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«International 17 Workshop th Nitrogen The was jointly organised by Teagasc and AFBI Printed by Print Depot Suggested citation Authors, 2012. Title ...»

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Can current and emerging molecular methods further expand our understanding of microbial mediation of soil nitrogen cycling? If so, what information can be provided that is relevant to understanding and predicting Nitrogen Workshop 2012 the rates of N-transformations in the field? What molecular information, at what detail and comprehensiveness is necessary or even useful?

2. Linking soil N transformations and microbial ecology This presentation explores a broad but important question: How changing environmental conditions will alter the characteristics and rates of soil nitrogen transformations. Can molecular characterization of soil microbial communities provide information and understanding useful to this exploration?

Environmental Changes Rates of N-processes

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Figure 2. Changes in environmental conditions control process rates directly and through affecting microbial mediation.

Two sets of examples are used to explore if and how information about genes that code for critical N-cycle processes provide information of use. Are there molecular microbial indices that are integrators of past conditions, indices that reflect present potentials, or possibly anticipate future response to changing environments?

The first example will assess the relative values of environmental characteristics and gene abundances for predicting the potential rates of denitrification and nitrification under relatively constant conditions in high latitude ecosystems. Using path analysis to assess the relative predictive value of soil characteristics such as water content, pH, substrate concentration and the values of four potentially relevant genes (coding for nitrite reduction, nitrous oxide reduction, and ammonium oxidation), we find that quantification of gene copies provides the strongest prediction of potential process rates (nitrification and denitrification).

In the second example, molecular analyses provide information about the origin of and mechanisms controlling bursts of N2O produced during wet up of dry soil. Wet up of dry soils (annual grassland soils) resulting from a California dry Mediterranean-type summer, causes rapid resuscitation of indigenous soil bacteria (Placella et al., 2012). The progression of the genetic control of the enzymes of nitrification and denitrification (at the level of m-RNA) can explain the basis of nitrous oxide production during these episodic events.

Environmental control of each microbial N-process occurs at a number of steps comprising the molecular foundation for the process. In a simplified model (Figure 3), occurrence of a process can be controlled by the environment at the levels of: 1) transcription of genes; 2) translation of

Nitrogen Workshop 2012

messenger RNA; 3) activity of enzymes. The net effect of the sum of these controls is the actual rate of the process in the field. Considering the points of control, what are the relationships among genes coding for a functional enzyme, transcripts carrying the information necessary for construction of the enzyme (protein), the suite of proteins comprising the functional enzymes, and finally the rate of the process in the field?

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Figure 3. Simplified model of environmental control of processes: from genes to field assays.

The two examples discussed suggest that molecular characterization of the mRNA transcripts coding for nitrification and denitrification genes can provide information predictive of and explaining the origin of pulsed events such as the production of atmospherically reactive trace gases (nitrous oxide and carbon dioxide) during wet up of dry soil. Different molecular indices (gene copy numbers) can provide predictive information about potential rates under relatively consistent conditions and when differences are relatively large. While gene copy numbers may predict potential rates, they will not provide information on actual process rates. Clearly the best way to determine process rates is to measure them.

Gene copy numbers may however reflect history, possibly providing an index of past process occurrence and hence integrating relevant characteristics of the past environment. Gene copy numbers (DNA) should integrate history of activity and environmental control because this characteristic: 1) results from recent usage; 2) reflects historical adaptation of populations within complex microbial assemblages that include more than 106 taxa spread across broad phylogenetic lineages; 3) incorporates sequestration of inactive cells and immobilized enzymes by soil matrices. If so, how long a time period may be reflected by gene copy numbers? The historical window of time likely reflects the turnover time of the cells or cell-free enzymes and Nitrogen Workshop 2012 the rate of environmental change. While the number of gene copies for N-processing may provide information about past conditions, there has been little work exploring the value of this molecular index for this purpose.

Regulatory networks in bacteria control the response of the organisms to environmental change.

Such regulation is achieved by a network of interactions among diverse array of molecules including DNA, RNA, and proteins. Bacterial regulatory networks result from extended evolution and adaptation of bacterial populations comprising complex soil assemblages. The environmental response characteristics coded in these regulatory networks can show anticipation of future environmental changes that have been repeatedly experienced in the past. Thus characteristics of regulatory networks may reflect historical conditions and also anticipate future environmental change. In photosynthetic bacteria, regulatory networks have been used to understand diurnal temporal patterns of photosynthesis. We present data suggesting that aggregate regulatory networks in soil microbial communities anticipate annual patters of precipitation in Mediterranean-type annual grasslands. Nitrifiers present in extremely dry soil carry transcriptional capacity to activate rapidly – arguably in anticipation of rainfall events (Placella and Firestone, in review).





3. Conclusions Molecular characterization of functional genes and the expression and translation of these genes

may provide useful understanding of:

1. How changing environment regulates rates of nitrogen transformations in soil

2. The basis for potential rates of processes under relatively constant environment.

–  –  –

References Peterson, D.G., Blazewicz, S., Herman, D.J., Firestone, M., Turetsky, M. and Waldrop, M. 2012. Abundance of microbial genes associated with nitrogen cycling as indices of biogeochemical process rates across a vegetation gradient in Alaska. Environmental Microbiol 14, 993-1008 Placella, S., Brodie, E. and Firestone, M. 2012. Rainfall-Induced CO2 Pulses Result from Sequential Resuscitation of Phylogenetically Clustered Microbial Groups. PNAS. (In Review). Placella, S.A. and M.K. Firestone.

Transcriptional Response of Nitrifying Communities to Soil Wet-Up. Environmental Microbiol. (In Review)

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Tracing of N Transformations in Soil and Gas Phases Using Isotopes and FTIR Spectroscopy Kira, O, D. Haroush, R. Tzulker, Y. Dubowski, R. Linker, and Avi Shaviv Division of Environmental, Water and Agricultural Engineering, CEE, Technion-IIT, Haifa

1. Background & Objective Knowledge regarding the pathways involved in N2O production is still limited despite efforts to quantify mechanisms and sources of its formation (dentrification, nitrification, nitrifierdenitrifictaion). Even calculations of gross rates of N-mineralization or nitrification are still limited and depend on complex, time consuming and destructive methods. These are essential for developing better management tools and mitigation measures. Techniques for quantitative investigation of N transformations and N2O source partitioning in soils are based on isotopic enrichment; can be improved by dual-isotope labelling (Baggs, 2008) or alternatively by tracing changes of N2O isotopomers or isotopologues (e.g. Baggs, 2008; Sutka et al., 2006) in the gas phase. These require the use of IRMS to get quantitative results. Yet, IRMS can not be used on line and demands laborious pre-treatment of soil samples. FTIR spectroscopy with the ability to monitor changes in N-gases (using LP, Long-Path gas cells; e.g. Esler et al, 2000) and in soil N-mineral species (using ATR, Attenuated Total Reflectance, e.g. Linker et al., 2006) offers powerful tools for in-situ investigations; particularly when combining smart labelling of 15N/14N and/or 18O/16O allowing direct measurements in the soil phase (Du et al., 2009) or changes in N2O isotopomer concentrations in gas phase (Esler et al., 2000). A new approach used for tracing changes in heterogeneous systems of air pollutants allows in situ investigation of changes in gas-liquid-soil phases (Segal-Rosenheimer and Dubowski, 2007). First efforts for developing a novel method based on FTIR spectroscopy for continuous monitoring of isotopic N-species directly in moist soil and gas phase are presented emphasizing their potential to serve as efficient tools to quantify Ndynamics and N2O source partitioning in complex systems.

2. Materials and Methods

Direct determination of N-isotopic species during soil incubation using FTIR-ATR:

Incubation experiments were performed by adding solutions of 15NH4Cl or 14NH4Cl to vessels containing Terra Rossa covered with a perforated lid, and incubated at 250C for 8 days. Soil were sampled as follows: 10g were mixed at a ratio of 1:1with KCl 1N solutions forming a paste that was placed on a Zinc/Se ATR crystal to obtain MIR spectra to determine 15NH4, 14NH4, 15NO3 & NO3 concentrations using a BRUKER Vector 22 FTIR spectrometer; Afterwards, the pastes were centrifuged, filtered and the clear solutions were placed again on the ATR crystal and MIR spectra taken again. Accordingly, 2 special calibration solutions containing mixtures of all the tested Nspecies were prepared with (i). soil pastes (1:1 KCl) as background or with (ii) the 1:1 KCL solution after filtration. A PLS algorithm was successfully used for the data processing and calibration of the14NH4Cl set. For the 15NH4Cl set a Neural Networks based algorithm was required. Additional samples of ~2 g were extracted at 1:10 ratio of soil:KCl 1N solutions, and used for determining total nitrate + nitrite and ammonium concentrations using an auto analyzer.

Tracing N2O emission from soils using LP-FTIR gas cells:

Saturated soil samples of a Grumosol were placed at the bottom of an LP-FTIR cell. The LP cell was connected to a FTIR spectrometer, allowing continuous collection of MIR spectra during 22 hrs of soil incubations at different condition (aerobic and non-aerobic conditions, with and without acetylene) and 2 soil thicknesses (~ 2 and ~10 mm). Concentrations were determined using the N2O peaks at the range of 2200-2250 cm-1.

N2O emissions, measured with the LP-FTIR system, with a Grumosol were significantly affected by aeration and soil thickness. Under aerobic conditions no N2O was formed, in the 2 mm saturated layer despite the saturation, presumably due to non restriced oxygen supply; yet under non-aerobic conditions ~ 60% of the initial nitrate was transformed to N2O, assumingly via denitrification. In the 10 mm saturated layer, exposed to aeration, ~ 30% of initial mineral-N was lost as N2O, possibly via nitrification and denitrification. In experiments performed with added acetylene, the losses of N2O from non-aerobic saturated 2mm soil layers were about ~40 to 50% larger than without acetylene addition. The results indicate the possibility of effective and fast/on line tracing of concentration changes of isotopic species of mineral-N in soils with no specific sample treatment/preparation. N2O emissions can be directly measured in incubated soils allowing measurement of continuous changes in the gas phase. The encouraging results observed with the separate set-ups indicate the potential for the next phase where a combined system consisting of FTIR-ATR and LPFTIR units will be used for on line measurements of changes in isotopic species both in soil and gas phases.

Yet, special emphasis should be put on efforts of increasing accuracy.

References Baggs E. M. 2008. A review of stable isotope techniques for N2O source partitioning in soils: Recent progress, remaining challenges and future considerations. Rapid Commun. Mass Spectrom 22, 1664-1672.

Du, C., Linker, R., Shaviv, A. and Jianmin, Z. 2009. In Situ evaluation of net nitrification rate in Terra Rossa soil using a Fourier transform infrared attenuated total reflection 15N tracing technique. Appl. Spectrosc. 63, 1168-1173.

Esler, M. B., Griffith, D. W. T., Turatt,i F., Wilson, S. R., Rahn, T. and Zhang, H. 2000. N2O concentration and flux measurements and complete isotopic analysis by FTIR spectroscopy. Chemosphere 2, 445-454.

Linker, R., Michal Weiner, Itzhak Shmulevich and Avi Shaviv. 2006. Nitrate determination in soil pastes using FTIRATR mid-infrared spectroscopy: Improved accuracy via soil identification. Biosystems Engineering 94 (1), 111-118.

Segal-Rosenheimer, M. and Dubowski, Y. 2007. Heterogeneous ozonolysis of cypermethrin using real-time monitoring FTIR techniques. J. Phys. Chem. C 111, 11682-11691.

Sutka, R. L., Ostrom, N. E., Ostrom, P. H. and Breznak, J. A. 2006. Gandhi, H.; Pitt, A. J.; Li, F. Distinguishing nitrous oxide production from nitrification and denitrification on the basis of isotopomer abundances. Appl. Environ.

Microbiol. 72, 638-644.

Nitrogen Workshop 2012

Effects of a nitrification inhibitor on soil nitrogen transformations and N2/N2O emissions after application of slurry to Irish grassland soils -- a microcosm study Ernfors, M.a,b, Brennan, F.a,c, McGeough, K.d, Müller, C.e, Laughlin, R.J.d, Watson, C.J.d, Griffiths, B.F.a, Philippot L.c, Richards, K.G.a a Teagasc, Johnstown Castle, Environmental Research Centre, Co Wexford, Rep. of Ireland b Swedish University of Agricultural sciences, Department of Agrosystems, Box 104, SE-230 53 Alnarp, Sweden c INRA, UMR 1347, Agroecology, 17 rue Sully BP 86510, 21065 Dijon, Cedex, France d Agri-Food and Biosciences Institute, Newforge Lane, Belfast, BT9 5PX, Northern Ireland e Department of Plant Ecology, Justus-Liebig University Giessen, Heinrich-Buff-Ring 26, 35392 Giessen, Germany



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