Localized Metal Solubilization in the Rhizosphere of Salix smithiana upon Sulfur Application

A metal-accumulating willow was grown under greenhouse conditions on a Zn/Cd-polluted soil to investigate the effects of sulfur (S0) application on metal solubility and plant uptake. Soil porewater samples were analyzed 8 times during 61 days of growth, while DGT-measured metal flux and O2 were chemically mapped at selected times. Sulfur oxidation resulted in soil acidification and related mobilization of Mn, Zn, and Cd, more pronounced in the rooted compared to bulk soil. Chemical imaging revealed increased DGT-measured Zn and Cd flux at the root-soil interface. Our findings indicated sustained microbial S0 oxidation and associated metal mobilization close to root surfaces. The localized depletion of O2 along single roots upon S0 addition indicated the contribution of reductive Mn (oxy)hydoxide dissolution with Mn eventually becoming a terminal electron acceptor after depletion of O2 and NO3–. The S0 treatments increased the foliar metal concentrations (mg kg–1 dwt) up to 10-fold for Mn, (5810 ± 593), 3.3-fold for Zn (3850 ± 87.0), and 1.7-fold for Cd (36.9 ± 3.35), but had no significant influence on biomass production. Lower metal solubilization in the bulk soils should translate into reduced leaching, offering opportunities for using S0 as environmentally favorable amendment for phytoextraction of metal-polluted soils.


Soil characteristics and plant metal concentrations S2
Rhizotron experiment and chemical mapping; Table SI-2; Table SI

Rhizobox experiment and soil porewater sampling
Prior to filling the treatments into the rhizobox compartments, they were moistened with lab water type 1 (0.055 µS cm −1 , TKA-GenPure, Thermo Electron LED GmbH, Niederelbert, Germany) using a spray bottle. The soils were filled into the compartments and carefully compacted using a plastic tamper to a soil bulk density of approximately 1.4 g cm -3 . A total of ~600 g soil was used per rhizobox. During filling, one pre-grown willow cutting was transplanted per rhizobox after washing off the potting mixture from the roots using lab water type 1. In the center of the rooted and bulk soil compartment, 50-mm long Rhizon samplers (Rhizosphere research products, Wageningen, Netherlands) were installed for collecting soil porewater during the experiment, ~20 mm above rhizobox ground. Soil porewater from the membrane compartment was not sampled. The S. smithiana cuttings were cut back to one single, vital twig to reduce leaf transpiration and ensure uniform growth conditions at the start of the experiment. For the maintenance of stable water content in soil, two PE-coated glass fiber wicks (TRIPP Kristallo Rundschnur, 4 mm, IDT, Frankfurt, Germany) were installed in each bulk S3 soil and the rooted compartment and connected to a lab water type 1 bottle. At the start of the experiment, the rhizoboxes were saturated gravimetrically to 80% water holding capacity (WHC). The rhizoboxes were wrapped in aluminum foil for preventing algal growth inside the transparent rhizobox compartments. The experimental plants were grown in the rhizoboxes for 61 days in a greenhouse at 60% rel. humidity (day/night cycle: 16/8 h). The rhizoboxes were randomized several times during the experimental period.

Soil porewater sampling
The soil porewater in the rooted and bulk soil compartments was sampled eight times during the growth period

Harvest and sample preparation
Twigs and leaves were washed with lab water type 1 in a stainless steel sieve (<2 mm) for three minutes, and dried at 65 °C for 72 h. For root harvesting, the rooted compartments were spread on plastic foils and air dried overnight. Large roots were manually collected using plastic tweezers. Smaller roots were separated from the soil by gentle sieving using a stainless steel sieve (<2 mm). Adsorbed metal cations from the root apoplasm were removed by washing the roots two times for ten minutes in 250 mL 0.05 mol L -1 CaCl2 (EMSURE® ACS, Merck, Vienna, Austria) in an ultrasonic bath (Sonorex RK510, Bandelin electronic GmbH & Co. KG, Berlin, Germany). 2 The washed roots were put on tissue paper and dried at 65 °C for 72 h. All plant samples were finely milled using a stainless steel mill (IKA A11, IKA®-Werke GmbH & CO. KG, Staufen, Germany).

DGT gel preparation and deployment
For localized metal cation sampling, 2 × 4 cm gel strips were used. After roots of ~10 cm length had developed (4 DAP), front plates were carefully opened, the PTFE foils removed and growth containers were placed horizontally on a lab stand while photographs were taken of the region of interest (ROI). In a laminar flow bench (class II, EN 12469:2000), DGT gel strips were placed on the rhizotron front plates containing a thin water film. As diffusive layer, and to avoid particle contamination of the DGT, 10 µm thick polycarbonate filter membrane strips (Nuclepore, Whatman, Maidstone, UK) were placed on top of each DGT gel. The assembly was then moistened using lab water type 1 to remove air bubbles and fixed to the front plate using water proof adhesive tape (Scotch Super 88, 3M TM ). The DGT-equipped front plates were carefully re-mounted to the rhizotrons and fixed using plastic clamps. The sampling time was 20 h, derived from a preliminary bulk DGT experiment, aiming at avoiding saturation of the SPR-IDA resin with metal cations.

S5
After detaching the front plates, the gels were retrieved and rinsed with lab water type 1. Membranes were carefully removed from the plates using plastic tweezers, DGTs were rinsed and subsequently transferred onto a clean stack of membrane (Supor 450, Pall Corporation, New York, USA) and gel blotting paper (GB005, Whatmann, Maidstone, UK), soil-exposed gel-side facing up which was covered with polyethylene (PE) foil.
For dehydration, the gel assemblies were placed into a gel dryer (Unigeldryer 3545, UNIEQUIP, Planegg, Germany) for 48 h at 55 °C. After drying, gels were retrieved, and carefully fixed onto glass plates using double-sided adhesive tape. Calibration standard and blank gel samples for LA-ICPMS were prepared as described in Warnken et al. 4 . The samples were then analyzed using laser ablation inductively coupled plasma mass spectrometry (LA-ICPMS).
Preparation of DGT-LA-ICPMS standards and blank gels

LA-ICPMS analysis
The laser ablation analysis of the samples and standards was carried out in line scanning mode according to For calibrating the O2 optodes, a modified Stern-Volmer equation was used (Eq. 2). 10

= 1
Eq. (2) Here, R is the (red-green/green) luminescent intensity ratio, R0 is the ratio in the absence of oxygen, C the O2 concentration, Ksv the Stern-Volmer quenching constant and α is the nonquenchable fraction of the luminescence. The calibration parameters were determined in in a gas-tight, transparent perspex container S8 which was purged several times with N2. Subsequently, O2 was mixed into the gas stream stepwise using a customized gas mixing device. The O2 concentration C was continuously measured by an oxymeter (GMH 3691 Oxymeter, GGO 370 sensor, Greisinger Electronic, Germany, SD ± 0.2 %) at ambient temperature and pressure. We measured ten sequences, while the O2 concentration was increased each step by ~1% O2(abs.). After ten minutes of equilibration, images were taken and average values were calculated for an optode area of ~8 cm 2 (1.94x10 8 pixel, RSD<10%). From these data we determined the fitting parameters, (-0.160) and ! "# (0.102 µmol L -1 ) and calibrated the optained images from the rhizotrons in ImageJ (http://imagej.nih.gov/ij/) after solving Eq. (2) for C. Figure SI-5 shows an example calibration.