Variation of isotopic composition of N sources and its effects on N transfer estimates (II, III)

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Nitrogenous root exudates

Root exudation is part of rhizodeposition, which is defined as the release of volatile non-particulate and particulate compounds from living plant roots (Wichern et al. 2008). Exudation of sugars and other carbonaceous compounds is increasingly understood as an important process in regulating the interactions of plants and soil microbial organisms (Jones et al. 2004). Nevertheless, plant roots also exude many nitrogenous compounds such as ammonium, amino acids or even proteins (Sawatsky and Soper 1991), although they have generally received much less attention than sugars. The exuded nitrogenous compounds may contribute to N nutrition of neighbouring plants. Exuded N may be absorbed by the associated plants if the root systems or mycorrhizal fungal symbionts of the plants are in close contact, or after short-distance mass flow of compounds with soil solution to the vicinity of roots (Fig. 1).
Ammonium has been observed to be exuded from living plant roots in significant amounts, and it seems to be the predominant form of exudate N of legumes (Brophy and Heichel 1989, Paynel and Cliquet 2003). From 3 to 23 % of total N in a non-N2-fixing ryegrass (Lolium perenne L.) was derived from the exudation of ammonium and to a lesser extent of amino acids by an associated N2-fixing clover with hydroponics cultivation (Trifolium repens L.; Paynel et al. 2001). Exudation of amino acids from legume roots has been observed in hydroponic cultivation (Ofosu-Budu et al. 1990, Shepherd and Davies 1994) and in solid growth medium (Rovira 1956, Brophy and Heichel 1989, Paynel and Cliquet 2003), but with the exception of Rovira (1956) the quantities have generally been small.
The factors controlling exudation are not well known. Membrane permeability of the solute, concentration differences within root and in soil, and biotic and abiotic stress have been suggested as driving forces (Jones et al. 2004). Jones and Darrah (1994) observed that exudation of amino acids from maize roots occurred by passive diffusion, but that their recapture processes were active. Especially in legumes exudation may act as a regulation mechanism for controlling the concentration of amino acids in cytoplasm and balancing N2 fixation with plant N requirements in the short term. Concentration of amino acids in cytoplasm regulates NH4+ and NO3- transporters (Amtmann and Blatt 2009), and if the concentration increases too much, e.g. as a result of excess N2 fixation, N transport within the root may slow down. Exudation of the excess N in such case could help to restore the N balance within roots. Defoliation of trees under a heavy fodder harvest regime may represent a stress factor which increases exudation. When N balance within a plant changes in pruning, flushes of N to the roots may be expected (Nygren 1995), and in herbaceous plants these have been associated with increased exudation (Eason and Newman 1990, Ayres et al. 2007). Shoot removal increases N transfer from a donor plant, both in mycorrhizal and non-mycorrhizal plants (Ikram et al. 1994, Johansen and Jensen 1996, Mårtensson et al. 1998), which may also imply increased exudation.
Relative competitive abilities between plant species in capturing exuded compounds from the soil solution may determine the net benefits of exudation for a species. Exudation could result in a net flux of N to the soil solution from plants which have low capacity for organic N uptake or low uptake rates (Lipson and Näsholm 2001). These plants would thus provide N to the associated plants that have a higher affinity for organic N. Legumes are generally less effective competitors for soil N than grasses (Ledgard and Steele 1992), and grasses may, thus, benefit from the exudates of legume species.

Direct and indirect N transfer

The underlying reasons for interplant N transfer are not well understood. Direct N transfer may be significant for plant nutrient acquisition especially in systems with high potential for nutrient immobilisation, because it can provide a shortcut to the nutrient mineralisation cycle and reduce competition with soil microbial organisms (cf. Owen and Jones 2001, Jones et al. 2005). Nitrogen transfer could contribute to species diversity and plant density in ecosystems, thus improving the overall resource use and increasing the stability and resilience of the systems. Such impacts have been associated to mycorrhizal fungal symbioses in general, since they are known to be able to alter plant competitive relationships and plant community composition as a consequence (Hart et al. 2003, van der Heijden et al. 2003).
It is important to note that N released by the N donor plant either via exudation or to AM fungal symbionts may also become subject to indirect N transfer, via microbial uptake or turnover of the mycelia which is known to be rapid (Staddon et al. 2003; Fig. 1). Suitable research methods to quantify whether N transfer via root exudation or mycorrhizal symbionts is direct or indirect are lacking, especially for CMN which are microscopic and fragile in nature. Consequently, the evidence of direct transfer of nutrients via these transfer pathways comes from indirect observations. For example, the amount of nutrients that plants are able to capture from the dying roots of associated plants has been shown to increase several-fold when the living and dying roots share or able to share the same mycorrhizal fungal symbionts (Hamel and Smith 1991). Such results also demonstrate the potential significance of direct N transfer.
In this study, direct N transfer is discussed theoretically, while making attempts to create experimental support to its role. Nitrogen transfer, in turn, is used to refer to both direct and indirect N transfer (via the mineralisation and immobilisation cycle in soil), as they are difficult to distinguish in experimental research. This use of the term is consistent with research literature where ‘N transfer’ commonly is used to refer to all N in the N recipient species originating from the N donor, without distinguishing between N mineralisation from organic residue and other N sources (Høgh-Jensen 2006).

The model fodder production system

An experimental fodder production site (agroforestry field site in the following) of the legume tree Gliricidia sepium (Jacq.) Kunth ex Walp (Papilionoideae) and fodder grass Dichanthium aristatum (Poir) C.E. Hubbard (Poaceae) in Guadeloupe, French Antilles, was used as model system for this study. The site is located at the Godet Experimental Station of the French National Institute for Agronomic Research (INRA) (16°25’ N, 61°30’ W, 10 m a.s.l.). The climate in the area is warm and subhumid, the annual mean air temperature being 26 °C and annual mean rainfall 1300 mm. Dry season lasts from February to July, and 30% of the annual rainfall occurs during that time (weather station of INRA situated on the site). The soil on the site is Vertisol with 80% of clay rich in smectite, developed over coral reef limestone and with pH of 7.8. Average soil depth is 0.5 m. Cation exchange capacity of Vertisol is 52 cmol(+) kg-1, with 75% of Ca2+ saturation. Soil aggregates, therefore, are stable, which ensures a good aeration of the soil despite of the high clay concentration.
The agroforestry field site comprises two systems: (i) tree-grass plots where D. aristatum is grown between rows of G. sepium, and (ii) the adjacent grass plots of D. aristatum, where the grass is in contact with G. sepium roots but not with the tree canopies. In addition, (iii) a separate grass monocrop of D. aristatum, located approximately 200 m away from the agroforestry field site, was selected for comparison as a system where the grass has no contact with G. sepium.
The tree-grass plots were established in 1989 by planting cuttings of G. sepium in natural grassland of D. aristatum. The trees were planted in North-South aligned rows at a spacing of 0.3 m x 2 m. The tree-grass plots were 20 x 13 m in size and separated from each other by the grass plots of 15 m of width. At the time of this study the actual tree density of the tree-grass plots was about 12 000 ha-1 because of mortality. The plots were not trenched, allowing contact of G. sepium roots with D. aristatum on the adjacent grass plots. The plots were managed since their establishment according to a cut-and-carry practice, with partial tree pruning every 2-6 months and grass cutting every 40-50 d. Grass on the grass plots and the grass monocrop was managed as on the tree-grass plot. All cut material was removed from the site.
Nitrogen fertilisers are not used on the agroforestry field site, and N2 fixation by G. sepium is the sole N input to the tree-grass plots and the adjacent grass plots. Nitrogen derived from the atmosphere constitutes 60-87% of total N in the above-ground biomass of the tree, depending on the season (Nygren et al. 2000). Phosphorus and K fertilisers were applied five times since the establishment of the site at 100 kg [P] ha-1 and 150 kg [K] ha-1, last time in 2003. Available soil P is lower on the tree-grass plot than on the adjacent grass plot (11.5 mg kg-1 vs. 18.1 mg kg-1; Dulormne 2001), probably because of large biomass exports as tree prunings. Soil organic N content decreases with distance from the tree rows (Sierra and Nygren 2006; Table 2). Fine root densities of the two plant species correlate negatively, with the root density of G. sepium decreasing and that of D. aristatum increasing with distance from the tree rows (Sierra and Nygren 2006; Table 2). Biomass production and N accumulation by the grass are negatively affected by competition from the trees to approximately 3 m from the tree rows (Daudin and Sierra 2008), although tree roots are found throughout the adjacent grass plot.


Identification of mycorrhizal fungal strains from root samples

DNA was extracted from the root samples by CTAB and proteinase K buffer (Timonen et al. 1997) after vigorous pulverisation in liquid nitrogen with pestle and mortar. Nested PCR with outer primers SSUmAf: 5′- TTG GTA ATC TTD TGA AAC TTY-3′ and LSUmAr: 5′- GCT CTT ACT CAA AYC TAT CRA -3′ and inner primers SSUmCf: 5′- TAT TGY TCT TNA ACG AGG AAT G -3′ and LSUmBr: 5′- AA CAC TCG CAY AYA TGT TAG A -3′ were modified from Krüger et al. (2009). The used primers are expected to have fairly good coverage of the most common families of Glomales (Krüger et al. 2009). The PCR was carried out as in Krüger et al. (2009) with the exception of using the annealing temperature of 58°C in the nested PCR. PCR reactions were run into 1% agarose-SYBR Green gel (InvitrogenTM, Life Technologies, NY, USA). The correct-size products were cut cleanly from the gel under black light and frosen at -20°C overnight. The liquid from the gel pieces was centrifuged into eppendorf tubes, precipitated with ethanol and sequenced at Macrogen Inc., Seoul, Republic of Korea. The acquired sequences were manually checked using Geneious Pro 4.5.5 (Biomatters Ltd., Auckland, New Zealand).
Reference sequences were retrieved from the GenBank database and aligned with Geneious Pro 4.5.5. The phylogenetic analyses were performed by WinClada Ver. 1.00.08 (Nixon 2002) according to Timonen and Hurek (2006), with the following exceptions: maximum number of trees to keep [hold] 1000 000, number of replications [mult*N] 5000, and starting trees per replications [hold/] 20. A combined sequence of Mortierella polycephala sequences AB476414 and AF113464 was created to act as a fixed outgroup. The sequences retrieved from the examined roots were deposited to GenBank with the accession numbers FR873155-FR873165.

Visual determination of mycorrhizal colonisation

The sampled root fragments were stained following the method of Phillips and Hayman (1970). The roots were cleared at 2.5% KOH in 85 ˚C water bath for 30 min and rinsed with distilled water. They were then stained in a lactoglycerol mixture (lactic acid, glycerol, and water 1:1:1) with 0.05% w/v Trypan Blue, in 85 ˚C water bath for 25 min. The samples were immersed in lactoglycerol without the stain to remove excess colour, and conserved in a mixture of 50% of glycerol and 50% of water until mounting.
Mycorrhizal colonisation in the roots was visually determined applying the magnified intersections method (McGonigle et al. 1990). Briefly, the roots were mounted directly from the glycerol-water mixture on microscope slides, and examined with 400 magnification at 150 cross-sections per sample. The number of sections where the AM formations arbuscules (including arbusculate coils), vesicles or hyphae were observed was noted separately for each formation type. Dark or septate hyphae were not counted (Jumpponen and Trappe 1998). The results were expressed as the proportion of root length colonised by arbuscules, vesicles, and hyphae, and the proportion of uncolonised root, where no AM formations were encountered.

Analyses of mycorrhizal colonisation, plant biomass and N isotopic composition

At the end of the experiments at week 10, all plants in treatments FI and MY, and D. aristatum in treatments EXMF and RA were harvested by biomass compartment (leaves, branches, stem, and roots of the trees; shoot, stubble, and roots of the grass). The trees were found to be effectively nodulated. It was also verified that the soil had remained well aggregated in the pots, and had not acquired a massive structure that would have restricted its aeration. In order to determine mycorrhizal colonisation within plant roots, 12 x 1 cm fragments of fine roots per pot and per species were sampled, prepared and analysed as described in section 2.2.3. All other plant material, sampled during and at the end of the experiment, was weighed, oven-dried at 70 ºC for 72 h, and ground to <0.2 mm for isotopic analysis. Sample N contents and their isotopic ratios were determined at the Stable Isotope Facility of the University of California-Davis, USA, using an element analyser (PDZ Europa ANCA-GSL) interfaced to an isotope ratio mass spectrometer (PDZ Europa 20-20; Sercon Ltd., Cheshire, UK). Carbon and N content of exudates for calculating their C:N ratio were determined with an element analyser at the Antillean Research Centre of INRA.

Table of contents :

Trees as nitrogen providers in agroforestry systems
Belowground pathways of N transfer
Common mycelial networks
Nitrogenous root exudates
Direct and indirect N transfer
Isotope techniques
Research needs
Objectives of the study
The model fodder production system
Arbuscular mycorrhizal colonisation (I)
Identification of mycorrhizal fungal strains from root samples
Visual determination of mycorrhizal colonisation
Nitrogen transfer pathways (II-IV)
Study design (II-IV)
Root exudation and exudate uptake by grass (III, IV)
Residue decomposition and N uptake by grass (IV)
Analyses of mycorrhizal colonisation, plant biomass and N isotopic composition
Calculations of N transfer
Nitrogen isotopic composition during residue decomposition and its effect on N transfer estimates (II)
Study approach
Model structure
Model parameterisation and simulations
Statistical analyses
Arbuscular mycorrhizal colonisation on the agroforestry field site (I)
Variation of isotopic composition of N sources and its effects on N transfer estimates (II, III)
Nitrogen transfer pathways (II-IV)
Nitrogen transfer in full root interaction and via mycelial networks (III)
Root exudation and exudate N uptake (III, IV)
Nitrogen uptake from decomposing residue (II)
Arbuscular mycorrhizal fungal symbionts on the agroforestry field site (I)
Methodological considerations in quantifying belowground N transfer
Quantifying N transfer with isotope techniques (II, III)
Competition in pot culture studies
Nitrogen transfer pathways (III-IV)
Common mycelial networks (III)
Root exudation (III, IV)
Nitrogen transfer in agroforestry systems
Future prospects


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