Floodplain soils polluted with high levels of mercury (Hg) are potential
point sources to downstream ecosystems. Repeated flooding (e.g., redox
cycling) and agricultural activities (e.g., organic matter addition) may
influence the fate and speciation of Hg in these soil systems. The formation
and aggregation of colloids and particles influence both Hg mobility and
its bioavailability to microbes that form methylmercury (MeHg). In this study,
we conducted a microcosm flooding–draining experiment on Hg-polluted
floodplain soils originating from an agriculturally used area situated in
the Rhone Valley (Valais, Switzerland). The experiment comprised two 14 d
flooding periods separated by one 14 d draining period. The effect of
freshly added natural organic matter on Hg dynamics was assessed by adding
liquid cow manure (+MNR) to two soils characterized by different Hg (47.3±0.5 or 2.38±0.01 mg kg-1) and organic
carbon (OC: 1.92 wt % or 3.45 wt %) contents. During the experiment,
the release, colloid formation of Hg in soil solution and net MeHg
production in the soil were monitored. Upon manure addition in the highly
polluted soil (lower OC), an accelerated release of Hg to the soil solution
could be linked to a fast reductive dissolution of Mn oxides. The manure
treatments showed a fast sequestration of Hg and a higher percentage of Hg bound by
particulate (0.02–10 µm). Also, analyses of soil
solutions by asymmetrical flow field-flow fractionation coupled with
inductively coupled plasma mass spectrometry (AF4–ICP–MS) revealed a
relative increase in colloidal Hg bound to dissolved organic matter (Hg–DOM)
and inorganic colloidal Hg (70 %–100 %) upon manure addition. Our
experiment shows a net MeHg production the first flooding and draining
period and a subsequent decrease in absolute MeHg concentrations after the
second flooding period. Manure addition did not change net MeHg production
significantly in the incubated soils. The results of this study suggest that
manure addition may promote Hg sequestration by Hg complexation on large
organic matter components and the formation and aggregation of inorganic
HgS(s) colloids in Hg-polluted Fluvisols with low levels of natural
organic matter.
Introduction
Mercury (Hg) is a pollutant of global concern due to its high toxicity and
to its global biogeochemical cycle which spans all environmental
compartments (atmosphere, oceans, soils, etc.) (Beckers and Rinklebe, 2017;
AMAP/UN Environment, 2019). Sediments and soils are major Hg pools with
relatively long residence times (Amos et al., 2013; Driscoll et al., 2013).
Legacy Hg from industrial sites (e.g., chlor-alkali plants or mining areas)
retained in soils is a key source for present-day atmospheric Hg (Amos et
al., 2013). However, this retained Hg pool can also be remobilized by
landscape alteration, land use (e.g., fertilization, manure addition) or
climate-induced changes such as drought–flood–drought cycles of soils
(Singer et al., 2016). These inputs are a threat to downstream ecosystems
and human health due to release of inorganic Hg and the formation and
bioaccumulation of toxic monomethylmercury (MeHg) in both aquatic and
terrestrial food chains (Bigham et al., 2017).
Mercury is redox sensitive and occurs mainly as elemental Hg0,
inorganic Hg2+ or in the form of MeHg in soils. In general, Hg
speciation in soils depends on the biogeochemical conditions. For example,
in boreal peatlands and forest soils rich in natural organic matter (NOM), Hg
is primarily bound to thiol groups of NOM (NOM–Hg), associated with
FeS(s) or found as cinnabar (HgS(s)) or metacinnabar
(β-HgS(s)). These species are the thermodynamically most favored
forms of Hg in these environments (Skyllberg et al., 2006; Skyllberg and
Drott, 2010; Biester et al., 2002). However, Hg sorbed on the surfaces of
manganese (Mn), iron (Fe) and aluminum (Al) oxy-hydroxides may also
represent important Hg pools in soils with low amounts of NOM (Guedron et
al., 2009).
The fate of Hg in soils is still not well characterized, and its
mobilization and sequestration in soil depend on a variety of factors and
mechanisms. The release of Hg to the soil solution and its further transport
have been associated with the mobilization of NOM (Kronberg et al., 2016;
Eklöf et al., 2018; Åkerblom et al., 2008), copper (Cu)
nanoparticles (Hofacker et al., 2013) or the reductive dissolution of
Fe / Mn oxyhydroxides (Frohne et al., 2012; Gygax et al., 2019; Poulin et al.,
2016). Earlier studies reported a relatively rapid decrease in dissolved Hg
after its release upon flooding in various riparian settings (Hofacker et
al., 2013; Poulin et al., 2016; Gygax et al., 2019). Possible pathways for
this decrease are Hg2+ reduction to Hg0, sorption to recalcitrant
NOM, formation of metacinnabar β-HgS(s) or co-precipitation of Hg
in sulfides (e.g., FeS(s)) or metallic particles.
Metallic colloids in soil may be formed by biomineralization during soil
reduction or precipitation in the root zone and potentially incorporate
toxic trace elements like Hg (Weber et al., 2009; Manceau et al., 2008).
These colloids may increase the mobility and persistence of toxic trace
metals in soil solution if they do not aggregate to bigger particles. During
a flooding incubation experiment, Hofacker et al. (2013) observed the
incorporation of Hg in Cu nanoparticles, which were shown to be formed by
fermentative bacteria species (Hofacker et al., 2015). Colloidal
β-HgS(s) has been reported to form abiotically in soils under oxic
conditions directly by interaction with thiol groups of NOM (Manceau et al.,
2015). In solution, dissolved organic matter (DOM) has a major influence in
the formation and aggregation of metallic colloids and particles. It may
promote the dissolution of HgS(s) phases, decelerate the
aggregation and growth of HgS(s) colloids, and affect the
crystallinity of HgS(s) phases (Miller et al., 2007; Ravichandran et
al., 1998; Gerbig et al., 2011; Poulin et al., 2017; Pham et al., 2014).
The same effects were also observed for other metal sulfide, oxide or
carbonate colloids (Aiken et al., 2011; Deonarine et al., 2011). In the case of
Hg, inhibition of β-HgS(s) formation may in turn increase its
mobility and bioavailability to MeHg-producing microorganisms (Deonarine and
Hsu-Kim, 2009; Ravichandran et al., 1999; Aiken et al., 2011; Graham et al.,
2012). Chelation of Hg with higher-molecular-weight NOM may also inhibit
the microbial availability of Hg (Bravo et al., 2017). Within Hg–NOM,
hydrophobic, thiol-rich NOM with higher molecular weight contains a higher
density of strong sorption sites (thiol groups) (Haitzer et al., 2002).
However, different ligand exchange reactions (e.g., carboxyl groups to thiol
groups) kinetically control this sorption and thus the bioavailability of
dissolved Hg in aqueous systems (Miller et al., 2007, 2009;
Liang et al., 2019). The partly contradicting statements above illustrate
the complex role of NOM and DOM in the Hg cycle and Hg bioavailability and
the need for more research in this field.
The formation of MeHg from inorganic Hg2+ has been shown to be
primarily microbially driven. Environments of redox oscillation (e.g.,
floodplains, estuaries) represent hotspots for Hg methylation
(Marvin-DiPasquale et al., 2014; Bigham et al., 2017). Mercury methylators
are usually anaerobe microbial species such as sulfate reducers (SRB), Fe
reducers (FeRB), archaea and some firmicutes (Gilmour et al., 2013).
Generally, Hg is bioavailable to methylators in the form of dissolved
Hg2+, Hg complexed by labile DOM or Hg-bearing inorganic nanoparticles
(e.g., FeS(s), HgS(s)) but is less available when complexed by
particulate organic matter (Hg–POM) or larger inorganic particles
(Chiasson-Gould et al., 2014; Graham et al., 2013; Rivera et al., 2019;
Zhang et al., 2012; Jonsson et al., 2012). Further, DOM is a main driver of
Hg methylation as it influences both bioavailability and microbial activity.
The role of DOM as an electron donor may enhance the microbial activity and
thus the cellular uptake. The composition and origin of DOM were reported to
change Hg methylation rates (Drott et al., 2007; Bravo et al., 2017). For
example, Bravo et al. (2017) showed that in lake sediments, terrestrially
derived DOM led to slower methylation rates than phytoplankton-derived DOM.
The addition of DOM in the form of organic amendments (e.g., manure, rice straw,
biochar) has been reported to have both an enhancing effect (Gygax et al., 2019;
Liu et al., 2016; Wang et al., 2019; Eckley et al., 2021; Wang et al., 2020)
and no effect (Zhu et al., 2016; Liu et al., 2016) on the net MeHg production
in soils. Further, organic amendments were reported to shift microbial
communities. Both the enhancement of Hg demethylators and Hg reducers (Hu et
al., 2019) and the enhancement of Hg methylators upon organic amendments
were reported (Tang et al., 2019; Wang et al., 2020). Environments of
elevated Hg methylation (riparian zone, estuary) are also places of elevated
NOM degradation and mineralization due to temporal changes in redox
conditions. The degradation of large NOM to more bioavailable low-molecular-weight (LMW) compounds promoted by microbial Mn oxidation, especially in
systems with neutral pH (Jones et al., 2018; Sunda and Kieber, 1994; Ma et
al., 2020), is also hypothesized to increase bioavailability of Hg–NOM.
However, amendments of Mn oxides were also shown to inhibit Fe- and
SO42--reducing conditions and thus MeHg formation in sediments
(Vlassopoulos et al., 2018).
Hg methylation and mobilization is intensively studied in paddy field soils
and peat soils due to their relevance in food production or the Hg global
cycle (Wang et al., 2019; Tang et al., 2018; Liu et al., 2016; Hu et al.,
2019; Wang et al., 2016; Zhao et al., 2018; Zhu et al., 2016; Kronberg et
al., 2016; Skyllberg, 2008; Skyllberg et al., 2006). However, only a few
studies focused on Hg methylation and mobility in temperate floodplain soils
(Frohne et al., 2012; Hofacker et al., 2013; Gilli et al., 2018; Poulin et
al., 2016; Lazareva et al., 2019; Wang et al., 2020; Beckers et al., 2019).
Also, few studies have examined the effect of flooding and/or land use
(NOM addition in the form of animal manure) in polluted soils with respect
to Hg release and methylation potential (Tang et al., 2018; Gygax et al.,
2019; Zhang et al., 2018; Hofacker et al., 2013; Frohne et al., 2012).
Furthermore, most of these studies were focusing on soils with rather high
OC levels (5 wt %–10 wt %), and only few researchers have addressed the
decrease in Hg in soil solution of flooded soils over time, including the
fate of colloidal Hg.
This work focused on the effect of agricultural practices on the Hg
mobility and methylation in a real-world contaminated Fluvisol with specific
emphasis on the flooding–draining cycle and manure addition. By conducting
microcosm experiments, we studied the effect of these cycles and manure
addition on (1) the release and sequestration of Hg, (2) the methylation of
Hg and (3) the evolution of colloidal and particulate Hg in soil solution.
The latter was studied by analyzing different soil solution filter fractions
(0.02 and 10 µm) as well as analyzing selected samples by asymmetric
flow field-flow fractionation coupled to a UVvis detector, a
fluorescence detector and an ICP–MS (AF4–ICP–MS). Based on the presented
state of knowledge, we hypothesize that the manure addition would accelerate
the release of Hg by accelerated reductive dissolution of Mn oxyhydroxides
in these soils and eventually change Hg speciation in the system towards
Hg–NOM complexes and β-HgS(s) colloids.
Methods and materialsSample collection
We sampled soil from agriculturally used fields in the alpine Rhone Valley
in Wallis, Switzerland, on 30 September 2019. The fields are situated
in a former floodplain next to the artificial “Grossgrundkanal” canal.
This canal was built in the 1900s to drain the floodplain and as a buffer
for the wastewater releases of a chemical plant upstream that historically
used Hg in different processes (chlor-alkali electrolysis, acetaldehyde
and vinyl chloride production). The soils on the floodplain were subjected
to Hg pollution from this plant between the 1930s and the 1970s, mostly
through the removal and dispersion of the canal sediments onto the
agricultural fields (Glenz and Escher, 2011). After heavy rain events, the
fields are subjected to draining–flooding cycles (Fig. S1 in the Supplement) and have been
identified as potential hotspots for Hg methylation and release (Gygax et
al., 2019). For this study, soil was sampled from a cornfield and a pasture
field next to the canal. A map and the coordinates of the sampling locations
are provided in the Supplement (Fig. S1, Table S1 in the Supplement). At each site, a composite
sample of approximately 10 kg of soil was sampled between 0–20 cm depth from
10 points on the fields. The soil samples were named after their relative
pollution and organic carbon levels (high mercury, low carbon (HMLC) and low
mercury, high carbon (LMHC); see Sect. 2.4 below for details on the soils).
After sampling, roots were removed, and the fresh soil was sieved to
<2 mm grain size, further homogenized, split in two parts and
stored on ice in airtight PE bags for transport to the laboratory.
Additionally, approximately 2 L of liquid cow manure was sampled from a close-by
cattle farm. One aliquot of the samples was stored at -20∘C
until further processing. The remaining part was used for the incubation
experiment within 12 h after sampling. A detailed description of the site
and sampling procedures is given in the Supplement (Sect. S1).
Microcosm experiments
An initial incubation was conducted in 10 L HDPE containers in the dark for
7 d in an atmosphere of 22 ∘C and 60 % relative
humidity (RH) in order to equilibrate the soils and to prevent a peak of
microbial respiration induced by the soil sieving before the onset of the
experiment (Fig. 1). After the initial incubation period soils were used in
the flooding and draining experiments, which were conducted in 1 L
borosilicate glass aspirator bottles (Fig. S2). The environment created
through soil flooding in these bottles will be called the microcosm (MC) in the
following text. Microcosm experiments were performed in experimental
triplicate and named after the relative Hg and organic carbon levels of the
used soil (HMLC and LMHC) and the treatment with or without manure addition
(added +MNR). The microcosms were equipped with an acid-washed suction cup
with a pore size of <10µm (model: 4313.7/ETH, ecoTech
Umwelt-Meßsysteme GmbH, Bonn, Germany). In the following sequence, 700 g
of artificial rainwater (NH4NO3 11.6 mg L-1/K2SO4
7.85 mg L-1/Na2SO4 1.11 mg L-1/MgSO4⚫7H2O 1.31 mg L-1/CaCl2 4.32 mg L-1) was added to the
microcosms. For the manure treatment, 0.6 % (w/w) (3 g) of liquid cow
manure was added to the microcosms corresponding to one application of
liquid manure on a cornfield following the principles of fertilization of
agricultural crops in Switzerland (Richner and Sinaj, 2017), and finally
fresh soil was added with a soildry : water ratio of 1 : 1.4 (w/w) (Fig. S3). Then, the microcosms were gently shaken for at least 1 min to
remove any remaining air bubbles in the soil and pore space. An additional
mixture of fresh soil artificial rainwater (1 : 1.4 (w/w)) was shaken for 6 h
to assess the equilibration of the solid and liquid phases during the
experiment. The microcosms were covered with Parafilm®,
transferred to the incubation chamber (APT.line™ KBWF, Binder,
Tuttlingen, Germany) and incubated in the dark for 14 d in an atmosphere of
22 ∘C and 60 % RH. The incubation temperature was chosen to be
close to the daily mean soil temperature at 10 cm depth during summer months
between 2015–2019 (21.4 ∘C) at the closest soil temperature
monitoring station (Sion, VS, provided by MeteoSwiss) situated downstream.
After the first flooding period, the supernatant water was pipetted off, and
remaining water was sampled through the suction cups to drain the
microcosms. They were weighted before and after water removal. Then,
approximately 25 g of moist soil was sampled by two to three scoops though
the whole soil column using a disposable lab spoon. The microcosms were
kept drained in an atmosphere of 22 ∘C and 10 % RH for 14 d. For the second flooding period, the microcosms were again flooded with
500 g of artificial rainwater and incubated for another 14 d in an
atmosphere of 22 ∘C and 60 % RH (Fig. 1). After the
incubation, the suction cups were removed, and the soils were homogenized and
then transferred from the MC to a PE bag and stored at -20∘C
until further processing.
Schedule of preformed incubation experiment, samplings and
measurements: blue bars indicate soil flooding periods. Gray bars represent
drained periods. The width of the columns is not proportional to the time of
incubation. In the treatments row the (+) symbol indicates the addition of
liquid manure to the microcosms specifically treated with manure (+MNR).
Triangles represent regular soil solution sampling points. Rectangles
represent soil solution sampling for colloid analyses. Diamonds represent
time points for soil sampling. At -7 d, soil was sampled from the pooled
soil directly before the pre-incubation.
Soil and manure characterization
Frozen soil and manure samples were freeze dried to avoid a loss of Hg prior
to analyses (Hojdová et al., 2015), ground using an automatic ball mill
(MM400, Retsch, Haan, Germany) and analyzed for the following chemical
parameters. Carbon (C), nitrogen (N) and sulfur (S) were measured with an
elemental analyzer (vario EL cube, Elementar Analysensysteme, Langenselbold,
Germany). Organic carbon (OC) was calculated by subtracting the C
concentration of a loss-on-ignition sample (550 ∘C for 2 h) from
the original C concentration. The pH was measured in an equilibrated 0.01 M CaCl2 solution (1 : 5 soil : liquid ratio). Mineral composition was
measured by X-ray diffraction (XRD, CubiX3, Malvern Panalytical,
Malvern, United Kingdom). Trace and major metals (e.g., Fe, Mn, Cu) and total Hg (HgT)
were extracted from soils using a 15.8 M nitric acid microwave digestion and
measured using an inductively coupled plasma – mass spectrometer (ICP–MS,
7700x, Agilent Technologies, Santa Clara, United States of America).
Methylmercury was selectively extracted with HCl and dichloromethane (DCM)
using an adapted method described elsewhere (Gygax et al., 2019). We
modified this method to achieve high throughput (64 samples per run) and
measurements by high-pressure liquid chromatography (HPLC, 1200 Series,
Agilent Technologies, Santa Clara, United States of America) coupled to the
ICP–MS. Details on laboratory materials, extractions, analytical methods
and instrumentation are provided in the Supplement (Sects. S2, S3). The
change in MeHg concentration in the microcosms was likely a result of the
simultaneous production and degradation of MeHg. Thus, the term “net MeHg
production” was used to represent these processes. We calculated the
relative net MeHg production during the incubation as the relative
difference of MeHg / HgT ratios between two time points (t) using Eq. (1).
Net MeHg production(%)=MeHgHgTti-1-MeHgHgTtiMeHgHgTti-1×100
Soil description
Both soils were identified as Gleyic Fluvisols. They have a silt loam texture and the same
mineral composition but differing Hg and organic carbon (Corg)
concentrations (Table 1). For elements relevant for Hg cycling, Hg molar
ratios (HgT : Cu, HgT : Corg, HgT : Mn) differ between samples and soils used in
similar incubation experiments (Hofacker et al., 2013; Poulin et al., 2016).
We note that the [Corg/ Mn]molar was 30 % higher in the LMHC
soil compared to HMLC. X-ray diffractograms of both soils is shown in Fig. S4. The soil diffractograms are overlapping each other, and the qualitative
analyses of the diffractograms show that the soils' parental material is
composed of the same five main mineral phases: quartz, albite, orthoclase,
illite/muscovite and calcite.
List of soil parameters for the two incubated soils (HMLC and LMHC)
and manure (MNR). Uncertainties are given as 1σ standard deviation
of triplicate experiments (method triplicates).
Parameter Cornfield (HMLC) Pasture field (LMHC) Cow manure (MNR) Land useCornfieldPasture–Depth0–20 cm0–20 cm–Soil type (WRB)Gleyic FluvisolGleyic Fluvisol–pH8.167.84–Water content(wt %)13.88.590.3Unit (dry.wt.)ConcentrationSDConcentrationSDConcentrationSDCorgwt %1.920.013.450.0145.220.09Ntotwt %0.1810.0010.3720.0023.680.08Corg/Ntot–10.61–9.29–––Sg kg-10.630.050.770.053.70.1HgTmg kg-147.30.52.40.30.0450.001MeHgµg kg-126.90.26.40.2<0.02–MeHg / HgT%0.06–0.28–––Alwt %0.910.051.050.040.01060.0003Fe1.950.072.380.050.03360.0009Mg1.250.071.390.050.490.03Mnmg kg-14932167238531P1169801044858245232Cr5646450.680.01Co10.750.0611.220.430.40.2Ni81.70.878.32.92.30.1Cu40.11.228.00.713.10.6Zn61.80.547.32.0813As11.740.0716.040.720.80.4Cd0.210.040.170.010.0420.004Pb20.80.518.340.5––V17.20.420.991.10.310.01Sr1372202645.91.6Cs1.990.021.520.04––Ba60.21.176.91.69.10.5Ce7.00.48.60.6––Gd0.940.031.000.050.0210.001U1.740.081.290.010.190.01HgT / Cu molar‰366.3–25.73–––HgT / Mn molar25.758–0.926–––HgT / Corg molar0.147–0.004–––Mn / Corg molar0.0056–0.0042–––Soil solution sampling and analyses
Soil solution was sampled 0.25, 1, 2, 3, 4, 5, 7, 9, 11 and 14 d after the
onset of each flooding period (Figs. 1, S5). It was sampled
though the tubing connected to the suction cup (<10µm pre
size). The first 2 mL was sampled with a syringe and discarded to prime the
system and condition the tubing. Then 4 mL was drawn through an airtight
flow-through system to measure the redox potential (Hg / HgCl ORP electrode)
and pH. Then, approximately 35 mL of soil solution was sampled using a
self-made syringe pump system allowing for a regular flow and minimal
remobilization of fine particles. Like this, 4 %–6 % of the added
artificial rainwater volume was sampled at each sampling point (Fig. S3).
Throughout the experiment the soils remained entirely submerged. At each
sampling time, sample splits were preserved without further filtration
(< 10 µm) and filtered at 0.02 µm
(Whatman® Anodisc 0.02 µm, Sigma-Aldrich, St. Louis,
United States of America). Additionally, at 2, 5 and 9 d an additional
sample split was filtered at 0.45 µm (Polytetrafluoroethylene
Hydrophilic, BGB, Böckten, Switzerland) for colloid characterization.
Incubation experiment blanks were taken by sampling Milli-Q water through
from an empty 1 L borosilicate aspirator bottle three times throughout the
experiment. Subsequently, the samples were subdivided and treated for
different analyses. They were preserved in 1 % HNO3 for multi-elemental analysis (Mn, Fe, Cu, As) and in 1 % HNO3 and 0.5 % HCl
for HgT analysis and analyzed by ICP–MS. For major anion (Cl-,
NO3-, SO42-) and cation (K+, Na+, Mg2+,
Ca2+) measurements, samples were diluted 1 : 4 in ultra-pure water and
analyzed by ion chromatography (Dionex Aquion™, Thermo Fisher
Scientific Inc., Waltham, United States of America). Samples for dissolved
organic carbon (DOC), particulate organic carbon (POC) and total bound nitrogen
(TNb) were diluted 1 : 5 and stabilized using 10 µL of 10 % HCl and measured using an elemental analyzer (vario TOC cube, Elementar
Analysensysteme, Langenselbold, Germany). Incubation experiment blanks were
below 4.75 mg L-1 and 22.4 µg L-1 for DOC and TNb,
respectively. These relatively high blank values might originate from either
the syringes or the suction cups (Siemens and Kaupenjohann, 2003).
Uncertainties of soil solution parameters are displayed as 1 SD of the
triplicate incubation experiments throughout the paper. HCO3-
concentrations were estimated based on the ionic charge balance of the soil
solution using VisMinteq (https://vminteq.lwr.kth.se/, last access: 1 October 2020). A detailed schedule
and list of analyses is provided in Fig. 1. Concentrations of specific
filtered fractions are labeled with subscripts (e.g., HgT<0.02µm)
for all measured metals. Particulate concentrations (0.02 µm <X< 10 µm) (e.g., P-Fe) and their proportion to the
total (e.g., P-Mnrel) were determined as the difference between
unfiltered and filtered concentration (Table 2).
Description of the symbols and terms used for different filter
fractions in the publication. The particulate fraction is calculated as the
difference of the 20 nm and the 10 µm filtrate concentrations.
Filter typeFilter sizeSymbol (e.g., HgTx)DescriptionSuction cup10 µmHgT<10µmSoil solution sampled directly from the suction cup contains a variety of particles (clay minerals, bacteria, Mn/Fe hydroxides, POM aggregates, etc.). We refer to this fraction by adding the subscripts <10µm to the analyte symbol.Syringe filter0.02 µmHgT<0.02µmSoil solution <0.02µm is a cutoff size that may still carry colloids. We refer to this fraction by adding the subscripts <0.02µm to the analyte symbol.––P-HgTParticulate Hg is calculated as P-HgT = HgT<10µm- HgT<0.02µm––P-HgTrel.Relative particulate HgT is calculated asP-HgTrel.= (HgT<10µm- HgT<0.02µm)/HgT<10µmAF4 membrane1 kDaHgT<1kDaMolecules in solution under this cutoff size are not expected to have colloidal properties. Therefore, this range is referred to as “truly dissolved” in the text.Characterization of colloids (AF4)
An aliquot of the soil solution was used for characterization of colloids in
one out of three replicate microcosms (Rep1) of each treatment on days 2, 5 and
9 after the onset of each flooding period. Right after
sampling, the aliquots were transferred to a N2 atmosphere in a glove
box. There, the samples were filtered to < 0.45 µm and
preserved in airtight borosilicate headspace vials at 4 ∘C.
Colloidal size fractions and elemental concentrations of the filtrates were
analyzed by asymmetrical flow field-flow fractionation (AF4, AF2000,
Postnova Analytics, Landsberg am Lech, Germany) coupled to a UV254nm
absorbance detector (UV, SPD-M20A, Shimadzu, Reinach, Switzerland), a
fluorescence detector (FLD, RF-20A, Shimadzu, Reinach, Switzerland) and an
ICP–MS (7700x, Agilent Technologies, Santa Clara, United States of America)
within 14 d after sampling. Colloids contained in 1 mL of samples were
separated in a channel made of a trapezoidal spacer of 350 µm
thickness and a regenerated cellulose membrane with a nominal cutoff of 1 kDa used as the accumulation wall. The mobile phase used for AF4 elution was 10 mM NH4NO3 at pH 7 and was degassed prior to entering the channel by
argon flowing. A linear decrease in crossflow from 2 to 0 mL min-1 over
20 min was used after injecting the samples at an initial crossflow of 2.7 mL min-1. At the end of a run, the crossflow was kept at 0 mL min-1 for 5 min in order to elute non-fractionated particles. Retention
times were transformed into hydrodynamic diameters (dh) by an external
calibration using Hemocyanin Type VIII from Limulus polyphemus hemolymph
(monomer dh=7 nm, Sigma-Aldrich) and ultra-uniform gold
nanoparticles (Nanocomposix) of known dh (19 and 39 nm).
Additionally, the elution of the smallest retention times (dh<10 nm) was converted into molecular masses (m) using polystyrene sulfonate standards (PSS) standards
(Postnova analytic, Landsberg am Lech, Germany) with m ranging from 1.1 to
64 kDa (Fig. S6), using AF4-UVD254nm.
Fractograms obtained in counts per second (CPS) from time-resolved analysis
(TRA) acquisition were converted to µg L-1 using external
calibrations made from a multi-element standard solution (ICP multi-element
standard solution VI, Merk, Darmstadt, Germany) diluted in 1 % HNO3
or a Hg standard (ICP inorganic Hg standard solution,
TraceCERT®, Sigma-Aldrich, St. Louis, United States of
America) diluted in 1.0 % HNO3 and 0.5 % HCl. The
different size fractions were obtained by multiple extreme-shaped peak
fitting, using OriginPro 2018 software (OriginLab Corporation). The peaks
obtained were then integrated individually, after conversion of elution time
to elution volume, to provide the quantity of Hg in each size fraction
(Dublet et al., 2019). The analytes passing the 1 kDa membrane are
considered to be the (< 1 kDa) truly dissolved fraction. It was
calculated by subtracting the concentrations of colloidal HgT recovered by
AF4–ICP–MS (total integration of the Hg signals) to the total dissolved
HgT concentrations measured separately by ICP–MS in corresponding acidified
samples. The concentration of truly dissolved Hg is displayed as
HgT<1kDa for the rest of the article (Table 1). AF4–ICP–MS,
UV254nm and fluorescence signals were used to further characterize Hg-bearing colloids, after hydrodynamic size separation by AF4. The
UV254nm light absorption is widely used to detect organic compounds, but
it should be noted that part of the UV254nm light signal can also
originate from Fe(II) or Fe hydroxides (Dublet et al., 2019). This was not
the case in this study since UV254nm signals co-eluted with C signals
recorded by ICP–MS and matched the fractograms obtained by the FLD detector
tuned at the wavelengths specific for humic-like fluorophores. It is
therefore assumed that the UV254nm signal represents organic compounds
throughout the paper.
ResultsSoil solution chemistry and Hg dynamics
In the HMLC microcosms, the pH of the soil solutions remained in a neutral-to-alkaline range of 8.0 to 8.4 during the incubation experiment (Fig. S7).
Soil solution conditions and concentrations of constituents support a
continuous reduction of soils with increased flooding time (Fig. 2a). Soil
solution NO3- depletion was observed during the first 7 d of
incubation (Fig. 2b). Nitrate was under the detection limit for the second
flooding phase. At day 7, Mn concentrations increased together with a
marginal increase in Fe (Fig. 2c–f). This was coincided with a decrease in
the relative particulate fraction (P-Mnrel. and P-Ferel.) of these
metals. Release of Mn and Fe was assumed to mark the onset of reductive
dissolution of Mn and Fe oxyhydroxides. The decrease in sulfate
(SO42-) concentration could not be used to assess the onset of
sulfate reduction. This is due to a chemical gradient between supernatant
water and soil solution demonstrated by the continuous decrease in
concentration of conservative ions (Cl-, Na+, K+) (Sect. 4.4). To monitor sulfate reduction, we use the molar ratios of
SO42- to Cl- (Fig. 2g). Sulfate-to-chloride ratios stayed
constant during the first flooding and slightly increased at the onset of the
second flooding phase. This suggests that no sulfate reduction took place
in the HMLC microcosms during the whole experiment. The DOC concentration
ranged between 37.5 and 106 mg L-1 (Fig. 2h). Both HgT<0.02µm
and HgT<10µm concentrations remained low between days 0–5 (Phase 0),
then increased together with the Mn release between days 5–11 (Phase 1) and
decreased between days 14–29 (Phase 2) during the draining period (Fig. 3a). The
relative fraction of particulate HgT (P-HgTrel.) gradually decreased
from a maximum of 88 % to a minimum of 25 % during phase 0 and phase 1, but increased again to 60 %–77 % during phase 2 (Fig. 3b–c).
Cu<0.02µm concentrations increased up to 88.2 ± 17.5 µg L-1 within the first 4 d and then gradually decreased to 30.6 ± 3.54 µg L-1 at day 14 (Fig. 4a). Arsenic concentrations
simultaneously increased with the release of Fe during the whole incubation
(Fig. 4b).
Soil solution dynamics in cornfield soil (HMLC) incubations for
redox potential (a), redox reactive elements (NO3-, Mn, P-Mn, Fe, P-Fe,
[SO42-] : [Cl-]) (b–g) and dissolved organic carbon (h). Lines
between points were plotted to improve readability. The gray area indicates
the drained period.
Soil solution dynamics in cornfield soil (HMLC) incubations for Hg (a–c) subdivided into phases (0–3). Lines between points were plotted to
improve readability. The gray area indicates the drained period. Red arrows
indicate sampling days for AF4–ICP–MS analyses.
Soil solution dynamics in cornfield soil (HMLC) incubations for Cu (a) and As (b). Lines between points were plotted to improve readability.
The gray area indicates the drained period.
During the second flooding period, individual microcosms behaved differently
in the HMLC run. The differences of soil solution Eh and redox-sensitive metals (e.g., Mn, Fe, Hg, Cu) were apparent from the start of the
second flooding (Figs. 2c–f, 3a–c, 4a). Contrastingly, DOC concentrations
and pH remained similar between incubators (Figs. 2h, S7). One replicate
(Rep1) showed a pronounced increase in redox potential after the draining
period (Fig. 2a). The Eh remained high (150 to 300 mV) for the whole
second flooding period. A depletion and subsequent release of Mn in soil
solution was observed, indicating the formation and redissolution of Mn
oxyhydroxide minerals (Fig. 2c–d). Subsequently, Mn<0.02µm increased
and peaked at 448 µg L-1 by the end of the experiment in Rep1.
The Eh of Rep2 was lower (between 28 and 120 mV), Mn concentrations did
not decrease during the draining phase and a release of Fe was observed
during the second flooding phase, indicating the reduction of Fe
oxyhydroxides. Rep3 had a Eh in the range of Rep2 but neither a
rerelease of Mn nor a release of Fe was observed during the second flooding
phase. Also, HgT behaved differently within incubators during the second
flooding period. Between days 29–42 (Phase 3), HgT<0.02µm and
HgT<10µm concentrations increased or remained at higher
levels for Rep1 and Rep3. During this phase P-HgTrel vastly decreased and was at a minimum of 1 %–7 % by the end of the incubation.
Contrastingly, HgT<0.02µm and HgT<10µm stayed constantly
low for Rep2 during phase 3, and P-HgTrel remained overall above 50 %.
The Rep1 was the only MC that showed an increase in Cu concentrations during
the draining phase (Fig. 4a).
In the HMLC + MNR microcosms, pH remained in the range of 8 to 8.35 with
minor fluctuations over both flooding periods (Fig. S7). The redox potential
decreased rapidly from approximately Eh 300 mV to 5.27 ± 14.4 mV within the first 14 d and remained constant at 14.3 ± 8.12 mV
during the second flooding period. Depletion of NO3- was observed
within the first day of incubation and was under the detection limit during the
second flooding period (Fig. 2b). A rapid release of Mn started at day 2, and
a slow release of Fe started at day 3 of the first flooding period (Fig. 2c–f).
The [SO42-] : [Cl-] ratios decreased from 0.57 ± 0.01 to
0.37 ± 0.02 between days 4–29. During the second flooding period
[SO42-] : [Cl-] ratios initially increased slightly between days
29–31 and then decreased to a minimum (0.12 ± 0.05) by the end of the
incubation (Fig. 2g). DOC concentrations were between 72.2 and 134 mg L-1 (Fig. 2h). This was significantly higher (3 to 43 mg L-1) than
in HMLC without manure. In these microcosms HgT<0.02µm and
HgT<10µm concentrations instantly increased together with the Mn
release between days 0–4 (phase 1), decreased during days 5–14 (phase 2)
and remained low between days 14–42 (phase 3) (Fig. 3a–c). The particulate
HgT (P-HgTrel.) decreased to 30 %–52.5 % in phase 1 and remained
overall above 50 % for the rest of the incubation. At the onset of phase 2 black precipitates were visually observed in the HMLC + MNR microcosms
(Fig. S13). Cu concentrations decreased gradually during the course of the
incubation experiment (Fig. 4a). Arsenic concentrations simultaneously
increased with the release of Fe during the whole incubation (Fig. 4b).
LMHC differed from HMLC in soil solution chemistry. In both treatments (LMHC
and LMHC + MNR), pH remained neutral but gradually decreased from 8.2 to
7.5 during the incubation (Fig. S7). Soil reduction progressed rapidly from
a max of 332 mV at day 3 to -14.3 mV at day 14 (Fig. 5a). During the second
flooding Eh stayed in the range of -2.3 to 34.5 mV. Nitrate was
exhausted within the first day of incubation and marked the onset of Mn
release. Mn as well as DOC concentrations gradually increased during the
first flooding period (Fig. 5b–c). Fe release started on day 4 and day 6 in
LMHC and LMHC + MNR (Fig. 5d). A decrease in
[SO42-] : [Cl-] ratio was observed after day 5 and remained
stable at 0.03±0.04 during the second flooding period. This is
indicative of sulfate reduction during the draining phase and the second
flooding phase (Fig. 5e). Soil solution HgT<0.02µm concentration (25–160 ng L-1) was 2 orders of magnitude lower than in the HMLC runs
(Figs. 2, 3a, 6a). Dissolved HgT<0.02µm degreased during the first
flooding period (phase 1), increased during the draining period (phase 2)
and gradually decreased again during the second flooding period (phase 3)
(Fig. 6a–c). No other soil solution parameter followed the trend of
HgT<0.02µm. Particulate HgT<10µm decreased during phase 1
and remained low during phases 2 and 3. In the LMHC microcosms P-HgTrel.
changed drastically between phase 1 (>65 %) and phase 3
(≪ 50 %) (Fig. 3c). In the LMHC + MNR microcosms the
P-HgTrel. was high during phase 1 (>65 %) and
fluctuated in phase 3 (≪ 50 %) (Fig. 3c). Cu
concentrations gradually decreased during the course of the experiment (Fig. 7a). Arsenic concentrations simultaneously increased with the release of Fe
during the whole incubation (Fig. 7b).
Soil solution dynamics in pasture field soil (LMHC) incubations for
redox potential (a), redox-reactive elements (NO3-, Mn, P-Mn, Fe, P-Fe,
[SO42-] : [Cl-]) (b–e) and dissolved organic carbon (f). Lines
between points were plotted to improve readability. The gray area indicates
the drained period.
Soil solution dynamics in pasture field soil (LMHC) incubations for
Hg (a–c) subdivided into phases (1–3). Lines between points were plotted to
improve readability. The gray area indicates the drained period.
Soil solution dynamics in pasture field soil (LMHC) incubations for
Cu (a) and As (b). Lines between points were plotted to improve readability.
The gray area indicates the drained period.
Colloidal Hg (AF4)
Hg-bearing colloids were detected in all soil solution samples of HMLC
incubations. Due to low signal-to-noise ratios (<3), we did not
detect colloidal Hg in samples of the LMHC incubations. Figure 8 shows the
evolution of concentrations and relative proportions of HgT size fractions.
Generally, changes in proportions were apparent during phases of Hg release
and decrease in soil solution, but little change was observed when Hg
concentrations were stagnant (HMLC + MNR, phase 3). The proportion of truly
dissolved HgT<1kDa varied between 0 % and 67 % in the HMLC
experiment and was high during Hg release to soil solution (phases 1 and 3)
(Fig. 8). In the HMLC + MNR treatment, HgT<1kDa values were lower and ranged
between 0 % and 29 %. The colloidal Hg can be divided into three main
fractions (Fig. 9). The first Hg colloidal fraction showed a main peak
ranging between 1–40 kDa (dh< 6 nm) and was associated with
UV254nm-absorbing compounds and various metals (Mn, Fe, Cu, Ni, Zn).
This fraction was interpreted as humic substance type Hg–NOM. The
proportion of this colloidal Hg fraction varied with no specific trends from
11.5 % to 23.3 % in HMLC and 13.6 % to 38.6 % in HMLC + MNR throughout
the course of the experiment. A second fraction of Hg colloids ranged
between 6 and 20 nm. This well-defined size fraction was eluting in the
tail of the first fraction for other metals (e.g., Fe, Mn, Cu) but did not
overlap with UV254nm and fluorescence signals (Fig. 9). This fraction
could not be chemically defined but is hypothesized to consist of
β-HgS(s) colloids. In the HMLC run, we observed a decrease in the
proportion of these inorganic colloids from 28 % in phase 0 to 15.3 %
at the end phase 3 (Fig. 9). In the HMLC + MNR treatment, the proportion of
this fraction ranged between 29.5 % and 41.9 % during phases 1 and
2 and could not be detected during phase 3. Further, we observed a third
colloidal fraction that continued to elute after the stop of the AF4
crossflow, and it included colloids in the range of 30–450 nm (effective
cutoff of the filter used for the sample preparation). In some cases, this
fraction was better fitted using two overlapping populations (Figs. 9,
S9–S12). In all the cases, HgT signal was associated with those of other
metals and a slight bump of the UV254nm signal but more specifically an
increase in fluorescence signal associated with protein-like fluorophores.
This fraction decreased continuously in the HMLC runs during the incubation
from 32.4 % in phase 2 to 5.6 % in phase 2 and remained under 9.1 %
during phase 3. By contrast, the HMLC +MNR showed an increase in the
proportion of this fraction from 7.3 % in phase 1 to 25.3 % by the end
of phase 3 (Fig. 8). The deconvolution of the fractograms included an
intermediate fraction of Hg-bearing colloids ranging between dh=6 nm and dh=450 nm depending on the sample. This fraction was added
to refine the fractogram fittings but could not directly be associated with
another measured metal. This indicates that this population represents a
polydispersed Hg particle population, although in some cases the presence of
small Hg particles dominates. This broad fraction was not detected in HMLC + MRN treatments during phases 1 and 2 but made up >30 %
during phase 3.
Size distribution of Hg estimated after AF4 fractogram
deconvolution for Rep1 of cornfield soil incubation (HMLC and HMLC + MNR)
subdivided into phases (0–3). The concentration of HgT in size fractions was
calculated using an external calibration of the ICP–MS directly after the
AF4 run. The concentration of HgT in “<1 kDa” was calculated by
subtracting the sum of the fractions from the HgT concentration in the same
sample measured separately by ICP–MS. The fractograms of all analyzed time
points are shown in the Supplement (Figs. S9–S12).
Hg, Cu, Mn and Fe concentrations (a) and C signals (ICP–MS) and
UV254nm absorbance and fluorescence signals (b) in colloids as a function of
hydrodynamic diameter (related to retention times on AF4) in a sample from
HMLC at day 9 after flooding. These fractograms were obtained at linearly
decreasing crossflow from 2 to 0 mL min-1 over 20 min. The red line
indicates the time point where the crossflow reached 0 mL min-1. Areas
(yellow to red color) indicate size fraction ranges assigned during
deconvolution.
Net MeHg production in soil
Soil MeHg levels fluctuated over the course of the incubation experiment
(Fig. 10 and Table 2). The highest net MeHg production was observed during the
first flooding period for the treatments with manure (up to +81 %) and
during the draining phase for the treatments without manure (up to +73.1 %). We observed a significant decrease in MeHg / HgT and absolute MeHg
concentrations in all incubators during the second flooding period (Fig. 10). In all microcosms, MeHg / HgT increased by a factor of 1.18 to 1.36
throughout the incubation (Table 2).
Soil MeHg concentrations and MeHg / Hg ratios over the course of the
experiment for cornfield soils (HMLC, yellow/red) and pasture field soils
(LMHC, lime/green). The highest net methylation was observed during first
flooding for +MNR treatments and during the draining period for microcosms
without manure addition. A significant decrease in MeHg / Hg was observed
during the second flooding for all treatments.
DiscussionMercury release and sequestration
Cornfield soil (HMLC) and pasture field soil (LMHC) behaved differently in
this incubation experiment and will be discussed separately. In the
cornfield soil (HMLC) Hg and Mn releases were simultaneous and started when
soil solution Eh entered the field of Mn reduction below approximately 300 mV
(Figs. 2c, 3a), strongly suggesting that this Hg pool was released by
reductive dissolution of Mn oxyhydroxides. During all experiments, low
Hg : DOM ratios (≪ 1 nmol Hg (mg DOM)-1) suggest that
strong binding sites of DOM were never saturated with respect to mercury,
assuming a binding site [RS22-] density of 5 nmol Hg (mg DOM)-1 and that DOC is 50 % of the DOM (Haitzer et al., 2002). The low
Hg : DOM ratio suggests that Hg is mainly present as complexed with DOM given
reported strong interaction with thiol sites of DOM. However, these
assumptions might not reflect the actual composition of DOM, which might
drastically differ in amended soils (Li et al., 2019). Reductive dissolution
of Mn oxyhydroxides drives both (1) the release of labile Hg–NOM complexes
and Hg2+ sorbed on the oxide's surfaces and/or (2) enhanced the
degradation and mineralization of unsubtle NOM binding Hg in soils (Jones et
al., 2018). After Hg release (phase 1), Hg concentrations remained high, and
the relative particulate Hg fraction was low throughout the experiment. This
illustrates that the released Hg pool mainly originated from
Mn oxyhydroxides or degradation of suspended POM during Mn reduction.
However, the released Hg pool is relatively small compared to the HgT levels of
the soil. We estimate that about 12.8±4.2µg kg-1 Hg (0.02 % of HgTsoil) was evacuated by sampling during the experiment.
In this Fluvisol, Hg mobilization is thus mainly driven by reductive
dissolution of Mn oxyhydroxides. Direct mobilization of DOM was reported to
govern Hg levels in peat soils, Histosols or Podsols in boreal environments
(Åkerblom et al., 2008; Kronberg et al., 2016; Jiskra et al., 2017), or
floodplain soils with higher OC levels (Beckers et al., 2019; Wang et al.,
2021) in temperate soils.
Further, Hg mobilization was not simultaneous to Cu release. This was
reported for polluted soils with high Cu levels (Hofacker et al., 2013) and a
comparably low Hg / Cumolar ratio in the soil matrix. In neighboring
soils, the main Hg pool was previously reported as HgS(s) and Hg
complexed by recalcitrant NOM (Grigg et al., 2018). Earlier studies assumed
that 0.1 % to 0.6 % (w/w) of NOM was reduced sulfur with a high affinity to
Hg (Grigg et al., 2018; Ravichandran, 2004). Following this assumption,
reduced sulfur groups of the cornfield soil NOM could sorb between 11.9 and
71.9 mg kg-1 of Hg. The soils' high Hg concentration (47.3±0.5 mg kg-1) suggests that soil NOM thiol sites are likely saturated in
terms of Hg. Therefore, saturated NOM sorption sites do not compete with
Mn oxyhydroxide sorption sites, resulting in a substantial Mn-oxyhydroxide-bound Hg pool. This leads to a higher mobility of Hg upon reductive
dissolution of Mn oxyhydroxide compared to Fluvisols used in other
incubation studies (Hofacker et al., 2013; Poulin et al., 2016; Beckers et
al., 2019).
During the second flooding phase, the cornfield soil (HMLC) runs showed a
higher variability in redox-sensitive soil solution parameters (Fig. 2).
This might be explained as (1) a shift in microbial communities, (2) disturbance of the soil column by invasive soil sampling in between the
flooding periods or (3) uneven draining of the pore space after the first
flooding. It can also reflect how the redox cycle can be easily affected in situ. We
suggest that the second release of Mn and Hg in Rep1 is due to Mn
re-oxidation during the draining period and a second reductive dissolution
of Mn oxyhydroxides upon reflooding. This is supported by the elevated
Eh at the onset of the second flooding. Further, Mn reduction oxidation
and reduction cycles were shown to enhance the degradation of NOM to more
labile forms (Jones et al., 2018), which might contribute to the
degradation/mineralization of recalcitrant Hg–NOM. The HMLC Rep3 showed a
second release of Hg without a remobilization of Mn. Changing redox
conditions have been shown to enhance microbial respiration and therefore
NOM degradation (Sunda and Kieber, 1994). Thus, we interpret the second Hg
release in Rep3 as a degradation/mineralization of NOM that bound Hg.
The carbon amendments were reported to decrease total Hg release in polluted
floodplain soils (Beckers et al., 2019) but may have a mobilizing effect in
NOM-depleted environments (Eckley et al., 2021). The addition of manure
accelerated the release of Hg through reductive dissolution of Mn
oxyhydroxides in the cornfield soil (HMLC). Mercury was released 4 d
earlier, as a result of additional labile carbon of the liquid manure (1) acting as an electron donor enhancing microbial soil reduction (Liu et al.,
2020) and (2) acting directly as a reductant of the Mn oxyhydroxides (Remucal and
Ginder-Vogel, 2014). In the manure treatment, we observed a fast decrease in
Hg concentration and a constantly high proportion of particulate
P-HgTrel. even after the plateau of Mn concentration in soil solution
and the relative decrease in particulate Mn. The addition of manure acts as a source
of POM (manure was sieved to <500µm) and increased DOC
by approximately 20 mg L-1. Sorption of Hg is directed towards thiol
rich high-molecular-weight NOM (Liang et al., 2019) following different
ligand exchange reactions (e.g., carboxyl groups to thiol groups) which
happen within days (Miller et al., 2009; Chiasson-Gould et al., 2014). The
constant of P-Hgrel proportion is suggested to be partly caused by the
complexation of dissolved Hg with the added POM of the manure.
In addition, we visually observed black precipitates (Fig. S13) and the
decrease in [SO42-] : [Cl-] ratios (Fig. 2g) at the onset of Hg
decrease (phase 2) in the microcosms with manure addition. This indicates
the precipitation of sulfide mineral particles. Although redox potential
measurements did not indicate sulfate reduction, the monitoring of Eh
in soil solution provides only a qualitative measure in a complex soil
system. We suggest that formation and aggregation of β-HgS(s)
explain the faster decrease in the manure-amended experiment. Furthermore,
formation of metacinnabar β-HgS(s) was observed under oxic
conditions by conversion of thiol-bound Hg(SR)2 (Manceau et al., 2015).
The formation and aggregation of β-HgS(s) are further supported by
AF4 results (Sect. 4.2).
Hofacker et al. (2013) reported a
quantitatively relevant incorporation of Hg into metallic Cu0
particles. However, we do not consider this a relevant pathway, due to the
relatively high Hg / Cumolar ratio in our soil compared to Hofacker et
al. (2013). Although the simultaneous decrease in Hg and Cu may be
interpreted as the immobilization of Hg through incorporation into metallic
Cu particles, (i) we did not observe the formation of colloidal Cu associated
with Hg, and (ii) relatively high Hg / Cu molar ratios indicate that
the decrease in Hg in the soil solution cannot be solely explained by this
mechanism as Hg would be marginally incorporated into metallic Cu0
particles.
Also, Hg in soil solutions is volatilized by reduction of Hg2+ to
Hg0 (Hindersmann et al., 2014; Poulin et al., 2016; Li et al., 2021).
Our experimental design did not allow for quantification of gaseous Hg0,
and it may have exited the microcosms since they were only sealed with
Parafilm. Reduction of Hg2+ may happen both biotically (Grégoire
and Poulain, 2018) and abiotically under UV light and in the dark (Allard
and Arsenie, 1991). Biotic reduction is a detoxication mechanism of bacteria
carrying merA genes in Hg-polluted environments. Biotic volatilization has been
observed in neighboring soils of our sampling site (Frossard et al., 2018).
Organic amendments and high Hg levels have been shown to increase the
abundance of Hg-reducing bacteria (Hu et al., 2019). Further, dark abiotic
reduction of Hg2+ complexed to functional groups of DOM in soils has
been demonstrated (Jiang et al., 2015). However, it is unlikely that Hg
reduction can solely explain the decrease in Hg in the soil solution in our
microcosms. We therefore interpret the decrease in Hg concentration to be
due to a combination of manure NOM complexation and sequestration together
with the formation of HgS(s) during flooding. Our data show that
manure addition may have an immobilizing effect on Hg in flooded soils. By
contrast, carbon amendments may increase Hg mobility and methylation in NOM-depleted and cinnabar-rich mountain soils (Eckley et al., 2021).
In the pasture field soil (LMHC), soil solution Hg concentrations remained
at low levels (<0.16µg L-1 Hg<0.02µm) during
the whole experiment in both treatments (Fig. 6a). Unlike in the cornfield
soil (HMLC), we did not observe a simultaneous release of Hg upon Mn
reduction (Fig. 5c). We explain this with the not completely Hg-saturated
NOM in this soil, if we assume that 0.1 %–0.6 % (w/w) of NOM was reduced
S with a high affinity to Hg (Grigg et al., 2018; Ravichandran, 2004;
Skyllberg, 2008). Thus, the pasture field soil has a rather limited pool of
labile Hg compared to the cornfield soil. Both Hg<0.02µm and
Hg<10µm negatively correlate with the sum of sampled soil solution
(R2=-0.841, p=<0.001) during both flooding periods and decreased fast. This suggests that the concentration gradient between
supernatant artificial rainwater and the soil solution contributed to the
fast exhaustion of the small labile Hg pool in pasture field soil. The
presence of this concentration gradient in our incubation setup is confirmed
by the continuously decreasing concentrations of conservative ions
(Cl-, Na+, K+) in soil solutions of the HMLC runs (Figs. S7, S8). The relatively high proportion of particulate Hg vastly
decreased during the draining period (Fig. 3b, c), and we speculate that this
change is a result of the mobilization of the POM–Hg pool by
mineralization/degradation of NOM which sorbed Hg during the draining period
(Jones et al., 2018). In summary, flooding of the pasture field soils
mobilized only a small pool of particulate-bound Hg, which was exhausted
within the first flooding period.
Colloidal Hg
For runs without manure, AF4 results show that the Hg released from
Mn oxyhydroxides (Sect. 4.1) was dominated by truly dissolved Hg
(Hg2+ or LMW–NOM–Hg) (Fig. 8). The high Cl- concentrations (up
to 800 mg L-1, Fig. S14) likely influenced the Hg speciation in the
soil solution, as chloride is a main complexant for Hg2+ (Li et al.,
2020; Gilli et al., 2018). During Hg release, the proportions of larger Hg
colloids (>25 nm) decreased. The stable proportion of humic-substance-bound Hg and inorganic Hg colloids between 6 and 25 nm
indicates that once released no major adsorption or aggregation of truly
dissolved Hg and larger colloidal Hg occurs. Additional complexation of Hg
by DOM can be excluded if we assume the saturation state of thiol sites of
the NOM pool in the soil (Sect. 4.1). These observations illustrate the
remarkably high Hg mobility and potentially increased bioavailability
(proportion of truly dissolved Hg) to Hg-metabolizing microorganisms
compared to other studies (Hofacker et al., 2013; Poulin et al., 2016).
The authors either did not observe Hg in the truly dissolved form, or they observed a
decrease to low levels within the first days of incubation. Overall, the
released Hg from cornfield soil (HMLC) shows a high mobility and might
represent a possible threat to downstream ecosystems and a source for Hg
methylating bacteria. However, the total Hg released and sampled from soil
solution represents a rather small pool (12.8±4.2µg HgT kg-1 soil) of the total Hg (47.3±0.5 mg kg-1). Further
work would be needed to establish a Hg flux model to better understand in situ soil
Hg mobility in these soils.
The manure addition had a key effect on the proportions of colloidal
fractions in soil solution and overall led to a low proportion of truly
dissolved fraction (Fig. 8). We suggest that the distinct fraction of
colloids with dh= 6–25 nm represents metacinnabar-like
HgS(s) colloids (Gerbig et al., 2011). This is supported by the onset
of sulfate reduction in phase 2 (Rivera et al., 2019; Poulin et al., 2016)
and reported Hg–NOM interactions that may cause the precipitation of
Hg-bearing sulfide phases (FeS(s), β-HgS(s)) (Manceau et al.,
2015). The size of β-HgS(s) nanoparticles formed
from free sulfide is dependent on the sulfide concentration as well as on
the Hg : DOM ratio (Poulin et al., 2017). The formation of a distinct size
fraction of HgS(s) has been experimentally observed at comparable Hg : DOM ratios
(Gerbig et al., 2011). The Hg colloidal distribution was dominated by the
presence of large fractions (dh= 30–450 nm). Larger organic acids
with high aromaticity usually contain higher proportions of thiol groups
than smaller molecules and selectively complex Hg (Haitzer et al., 2002).
This suggests that Hg complexation is kinetically driven, and it can shift
from LMW–DOM to larger NOM and larger aggregates of POM as supported by
earlier incubation experiments (Poulin et al., 2016). We therefore interpret
that the relative increase in Hg colloids with dh= 30–450 nm
(Fig. 8) is caused by (1) complexation of the released dissolved
Hg<1kDa by strong binding sites of thiol-rich NOM in larger
clay–organic–metal complexes and (2) the aggregation of HgS(s) colloids
during the experiment. Although the presence of humic substances and
larger NOM was shown to narrow the size range of HgS(s) nanoparticles
precipitating from solution (Aiken et al., 2011), through time, these
colloids may grow, aggregate and form clusters in a wide size distribution
(Deonarine and Hsu-Kim, 2009; Poulin et al., 2017). Thus, their aggregation
during the draining period may explain the decrease in monodisperse Hg-bearing colloids, also leading to sequestration of Hg in the soil matrix,
without remobilization during the second flooding. Our data suggest metacinnabar formation (β-HgS(s)) in a distinct size fraction
(dh= 6–25) and their aggregation to large fractions (dh= 30–450 nm) at environmental conditions in real-world samples.
Net MeHg production in soil
The studied soils show uncommonly high initial MeHg levels (6.4–26.9 µg kg-1) when compared to other highly polluted mining or
industrial legacy sites (Horvat et al., 2003; Neculita et al., 2005; Qiu et
al., 2005; Fernández-Martínez et al., 2015), supposedly as a result
of a flooding event prior to sampling resulting in a net MeHg production.
Still, we observed significant net MeHg production during the first 28 d
of the incubation, resulting in even higher MeHg concentrations of up to
44.81 µg kg-1 (Table 3; Fig. 10). Soils treated with manure
showed a faster net MeHg production, with the highest increase in MeHg during the
first flooding period. Controls showed the highest net MeHg production during
the draining period and reached similar levels of MeHg at the start of the
second flooding on day 28 (Fig. 10). For cornfield soil (HMLC), both
treatments show a high concentration of bioavailable Hg2+ or Hg
associated with labile NOM (HgT<0.02µm>15µg L-1) in soil solution during the first flooding. Net MeHg production is
therefore rather limited by cellular uptake of Hg or the microbial activity
of methylating microorganisms than bioavailability. Thus, we interpreted the
addition of labile carbon in the form of manure to result in a higher
microbial activity and net MeHg production during the first flooding period.
However, we neither assessed the activity nor the abundance of Hg
methylating bacteria such as sulfate reducers (SRB), Fe reducers (FeRB),
archaea or firmicutes (Gilmour et al., 2013). In the runs without manure
addition, a substantial part of Hg was methylated during the draining
period. This indicates that even if low concentrations of Hg are released
(LMHC microcosms day 14: HgT<0.02µm<50 ng L-1) a
substantial amount of Hg can be methylated. Micro- and meso-pore spaces with
steep redox gradients act as ideal environments for microbial methylation
even in a drained and generally aerobic system (e.g., HMLC without manure
during the draining period).
Soil MeHg concentrations and net methylation (MeHg / Hg) over the
time of the experiment.
Further, we observed a decrease in absolute MeHg concentrations in all
microcosms during the second flooding period. Oscillating net
de-/methylation in environments characterized by flood–drought–flood cycles
has been reported earlier (Marvin-DiPasquale et al., 2014). Degradation of
MeHg was reported to happen either abiotically by photodegradation or
biotically by chemotrophic reductive or oxidative demethylation by
microorganisms carrying the mer operon (Grégoire and Poulain, 2018).
Photodegradation of MeHg can be excluded as the experiment was conducted in
the dark. However, demethylation could have happened as biotic reductive
demethylation. A possible explanation is a MeHg detoxification reaction by
microorganisms carrying the mer operon (merB) (Hu et al., 2019; Frossard et al.,
2018; Dash and Das, 2012). However, we can only hypothesize about
demethylation mechanisms, as neither communities (DNA) nor gene expression
(mRNA) dynamics in the soils were analyzed during the experiment.
Experimental limitations
Incubation experiments on a laboratory scale are a common way to study the
changes in mobility of trace elements in floodplain soils (Gilli et al.,
2018; Frohne et al., 2011; Poulin et al., 2016; Abgottspon et al., 2015).
These study designs allow for controlled conditions and replicable results.
However, controlled experiments usually fail to cover the complexity of a
real floodplain soil system (Ponting et al., 2020). Our study design did not
involve temperature gradients, realistic hydrological flow conditions or
intact soil structure. In this study, the artificial rainwater and the soil
were equilibrated by shaking for a few minutes. However, the equilibration
appeared to be incomplete with respect to highly soluble chloride-bearing
minerals for the experiment with cornfield soil (Fig. S14). The incomplete
equilibration is indicated by the temporal patterns of conservative ions
(Cl-, K+ and Na+) in soil solution (Figs. S7, S8) and the
difference in Cl- concentration between the soil solutions at t=6 h
and the same water–soil mixture shaken for 6 h (Fig. S14). These patterns
are a result of a concentration gradient between supernatant water and the
solution in the soil pore space. They only became visible due to high
levels of conservative ions to start with, which most likely stem from a
fertilization event prior to sampling the soil. Infiltration of supernatant
water was facilitated by the sampling of 4 %–6 % of the total added water
at each time point. This resulted in a dilution of the soil solution.
Consequently, the continuous decrease in sulfate was not directly
indicative of sulfate reduction but rather the result of this dilution effect.
However, this effect did not directly affect the release of soil bound
elements (e.g., Hg, Mn, Fe, As) by reductive dissolution (Figs. 2, 3, 4).
It should also be noted that high initial Cl- concentrations in the
soil solution may influence Hg solubility since Cl- is a complexant
for Hg2+ (Li et al., 2020), and this warrants further studies on the
role of inorganic fertilization in Hg mobility.
Conclusions
We studied the effect of manure addition on the mobility of Hg in soil
during a flooding–draining experiment. We observed formation and size
distribution changes of Hg colloids (β-HgS(s), Hg–NOM) at
environmental conditions in soil solution by AF4–ICP–MS. The results of
this study show that manure addition (1) diminished HgT mobility, (2) facilitated Hg complexation with fresh NOM and formation of
β-HgS(s), and (3) had only a limited effect on net MeHg production in
polluted and periodically flooded soils.
Mercury was mobilized upon reductive dissolution of Mn oxyhydroxides in
highly Hg-polluted (47.3±0.5 mg kg-1) and NOM-poor soils. The
application of manure accelerated the release of Hg, facilitated the
formation of colloidal Hg and exhausted the mobile Hg pool within the first
7 d of flooding. This prevented Hg remobilization during the second
flooding period. Contrastingly, Hg was mainly released as particulate-bound
Hg in soils with moderate Hg pollution (2.4±0.3 mg kg-1) and
high NOM levels, presumably due to its higher soil organic carbon content.
This relatively small pool of particulate Hg was exhausted within the first
flooding period. In both soils, soil reduction enhanced net MeHg production
of a substantial part of the Hg pool as confirmed by MeHg formation upon
flooding–draining cycles. However, MeHg was either subsequently removed from
the soil by advective transport of dissolved MeHg in the soil column or
transformed by reductive demethylation. We suggest that the temporal changes
in net MeHg production are limited by microbial activity of Hg methylators,
given the similar net MeHg production in treatments and soils with variable
dissolved Hg levels. Microbial activity is likely to be stimulated by manure
addition.
The release of Hg from polluted soils to downstream ecosystems does depend
on both biogeochemical conditions and hydrological transport. Our
experiment shows that redox oscillations (flooding–draining–flooding cycles)
of a polluted floodplain soil are likely to induce pulses of both Hg and
MeHg to the downstream ecosystems. This is supported with earlier studies
(Poulin et al., 2016; Frohne et al., 2012; Hofacker et al., 2013). In
contrast to NOM-rich soil systems, we show that the Mn dynamics may govern
the release of Hg in highly polluted soil systems low in NOM. Further, the
application of additional NOM in the form of manure facilitated soil reduction,
contributed to the transformation of Hg towards less mobile species and reduced
the Hg mobilization. However, effects of carbon amendments (organic
amendments or biochar) are contrasting between enhancing (Li et al., 2019;
Eckley et al., 2021) and diminishing (Beckers et al., 2019; Wang et al.,
2020, 2021) Hg mobility. We therefore stress the need for
characterization of soil properties and especially NOM in future studies
focusing on Hg mobility upon organic amendments (Li et al., 2019). We
further emphasize the need for field trials integrating biogeochemical
processes, hydrological transport and Hg soil–air exchange in order to
establish Hg flux models to better understand in situ soil Hg mobility.
Data availability
Details of analytical methods and AF4–ICP–MS fractograms are given in the
Supplement. A complete dataset of the data used in this study is accessible
at 10.5281/zenodo.4715110 (Gfeller et al., 2020).
The supplement related to this article is available online at: https://doi.org/10.5194/bg-18-3445-2021-supplement.
Author contributions
AM and LG designed the study. LG and AW performed the incubation
experiments. LG and IW performed laboratory analyses. LG and IW performed
the data analysis. AM and VIS supervised and financed the study. LG prepared
the manuscript with contributions from all co-authors.
Competing interests
The authors declare that they have no conflict of interest.
Acknowledgements
We acknowledge Patrick Neuhaus, Jaime Caplette, Kevin Trindade, Killian Kavanagh and Daniela Fischer for the help in the laboratory. We thank Tobias Erhardt at the Climate
and Environmental Physics (CEP) at University of Bern for the ICP–TOF–MS
analyses, Urs Eggenberger for the access to the X-ray diffraction spectrometer, and Stephane Westermann at the Dienststelle für Umweltschutz
(DUS) of the Canton Wallis for the help with site selection and sampling
permissions. Soil temperatures have been provided by MeteoSwiss, the Swiss
Federal Office of Meteorology and Climatology. Klaus Jarosch and Moritz Bigalke of the soil science group at the Institute of Geography at
University of Bern gave valuable advice during the writing process.
Financial support
This work was funded the Swiss National Science Foundation (SNSF, grant no.
163661). Vera I. Slaveykova and Isabelle Worms were supported by the SNSF R'Equip
project no. 183292.
Review statement
This paper was edited by Perran Cook and reviewed by Brett Poulin and Jan G. Wiederhold.
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