Ion-Selective Electrodes

Urine samples from five different individuals were collected in 1 day between 10 am and 3 pm, and placed on ice. The collected samples were pooled, and then frozen at −80 °C in aliquots of approximately 150 ml. Prior to the NMR experiments, urine was passed through a column of Chelex-100 ion exchange resin (BioRad) to remove the majority of Ca²⁺ and Mg²⁺ ions, and the pH indicator standards imidazole, formate, Tris and piperazine were added to a final concentration of 1 mM, 1 mM, 0.25 mM and 0.25 mM, respectively. We measured the main metal ion concentrations in urine (Ca²⁺, Mg²⁺, Na⁺ and K⁺) with ion selective electrodes and pH measurements were performed with a glass electrode with inbuilt temperature sensor (Fisher Scientific).
For the pH titration experiment, two volumes of Chelex-treated urine (200 ml) were treated dropwise with either 1 M HCl or 1 M NaOH while stirring. Samples were taken for NMR (400 μl) at 0.2 pH unit intervals and were prepared for NMR with addition of H₂O (180 μl) and ²H₂O (20 μl) containing 4,4-dimethyl-4-silapentane-1-sulfonic acid-²H₆ (DSS) to give a final concentration of 0.1 mM. For the ion titration experiment, Chelex-treated urine (400 μl), was supplemented with H₂O containing various concentrations of CaCl₂, MgCl₂, NaCl or KCl, ranging from 0 to 1 M, and D₂O (20 μl) containing DSS to give a final concentration of 0.1 mM. NMR samples were centrifuged for 5 min at 13,000 rpm and 550 μl was transferred to a 5 mm NMR tube. Spectra were acquired on a Bruker Avance DRX600 NMR spectrometer (Bruker BioSpin, Rheinstetten, Germany), with ¹H frequency of 600 MHz, and a 5 mm inverse probe at a constant temperature of 300 K. Samples were introduced with an automatic sampler and spectra were acquired following the procedure described by Beckonert et al. (2007 (link)). Briefly, a one-dimensional NOESY sequence was used for water suppression; data were acquired into 64 K data points over a spectral width of 12 kHz, with 8 dummy scans and 128 scans per sample. Spectra were processed in iNMR 3.4 (Nucleomatica, Molfetta, Italy). Fourier transform of the free-induction decay was applied with a line broadening of 0.5 Hz. Spectra were manually phased and automated first order baseline correction was applied. Metabolites were assigned at Metabolomics Standards Initiative (MSI) level 2 using the Chenomx NMR Suite 5.1 (Chenomx, Inc., Edmonton, Alberta, Canada). Metabolite peak positions from the different samples relative to DSS were obtained using MATLAB scripts written in-house by Dr Gregory Tredwell, and appropriate chemical shifts were determined for multiplets. A version of the scripts for peak picking and spline fits are part of the BATMAN project (batman.r-forge-project.org) (Liebeke et al. 2013 (link)). The observed chemical shifts of the various metabolite peaks were modelled with respect to pH with the general formula (Eq. 2) for multibasic acids (H_nL), using the nlinfit function within MATLAB. The number of sites was assumed from the chemical structure. This enabled the estimation of pK_a values and acid and base chemical shift limits for individual metabolite peaks.

These involved the administration of a questionnaire concerning family and personal history, habitual physical activity and dietary habits, a standard physical examination and an anthropometric evaluation (height, weight, body-mass index). BMI was calculated for each subject and BMI z-score was assessed, according to Centers for Disease Control and prevention (CDC) growth charts (http://www.cdc.gov/nchs/data/series/sr_11/sr11_246.pdf) [14 ].
At the end of the visit, the participants (or their caregivers in the case of younger children) received a plastic container for 24h urine samples together with detailed oral and written instructions on how to collect complete 24h urines, as previously described [13 ]. Once the collection was returned, the subject was required to confirm completeness of the collection, the total urine volume was recorded and two samples were extracted, immediately stored in plastic containers and frozen at -30°C to be later analysed by the central laboratory at Federico II University of Naples, as previously reported [13 ]. 24 h urinary excretions were used as proxies for the respective dietary intakes, according to WHO recommendation [15 ]. Urinary sodium and potassium concentrations were measured by ion selective electrode potentiometry and urinary creatinine by a kinetic Jaffe′ reaction using an ABX Pentra 400 apparatus (HORIBA ABX, Rome, Italy). Quality control was performed using urine specific reference samples from UrichemGol BIO-DEV (Milan, Italy). The inter-assay technical error was 0.73% for sodium, 1.16% for potassium and 1.12% for creatinine.

Annual fluxes of C and N were estimated from the measured monthly fluxes. Biomass stocks were estimated annually, while soil stocks were estimated only at the end of the investigation.
Soil C fluxes were characterized by determining the total soil CO₂ efflux and heterotrophic respiration, whereas soil N fluxes were characterized by determining the net mineralization and nitrification rates using an in situ incubation method. Total soil CO₂ efflux was separated into root- and heterotrophic respiration using a trench method^{59 (link)}. A root exclusion barrier (50 cm diameter, 30 cm depth, PVC) was installed in each plot, 4 weeks before the treatments were initiated. Two soil collars (10 cm diameter, 5 cm depth, PVC) were installed inside the rooting barrier to measure heterotrophic respiration and another two were installed outside the rooting barrier to measure heterotrophic + root respiration. Vegetation was removed within the rooting barriers but litterfall was kept. Soil CO₂ efflux was measured once a month using an infrared gas analyzer (Model EGM-4, PP-Systems, Hitchin, Hertfordshire, UK) equipped with a flow-through closed chamber (Model SRC-2). Measurements were performed between 10:00 and 13:00 over the study period. Soil temperature was also measured at a depth of 8 cm near the soil CO₂ efflux collar using a digital soil temperature probe (K-type, Summit SDT 200, Seoul, Korea).
On each day when the soil CO₂ efflux was measured, two soil cores were collected from each plot at a depth of 5 cm using a 100 cm³ core soil sampler. One sample was placed in a plastic bag and the other was returned to the soil and incubated to estimate the net N mineralization rate. We thus collected two soil samples from each plot every month, one fresh and one incubated. Both samples were transported to the laboratory and their fresh weight was measured. A 10 g portion of the fresh soil sample was oven-dried for 48 h at 105 °C to quantify the soil’s gravimetric water content and the rest was kept to determine the concentrations of nitrate and ammonium and the soil pH (measured using a 1:5 soil water suspension with an ion-selective glass electrode; Istec Model pH-220L, Seoul, Korea). The bulk density of each soil sample was calculated from its gravimetric water content and fresh weight.
Ammonium and nitrate were extracted from soil samples (5 g) using 50 ml of a 2 M KCl solution in a mechanical vacuum extractor (Model 24VE, SampleTek, Science Hill, KY, USA). The resulting solutions were then immediately placed in a cooler at 4 °C for storage. The ammonium and nitrate concentrations of the solutions were determined using an auto analyzer (AQ2 Discrete Analyzer, Southampton, UK). The mineral N concentration was measured throughout the period when fertilizer was applied, from April 2011 to April 2014. The net rates of ammonification and nitrification were estimated based on the differences in the ammonium and nitrate contents of the soil, respectively, between before and after the incubation. The sum of these two rates was taken as the net mineralization rate. If the net mineralization rate was negative, it was regarded as a net immobilization rate.
Additional soil samples were collected from four randomly selected points in each plot at depths of 0–15 cm both 4 weeks before starting the treatments and at the end of the investigation (2014). Samples were pooled within a plot and their concentrations of C, N, and available P were determined to capture the soil’s initial condition and the treatment response. C and N were analyzed with an elemental analyzer (vario Macro cube, Elementar Analysensysteme GmbH, Germany). The available P concentration was determined by extraction using NH₄F and HCl solutions⁶⁰ and analyzed using a UV spectrophotometer (Jenway 6505, Staffordshire, UK). C and N stocks were estimated by measuring the concentrations of both elements in a 100 cm³ core and dividing by the bulk density of the soil in the 0–5 cm layer.