Reviewers: Berauer B3, Verbruggen E4,
Measurement unit: μg N or P g-1 soil (or g m-2; ug N g-1 SOM); Measurement scale: plot; Equipment costs: None – €; Running cost: € – €€; Installation effort: low; Maintenance effort: medium; Knowledge need: medium to high; Measurement mode: manual
Nutrients are essential for plant growth. Mineralisation is the conversion of dead organic matter (OM) to inorganic forms of, for example, N, P, K, and Ca by soil microbes. Consequently, mineralisation is the process responsible for making organic compounds re-available for plant uptake, and thus the rate of mineralisation is a critical component of ecosystem productivity. A measurement of soil nutrient pools merely supplies a snapshot of the available nutrient pool at a given time; it does not provide information on nutrient flow-rates from OM to microorganisms and plants.
It is difficult to quantify gross mineralisation rates directly in soils, as soil microbes and plants may take up a substantial part of the mineralised nutrients immediately after nutrients become available within the soil. Consequently, we focus on net mineralisation methodologies in this chapter.
Net mineralisation is the balance between a) gross mineralisation of organic matter to inorganic mineral forms and b) immobilisation of mineral nutrient forms. Immobilisation is the inverse process of mineralisation, where soil microbes take up inorganic nutrients and incorporate them into microbial biomass, making those nutrients unavailable to plants. However, immobilisation can also be physical, for example when nutrients form in mineral complexes within the soil.
Climate change directly affects microbial activity through changes in soil temperature and moisture regimes and thus induces changes in microbial decomposition activities and availability of soil nutrients for plants. Therefore, measurements of net mineralisation rates are critical for linking soil–plant interactions in climate- or global-change manipulation experiments as well as in non-manipulated natural experiments across space (i.e. along gradients) and time (i.e. observational studies).
18.104.22.168 What and how to measure?
Basically, net nutrient mineralisation is the difference in nutrients in the soil or on an ion exchange membrane at two time points. There are different methods available for estimating mineralisation but none of them are optimal (see Table 22.214.171.124). Most commonly used include the buried bag technique, ion resin exchange, and pool dilution.
Table 126.96.36.199. Comparison of three methods for estimating nutrient mineralisation.
|Buried bags||Two soil cores are sampled. One goes to the laboratory for analysis of initial nutrient content. The other is incubated in the soil in a plastic bag and nutrients in the core are measured after incubation. Mineralisation is the difference in nutrient content before and after incubation.||Cheap and easy to use. The method only takes fluctuation in soil temperature into account but soil moisture is kept constant during incubation. Microbial immobilisation of nutrients in the bag may mask the results. Mineralisation is uncoupled from plant processes.||Two soil cores per mineralisation period.|
|Ion exchange membranes||Membranes for absorption of anions and cations, respectively, are inserted in slits in the soil made by a knife.||All the major nutrient elements can be extracted simultaneously with one extraction. Includes spatiotemporal variation in temperature and soil water.
Removal of nutrients from the resins has been reported from high pH soils.
|Less destructive. The same slit can be used for sequential incubation and harvesting of membranes.|
|15N pool dilution||15N pulse is added to a soil as 15NH4, 15NO3, or other N forms. The excess 15N to the natural abundance is followed in different pools over time by soil sampling. The dilution of 15N mirrors the mineralisation and release of nitrogen.||The pool dilution method gives a more mechanistic understanding of the underlying processes as the fate of a cohort of 15N can be followed over time. Time-consuming and more expensive analysis.||At least 3 soil cores at three different times: T0, T1, and T2.|
Buried bag method
Net mineralisation rates of, for example, N and P can be measured in the field by the buried bag method (Eno, 1960). Net mineralisation measured in buried bags is an estimate of the balance between mineralisation and microbial immobilisation of nutrients in the absence of plant roots as the technique prevents plant uptake of mineralised nutrients, but allows uptake by microorganisms. The method accounts for the differences and fluctuations of in situ soil temperatures during the incubation period, while the water content is kept constant inside the bag. Thus, one should note that the net mineralisation rates obtained via this method incur destructive soil sampling and do not account for plant interactions, leaching processes, and lateral flow.
Duplicate soil cores are sampled per plot or across a site per mineralisation period. One intact core is carefully placed in a sealed low-density polyethylene bag (commonly available as lunch bags in supermarkets), making sure that the core does not break, and the bag is then placed back into the ground for a period of weeks to months, depending on the season and research objectives. The remaining soil core is transferred to the laboratory where inorganic N (NH4+ and NO3–) and P (PO4–) are measured following extraction (see section 188.8.131.52 in protocol 2.2.1 Soil microbial biomass – C, N, and P for handling of soil samples and extraction). After the desired incubation period has passed, the bag is retrieved from the ground and inorganic nutrients are extracted in the same way as the initial soil core. The net mineralisation rate is the difference in inorganic N (NH4-N, NO3-N) in the non-incubated soil core relative to the bag-incubated core, and it is expressed per unit time (e.g. μg NH4-N month-1). Seasonal and annual mineralisation rates can be estimated from sequential incubations running over a growing season or a full year.
In nutrient-poor soils, the microorganisms may immobilise a substantial amount of the mineralised nutrients in the bags (Schmidt et al., 2002). Therefore, including measurements of microbial immobilisation, i.e. microbial biomass changes, within the incubated bags enables a better estimate of total mineralisation. However, it is also a substantial amount of extra work. This can be done by measurement of microbial biomass C, N, and P before and after incubation. For more details see Schmidt et al. (1999, 2002) and protocol 2.1.1 Aboveground plant biomass and 2.2.1 Soil microbial biomass – C, N and P.
Ion exchange resin membranes (IEMs)
Plant-available soil nutrients can be estimated using ion exchange membranes (IEMs). Previously, ion exchange resin beads were used to adsorb soil solution cations and anions, simulating plant-root surface properties with adsorption of nutrients directly from the soil (Giblin et al., 1994). However, using resin beads is destructive, similar to the buried bag method, and resin bead measurements do not always correlate well with other mineralisation measurements (Giblin et al., 1994). Recently, ion-exchange membranes (IEMs) have emerged as the preferred method for estimating mineralisation: its 2-dimensional structure imposes minimal disturbance to plots and soil, while allowing for full membrane contact with the surrounding soil medium (Harrison & Maynard, 2014). Consequently, strong relationships have been reported for soil N availability obtained using IEMs and classic soil extraction and mineralisation methods (Duran et al., 2013). Furthermore, IEMs are relatively easy to prepare in large numbers, can be cut to any size, and they are cost-effective to produce in bulk, thus making it possible to cover a large area in situ and perform long-term continuous mineralisation measurements by repeated insertion within the same micro-site (Harrison & Maynard, 2014).
IEMs work much like their resin bead counterparts (Qian & Schoenau, 2002). Large anion and cation membrane sheets are commercially available as they are commonly used in water treatment facilities. Once cut to the desired size, anion and cation strips are kept separate and charged in an appropriate solution prior to insertion. Typically, cation strips are charged with 0.5 M HCl and anion strips are charged with 0.5 M NaHCO3 (Lajtha et al., 1999). After charging, the IEM strips should be rinsed with H2O to remove any excess chemical from the IEM surface. To maximise soil contact, the IEM strips are inserted into the ground at a slight 15–30 ᵒ angle (Lajtha et al., 1999). Note that the ion-exchange method is sensitive to moving soil water, as nutrients are adsorbed from passing soil water (Binkley, 1984).
Depending on the research objective and ecosystem type, the IEMs can be inserted for days, weeks, or months before retrieval, but note that Lajtha et al. (1999) recommends relatively short incubation times in order to minimise potential desorption occurring if the IEM nutrient concentration becomes greater than the soil surroundings. This can take weeks during summer but several months during winter. This reverse gradient phenomenon has previously been observed for ion-exchange resin beads, suggesting either microbial removal of adsorbed nutrients or a reverse gradient caused by prolonged IEM immobilisation (Giblin et al., 1994). If longer-term mineralisation rates are desired, sequential insertion of IEM strips into the same slit in the ground is possible.
Once retrieved from the field, IEM strips should be kept cool or frozen before desorbing anions and cations by extraction in the lab.
Plant root simulator (PRS) probes are an alternative commercial option to custom-made IEM strips. PRS probes have similar properties to custom IEMs, and they are usually bought with a prepaid nutrient extraction from the manufacturer, omitting the labwork component for the user. Similar to custom IEMs, PRS probes consist of paired negatively and positively charged IEM strips that are inserted into the soil. Once retrieved, the probes are sent to a laboratory where a large range of nutrients can be measured.
Isotopic tracer – 15N fate or pool dilution method
It is possible to determine gross N transformation rates by applying a trace amount of inorganic 15N to a soil using the pool dilution method (Davidson et al., 1991; Murphy et al., 2003; Poertl et al., 2007). Based on a time series of the isotopic signature of the different soil N pools over time, gross mineralisation can be calculated using either an equation related to the decline of the tracer (Kirkham & Bartholomew, 1954, 1955) or with a simulation model (Wessel & Tietema, 1992; Mary et al., 1998).
The knowledge on gross transformation rates using the pool dilution technique gives a better mechanistic understanding of the underlying processes (Verchot et al., 2001). However, the method is very time-consuming and application throughout a year for annual estimates of nutrient mineralisation has never, to our knowledge, been applied.
Where to start
For details on the classic buried bag method, see Eno (1960). For a comprehensive review on ion-exchange techniques, see Qian & Schoenau (2002), and for modern IEM-specific methodologies, see Duran et al. (2013) or Lajtha et al. (1999). Murphy et al. (2003) provide details on the 15N pool dilution method and a good overview on the N transformation processes and key references.
184.108.40.206 Special cases, emerging issues, and challenges
Comparison of buried bag and IEM methods
The different approaches for estimating net mineralisation rates in soil have their own pros and cons – none of the methods are generally considered optimal (Table 220.127.116.11; Binkley, 1984; Binkley et al., 1986). IEM strips are inserted into the ground so that they are in direct contact with the surrounding soil environment. This is in contrast to the buried bag method, where the incubated soil is isolated from any fluctuations in water content, precipitation, or movement of soil water. Additionally, the buried bag method disregards root activity, whereas IEMs are inserted into the soil matrix where nutrient sorption “competes” with roots and microbes alike. In contrast to IEMs, the buried bag method can be used to estimate and account for microbial immobilisation of nutrients. Furthermore, the buried bag method is not subject to the reverse-gradient bias, described above, which may occur during prolonged IEM incubation times. Compared to the buried bag and ion-exchange resin bead methods, IEM strips are minimally destructive, and can be inserted repeatedly at the microsite level. The buried bag method requires duplicate cores to be compared, which in heterogeneous soil environments can add potential noise to the data. In contrast, when dealing with small-scale heterogeneity, multiple small IEM strips can be relatively easily inserted across plots or sites to account for spatial variability.
The methods described here were originally developed for measuring inorganic nutrients. However, experiments have demonstrated that many plants are able to take up amino acids (Kielland, 1994; Näsholm et al., 1998; Nordin et al., 2004; Schimel & Bennett, 2004), and ericaceous species can even use more complex N-containing compounds due to their mycorrhizal association (Read & Perez‐Moreno, 2003). Therefore, quantifying amino acid availability or flux-rates may be desired. The buried bag method allows for extraction and quantification of dissolved organic N (DON), which is total dissolved N minus inorganic N forms. DON includes amino acid components and DON can therefore be used as proxy for amino acid availability. Similarly, extractions of IEM strips can also be analysed for DON.
Theory, significance, and large datasets
For details on the classic buried bag method, see Eno (1960). For a comprehensive review on ion-exchange methods, see Qian & Schoenau (2002), and for modern IEM-specific methods, see Lajtha et al. (1999) and Duran et al. (2013). Murphy et al. (2003) provide details of the 15N pool dilution method and a good overview of the N transformation processes and key references.
Binkley, D. (1984). Ion-exchange resin bags – Factors affecting estimates of nitrogen availability. Soil Science Society of America Journal, 48(5), 1181-1184.
Binkley, D., Aber, J. D., Pastor, J., & Nadelhoffer, K. (1986). Nitrogen availability in some Wisconsin forests: comparisons of resin bags and on-site incubations. Biology and Fertility of Soils, 2, 77-82.
Davidson, E. A., Hart, S. C., Shanks, C. A., & Firestone, M. K. (1991). Measuring gross nitrogen mineralization, immobilization, and nitrification by 15N isotopic pool dilution in intact soil cores. Journal of Soil Science, 42, 225-349.
Duran, J., Delgado-Baquerizo, M., Rodríguez, A., Covelo, F., & Gallardo, A. (2013). Ionic exchange membranes (IEMs): A good indicator of soil inorganic N production. Soil Biology and Biochemistry, 57, 964-968.
Eno, C. F. (1960). Nitrate production in the field by incubating the soil in polyethylene bags. Soil Science Society of America Journal, 24, 277-279.
Giblin, A. E., Laundre, J., Nadelhoffer, K., & Shaver, G. R. (1994). Measuring nutrient availability in arctic soils using ion-exchange resins: A field test. Soil Science Society of America Journal, 58, 154-1162.
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Read D. J. & Perez‐Moreno J. (2003). Mycorrhizas and nutrient cycling in ecosystems – a journey towards relevance? New Phytologist 157(3), 475-492.
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Authors: Schmidt IK1, Christiansen CT2
Reviewers: Berauer B3, Verbruggen E4,
1 Department of Geosciences and Natural Resource Management, University of Copenhagen, Frederiksberg, Denmark
2 NORCE Norwegian Research Centre and Bjerknes Centre for Climate Research, Bergen, Norway
3 Centre of Excellence PLECO (Plants and Ecosystems), Biology Department, University of Antwerp, Wilrijk, Belgium
4 University of Bayreuth, Department of Disturbance Ecology, Bayreuth, Germany