Development of liquid chromatography mass spectrometry method for analysis of organic N monomers in soil

Soil Biology & Biochemistry 78 (2014) 233e242

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Development of liquid chromatography mass spectrometry method for analysis of organic N monomers in soil Charles R. Warren* School of Biological Sciences, The University of Sydney, NSW 2006, Australia

a r t i c l e i n f o

a b s t r a c t

Article history: Received 10 July 2014 Received in revised form 8 August 2014 Accepted 9 August 2014 Available online 27 August 2014

The aim of this study was to develop an analytical procedure based on liquid chromatography-mass spectrometry (LCeMS) for analysis of monomeric organic N compounds in soil extracts. To benchmark the developed LCeMS method it was compared with a capillary electrophoresisemass spectrometry (CE eMS) method recently used for analysis of small organic N monomers in soil. The separation was optimized and analytical performance assessed with 69 purified standards, then the LCeMS method was used to analyse soil extracts. Sixty-two out of 69 standards were analysable by LCeMS with separation on a hydrophilic interaction liquid chromatography column. The seven compounds that could not be analysed were strongly cationic polyamines. Limits of detection for a 5 mL injection ranged between 0.002 and 0.38 mmol L
1, with the majority (49 out of 62) having limits of detection better than 0.05 mmol L
1. The overall profile and concentration of small organic N monomers in soil extracts was broadly similar between LCeMS and CEeMS, with the notable exception of four ureides that were detected by LCeMS only. In soil extracts that had been concentrated ten-fold the detection and quantification of (some) organic N compounds was compromised by the presence of large amounts of inorganic salts. The developed LCeMS method offered advantages and disadvantages relative to CEeMS, and a combination of the two methods would achieve the broadest possible coverage of organic N in soil extracts. © 2014 Elsevier Ltd. All rights reserved.

Keywords: LCeMS HILIC CEeMS Soil extract Organic N Microbial biomass

1. Introduction The profile and concentration of organic N molecules in soil solution and soil microbes are information-rich signals indicative of soil function. This is because organic N compounds in soil solution reflect biological processes such as uptake and efflux of organic molecules by plant roots and free-living and symbiotic microbes, activity of extracellular enzymes, and leaching of organic molecules from litter. In recent times the profile of organic N molecules in the soil solution has become of particular interest with the demonstration that plants can directly take up various small organic forms €sholm et al., of N (Chapin et al., 1993; Jones and Darrah, 1993; Na 2009; Warren, 2013b) and recognition that knowledge of the identity and concentration of compounds in soil solution is key to designing and interpreting experiments on organic N uptake (Warren, 2014a). Knowledge of organic N in soil microbes holds great promise as an indicator of function of the microbial community. For example, stresses such as water deficits and

* Corresponding author. Tel.: þ61 2 9351 2678; fax: þ61 2 9351 4119. E-mail address: [email protected]. http://dx.doi.org/10.1016/j.soilbio.2014.08.008 0038-0717/© 2014 Elsevier Ltd. All rights reserved.

freezeethaw cycles that have strong effects on the physiology and composition of the microbial community (Schimel et al., 2007) ought to also affect the composition of organic compounds in soil microbes (e.g. concentrations of osmolytes: Warren, 2014c). The most common approach for exploring organic N monomers in soil has involved use of methods for identification and quantification of amino acids. Various chromatographic and electrophoretic methods have been used successfully for separation and quantification of primary (and in some cases secondary) amino acids in soil extracts (e.g. Kielland, 1995; Yu et al., 2002; Warren, 2008; Farrell et al., 2011). These methods are generally based upon highly selective derivatization and/or detection schemes. The downside of this selectivity is that the methods are effectively blind to many of the other compound classes of organic N present in soil (Warren, 2013a). To address questions such as what organic N compounds are used for osmotic adjustment or what organic N compounds are taken up by plants requires a broader exploration of organic N than is possible with methods that target amino acids only. Few methods have been developed for the comprehensive analysis of organic N monomers in soil-based samples, at least in part because the properties of organic N in soil limit the application

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of common analytical platforms such as nuclear magnetic resonance (NMR) and gas chromatography (GC). NMR spectroscopy shows inherent limitations in terms of detection sensitivity (Gika et al., 2012) and thus when applied to soil will be able to detect only the most abundant compounds. Gas chromatography is unsuitable for broad-based exploration of organic N in soil because many organic N molecules (e.g. quaternary ammonium compounds, oligomers and polymers) are not volatile and cannot be derivatized using common reagents, or are degraded during passage through the hot injection port and column (Kaspar et al., 2009). Capillary electrophoresis in combination with electrospray ionization and mass spectrometry (hereafter referred to as CEeMS) has recently been demonstrated as a useful tool for profiling small organic N compounds in soil extracts (Warren, 2014a). The major advantages of CEeMS are that compounds do not need to be volatile or derivatized, and it can separate molecules of widely varying polarity while also resolving isomeric and isobaric ions (Kuehnbaum and Britz-McKibbin, 2013). However, the widespread adoption of CEeMS may be prevented by some limitations. First, the described CEeMS separation (Warren, 2014a) was limited to those compounds that were cationic at the electrolyte's pH (~2). Organic N compounds that are weakly cationic (e.g. ergothioneine) yield broad peaks and poor detection limits, while compounds that are neutral (e.g. N-acetylglucosamine) or anionic (e.g. ureides) cannot be analysed. The second factor that may limit the more widespread adoption of CEeMS is the comparatively poor reproducibility of absolute migration time (Ramautar et al., 2009; Sugimoto et al., 2010), which complicates data analysis because the same peak can appear at (slightly) different migration times in replicate analytical runs. The final factor limiting use of CEeMS is that the installed base of CEeMS instruments remains small, and thus there is limited access to instrumentation and expertise. Liquid chromatography in combination with electrospray ionization and mass spectrometry (hereafter referred to as LCeMS) may offer an alternative to CEeMS for analysis of small organic N compounds. One of the biggest assets of LCeMS is that the installed base of LCeMS instruments is large, and thus instrumentation and expertise are widely accessible. Unfortunately, LCeMS of hydrophilic compounds such as small organic N compounds is far from straightforward. The challenge arises because organic N compounds exhibit negligible retention or separation on the reversed phase columns commonly used for LCeMS. Retention and separation can be improved by adding ion pairing reagents to the mobile phase (Hakkinen et al., 2007; Lu et al., 2008; Sanchez-Lopez et al., 2009), but methods utilizing ion pairing reagents are generally unsuitable because ion pairing reagents suppress mass spectrometry signals and lead to persistent contamination of the mass spectrometer's ion source (Rutters et al., 2000; Holcapek et al., 2004). Another alternative is to derivatize compounds so as to improve their chromatography (and in many cases also detection limits) (Inagaki et al., 2010; Murphy et al., 2014), but not all compounds can be derivatized and thus derivatization cannot achieve a sufficiently large breadth of coverage. Two separation modes capable of LCeMS of hydrophilic compounds without ion pairing or derivatization are hydrophilic interaction liquid chromatography (HILIC) (Alpert et al., 1994) and aqueous normal phase liquid chromatography (Pesek and Matyska, 2007). In recent times LCeMS with HILIC separations have been used by numerous groups for analysis of complex mixtures of hydrophilic compounds (Lu et al., 2008; Kato et al., 2009; Schiesel et al., 2010; Creek et al., 2011; Iwasaki et al., 2011; Rappold and Grant, 2011; Boudra et al., 2012; Buszewski and Noga, 2012; Chen et al., 2012; Fraser et al., 2012; Zhang et al., 2012) but it is unclear if methods developed for other biological samples (e.g. animals, plants, microbial

cultures) can be directly applied to soil. Soil-based samples are particularly challenging because concentrations of organic N compounds are typically several orders of magnitude smaller than concentrations of (potentially interfering) inorganic ions (Oikawa et al., 2011; Warren, 2014a). The aim of this study was to develop an analytical procedure based on LCeMS with HILIC separation for analysis of monomeric organic N compounds in H2O extracts and 2.5% CHCl3 extracts of soil. To benchmark the developed LCeMS approach it was compared with a CEeMS approach recently demonstrated as useful for broad-based analysis of small organic N monomers in soil (Warren, 2013a). To provide a fair benchmarking of performance, common samples and standards were analysed on both analytical platforms and the same mass spectrometer was used for LCeMS and CEeMS. 2. Methods 2.1. Experimental design To characterize analytical performance the same standards and samples were analysed by LCeMS and CEeMS. The first test of performance involved injecting 69 purified standards (see Section 2.2) so as to determine which organic N monomers could be analysed, their detection limits, and the shape and width of peaks. Special attention was paid to the separation of five pairs of structural isomers (Fig. 1). To examine applicability of LCeMS and CEeMS to soil-based samples, two types of soil extract (see Section 2.3) were analysed. Soil extracted with ultra-pure water was used to provide a sample indicative of compounds in free solution, while soil extracted with aqueous 2.5% (v/v) CHCl3 was used to provide a sample indicative of compounds present in microbial biomass. These two types of soil extract were then analysed a) after preparation in appropriate injection buffer/solvent but with no pre-concentration and, and b) after 10-fold pre-concentration. The final tests of analytical performance involved determining the repeatability of LCeMS and CEeMS analyses of a soil extract. Five replicate injections of the same 2.5% CHCl3 extract of soil were used to calculate for each peak the precision of concentration estimates, and precision of retention time (LCeMS) or migration time (CEeMS). 2.2. Chemicals and standards Methanol, acetonitrile and formic acid were LC/MS grade; while ammonium formate, ammonium hydroxide (28e30% NH3), and chloroform were analytical grade. All electrolytes, rinsing solutions, standards and extracts were prepared with 18.2 MU cm ultra-pure water (Arium 611UV, Sartorius, Goettingen, Germany). The majority of method development was carried out with 69 purified standards of monomeric organic N compounds. Standards were prepared from their free acids or salts purchased from Sigma. All standards of chiral amino acids were L enantiomers. Stock solutions of 1 mg mL
1 were made in ultra-pure water or 0.1 M HCl. To determine which compounds were analysable, the individual stock solutions were combined into five different mixtures at concentrations of 20 mg mL
1 (Table S1). Mixtures were prepared ensuring that there were no compounds with the same nominal mass within each mixture, and thus assignment of peaks was generally unambiguous. In cases where peaks could not be assigned unambiguously, individual standards were analysed separately. Finally, in a few instances where compounds (apparently) present in a mixture did not appear in a chromatogram or electropherogram, the mixture was subsequently analysed by direct infusion-mass

C.R. Warren / Soil Biology & Biochemistry 78 (2014) 233e242

sample of approximately 500 g of soil was collected from 0 to 15 cm depth by taking and combining multiple cores from each of eight mesocosms. The mesocosms from which soil was collected were not involved in a previous drought experiment (Warren, 2014c), but had instead been watered every 7e21 days since 2009. The composite soil sample was mixed and roots were removed prior to extraction. To provide a sample indicative of compounds in free solution, soil was extracted with ultra-pure water; whereas to provide a sample indicative of compounds present in microbial biomass, soil was extracted with aqueous 2.5% (v/v) CHCl3. Direct extraction with an aqueous solution containing chloroform leads to lysis of microbial membranes and concomitant extraction of microbial metabolites, and avoids some of the problems that can arise from the more traditional chloroform gas fumigation (Setia et al., 2012). Aliquots of 5.0 g of soil were extracted with 25.0 mL of ultra-pure water or 25.0 mL of 2.5% (v/v) CHCl3 in 50-mL centrifuge tubes (62.559, Sarstedt, Nümbrecht, Germany). Extracts were shaken at 100 rpm for 10 min at room temperature. Samples were pipetted into multiple 2-mL microfuge tubes (Safe-Lock, Eppendorf, Hamburg, Germany), rapidly centrifuged (1 min at 16,000 g) and then supernatants were frozen at
80  C. Blanks were also extracted and carried through the analysis procedures. The multiple replicates of the composite soil extracts and blanks were kept at
80  C and defrosted immediately before analysis.

a) 90.05 ± 0.1 Da Ala

β-Ala

b) 104.10 ± 0.1 Da

GABA

dimethylglycine

intensity

c) 118.10 ± 0.1 Da

betaine

Val

d) 132.10 ± 0.1 Da Leu

235

Ile

2.4. Analysis by capillary electrophoresisemass spectrometry e) 146.10 ± 0.1 Da acetylcholine

5

10

γ-butyrobetaine

15

20

25

Retention time (minutes) Fig. 1. Extracted ion chromatograms demonstrating the LCeMS separation of five pairs of structural isomers.

spectrometry (infusion at 4 mL min
1 of standard at 5 mg mL
1 in 50% methanol 0.1% formic acid or 50% acetonitrile 0.1% formic acid) to confirm that the missing compound was present within the mixture and could be ionized under the measurement conditions. An unknown peak in LCeMS chromatograms was tentatively identified as allantoic acid based on MHþ and MS2, but it was not initially possible to confirm ID because a purified standard could not be sourced commercially. ID of the unknown was subsequently confirmed by synthesizing potassium allantoate from allantoin, essentially as described previously (Young and Conway, 1942): 1.12 g of allantoin was added to 8.0 mL of 1 M KOH, the solution was heated at 75  C for 30min, cooled, 80 mL of 95% (v/v) ethanol was added and the solution was crystallised by refrigerating at 4  C overnight, the potassium allantoate was dissolved in a minimal volume of warm water, recrystallized, and finally evaporated to dryness. 2.3. Soil samples and extraction To evaluate the analytical performance with soil extracts, soil was collected from mesocosms containing soil from Themeda triandra grassland. Previous studies have described the mesocosms (Warren, 2014c), and the intact soil (an abruptic lixisol) from which they were derived (Warren, 2013b). In March 2014, a composite

Prior to injection standards and samples for CEeMS were made up in 100 mM ammonium formate 25% acetonitrile. In the case of soil extracts analysed without pre-concentration this was achieved by diluting an aliquot of sample with an equal volume of 200 mM ammonium formate (pH 10) in 50% acetonitrile. To analyse soil extracts concentrated ten-fold, 1000 mL was evaporated under reduced pressure (Vacufuge, Eppendorf) and then taken up in 100 mL of 100 mM ammonium formate in 25% acetonitrile. Capillary electrophoresisemass spectrometry (CEeMS) was used for profiling of organic N monomers in extracts, essentially as described previously (Warren, 2013a). CEeMS was performed with a capillary electrophoresis system (P/ACE MDQ, BeckmaneCoulter, Fullerton, USA) equipped with a bare fused silica capillary (50 mm i.d.  100 cm long, Polymicro Technologies, Phoenix, USA) interfaced via a co-axial sheath-flow sprayer (G1607A, Agilent, Waldbronn, Germany) to an ion trap mass spectrometer (AmaZon SL, Bruker Daltonics, Bremen, Germany). Sheath liquid of 50% (v/v) methanol with 0.1% (v/v) formic acid was delivered at 4 mL min
1 by a syringe pump (NE-1002X, New Era Pump Systems, Farmingdale, USA) driving a 10-mL PTFE-tipped gas tight syringe (SGE, Ringwood, Australia). Ion source parameters were: dry gas 4 L min
1 N2 at 200  C, nebulizer 6 psi, electrospray in positive mode at 4.5 kV. Ion transmission was optimized for a target mass of 100 m/z using smart parameter setting. The ion trap was set to scan a range of 50e255 m/z in enhanced resolution mode (8100 u/s). Ion accumulation time was adjusted automatically by setting the ion trap's ion charge control to 100,000 with a maximum accumulation time of 10 ms. Samples were injected by pressure (3 psi for 30 s) and separated with an electrolyte of 2 M formic acid with 20% (v/v) methanol under 30 kV positive polarity. Between runs the capillary was flushed with electrolyte for 10 min at 30 psi. 2.5. Analysis by liquid chromatographyemass spectrometry Aqueous extracts cannot be directly analysed by HILIC because injection of samples (or standards) in 100% water leads to a complete loss of resolution. Samples instead had to be brought to 80%

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(v/v) acetonitrile prior to injection. In the case of soil extracts analysed without pre-concentration this was achieved by diluting five-fold with 100% acetonitrile. To analyse soil extracts concentrated ten-fold, 1000 mL was evaporated under reduced pressure (Vacufuge, Eppendorf) and then taken up in 100 mL of 80% acetonitrile. The LC separation was performed using a 150 mm long  2.0 mm i.d., 3.5 mm ZIC-HILIC column (Merck SeQuant, Umeå, Sweden) operated by a HPLC system comprising a degasser (2003 degasser, Biotech, Onsala, Sweden), quaternary HPLC pump (Rheos 2000, Flux Instruments, Basel, Switzerland), and column heater (Cera Column Heater, SPEware Corporation, Baldwin Park, USA). During method development injections were made with a low-dispersion manual injector (model 8125, Rheodyne, Rohnert Park, USA) while for routine analysis an autosampler was used (model 430, Varian, Walnut Creek, USA). During method development different mobile phase modifiers were examined, namely, 5 mM ammonium acetate, 5 mM ammonium acetate with 0.1% formic acid, and 0.1% formic acid. The best overall performance for organic N monomers was achieved with mobile phases comprising water and acetonitrile modified with 0.1% formic acid. The final mobile phase composition was solvent A ¼ 0.1% (v/v) formic acid in water, solvent B ¼ 0.1% (v/v) formic acid in acetonitrile. The LC mobile phase was held at 80% B for 1 min, then a linear gradient from 80% B to 15% B over 35 min, a 10 min hold at 5%B, a 1 min ramp back to 80% B, then 18 min reequilibration at 80% B. The flow rate was 150 mL min
1, the column was maintained at 25  C. Injections volumes were generally 5 mL. The column was directly interfaced with no splitting into an ion trap mass spectrometer (AmaZon SL, Bruker Daltonics, Bremen, Germany). Ion source parameters were: dry gas 7 L min
1 N2 at 300  C, nebulizer 20 psi, electrospray alternating between positive mode at 3.0 kV and negative mode at
2.5 kV. Ion transmission was optimized for a target mass of 100 m/z using smart parameter setting. The ion trap was set to scan a range of 50e255 m/z in enhanced resolution mode (8100 u/s). Ion accumulation time was adjusted automatically by setting the ion trap's ion charge control to 100,000 with a maximum accumulation time of 10 ms. 2.6. Data analysis The aim of this study was not to identify as many compounds as possible, but instead compare the performance of the two analytical platforms for 69 standard compounds and two types of soil extract. To estimate how many organic compounds could be resolved by the two analytical platforms for the two extract types involved three steps. The first step involved integrating all peaks with a signal to noise ratio greater than 5. This was achieved by creating extracted ion chromatograms (±0.2 Da) for each nominal mass from 50 to 254 Da and then automated peak picking with a smoothing width of 2 and signal to noise ratio of 5 (DataAnalysis version 4.0, service pack 4, Bruker). In the second step peaks from exogenous and inorganic compounds were removed by manually curating the peak list and deleting peaks arising from compounds present in blanks and peaks originating from inorganic ions (e.g. salt-formate clusters). The final step of data analysis involved ensuring that each organic compound was represented by one peak only (typically [MþH]þ). This was achieved by manually inspecting the data and deleting those peaks that were in-source fragments, isotopes, adducts with alkali metal ions, or multimers. To calculate the precision of LCeMS and CEeMS analyses of soil extracts, 27 compounds in the 2.5% CHCl3 extract were identified and integrated in each of five replicate samples. Compounds in soil extracts were identified based on comparison of retention times or migration times, [MþH]þ, MS2 and (for some compounds) MS3 with authentic standards run under the same conditions on the

same instrument. For the majority of compounds quantification was based on 4-point standard curves constructed from 0.01 mg mL
1 to 1 mg mL
1. For each peak I calculated the standard deviation and relative standard deviation (sd/mean) of concentration, and retention time (LCeMS) or migration time (CEeMS). 3. Results 3.1. Separation and detection of organic N monomers by LCeMS Sixty-two out of the 69 standards were analysable by LCeMS with HILIC separation (Table S2). The seven compounds that could not be analysed were amines or polyamines. The 62 analysable compounds eluted between five and approximately 27 min, but the total analysis time was approximately one hour because of the need for lengthy re-equilibration of the column before injection of subsequent samples. Peak shapes were approximately Gaussian for most compounds, though there was some tailing for most quaternary ammonium compounds. Peak widths at half the peak height (FWHM) ranged between 0.1 and 0.3 min for a 5 mL injection of 0.01 mg mL
1 standards. Among the 69 standards there were five pairs of structural isomers for which chromatographic separation was important because they were detected at the same mass and thus could not be resolved by (single-stage) mass spectrometry. Three of the pairs of structural isomers were well resolved chromatographically (Ala from b-Ala, dimethylglycine from GABA, acetylcholine from gbutyrobetaine) (Fig. 1). Leu was not baseline resolved from Ile, but quantification and identification was not problematic because in standards and samples peak apexes were always separated by a valley. Peak apexes of valine and betaine were separated from one another in standards, but in samples valine sometimes appeared as an unresolved lump on the side of the much larger betaine peak (see Section 3.3). Nevertheless, valine and betaine could be successfully quantified by MS2 since they have contrasting fragmentation patterns (Fig. 2). Compounds were detected in positive and negative ionization mode, but subsequent discussion refers only to detection of ions formed in positive mode owing to generally better detection limits. Compounds were detected and quantified as their quasi-molecular ions [MþH]þ. For many compounds adducts with alkali metals (i.e. [MþNa]þ and [MþK]þ) were also detected, but the intensity of adduct ions was typically less than 10% of quasi-molecular ions. Limits of detection for a 5 mL injection ranged between 0.002 and 0.38 mmol L
1, with the majority (49 compounds) having limits of detection better than 0.05 mmol L
1 (Table S2). Limits of detection were poor for allantoin (0.38 mmol L
1) and uric acid (0.34 mmol L
1). 3.2. Comparison of LCeMS with CEeMS Separation by CEeMS has been described previously (Warren, 2013a), so only brief data are given here. Of the 69 standards, 62 were analysable by CEeMS. The weakly cationic compound ergothioneine migrated slowly and produced a broad peak with poor detection limits; five neutral compounds (urea, thymine, uracil, allantoin, N-acetyl-glucosamine) were detectable but not analysable because they yielded very broad unresolved peaks with very poor detection limits; while the anionic compound uric acid was not detected at all. The majority of compounds had migration times between approximately 10 and 25 min, with another 10 min required for neutrals to migrate from the capillary. The total analysis time for one sample, including a 10-min washing of the capillary, was 50 min. Peak shapes were approximately Gaussian for all compounds. For most compounds peaks in CEeMS were

C.R. Warren / Soil Biology & Biochemistry 78 (2014) 233e242

237 118

a) 10.8 min 98

72

intensity

55

60

b) 10.8 min

101

110

MS2 of 118

72

118

c) 11.5 min

d) 11.5 min

50

MS

MS2 of 118

58 59

60

MS

70

80

90

100

110

120

m/z Fig. 2. Mass spectra of valine (a and b) and betaine (c and d). A mixture of valine and betaine was injected, partially separated by LC (see Fig. 1c), then autoMSn was used to isolate and fragment the quasi-molecular ions ([MþH]þ ¼ 118 Da) thereby yielding MS2 spectra (c and d).

approximately five times narrower than in LCeMS, with FWHM of most CEeMS peaks being 0.02e0.05 min. All of the five pairs of structural isomers were baseline resolved by CEeMS (data not shown). Limits of detection for a 3 psi  30 s injection (~80 nL calculated based on Poiseuille's law) were on average similar to those measured by LCeMS, although for individual compounds there was no correlation between detection limits measured by LCeMS and CEeMS (Fig. 3). In CEeMS limits of detection ranged between 0.004 and 0.25 mmol L
1, with the majority (53 compounds) having limits of detection better than 0.05 mmol L
1 (Table S2). Limits of detection were notably poor for cadaverine (0.25 mmol L
1) and ergothioneine (0.13 mmol L
1).

3.3. Separation and detection of monomeric organic N compounds in soil extracts The first tests of performance with soil extracts involved examining if it was possible to analyse soil extracts with the bare minimum of sample preparation, i.e. without pre-concentration or other lengthy preparative procedures. Unfortunately some sample preparation was necessary because optimal performance of LCeMS and CEeMS required that samples were prepared in appropriate injection solvents/buffers prior to injection. In the case of LCeMS soil extracts were brought to 80% acetonitrile by diluting 5-fold with 100% acetonitrile, while in the case of CEeMS soil extracts were brought to 100 mM ammonium formate 25% acetonitrile by diluting 1:1 with 200 mM ammonium formate 50% acetonitrile. When water extracts of soil were analysed without any preconcentration, there were 24 compounds that were identified by

both LCeMS and CEeMS while an additional 4 compounds were identified by LCeMS only (Table 1, Figs. 4a and 5a). The 24 compounds in common comprised 16 protein amino acids, 5 quaternary ammonium compounds, ectoine and hydroxyectoine. The four compounds detected by LCeMS only were the ureides urea, allantoin, uric acid and allantoic acid. In 2.5% CHCl3 extracts of soil concentrations of organic N monomers were greater than in water extracts and an additional three compounds were detected by both LCeMS and CEeMS, while a fourth additional compound (N-acetylglucosamine) was detected by LCeMS only (Table 1). When samples were pre-concentrated 10-fold prior to injection 5e10 times as many peaks were detected (Table 1, Figs. 4bc, and 5bc). The better separation performance of CEeMS meant that in 10-fold concentrated extracts CEeMS resolved an additional 50e60 compound peaks than LCeMS. When analysing 10-fold concentrated samples by LCeMS it became apparent that inorganic salts present in the sample were separated on the column and eluted among the organic compounds. Inorganic salts and other components of the sample matrix can adversely affect analysis by inhibiting the ionization and detection of organic compounds. This phenomenon, known as ion suppression (Annesley, 2003), can be a substantial problem for quantification by altering the relationship between concentration and peak area such that a standard curve determined with purified standards is inapplicable for samples. To assess if ionization and detection of ions was affected by the complex matrix of soil extracts, a syringe pump and “T” union was used to infuse a solution of 2 mg mL
1 13C3 15N alanine at 1 mL min
1 into the mobile phase downstream from the column. When blanks or standards were injected the signal from 13C3 15N alanine decreased gradually from 6 to 17 min owing to the composition of the mobile phase changing

Table 1 Number of endogenous organic compounds detected in soil extracts analysed by LCeMS and CEeMS. Samples were analysed after dilution in injection buffer/solvent (5-fold dilution in the case of LCeMS, 2-fold dilution in the case of CEeMS), or analysed after 10-fold concentration. Compounds were integrated and identified only if they occurred in at least 2 out of 4 replicates and had a signal to noise ratio great than 5. Data were curated by hand to remove peaks related to compounds present in blanks and inorganic ions. Moreover, to ensure each organic compound was represented by one peak only (typically [MþH]þ), I deleted peaks that were insource fragments, isotopes, adducts with alkali metal ions, or multimers. H2O extract 2.5% CHCl3 2.5% CHCl3 extract H2O extract (diluted) extract (diluted) 10  concentrated 10  concentrated Fig. 3. Limits of detection (at a signal-to-noise ratio of three) for purified standards analysed by LCeMS and CEMS. LODs were determined for 5 mL LCeMS injections and CEeMS injections of 3 psi  30 s (calculated as 80 nL based on Pouseuille's law).

LCeMS 28 (5-fold) CEeMS 24 (2-fold)

32 (5-fold) 27 (2-fold)

75 135

159 208

238

C.R. Warren / Soil Biology & Biochemistry 78 (2014) 233e242

Fig. 4. Overlayed extracted ion chromatograms of LCeMS separations of a) a H2O extract of soil diluted 5-fold prior to injection, b) a H2O extract concentrated 10-fold prior to injection, and c) a 2.5% CHCl3 extract concentrated 10-fold prior to injection. In a) and c) most of the readily-identifiable peaks have been annotated: Amino acids are given standard 3-letter abbreviations, g-butbet ¼ g-butyrobetaine, Ac-carn ¼ acetyl-carnitine, ergo ¼ ergothioneine. Note that the y-axis scale is 10 times greater for b) and c) than a).

over time and affecting ionization efficiency (Fig. 6a). When water or CHCl3 extracts were analysed without pre-concentration there was the same gradual decline in signal intensity from 13C3 15N alanine but superimposed on it were three modest negative responses (dips) around 7, 10 and 17 min (Fig. 6b). With ten-fold concentrated extracts there was an almost complete loss of the 13 C3 15N alanine signal around 7, 10 and 17 min (Fig. 6c). These zones of ion suppression coincided with retention times of inorganic salts.

3.4. Quantification and analytical precision Facile quantification was not possible in ten-fold concentrated extracts due to the strong ion suppression effect (Fig. 6c). Hence, quantitative performance and reproducibility was assessed with a 2.5% CHCl3 extract injected without any pre-concentration since in such extracts ion suppression was modest (Fig. 6b). The 27 compounds quantified by LCeMS and CEeMS spanned an almost 200-

Fig. 5. Overlayed extracted ion electropherograms of CEeMS separations of a) a H2O extract of soil diluted 2-fold prior to injection, b) a H2O extract concentrated 10-fold prior to injection, and c) a 2.5% CHCl3 extract concentrated 10-fold prior to injection. In a) and c) most of the readily-identifiable peaks have been annotated. Abbreviations are as in Fig. 4. To enable easy comparison with the same samples analysed by LCeMS the scale of the y-axes are the same as in Fig. 4.

C.R. Warren / Soil Biology & Biochemistry 78 (2014) 233e242

239

a) blank

3 2

1

Intensity (x106)

0 b) CHCl3 extract diluted 5-fold

3 2 1 0

Cl

2

c) water extract concentrated 10-fold

PO4, SO4

3

Na K

1 0

5

10

15

20

25

Retention time (minutes) Fig. 6. The effects of injecting a) a blank, b) a 2.5% CHCl3 extract diluted 5-fold, or c) an H2O extract concentrated 10-fold on ionization and detection of a standard (13C3 15N Ala) infused at a constant rate in-between the LC column and mass spectrometer ion source. The pink line is the signal from the constantly infused 13C3 15N Ala, while the green line is a basepeak chromatogram (the intensity of the most intense peak at every point in the analysis).

fold concentration range from 0.05 to 8.1 mmol L
1 (Fig. 7). Concentrations of individual compounds measured by LCeMS and CEeMS were strongly positively correlated. The total concentration of the 27 compounds quantified with both analytical platforms was 26.3 ± 0.3 mmol L
1 (mean ± sd, n ¼ 5) by LCeMS and 28.3 ± 0.5 mmol L
1 by CEeMS. The four ureides (urea, allantoin, uric acid and allantoic acid) that were quantifiable by LCeMS had a total concentration of 9.4 ± 0.4 mmol L
1. For the 27 compounds quantified by LCeMS and CEeMS the reproducibility of concentration was, on average, similar at 6.8%RSD for LCeMS and 6.9% RSD for CEeMS. The reproducibility of retention time was, on average, approximately six times better for

Fig. 7. Comparison of LCeMS and CEeMS measured concentration of 27 organic N compounds in a 2.5% CHCl3 extract of soil. LCeMS samples were diluted 5-fold prior to injection, CEeMS samples were diluted 2-fold prior to injection. Concentrations reported here are those in the original extract (i.e. after accounting for dilution by injection solvent/buffer). The same sample was analysed five times by each of LCeMS and CEeMS. Error bars are one standard deviation.

LCeMS (mean retention time reproducibility ¼ 0.25%RSD) than CEeMS (mean migration time reproducibility ¼ 1.6%RSD).

4. Discussion 4.1. Analytical performance of LCeMS A variety of targeted methods are already in widespread use for analysis of compound classes such as amino acids in soil extracts (e.g. Kielland, 1995; Yu et al., 2002; Warren, 2008; Farrell et al., 2011). These methods work admirably and the intention here was not to supplant these methods, but instead develop a LCeMS method that could yield information about known and unknown compounds from multiple compound classes within one analytical run. In the present study the HILIC separation mechanism was examined because it has been used successfully by numerous groups for analysis of complex mixtures of hydrophilic compounds (Lu et al., 2008; Kato et al., 2009; Schiesel et al., 2010; Creek et al., 2011; Iwasaki et al., 2011; Rappold and Grant, 2011; Boudra et al., 2012; Buszewski and Noga, 2012; Chen et al., 2012; Fraser et al., 2012; Zhang et al., 2012). As with other studies we found that HILIC separation with a generic formic acid based mobile phase provided broad coverage of multiple classes of organic N, namely, amino acids, quaternary ammonium compounds, ureides, and nucleobases. There was massive overlap in the compounds that could be analysed by LCeMS and CEeMS, but neither analytical platform was on its own truly comprehensive for organic N compounds. Most compounds could be detected by both analytical platforms, but there were some that could be detected by LCeMS only and some by CEeMS only. LCeMS was able to analyse neutral and anionic compounds (e.g. N-acetyl-glucosamine and ureides) that could not be detected by CEeMS of cationic compounds (Warren, 2013a). However, LCeMS was unable to analyse polyamines e a compound class that could be analysed by CEeMS and can be abundant in some soil extracts (Fujihara and Harada, 1989;

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Fujihara, 2009; Warren, 2014b). Additional testing showed polyamines could not be effectively eluted from the HILIC column even with stronger eluents (i.e. lower % organic and higher salt concentration), and thus it is plausible that polyamines were irreversibly retained by electrostatic interactions with the column's zwitterionic stationary phase. Hence, while there is considerable overlap between LCeMS and CEeMS the two analytical platforms are complementary and use of both platforms would yield the broadest coverage of organic N compounds. The other aspect to knowing which compounds are analysable is determining if detection limits are sufficiently low for the specific samples to be analysed. Detection limits are especially important for soil due to the modest concentrations of organic N compounds in soil extracts (Warren, 2014a). As it happens, neither analytical platform had an overall advantage in terms of detection limits with the overwhelming majority of purified standards having limits of detection better than 0.05 mmol L
1 when analysed by LCeMS or CEeMS. This is an important outcome because one of the criticisms commonly levelled at CEeMS is that detection limits are too poor for use with Real-world samples (Shen and Smith, 2002). However, this is clearly not the case with CE methods such as the one here that incorporate on-line sample stacking procedures (Simpson et al., 2008) having detection limits comparable to LCeMS. Independent of whether LC or CE separation is used, the limits of detection reported here could be improved upon by modifying the means of mass spectrometry detection to a targeted MS/MS approach. For example, use of a triple quadrupole mass spectrometer in multiple reaction monitoring mode would achieve better detection limits than reported here (Soga et al., 2004; Büscher et al., 2009). The overall separation efficiency was poorer for LCeMS than CEeMS. LCeMS peaks were, on average, about five times as wide as peaks in CEeMS, and thus all else being equal CEeMS ought to be able to resolve five times as many peaks as LCeMS. This is supported by data for ten-fold concentrated soil extracts showing CEeMS resolved more compound peaks than LCeMS (Table 1). This was not surprising because separation efficiency in CE is known to be up to an order of magnitude greater than conventional LC (Kuehnbaum and Britz-McKibbin, 2013). One way of working around the poorer separation performance of LCeMS is to rely upon the selectivity provided by MS/MS and/or high resolution mass spectrometry. For example, use of MS/MS would enable the mass spectrometer to resolve compounds that have the same (nominal or exact) mass but different fragmentation patterns (e.g. valine and betaine for which [MþH]þ ¼ 118.0862 Da, Fig. 2), while high resolution enables resolution of peaks that have the same nominal mass but different exact mass (e.g. guanidinoacetic acid with [MþH]þ ¼ 118.0611 Da from valine with [MþH]þ ¼ 118.0862 Da). Many of the recent studies using LCeMS with HILIC separation to examine complex samples of biological origin have used high resolution mass spectrometry and/or MS/MS (Gika et al., 2012; Zhang et al., 2012), and this approach is to be recommended for future studies to at least partially offset the poorer separation performance of LCeMS. One area in which LCeMS has a distinct advantage over CEeMS is in the reproducibility of retention time. With LCeMS, peaks had very consistent retention times and thus automated peak detection, ID and integration were feasible. With CEeMS the order of peaks and their migration times relative to an internal standard (relative migration time) did not vary between injections (data not shown), but there was significant variation between injections in the absolute migration time of peaks. The variation in absolute migration time is problematic because absolute times are used for peak detection, ID and integration in the software supplied with mass spectrometry instrumentation. Variation in absolute migration

times can be addressed via calculation of relative migration time (Kuehnbaum and Britz-McKibbin, 2013) or use of peak alignment software (Ramautar et al., 2009; Sugimoto et al., 2010), but both approaches take time and require use of third-party software. Hence, for analysis of large numbers of samples LCeMS will offer distinct advantages due to the ease with which one can automate data analysis (i.e. peak detection, identification and integration) without having to perform additional calculations and use multiple software suites. 4.2. Dealing with inorganic salts and ion suppression The comparatively poor salt tolerance of LCeMS is problematic because it makes it difficult to analyse samples with an unfavourably high ratio of inorganic to organic compounds and precludes analysis of strong salt extracts (e.g. 0.5 M K2SO4). Inorganic salts are a problem because they co-elute with (target) organic N compounds and suppress their ionization and/or detection (Fig. 6c) (socalled “ion suppression”, Annesley, 2003; Antignac et al., 2005; Remane et al., 2010). Ion suppression and other matrix effects may cause limits of detection to be poorer in samples than was found for standards (Fig. 3 and Table S2) and can massively complicate quantification. This complication arises because ion suppression can affect the relationship between analyte concentration and peak area (i.e. the standard curve established with standards is not applicable to samples). It is possible to work around the problem by using calibration strategies such as matrix matching, standard addition and isotope-labelled internal standards (Sojo et al., 2003; Rychlik and Asam, 2008; Hewavitharana, 2011; Hewavitharana et al., 2014); however, such strategies typically require a large investment in time and money. Desalting samples with solid-phase extraction (SPE) was tested but rejected as a means of avoiding ion suppression. The problem is that of the SPE stationary phases tested (strong cation exchange, porous graphitic carbon, and C18) none yielded consistently high recovery of all organic N compounds (data not shown). For example, strong cation exchange led to good recovery of amino acids, poor recovery of quaternary ammonium compounds and irreversibly retained polyamines. These findings are not unprecedented with a previous study reporting similarly poor recovery of small organic compounds from soil extracts de-salted with various SPE stationary phases (Oikawa et al., 2011). A solution could be offered by more sophisticated desalting strategies involving combinations of different SPE stationary phases, but this would add an extra layer of complexity to sample preparation. In any case, the challenge for all desalting strategies is that inorganic salts and many small organic N compounds share similar physico-chemical properties, and thus it is difficult to desalt samples without simultaneously affecting relative and absolute concentrations organic N compounds. Consequently, to achieve accurate quantification of organic N compounds in desalted samples would require the liberal use of isotope-labelled internal standards to check and correct for recovery. The solution used here was to visualize and quantify the occurrence of ion suppression using a post-column infusion (Fig. 6), and then perform quantification only with dilute soil extracts for which ion suppression was negligible. The strong agreement between concentrations estimated by LCeMS and the almost completely independent technique of CEeMS (Fig. 7) strongly supports the validity of quantitative LCeMS estimates made with dilute soil extracts. One obvious drawback of being restricted to working with dilute extracts is that only the most abundant organic N compounds will be quantified. The second drawback is that to quantify and visualize ion suppression before attempting quantification requires the investment of additional time and

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instrumentation (namely a syringe pump, and various pieces of HPLC tubing and connectors). However, the investment in time and instrumentation is modest and absolutely necessary given the widespread occurrence (Annesley, 2003; Antignac et al., 2005; Remane et al., 2010) and deleterious effects of ion suppression (e.g. Fig. 6c). The most elegant way to avoid the problems posed by inorganic salts is to use a separation such as capillary electrophoresis that ensures salts do not enter the mass spectrometer at the same time as organic N compounds. One of the largest advantages of CEeMS for analysis of cationic compounds is that the electrophoretic separation functions as a form of on-line de-salting (Lee et al., 2007; Kuehnbaum and Britz-McKibbin, 2013). After injecting a sample the inorganic anions back-migrate into the buffer vial upon application of a voltage, while inorganic cations migrate in the same direction as organic cations but at a faster rate. Hence, inorganic salts are effectively removed from the sample before the organic N compounds migrate from the CE column into the mass spectrometer. As a consequence samples with an unfavourably high ratio of inorganic salts to organic N and even strong salt extracts (e.g. 0.5 M K2SO4) are analysable by CEeMS (Warren, 2014b, 2014c) and CEeMS of cationic compounds suffers very little ion suppression or other matrix effects (Büscher et al., 2009). 4.3. LCeMS analysis of soil extracts The overall profile of small organic N in soil extracts was similar to previous reports of the same soil analysed by CEeMS (Warren, 2014b) and differed little between LCeMS and CEeMS, with the notable exception of ureides that were detected by LCeMS only. In a 2.5% CHCl3 extract the pool of four ureides comprised approximately 25% of the pool of LCeMS detected organic N monomers, while individual ureides were the 2nd (uric acid), 4th (allantoic acid), 7th (allantoin), and 10th (urea) most abundant compounds. Few of the contemporary studies on organic N in soil have reported the presence of ureides, yet the occurrence and metabolism of ureides in soil may be important for soil function and plant nutrition. Ureides are produced during catabolism of basic amino acids and purines (Wang et al., 2008; Witte, 2011), and occur in plants (Wang et al., 2008), mycorrhizae (Larsen et al., 2011) and other microorganisms (Vogels and Vanderdrift, 1976). Moreover, at least some ureides can serve as an N source for plants (Molliard, 1910; Brigham, 1917; Wang et al., 2008), mycorrhizae (Sharples and Cairney, 1997) and other microorganisms (Vogels and Vanderdrift, 1976). The ability of LCeMS to measure ureides alongside precursors and products of ureide metabolism may help uncover the role of this compound class in soil function and plant nutrition. 4.4. Conclusions The LCeMS procedure developed here provided good coverage of small organic N compounds and demonstrated three crucial advantages over CEeMS. The first advantage was being able to analyse neutral and anionic compounds such as N-acetyl-glucosamine and ureides that are not amenable to analysis by CEeMS of cationic compounds. The second advantage is that the reproducibility of retention times of LCeMS peaks was 6 times better than the reproducibility of migration times of CEeMS peaks, and thus analysis of LCeMS data was more straightforward and easier to automate than for CEeMS. The final advantage of LCeMS is that the installed base of LCeMS instruments is orders of magnitude larger than for CEeMS, and thus instrumentation and expertise are readily accessible. There were, however, areas in which LCeMS performed comparatively poorly. In LCeMS quantitative analysis was possible only with quite dilute extracts in which the majority of

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organic N compounds were below detection limits. The overall separation efficiency in LCeMS was approximately five times poorer than CEeMS, and thus many more peaks could be resolved in a CEeMS electropherogram than in a LCeMS chromatogram. Finally, LCeMS was unable to analyse strongly cationic compounds such as polyamines. This study has highlighted that the comprehensive analysis of low-molecular weight organic N compounds in soil is challenging and cannot be achieved with any one analytical platform. Neither LCeMS nor CEeMS offered demonstrably better performance than each other, and a combination of the two would provide the broadest possible coverage of organic N in soil. Acknowledgements Charles Warren is supported by a Future Fellowship from the Australian Research Council. The University of Sydney is thanked for financial support via the major equipment scheme. Appendix A. Supplementary data Supplementary data related to this article can be found at http:// dx.doi.org/10.1016/j.soilbio.2014.08.008. References Alpert, A.J., et al., 1994. Hydrophilic-interaction chromatography of complex carbohydrates. Journal of Chromatography A 676, 191e202. Annesley, T.M., 2003. Ion suppression in mass spectrometry. Clinical Chemistry 49, 1041e1044. Antignac, J.P., de Wasch, K., Monteau, F., De Brabander, H., Andre, F., Le Bizec, B., 2005. The ion suppression phenomenon in liquid chromatographyemass spectrometry and its consequences in the field of residue. Analytica Chimica Acta 529, 129e136. Boudra, H., Doreau, M., Noziere, P., Pujos-Guillot, E., Morgavi, D.P., 2012. Simultaneous analysis of the main markers of nitrogen status in dairy cow's urine using hydrophilic interaction chromatography and tandem mass spectrometry detection. Journal of Chromatography A 1256, 169e176. Brigham, R.O., 1917. Assimilation of organic nitrogen by Zea mays and the influence of Bacillus subtilis on such assimilation. Soil Science 3, 155e200. Büscher, J.M., Czernik, D., Ewald, J.C., Sauer, U., Zamboni, N., 2009. Cross-platform comparison of methods for quantitative metabolomics of primary metabolism. Analytical Chemistry 81, 2135e2143. Buszewski, B., Noga, S., 2012. Hydrophilic interaction liquid chromatography (HILIC) e a powerful separation technique. Analytical and Bioanalytical Chemistry 402, 231e247. Chapin, F.S., Moilanen, L., Kielland, K., 1993. Preferential use of organic nitrogen for growth by a nonmycorrhizal arctic sedge. Nature 361, 150e153. Chen, J., et al., 2012. Urinary hydrophilic and hydrophobic metabolic profiling based on liquid chromatography-mass spectrometry methods: differential metabolite discovery specific to ovarian cancer. Electrophoresis 33, 3361e3369. Creek, D.J., Jankevics, A., Breitling, R., Watson, D.G., Barrett, M.P., Burgess, K.E.V., 2011. Toward global metabolomics analysis with hydrophilic interaction liquid chromatography-mass spectrometry: improved metabolite identification by retention time prediction. Analytical Chemistry 83, 8703e8710. Farrell, M., Hill, P.W., Farrar, J., Bardgett, R.D., Jones, D.L., 2011. Seasonal variation in soluble soil carbon and nitrogen across a grassland productivity gradient. Soil Biology & Biochemistry 43, 835e844. Fraser, K., et al., 2012. Non-targeted analysis of tea by hydrophilic interaction liquid chromatography and high resolution mass spectrometry. Food Chemistry 134, 1616e1623. Fujihara, S., 2009. Biogenic amines in rhizobia and legume root nodules. Microbes and Environments 24, 1e13. Fujihara, S., Harada, Y., 1989. A novel polyamine, aminobutylhomospermidine, in Japanese volcanic ash soils. Soil Biology & Biochemistry 21, 449e452. Gika, H.G., Theodoridis, G.A., Vrhovsek, U., Mattivi, F., 2012. Quantitative profiling of polar primary metabolites using hydrophilic interaction ultrahigh performance liquid chromatographyetandem mass spectrometry. Journal of Chromatography A 1259, 121e127. Hakkinen, M.R., et al., 2007. Analysis of underivatized polyamines by reversed phase liquid chromatography with electrospray tandem mass spectrometry. Journal of Pharmaceutical and Biomedical Analysis 45, 625e634. Hewavitharana, A.K., 2011. Matrix matching in liquid chromatography-mass spectrometry with stable isotope labelled internal standards e is it necessary? Journal of Chromatography A 1218, 359e361. Hewavitharana, A.K., Tan, S.K., Shaw, P.N., 2014. Strategies for the detection and elimination of matrix effects in quantitative LCeMS analysis. LC GC North America 32, 54e64.

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