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Appl Environ Microbiol. Oct 2001; 67(10): 4694–4700.
PMCID: PMC93221

Response of the Endophytic Diazotroph Gluconacetobacter diazotrophicus on Solid Media to Changes in Atmospheric Partial O2 Pressure

Abstract

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) 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 cells−1 h−1) 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.

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−1 year−1 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).

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.

MATERIALS AND METHODS

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−1, 0.1 mg of biotin liter−1, 0.2 mg of pyridaxol HCl liter−1, and 5 ml of sugarcane juice (pressed from fresh sugarcane stem) liter−1, 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 liter−1 plus 50 mg of yeast extract liter−1). 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.

G. diazotrophicus was grown on solid LGI-P medium in petri plates for 5 or 6 days at 30°C prior to gas exchange measurements. Cultures were grown under ambient atmospheric pO2 (approximately 20 kPa) or in a gas exchange chamber (see below) with 10 or 30 kPa of O2 flowing through the chamber at approximately 65 ml per min. For cultures grown in the chamber, input air was bubbled through a flask of water prior to being fed into the chamber to avoid desiccation of the cultures.

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 min−1. 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 ml−1 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 min−1 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 min−1 (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 min−1. 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 min−1 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).

RESULTS

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. (Fig.1).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.

FIG. 1
Effect of atmospheric pO2 on total nitrogenase activity (H2 evolution in Ar-O2) of G. diazotrophicus colonies grown at 20 kPa of O2. Data are means plus standard errors (n = 4). Results with different letters are significantly different at a P value of ...

Stepped increases of 10 kPa of O2 above 20 kPa had no significant effect on the EAC of nitrogenase activity (Fig. (Fig.2).2). However, as atmospheric pO2 was lowered from 20 kPa to10 and 5 kPa of O2, the EAC decreased.

FIG. 2
Effect of atmospheric pO2 on EACs of nitrogenase of G. diazotrophicus grown at 20 kPa of O2. Data are means plus standard errors (n = 4). Data with different letters are significantly different at a P value of ≤0.05.

As atmospheric pO2 was increased from 20 to 60 kPa of O2 in 10-kPa steps, the respiration rate of G. diazotrophicus cells increased marginally (Fig. (Fig.3).3). For example, the threefold increase in atmospheric pO2 from 20 to 60 kPa resulted in an 11% increase in CO2 evolution per cell. In contrast, decreasing atmospheric pO2 from 20 to 10 kPa and then 5 kPa resulted in severe decreases in respiration rates of 39 and 51%, respectively.

FIG. 3
Effect of atmospheric pO2 on respiration rate (CO2 evolution in N2-O2) of G. diazotrophicus colonies grown at 20 kPa of O2. Data are means plus standard errors (n = 4). Data with different letters are significantly different at a P value of ≤0.05. ...

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. (Fig.4).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. (Fig.4)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. (Fig.11 and and3).3).

FIG. 4
Time course of the response of nitrogenase activity (H2 evolution in Ar-O2) of G. diazotrophicus colonies to large, sudden changes in O2 concentration. The gas composition (top axis) was changed from 20 to 60 kPa of O2, maintained for 15 min, then changed ...

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. (Fig.55 and and6).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. (Fig.5).5). For colonies grown under 30 kPa of O2, nitrogenase activity was maximized at 40 kPa of O2 (Fig. (Fig.6).6).

FIG. 5
Effect of atmospheric pO2 on nitrogenase activity (H2 evolution in Ar-O2) of G. diazotrophicus colonies grown at 10 kPa of O2. Data are means plus standard errors (n = 4). Data with different letters are significantly different at a P value of ≤0.05. ...
FIG. 6
Effect of atmospheric pO2 on nitrogenase activity (H2 evolution in Ar-O2) of G. diazotrophicus colonies grown at 30 kPa of O2. Data are means plus standard errors (n = 4). Data with different letters are significantly different at a P value of ≤0.05. ...

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 cells−1 h−1. 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 cells−1 h−1 and H2 consumption rate by the Hup+ aerobe R. eutropha was 0.080 ± 0.003 μmol of H2 1010 cells−1 h−1.

DISCUSSION

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. (Fig.1).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 cells−1 h−1 (Fig. (Fig.1).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 cells−1 h−1. 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. (Fig.2),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. (Fig.1).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. (Fig.1).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. (Fig.4)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. (Fig.1,1, ,5,5, and and6).6). This appears to be due to a generalized O2 limitation of cellular metabolism (Fig. (Fig.3)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. (Fig.11 and and5);5); the optimal atmospheric pO2 for nitrogenase activity for cultures grown at 30 kPa of O2 was 40 kPa of O2 (Fig. (Fig.6)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.

ACKNOWLEDGMENTS

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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