Climate induced microbiome alterations increase cadmium bioavailability in agricultural soils with pH below 7

Abstract

Climate change and metals independently stress soil microbiomes, but their combined effects remain unresolved. Here we show that future climate affects soil cadmium through altered soil microbiome and nutrient cycles, with soil pH as critical factor. In soils with pH<7 and during summer temperatures, future climate increased porewater cadmium, shifting total and potentially active taxonomic microbiome structures. Microbial ammonium oxidation released protons liberating cadmium through cation exchange from mineral surfaces. When porewater cadmium levels became toxic to non-cadmium-tolerant bacteria, microbial activity, and nutrient cycling decreased, reducing carbon and nitrogen emissions. In contrast, pH>7 soil show no climate impacts on cadmium mobilization, though imprints on microbiome structure were apparent. Subsequent nutrient cycling increased under future climate, stimulating soil respiration and nitrous oxide release. These findings underscore complex interactions between climate change and soil contaminants affecting the soil microbiome and its activity and highlights potential impacts on crop production, groundwater quality, and climate feedback.

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communications earth & environment Article
https://doi.org/10.1038/s43247-024-01794-w
Climate induced microbiome alterations
increase cadmium bioavailability in
agricultural soils with pH below 7
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Sören Drabesch1,2,3, Oliver J. Lechtenfeld 4,5, Esmira Bibaj1,3, José M. León Ninin 6,
Juan Lezama Pachecco7, Scott Fendorf 7, Britta Planer-Friedrich 6, Andreas Kappler 2&
E. Marie Muehe1,3
Climate change and metals independently stress soil microbiomes, but their combined effects remain
unresolved. Here we show that future climate affects soil cadmium through altered soil microbiome
and nutrient cycles, with soil pH as critical factor. In soils with pH<7 and during summer temperatures,
future climate increased porewater cadmium, shifting total and potentially active taxonomic
microbiome structures. Microbial ammonium oxidation released protons liberating cadmium through
cation exchange from mineral surfaces. When porewater cadmium levels became toxic to non-
cadmium-tolerant bacteria, microbial activity, and nutrient cycling decreased, reducing carbon and
nitrogen emissions. In contrast, pH>7 soil show no climate impacts on cadmium mobilization, though
imprints on microbiome structure were apparent. Subsequent nutrient cycling increased under future
climate, stimulating soil respiration and nitrous oxide release. These findings underscore complex
interactions between climate change and soil contaminants affecting the soil microbiome and its
activity and highlights potential impacts on crop production, groundwater quality, and climate
feedback.
Taxonomically and functionally diverse soil microbiomes are indispensable
for agricultural productivity under global food demands1–10. Nevertheless,
environmental stresses, such as climate change and metal contamination,
can compromise soil health and microbial dynamics11,12. Metals, like the
common cation contaminant cadmium (Cd), naturally occur at background
quantities in all soils13, but are rising due to intensified cultivation, and
(legacy) atmospheric depositions due to mining or industry14.Cadmium
holds no known metabolic function15, is considered toxic to any life form by
inhibiting physiological processes in cells16–18, and ranked at position seven
by the Agency for Toxic Substances and Disease Registry.Yet,moderate
concentrations have been observed to stimulate microbial growth19 and
activity8,9,19, which is likely due to a stress response that enhances metabolic
processes19 and the activation of defense mechanisms9. Whether soil biota is
affected or not by heavy-metal-related toxicity depends on the bioavailable
fraction (i.e., dissolved or weakly adsorbed), rather than the total
concentration18,20. Metal bioavailability is governed by pH, organic matter
content and composition, and the type of minerals present. However, cli-
matic conditions also play a crucial role, not only by affecting these soil
properties21 but also by impacting microbial communities5and mineral
solubility22. Current evidence suggests that elevated temperature increases
physical Cd retention23,24, whereas impacts of increased atmospheric CO
2
or
changes in soil moisture have yet to be considered. Earth’schangingclimate
with shifts in the partial pressure of CO
2
, temperature, and precipitation
frequency and intensity has been investigated extensively for impacts on
microbial community composition, abundance, activity, and function5,11
Elevated temperature generally enhances microbial activity6,11,25,while
evaporation decreases soil moisture exerting drought stress on
microorganisms7. For elevated atmospheric CO
2
, contrasting findings were
reported with more studies observing an increase2,26, and less of a decrease27,
in microbial activity. While there is a growing body of research examining
1Plant Biogeochemistry, Department of Geosciences, University of Tuebingen, 72076 Tuebingen, Germany. 2Geomicrobiology, Department of Geosciences,
University of Tuebingen, 72076 Tuebingen, Germany. 3Plant Biogeochemistry, Department for Applied Microbial Ecology, UFZ –Helmholtz Centre for Environ-
mental Research, 04318 Leipzig, Germany. 4BioGeoOmics, Department of Analytical Chemistry, UFZ –Helmholtz Centre for Environmental Research, 04318
Leipzig, Germany. 5ProVIS –Centre for Chemical Microscopy, UFZ –Helmholtz Centre for Environmental Research, 04318 Leipzig, Germany. 6Environmental
Geochemistry, Bayreuth Center for Ecology and Environmental Research (BAYCEER), University of Bayreuth, 95447 Bayreuth, Germany. 7Soil and Environmental
Biogeochemistry, Stanford University, Stanford, CA, 94305, USA. e-mail: marie.muehe@ufz.de
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the singular effects of climate change or metals on the microbiome, there is a
lack of studies investigating the combined impacts of these factors on soil
microbiomes and the interplay between climate-induced microbiome
alterations affecting Cd bioavailability. Understanding this relationship is
crucial, as it directly influences the resilience of the global food system and
the broader ecosystem28,29.
Here, we evaluated the combined impacts of soil Cd and climate
change on Cd bioavailability and microbial community dynamics in
agricultural soils as a proof of concept study. Three European agricultural
soils were examined under a multifactorial design of the two stressors
(Supplementary Fig. S1 and Supplementary Table S1). The soils’native
Cd content under 0.2mg Cd kg−1dry soil represents uncontaminated
soils (termed low Cd). To vary total soil Cd contents, each soil was
additionally amended with two Cd amounts: +0.3 (termed moderate Cd)
and +2.2 mg Cd kg−1dry soil (termed high Cd) (Supplementary
Table S1). Moderate Cd mimicked subtle contamination, embodying
~80% of European and US topsoils13,30, while high Cd represented con-
taminated areas sparsely used for food production but relevant for
bioenergy production13,30,31. Given that the soils were from a similar cli-
matic region and parent rock, soil pH is isolated as the primary geo-
chemical variablein this study. Soil pH rangedfrom 6.3, 6.7 to 7.3, which
falls into the pH range suitable to most cultivated crops32,33 and should
cover the pH ranges of at least 40% of (agricultural) soils34,35. In these
soils, cadmium transitions from an increased mobility in the porewater to
almost all Cd adsorbed to SOM and minerals, respectively36. Incubated in
columns in exterior climatic controlled chambers with day and night
temperature cycles of a marine west coast climate (Cfb)37, soils were
exposed to ambient and projected future climatic conditions. Future
conditions simulated in the study considered a worst-case scenario,
comparing current CO
2
levels of 430 ppm
v
and temperatures with pro-
jected values of 800 ppm
v
and elevated temperature by 4 °C respective to
ambient temperature for the year 210038–40, with Liu & Raftery, 2021
proposing the current trajectory for a temperature rise until 2100
between 2.1 and 3.9 °C41. Under both climatic conditions, soils were
irrigated with sterilized artificial rainwater at fixed intervals. Thus, soils
exposed to future climatic conditions experienced lower minimum water
contents between irrigations due to higher evaporative losses (Supple-
mentary Fig. S2).
Results
Climate impact on soil cadmium
For pH 6.3 soils with a low Cd content, average porewater Cd was
approximately 0.11 ug L−1under today’s climatic conditions, stabilizing
after 35 days (Fig. 1A, Supplementary Table S2, Supplementary Fig. S3).
Future climatic conditions significantly raised average porewater Cd to
approximately 0.16 ug L−1, being 45% higher during summer temperatures
above 20 °C (Supplementary Fig. S2). Moderate and high Cd amendment
disproportionally increased porewater Cd to respective 1.03 and 8.30 ug L−1
as amended Cd associated to mineral surfaces rather than interiors17.Future
conditions caused significant 40% and 27% increases in porewater Cd,
respectively, comparable to low and moderate Cd soils. According to a
geochemical model42 climatic conditions did not affect the relative con-
tribution of DOC association versus free Cd2+in the porewater (Supple-
mentary Table S3). Observed climatic impacts on Cd porewater imprinted
also in 0.01 M CaCl
2
- and 0.1 M HCl-extractable Cd (Supplementary
Fig. S4A–F) and in more pronounced shifts from mineral-associated to
organic-bound Cd associations according to Cd K-edge EXAFS (Supple-
mentary Fig. S5).
For soils with a higher pH, lower porewater Cd concentrations were
observed and expected43 (Fig. 1C, D, Supplementary Table S2, Supple-
mentaryFig.S3D,E,andSupplementaryFig.S4G–I), as Cd can additionally
be incorporated in the crystal structure of carbonates as an impurity18,44 and
Fig. 1 | Porewater cadmium concentration in agricultural soils exposed to dif-
ferent climatic conditions. Porewater Cd concentration across an entire incubation
season for three agricultural soils with varying pH: (A,B) pH 6.3, (C) pH 6.7, and (D)
pH 7.3; and three different total soil Cd contents (low, moderate and high; for exact
Cd contents per soil see Supplementary Table S1)). Please note the difference in
y-axes scale due to soil pH impacts on overall porewater Cd levels. The data for the
soil with pH 6.3 is only shown for the warm period after day 30 of the season as
microbial impacts on porewater Cd at lower temperatures found in spring are less
pronounced. The grey shaded area highlights aqueous Cd concentrations above
1μgL
−1, relating to roughly the porewater concentration at which Cd toxicity
outweighs Cd stimulation effects on the microbiome. Soils were exposed to ambient
(430 ppm
v
CO
2
and ambient temperature) and future (800 ppm
v
CO
2
and +4°Cin
atmospheric temperature respective to the ambient set-up) climatic conditions.
Biological replicates: A, B = 5, C = 4, and D = 3 with B = 7, C = 10, and D = 15
timepoints across season. Squares represent the mean and the line in the box the
median; whiskers give the Q10 and Q90, and N the numbe r of datapoints per whisker
plot. Significant differences are indicated with p-values above plots. All p-values can
be found in Supplementary Table S2. Results of a generalized mixed linear model of
individual and interactive effects on porewater Cd are shown in Supplementary
Table S10.
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less H
3
O+ions compete for binding sites on SOM or oxide minerals. For the
pH 6.7 soil, porewater Cd remained below 4 μg L−1upon amendment and
significantly increased by 20 and 40% in low and moderate Cd soil after
9 days of future conditions. No statistically significant increase in porewater
Cd was observed for the high Cd soil, though a distinct increased trend is
seen in the time-resolved data. With the pH 7.3 soil containing carbonates
(Supplementary Table S1), porewater Cd remained below 1.0 ug L−1with no
observable climatic impact.
Alterations of the soil microbiome
Cadmium addition, after a 14-week acclimatization period before climate
onset, distinctly altered total and active microbial communities on phyla and
genera levels (Fig. 2A,F,J,SupplementaryFig.S6A,F,J,Supplementary
Table S4 with increasing distances, Supplementary Fig. S7A, and Supple-
mentary Fig. S8A). For pH 6.3 soils, Cd amendment caused a stepwise
divergence to significantly different potentially active and total micro-
biomes, while it developed to similar degrees in the pH 7.3 soil upon Cd
amendment (indicated by arrows in Fig. 2A, J and Supplementary Fig. S6A,
J; highlighted in red in Supplementary Table S4). Broad community
descriptors and 16S rRNA gene and transcript copy numbers indicated
recovery of microbiomes from Cd amendment after 14-week acclimatiza-
tion (Supplementary Fig. S9). Universally increasing or decreasing trends
for phyla across all soils were not identifiable, though trends within each soil
were consistent with increasing soil Cd contents pointing to metal tolerance/
stimulation45 and resistance genes46 (Supplementary Fig. S7A, Supple-
mentaryFig.S8A,SupplementaryTablesS5–7).
Upon imposing future climatic conditions, compositional differences
in taxonomic and potentially active microbial diversity evolved for all soils
andCdcontents(indicated by grey dashed arrows on the PCoA 1 coordinate
andblueandredcirclesinFig. 2, Supplementary Fig. S6). Notably, a distinct
separation of microbial communities due to climate was observed at day 32
of the pH 6.3 soil, becoming more pronounced over time. The degree of
development after 110 days relative to day 0 was significantly further for the
ambient compared to the future climate (indicated by yellow, green, and
blue color-coding in Supplementary Table S4). Differences between com-
munities exposed to ambient and future conditions were particularly sig-
nificant for the two Cd treatments comparedtothelowCdsoil(indicatedby
grey color-coding in Supplementary Table S4). For the pH 6.7 soil,
separation occurred at day 11 for future communities in moderate and high
Cd soils followed by communities in the low Cd soil at day 41. In the pH
7.3 soil, climate-driven separation happened around day 7 for moderate and
high Cd contents and started to occur slightly for the low Cd soil from day 12
onward.
Soil carbon usage
Porewater organic carbon levels ranged from 15 to 110 mg L−1for all three
soils and Cd contents and were not impacted by future climatic conditions
(Supplementary Fig. S10). LC-FT-ICR-MS fingerprinting of the organic
carbon in the pH 6.3 soil with low and moderate Cd contents revealed a
temporal evolution of distinct signatures, with Cd content exerting a
stronger influencethanclimaticcondition(Fig.3). Cadmium amendment
increased the mass to charge ratio. It decreased the nominal oxidation state
of the remaining carbon from day 35 onward. While future conditions
decreased the nominal oxidation state of the remaining carbon at day 67,
they did not affect the mass-to-charge ratio. Clustering 640 organic carbon
compounds into three size-polarity clusters revealed climatic sensitivity but
Temporal change
PCoA2[12.4%]
PCoA 1 [15.4 %]
Total soil Cd increase
PCoA2[12.4%]
PCoA 1 [15.4 %]
PCoA2[12.4%]
PCoA 1 [15.4 %]
day 0
PCoA2[12.4%]
PCoA 1 [15.4 %]
day 4
day 2 day 11 day 41
day 7 day 12 day 55
PCoA2[12.4%]
PCoA 1 [15.4 %]
PCoA 2 [10.5 %]
PCoA 1 [16.5 %]
F
PCoA 2 [10.5 %]
PCoA 1 [16.5 %]
MLKJ
GH I
PCoA 2 [10.5 %]
PCoA 1 [16.5 %]
PCoA2[8.3%]
PCoA 1 [14.9 %]
Total soil Cd increase
PCoA2[8.3%]
PCoA 1 [14.9 %]
day 32 day 110day 65 EDCA
soil pH 7.3
soil pH 6.7
PCoA2[8.3%]
PCoA 1 [14.9 %]
soil pH 6.3
B
day 0
PCoA2[8.3%]
PCoA 1 [14.9 %]
High Cd
Moderate Cd
Low Cd
FutureAmbient
Start
Cd
level
Climatic
condition
day 0
PCoA 1 [16.5 %]
PCoA 2 [10.5 %]
Fig. 2 | Principal coordinates analysis (PCoA) visualizing compositional whole
bacterial and archaeal community differences in Cd-bearing agricultural soils
exposed to different climatic conditions. PcoAs are based on 16S rDNA-calculated
Bray-Curtis dissimilarities and the first two principal coordinates are displayed for
three different agricultural soils with varying pH; (A–E) soil pH 6.3, (F–I) soil pH 6.7
and (J–M) soil pH 7.3 and three different total soil Cd contents (low (square),
moderate (circle) and high (triangle); for exact Cd contents per soil see Supple-
mentary Table S1) across incubation time. The soils were exposed to ambient (430
ppm
v
CO
2
and ambient temperature) (blue) and future (800 ppm
v
CO
2
and +4°Cin
atmospheric temperature respective to the ambient setup) (red) climat ic conditions.
Grey arrows in (A) through (E) indicate a community shift along the PCoA 1
coordinate due to time. Colored circles indicate community shifts due to climate.
PCoA was calculated for each soil independently considering all time points and
respective biological replication (soil pH6.3 = 5, soil pH 6.8 = 4, and soil pH7.3 = 3).
Average distance matrices are provided in Supplementary Table S4 for soil pH 6.3.
Data points with low sequencing depth were sorted out leading to less replication for
some time points (Methods).
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not to Cd amendment (Fig. 3D–G). Especially at day 67, clusters 2 and
3 showed a higher normalized intensity but also small differences between
climatic conditions earlier on. Carbon usage is reflected in respirational CO
2
output. In the pH 6.3 soil, 210 ± 11 g of CO
2
m−2were emitted within
92 days under low Cd and ambient climatic conditions versus 193 ± 5 g
CO
2
m−292 d−1under respective future conditions (Fig. 3C, Supplementary
Fig. S11, Supplementary Table S8). CO
2
emissions also decreased with Cd
amendment in this soil. During the initial weeks of the pH 6.3 soil incu-
bation, with moderate ambient temperatures and less drought stress, future
conditions stimulated CO
2
emissions across all Cd-treatments. However, in
the later experimental phase during summer with more pronounced
drought stress and temperatures above 25 °C47, there was an overall
reduction in soil respiration. In the pH 6.7 soil less CO
2
was emitted from the
high compared to the low and moderate Cd soil, though future conditions
increased respirational output from the low Cd soil only. Also in the pH
7.3 soil, CO
2
emissions decreased with increased soil Cd and future
conditions further decreased respiration for the native and low Cd but
enhanced CO
2
emissions for the high Cd.
Soil nitrogen cycling
Porewater ammonium and nitrate concentrations in the pH 6.3 soil
remained below 0.4 mg L−1and 80 mg L−1for all Cd contents and climatic
conditions, respectively, with similar trends in the pH 6.7 soil and no
changes in the pH 7.3 soil (Fig. 4, Supplementary Figs. S12, 13). Future
conditions resulted in decreased porewater ammonium and 1.8-fold higher
nitrate under all three Cd contents after 50 days. N
2
O emissions almost
significantly decreased due to the presence of more Cd in the pH 6.3 soil and
even further under future climatic conditions (Supplementary Fig. S11D–F,
Supplementary Table S9). In the pH 6.7 soil, ANOVA indicated a strong
interaction between climate and soil Cd at the 95% confidence level (Sup-
plementary Table S9). The exemplary ammonium-oxidizing microorgan-
ism Bacilliae increased its active relative abundance 1.2-fold under future
Fig. 3 | Dynamics of porewater organic carbon in an agricultural soil exposed to
two climatic conditions. Dissolved organic carbon of the pH 6.3 soil was char-
acterized with LC-FT-ICR-MS at selected time points (days 3, 12, 35, 67).
AWeighted average mass over charge (m/z) ratio, (B) weighted average aromati city
index, and (C) corresponding CO
2
emissions for the pH 6.3 soil with a low
(154 ± 44 μg kg−1dry soil) and moderate (486 ± 12 μg kg−1dry soil) Cd content.
DClustering of key molecular features (n=640) based on standardized mass peak
intensities and Euclidian distances. Cluster I (n=78) contains features of low
molecular weight and variable polarity (represented by the RP-LC retention time),
Cluster II (n=230) and III (n=332) are molecules with higher mass but contrasting
polarity and NOSC. E–GTemporal trends of clusters from (D) using summed
intensities of key features according to their polarity. In each panel the summed
intensities are normalized to 1 for each segment at the first sampling day. Soils were
exposed to ambient (430 ppm
v
CO
2
and ambient temperature) and future
(800 ppm
v
CO
2
and +4 °C in atmospheric temperature respective to the ambient
setup) climatic conditions. Averaged data represents the mean ± standard error of 3
biological replicates or minimum-maximum levels when 2 biological replicates
are taken.
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respective to today’s conditions. In connection with ammonium oxidation,
which liberated protons, the pH of the pH 6.3 soil stabilized be tween 6.3 and
6.8 during incubation and was reduced by 0.2 pH units for all Cd contents
under future climatic conditions (Fig. 4C, Supplementary Fig. S14). A
similar trend with a 0.1 pH unit decrease was observed for the pH 6.7 soil
under low and moderate Cd contents, and no difference in the pH 7.3 soil.
Discussion and Implications
Future climatic conditions, featuring doubled atmospheric CO
2
,4°Ctem-
perature increase, and reduced soil moisture, notably increased porewater
Cd in soils with pH below 7 (scheme in Fig. 5). This interactive effect
between soil Cd and climatic conditions (Supplementary Table S10)
occurred in native low Cd soils and Cd-amended soils at concentrations
relevant to agricultural and industrial contamination scenario13,30,31.The
heightened Cd bioavailability reflected in free Cd2+increases and extends to
deeper mineral-associated fractions, impacting HCl-extractable Cd and
altering Cd binding from oxide minerals to organic matter. Thus, it may
hold relevance for plant uptake48.
The observed changes in Cd mobility were season-dependent, pro-
minent during the transition from spring to summer with temperatures
exceeding 20 °C in pH 6.3 and 6.7 soils. Despite higher temperatures
enhancing Cd adsorption-capacity on minerals49, they can also increase
chemical reactions in the soil and therefore enhance weathering of
minerals50 that have Cd either adsorbed or incorporated in their crystal
structure. However, since a drop in pH and increased porewater Cd (Sup-
plementary Fig. S3 A–E) occur simultaneously, the rise in porewater Cd is
primarily attributed to soil acidification.Thisconclusionisfurtherrein-
forced by a principal component analysis of the geochemical porewater
parameters, which reveals a positive correlation between NO₃⁻and Cd, and a
negative correlation of both NO₃⁻and Cd with porewater pH (Supple-
mentary Fig. S15). The PCA results indicate that the nitrogen cycle plays a
pivotal role in regulating soil pH under future climatic conditions
(Supplementary Fig. S15), and minorly the dissolution of doubled atmo-
spheric CO
2
in soil solution51. Ammonium-oxidizing microorganisms, like
Bacilliae, were stimulated under future conditions (Fig. 4D), leading to
increased NH
4
+turnover to NO
3
−(Fig. 4A, B). This process released pro-
tons (Fig. 4C), facilitating cation exchange with mineral surface-bound Cd.
Released Cd most likely manifested to equal degrees in the free Cd2+and
DOC-associated fraction (Supplementary Table S3), indicating that parti-
tioning between DOC associated and free Cd2+is climate insensitive once
climate generally released more Cd into the porewater. Agricultural soils
with a more acidic pH than 6.3 could feature such a high mobility of Cd36,
that they may prove to be climate insensitive. However, the used empirical
geochemical model42 is based on sodium and DOC concentrations, as well as
solution pH, but it does not account for DOC composition. As a result, it
may overestimate the free Cd²⁺fraction and should be considered an
approximation. In soils with high carbonate contents (e.g. pH 7.3 soil with
0.3 %
w
inorganic carbon), cadmium mobilization was limited by the car-
bonate buffer system capturing protons. Additionally, cadmium associates
and integrates more with carbonates, forming calcite-otavite (Ca-CdCO
3
)
mixed minerals52, rather than binding to silicate and oxide mineral surfaces
asknownforlowerpHsoils
18,53. Given that approximately 30% of global
soilshaveapHabove7.3
34,35, climatic change is less likely to affect Cd
partitioning in these soils. This proof of concept study covered soils from pH
7.3 to 6.3, representing a pH range, relevant for crop production34,but
missing global soils beyond this range.
Shifts in microbial community taxonomy, diversity, metabolisms, and
activity were induced by climatic stressors, influencing observed changes in
Cd mobility. The introduction of Cd into the soil, followed by a 14-week
acclimatization period, led to distinct microbial communities upon the
onset of climate incubation (Fig. 2). The presence or a bsence of Cd-tolerance
genes54 likely caused this divergence, decreasing activity or cell death among
non-tolerant microbial members. As a result, lysed non-Cd-tolerant cells
became a primary food and energy source for surviving microbes, while cell
Fig. 4 | Snapshot of microbial nitrogen cycling in
the pH 6.3 soil exposed to different climatic con-
ditions. A Porewater ammonium (NH
4
+), (B)
porewater nitrate (NO
3
−), (C) porewater pH for the
pH 6.3 agricultural soil, bearing three different total
Cd contents (low (squares), moderate (circles), and
high (triangles), for exact Cd contents per soil see
Supplementary Table S1) and (D), relative potential
activity changes of the ammonium-oxidizer Bacil-
lales based on 16S rRNA transcript sequencing. The
soils were exposed to ambient (430 ppm
v
CO
2
and
ambient temperature) (blue, open/striped symbols)
and future (800 ppm
v
CO
2
and +4 °C in atmo-
spheric temperature respective to the ambient setup)
(red, filled symbols) climatic conditions. Relative
changes of the potential activity to microbial groups
are based on 16S rRNA transcript sequencing and
are calculated relative to the equivalent Cd con-
centration under ambient climatic conditions. Bio-
logical replicates = 5, error bars represent the
standard error. N represents the number of data-
points per whisker plot. Significant differences are
indicated with p-values above plots.
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walls and other microbially derived compounds with slower turnover were
enriched55,56. This process likely contributed to the shift towards larger and
more oxidized organic substances (Fig. 3)asalsoobservedinsoilprofiles57.
This adaptation allowed the microbial community to reach similar absolute
16S rRNA copy numbers within the 14-week acclimatization period.
Microbial adaption to climatic changes depends on soil history and
temperature58 and is promoted by temperature fluctuations, as imple-
mented in this incubation through natural day and night cycles59. However,
once future climatic conditions were imposed on the soils, compositional
differences in taxonomic and potentially active microbial diversity con-
tinued evolving for all soils, as indicated by soil-specific movements on
PCoA1and2coordinatesuntiltheend of incubation. While we imple-
mented sudden climatic changes in this study, which may not fully represent
the gradual nature of real-world climate change, the data indicate that the
microbial community acclimatized and continued adapting to these chan-
gesasexpected(Fig.2). These differences first appeared at different time
points in each soil depending on transitions to summertime temperatures,
under which higher Cd mobilities were observed. Taxonomic changes in the
total and potentially active microbiome were less rapid and pronounced
under different climates than Cd amendment. The 4 °C climate differences
had a more interfering impact on microbial activity and function rather than
survival contrasting the drastic effects of Cd amendment. Upon climate-
induced Cd mobilization, microorganisms lacking metal resistance genes
were inactivated, and some potentially perished, becoming a carbon source
of small and aliphatic substrates for thriving Cd-tolerant microorganisms1
(Fig. 3). This gradual shift in organic carbon signatures towards higher
aromaticity and m/z ratio has implications for the energy content of the soil
organic matter.
The potential link between a climate-induced shift in microbiome
dynamics and increased soil Cd mobility is evident through similar but less
pronounced shifts in PCoA 1 and 2 of native low Cd microbial communities
compared to Cd-amended soils (statistically significant average distances of
the Bray Curtis dissimilarity relative to the low Cd soil in Supplementary
Table S4). Notably, in the pH 7.3 soil, the microbial community responded
firmly even to low amounts of added Cd. While future climatic conditions
did not impact porewater nor bioavailable Cd, they promoted changes in
microbiome dissimilarity from day seven onwards for the moderate and
high Cd treatment, followed later by the low Cd soil. This suggests that an
interaction between climatic conditions and elevated Cd levels in these soils
stimulated the microbiome to adapt sooner than when only climatic con-
ditions were applied.
The overall porewater Cd level was crucial in determining whether
climate-affected Cd stimulated8,9,16,60 or adversely impacted16–18 microbiome
balance and functioning. In soils where porewater Cd remained below
approximately 0.8 to 1 μg L−1(non-grey shaded data in Fig. 1), microbial
diversity adapted, and activity was stimulated, as indicated by low impacts
on Faith phylogenetic diversity (Supplementary Table S15, 16) and
increased CO
2
and N
2
O emissions (Fig. 4). Conversely, when porewater Cd
exceeded approximately 1 μg L−1, notably in highly contaminated soils or
those with pH < 7, cadmium reached toxic levels, reducing initial micro-
biome phylogenetic diversity, which later recovered. Broad changes in
microbiome dissimilarity (Fig. 2) along with the shift toward larger and
oxidized organic molecules (Fig. 3) suggest that dying microorganisms
served as a food source for a subsequently thriving microbiome with spe-
cialized metabolic pathways55. In the specific light of the nitrogen cycle in the
pH 6.3 soil, climate-induced Cd mobility increases were associated with
increased nitrate due to the increased oxidation of ammonium (Fig. 4)and
likely enhanced nitrite oxidation towards nitrate.
Climate’s impact on metal mobility in soils holds significant implica-
tions, especially in cropland systems. We partially assessed this impact by
monitoring CO
2
and N
2
O emissions from agricultural soils. With ~30% of
global agricultural soils exhibiting a pH > 734,35, coupled with the rising trend
of nitrogen-based fertilizer application61, and the substantial radiative for-
cing of N
2
O21, the potential exists for higher agricultural greenhouse gas
Soil pH
8.07.57.06.56.0
Cd
Cd
Cd
Cd
pH
high buffer capacity
Carbonate minerals
Mineral (hydr)oxides
Cd Adsorbed /
bioavailable Cd
Porewater Cd
(blue ambient / red future
climatic conditions)
Bacteria
Dead bacteria
Cd
Cd
Cd
Cd
Cd Cd
Cd
Cd
NH4+NO3-
H+
H+
H+
Changing microbiome
due to climate and Cd
Cd mobility /
bioavailability
Shifts in microbial
taxonomy and activity
Toxicity feedback ~ 1 µg Cd L-1
A
D
Microbial
nitrogen cycling C
Porewater pH B
Microbial
respiration E
H+
Fig. 5 | Scheme illustrating climate change impacts on soil Cd bioavailability with
subsequent implications for microbiome dynamics. A Future climatic conditions
increase porewater and bioavailable Cd in acidic but not in alkaline soils as indicated
by blue colors for ambient and red colors for future climatic conditions (B) through
alterations in soil pH upon climate-driven differences in microbial activity such as
ammonium oxidation (C), cell death (indicated by broke bacteria), and altered
taxonomic and active microbial composition (D) (indicated by different colors and
shapes of bacteria). Porewater Cd concentrations below approximately 1 μg L−1have
a stimulatory effect on the soil microbiome (E), most prominent for neutral to
alkaline soils. At porewater Cd concentrations above approximately 1 μg L−1, sti-
mulatory Cd effects on the microbiome may be outweighed by toxic effects, which is
most prominent for soils with a pH < 7 or with high total Cd contents. Figure was
created with BioRender.com.
https://doi.org/10.1038/s43247-024-01794-w Article
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emissions globally than currently anticipated. Accurate projections are
crucial given that agriculture contributes 3.7% to anthropogenically-derived
greenhouse gas emissions62. Conversely, with 70% of global agricultural soils
exhibiting pH < 734,35, and considering a small fraction of agricultural soils
withpHbelow6andthuslesslikelyto be impacted by climate, there is a
heightened risk of increased mobile Cd transferring into the food chain even
at overall background total soil Cd contents, thereby endangering yields and
food quality. While current climate and crop production simulations con-
sider temperature, elevated atmospheric CO
2
, and rainfall patterns21,they
neglect the presence and bioavailabilityofmetalsinsoils.Thisoversightmay
exacerbate reduced ecosystem health, groundwater quality, food produc-
tion, and climate feedback through altered microbiome dynamics.
Material and Methods
Soil preparation and acclimatization
Three agricultural soils from the same parent rock material and the same
climatic region were sampled from farms in South-west Germany. Soils
were chosen for their difference in CaCl
2
-based pH and are subsequently
named by their pH in this manuscript; soil with pH 6.3 (Tuebingen), soil
with pH 6.7 (Dusslingen), and soil with pH 7.3 (Ehningen) (Supplementary
Table S1). According to Bradl et al. 2004, increased partitioning of Cd into
the porewater is especially relevant for soils with a pH from 5.5 to 7 as Cd
mobility drastically changes from almost everything mobile to immobilized
through adsorption to organic matter, minerals and incorporation into
mineral structures36.Weconfirmed this with a geochemical model for
free Cd2+in the porewater42, suggesting that up to 96% of the dissolved
Cd are present as free Cd2+available for plants48 in the pH 6.3 soil while in
the pH 7.3 soil only 60% are present as free Cd2+. At pH below ~5.5, most Cd
is expected to be dissolved; therefore, climatic effects are less likely to
imprint.
To characterize the soil, soil pH was determined with 2 g of air-dried
soilsin10mLof0.01MCaCl
2
solution after 1, 24, and 48 hours. Soil texture
was determined by wet-sieving and sedimentation analysis with PARIO soil
texture analyzer (Meter, USA). Total (organic) carbon and nitrogen con-
tents were determined on air-dried, ground soils by combustion on a soli
TOC cube (Elementar, Germany). Cation exchange capacity was deter-
mined from 1 g of wet soil in 25 mL 0.1 M BaCl
2
for 4 h (horizontal shaker
150 rpm, filtered through 0.45 μm PES filter), followed by cation quantifi-
cation with microwave plasma atomic emission spectroscopy (4200 MP-
AES, Agilent Technologies, USA). Total elemental contents, exchangeable
and bioavailable metal contents were quantified according to standard
protocols by aqua-regia-based microwave digestion63,0.01MCaCl
2
64 and
0.1 M HCl-extractions65, respectively, followed by quantification on an
Agilent 7900 ICP-MS (instrumental details under porewater analysis).
Water-extractable carbon and nitrogen were extracted from 5 g of air-dried
soil with 40 mL H
2
O for 1 h (horizontal shaker, 150 rpm, filtered through
0.45 μm PES filter)66. Organic carbon was quantified on a multi-N/C ana-
lyzer (Analytik Jena, Germany) and inorganic nitrogen species on an
AutoAnalyzer3 (SEAL Analytical, Germany).
All soils were prepared in the same way for experimental runs. First, the
soil was air-dried in the dark at 20 °C and sieved with a 2 mm mesh com-
bined with manual removal of plant and rock material67. Subsequently, the
soil was mixed and separated into equal parts using a sample splitter (RT 50,
Retsch, Germany). Next, soils were rewetted with artificial rainwater con-
taining either CdCl
2
(trace metal grade, Sigma Aldrich) for metal-bearing
set-ups or with CaCl
2
for low-Cd soils to account for chlorine toxicity effects
and equal ion amendments. Finally, metal stock solutions were applied with
a pressure sprayer and thoroughly mixed with a stirring adapter of a drilling
machine. The following Cd amendments were prepared for each soil: low
Cd with 2.5 mg Ca kg−1(CaCl
2
addition), moderate Cd with the addition of
0.3 mg Cd kg−1(CdCl
2
), and 2.2 mg Ca kg−1dry soil (CaCl
2
), and high Cd
with the addition of 2.5 mg Cd kg−1dry soil (CdCl
2
). Soils were acclimated
to metal amendments for 14 weeks, allowing Cd to establish a realistic
binding environment to the soil matrix68 and for the soil microbiome to
adapt to metal stress17. To do so, soils were run through three dry (5 %
gravimetric water content) and wet (20 % gravimetric water content) cycles
for acclimatization until the start of the experiment.
Experimental set-up
Black PVC-U tubes (inner diameter: 7.3 cm height: 33 cm) (Supplementary
Fig. S1), washed with 1 M HCl, MilliQ water and wiped with 80% ethanol,
were filled with 800 g of air-dried soil, and randomly distributed in climate-
controlled chambers (110 x 50 x 50 cm) made of poly(methylmethacrylate)
(front and top) and polyvinylchloride (ground, back and sides). Within each
tube, a rhizonsampler (RhizonFlex, 5 mm, 0.22 μm, Rhizosphere Research
Products, Netherlands) was installed horizontally, 8 cm below the soil
surface (Fig. S1). The three soils were run individually between 2019 to 2021.
The replication for each experiment was three, four, and five tubes for pH
7.3, 6.7, and 6.3 soils, respectively, as with each experiment our experience
and pipeline in sample handling allowed for more replication. The gravi-
metric water content of the soil was monitored every five days and weight-
adjusted to 25% with autoclaved artificial rainwater. Chambers were placed
outside to ensure natural day/night cycles (for detailed climatic conditions,
see Supplementary Fig. S2). Each chamber was equipped with a temperature
control unit using a raspberry Pi 3b+(Raspberry Trading., United King-
dom) and DS18B20 T-sensor (Analog-devices, USA) monitoring tem-
perature in four different spots within the chamber in minute intervals. The
ambient chambers featured the temperature and 430 ppmv atmospheric
CO
2
conditions of the surrounding environment, albeit on average about
2 °C higher than outside due to the greenhouse effect within the chamber
(Supplementary Fig. S2). The future chamber implemented future climatic
conditions by adding +4°Cand+370 ppmv CO
2
compared to the ambient
chamber through heater fans and CO
2
infusion, respectively. When the
mean difference in atmospheric temperature inside the ambient and future
chambers dropped below 3.9 °C, heating fans (HP8232, Phillips, Germany)
re-adjusted the temperature in the future chamber with simultaneous air
mixing without heating in the ambient chamber. Increased atmospheric
CO
2
concentration was achieved by mixing pumped (HiBlow HAP-100)
ambient air with CO
2
(industrial grade 99.5 %, Westfalen Gas, Germany)
using a two-tube gas proportioner (ColeParmer, USA) with flow tube 044-
40 with stainless steel float (Cole-Parmer) for ambient air and flow tube 042-
15 with glass float (Cole-Parmer, USA) for CO
2
.Theambientset-upwas
constructed the same, excluding the CO
2
flow. The flow was 36.6 L min−1
for ambient air and 14.66 mL min−1for CO
2
. By this, the chamber volume
was exchanged eight times per hour.
While climatic-controlled incubation studies allow for evaluating
subtle differences in biogeochemical processes, they also pose constraints,
such as a potential overestimation of mobilized Cd or greenhouse gas
emissions due to the homogenization of soils, amendment of Cd, and lack of
horizontal and vertical water flow. By acclimating the soil to Cd inputs for
weeks before the experiments, we aimed at a more realistic Cd binding68 and
an adaptation and recovery of the soil microbiome to the sudden stress20,
reflected by porewater Cd concentrations in the expected range of field
concentrations14. We could not account for an evolutionary adaptation of
the microbiome to a changing climate for decades into the future and are
constrained to using today’s soils with the current microbial community.
Porewater analysis
Porewater was extracted from each column through rhizon sampler using
20 mL syringes (Braun, Germany). Porewater pH was determined directly
in technical unicates per soil column in 1.5 mL of extracted soil solution with
an InLab Easy pH probe (Mettler Toledo, USA). The remaining porewater
was filtered through a 0.22 μm PES filter (pre-washed with 10 mL MilliQ to
remove organic carbon impurities). For quantification of dissolved Cd,
filtered porewater was diluted and acidified with nitric acid (trace metal
grade), stored at 4 °C in darkness, and analyzed in technical triplicates on an
Agilent 7900 ICP-MS (samples of pH 6.3 soil) and a Thermo Scientific
xSeries 2 ICP-MS (samples of pH 6.7 and 7.3 soil). For both, certified quality
controls at different concentrations were measured every 20th sample; for
instrumental drifts, Rhodium as an internal standard was used for
https://doi.org/10.1038/s43247-024-01794-w Article
Communications Earth & Environment | (2024) 5:637 7
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corrections. Porewater organic carbon (DOC) and nitrogen species (NO
3
−,
NH
4
+)werequantified on a multi N/C 2100 (Analytik Jena, Germany) and a
continuous flow analyzer (AA3, Seal analytical, USA), respectively. Details
on LC-FT-ICR-MS analysis are found in Methods.
Gas analysis
For determining greenhouse gas emissions from soils, columns were closed
air-tight with rubber stoppers (64.5–75.5, Deutsch Neumann, Germany)
from outside the chambers by leveling down screw rods. A cannula was
passed through the plug and connected to Tygon tubing (ID 1 mm, Isma-
tec), which ended at a 3-way valve (Braun, Germany) with two syringes. A
sample was drawn with one syringe to flush Tygon tubes with headspace gas,
and the sample was discarded. Subsequently, a headspace gas sample was
drawn with the other syringe after 0, 15, and 30 min and injected into
helium-flushed 20 mL crimped headspace vials. In total, 5% of the gas
headspace was removed. After sampling, tubes were opened again, allowing
the headspace to be exchanged passively by air circulation and diffusion. Gas
samples were analyzed immediately after sampling on a TraceGC1300
(ThermoFisher Scientific, modified by S +HA analytics, Germany), in
which the sample is split into two different column configurations (first
configuration: 30mlong,0.53mmID TGBondQcolumnand30m long,
0.53 mm ID Molsieve column; second configuration: 30 m long, 0.53 mm
ID TGBondQ column and a 30 m long 0.25 mm ID TGBondQ+column
(all ThermoFisher Scientific, USA), each connected to a Pulse Discharged
Detectors. This allowed the simultaneous quantification of CO
2
,N
2
O, and
CH4. Gas concentrations were quantified with external calibrations, and gas
fluxes were calculated by linear regression and cumulated throughout the
experiment. The cumulation was only performed for 12 h per day, as fewer
greenhouse gas emissions are expected during the night, which are not
included in the cumulative data. For the entire experiment, CH
4
emissions
were below the detection limit.
Soil sampling
Soil was sampled from the top 5 cm of soil in each column throughout the
experiment. Soil was homogenized with a sterile spatula (baked at 300 °C
for 3 h) and aliquoted into a sterile 2 mL microcentrifuge tube(Eppendorf,
Germany) for molecular ecology analysis, into a 15 mL centrifuge tube for
geochemical analysis, and a 2 mL microcentrifuge tube for water content
determinations. Geochemical samples were stored at −20 °C if samples
were not processed immediately. Samples for molecular ecology analysis
were stored at −80 °C. Soil selective extractions were performed to tease out
Cd binding differences due to climatic impacts and included 0.1 M HCl (all
soils) and 0.01 M CaCl
2
(only pH 6.3 soil) extractants. To do so, 1 g of fresh
soil was extracted with 9 mL of the respective extractant for 30min (0.1 M
HCl) or 60 min (0.01 M CaCl
2
) by 140 rpm horizontal shaking at room
temperature in darkness. Supernatants containing extracted elements were
collected after centrifugation at 7000 g for 5 min, 0.22 μm filtration, diluted
in 2% nitric acid, and stored at 4 °C and darkness until measurement on an
Agilent 7900 ICP-MS.
Extended X-ray absorption fine structure analysis
To identify Cd binding environments, soil samples were frozen,
freeze-dried, ground, and stored anoxically until measurement.
Samples were placed in aluminum sample holders (window 3 mm by
13 mm) and sealed with 0.5 mil Kapton tape from both sides. Cad-
mium K-edge extended X-ray absorption fine structure (EXAFS) data
were obtained at beamline 11-2 and 7-2 at the Stanford Synchrotron
RadiationLightsource(SSRL),MenloPark,USA.DuetothelowCd
concentrations, measurements were conducted using only the high
soil Cd samples. Additionally, up to 29 scans per sample were per-
formed to minimize noise and ensure accuracy. To overcome the low
Cd levels Spectra were calibrated against a Cd-foil, aligned, and
merged subsequently in Athena. Linear combination fittings were
performed in Athena with all samples and standards (Supplementary
Table S11).
Microbial community assessments
DNA/RNA were co-extracted from 0.6 g of wet soil with a phenol/chloro-
form/isoamyl alcohol phase separation extraction69. The DNA/RNA pellet
was dissolved in a 100 μL TE buffer. DNA and RNA contents were quan-
tified with a Qubit 2.0 fluorometer (Thermo Fischer Scientific), quality
checked for 260/280 ratios on NanoDrop (Thermo Fisher Scientific), and
aliquots were stored at −80 °C. For rRNA analysis, the extracts we re purified
from DNA with the InvitrogenTM TURBO DNA-freeTM Kit (Thermo
Fisher Scientific) and reversely transcribed into complementary DNA
(cDNA) using the InvitrogenTM SuperScriptTM III Reverse Transcriptase
Kit (Thermo Fischer Scientific). A PCR followed by a 1% (w/v) ethidium-
bromide-died agarose gel was performed to check successful DNA removal
and subsequent cDNA synthesis.
Gene and transcript copy numbers were quantified with quantitative
PCR (qPCR) using 1× SsoAdvanced Universal SYBR Green Supermix (Bio-
Rad Laboratories, Hercules, CA, USA). In a 10 μL reaction volume, 1μL of
template DNA ( ~4 ng) and cDNA ( ~ 0.5 ng) or a tenfold dilution series of
the standard plasmid DNA were annealed with 125 nM of primers 515-F
(5′-GTGYCAGCMGCCGCGGTAA-3′)70 and 806-R (5′-GGACTAC
NVGGGTWTCTAAT-3′)71 and 5% DMSO. As standard, a 16S rRNA
Thiomonas gene fragment was amplified. The qPCR program ran on the
CFX Connect Real-Time PCR Detection System (Bio-Rad Laboratories,
Hercules, USA) with 3 min at 95 °C, 40 cycles of 10 s at 95 °C and 30 s at
55 °C, followed by melting curve analysis. The data analysis was performed
using the Bio-Rad CFX Maestro software (Bio-Rad Laboratories,
Hercules, USA).
Microbial 16S rRNA genes and transcripts were amplicon-sequenced
using primers 515 F and 806R72. Quality and quantity of the purified
amplicons were determined using agarose gel electrophoresis. Subsequent
library preparation steps (Nextera, Illumina, USA) and sequencing were
performed using the 2 × 250 bp MiSeq Reagent Kit v2 on an Illumina MiSeq
sequencing system (Illumina, San Diego, USA). Adapters were trimmed
using the MiSeq Reporter Software. Reconstruction of 16S rRNA gene and
transcript sequences and taxonomic annotation was performed with
QIIME2 version 2019.10.073. Initially, data from the three sequencing runs
were treated separately and merged for each experiment after annotation.
Adapter-free sequences were imported into QIIME2 version 2019.10.0,
processed with DADA2 version 1.10.074 to eliminate PhiX contamination,
trim reads (before the median quality drops below 30), merge read pairs, and
remove PCR chimeras. Alpha rarefaction curves were produced with the
QIIME2 diversity alpha-rarefaction plugin, indicating that the samples’
richness had been fully observed (Supplementary Table S12). A Naive Bayes
classifier was fitted with 16S rRNA gene sequences extracted with the PCR
primer sequences from the QIIME compatible, 99%-identity clustered
SILVA v132 database75. ASVs were classified by taxon using the fitted
classifier76. Relative abundance per sample and the remaining ASVs were
extracted using the feature table.
Before alpha and beta diversity analyses, the data were rarefied using a
specific sampling depth for each experiment. Diversity metrics (alpha
diversity: Shannon index, Pielou’s evenness, and observed ASVs; beta-
diversity: Bray-Curtis dissimilarity) were calculated using the core-diversity
and emperor plugins within QIIME2 (Supplementary Tables S13–18).
LC-FT-ICR-MS. Original filtered porewater samples from the pH 6.3
treatments with low (three replicates) and moderate Cd (two replicates)
and four time points (days 3, 12, 35, 67) were stored at −20 °C until
analysis. After thawing, samples were filtered with 0.2 μm regenerated
cellulose filters (Minisart RC4, Sartorius, Germany) before directly
injecting 100 μL into a UPLC system coupled to an FT-ICR-MS. The
method was adapted from Han et al. 77 and is suitable to separate and
detect dissolved organic matter. Briefly, the chromatographic separation
was performed on reversed phase polar end-capped C18 column
(ACQUITY HSS T3, 1.8 μm, 100 Å, 150 × 3 mm, Waters, Milford, MA)
equipped with guard column (ACQUITY UPLC HSS T3 VanGuard
Precolumn, 100 Å, 1.8 μm, 2.1 mm × 5 mm, Waters). An FT-ICR mass
https://doi.org/10.1038/s43247-024-01794-w Article
Communications Earth & Environment | (2024) 5:637 8
Content courtesy of Springer Nature, terms of use apply. Rights reserved

spectrometer equipped with a dynamically harmonized analyzer cell
(solariX XR, Bruker Daltonics Inc., Billerica, MA, USA) and a 12 T
refrigerated actively shielded superconducting magnet (Bruker Biospin,
Wissembourg, France) was coupled to the LC system. An electronspray
ionization source (Apollo II, Bruker Daltonics Inc., Billerica, MA, USA)
was used in negative mode (capillary voltage: 4.3 kV, nebulizer gas
pressure: 1.0 bar, dry gas temperature: 250 °C, dry gas flow rate: 8.0 L/
min). Mass spectra for LC-MS measurements were acquired in broad-
band mode (147.41 to 1000 m/z) with a transient size of 2
MWord(∼0.84 s FID) and full profile mode. The ion accumulation time
(IAT) was set to 8 ms.
DOM from the porewater eluted between 9.8 and 19.9 min. The LC-
FT-ICR-MS chromatogram were segmented into 1min bins, resulting in
12 segments per sample (nSamples = 40). Each segment was treated as an
individual spectrum and internally calibrated (DataAnalysis 5.0, Bruker
Daltonics) with masses commonly found in DOM (m/z 150–1000, n
calibrants > 186). Assuming singly charged ions (based on12C–13 C1 m/
z spacing), molecular formulas (MFs) were assigned to each m/z value
with a signal/noise ratio > 4 in the mass range m/z 150–1000 using an in-
house software considering the following elements 12C0–60, 13C0–1,
1H0–122, 16O0–40, 14N0–2, 32S0–1, 34S0–1 and a maximum assign-
ment error of <abs(0.5) ppm. Only formulas with 0.3 ≤H/C ≤2.5, 0 ≤O/
C≤1, 0 ≤N/C ≤0.5, 0 ≤DBE ≤20 (double bound equivalent, DBE = 1
+0.5 *(2 c - h +n) and −10 ≤DBE-O ≤+10 were considered for
further data evaluation78,79. Isotopologue formulas (13C, 34S) were used
for quality control but removed from the final data set as they represent
duplicate chemical information. All MF found in blanks (ultrapure water
measured with LC-FT-ICR-MS, n Blanks = 4) were removed from the
corresponding segments in the samples.
Based on the mass peak magnitudes of all assigned peaks, intensity-
weighted mean (wa) molecular descriptors were calculated for aromaticity
index (AI = [1 +c-o-s−0.5 *(h +n)] / [c - o - s –n])80, nomina l oxidation
state of carbon (NOSC = 4 –[4c +h−3n −2o +5p −2s]/c)andm/z
according to waX = ∑(Xi *Intensity) / ∑(Intensity), where X is a molecular
descriptor, and i is the i-th assigned peak in the spectrum.
For Fig. 3A, B, all segments of a respective biological replicate were
averaged into a single spectrum considering MF which occurred at least in
two segments, with average peak magnitudes calculated as simple means,
and weighted average molecular descriptors calculated as above. For
Fig. 3D-G, key features were extracted from the data set as follows: First,
each segment of the biological replicates was averaged using only MF
occurring in all replicates, and which have a peak magnitude variability
(calculated as median +−IQR from 2-3 replicates) < 20%, resulting in
12 segments per sample and 12970 unique MF. Second, MF were extracted
from each segment which show a peak magnitude variability (calculated as
coefficient of variance from 40 samples) > 25%, resulting in 640 key features
(MFxSegment, 14≤n≤116 key features per segment and 563 unique MF)
for the entire data set. Key features thus represent MF, which show the
largest differences within the data set.
Data analysis
Mean and standard errors were calculated for all data sets. Interactions
between climatic conditions and Cd were evaluated using a two-factorial
ANOVA at a 95% confidence interval, specifically for the greenhouse gas
emissions data. Differences between individual treatments were assessed
with a Student’st-testata95%confidence interval. To analyze porewater Cd
mobility, a generalized mixed linear model was employed, with climatic
conditions, time, and soil Cd levels treated as fixed factors. To address which
factors influenced porewater Cd mobility a principle component analysis
was performed on the geochemical porewaterdata.Forthemicrobialdis-
similarity, a principle coordinates analysis was performed based on the
Bray-Curtis dissimilarity of the 16S rRNA gene and transcript amplicon
sequencing. LC-FT-ICR-MS key features were clustered based on Euclidian
distance and complete clustering using the heatmap function in R
(version 4.3.0).
Reporting summary
Further information on experimental design is available in the Nature
Research Reporting Summary linked to this paper.
Data availability
The data that support the findings of this study are included in a
compressed Source Data file at figshare (https://doi.org/10.6084/m9.
figshare.27165732.v1), FT-ICR-MS data is available via the UFZ data
depository (10.48758/ufz.14199) and raw sequencing data were
deposited at the Sequence ReadsArchive(SRA,BioProject:
PRJNA1022319, (https://www.ncbi.nlm.nih.gov/bioproject/PRJNA-
1022319).
Received: 14 May 2024; Accepted: 16 October 2024;
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Acknowledgements
We thank A. Gloeckle, C. Glotzbach, C. Leven, S. Cafisso, D. Buchner, and
the Geomicrobiology group members for help in the laboratory. We also
thank F. Schaedler, R. Kallies, and S. Schreiber for their supportand advice
on sequencing and data evaluation and E. Baeurle, E. Wizemann and J.
Bodemer for providing the soil. For FT-ICR-MS measurement, we thank J.
Kaesler. We appreciate using the analytical facilities of the Centre for Che-
mical Microscopy (ProVIS) at the Helmholtz Centre for Environmental
Research, Leipzig, which is supported by the European Regional Develop-
ment Funds (EFRE—Europe funds Saxony) and the Helmholtz Association.
This work was financed by the Baden Wuerttemberg Stiftung’sExcellence
Programme for Postdocs and the Helmholtz Young Investigator Grant
RhizoThreats. We further acknowledge infrastructural support by the
Deutsche Forschungsgemeinschaft (DFG, German Research Foundation)
under Germany’s Excellence Strategy, cluster of Excellence EXC2124,
project ID 390838134. We thank M. Latimer and R. Davis for beamline
support at SSRL (proposal number 5587). Use of the Stanford Synchrotron
Radiation Lightsource, SLAC National Accelerator Laboratory, is supported
by the U.S. Department of Energy, Office of Science, Office of Basic Energy
Sciences under Contract No. DE-AC02-76SF00515. The SSRL Structural
MolecularBiology Program is supported by the DOE Office of Biological and
Environmental Research and by the National Institutes of Health, National
Institute of General Medical Sciences (including P41GM103393). The con-
tents of thispublication are solely the responsibility ofthe authors and do not
necessarily represent the official views of NIGMS or NIH.
Author contributions
Funding for this work was acquired by E.M.M.; This work was conceptuali-
zed by E.M.M.; Laboratorywork was plannedby S.D. with primary inputfrom
E.M.M. and advised by A.K. and S.F.; Laboratory work was carried out by
S.D. and E.B.; Synchrotron work was carried out by S.D., E.M.M., and J.L.P.
and analyzed by J.L.P.; Cd quantification was carried out by B.P.F., J.M.L.,
and S.D.;FT-ICR-MS data wereprocessed by O.L.and S.D.; The manuscript
was written by S.D. with primary input from E.M.M. and overall input and
discussion from all co-authors.
Funding
Open Access funding enabled and organized by Projekt DEAL.
Competing interests
The authors declare no competing interest.
Additional information
Supplementary information The online version contains
supplementary material available at
https://doi.org/10.1038/s43247-024-01794-w.
Correspondence and requests for materials should be addressed to
E. Marie Muehe.
Peer review information Communications Earth & Environment thanks
Chaolei Yuan and Mallavarapu Megharaj for their contribution to the peer
review of this work. Primary Handling Editors: Alice Drinkwater and
Somaparna Ghosh A peer review file is available.
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