Journal of Experimental Botany, Vol. 54, No. 381, pp. 325-334,
January 2, 2003
© 2003 Oxford University Press
Biosensor reporting of root exudation from Hordeum vulgare in relation to shoot nitrate concentration
Received 20 December 2001; Accepted 26 August 2002
1 The Macaulay Institute, Craigiebuckler, Aberdeen AB15 8QH, Scotland, UK
2 Department of Plant and Soil Science, University of Aberdeen, Aberdeen AB24 3UU, Scotland, UK
3 School of Biological Sciences, University of Wales, Bangor LL57 2UW, Wales, UK
4 To whom correspondence should be addressed. Fax: +44 (0)1224 311 556. E-mail: eric.paterson{at}macaulay.ac.uk
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Abstract |
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The aim of this study was to determine the relationship between shoot nitrate concentration, mediated by nitrate supply to roots, and root exudation from Hordeum vulgare. Plants were grown for 14 d in C-free sand microcosms, supplied with nutrient solution containing 2 mM nitrate. After this period, three treatments were applied for a further 14 d: (A) continued supply with 2 mM nitrate (zero boost), (B) supply with 10 mM nitrate (low boost), and (C) supply with 20 mM nitrate (high boost). At the end of the treatment period, a bacterial biosensor (Pseudomonas fluorescens 10586 pUCD607, marked with the lux CDABE genes for bioluminescence) was applied to the microcosms to report on C-substrate availability, as a consequence of root exudation. The nitrate boost treatments significantly affected shoot nitrate concentrations, in the order C>B>A. In treatments receiving a nitrate boost (B, C), increased shoot nitrate concentration was correlated with increased plant biomass, reduced root length, reduced number of root tips, and increased mean root diameter, relative to the no boost treatment (A). Imaging of biosensor bioluminescence (proportional to metabolic activity in response to availability of root exudates) indicated that root exudation increased with decreasing shoot nitrate concentration. Biosensor reporting of root C-flow indicated that exudation was greater from root tip regions than from the whole root, but that specific exudation rates for all sites were unaffected by treatments. Total root exudation across treatments was found to be closely correlated with total root length, indicating that increased root exudation, per unit root biomass, with decreasing nitrate supply was associated with altered root morphology, as a consequence of systemic plant responses to internal N-status.
Key words: Barley, biosensor reporting, root exudation, shoot nitrate.
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Introduction |
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Plasticity of root system functioning and structure has been demonstrated across a broad range of plant species in response to variations in nitrogen supply. This plasticity is manifested at a number of levels, from gross alterations in biomass partitioning between roots and shoots (Reynolds and D’Antonio, 1996), altered root morphology and architecture (Fitter, 1982; Robinson, 1994), and the physiological regulation of N-uptake processes (Jackson et al., 1990). These responses are considered adaptive with respect to the maintenance of a functional equilibrium between C and N acquisition (Thornley, 1995). In soil, the efficiency of these strategies to acquire N is mediated by competition with co-existing plants (Hodge et al., 1999) and by interactions with soil microorganisms (Hodge et al., 2000).
Microbial growth and activity in the rhizosphere is driven by organic compounds released from roots (Lynch and Whipps, 1990). Therefore, around roots there is an active microbial community, which is in competition with plant roots for available N. It could be hypothesized that this would reduce N-acquisition by roots, particularly under limiting N-supply. Indeed, this prediction is supported by modelling approaches for root acquisition of phosphate in competition with the rhizosphere microbial biomass (Darrah, 1998). It would also be expected that this effect would be exacerbated at low N supply due to reduced N-content of rhizodeposits (Janzen, 1990), which would increase the microbial N-requirement in the rhizosphere. However, an alternative hypothesis is that rhizodeposition by stimulating microbial activity would consequently increase microbial mineralization of N from soil organic matter (Kuzyakov et al., 2000). Although much of this mineralized N would initially be incorporated into the microbial biomass, it is suggested that due to the rapid microbial turnover (relative to roots) and feeding activity of protozoa, mineralized N would ultimately become available to plant roots (Clarholm, 1985a, b). Consequently, microbially immobilized N in the rhizosphere, although not immediately available for root uptake, could function as a ‘slow-release store’ for the root.
To gain a better understanding of these interactions, it is necessary to determine the effects of N-supply on rhizodeposition in detail. Studies in hydroponics and axenic sand culture systems suggest that loss of C per unit weight of root is increased with low N-supply (Paterson and Sim, 1999, 2000). However, it is unclear whether this is due to altered exudation intensity from the whole root, or due to exudation from sites which are more abundant under low N supply (e.g. junctions of lateral roots and root tips) and have inherently greater rates of exudation. In soil, the effects of N supply on rhizodeposition have been studied through the use of 14C-tracers. The results of these studies are inconclusive, with both positive (Johansson, 1992) and negative (Merckx et al., 1987; Liljeroth et al., 1990) effects of low N supply on rhizodeposition reported. Contradictory findings may partly be due to methodological difficulties with the use of 14C-labelling. Pulse-labelling results in incomplete labelling of plant C-pools, with the quantified release from roots biased to those pools which receive the greatest contribution from recent assimilate (Meharg, 1994; Paterson et al., 1997). Consequently, the duration of the labelling and chase periods are critical in determining the outcome of these experiments. In addition, 14C-labelling methodologies cannot distinguish between root and microbial respiration. Concurrent with changes in N-supply, the respiratory costs of root uptake, construction and maintenance would be expected to change (Fitter, 1994), resulting in quantitative shifts in root respiration. Therefore, for the consideration of the supply of substrates to the rhizosphere microbial biomass, 14C-labelling has considerable difficulties.
Recently, the potential of utilizing bacterial gene reporter systems to investigate C-release from roots has been demonstrated (Kragelund et al., 1997; Jaeger et al., 1999; Yeomans et al., 1999). This approach has several advantages over the chemical characterization of exudates from hydroponic cultures and 14C-labelling methods. Firstly, because bacterial reporter systems respond to the organic substrate and the quantification of activity is independent of microbial CO2 production, root respiration does not interfere with interpretations of microbial substrate use. Secondly, reporters (lux, lacZ, gus, and ice nucleation genes) can be used to characterize substrate use spatially, relative to roots and the growth matrix, providing information on the location and intensity of root exudation sites. Finally, reporter systems can be constructed to report on whole-cell metabolic activity or the activity of single catabolic pathways, yielding information on total exudation or on a single component (Jaeger et al., 1999).
The partitioning of assimilate within plants is strongly affected by the balance of C acquisition by shoots and nutrient acquisition by roots (Reynolds and Chen, 1996; Ingestad and McDonald, 1989). Typically, this results in allocation balanced to optimize the acquisition of these resources for growth, described as establishing a functional equilibrium (Thornley, 1977). During the consideration of nitrogen acquisition, the functional equilibrium hypothesis predicts proportionately greater root growth under low N-supply, and greater shoot growth with plentiful N-supply or low light. For individual plants this prediction is well supported by published data (Minchin and Thorpe, 1996; Smolders and Merckx, 1992). The functional equilibrium hypothesis does not, however, provide a physiological explanation of how the plant regulates the relative allocation of assimilate between roots and shoots. Several hypotheses have been proposed to describe where the plant control of assimilate allocation resides (reviewed by Farrar and Jones, 2000) and a number of mechanisms by which this control is exerted have also been proposed (reviewed by Marschner et al., 1996). Currently, there is no generally accepted theory in relation to these processes and their control. It has been demonstrated that high concentrations of nitrate in leaves, indicative of nitrate uptake in excess of immediate reduction and N usage in growth processes, is negatively correlated with assimilate allocation to roots (Scheible et al., 1997b). As root exudation is closely related to assimilate allocation to roots (Rattray et al., 1995), it is hypothesized that C-efflux from roots may also be inversely related to high shoot nitrate concentrations and high soil nitrate availability.
The aim of this study was to test the hypothesis that root exudation from Hordeum vulgare is negatively correlated with leaf nitrate concentration. In addition, the study aimed to characterize exudate release from roots spatially, concurrent with root structural changes in response to N-supply treatments. The approach was to generate plants with differing leaf nitrate concentrations by the manipulation of N-supply to the roots and to identify sites of exudation with a lux-marked rhizobacterial reporter system, Pseudomonas fluorescens 10586 pUCD607 (Yeomans et al., 1999), through quantification of light with a Charge Coupled Device (CCD). The reporter construct contains the lux CDABE gene cassette from Vibrio fischeri. These genes encode both the luciferase heterodimer responsible for oxidation of FMNH2 and a fatty acid aldehyde (light-producing reaction), and the regeneration of the aldehyde. The construct was originally selected on the basis of light output proportional to metabolic activity (as quantified by dehydrogenase activity), for a range of substrates.
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Materials and methods |
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Plant material
Seeds of winter barley (Hordeum vulgare L. cv. Melanie) were surface-sterilized in 1% peracetic acid for 2 min, rinsed in distilled water (repeated five times), germinated between sterile filter paper, and then transferred to sterile tissue lined cylinders for 48 h until the roots were c. 2 cm in length. Individual seedlings were transferred to sterile sand-filled plate microcosms (see below) after this period. Microcosms were sited in a growth cabinet (ConvironTM S10H growth cabinet, Winnipeg, Canada) and angled at 30° above horizontal. Plants were maintained at 20 °C with a photoperiod of 14/10 h (light/dark) and a PAR of 350 µmol m–2 s–1 for 28 d.
Microcosms
The microcosms (Fig. 1) were constructed through the adaptation of transparent polystyrene boxes (Norlab Instruments Ltd, Loughborough, UK, dimensions 27.9x15.9x10.2 cm). The lid was inverted and two strips of TerostatTM elastomer (Teroson, NDA Engineering Equipment Ltd., Kempston, Bedford, UK) were secured along its length 7.5 cm apart. These strips were used to enclose a layer (2–3 mm) of C-free sand (muffle-furnace 800 °C for 4 h). The sand was saturated with nurient solution and allowed to drain by slanting the assembly to 30° above the horizontal. Drainage was facilitated by grooves cut into the lower end of the lid. Shoots of the seedlings were supported in a plastic tube (2 cm length x 2 cm diameter) and held securely with non-absorbent cotton wool. Roots were placed into the sand, and seven glass microscope slides (2.5x7.5 cm) were placed across the area of sand, resting on and secured by the TerostatTM strips. The base of the polystyrene box was placed on top of the lid, with a small opening (2 cm diameter) cut out to allow the extension of the shoot. The whole assembly was supported at 30° above the horizontal to encourage downward extension of the root through the sand and to facilitate drainage. Nutrient additions were applied by injection through the TerostatTM strips with a hypodermic syringe. The plastic and glass components of the assembly were acid-washed to remove residual carbon, and sterilized by immersion in 70% (w/v) ethanol or autoclaving prior to use.
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Nitrate treatments
Microcosms were supplied with nutrient solution consisting of CaCl2 (2 mM), MgSO4 (0.75 mM), NaH2PO4 (0.58 mM), FeNaEDTA (0.045 mM), KCl (2 mM), and micronutrients. The nitrate component of the treatment was supplied in the form of additions of KNO3– to this nutrient solution. Three nitrate treatments were applied to the planted microcosms, with five replicate microcosms for each treatment. Treatment A was a supply of nutrient solution containing 2 mM nitrate for 28 d. Treatments B and C were supplied with 2 mM nitrate for the first 14 d after which they were supplied with either a low (10 mM) or a high (20 mM) boost of NO3– for the following 14 d. Each microcosm was supplied with nutrient solution, at a rate of 2.5 ml d–1, injected directly into the sand layer through the upper TerostatTM wall of the base plate as a single application. The citrate present to solubilize Fe contributed 0.72 mg C l–1, and the nutrient solution drained from the microcosms (in the absence of plants) contained 0.93±0.09 mg C l–1.
Biosensor
The biosensor used was Pseudomonas fluorescens 10586 pUCD607 (lux CDABE from Vibrio fisheri, kanr, ampr) originally described by Amin-Hanjani et al. (1993). The organism had previously been characterized with respect to the bioluminescence response to a range of compounds present in the root exudate, and it was demonstrated that bioluminescence was proportional to substrate availability and use (Yeomans et al., 1999). Carbon starved cells of the biosensor were prepared according to the methods outlined by Yeomans et al. (1999). In brief, P. fluorescens pUCD607 was grown in LB broth, containing kanamycin (Sigma) at 50 µg ml–1 at 25 °C, shaken at 200 rpm. Cells were harvested in the late exponential phase, determined by measurement of the optical density after reference to a standard growth curve, followed by centrifugation (10 min, 3500 g). The supernatant was then decanted and cells resuspended in an equal volume of C-free M9 minimal medium (Difco) also containing 50 µg ml–1 kanamycin. The process was repeated and a final suspension of the centrifuged pellet of washed cells shaken in C-free M9 minimal medium at 200 rpm for 3 h at 25 °C. During this period of C-starvation, the decline in activity in the form of bioluminescence was monitored.
Application of the biosensor
After 28 d growth in the microcosms, starved biosensor inocula were applied to the sand matrices in which the roots were growing. Each microcosm was subdivided into nine zones (2x7 cm) comprising the region beneath each of the nine covering plates. A 0.5 ml aliquot of starved cells (3.25x107 cfu) in M9 medium was applied evenly across the surface of each zone using a pipette.
CCD imaging of plate microcosms
Images of the plate microcosms and of light output from the lux-marked biosensor were captured using a nitrogen-cooled, charged coupled device type camera (type EEV CCD 47–10 grade 1 CCD, Pixcellent Imaging Ltd, Cambridge, UK) sited within a light-tight box. A 50 mm F1.4 Nikon lens was attached to the camera set at an aperture of 2.8. Two sets of images were recorded: digital images of the plate microcosm (bright-field image), and CCD images of light output (during a 5 min exposure) from the biosensor (dark-field image) (PixCel software package, Pixcellent Imaging Ltd, Cambridge, UK). Each CCD image consisted of a large array of pixels (1029x1029). Relative light unit values (RLU) were recorded for each pixel in the array and exported to a spreadsheet, with cell values corresponding to RLU values for each pixel-address. The bright- and dark-field images were stacked to view the spatial distribution of light output of the biosensor in relation to root system architecture (PerkinElmer UltraPlus software package). Total bioluminescence (light output in RLU from all pixels) was recorded from each microcosm on two occasions, 15 min (t15min) after the initial application of starved cells and 24 h (t24h) after application. These digital images were stored for subsequent analysis.
After completing CCD image capture, a fine-tipped Pasteur pipette was used to sample nutrient solution from the sand matrices. Samples were taken from locations along the root axes, branch points and regions in the matrices more than 3 cm from roots. The samples were then spotted onto pH reactive paper, and the pH determined from the subsequent colour development.
Analysis of CCD images
The total bioluminescence (summation of light output (RLU) from all pixels in each array) was determined from the dark-field images recorded from each microcosm as follows. The background signal level detected by the system in the absence of any input signal (the dark current) was determined from readings taken during a 5 min exposure within the light-tight box (dark reference image). This background signal was subtracted from the RLU recorded for, and associated with, each pixel within the digital image. A spreadsheet macro was used to filter the pixel array produced for each image, eliminating all pixels below that of the dark reference image so that only RLU values associated with biosensor activity remained. The total bioluminescence was then calculated by summing the RLU values for the remaining pixels. Total light output at specific sites (e.g. root tips) was determined after light- and dark–field images were overlain to produce a stacked image, combining the root system image with that of the bioluminescence recorded within each microcosm. The total light output (RLU) within a set of quadrats consisting of a grid of pixels (25x25) was calculated. Within each series of images, recorded at both sampling occasions, the total RLU at 150 selected sites (30 sites for each replicate) was determined. These data were used to evaluate treatment effects on the light output from the biosensor associated with identifiable root sites.
Harvest and nitrate analysis
On harvest, 28 d after planting into the microcosms, shoot material was removed and dried at 80 °C for 48 h. Roots were carefully removed from the sand matrix, and stored in 70% ethanol prior to morphological analysis. The shoot material was weighed and then ground (Retsch Mixer Mill type, MM-2) in preparation for NO3– analysis. The ground powder was shaken with 5 ml of deionized water and incubated in a water bath for 1 h at 45 °C (Cataldo et al., 1975). The suspension was then filtered through No. 42 Whatman filter papers and the nitrate concentration of the extract determined by flow injection analysis (FIAstar).
Morphological analysis
Root systems were analysed for morphological parameters using a WinRHIZOTM scanner, (Régent instruments Inc, Quebec). The storage of the root systems in ethanol (>48 h) was sufficient for complete killing of the genetically modified biosensor. Individual root systems were spread out on a clear tray in distilled water and placed on a flat-bed scanner. Analysis of the scanned images provided size-class distributions and quantification of parameters including root length, surface area, average diameter, root volume, and number of tips.
Response of biosensor to carbon substrates and nutrient solutions
To assess the effect of availability of NO3– in the nutrient solution on biosensor responses to C-substrate, bioluminescence in response to a range of glucose concentrations (1–100 mM) was determined in nutrient solutions containing 0, 2, 10, and 20 mM NO3–, respectively. Bioassays were undertaken in opaque micro-titre plates (96 well, flat-bottom, Bibby Sterilin Ltd, Staffordshire, UK). Into each well in the series, a 100 µl aliquot of carbon-starved cells was added to 100 µl of the C-solution (0, 1, 5, 10, 25, 75 or 100 mM glucose) and 0.1 g sterile carbon-free sand. Biosensor bioluminescence in each well was quantified by the determination of mean pixel intensities using the CCD camera, as described previously. Images were captured and recorded over a 24 h period, to provide a time-series of images that could be used to determine temporal changes in metabolic activity of the biosensor.
Statistical analysis.
One-way ANOVAs were used to compare differences in values for biomass (plant, root and shoot), root morphological parameters (root length, specific root length, average diameter, number of tips) and bioluminescence measured between treatments. The Tukey test was used to compare significant differences between the means at the <0.05% level. One-way unstacked ANOVAs were used to compare bioluminescence from root tip sites and data were normalized using Box-Cox transformations where appropriate.
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Effects of treatment on shoot NO3–-N and plant biomass partitioning
The nitrate-boost treatments were found to affect shoot nitrate concentrations strongly (Table 1). Barley seedlings not exposed to a boost of NO3– at day 14 (A), had a very low mean nitrate concentration of 0.004 mg NO3–-N g–1 shoot FW. The two treatments to which a NO3– boost was applied at day 14, developed shoot systems which contained significantly (P <0.05) higher nitrate concentrations on harvest: 0.41 and 1.77 mg NO3–-N g–1 shoot FW for treatments B and C, respectively.
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On a whole plant basis, the two treatments provided with a NO3– boost had significantly (P <0.05) greater dry weight biomass. However, there was no significant difference in root biomass between the treatments. The effects on plant biomass were due to an increase in shoot biomass accumulation with increasing NO3– supply in the boost treatment. This was apparent from the decline in root weight ratio (RWR) (Table 1).
Effect of treatment on root morphology
Despite the lack of treatment effects on root biomass, significant effects of treatments on root morphology were found. Total root length decreased as shoot NO3– concentration increased (Table 2). Analysis of root systems with the WinRHIZOTM system indicated that NO3– exposure differentially affected the size class (root diameter) distributions within root systems. Root systems not provided with a NO3– boost (treatment A), had a greater proportion of their total root length in finer root classes. The proportion of root length less than 0.4 mm in diameter was 69.8%, 28.0% and 17.5% for the zero boost (A), low boost (B) and high boost (C) treatments, respectively. For treatment A, the most frequent size class was 0.2–0.4 mm (60.0%), whereas the modal frequency was shifted to the 0.4–0.6 mm size class for treatments B (34.6%) and C (34.9%). In treatment A, 95.5% of the root length was less than 0.6 mm diameter, whereas 37.5% and 47.6% of root length had diameter above 0.6 mm in treatments B and C, respectively.
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Regression analysis indicated that there was a significant (R2=0.88, P <0.01) negative correlation between shoot nitrate concentration and the number of root tips per root system. The mean number of tips per root system was significantly (F(1,14) 26.3, P <0.05) reduced in the high boost treatment (C), compared with the no boost (A) and low boost treatments (B) (Table 2).
The microcosm design restricted, but did not entirely eliminate, contamination from air-borne bacteria. Contaminant numbers were determined in replicate microcosms (3) of each treatment. No treatment differences in bacterial numbers were found, and in all cases culturable bacterial numbers were less than 102 cells g–1 sand. This level of contamination was insignificant relative to the numbers of biosensor cells applied to the microcosms (approximately 108 cells g–1 sand).
Biosensor reporting of root C-flow
Total bioluminescence from each microcosm was quantified on two occasions after application of the biosensor (Figs 2, 3). Total bioluminescence quantified for each treatment was found to decrease at t15min and t24h with increasing NO3– supply in the boost treatment (Tables 3, 4). In all cases, total bioluminescence declined between t15min and t24h. At t24h, bioluminescence was seen to be more localized around roots (in particular root tips). The only morphological parameter with which there was a significant correlation (R2=0.83, P <0.05) with total bioluminescence at both sampling times (when replicate values were plotted individually) was total root length. Quantification of biosensor bioluminescence associated with root tips indicated that these regions exhibited elevated bioluminescence relative to bioluminescence associated with the root system as a whole. Elevated biosensor responses associated with root tip regions were most evident at t24h (Table 4). The specific bioluminescence for root tip sites was found to vary between treatments (increasing with increasing shoot nitrate concentration) at t15min, but not at t24h.
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The pH of solution within the microcosms, a potentially important determinant of biosensor activity, were found not to be affected by location along root axes, distance from roots or level of nitrate applied in the nutrient solutions. In all cases pH was found to be within the range 5.8 to 6.1, values optimal for biosensor activity.
Effect of NO3– concentration on response of the biosensor to glucose concentration
Nitrate concentration (0, 2, 10, and 20 mM) was found not to significantly affect biosensor response to glucose concentration (0, 1, 5, 10, 25, 75, and 100 mM). In each case, the biosensor response to glucose concentration was linear with the line gradients unaffected by nitrate concentration (P >0.1) (Fig. 4).
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Discussion |
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To test the hypothesis that shoot NO3– concentration, as a systemic indicator of plant N-status (Forde and Lorenzo, 2001), is inversely correlated with quantitative exudation of organic C from roots, it was necessary that the NO3– supply treatments resulted in contrasting shoot NO3– concentrations. This condition was clearly met, with shoot NO3– concentration approaching zero in Treatment A, and reaching values indicating significant storage of excess NO3– in Treatment C (Table 1). The effects of treatment on shoot NO3– concentration were concurrent with shifts in dry matter partitioning (RWR, Table 1), consistent with the hypothesis that shoot NO3– concentration is a primary signal in systemic plant responses to internal N-status (Scheible et al., 1997a, b). However, these effects on RWR were primarily mediated by effects on shoot biomass production, with no significant effect of treatments on root biomass.
Impacts of treatments on roots were apparent in their morphology, but not their architecture (Table 2). With increasing NO3– supply, total root length and number of root tips decreased, while root diameter increased. These responses are consistent with previous studies which demonstrated increased production of fine roots under low N-supply. This is considered to maximize the efficiency of plant resource-use in foraging for scarce nutrients (Fitter, 1985). It is notable that the increased number of root tips with decreasing NO3– supply is directly proportional to the increase in root length (Table 2), indicating that lateral production per unit root length was unaffected by treatment. Therefore, the effects of treatments on root growth in this study are consistent with systemic plant responses to internal N-status, as opposed to responses to localized NO3– additions, which are typically characterized by shifts in frequency of lateral root production (Robinson, 1994).
The biosensor used in this study (P. fluorescens pUCD607) has been characterized with respect to bioluminescence in response to C-substrates present in root exudates (Yeomans et al., 1999). In the present study, a range of amino acids, organic acids and sugars were supplied as substrates to the biosensor, and it was confirmed that bioluminescence was proportional to substrate concentrations (data not shown). Further to this, it was demonstrated that the contrasting nitrate concentrations applied to the plants did not affect significantly the response of the biosensor to glucose concentration (Fig. 4). The lack of dependence on nitrate concentration in this assay supports the view that biosensor bioluminescence is affected only by exudate concentration across treatments. This condition was achieved through the C-starvation of the organism in M9 media prior to application to the microcosms. The lack of nutrient limitation on the biosensor was confirmed after the assay period through the exogenous addition of glucose—bioluminescence was stimulated across all treatments.
Total biosensor bioluminescence per microcosm was greater when determined 15 min after inoculation than after 24 h (Table 3 compared with Table 4). This was observed as more diffuse bioluminescence associated with the roots at t15min (Figs 2A, 3A compared with Figs 2B, 3B). This can be explained as the biosensor response to C-substrates released from roots during the whole growth period at t15min, but after 24 h, when accumulated root exudate substrates have been exhausted, the response is seen to be localized to root surfaces active in the exudation of substrates. Consequently, the determination of biosensor bioluminescence at t24h provided a better assay of current exudation from roots, with respect to NO3– treatment and spatial distribution of exudation intensity for each root system. In contrast to total microcosm-bioluminescence, specific bioluminescence localized to root tips was seen to increase between t15min and t24h. This was likely to have been due to activation of the biosensor (induction of catabolic pathways) and probably also cell growth associated with sites of active exudation. At t24h, total microcosm bioluminescence decreased with increasing NO3– supply to the plants, as was also seen at t15min. This indicates that the total substrate available to the biosensor (on a whole microcosm basis) was greater with reduced NO3– supply. This effect can be directly attributed to greater exudation from the root systems at low NO3– supply, as no significant biosensor activity occurred at sites not associated with roots (Figs 2, 3).
Increased root exudation per unit root biomass at low N-supply has been reported in previous studies under axenic conditions (Bowen, 1969; Xu and Juma, 1994; Hodge et al., 1996; Paterson and Sim, 2000). However, the basis for this response has remained unclear. It can be hypothesized that increased release of organic compounds from roots under low N is due to elevated specific exudation rates of the whole root system, elevated exudation from particular root sites, or a relative increase in root sites that are intrinsically more active with respect to exudation (i.e. shifts in exudation concurrent with shifts in root architecture). Specific bioluminescence of the biosensor was greater in response to root tip regions, with increasing shoot nitrate at t15min. However, this effect was not apparent at t24h. Here, bioluminescence at t15min is in response to the cumulative release of root exudates during previous growth. Consequently, it is likely that increased bioluminescence (t15min) associated with root tips was due to a greater accumulation of exudates per unit area of sand (i.e. a function of lower root extension rates, but equivalent exudation intensity, with increasing shoot nitrate concentrations). The results presented for biosensor bioluminescence at t24h provide a better estimate of current exudation rates, and suggest that exudation per unit root length (whole root and root tip sites) does not increase with decreasing NO3– supply (Table 4). The biosensor did report significantly elevated exudation from root tips, relative to the whole root (Table 4). However, the frequency of root tips did not increase per unit root length, suggesting that increased exudation under low NO3– supply was not mediated by a shift in root architecture. Root tips did contribute to increased total root exudation, but in proportion to the increased length of the root system with low NO3– supply. Consequently, the results indicate that root exudation from barley increases as a function of increased root length, in response to decreasing NO3– supply. This can also be stated as: increased specific root exudation (on a below-ground biomass allocation basis) was a consequence of construction of a root system with increased specific root length. This result provides direct evidence to support previous work (Xu and Juma, 1994; Paterson and Sim, 2000) that had identified a correlation between the quantity of root exudation and total root length, but had been unable to determine that this relationship was causal.
A potential confounding factor in this study is the influence of exudate quality on biosensor activity. The use of a general metabolic reporter, as in this study, has the advantage that it reports on cellular C-flow through cells of the biosensor, independently of substrate form. However, the use of individual substrates is dependent on the physiological capacity of the bacterial species (biosensor) to metabolize these substrates. It would be expected that this capacity would vary both with the chemical form of substrate and the bacterial species used as the biosensor. There are several lines of evidence to suggest that exudate quality did not greatly influence biosensor activity in this study. (1) P. fluorescens is a ubiquitous rhizosphere organism, characterized by its adaptation to the use of a broad range of substrates present in root exudates. Consequently, it is ideally suited to report on changes in flux of these substrates. (2) Yeomans et al. (1999) demonstrated that the biosensor used in this study responded to the major components of root exudate (sugars, amino acids and organic acids), and that the Vmax response was equivalent for sugars and amino acids (lower for organic acids). (3) Previous studies of whole-root exudation in response to N-supply have found changes in the amino acid component only, with increased amino acid exudation at high N supply (Bowen, 1969; Ofosubudu et al., 1995). However, amino acid exudation is a relatively small fraction of total root exudation (1–5%; Lynch and Whipps, 1990) and is unlikely to affect biosensor responses greatly. Approaches to address this factor directly are, (1) to study the Michaelis–Menten kinetics of bioluminescence in response to root exudation in situ through sequential image capture, with reference to Michaelis–Menten parameters for known substrates and (2) through the use of substrate-specific biosensors to identify spatial variation in exudation of single compounds (Jaeger et al., 1999).
In conclusion, this study demonstrated the applicability of biosensor approaches to study C-flow from roots, particularly in the context of separating effects of treatments on root physiological processes and root morphological development. Increased root exudation from barley in response to decreased NO3– supply was found to be a consequence of increased root length, with no effect on specific root exudation rates. This finding highlights the importance of root morphological and architectural responses to environmental factors (as opposed to biomass allocation alone) in mediating exudation inputs to soil.
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Acknowledgements |
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We would like to thank Professor Ken Killham for providing the biosensor, and Allan Sim for technical support. The work was funded by BBSRC and SEERAD.
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References |
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