# Joseph Adu-Gyamfi · Verena Pfahler Editors

# Oxygen Isotopes of Inorganic Phosphate in Environmental Samples

Purification and Analysis

Oxygen Isotopes of Inorganic Phosphate in Environmental Samples

Joseph Adu-Gyamfi · Verena Pfahler Editors

# Oxygen Isotopes of Inorganic Phosphate in Environmental Samples

Purification and Analysis

*Editors* Joseph Adu-Gyamfi Soil and Water Management and Crop Nutrition Section, Joint FAO/IAEA Centre of Nuclear Techniques in Food and Agriculture International Atomic Energy Agency Vienna, Austria

Verena Pfahler Formerly Smithsonian Tropical Research Institute Balboa, Ancon, Panama

ISBN 978-3-030-97496-1 ISBN 978-3-030-97497-8 (eBook) https://doi.org/10.1007/978-3-030-97497-8

© IAEA: International Atomic Energy Agency 2022. This book is an open access publication. The opinions expressed in this publication are those of the authors/editors and do not necessarily reflect the views of the IAEA: International Atomic Energy Agency, its Board of Directors, or the countries they

represent.

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# **Preface**

The Soil and Water Management & Crop Nutrition (SWMCN) Subprogramme of the Joint Food and Agriculture Organization (FAO)/International Atomic Energy Agency (IAEA) Centre of Nuclear Techniques in Food and Agriculture supports IAEA Member States and FAO Member Countries in the use of stable isotope techniques for assessing the hydrological cycle in agro-ecological systems, evaluate evaporative losses and mixing of different water sources with the aim to improve water management and quality. Multi-tracer stable isotopes of nitrogen (δ15N, δ15N-NO3, δ18O-NO3), water (δ2H-H2O, δ18O-H2O), sulphur (δ34S-SO4, δ18O-SO4) and carbon (δ13C), have been used as forensics in water quality investigations. However, identifying sources and transport of phosphorus (P), as an agro-contaminant in agricultural catchments and environment has a drawback because it has only one stable isotope ( 31P) but many radioisotopes with short half-lives, making it difficult to use for long-term P cycling studies.

Phosphorus in most organic and inorganic forms is strongly bound to oxygen (O), which has three stable isotopes to enable tracking P cycling and transformation using the stable isotopes of O in phosphate (PO4), (δ18OP). In recent years, various studies have indicated that the analysis of the stable isotopic composition of oxygen (O) bound to P (δ18OP) to better understand P cycling in the environment, has become a promising tracer (surrogate) to investigate soil P transformation, plant P uptake, and to trace the sources of P from the soil to water bodies and the environment.

The SWMCN Subprogramme through a Coordinated Research Project (CRP) has actively sought to modify the methodological problems of the δ18OP protocols encountered in the field such as high loads of organic matter in the samples which makes purification challenging, and the need to improve sampling strategies in the field. This was done in collaboration with the Swiss Federal Institute of Technology in Zurich (ETH), the Trinity College, Dublin, Ireland, and scientists from developing countries. This guideline reflects the latest development and the state of art related to the use of stable isotopes of oxygen in phosphate to address the challenges encountered by scientists in the laboratory and in the field.

This publication is structured into six chapters outlines the background and examples of δ18Op studies in sediments, soils, fresh water, mineral fertilizers and plants, it presents a modified stepwise extraction and purification protocols, sample preparation for analysis, planning and designing of a study using δ18Op. The aspect of external quality assurance was provided with an example of an inter-laboratory study for silver phosphate standards and data interpretation, and finally the conclusions, future trends and opportunities for scaling out of the method from laboratory to field studies. It is expected that this guideline would be extensively applied in research geared to understand phosphorus dynamics in different agro-environments. The SWNCM Subprogramme wishes to thank all the contributors involved in the preparation of this publication.

Vienna, Austria Vienna, Austria Panama City, Panama Joseph Adu-Gyamfi Lee Heng Verena Pfahler

# **Contents**


# **Chapter 1 The Use of the δ18OP to Study P Cycling in the Environment**

#### **V. Pfahler, J. Adu-Gyamfi, D. O'Connell, and F. Tamburini**

**Abstract** Phosphorus (P) fertilizers are known to increase crop productivity; however, when applied in excess, it can cause serious environment pollution. Monitoring P pollution in natural environments using stable isotopes has been difficult because P has only one stable isotope (31P) making the use of P stable isotope tracing not an option. Radioactive P isotopes (32P and 33P) have been used but its drawbacks are the short half-life, health risks and safety procedures required to apply them in agricultural catchments. Phosphorus in organic and inorganic P forms is strongly bonded to oxygen (O), which has three stable isotopes, providing a system to track P cycling in agricultural catchments and environment using the stable isotopes of O in phosphate (δ18O-PO4). In recent years, various studies have indicated that the analysis of the stable isotopic composition of oxygen (O) bound to P (δ18Op) to better understand P cycling in the environment, has become a promising tracer (surrogate) to investigate soil P transformation, plant P uptake and to trace the sources of P from the soil to water bodies and the environment. The chapter outlines the background and examples of δ18Op studies in sediments, soils, fresh water, mineral fertilizers and plants.

#### D. O'Connell Department of Civil, Structural and Environmental Engineering, Trinity College Dublin, Dublin, Ireland

V. Pfahler Formerly Smithsonian Tropical Research Institute, Ancón, Panama

J. Adu-Gyamfi (B)

Soil and Water Management & Crop Nutrition Section, Joint FAO/IAEA Centre of Nuclear Techniques in Food and Agriculture, Vienna, Austria e-mail: J.Adu-Gyamfi@iaea.org

F. Tamburini Group of Plant Nutrition, D-USYS, ETH, Zurich, Switzerland

# **1.1 Background**

Phosphorus (P) is an essential nutrient for all living organisms as it is part of many biochemical compounds like DNA and ATP. Available P levels in the soil are, however, often limiting crop production and farmers need to apply P fertilizer. The primary source for P fertilizer is rock phosphate, which is widely distributed throughout the world both geographically and geologically with USA, China, Morocco and Western Sahara and Russia accounting for 72% of the world total (Zapata and Roy 2004). In addition, farmers in many developing countries cannot always afford P fertilizer, but the soils in those countries are often amongst those with the lowest P availability. Besides the positive impacts of P fertilizers increasing crop productivity, P can cause negative impacts on the environment when applied in excess. Algal blooms in aquatic systems due to large input of P are still an issue in many aquatic systems including the Great Lakes in North America (Paytan et al. 2017).We therefore need to better understand P cycling in the environment. Radioisotopes and stable isotopes are a useful tool to investigate nutrient cycling in the environment (Di et al. 1997; Hogberg 1997; Barbour 2007). In regard to P, researchers relied for a long time mainly on the two artificial radioisotopes of P, 32P and 33P, since P only has one stable P isotope (31P) and several radioisotopes (from 26P to 30P and from 32P to 38P). The radioisotope approach is often used to study the uptake of P from different phosphate rocks and fertilizers by plants (Zapata and Roy 2004; Frossard et al. 2011; IAEA 2013, 2016a, b; Nanzer et al. 2014; Nguyen et al. 2017; Scrase et al. 2020; Wolff et al. 2020), the effects of organic manure and inorganic fertilizer application on soil P cycling (Ma et al. 2020) or the residence time of P in different P pools in soils or plants (Mimura et al. 1996; Helfenstein et al. 2018). It is a useful tool to study P cycling, but its drawbacks are the short half-life of the two radioisotopes (14.3 days for 32P and 25.4 days for 33P) and possible health risks, which has led to the restrictions of applying them in the field, and the need for special safety equipment and national regulations to monitor radiation. Obtaining the licence to work with radioisotopes of P that emit high beta (β) radiation will require operational radiation protection and safety procedures, as well as adequate training of personnel to handle these nuclides safely (IAEA 2016a, b). There is therefore a need for a stable isotope approach to investigate P cycling in the environment.

In recent years, various studies have indicated that the analysis of the stable isotopic composition of oxygen (O) bound to P (δ18OP) to better understand P cycling in the environment, has become a promising tracer (proxy) to investigate soil P transformation and to trace the source of P from the soil to water bodies and the environment. The two main underlying facts for using this method to study P cycling are (1) P is mainly bound to O in the environment and (2) the bond between P and O is stable under earth surface conditions and in the absence of biological activity (Winter et al. 1940). The two ways of altering the δ18OP signature are by (a) sorting between heavier and lighter isotopologues or (b) cleaving the P–O bond.

# *1.1.1 Sorting of Different Isotopologues*

The sorting of heavier and lighter isotopologues could happen due to biological P uptake or inorganic processes, like the preferential sorption onto iron oxides (Jaisi et al. 2010). There are only a few studies which investigated the effect of P uptake by biota. Blake et al. (1997) report a fractionation factor of around −3‰ for *Escherichia coli*. More recently Ferrera et al. (2015) investigated the effect of P uptake by different coral species on the δ18OP. Depending on the coral species, fractionation factors varied between −18.80 and +0.91‰, with the majority of the investigated species having negative fractionation factors. If an uptake effect, however, is visible in environmental systems, is questionable as a significant amount of P (>50%) must be removed from a P pool without being replenished in order to observe an effect on δ18OP values. This seems rather questionable in case of P in the soil solution since it is usually constantly replenished. Another kind of sorting can occur due to inorganic processes like sorption/desorption of P onto iron or aluminium oxides and precipitation/dissolution of P minerals (Jaisi et al. 2010). These processes play an important role in the environmental P cycle (Arai and Sparks 2007), but fractionation factors associated with inorganic processes seem to be rather small and are still debated (Jaisi et al. 2010; Melby et al. 2013).

# *1.1.2 Cleaving the P–O Bond*

Biological activity like the hydrolysis of organic P by enzymes, can cleave the P–O bond and thus lead to an exchange of O between phosphate and water (Blake 2005). This leads to a change of the initial δ18OP value (Table 1.1).

Inorganic pyrophosphatase (PPase), an ubiquitous enzyme catalyses the hydrolysis of pyrophosphate (= diphosphate) into phosphate (Cohn 1958; Blake et al. 1997) and leads to a temperature-dependent equilibrium between O in water and in phosphate. It can be calculated by the equation of Chang and Blake (2015), as rearranged by Pistocchi et al. (2017):

$$\rm{EQ\delta}^{18}\rm{O}\_{\rm P} = -0.17 \cdot T + 26.5 + \delta^{18}\rm{O}\_{\rm w} \tag{1.1}$$

where EQδ18OP is the δ18OP value of phosphate at equilibrium with O in the ambient water in ‰,

T is the ambient temperature in °C and δ18Ow is the isotopic composition of O in water in ‰.

What makes the inorganic PPase so important for the δ18OP is that O exchange can not only occur during the hydrolysis of pyrophosphate but also by locking a phosphate molecule into the active site of the enzyme (Cohn 1958; Blake et al. 1997). When the inorganic PPase is involved, the initial δ18OP signature is erased completely (see Table 1.1).


**Table 1.1** Examples for the effect of different enzymes on δ18OP values

Inorganic PPase = inorganic pyrophosphatase; EQ = Equilibrium fractionation; nr: not relevant (reaction is temperature-independent); DNase 1= deoxyribonuclease 1; AlPase = alkaline phosphatase

Hydrolytic enzymes like acid and alkaline phosphatases only lead to the exchange of 1–2 oxygen atoms and are temperature-independent (Liang and Blake 2006a; von Sperber et al. 2014, 2015). The associated fractionation factors vary greatly, from + 20 to −30‰, and consequently the δ18OP of the released phosphate varies depending on the involved enzymes. Some fractionation factors like the one of phytase and acid and alkaline phosphatase are substrate dependent (von Sperber et al. 2015; Bai et al. 2020).

# **1.2 From Past to Present**

Early 18O studies worked with 18O-enriched samples rather than natural abundance (Winter et al. 1940; Cohn and Hu 1978; Larsen et al. 1989) and were designed to understand the mechanism of the studied enzymes. Natural abundance studies only came later and were not that common, most likely due to the challenges of purifying extracts for δ18OP analysis. At the beginning, bismuth phosphate (BiPO4) was used as analyte (Tudge 1960), however, BiPO4 is highly hydroscopic, making it more difficult to handle and store (Vennemann et al. 2002). Nowadays, Ag3PO4 is used as analyte (Firsching 1961; Crowson et al. 1991), which is easier to handle and faster to precipitate. Advancements have also been made in the analysis of Ag3PO4 (Vennemann et al. 2002). The analysis with a high temperature conversion elemental analyser (TC/EA) coupled to an isotope ratio mass spectrometer (IRMS) is now the most common method.

The origins of the 18O method lie in chemical, biochemical and paleotemperature studies (Tamburini et al. 2014). In the early 2000s, Blake et al. (2001) postulated that the δ18OP method could be used to study biological P cycling. From then onwards, the δ18OP method became more widespread and scientists used it in a diverse range of environmental samples, including lake and marine sediments, salt and fresh water, soils, mineral and organic fertilizers, rocks, dust and plants (Tamburini et al. 2014). Most naturally occurring δ18OP values, reported so far, range between 10 and 30‰, with the exception of igneous rocks, which range between −0.8 and +12‰ (Smith et al. (2021)). For a more detailed overview of δ18OP values in the environment, the reader is referred to Bauke (2020) for δ18OP values in soils; Smith et al. (2021) for δ18OP values from rocks; for aquatic ecosystems, see for example Davies et al. (2014) and Gooddy et al. (2015). General overviews of δ18OP values can, for example, be found in Tamburini et al. (2014) and Jaisi & Blake (2014). Figure 1.1 shows examples

**Fig. 1.1** Examples for δ18OP values of potential phosphorus (P) inputs into soils and aquatic systems. Vegetation Pi = inorganic P in vegetation; vegetation Porg = organic P in vegetation; org. fertilizer = organic fertilizer; min. fertilizer = mineral fertilizer; WWTP = wastewater treatment plant

for δ18OP values of potential P inputs like animal faeces and mineral P fertilizers into soils and aquatic systems (see Annex Table 1.2 for references).

The δ18OP method was also considered ideal to proof life on other planets, for example, on Mars (Greenwood et al. 2003). To be able to analyse the δ18OP from such a diverse range of substrates, several obstacles had to be overcome. The main obstacles for the δ18OP method are low P concentrations and/or high concentrations of interfering compounds like organic matter (Tamburini et al. 2010; Goldhammer et al. 2011b; Nisbeth et al. 2019). Therefore, modern protocols of the δ18OP method involve several purification steps before the final precipitation of silver phosphate (Ag3PO4) which is then used to determine the δ18OP via TC/EA-IRMS. McLaughlin et al. (2004) adapted the method for seawater samples adding a MAGIC step (precipitation of Mg (OH)2) and purifying the samples using cerium phosphate. Tamburini et al. (2010) applied this protocol to soil extracts but found that the Ag3PO4 could be still contaminated with organic matter. It was therefore suggested to use precipitation of ammonium phosphomolybdate and magnesium ammonium phosphate to purify extracts (Kolodny et al. 1983), adding an additional purification step (DAX-resin) at the beginning of the protocol.

Nowadays, the δ18OP method is more and more used to investigate P cycling in the environment, including sediments (Jaisi and Blake 2010; Goldhammer et al. 2011a; Pistocchi et al. 2017; Liu et al. 2019), soils (Amelung et al. 2015; Gross and Angert 2015; Helfenstein et al. 2018; Pistocchi et al. 2020; Siegenthaler et al. 2020; Pfahler et al. 2020) and plants/algae (Pfahler et al. 2013; Mellett et al. 2018). The focus in case of aquatic systems is often on source apportionment (McLaughlin et al. 2006a; Gross et al. 2013; Granger et al. 2017). Other studies characterise the δ18OP of different P inputs into the environment like mineral fertilizers and farm-yard manure (Granger et al. 2018) or investigate the effect of enzymes on δ18OP values (Blake et al. 1998; Liang and Blake 2006b; von Sperber et al. 2014).

# **1.3 Examples of δ18OP Studies**

# **1.4 Sediments**

Goldhammer et al. (2011a) analysed dissolved P extracted from sediment pore water, using marine sediments from the Benguela upwelling system of the coast of Namibia in Africa. They found dissolved P δ18OP values in and out of equilibrium, ranging between 12.8 and 26.6‰, and attributed these values to different P uptake strategies of microorganisms at low and high inorganic P availability.

Investigating river sediments from the Redon River in France, Pistocchi et al. (2017) showed that the δ18OP can potentially be used to trace P sources and study P cycling in river sediments. They also adapted the method by Tamburini et al. (2010) successfully to river sediments. Liu et al. (2019) explored a pre-treatment method for the δ18OP of different P fractions in sediments. They also concluded that the δ18OP can be a promising tool to trace P and study P cycling in freshwater sediments.

# **1.5 Soils**

With the δ18OP approach, Tamburini et al. (2012) investigated the importance of microorganisms for soil P cycling. They extracted available, microbial, vegetation and mineral P and analysed the corresponding δ18OP values from soils taken along a soil chronosequence at the Damma glacier fore field in Switzerland. Regardless of the contribution of vegetation, mineral or P released by organic P hydrolysis, available P δ18OP values were always close to microbial P δ18OP values. This showed, for the first time under field conditions, the importance of microbial P cycling for the available P.

Recently, Bi et al. (2018) combined the δ18OP method with measuring the abundance of genes related to the P cycle like *phoX* (acid phosphatase) and *phoD* (alkaline phosphatase D) to investigate P cycling in agricultural soil from the Fengqiu State Key Experimental Station for Ecological Agriculture (Henan, China)*.*Like Tamburini et al. (2012), they measured acid phosphatase activity and additionally also alkaline phosphatase, phosphodiesterase and dehydrogenase activity. They showed the importance of microbial P cycling in agricultural soil with δ18OP values of the more labile P pools (water, sodium bicarbonate and sodium hydroxide extractable inorganic P) tending towards equilibrium.

# **1.6 Salt and Fresh Water**

After McLaughlin et al. (2004) successfully developed a method to analyse the δ18OP of phosphate in seawater samples, Elsbury et al. (2009) applied this method to investigate P cycling and inputs into Lake Erie. Along with water samples from different locations at Lake Erie, they also analysed water samples from seven rivers feeding into Lake Erie. The δ18OP values ranged between 10 and 17‰, with an expected equilibrium value of 14‰. The δ18OP values of the rivers were around 11‰ and could thus be one source of P, however, they did not find a P source with higher δ18OP values (Elsbury et al. 2009). They proposed that P released from sediments might cause the higher δ18OP values found in Lake Erie. Gooddy et al. (2016) used the δ18OP along with δ15N and δ18O of nitrate and δ15N of ammonium to investigate eutrophication. They took water samples from River Beult (Kent, UK) twice within six months and at seven sampling sites along the river. One of their conclusions was that abiotic processes in the river cause the P concentration changes as the isotopic values did not change dramatically along the river.

# **1.7 Dust**

The main goal of the study by Gross et al. (2013) was to trace the input of P into Lake Kinneret in Israel through atmospheric deposition of dust. They analysed the δ18OP values of available P from soil and dust samples and found that in natural soils, the δ18OP values were between 17.4 and 18.2‰ and between 19.3 and 22.1‰ in agricultural soils. Resin P extracted from dust samples had δ18OP values of around 22‰, indicating that agricultural soils were the main source for dust in the region around Lake Kinneret. Following this study, Gross et al. (2016) found that P associated with Saharan dust can be traced to South America.

# **1.8 Mineral Fertilizers, Manures and Rocks**

The δ18OP values of mineral fertilizers range from 6.4 to 25.9‰ (Fig. 1.1). This wide range is most likely caused by the bedrock material used to produce the mineral fertilizers (Gruau et al. 2005), which δ18OP values vary similarly to the ones of mineral fertilizers (Smith et al. 2021). Phosphorus leached from fresh cattle faeces can be an important P input into aquatic systems, but is sometimes hard to quantify (Bond et al. 2014). Granger et al. (2018) investigated how variable faeces δ18OP values are. They collected faeces samples from seven different cows, grazing on different pastures in Devon (UK) and differing in gender, race and age. Values ranged between 13.2 and 15.3‰ without correlation with any of the variables (pastures, gender, race and age), but they were within equilibrium range calculated with the δ18OP of groundwater and cattle body temperature (Granger et al. 2018).

In one of the early δ18OP studies, Mizota et al. (1992) determined the δ18OP of a range of lithogenic material and soil samples from the Great Rift Valley in Africa and Java (Indonesia). Volcanic ashes had δ18OP values between 5.3 and 6.2‰, apatite from hydrothermal deposits had δ18OP values between 2.4 and 12.2‰, and apatite from carbonatites δ18OP values between 0.2 and 10‰. The soil samples were extracted sequentially, first with 2.5% acetic acid and then with 1M ammonium fluoride (NH4F). Their soil δ18OP values ranged between 10.1 and 20.6‰ (acetic acidextractable P) and between 12.7 and 24.8‰ (NH4F-extractable P). Their conclusion was that available soil P δ18OP values are due to biogenic and volcanic inputs.

# **1.9 Plants**

Inorganic P in plants tends to have higher δ18OP values compared to the soil due to the 18O-enriched plant water compared to the soil water (Pfahler et al. 2013). However, as Pfahler et al. (2017) revealed, P limitation lowers inorganic P δ18OP values in soybean leaves. Using 33P along with measurement of the δ18OP, the authors could also calculate P fluxes to and from different plant parts. Qin et al. (2018) used the δ18OP method to investigate P uptake by maize plants. They found that the δ18OP value of the P applied to the soil and taken up via the roots directly was found in maize shoots, whereas the δ18OP value had changed if P was taken up via arbuscular mycorrhizal fungi (Qin et al. 2018).

# **Glossary**


# **Annex**


**Table 1.2** References included in Fig. 1.1

(continued)


(continued)

# **References**


The opinions expressed in this chapter are those of the author(s) and do not necessarily reflect the views of the IAEA: International Atomic Energy Agency, its Board of Directors, or the countries they represent.

**Open Access** This chapter is licensed under the terms of the Creative Commons Attribution 3.0 IGO License (https://creativecommons.org/licenses/by/3.0/igo/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the IAEA: International Atomic Energy Agency, provide a link to the Creative Commons license and indicate if changes were made.

Any dispute related to the use of the works of the IAEA: International Atomic Energy Agency that cannot be settled amicably shall be submitted to arbitration pursuant to the UNCITRAL rules. The use of the IAEA: International Atomic Energy Agency's name for any purpose other than for attribution, and the use of the IAEA: International Atomic Energy Agency's logo, shall be subject to a separate written license agreement between the IAEA: International Atomic Energy Agency and the user and is not authorized as part of this CC-IGO license. Note that the link provided above includes additional terms and conditions of the license.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# **Chapter 2 Extraction Protocol**

#### **V. Pfahler, J. Adu-Gyamfi, D. O'Connell, and F. Tamburini**

**Abstract** Studies showed that the δ18OP is a useful tool to study P in the environment. Adequate extraction protocols for the targeted P pools of the study are a prerequisite for a successful study. Likewise, for most environmental samples, including water, soil, sediment and plant samples, it is crucial that the samples are processed as soon as possible after they have been taken to avoid any alterations of the original δ18OP signature. This is especially true when more bioavailable P pools, like soluble reactive P (SRP) in water samples, are extracted and analysed. Brucite precipitation of water samples should be directly done in the field, fresh soil and sediment samples have to be extracted within 7 days (if microbial P is targeted, on the day of sampling), and plant samples have to be extracted within a few hours of sampling or be frozen. The chapter briefly describes the P cycle in aquatic and terrestrial ecosystems and give an overview about extracting the most common P pools for δ18OP analysis: soluble reactive P in water samples, sequentially extracted P pools of soil, sediment, fertilizer and plant samples.

# **2.1 An Overview of the Extraction Protocols**

Studies showed that the δ18OP is a useful tool to study P in the environment (Helfenstein et al. 2018). Adequate extraction protocols for the targeted P pools of the study

V. Pfahler

#### D. O'Connell Department of Civil, Structural and Environmental Engineering, Trinity, College Dublin, Dublin, Ireland

#### F. Tamburini Group of Plant Nutrition, D-USYS, ETH, Zurich, Switzerland

Formerly Smithsonian Tropical Research Institute, Ancón, Panama

J. Adu-Gyamfi (B)

Soil and Water Management & Crop Nutrition Section, Joint FAO/IAEA Centre of Nuclear Techniques in Food and Agriculture, Vienna, Austria e-mail: J.Adu-Gyamfi@iaea.org

are a prerequisite for a successful study. Likewise, for most environmental samples, including water, soil, sediment and plant samples, it is crucial that the samples are processed as soon as possible after they have been taken in order to avoid any alterations of the original δ18OP signature. This is especially true when more bioavailable P pools, like soluble reactive P (SRP) in water samples, are extracted and analysed. Brucite precipitation of water samples should be directly done in the field (see Sect. 2.2), fresh soil (see Sect. 2.3) and sediment (see Sect. 2.4) samples have to be extracted within 7 days (if microbial P is targeted, on the day of sampling), and plant samples (see Sect. 2.5) have to be extracted within a few hours of sampling or be frozen.

The following chapters briefly describe the P cycle in aquatic and terrestrial ecosystems and give an overview about extracting the most common P pools for δ18OP analysis: soluble reactive P in water samples, sequentially extracted P pools of soil, sediment, fertilizer and plant samples. In general, P occurs in microbial, organic and inorganic forms in the environment (Condron and Newman 2011). These forms are interlinked with each other through inorganic and biological processes and are often clustered together according to their extractability with different chemicals. Which of these pools needs to be extracted and analysed for its δ18OP value depends on the research question.

# **2.2 Aquatic Systems**

Phosphorus inputs into aquatic systems are mainly from non-point sources like runoff from roads and fields or point sources, for example, from wastewater treatment plants. Direct disposal of untreated wastewater, effluents and solid wastes to water ways from households and small-scale industries, and leachates from septic tanks also contributed to the enrichment of P in water ways. Atmospheric inputs of P into aquatic systems are often low but can still negatively affect water quality in some cases. Since bioavailability of P in aquatic systems is often low and hence aquatic biota like plants are efficient in taking up P (Reynolds and Davies 2001), those P inputs can have a drastic effect on aquatic ecosystems like eutrophication (Schindler et al. 2016). In marine ecosystems, P is often considered as the nutrient which is limiting primary production (Filippelli 2008).

Soluble reactive P (SRP) is considered most bioavailable and many studies and water quality guidelines focus on this pool (Fig. 2.1) (Directive 2000).

Particulate P in water is also important as it can contribute significantly to the bioavailable P in aquatic systems (Ellison and Brett 2006). Both particulate P and SRP contribute to sedimentary P, either by deposition or adsorption onto particles. Through resuspension and desorption processes their bioavailability for aquatic organisms increases again. Once taken up by the biota, inorganic P can be incorporated into organic P (Porg). The P bond in organic forms becomes available through the hydrolysis by enzymes. Organic P like organic matter, in general, can also be buried into the sediment.

#### 2 Extraction Protocol 19

**Fig. 2.1** Simplified phosphorus (P) cycling in aquatic systems. WWTP = wastewater treatment plant

# *2.2.1 Extraction of Soluble Reactive Phosphorus (SRP)*

Concentrations of SRP can be very low in water bodies and thus it might be necessary to collect several litres of water for the determination of the δ18OP as 20–30 μmol P (= 0.65–0.97 μg P) which are necessary for the purification protocol. Granger et al. (2017b) sampled, for example, between 25 and 50 L of water. It is therefore recommended to measure the SRP concentration in the water before the actual sampling. The below-described sampling protocol is based on the method by Nisbeth et al. (2019) (Fig. 2.2).

As SRP concentrations are often low, it is recommended to thoroughly clean all equipment/consumables like polypropylene bottles with P-free detergent and rinse them with diluted HCl and ultrapure water before using.

**Fig. 2.2** Scheme of water collection and processing for δ18OP analysis

# *2.2.2 Equipment and Consumables*


# *2.2.3 Reagents*

# 1. **3 M magnesium brine**

Weigh out 1.6 kg of MgCl2 (hexahydrate; MW: 203.3 g/mol). Add 2.5 L of ultrapure water. After the salt has dissolved, filter the brine on a GF/F filter.

# 2. **1 M NaOH**

Weigh out 40 g of NaOH pellets. Dissolve it in 1 L of ultrapure water.

# 3. **1 M HNO3**

In a volumetric flask add 66 mL concentrated HNO3 to 800 mL ultrapure water and make up to 1 L.

# **Step SRP\_1—Water collection (field)**


# **Step SRP\_2—First Brucite precipitation (field)**

• Measure out 1 L of the Mg brine and pour it into the 50 L bottle. Shake or stir using a rod well. Measure out 250 mL of 1 M NaOH and add it. Shake again.

# **Step SRP\_3—Discarding supernatant (field)**

• If it is cold, leave the 50 L bottle in the field and let the Mg(OH)2 (brucite) floc settle. After about 1 h, siphon out the supernatant by using the plastic tube. Thick pipes might be faster; however, thinner tubes can reduce losses and are therefore preferable. About 1/10 of the initial volume will remain. If it is warm, the bottle

#### 2 Extraction Protocol 21

should be transferred to the lab, put into a fridge and shaken. After a couple of hours, the floc is settled and no brucite remains in suspension, the supernatant is discarded.

#### **Step SRP\_4—Centrifuging brucite flocs (laboratory)**

• The remaining brucite floc should be centrifuged at about 3000 rpm for 15 min. The supernatant is then eliminated.

#### **Step SRP\_5—Dissolution of brucite flocs (laboratory)**

• Brucite is removed from the centrifuging bottles by dissolving it with 1 M HNO3. Use the minimum amount of acid to dissolve the brucite from each bottle. Combine the solutions. pH should be around 1. If needed, the volume can be further reduced by performing additional MAGIC (steps SRP\_2 to SRP\_5) steps. This is done only by raising the pH to about 10–11 (e.g. by adding NaOH). Then, repeat step SRP\_2–5. Generally, after three MAGIC steps, the volume is reduced from 50 L to about 250–300 mL.

#### **Step SRP\_6—Filtration**

• Once the desired volume is reached (generally around 100–150 mL), the solutions are filtered using GF/F filters.

# **2.3 Soils**

Phosphorus occurs in inorganic and organic forms in soils (Fig. 2.3). The concentration of inorganic P in the soil solution (available P) is often relatively low, around 0.1 mg P L−1, and is controlled by several biotic and abiotic processes (Fig. 2.3).

Immobilization of inorganic P by microorganisms, precipitation of P minerals, and sorption of P onto iron oxides or clay minerals reduce the concentration of inorganic P in the soil solution (Arai and Sparks 2007). Mineralization of organic P, dissolution of P minerals and desorption of Pi from soil particles increase the concentration of P in the soil solution (Shen et al. 2011). Organic P in soils typically makes up 30– 65% of total P (Harrison 1987) and includes phosphomonoesters and diesters, such as DNA and RNA. These forms of organic P cannot directly be taken up by plants or microorganisms. Plants and microorganisms developed several strategies to make organic P and other immobilized P available again, such as exudation of organic acids and phosphoenzymes like acid phosphatase. Association with mycorrhizal fungi is also a common strategy among plants to increase P availability (Shen et al. 2011).

The extraction methods for different soil P pools vary greatly from country to country and soil to soil (Nawara et al. 2017). Sodium bicarbonate, anion exchange resin membranes and water are only some of the extractants used for available P. Not all existing methods for extraction different soil P pools are suitable for the analysis of the δ18OP. In some cases, P concentrations in the extracts would be too low so that a large amount of soil would be necessary, or the extraction would

**Fig. 2.3** Abiotic processes with no/little fractionation 2. Inorganic hydrolysis of condensed phosphates 3. Preferential uptake of P by, for example, microorganisms 4. Intracellular cycling of P leading to equilibrium between O in phosphate and water, mediated my inorganic pyrophosphatase 5. Hydrolysis of organic P by phosphoenzymes like acid and alkaline phosphatase

lead to an uncontrolled O exchange between phosphate and the extracting reagent. Other extracts might cause issues during the purification protocol. A new extraction method should therefore first be tested for its suitability for the δ18OP analysis. The most tested and used method is the extraction with 1 M HCl (Tamburini et al. 2010; Amelung et al. 2015; Granger et al. 2017a). It is also often the easiest when it comes to sample handling as the samples do not need to be fresh but can be dried. It might, however, not always be the most meaningful method, depending on the research hypothesis. When investigating the impact of biological processes on soil P cycling, the available (and microbial) P pool is better suited (see also Chap. 5). The following sequential extraction protocol is based on Tamburini et al. (2018) and consists of the extraction of resin and hexanol P, NaOH-EDTA inorganic and organic P and HCl P (Fig. 2.4). Tamburini et al. (2018) developed the method to purify inorganic and organic P in NaOH-EDTA extracts and follows the protocols by Weiner et al. (2011) and Kouno et al. (1995) for resin and hexanol P and Tamburini et al. (2010) for HCl P. The microbial P is calculated with a simple mass balance using the resin and hexanol P concentrations and δ18OP values (Tamburini et al. 2012).

#### **Equipment and consumables**



#### **Reagents**

1. **Anion exchange resin membranes**

The resin membranes are stored in 1 M HNO3. Take out the number of membranes you need and wash with ultrapure water. Shake for 1 h in 0.5 M NaHCO3. Wash with ultrapure water. Shake for 1 h in 0.5 M NaHCO3. Wash three times with ultrapure water and store in ultrapure water before use.

# 2. **0.5 M NaHCO3**

Weigh out 42.3 g of sodium bicarbonate (NaHCO3; MW = 84.01 g/mol). Add 1 L of ultrapure water. Dissolve well by stirring on magnetic plate. Prepare fresh.

# 3. **Hexanol**

# 4. **0.2 M HNO3**

Measure 986 mL ultrapure water and add 14 mL concentrated HNO3 (65% HNO3).

# 5. **1 M HNO3**

In a volumetric flask add 66 mL concentrated HNO3 to 800 mL ultrapure water and make up to 1 L.

## 6. **0.25 M NaOH–0.05 M EDTA solution**

Weigh out 10 g NaOH and 18.612 g EDTA disodium salt and dissolve in 1 L ultrapure water.

#### 7. **1 M HCl**

In a volumetric flask add 82.7 mL concentrated HCl (37%) in 900 mL ultrapure water and make up to 1 L. Prepare two batches of 1 M HCl (18O-labelled and unlabelled HCl solutions). Take a subsample from each batch for the determination of the δ18Ow value of each batch.

Use a soil: solution ratio of 1:10 throughout the protocol.

#### **Step S\_1 Sample preparation**

	- Sample 1: Resin P.
	- Sample 2: Hexanol P.

#### **Step S\_2 Resin and hexanol P extraction**

• Add ultrapure water to sample 1 and 2.

#### 2 Extraction Protocol 25


# **Step S\_3 Recovering resin strips**


# **Step S\_4 NaOH-EDTA extraction**


# **Step S\_5 HCl extraction**


# **2.4 Extraction of P from Sediments**

Extracting P from sediments for δ18OP analysis is similar to the extraction of P from soils (see Sect. 2.3; Fig. 2.4). Sediment samples can be extracted sequentially including extraction with a citrate-dithionite-bicarbonate (CDB) solution and a sodium acetate buffer prior to the extraction with 1 M HCl (Ruttenberg 1992; Ruttenberg et al. 2009). The CDB and acetate fractions can be discarded and the 1 M HCl fraction purified is described in Chap. 3.

# **2.5 Plants and Fertilizers**

Plants and fertilizers are a source of P into aquatic and terrestrial ecosystems. They are therefore important endmembers in δ18OP studies. Both are relatively easy to extract since concentrations are usually higher than in other samples like soils and thus less material is necessary.

# *2.5.1 Fertilizers*

Phosphorus fertilizers are divided into mineral (e.g. triple superphosphate (TSP)) and organic fertilizers (e.g. manure). In mineral P fertilizers most of the P is present as inorganic P and is readily available. Organic P fertilizers also contain less readily available P forms. It is thus useful to extract especially organic P fertilizers sequentially. 18O-labelled and unlabelled 1 M HCl have to be used for the extraction as 1 M HCl could hydrolyse some organic P and/or condensed phosphate, which can be present especially in organic fertilizers.

# **Equipment and consumables**


# **Reagents**

1. **1 M HCl**

In a volumetric flask add 82.7 mL concentrated HCl (37%) in 900 mL ultrapure water and make up to 1 L. Prepare two batches of 1 M HCl (18O-labelled and unlabelled HCl solutions). Take a subsample from each batch for the determination of the δ18Ow value of each batch.

# **Step Fert\_1—Sample preparation**


# **Step Fert\_2—Water extraction**

• Add ultrapure water and shake overnight.

#### **Step Fert\_3—Filtration**

• The next day, filter extract using GF/F filters.

#### 2 Extraction Protocol 27

This will yield the readily available P of a fertilizer. For less available P, extract the fertilizer samples also with HCl. Ideally this is done sequentially; however, some fertilizers will almost completely dissolve during the water extraction. In that case, HCl extraction needs to be done on a separate subsample without prior water extraction.

#### **Step Fert\_4—HCl extraction**


# *2.5.2 Plants*

Plants contain inorganic and organic P. Inorganic P can be leached from plant material and contributes directly to the available P pool in soils. Organic P needs to be mineralized by enzymes first. Especially in low P soils with high biomass production like tropical rainforest, P from plants can contribute significantly to the P nutrition of the rainforest. Therefore, both inorganic and organic P from plants are important pools in δ18OP studies. The main challenge for extracting P from plant material is to stop enzymatic activity during the extraction (Bieleski 1964; Adu-Gyamfi et al. 1990). As enzymatic activity leads to O exchange between phosphate and water, it is essential to choose extraction conditions which minimize it. Trichloroacetic acid (TCA) is efficient in halting enzymatic activity and it has been successfully used to analyse the δ18OP in plant material (Pfahler et al. 2013). TCA could also extract some organic P and therefore 18O-labelled and unlabelled TCA solutions are used, similar to extracting samples with 1 <sup>M</sup> HCl. After the extraction with TCA, plant material can be extracted with NaOH-EDTA for the determination of the δ18OP of structural P (mainly organic P) (Helfenstein et al. 2018; Tamburini et al. 2018).

#### **Equipment and consumables**


# **Reagents**

# 1. **0.3 M Trichloroacetic acid**

Weigh 49.02 g TCA (MW 163.39 g/mol) in a volumetric flask and fill it up to 1000 mL with ultrapure water. Prepare two batches of 0.3 M TCA (18Olabelled and unlabelled TCA solutions). Take a subsample from each batch for the determination of the δ18Ow value of each batch.

# 2. **0.25 M NaOH—0.05 M EDTA**

Weigh out 10 g NaOH and 18.612 g EDTA disodium salt and dissolve in 1 L ultrapure water.

# **Step Pl\_1—Sample preparation**


# **Step Pl\_2—Extraction**


# **Step Pl\_3**


If the δ18OP of the organic P in the plants needs to be analysed, proceed with steps Pl-4 and Pl\_5.

# **Step Pl\_4**

• Carefully remove the plant material remaining on the GF/F filter with tweezers and extract it with NaOH-EDTA (see step S\_4 for more details).

#### 2 Extraction Protocol 29

#### **Step Pl\_5**


# **Glossary**


# **References**


#### 2 Extraction Protocol 31

The opinions expressed in this chapter are those of the author(s) and do not necessarily reflect the views of the IAEA: International Atomic Energy Agency, its Board of Directors, or the countries they represent.

**Open Access** This chapter is licensed under the terms of the Creative Commons Attribution 3.0 IGO License (https://creativecommons.org/licenses/by/3.0/igo/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the IAEA: International Atomic Energy Agency, provide a link to the Creative Commons license and indicate if changes were made.

Any dispute related to the use of the works of the IAEA: International Atomic Energy Agency that cannot be settled amicably shall be submitted to arbitration pursuant to the UNCITRAL rules. The use of the IAEA: International Atomic Energy Agency's name for any purpose other than for attribution, and the use of the IAEA: International Atomic Energy Agency's logo, shall be subject to a separate written license agreement between the IAEA: International Atomic Energy Agency and the user and is not authorized as part of this CC-IGO license. Note that the link provided above includes additional terms and conditions of the license.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# **Chapter 3 Purification Protocol**

#### **V. Pfahler, J. Adu-Gyamfi, D. O'Connell, and F. Tamburini**

**Abstract** The five stepwise purification of extracts and final precipitation of silver phosphate (A1–A5) are described. The first two steps (A1 and A2) are removing organic matter and are concentrating the phosphate in the extract by reducing the volume. Certain cations could interfere with the precipitation of silver phosphate and are removed in step A3. Silver chloride, which, if not removed, could co-precipitate with silver phosphate, is removed in step A4. The final analyte is then precipitated in step A5. The filtration steps can be quite tedious, using vacuum filtration equipment is therefore recommended. Following step A5, the silver phosphate samples need to be weighed in for the measurement with a thermal conversion elemental analyser (TC/EA) coupled to a continuous-flow isotope-ratio mass spectrometer (IRMS).

# **3.1 Introduction**

The main difference between the protocol by Tamburini et al. (2010) and other methods is the precipitation of cerium phosphate to purify extracts (McLaughlin et al. 2004). An issue with the precipitation of cerium phosphate is that it precipitates around pH 5.5, which is the same range where iron oxides flocculate. While flocculating iron oxides can strip out other components like dissolved organic matter (DOM) from the solution (Hiradate et al. 2006). This combination of iron oxides

V. Pfahler

#### D. O'Connell Department of Civil, Structural and Environmental Engineering, Trinity College Dublin, Dublin, Ireland

F. Tamburini Group of Plant Nutrition, D-USYS, ETH, Zurich, Switzerland

Formerly Smithsonian Tropical Research Institute, Ancón, Panama

J. Adu-Gyamfi (B)

Soil and Water Management & Crop Nutrition Section, Joint FAO/IAEA Centre of Nuclear Techniques in Food and Agriculture, Vienna, Austria e-mail: J.Adu-Gyamfi@iaea.org

and OM stays until the end and the obtained silver phosphate will be contaminated (Tamburini et al. 2010).

The following purification protocol is based on the original protocol published by Tamburini et al. (2010) and the protocol described in Chap. 2 of the IAEA-TECDOC-1783 (IAEA 2016). The original protocol was developed for 1 M HCl extracts of soils and fertilizers but has been adapted since its publication to a range of other extracts including NaOH-EDTA from soils (Tamburini et al. 2018) and trichloroacetic acid (TCA) extracts from plants (Pfahler et al. 2013). Those modifications are described in Chap. 4.

# **3.2 Reagents**

# (1) **Ammonium nitrate solution, 35% (w/v) (= 4.2 M)**

Weigh out 538.5 g of ammonium nitrate salt (NH4NO3; molecular weight (MW) 80.052 g/mol). Add 1000 mL of ultrapure water and dissolve the salt, stirring well. Store at room temperature.

# (2) **Ammonium nitrate solution, 5% (w/v) (= 0.6 M)**

Weigh out 105.3 g of NH4NO3 salt. Add 2000 mL of ultrapure water. Stir well to dissolve salt. Store at room temperature.

#### (3) **Ammonium heptamolybdate solution, 10% (w/v)**

Weigh out 53.3 g of ammonium molybdate salt (NH4Mo x7H2O; tetrahydrate form; MW 1235.86 g/mol). Add 480 mL of ultrapure water. Dissolve well. Enough for 12 samples. Prepare fresh before each use.

# (4) **Ammonium-citrate solution**

Weigh out 10 g of citric acid. Add 300 mL of ultrapure water and 140 mL of concentrated ammonia solution (NH4OH). Prepare and use under hood. Stable at room temperature.

# (5) **Magnesia solution**

Weigh out 50 g of magnesium chloride (MgCl2 <sup>×</sup> 6H2O; hexa-hydrate salt; MW 203.3 g/mol) and 100 g of ammonium chloride (NH4Cl; MW 53.49 g/mol). Dissolve them in 500 ml of ultrapure water. Acidify to pH 1 with conc. HCl. Volume is then adjusted to 1 L with ultrapure water. Stable indefinitely at room temperature. Caution: MgCl2 can be contaminated with P.

# (6) **1:1 and 1:20 ammonia solutions**

Measure in a volumetric cylinder the concentrated ammonia (NH4OH; 50 mL for the 1:1 and 100 mL for the 1:20). Pour into an appropriate glass bottle and dilute with ultrapure water (50 mL for the 1:1 and 1900 mL for the 1:20). Store at room temperature.

(7) **1 M HCl**

In a volumetric flask add 82.7 mL concentrated hydrochloric acid (HCl; 37%) in 900 mL ultrapure water and make up to 1 L.

# (8) **0.5 M HNO3**

Measure 967 mL of ultrapure water and add 33 mL of concentrated nitric acid (HNO3).

# (9) **1 M HNO3**

In a volumetric flask add 66 mL concentrated HNO3 to 800 mL ultrapure water and make up to 1 L.

# (10) **7 M HNO3**

Add 463 mL concentrated HNO3 to 537 mL ultrapure water.

# (11) **AG 50 × 8 cation resin**

Take the equivalent of 6 mL of resin per sample and place it in a plastic bottle. Add 1.5 BV of 7 M HNO3 and shake well and let rest overnight. The next day, discard the acid and wash it thoroughly using ultrapure water till pH is close to neutrality. After use, collect the resin and store it in 1 <sup>M</sup> HNO3 (66 mL concentrated HNO3 + 934 mL ultrapure water).

# (12) **Silver ammine solution**

Weigh out 10.2 g of silver nitrate (AgNO3; MW 169.87 g/mol) and 9.6 g of NH4NO3. Dissolve in 81.5 mL of ultrapure water and add 18.5 mL of concentrated NH4OH. Store in amber bottle in the dark.

Some of the above-mentioned chemicals are quite hazardous. According to the Globally Harmonized System of Classification and Labelling of Chemicals (GHS), concentrated HCl, for example, has the following hazardous statements: H290 may be corrosive to metals, H314 causes severe skin burns and eye damage and H335 may cause respiratory irritation. It is therefore advised to check the GHS hazardous and precautionary statements and the safety data sheets for the chemicals before using the protocol. It is also recommended to follow the good laboratory practice which includes wearing personal protective equipment (PPE) like lab coats, safety glasses and gloves.

# **3.3 Equipment and Consumables**


# **3.4 Procedure**

The following describes the ideal case, i.e. 1 M HCl extract with around 20 μmol P, low in organic matter and no other compounds which might interfere. How to deal with other extracts and potential problems during the purification is described in Chap. 5. The extractants are first purified in four consecutive steps (A1–A4; Fig. 3.1), before the precipitation of the final analyte in Step A5 (Fig. 3.1).

**Fig. 3.1** Stepwise purification of extracts and final precipitation of silver phosphate

#### 3 Purification Protocol 37

The first two steps (A1 and A2) are removing, for example, organic matter and are also concentrating the phosphate in the extract by reducing the volume. Certain cations could interfere with the precipitation of silver phosphate and are removed in Step A3. Silver chloride, which, if not removed, could co-precipitate with silver phosphate is removed in Step A4. The final analyte is then precipitated in Step A5. The filtration steps can be quite tedious, and using vacuum filtration equipment is therefore recommended.

# **Step A1: Ammonium phosphor-molybdate (APM) mineral precipitation and dissolution**


# **Step A2: Magnesium ammonium phosphate (MAP) mineral precipitation and dissolution**


# **Step A3: Cation removal**


# **Step A4: Silver chloride (AgCl) removal**

• Check if Cl<sup>−</sup> is still present in solution by adding a little amount of AgNO3. If a white cloud forms (AgCl crystals), wait 10 min and filter again.

# **Step A5: Ag3PO4 precipitation**


# **3.5 Preparation for TC/EA-IRMS and Data Analysis**

Following Step A5, the silver phosphate samples need to be weighed in for the measurement with a thermal conversion elemental analyser (TC/EA) coupled to a continuous flow isotope ratio mass spectrometer (IRMS). In the TC/EA the silver capsules containing the samples and standards are pyrolysed at 1450 °C and the oxygen is converted into CO. The CO is then, via a helium stream, transported through a GC column which is separating CO from any nitrogen (N2) that might be present to the IRMS. In the IRMS the isotopic composition of CO is then measured. Before each run, the linearity of the IRMS should be checked and this will inform us about the range of weight that should be used for the silver phosphate standards and samples. In general, most instruments are linear between 250 and 350 μg of silver phosphate; however, linearity should still be checked (Carter and Barwick 2011). Likewise, a stability check of the IRMS should be done before each run (Carter and Barwick 2011). Further information about normalization and selection of reference materials can also be found in Skrzypek (2013) and Skrzypek and Sadler (2011).


#### 3 Purification Protocol 39

even if it was added to the silver phosphate samples and standards. Close tightly by forming little balls using tweezers and gloves (Werner and Brand 2001).


During the whole preparation process neither the crystals nor the silver capsules should be touched with bare hands in order to avoid contamination with any Ocontaining compounds.

A sample sequence might look as follows (see the Annex for an example):

	- 12 samples,
	- 1 silver phosphate standard,
	- 1 IAEA-601 and
	- 12 samples.
	- 4× certified silver phosphate standard and
	- Additional standard.

Once the δ18OP values are analysed with the TC/EA-IRMS, a quality check needs to be done. The first three steps of the data analysis therefore are


Oxygen yield above or below expectation implicates that the silver phosphate sample was not pure silver phosphate and hence the obtained δ18OP value might be erroneous. To check for the purity of the Ag3PO4, we control the O% given by the TC/EA. The silver phosphate standards weighed at different amounts provide a calibration curve used to determine the O content of all samples. If the analysed Ag3PO4 samples are pure, their O content should be in the range of the Ag3PO4 standard.

Similarly, a contamination with N2 could also lead to erroneous δ18OP values. Either due to interference during the mass spectrometer measurements or because of N compounds containing oxygen (Pederzani et al. 2020). Sometimes, if the final wash of the Ag3PO4 is not done properly, NO3 could remain on the crystals. The presence of N2 is checked by looking at the chromatogram from the TCEA and from the IRMS. The N2 peak should be identified before the CO peak. Only when the first two checks have been passed, an average δ18OP value can be calculated.

The isotopic values of the certified standards should remain stable along the run. If this is not the case, a drift correction needs to be done (Carter and Barwick 2011). First the drift needs to be calculated:

$$\text{Dift} = \left( \text{Average}\_{\text{STD10}-12} - \text{Average}\_{\text{STD1}-4} \right) / \left( \text{Position}\_{\text{STD10}} - \text{Position}\_{\text{STD1}} \right), \tag{3.1}$$

where AverageSTD10–12 is the average δ18OP value of the last three replicates of the certified silver phosphate standard in a run, AverageSTD1–4 is the average δ18OP value of the first four replicates of the certified silver phosphate standard in a run, PositionSTD10 is the position of the 10th certified silver phosphate standard in a run and PositionSTD1 is the position on the 1st certified silver phosphate standard in a run.

The drift can then be used to perform a drift correction of the δ18O values:

$$\delta^{18}\mathcal{O}\_{\text{D\\_corr}} = \delta^{18}\mathcal{O}\_{\text{meas}} - \text{drift} \cdot \left(\text{Position}\_{\text{sample}} - \text{Position}\_{\text{STD1}}\right),\tag{3.2}$$

where δ18Omeas is the measured δ18O value of a sample/standard, drift is the previously calculated drift value, Positionsample is the position of a sample/standard in a run and PositionSTD1 is the position on the 1st certified silver phosphate standard in a run.

After drift correction of samples and standards, average measured and drift corrected values of the standards should be correlated to the reported values (Brand et al. 2009). Using the slope and intercept of this correlation, calculate the real value of the samples.

A good practice is to compare the standard deviation of each sample to the standard deviations of the silver phosphate and benzoic acid standards within the same run. Standard deviations are around ±0.3‰ (Brand et al. 2009; Halas et al. 2011; Watzinger et al. 2021). Only if the standard deviation of a sample is equal or smaller than the standard deviations of the standards, a δ18OP value can be trusted. In case oxygen yield is outside the acceptable range and/or N2 contamination was found, the silver phosphate sample can be treated with H2O2 (removing organic compounds), thoroughly rinsed again with ultrapure water (removing N-containing compounds) and/or homogenized better (in case of crystals with different sizes). If the oxygen yield and N2 are ok, but the standard deviation (SD) of the replicates is still high (higher than SD of certified standard), then repeat analysis and possibly homogenize sample better. Vacuum roasting is also sometimes used to remove contamination with other O-bearing compounds; however, this procedure also has some issues including potential reoxidation when samples are stored too long (Mine et al. 2017).

# **Glossary**


# **Appendix**

See Fig. 3.2 below.

**Fig. 3.2** Example for a TC/EA-IRMS run. This run includes a certified silver phosphate standard (Ag3PO4\_certified), two internal silver phosphate standards (Ag3PO4\_internal A and B) and one benzoic acid standard (IAEA-601). Each sample is weighed in triplicates (rep1-3) and six empty silver capsules are included as blanks at the beginning of the run


# **References**


The opinions expressed in this chapter are those of the author(s) and do not necessarily reflect the views of the IAEA: International Atomic Energy Agency, its Board of Directors, or the countries they represent.

**Open Access** This chapter is licensed under the terms of the Creative Commons Attribution 3.0 IGO License (https://creativecommons.org/licenses/by/3.0/igo/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the IAEA: International Atomic Energy Agency, provide a link to the Creative Commons license and indicate if changes were made.

Any dispute related to the use of the works of the IAEA: International Atomic Energy Agency that cannot be settled amicably shall be submitted to arbitration pursuant to the UNCITRAL rules. The use of the IAEA: International Atomic Energy Agency's name for any purpose other than for attribution, and the use of the IAEA: International Atomic Energy Agency's logo, shall be subject to a separate written license agreement between the IAEA: International Atomic Energy Agency and the user and is not authorized as part of this CC-IGO license. Note that the link provided above includes additional terms and conditions of the license.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# **Chapter 4 Modifications and Issues During Purification**

**V. Pfahler, J. Adu-Gyamfi, A. Watzinger, and F. Tamburini**

**Abstract** Depending on the extract, it is necessary to modify the purification protocol slightly. Each sample is different and despite a thorough testing of the purification protocol, issues might occur. The three modifications suggested include (1) adjustments in pH, (2) magnesium ammonium phosphate (MAP) precipitation and (3) reductions, prior to A1, of cations like iron (Fe), silica (Si) and calcium (Ca) which could cause interferences during the purification process. Some of the major issues often encountered are (1) no APM precipitation due to the presence of high carbonate concentrations, (2) the presence of high organic matter that requires additional steps in the protocol, (3) crystals not dissolving and (4) discoloration of solution.

# **4.1 Method Modifications During the Purification Process**

The original purification protocol is using 1 M HCl extracts but it is nowadays used for a diverse set of extracts (Tamburini et al. 2010, 2018; Pistocchi et al. 2017; Granger et al. 2018; Pfahler et al. 2020). Depending on the extract it is necessary to modify the purification protocol slightly. Also, each sample is different and despite thorough testing of the purification protocol, issues might occur.

V. Pfahler

J. Adu-Gyamfi (B)

A. Watzinger

Formerly Smithsonian Tropical Research Institute, Ancón, Panama

Soil and Water Management & Crop Nutrition Section, Joint FAO/IAEA Centre of Nuclear Techniques in Food and Agriculture, Vienna, Austria e-mail: J.Adu-Gyamfi@iaea.org

Department of Forestry and Soil Sciences, Institute of Soil Research, University of Natural Resources and Life Sciences, Tulln, Austria

F. Tamburini Group of Plant Nutrition, D-USYS, ETH, Zurich, Switzerland

J. Adu-Gyamfi and V. Pfahler (eds.), *Oxygen Isotopes of Inorganic Phosphate in Environmental Samples*, https://doi.org/10.1007/978-3-030-97497-8\_4

# *4.1.1 Modification 1*

The ideal pH for the precipitation of ammonium phosphomolybdate (APM; step A1 of purification protocol) is 1. When using acidic reagents that are weaker than 1 M HCl, for example, 0.2 M HNO3 for eluting resin P, it is therefore necessary to add concentrated sulphuric acid (H2SO4). Concentrated H2SO4 is a strong acid that could potentially hydrolyse any organic or condensed P present in the extract. Add concentrated H2SO4 slowly to the sample only after ammonium molybdate and ammonium nitrate are added to the extract in step A1 of the purification protocol. Usually, 1 ml concentrated H2SO4 is sufficient to facilitate the precipitation of APM.

# *4.1.2 Modification 2*

If the initial extract, e.g., 0.5 M NaHCO3, has an alkaline pH, start with step A2 to avoid unnecessary pH adjustment. Indeed, step A2 of the purification protocol (magnesium ammonium phosphate (MAP) precipitation) requires an alkaline pH (>7). In these cases, the crystals obtained from the MAP step are never properly clean. So, after the dissolution of the MAP crystals, proceed with step A1, repeat step A2 and continue with the remaining steps.

# *4.1.3 Modification 3*

Certain cations like iron (Fe), silica (Si) and calcium (Ca) can interfere with the purification protocol and therefore their concentrations need to be reduced prior to Step A1. High concentrations of Si can be present in volcanic soils and can interfere with the molybdate complexation in step A1 of the purification protocol. Ca can also interfere in step A1 due to the formation of crystals with molybdate. Fe could form colloids and co-precipitated with organic P. Brucite precipitation should be added before step A1 to reduce the concentrations of interfering cations. Alternatively, cation exchange resins could help in reducing the cation concentrations; however, this has, to the best of our knowledge, not yet been tested in case of the purification protocol.

# **4.2 Major Issues**

# *4.2.1 Issue 1: No APM Precipitation*

This is often the case when the initial extract is not acidic enough. In most cases modification 1 will help. In the presence of high carbonate concentrations, even the 1 M HCl extract might not be acidic enough (Pistocchi et al. 2017). Pistocchi et al. (2017) therefore adjusted the liquid-to-solid ratio for their samples during the extraction to 100:1 instead of 50:1. Too low P concentrations (<10 μmol) can also be a reason for no APM precipitation. In that case, the sample needs to be extracted again and a higher amount of material needs to be used. In this case, it is possible that multiple subsamples are extracted and then the extracts are combined. A brucite step is most likely needed in order to reduce the extract volume prior to APM precipitation.

# *4.2.2 Issue 2: High Organic Matter*

In most cases, for example with low organic matter but high inorganic P concentrations, steps A1 and A2 are sufficient to remove organic matter from the initial extracts. Sometimes steps A1 and A2 are not sufficient and it is necessary to use the DAX-8 resin or brucite precipitation before proceeding with the purification protocol. Organic matter or colouration is also an issue in Olsen P extracts and charcoal is used to deal with this issue. However, charcoal is often contaminated with P and hence not ideal for the δ18OP method, unless acid cleaned and checked for P.

**For conditioning the DAX-8 resin, proceed as follows**: Take the equivalent of 10 ml of resin per sample and place it in a 500 ml plastic bottle. To condition the new resin, use 1.5 bed volumes (BV) of methanol, shake well, and let rest for 15 min. Discard methanol, rinse with 1.5 BV of ultrapure water, shake and let rest for 15 min. Carefully discard water. Add ultrapure water just to cover the surface of the resin. After use, the resin should be collected and washed with 1 M HCl + methanol. Store in methanol and at room temperature.

# *4.2.3 Issue 3: Crystals not Dissolving and Discolouration of Solution*

Sometimes the ammonium phosphomolybdate (APM), which formed during step A1, does not dissolve immediately in the ammonium citrate solution (step A2). In this case, leave the solution for about 1 h and if it did not dissolve by then, filter it before continuing with the remaining parts of step A2. Likewise, magnesium ammonium phosphate (MAP), which formed during step A2, does not dissolve immediately

**Fig. 4.1** Dissolved, but decoloured ammonium phosphomolybdate (APM)

when adding 0.5 M HNO3. Leave the solution for one hour; if not dissolved by then, filter the solution.

Another issue, which might occur, is the discolouration of the solution (Fig. 4.1). Dissolved APM and MAP should yield clear solutions. Filtering the solutions prior to continuing with the respective step (A2 or A3), usually reduces the discolouration.

# **Glossary**


4 Modifications and Issues During Purification 49

# **References**


The opinions expressed in this chapter are those of the author(s) and do not necessarily reflect the views of the IAEA: International Atomic Energy Agency, its Board of Directors, or the countries they represent.

**Open Access** This chapter is licensed under the terms of the Creative Commons Attribution 3.0 IGO License (https://creativecommons.org/licenses/by/3.0/igo/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the IAEA: International Atomic Energy Agency, provide a link to the Creative Commons license and indicate if changes were made.

Any dispute related to the use of the works of the IAEA: International Atomic Energy Agency that cannot be settled amicably shall be submitted to arbitration pursuant to the UNCITRAL rules. The use of the IAEA: International Atomic Energy Agency's name for any purpose other than for attribution, and the use of the IAEA: International Atomic Energy Agency's logo, shall be subject to a separate written license agreement between the IAEA: International Atomic Energy Agency and the user and is not authorized as part of this CC-IGO license. Note that the link provided above includes additional terms and conditions of the license.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# **Chapter 5 How to Design a Study Including the Analysis of δ18OP**

**V. Pfahler, J. Adu-Gyamfi, and F. Tamburini**

**Abstract** To plan a research study, one needs to (1) establish a research question, (2) make a set of observations, (3) form a hypothesis in an attempt to explain the observations and (4) test the hypothesis based on the data collected. The following questions should be addressed when designing a study including the analysis of δ18OP: (i) what is the research hypothesis? (ii) what is the main objective of the study? (iii) what are the aims to address these objectives? and (iv) which techniques are appropriate to address such research question. In addition, one needs to consider (1) which kind of samples needs to be collected, e.g. soil, vegetation or water? (2) in case of soil and sediment samples, which sampling depths and increments need to be sampled? (3) which P pools need to be extracted and analysed for the corresponding δ18OP values? (4) when and how often should samples be taken and (5) how many samples can be processed per week?

# **5.1 Planning or Designing a Study**

To plan a research study, one needs to (1) establish a research question, (2) make a set of observations, (3) form a hypothesis in an attempt to explain the observations and (4) test the hypothesis based on the data collection.

The following questions should be addressed when designing a study including the analysis of δ18OP:

V. Pfahler

J. Adu-Gyamfi (B)

Soil and Water Management & Crop Nutrition Section, Joint FAO/IAEA Centre of Nuclear Techniques in Food and Agriculture, Vienna, Austria e-mail: J.Adu-Gyamfi@iaea.org

Formerly Smithsonian Tropical Research Institute, Ancón, Panama

F. Tamburini Group of Plant Nutrition, D-USYS, ETH, Zurich, Switzerland

J. Adu-Gyamfi and V. Pfahler (eds.), *Oxygen Isotopes of Inorganic Phosphate in Environmental Samples*, https://doi.org/10.1007/978-3-030-97497-8\_5


In addition to the above questions, (1) monitoring of progress towards results, resources consumed and budget, (2) a reflection and a shared experience and lessons drawn from success and failure, should be considered a set of observations, (3) form a hypothesis in an attempt to explain the observations and (4) test the hypothesis based on the data collection.

# **5.2 Comments to the Questions**


Once questions 1 to 6 are answered, one could think about analyses that complement the δ18OP analysis. Essential analyses which should always be included are:


Depending on the hypothesis, it might also be useful to include further analyses. The δ18OP has been successfully combined, in incubation and glasshouse studies, with 33P in soils (Helfenstein et al. 2018; Siegenthaler et al. 2020) and plants (Pfahler et al. 2017). Also, the combination with metagenomics and other microbial analyses becomes more popular (Bi et al. 2018; Shen et al. 2020). Table 5.1 shows examples of analyses and data typically used along the δ18OP for investigating P cycling in the environment.

# **5.3 External Quality Assurance/Control**

Quality assurance of isotope measurements by TC/EA-IRMS is based on the trueness and precision of the values from external standards that are analysed along with the samples in daily sequences (Watzinger et al. 2021). Most commonly, researchers use two benzoic acid standards provided by the IAEA (IAEA-601 with a δ18O value of 23.14‰ and IAEA-602 with a δ18O value of 71.28‰), a commercially bought silver phosphate, and sometimes also silver phosphate produced in-house. Before a new batch of silver phosphate, bought or produced in-house, is used as a standard, it should be sent to at least one additional laboratory for cross-validation of the δ18OP value.

An inter-laboratory study for silver phosphate standards was conducted by Watzinger et al. (2021). A silver phosphate reference material was produced by the University of Natural Resources and Life Science (Austria) and sent to four other laboratories: The University of Western Australia (Australia), the ETH Zurich (Switzerland), the University of Helsinki (Finland) and the Helmholtz Centre for Environmental Research (Germany). This new reference material has an δ18OP value of 13.8 ± 0.3‰ and is available for research laboratories.

In addition, the purification protocol should be tested in an inter-laboratory study using different materials: pure potassium dihydrogen phosphate and silver phosphate as a control and a set of different dried soil and sediment samples. A water sample would also be nice; however as water samples should be processed right away after sampling, storing and sending water samples to different laboratories might alter the δ18OP signature due to an extended storage period.


# **5.4 Interpretation of Isotopic Data from the TC/EA-IRMS and Its Applications**

There is no one-fits-all approach when it comes to the interpretation of δ18OP data especially since not all variables influencing δ18OP values are known. How to interpret δ18OP data strongly depends on the research question, the research subject (soil, lake water, marine sediments etc.), and the data itself. However, there are some general steps/rules which can help to interpret the δ18OP data:


A good starting point for the interpretation of δ18OP data is the comparison with the theoretical equilibrium value. Values lower than the equilibrium value are often an indication for hydrolysis of organic P but could also be due to other P inputs with a lower δ18OP value like P from igneous rocks. Values higher than the equilibrium value could for example be caused by inorganic P leached from plants as plant inorganic P δ18OP values tend to be enriched in 18O compared to other P pools (Table 5.1).

The following examples are very simple case studies and only show the general workflow when designing a δ18OP study.

# **5.5 Example Research Study—Small Lake**

A small lake, surrounded by five agricultural fields, is suffering from eutrophication during the summer. Each of the agricultural fields has a drainage didge which flows into the lake. Three of those agricultural fields are fertilized with mineral P fertilizers, the other two with farm-yard manure. One river, with a wastewater treatment plant (WWTP) upstream, is fed into the lake.

	- a. Eight water samples
	- b. Five fertilizer samples

To calculate the total number of samples for δ18OP analysis, one needs to now consider the following:

	- a. **Eight** water samples (SRP only)
	- b. Two P pools for each fertilizer sample, including a factor 2 for the HCl P due to using 18O-labelled and unlabelled HCl: **15 samples**
	- c. Two P pools for each soil sample, including a factor 2 for the HCl P due to using 18O-labelled and unlabelled HCl: **15 samples**.

In total, 76 samples need to be analysed and afterwards purified, with **38 samples** at each sampling time point, and 38 samples cannot be analysed within one week. It is recommended to first deal with the samples/extracts which are more susceptible to changes due to biological activities. One would therefore start with the water samples and extract the more labile P pool (water-extractable or resin P) from the soils and farm-yard manure.

# **5.6 Example Research Study—Agricultural Fields**

To investigate P cycling at agricultural fields where rapeseed is cultivated, five agricultural fields are selected as study sites. Those five fields are in the same climatic zone, but differ in their soil properties, like soil pH and P saturation index. All five agricultural fields are fertilized with the same mineral P fertilizers.

	- a. One fertilizer sample
	- b. Soil samples from each field at different depths
	- c. Only above-ground plant sample

To calculate the total number of samples for δ18OP analysis, one needs to now consider the following:

	- a. Five P pools for each soil sample, including a factor 2 for the HCl P due to using 18O-labelled and unlabelled HCl: **60 samples**
	- b. One bulked plant sample; two P pools; including a factor 2 for the TCA P due to using 18O-labelled and unlabelled TCA: **Three samples**.

In total, 318 samples need to be analysed and afterwards purified, with **63 samples** at each field and three samples from the fertilizer. Those samples cannot be analysed within one week. It is recommended to first deal with the samples/extracts which are more susceptible to changes due to biological activities. One would therefore start with extracting the most labile P pool (resin and microbial P) from the soils. It might also be useful to analyse other parameters relevant for P cycling like enzyme activities.

# **Glossary**

δ**18O** The oxygen isotope ratio is conventionally given in the delta notation: δ18O = (Rsample/Rstandard) − 1, where Rsample is the 18O/16O ratio of a sample and Rstandard is the 18O/16O ratio of the Vienna Standard Mean Ocean Water (V-SMOW). δ18OP is the δ18O value of a P compound or pool.

**Soluble reactive P** SRP; considered the most bio-available P pool in water samples.


# **References**


The opinions expressed in this chapter are those of the author(s) and do not necessarily reflect the views of the IAEA: International Atomic Energy Agency, its Board of Directors, or the countries they represent.

**Open Access** This chapter is licensed under the terms of the Creative Commons Attribution 3.0 IGO License (https://creativecommons.org/licenses/by/3.0/igo/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the IAEA: International Atomic Energy Agency, provide a link to the Creative Commons license and indicate if changes were made.

Any dispute related to the use of the works of the IAEA: International Atomic Energy Agency that cannot be settled amicably shall be submitted to arbitration pursuant to the UNCITRAL rules. The use of the IAEA: International Atomic Energy Agency's name for any purpose other than for attribution, and the use of the IAEA: International Atomic Energy Agency's logo, shall be subject to a separate written license agreement between the IAEA: International Atomic Energy Agency and the user and is not authorized as part of this CC-IGO license. Note that the link provided above includes additional terms and conditions of the license.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# **Chapter 6 Conclusions and Way Forward**

**J. Adu-Gyamfi, V. Pfahler, L. Heng, and F. Tamburini**

**Abstract** With an increasing number of researchers using the δ18OP method to investigate P cycling in the environment, it is necessary to conduct an inter-laboratory comparison study for the purification protocol as well as the measurement of silver phosphate with the TC/EA-IRMS like Watzinger et al. (2021) did. For the δ18OP method to progress, further fundamental research as well as field and laboratory studies need to be conducted. To the best of our knowledge, the effect of synthesizing enzymes on the δ18OP has not been investigated yet, despite the importance of those enzymes in the P cycle.

# **6.1 Future Trends and Opportunities**

With increasing fertilizer prices and the negative environmental issues such as eutrophication of aquatic systems and phosphorus contaminations in surface and groundwater that affect water quality, it is now, more than ever, vital to better understand the environmental P cycle. Studies during the last 10 years showed that the δ18OP method is a promising tool to study the environmental P cycle under field conditions and the importance of microbes for available P in soils (Tamburini et al. 2012). The oxygen isotopes in phosphates have helped in identifying P inputs in Lake Erie and Chesapeake Bay, some of the aquatic systems most struck by eutrophication (Elsbury et al. 2009; Joshi et al. 2015; Paytan et al. 2017; Depew et al. 2018). It has the potential to track P derived from glyphosate in the environment (Li et al. 2016),

J. Adu-Gyamfi (B) · L. Heng

Soil and Water Management & Crop Nutrition Section, Joint FAO/IAEA Centre of Nuclear Techniques in Food and Agriculture, International Atomic Energy Agency (IAEA), Vienna, Austria

e-mail: J.Adu-Gyamfi@iaea.org

V. Pfahler Formerly Smithsonian Tropical Research Institute, Ancón, Panama

F. Tamburini Group of Plant Nutrition, D-USYS, ETH, Zurich, Switzerland

J. Adu-Gyamfi and V. Pfahler (eds.), *Oxygen Isotopes of Inorganic Phosphate in Environmental Samples*, https://doi.org/10.1007/978-3-030-97497-8\_6

to name just a few highlights. The δ18OP could also provide parameters for environmental models (Jaisi et al. 2017). Modeling P cycling and including P in carbon and nitrogen models is becoming more important and the interest to also include P in life cycle assessments is increasing, especially the mineralization of organic P and its quantification are of interest for modelers (Vereecken et al. 2016; Helfenstein et al. 2018; Thum et al. 2019).

# **6.2 Need for Inter-laboratory Studies for Quality Control**

With an increasing number of researchers using the δ18OP method to investigate P cycling in the environment, it is necessary to conduct an inter-laboratory comparison study for the purification protocol as well as the measurement of silver phosphate with the TC/EA-IRMS like Watzinger et al. (2021) did. While nowadays silver phosphate is the analyte of choice, there are different protocols for purifying extracts and precipitating silver phosphate (see, e.g., Paytan and McLaughlin 2011). Even when using the same protocol, every laboratory is most likely amending the protocol for their needs, using different equipment, consumables, and chemicals. For better comparability of different δ18OP studies, there is a need for an inter-laboratory study similar to other protocols, like for the extraction of microbial P (Bergkemper et al. 2016). Equally important is an international silver phosphate standard which should be easily available to most laboratories (Watzinger et al. 2021).

# **6.3 From Laboratory to Field Studies in Different Ecotones**

For the δ18OP method to progress, further fundamental research, as well as field and laboratory studies, need to be conducted. To the best of our knowledge, the effect of synthesizing enzymes on the δ18OP has not been investigated yet, despite the importance of those enzymes in the P cycle. Furthermore, the effects of some processes like P uptake on the δ18OP were only shown in laboratory studies but are missing proof from field studies. The majority of δ18OP studies so far was conducted in countries in the northern hemisphere. An increase of studies in the southern hemisphere is therefore desirable. This will advance the knowledge about the variables influencing δ18OP values as countries in the global South tend to have different ecosystems, climates, and soil types compared to the North. Two aspects that need to be addressed in further field studies are the importance of soil P concentrations, i.e., P limitation and surplus, and microorganisms (community structure and activity) for the δ18OP of different soil P pools. Is there for example a relationship between P concentrations and δ18OP values and could thus the δ18OP be used as an indicator for P limitation? This will not only help with the interpretation of δ18OP data but will ultimately lead to a better understanding of P cycling in the environment.

# **References**


The opinions expressed in this chapter are those of the author(s) and do not necessarily reflect the views of the IAEA: International Atomic Energy Agency, its Board of Directors, or the countries they represent.

**Open Access** This chapter is licensed under the terms of the Creative Commons Attribution 3.0 IGO License (https://creativecommons.org/licenses/by/3.0/igo/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the IAEA: International Atomic Energy Agency, provide a link to the Creative Commons license and indicate if changes were made.

Any dispute related to the use of the works of the IAEA: International Atomic Energy Agency that cannot be settled amicably shall be submitted to arbitration pursuant to the UNCITRAL rules. The use of the IAEA: International Atomic Energy Agency's name for any purpose other than for attribution, and the use of the IAEA: International Atomic Energy Agency's logo, shall be subject to a separate written license agreement between the IAEA: International Atomic Energy Agency and the user and is not authorized as part of this CC-IGO license. Note that the link provided above includes additional terms and conditions of the license.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.