Spatial decoupling of soil nitrogen cycling in an arid chenopod pattern ground.
Shrub-steppe covers 7.5% (451 000 [km.sup.2]) of Australia's arid and semi-arid rangelands, and is dominated by various annual and perennial species of the family Chenopodiaceae (Leigh 1972). These chenopod rangelands are characterised by patterned ground, an arid-zone landscape feature where the spatial variation of grove (vegetated) and intergrove (bare) areas is rhythmically repeated (Macdonald et al. 1999). Patterned ground occurs wherever rainfall is low (50-700 mm) and has been found on every continent (Valentin et al. 1999).
The pattern-ground chenopod shrubland complexes in Australia are predominantly vegetated by the plant genus Atriplex if the slope is 1-5% (Macdonald et al. 1999). Chenopod pattern ground is most extensive in arid regions of South Australia but also occurs widely in western New South Wales and in south-western Queensland. The patterning is caused by the redistribution of rainfall on the land surface, where runoff from the intergrove areas is concentrated into the downslope grove areas (Valentin and d'Herbes 1999; Valentin et al. 1999) because of gilgai-induced microtopography (Charley and McGarity 1964). The patterning is distinctive, and from the air, the grove-intergrove system looks like tiger stripes (Fig. 1).
The soils of the intergrove and grove areas are chemically distinct, with soluble salts concentrated in the intergrove soils, a result of the limited infiltration and subsequent redistribution of runoff water into the grove soils (Macdonald and Melville 2000). Therefore, sodium and chloride concentration gradients 10-100fold have been built up between the grove and intergrove soils (Macdonald and Melville 1999, 2000; Macdonald et al. 1999). Charley and McGarity (1964) found accumulation of nitrate (N[O.sub.3]) in the intergrove soils of chenopod pattern ground, which may due to redistribution of N[O.sub.3] from the intergrove and/or due to the deposition of solutes or dry matter over time. It has long been recognised that large, near-surface N[O.sub.3] pools, in some cases >104kg N[O.sub.3] [ha.sup.-1], exist in desert environments (Walvoord et al. 2003; Graham et al. 2008). Graham et al. (2008) stated: 'that the soil conditions coincident with desert environments favour the retention and accumulation of N[O.sub.3] delivered by atmospheric deposition or in situ fixation'. Equally, denitrification (nitrous oxide ([N.sub.2]O) reduction to nitrogen gas ([N.sub.2])) losses are assumed to occur (Walvoord et al. 2003) in desert ecosystems but have not been directly quantified (Peterjohn and Schlesinger 1990) and could be a significant source or sink of [N.sub.2]O. In general, microbial denitrification is assumed to be the main biological process of [N.sub.2]O production or consumption in soils (Rudaz et al. 1999; Ruser et al. 2006; Chapuis-Lardy et al. 2007), but evidence is lacking for different soil systems, and other processes may be involved.
This paper investigates the nitrogen (N) and carbon (C) cycles in intergrove and grove soils within the arid zone of Australia. We quantify emission and consumption of [N.sub.2]O and discuss the source of [N.sub.2]O, with a focus on the denitrification process and its controlling factors within a chenopod pattern ground system. We hypothesise that spatial decoupling of the N cycle is exists between the grove and intergrove soils in chenopod pattern systems.
Materials and methods
The chenopod pattern-ground site used for this study is an enclosure, in Hotel Paddock at the University of New South Wales Fowlers Gap Arid Zone Research Station (31.087104[degrees]S, 141.704049[degrees]E). The enclosure is on the foot-slopes to the east of the Barrier Range on an alluvial lobe, with a slope of 1%, within the Gap Hills land system (Mabbutt 1973). The aerial photograph of the site (Fig. 1) shows the clear vegetation patterning, where the bare intergrove areas alternate with the vegetated grove areas.
The vegetation analysis and soil surface mapping determined that the patterned ground is characterised by eight different vegetation units ranging from completely bare stony ground to that dominated by A. vesicaria (Macdonald and Melville 2000). The enclosure is characterised by A. vesicaria and Astrebla pectinata association, which covers 26.2% of the enclosure. Stone litter is not uniform across the site but is concentrated in the intergrove areas across the landscape (Macdonald and Melville 2000). The intergrove areas are saline (3()dS[m.sup.-1]) relative to the grove areas (<0.7 dS[m.sup.-1]), and the hydrological and vegetation patterns control the distribution of soil cations within patterned ground by leaching and vegetation-induced salt turnover (Macdonald and Melville 2000). The soils at this location are extremely stable and the age of the dust mantle at the site is at least 0.9 [+ or -] 0.2 Ma and possibly as old as 1.8 [+ or -] 0.3 Ma (Fisher et al. 2014).
All of the sampling and measurements occurred between 2 and 8 November 2010. This was a wet year, being 280 mm above the long-term average rainfall of 259 mm.
Soil sampling and analysis
Triplicate intact soil cores (0-500 mm) were taken in the centre of both the grove and intergrove soils in line with the soil chamber placements (Fig. 1). Each core was divided in the field into the following discrete depths: 0-25, 25-50, 50-75, 75-100, 100-150, 150-200, 200-300, and 300-400 mm. Samples were placed in sealed plastic bags and stored at ~3[degrees]C until analysis. Once at the laboratory, a subsample was taken and used to determine soil N[O.sub.3.sup.-] and ammonium (N[H.sub.4.sup.+]) content using 1 :5 soil :2m KC1 extract. Concentrations of N[O.sub.3] and N[H.sub.4.sup.+] were determined by the cadmium reduction and phenate method (Rice et al 2012), respectively, by using an Alpkem Autoanalyzer (OI Analytical, Texas, USA). The remaining soil samples were oven-dried and the gravimetric moisture content was determined. A subsample was collected and finely ground using a puck-mill, and the total C and N and [delta][sup.15]N and [[delta].sup.13]C were determined using a PDZ Europa 20/20 stable isotope mass spectrometer (Sercon Ltd, Crewe, UK) with an ANCA solid liquid preparation module. TNrcs equates to the residual soil N and it was calculated by subtracting the mineral N from the total N concentration.
Samples for bulk density measurements were collected in brass cores (75 mm diameter by 100 mm length) to a depth of 400 mm at three locations within grove and intergrove soils. Bulk density samples were scaled in double plastic bags and stored at ~3[degrees]C until analysis. Once at the laboratory, the bulk density of the samples was determined following the protocol of McKenzie et al. (2002).
Plant sampling and analysis
Atriplex vesicaria (n = 7) was collected from the grove soils and Sclerolaena brachyptera (n = l) from both the grove and intergrove sites. Aboveground biomass was collected, placed in paper bags and oven-dried in the laboratory. Prior to grinding in a puck mill, any mineral soil was removed from the aboveground biomass. A subsample was collected, and total C and N and [delta][sup.15]N and [delta][sup.13]C were determined using the same instrumentation as for the soils (above).
Field nitrous oxide, carbon dioxide and methane measurements
Measurements of [N.sub.2]O, C[O.sub.2] and C[H.sub.4] were made using 12 static chambers (mean headspace volume 1.69 [+ or -] 0.02 L) located in the grove and intergrove areas. The chambers were installed on 2 November and measurements commenced on 3 November and continued until 8 November. Emission measurements were made over a 1-h period at 07:00; 11:00; 13:30 and 20:00 each day. Gas samples were collected using a syringe and injected into an evacuated Labco Exetainer (Labco, Lampeter, UK), and were analysed on a Shimadzu GC-2014 gas chromatograph (Shimadzu Corp., Kyoto, Japan) fitted with an ECD and an FID.
Factors affecting nitrous oxide consumption and production
A laboratory soil-incubation experiment was undertaken to assess factors that influence denitrification and consumption of [N.sub.2]O. Intact cores (n=12; 200mm diameter by 100mm depth) were collected for the grove and intergrove soils and stored at ambient temperature in the laboratory. Samples were collected from the 0-50 mm depth and gently homogenised, and 35 g was placed in 120-mL jars. Prior to the experiment, each soil received 7 mL of distilled water; the jars were not closed and were then pre-incubated at 25[degrees]C. After 48 h, the jars + soils were reweighed and deionised water was added so that the soil in all the jars had the same water content. One subset of jars was sealed with rubber septa and gas-tight crimp seals and the other subset was autoclaved before being sealed.
After sealing 6mL of 0.01% [N.sub.2]O was injected into the headspace of the jars used for the elevated [N.sub.2]O experiment and remainder were injected with 6mL of ultrapure He. Because both available C and N[O.sub.3.sup.-] are known to affect [N.sub.2]O flux, two sets of soils jars (35 g) were incubated at 35[degrees]C with either ambient or elevated (0.01%) [N.sub.2]O concentration and were amended with N[O.sub.3] (2mL of 10 g KN[O.sub.3] [L.sup.-1]), glucose (2mL of 20g D-glucose [L.sup.-1]), a combination (2mL of 10 g KN[O.sub.3] + 20 g D-glucose [L.sup.-1]) or deionised water as a control. The different amendments were injected into the jars and then the jars were placed directly in the incubator.
Headspace gas samples (6 mL) were removed with a gas-tight polypropylene syringe at 1, 10, 18, 97, 145 h after [N.sub.2]O injection into pre-evacuated Labco Exetainers. An equal volume of helium gas was then injected into the headspace to maintain a similar gas pressure, resulting in a final 7% dilution of the headspace. The C[O.sub.2], C[H.sub.4] and [N.sub.2]O concentrations were determined by a Shimadzu GC-2014.
Differences between the grove and intergrove soil properties, and greenhouse-gas emissions, were determined using paired r-test analysis. All statistical analyses including descriptive statistics were undertaken using SPSS Statistics version 21.0 (IBM, Armonk, NY, USA).
Soil nitrogen and ,SN natural abundance
The distribution of N[O.sub.3] and residual N was not uniform across the pattern landscape (Fig. 2). The distribution of N[O.sub.3] was significantly different at each measured depth in the soil profile except in the top 25 mm (0.06 g [m.sup.-2]). Nitrate was concentrated in the intergrove area compared with the grove area. Residual N was significantly greater in the surface soil of the grove soil (0-50 mm) than the intergrove. However, at 100-300 mm, residual N was greater in the intergrove soils than the grove soils. The distribution of the N[H.sub.4.sup.+] was not significantly different between the grove and intergrove soils.
In total, 136 kg N[O.sub.3.sup.-]-N [ha.sup.-1] was stored in the top 400 mm of the grove soils, which was significantly greater than the concentration of N[O.sub.3.sup.-]] in the intergrove soils (Table 1). There was no difference in the distribution of N[H.sub.4.sup.+] (32 kg N[H.sub.4.sup.+]-N [ha.sup.-1]) or residual N (2.11N [ha.sup.-1]) within chenopod pattern ground (Table 1). The grove soils were significantly enriched in [sup.15]N at 75-150 mm and at 300-400 mm relative to the intergrove soils (Fig. 2, 10.3-10.6 [delta][sup.15]N).
Soil carbon and [sup.13]C natural abundance
The distribution of total soil C was significantly greater in the surface soil (0-50 mm) and at 100-150 mm in the grove relative to the intergrove. Overall, total soil C (26.3 t C [ha.sup.-3]) increased in concentration down to 400 mm (Fig. 3). There was no difference in the total amount of C stored in 0-400 mm of soil in the grove and intergrove soils. There was no significant difference in the [delta] [sup.13]C distribution between grove and intergrove soil. The overall trend was enrichment to a depth of 400 mm (Fig. 3).
At the time of sampling, there was significantly more soil water (3.11 [ha.sup.-3]) in the grove soils than the intergrove soils (Table 1).
Plant isotopic signatures
The [delta][sup.15]N natural abundance of the S. brachyptera (n = l) was significantly different between the grove and intergrove (Table 2), but grove and intergrove vegetation exhibited a similar [delta][sup.13]C signature. The S. brachyptera in the grove was enriched in [delta][sup.15]N and had similar [delta][sup.15]N content to A. vesicaria. Atriplex vesicaria is a CAM (crassulacean acid metabolism) plant and is therefore more enriched in [sup.13]C than S. brachyptera.
Nitrous oxide carbon dioxide and methane measurements
There was a significant difference in the average [N.sub.2]O, C[O.sub.2], C[H.sub.4] emission between the grove and intergrove areas. Nitrous oxide was consumed from the atmosphere in the intergrove areas (-0.1334 [+ or -] 0.0587 ng [N.sub.2]O-N[m.sup.-2][s.sup.-1]) and emitted from the grove areas (0.26[+ or -]0.12 ng [N.sub.2]O-N [m.sup.-2][s.sup.-1]) (Fig. 4). The emission of C[O.sub.2] from the grove area was ~31 777.71 [+ or -] 3052.54 ng C[O.sub.2]-C [m.sup.-2][s.sup.-1] and two orders of magnitudes greater than the release of C[O.sub.2] from the intergrove soils. Intergrove and grove areas consumed -0.55 [+ or -] 0.08 and 2.92[+ or -]0.12ng C[H.sub.4]-C[m.sup.-2][s.sup.-1], respectively (Fig. 4). Laboratory studies revealed no significant difference between [N.sub.2]O consumption rates of autoclaved and non-autoclaved intergrove soils exposed to an elevated [N.sub.2]O atmosphere. Furthermore, addition of glucose did not change the [N.sub.2]O consumption significantly (Fig. 5). The consumption rates of [N.sub.2]O from the laboratory incubations range from 0.3 [+ or -]0.1 to 0.7 [+ or -] 0.2 pg N [h.sup.-1] [g.sup.-1] within the first 17 h of sampling.
Factors controlling [N.sub.2]O emission
Incubation of intergrove soils with different controlling factors for denitrification, including amendments with glucose, N[O.sub.3.sup.-] and glucose+ N[O.sub.3.sup.-], showed no significant differences. The average denitrification rate (DR) of the treated intergrove soils was 24[+ or -]5ng N [h.sup.-1] [g.sup.-1] and the unamended intergrove soil showed an average DR of 9[+ or -] 1 ng N [h.sup.-1] [g.sup.-1]. The DR of unamended grove soils was 10 times greater than that of intergrove soil. Amendment with C and C + N showed a trend of increasing DR (up to 0.56 [+ or -]0.001 [micro]g N [h.sup.-1] [g.sup.-1]). However, this trend was not significant. Amendment with only NO, to grove soils showed no trend (Fig. 6).
Nitrogen cycling and landscape function
There was a clear partitioning of the N cycle within the patterned landscape at this site (Table 1, Fig. 2). The grove areas, relative to the intergrove areas, do not have significant amounts of soil N[O.sub.3.sup.-]-N and N[H.sub.4.sup.+]-N in the profile (Table 1). The soils of the bare and vegetated areas in patterned ground have different properties and many researchers (e.g. Greig-Smith 1979) have attributed the differences to the presence of vegetation and differential penetration of water. The vegetation pattern of the chenopod shrub land pattern ground is due to the runoff of rainwater from the bare, impermeable intergrove soils to the vegetated groves (Hunter and Melville 1994; Dunkerley and Brown 1995; Macdonald et al. 1999). The redistribution of rainfall is evident in the measured water mass in each area Table 1. Charley and McGarity (1964) found that N[O.sub.3.sup.-] and salinity gradients run parallel to each other within chenopod pattern ground, such that the levels of N[O.sub.3.sup.-]-N is greatest in the saline bare areas. A similar partitioning within the patterned landscape is evident at this site. The intergrove areas do not have significant amounts of soil N[O.sub.3]-N and significantly less N[H.sub.4.sup.+]-N in the profile relative to the grove areas (Table 1). The soil mantle at the site is very stable and ancient, being laid down 0.7 million years ago (Fisher et al. 2014). If the soil accumulation rate was constant, the intergrove soils have been storing 0.2 g N[O.sub.3]-N [ha.sup.-3] [year.sup.-1] through fixation and from dry and wet atmospheric deposition.
The soil N cycle in the intergrove is tightly coupled to the plant primary production. The mean N content of the dry biomass of an A. vesicaria plant is 12.5 g [kg.sup.-1] (Table 2), using the relationship between plant biomass and area, which equates to 215 kg N [ha.sup.-1]. This would explain the low concentrations of soil N[O.sub.3.sup.-] in the intergrove soil because most available N has already been taken up by the vegetation, depleting the N pools within the grove areas. This is reflected in the reduced total residual N concentration (Fig. 2).
The distribution of [sup.15]N in the pattern ground system (Fig. 2) indicates that grove soils lose N via plant uptake and denitrification, which results in isotopic [sup.15]N enrichment relative to the intergrove. Further, biological [N.sub.2] fixation is known to occur in desert environments (Billings et al. 2003), and this appears to be occurring in the pattern-ground systems. Plants that were growing in the grove had double the [delta][sup.15]N of the S. brachyptera growing in the intergrove (Table 2); S. brachyptera is a shallow-rooted plant and the rooting system is confined to the 0.05 m depth of the soil profile. The sampled plants within the grove and the intergrove were of the same size, but some differences do exist in the soil profiles. The effect of biological [N.sub.2] fixation was not detected in the soil 15N pool (Fig. 2).
The soils of both grove and intergrove are characterised by mosses and lichens, which become active after rainfall, and cyanobacteria colonies are present under the quartz stones. At the site and within many chenopod pattern-ground complexes, the majority of the quartz stones and, hence, the cyanobacteria colonies are concentrated in the intergrove areas. The crusts and cyanobacteria colonies are known to fix [N.sub.2] into soil (Billings et al. 2003; Abed et al. 2010) and they would be an important source of N[O.sub.3.sup.-] in the grove and intergrove areas. The 513C signature of the surface soil (Fig. 3), which comprised the crust, indicated that in both areas, lichens and mosses dominate (Billings et al. 2003). The [sup.13]C enrichment and total C content (Fig. 3) at depth does not reflect cyanobacteria inputs but rather diagenetic calcite formation.
Denitrification and [N.sub.2]O emission dominated the grove soils during the period of field measurement (Fig. 4). The [N.sub.2]O emissions from these grove soils were relatively high and in the lower range of [N.sub.2]O emission studies in wetlands (c.g. Teiter and Mander 2005). Generally, denitrification is controlled by several environmental factors including temperature, pH, concentration of C, oxygen, N[O.sub.3.sup.-] , nitrite (N[O.sub.2.sup.-]) and sulfide (Firestone and Davidson 1989; Seitzinger et al. 2006). Our laboratory incubations showed a trend that the denitrification process is C-limited rather than [O.sub.3.sup.-] limited in grove soils (Fig. 6), which is usual in soils with sufficient [O.sub.3.sup.-] load (Knowles 1982; Reddy et al. 1982); however, this trend was not significant. Consumption of [N.sub.2]O was observed from the intergrove soils (Fig. 4). The observed consumption rate of 0.2 ng [N.sub.2]O-N [m.sup.-2] [s.sup.-1] is in the midrange of data from [N.sub.2]O consumption studies (Wu et al. 2013). However, Wu et al. (2013) reported an [N.sub.2]O consumption rate from another desert soil (~317 ng [N.sub.2]O [m.sup.-2] [s.sup.-1]) that was magnitudes greater than the [N.sub.2]O emission observed in this study. Generally, [N.sub.2]O reduction to [N.sub.2] by denitrifiers is regarded as the main biological [N.sub.2]O consumption process in soils (Rudaz et al. 1999; Ruser et al. 2006; Chapuis-Lardy et al. 2007). Vieten et al. (2008) proposed direct [N.sub.2]O fixation to N[H.sub.4.sup.+], which would be energetically more favourable than [N.sub.2] consumption. The uptake of [N.sub.2]O would be additional to the uptake of [N.sub.2] as proposed by Walvoord et al. (2003) for desert ecosystems. Additionally, diffusion and heat convection of microbially produced and atmospheric [N.sub.2]O in deeper soil systems, as proposed by Clough et al. (2005), could contribute to the consumption of [N.sub.2]O in semi-arid soils or the consumption of [N.sub.2]O by soil water (Weiss and Price 1980). Our laboratory experiments show that the [N.sub.2]O consumption in the intergrove soil was abiotic (Fig. 5). Autoclaved and biologically active intergrove soils showed no significant differences in [N.sub.2]O consumption when exposed to an elevated [N.sub.2]O atmosphere. Furthermore, the addition of C to enhance potential microbial [N.sub.2]O consumption within the [N.sub.2]O-elevated flasks had no effect (Fig. 5). These results are supported by our observation that denitrifying activity was not stimulated by any of our laboratory treatments in the intergrove soils (Fig. 6). Addition of C or C + N[O.sub.3] to intergrove soil in ambient atmosphere enhanced bacterial activity as seen by an increase in C[O.sub.2] emissions (data not shown), but only increased denitrification rate slightly. Additionally, the [delta] [sup.815]N of plant material and topsoil from intergrove area was significantly depleted relative to the [delta][sup.15]N of plant material and topsoil from grove soils (Table 2, Fig. 2), which is likely due to lower denitrification rates in intergrove soils. Denitrification discriminates against the heavier [sup.15]N atom, and thus enriches the soil with [sup.15]N, which is assimilated by the plants. Therefore, we assume that the intergrove soils arc limited in denitrifying bacterial activity, which is unusual because up to 10% of ubiquitous soil bacteria are able to denitrify (Schmider and Ottow 1984). It is likely that intergrove soils show a low diversity of microorganisms and a low bacterial density; however, further microbial community analyses are required to verify this presumption.
Although the intergrove and grove soils are rhythmically repeated in arid soils, we could show that both geochemical cycles C and N are distinct between intergrove and grove areas within the patterned ground; hence, the cycles are decoupled from each soil.
The C[O.sub.2] emission from intergrove soils, compared with grove soils, is two orders of magnitudes lower (Fig. 4). This is likely due to a substantially downsized bacterial density in intergrove soils and lower total C content in intergrove soils (Fig. 3). Both soils show, on average, more methanotrophic than methanogcnic activity, which resulted in C[H.sub.4] consumption. It is likely that the ~5 times greater C[H.sub.4] consumption in grove soils compared with intergrove soils is due to a greater bacterial density in the grove soil. Furthermore, differences in the bacterial density between intergrove and grove soils could explain the differences in [N.sub.2]O emissions of these soils. Abiotic [N.sub.2]O consumption is overwhelmed by microbial [N.sub.2]O production in grove soils, whereas abiotic [N.sub.2]O uptake in intergrove soils was almost the sole fate of [N.sub.2]O from the atmosphere and the atmosphere is the sole source of [N.sub.2]O in intergrove soils.
Intergrove and grove soils showed distinct differences in key elements of C and N cycling. Intergrove soils consume [N.sub.2]O because of abiotic processes, whereas microbial denitrification in grove soils leads to emissions of [N.sub.2]O. Additionally, key N cycle elements in grove soils were closely linked to the N uptake. The C[O.sub.2] emission and C[H.sub.4] consumption was low in intergrove soils relative to grove soils, presumably resulting from the low quantity of bacteria in intergrove soils.
Funding was provided by CSIRO Land and Water Capability Grant. BCTM and MF were supported by the CSIRO OCE Julius Career Award.
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B. C. T. Macdonald (A,C), S. Warnckc (A), E. Maison (A), G. McLachlan (A), and M. Farrell (B)
(A) CSIRO Agriculture Flagship, Black Mountain, ACT 2600, Australia.
(B) CSIRO Agriculture Flagship, Waite Campus, SA 5064, Australia.
Corresponding author. Email: email@example.com
Table 1. Average soil nitrogen, carbon and water contents of the grove and intergrove soils ** P<0.01, *** P<0.001 Intergrove Grove Significance Mean s.e. Mean s.e. kg N [ha.sup.-1] Nitrate 136.0 6.8 0.6 0.2 *** Ammonium 42.0 9.2 22.9 5.8 t [ha.sup.-1] Total carbon 24.9 2.6 27.7 1.8 Water 4.1 0.6 7.2 0.6 ** Residual nitrogen 2.1 0.1 2.1 0.1 Table 2. [[delta].sup.15]N and [[delta].sup.13]C for main plants within the Hotel Paddock site enclosure Location Plant [[delta].sup.15]N Grove Atriplex vesicaria 7.84 Sclerolaena brachyptera 8.40 Intergrove Sclerolaena brachyptera 4.06 Location Plant s.e. [[delta].sup.15]N Grove Atriplex vesicaria 0.19 Sclerolaena brachyptera 0.08 Intergrove Sclerolaena brachyptera 0.19 Location Plant [[delta].sup.13]C Grove Atriplex vesicaria 14.09 Sclerolaena brachyptera -26.15 Intergrove Sclerolaena brachyptera -24.19 Location Plant s.e. [[delta].sup.13]C Grove Atriplex vesicaria 0.04 Sclerolaena brachyptera 0.07 Intergrove Sclerolaena brachyptera 0.06
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|Author:||Macdonald, B.C.T.; Warnckc, S.; Maison, E.; McLachlan, G.; Farrell, M.|
|Date:||Feb 1, 2015|
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