Department of Biology, Carleton University, Ottawa, ON, CanadaK1S 5B61
Department of Plant Science, University of Manitoba, Winnipeg, MB, CanadaR3T 2N22
Department of Physiology, University of Manitoba, Winnipeg, MB, CanadaR3E 3J73
Author for correspondence: J. K. Vessey. Tel: +1 204 474 8251. Fax: +1 204 474 7528. e-mail: k_vessey{at}umanitoba.ca
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ABSTRACT |
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Keywords: Acetobacter diazotrophicus, diffusion resistance, N2 fixation, nitrogen, oxygen
Abbreviations: CLSM, confocal laser scanning microscopy; DAI, days after inoculation; pO2, partial pressure of oxygen
a Present address: Department of Biology, St Marys University, Halifax, NS, Canada B3H 3C3.
b Present address: Division of Plant Industry, CSIRO, GPO 1600, Canberra, ACT 2601, Australia.
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INTRODUCTION |
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Sugarcane does not form any specialized structure to host G. diazotrophicus (Dong et al., 1994 ) that may aid in the regulation of O2 flux as root nodules do in legume plants (Hunt & Layzell, 1993
). Nitrogenase activity by G. diazotrophicus in liquid medium was optimized when the dissolved oxygen content of the medium was equilibrated with 0·2 kPa O2 in the gas phase (Reis & Döbereiner, 1998
). However, G. diazotrophicus is able to use N2 as the its sole nitrogen source under 21 kPa O2 on a semi-solid medium (Cavalcante & Döbereiner, 1988
). Under these conditions, distinct colonies are not formed; rather, the bacteria grow just below the surface of the media. This behaviour may help to optimize the O2 flux to the bacterium as seen in other aerotactic diazotrophs (Zhulin et al., 1996
). On solid medium, distinct, superficial colonies of G. diazotrophicus form thick, mucilaginous matrices and are able to grow on N2 as the sole nitrogen source at 20 kPa O2 (Dong et al., 1994
). Pan & Vessey (2001)
showed that bacterial respiration and nitrogenase activity by G. diazotrophicus in colonies adapted over long-term exposures (i.e. several days) to different atmospheric pO2 (10, 20 and 30 kPa). Optimal nitrogenase activity by G. diazotrophicus colonies occurred at or slightly above (i.e. +10 kPa) the O2 concentrations at which they were grown (Pan & Vessey, 2001
).
Since nitrogenase is active in G. diazotrophicus colonies grown on solid media, the bacterium must have means of ensuring an appropriate concentration and flux of O2 to balance aerobic respiration and nitrogenase activity. In this study, we test the hypothesis that G. diazotrophicus positions itself within the mucilaginous matrix of its colony to achieve an appropriate O2 environment for nitrogenase activity, and that an intact colony structure is required to maintain this nitrogenase activity. This hypothesis was tested by comparing G. diazotrophicus colony structure when grown on solid medium under 2 and 20 kPa pO2, correlating nitrogenase activity to colony development, and observing nitrogenase activity in response to disruption of colony structure.
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METHODS |
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Colony structure was examined in both live, intact colonies and fixed, sectioned colonies. Cells of G. diazotrophicus accumulated the pH indicator bromothymol blue from the LGI-P medium. Fluorescence of the pH indicator permitted the visualization of cells within live colonies using confocal laser scanning microscopy (CLSM). A minimum of six randomly selected 4-day-old colonies of G. diazotrophicus PAL 5 from each of the two pO2 treatments were examined using a Bio-Rad MRC600 CLSM equipped with a 514 Argon laser utilizing a GHS filter block (514 nm DF excitation/550 nm LP emission). An inverted stage and 32x open air objective were used to view the colonies, which were removed with a thin layer of subtending agar from the plates on which they were grown. Optical sections (Z-series) were initiated at the top of the colony mucilage and collected at 5 µm intervals down through each colony toward the agar substrate, to the maximum penetration depth of the laser (120 µm). A montage of the Z-series was produced using Bio-Rads Confocal Assistant v. 4.02.
For light microscopy, a freeze-substitution method was used because standard fixatives flooded over the agar surface caused the colonies to rupture. Five-day-old colonies of G. diazotrophicus JO-2, supported by small pieces of the subtending LGI-P agar, were plunged into the freezing mixture (isopentane/methyl cyclohexane, 1:1, at the melting point). Freeze substitution in dry acetone resulted in the formation of sucrose crystals (from the medium), which damaged the structure of the colonies. The colonies were therefore freeze-substituted in methanol/acrolein (10:1) at -80 °C for 7 days. Under these conditions, sucrose crystals formed at the bottom of the vial or on the surface of the agar, from which they were easily removed.
Freeze-substituted colonies were gradually warmed (-20 °C overnight, then +5 °C for 24 h), rinsed (3x10 min each) in methanol on ice, post-fixed with 1% OsO4 in methanol for 1 h on ice, then rinsed again with three changes of methanol. The methanol was replaced by the transition solvent acetone in a graded series (5, 10, 20, 50, 70, 90 and 100% acetone; two 10 min changes per step). Colonies were gradually infiltrated in Spurrs resin monomer mixture (Spurr, 1969 ), with the concentration of the resin in acetone reaching 5% at 90 min, 10% at 150 min, 25% at 210 min and 75% at 330 min. The vials were then covered with perforated foil to allow evaporation of the remaining acetone overnight. The next day, resin in the vials was replaced with 100% resin and polymerized at 70 °C overnight. Mid-colony transverse sections were cut with glass knives, stained with toluidine blue O (0·05% in benzoate/borate buffer at pH 4·4), mounted in immersion oil and viewed with an Olympus Vanox microscope using phase-contrast and brightfield optics. Transmission electron microcopy was performed to confirm that stained bodies seen in the light microscopy corresponded to bacteria (data not shown).
Observation of nitrogenase activity.
Nitrogenase activity was assayed for developing G. diazotrophicus colonies and for mature colonies before and after physical disruption. Nitrogenase activity was measured by H2 evolution in the presence of Ar/O2 (Hunt & Layzell, 1993 ) in a flow-though gas-exchange system (Pan & Vessey, 2001
). To assess the effect of physical disruption of colony structure on nitrogenase activity, G. diazotrophicus PAL 5 was grown on solid, modified LGI-P medium (Pan & Vessey, 2001
). At 6 days after inoculation (DAI), 20 Petri dishes containing 100150 colonies per dish were placed in the gas-exchange system. A gas mixture of Ar/O2 (80:20) was passed through the chamber at the rate of 500 ml min-1. Hydrogen evolution of the colonies was recorded after 1 h, then the plates were removed from the chamber. The colonies on each plate were gently disrupted by using a glass rod to smear the colonies on the surface of the agar to approximately twice their original surface area. The plates were returned to the chamber along with the bent glass rod, and once again exposed to the Ar/O2 mixture. Hydrogen evolution by the disrupted colonies was recorded after 1 h and reported as nmol H2 h-1 per colony±SEM.
To assess the effect of colony development and morphology on nitrogenase activity, G. diazotrophicus PAL 5 was inoculated onto Petri dishes containing solid, modified LGI-P medium (Pan & Vessey, 2001 ) and incubated at 30 °C. Discrete colonies were visible at 2 DAI. Nitrogenase activity of the intact colonies was measured daily from 3 to 8 DAI by H2 evolution in the flow-through gas-exchange system. Nitrogenase measurements were performed daily on four replicates of 20 Petri dishes each (100150 colonies per dish) as described above. Because of increasing colony size over time, bacterial titre per colony was quantified and nitrogenase activity is reported as H2 evolution rate per cell number (i.e. µmol H2 per 1010 cells h-1). Concentrations of bacteria per colony were assessed by plate counting as detailed by Pan & Vessey (2001)
. Colonies were visually assessed daily from 3 to 8 DAI for breakdown of colony structure. Breakdown of colony structure was indicated by a slumping of the upper layer of mucilage to one side of the colony (easily visible due to the brightly yellow stained bacteria) and a flattening of the colony profile.
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RESULTS AND DISCUSSION |
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Influence of pO2 on colony structure
Colonies grown for 4 days under 2 and 20 kPa pO2 were lens-shaped in transverse section. In both treatments, two bacterial populations were seen embedded in the matrix of each colony, one adjacent to the agar medium, the other in a layer closer to the upper surface of the colony, with a low density of cells in between the two (Figs 1 and 2
). The toluidine-blue-stained colonies (Fig. 1
) clearly demonstrate that the position of the upper population varied with pO2 treatment. At 20 kPa O2, the uppermost population of bacteria was midway between the surface and the base of the colony (Fig. 1a
). In contrast, the upper population of bacterial cells in colonies grown at 2 kPa O2 was positioned just below the surface of the colony (Fig. 1b
). The direction of the knife blade sectioning these colonies was from the top of the colony to the bottom. The vertical striations in these sections (particularly Fig. 1a
) were due to fine crystals of sucrose remaining in the colony after fixation.
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Because fixation processes have the potential of disrupting the structure of what is being observed, live colonies grown at 2 and 20 kPa O2 were also observed by CLSM. The difference in location of the upper population of bacteria between the pO2 treatments was also evident in intact colonies of G. diazotrophicus (Fig. 3). Each panel in these images represent a 5 µm optical section through a living colony, starting at the top of the dome of the colony (upper left panel) and moving down towards the base of the colony (lower left panel). The whitish florescence indicates the presence of bacteria due to laser-induced excitation of bromothymol blue bound to bacterial capsular material. For colonies grown at 20 kPa O2, the highest density of the upper population was typically located at a depth of 85100 µm below the highest point of the colony surface (Fig. 3a
). At 2 kPa O2, the majority of cells of the upper population was typically located only 4560 µm below the top surface of the mucilage (Fig. 3b
). The haloing effect seen in the sections from 60 to 95 µm (Fig. 3b
) shows that this upper population of bacteria is following the contour of the dome-shaped colony. Because the penetration of the CLSM system was limited to just beyond the upper 100 µm of the colony, the lower population of cells near the base of the colonies (Fig. 2
) could not be imaged.
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Nitrogenase activity as influenced by physical disruption of colonies and by colony development
Nitrogenase activity of intact G. diazotrophicus colonies grown at 20 kPa O2 at 6 DAI was 1·14±0·07nmol H2 h-1 per colony. After physical disruption of colony structure by smearing colonies across the agar surface with a glass rod, nitrogenase activity was decreased by 96·7% to 0·038±0·006 nmol H2 h-1 per colony. Likewise, breakdown in colony structure due to development/ageing (Figs 4 and 5
) also resulted in declines in nitrogenase activity.
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Nitrogenase activity of the colonies grown at 20 kPa O2 (Fig. 5b) was detectable at 3 DAI, although at a very low rate of <0·2 µmol H2 per 1010 cells h-1. Nitrogenase activity increased daily to a maximum value of 0·697 µmol H2 per 1010 cells h-1 at 6 DAI. After this time, nitrogenase activity decreased, declining to only 76% of the maximum at 8 DAI. Hence, the increase in the breakdown in colony structure due to ageing (Fig. 5a
) was coincident with the decline in nitrogenase activity per cell within the colonies (Fig. 5b
).
Disruption of colony structure due to either manipulation or ageing would compromise all spatial relationships between the bacterial cells and path-length of mucilage to the open atmosphere. Decline in the path-length between bacteria and the atmosphere would result in a dramatic increase in O2 flux to the bacteria and in the concentration of free O2 at the sites of nitrogenase activity. Pan & Vessey (2001) demonstrated that G. diazotrophicus displays a rapid switch-off protection phenomenon when O2 flux rapidly increases to G. diazotrophicus within colonies. Alternatively, a rapid increase in O2 flux to G. diazotrophicus with disruption of colony structural integrity could result in a reversible inhibition of nitrogenase activity (Burris, 1991
).
Physical disturbance of the colonies by manipulation with a glass rod resulted in a much greater decline in nitrogenase activity (96·7%) than that caused by the slumping of colonies due to ageing between 6 and 8 DAI (Fig. 5b; a 24% decline in nitrogenase activity). However, this is not unexpected as the smearing of the colony with the glass rod is a much more severe physical disturbance than that induced by colony slumping.
Conclusion
The results of both the imaging of colonies grown at different pO2 values and the correlation of nitrogenase activity with colony intactness are consistent with the hypothesis that the mucilaginous matrix of G. diazotrophicus colonies is important in the protection of nitrogenase activity by the bacterium from excessive O2 flux. Likewise, the position of G. diazotrophicus within the colony appears to be a component of the bacteriums long-term adaptation to changes in pO2 in the surrounding atmosphere. However, the actual concentration of free and dissolved O2 within the colonies at the sites of nitrogenase activity is as yet unknown. Reis & Döbereiner (1998) found that nitrogenase activity of G. diazotrophicus in liquid cultures was maximal when the culture was at equilibrium with 0·2 kPa O2 in the gas phase. However, the actual concentration of dissolved O2 at the site of nitrogenase activity in any medium is dependent upon the concentration of O2 in the gas phase, the diffusion rate through the medium, and the rate of O2 consumption by bacterial respiration (Hunt & Layzell, 1993
). The current study supports the hypothesis that G. diazotrophicus utilizes the path-length of colony mucilage between the atmosphere and the bacteria to achieve this optimal flux and concentration of O2 for respiration and nitrogenase activity.
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ACKNOWLEDGEMENTS |
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Received 5 December 2001;
revised 11 March 2002;
accepted 29 April 2002.
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