occurred at 10 to 20 kPa of O2 above the
atmospheric pO2 at which the colonies had been
grown. For colonies grown under 10 kPa of O2,
nitrogenase activity was maximized at 20 and 30 kPa of
O2 (Fig. 5). For colonies grown under 30 kPa of
O2, nitrogenase activity was maximized at 40 kPa
of O2 (Fig. 6).
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Hydrogenase uptake activity of G.
diazotrophicus.
There was no detectable
H2 consumption by the Nif
mutant of G. diazotrophicus (MAD3A) under typical conditions
experienced by the wild-type strain when nitrogenase activity was
assayed (2 ppm of H2 in air in a flowthrough
system at ambient atmospheric pO2). However, when
the strain was supplied with 50 ppm of H2 in air
in a closed-assay system, we detected a H2
consumption rates of 0.016 ± 0.009 µmol of
H2 1010
cells1 h1. This rate is
low, as the H2 evolution rate by wild-type
G. diazotrophicus assayed under the same conditions was
0.362 ± 0.027 µmol of H2 1010 cells1
h1 and H2 consumption
rate by the Hup+ aerobe R. eutropha
was 0.080 ± 0.003 µmol of H2
1010 cells1
h1.
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DISCUSSION |
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Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
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The concentration of O2 at the sites of nitrogenase activity in aerobic and microaerophilic diazotrophs is the result of the interplay among (i) the concentration of O2 in the surrounding atmosphere, (ii) the diffusion rate of O2 from the surrounding atmosphere to the sites of nitrogenase activity, (iii) the consumption rate of O2 in the vicinity of nitrogenase (predominantly via oxidative phosphorylation), and (iv) the role of carriers of O2 which facilitate diffusion in some systems (such as leghemoglobin in legume nodules) (21). The present study investigated responses in nitrogenase activity to changes in atmospheric pO2 around colonies in a flowthrough system; previous studies (5, 40) injected enough pure O2 into a previously anaerobic, closed liquid system to achieve target pO2. All these studies enable observation of changes in nitrogenase activity in response to relative changes in O2 flux to the bacteria; however, neither the actual flux of O2 to the diazotrophs or the actual concentration of O2 at the sites of nitrogenase activity was determined.
In our study, nitrogenase activity by G. diazotrophicus was tolerant of atmospheric pO2 as high as 60 kPa (Fig. 1). These findings are not in conflict with previous studies (5, 40) that found that nitrogenase activity by G. diazotrophicus in liquid culture was totally inhibited when the culture was at equilibrium with 6 kPa of O2 in the gas phase. These findings simply reflect that in our study, atmospheric pO2 surrounding the colonies was changed, and in the previous studies, the partial pressure of dissolved oxygen in liquid cultures was changed. However, comparison of these studies indicates that G. diazotrophicus can use the milieu of a colony as an effective resistance to O2 diffusion, resulting in an O2 concentration and O2 flux within the colony which enable the bacteria to fix N2 in a broad range of atmospheric pO2 surrounding the colony.
Using H2 production as a measure of nitrogenase activity in closed-assay systems, Dong et al. (8, 10) showed that G. diazotrophicus could fix N2 in colonies with 2 and 20 kPa of O2 in the surrounding atmosphere and suggested that colony structure and location of bacteria within the colony played a role in the protection of nitrogenase from excessive O2 flux. Bacterial mucilage is known to decrease the rate of oxygen diffusion to cells (3). The presence of extracellular polysaccharide surrounding Beijerinckia derxii cells is necessary to maintain nitrogenase in this organism (1). Derxia gummosa forms small nonfixing colonies if grown at 20 kPa of O2; however, if grown at 5 kPa of O2, the bacterium forms large, highly mucilaginous colonies which fix N2 (17, 18). The motile diazotroph Azospirillum brasilense (50) is known to display aerotaxis within suspensions to achieve the appropriate O2 environment for N2 fixation.
For colonies grown at 20 kPa of O2 and assayed at
the same atmospheric pO2, total nitrogenase
activity was approximately 0.5 µmol of H2
1010 cells1
h1 (Fig. 1). Is this a relatively low or high
rate of nitrogenase activity? We have compared the level of nitrogenase
activity in G. diazotrophicus to that of
Bradyrhizobium japonicum in a typical soybean (Glycine
max [L.] Merr.) nodule. Based on measurements of nodules on
5-week-old soybean plants, Lin et al. (28) found that
nodules contained approximately 109 B. japonicum bacteroids each and that total nitrogenase activity was
in the range of 2.0 to 4.0 µmol of H2
1010 cells1
h1. Nitrogenase activity for G. diazotrophicus colonies at ambient atmospheric
pO2 in our study was approximately 12 to 25% of
the rates calculated for B. japonicum in soybean nodules.
We consider such a level of nitrogenase activity by G. diazotrophicus in colonies to be remarkably high considering that
a soybean nodule is a highly sophisticated organ designed to provide a
highly conducive milieu (in terms of O2 flux,
carbon supply, assimilation of fixed N, etc.) for bacteroids to fix
N2.
Our study is the first measure of EACs for G. diazotrophicus. We found that the EAC of G. diazotrophicus at 20 kPa of O2 was approximately 0.6 (Fig. 2), meaning that in air, approximately 60% of electron flow through nitrogenase would be allocated to reduction of dinitrogen and 40% would be allocated to proton reduction. Again, the EAC of G. diazotrophicus can be put into context by comparing it to that of rhizobia in legume nodules. The theoretical maximum for EAC is 0.75 (i.e., at least one H2 produced for every N2 fixed by nitrogenase) (43). The EACs of legume symbioses are commonly between 0.59 and 0.70 (21). The reason for the variability in the EAC is not clearly understood (19, 26). Our measurements of the EAC for nitrogenase in G. diazotrophicus grown on solid medium at ambient atmospheric pO2 is in the same range as EACs commonly observed in legume nodules.
The accuracy of measurements of nitrogenase activity by
H2 evolution is dependent upon the lack of
hydrogenase uptake activity (21, 26). Although the
Nif mutant of G. diazotrophicus was
seen to have a low level of Hup+ activity when
supplied with relatively high levels of H2 (50 ppm) in a closed-assay system, under the standard conditions in our
flowthrough assay system (i.e., 2 ppm of H2),
Hup+ activity was not detectable.
Small (5- to 10-kPa) decreases in atmospheric pO2 resulted in declines in nitrogenase activity and respiration rate in G. diazotrophicus grown at 20 kPa of O2 (Fig. 1). This is clearly representative of an O2 limitation of cellular metabolism and has been seen in other aerobically functional N2-fixing systems, such as Azotobacter (48) and soybean nodules (25). However, small stepwise increases in atmospheric pO2 above 20 kPa also resulted in declines in nitrogenase activity (Fig. 1). The declines in nitrogenase activity with small (10-kPa) increases in atmospheric pO2 could occur for one of two reasons: (i) an irreversible O2-induced denaturation of the nitrogenase enzyme or (ii) a reversible controlled down-regulation of nitrogenase activity (14). The time course of nitrogenase activity in response to single-step, 40-kPa changes in atmospheric pO2 (Fig. 4) indicates that the latter and not the former mechanism is at work in G. diazotrophicus. The rapid decrease in nitrogenase activity when atmospheric pO2 was switched from 20 to 60 kPa, and the subsequent rapid recovery when the bacteria returned to 20 kPa, indicate that G. diazotrophicus has a switch-off/switch-on protection mechanism in response to changes in atmospheric pO2.
Reversible inhibition of nitrogenase activity has been seen in a number of diazotrophs in response to increases in pO2 and to the addition of ammonium. Three underlying physiological mechanisms have been associated with switch-off/switch-on kinetics of nitrogenase in diazotrophs. The switch-off/switch-on mechanism can be the result of an O2-induced conformational change in nitrogenase as seen in the Mo-dependent nitrogenase of Azotobacter (11, 32, 35, 36, 41). Switch-off/switch-on mechanisms can also be facilitated by an ADP-ribosylation of dinitrogenase reductase which halts nitrogenase activity. This mechanism is coded for by the draT and draG genes and has been observed in a number of diazotrophs, including Rhodobacter capsulata (46), Rhodospirillum rubrum, Azospirillum brasilense, and Azospirillum lipoferum (34, 49). Finally, the nitrogenase switch-off/switch-on mechanism in a number of diazotrophs may involve diversion of electrons from nitrogenase to other (unidentified) electron acceptors (16) and/or an ATP limitation of nitrogenase activity, possibly due to a switch to uncoupled respiratory chain as proposed for Azotobacter vinelandii (29).
Which if any of the above identified switch-off/switch-on mechanisms are at work in G. diazotrophicus was not investigated in our study. However, it is highly unlikely that the mechanism involves the ADP-ribosylation of dinitrogenase reductase. Although Burris et al. (5) found that G. diazotrophicus had "a rather sluggish" response to ammonium addition and required 10 µM NH4+ to switch off nitrogenase, they found no evidence of ADP-ribosylation of dinitrogenase reductase or of the draT-draG gene complex in G. diazotrophicus. Recently, S. Norlund (personal communication) also found evidence of a switch-off/switch-on phenomenon in G. diazotrophicus in response to changes in pO2, possibly involving a conformational change in nitrogenase medium by a Shethna-like protein (32).
In this study, long-term adaptation of G. diazotrophicus to different atmospheric pO2 was tested by growing the bacterium for 5 or 6 days at 10, 20, or 30 kPa of O2 before nitrogenase activity was measured. Although culture conditions were not exactly the same for all the cultures (see Material and Methods), some trends are consistent in all three treatments. During assays of nitrogenase activity, when atmospheric pO2 was decreased below the concentration at which the bacteria were cultured, nitrogenase activity was always lower (Fig. 1, 5, and 6). This appears to be due to a generalized O2 limitation of cellular metabolism (Fig. 3) (25, 48). G. diazotrophicus cultures which were grown under different atmospheric pO2 also showed different optimal atmospheric pO2 for nitrogenase activity. The optimal atmospheric pO2 for cultures grown at 10 and 20 kPa of O2 was 20 to 30 kPa of O2 (Fig. 1 and 5); the optimal atmospheric pO2 for nitrogenase activity for cultures grown at 30 kPa of O2 was 40 kPa of O2 (Fig. 6) The fact that the cultures grown at the highest atmospheric pO2 showed a higher optimal pO2 for nitrogenase activity indicates a long-term adaptation of G. diazotrophicus colonies to different pO2. Other aerobically functional N2-fixing systems such as A. vinelandii (30) and the B. japonicum-soybean symbiosis (7) are known to make long-term adaptations of nitrogenase activity to nonambient pO2. Dong (8) noted differences in colony morphology of G. diazotrophicus between cultures grown long-term on 2 and 20 kPa of atmospheric pO2. We are currently investigating whether morphologic and structural characteristics of the colonies contribute to these long-term adaptations.
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ACKNOWLEDGMENTS |
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We thank B. Luit and J. Foidart for their technical assistance,
C. Kennedy for donating the Nif strain G.
diazotrophicus MAD3A, and P. Hallenbeck, Université de
Montréal, for useful discussions on this paper.
This study was funded by grants from Cargill Inc. and the AAFC/NSERC (Canada) Research Partnership Program.
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FOOTNOTES |
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* Corresponding author. Mailing address: Department of Plant Science, University of Manitoba, Winnipeg, MB, R3T 2N2, Canada. Phone: (204) 474-8251. Fax: (204) 474-7528. E-mail: k_vessey{at}umanitoba.ca.
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REFERENCES |
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Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
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Applied and Environmental Microbiology, October 2001, p. 4694-4700, Vol. 67, No. 10
0099-2240/01/$04.00+0 DOI: 10.1128/AEM.67.10.4694-4700.2001
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