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ARTICLE
Organic coating on biochar explains its nutrient
retention and stimulation of soil fertility
Nikolas Hagemann 1,17, Stephen Joseph2,3,4, Hans-Peter Schmidt5, Claudia I. Kammann6, Johannes Harter1,
Thomas Borch 7, Robert B. Young 7, Krisztina Varga 8, Sarasadat Taherymoosavi2, K. Wade Elliott8,
Amy McKenna9, Mihaela Albu10, Claudia Mayrhofer10, Martin Obst11, Pellegrino Conte 12,
Alba Dieguez-Alonso13, Silvia Orsetti14, Edisson Subdiaga14, Sebastian Behrens15,16 & Andreas Kappler1
Amending soil with biochar (pyrolized biomass) is suggested as a globally applicable
approach to address climate change and soil degradation by carbon sequestration, reducing
soil-borne greenhouse-gas emissions and increasing soil nutrient retention. Biochar was
shown to promote plant growth, especially when combined with nutrient-rich organic matter,
e.g., co-composted biochar. Plant growth promotion was explained by slow release of
nutrients, although a mechanistic understanding of nutrient storage in biochar is missing.
Here we identify a complex, nutrient-rich organic coating on co-composted biochar that
covers the outer and inner (pore) surfaces of biochar particles using high-resolution spectro
(micro)scopy and mass spectrometry. Fast field cycling nuclear magnetic resonance, elec-
trochemical analysis and gas adsorption demonstrated that this coating adds hydrophilicity,
redox-active moieties, and additional mesoporosity, which strengthens biochar-
water interactions and thus enhances nutrient retention. This implies that the functioning
of biochar in soil is determined by the formation of an organic coating, rather than biochar
surface oxidation, as previously suggested.
DOI: 10.1038/s41467-017-01123-0 OPEN
1Geomicrobiology, Center for Applied Geosciences, University of Tuebingen, Sigwartstrasse 10, Tuebingen 72076, Germany. 2School of Environmental and
Life Sciences, Chemistry, University of Newcastle, Callaghan NSW 2308, Australia. 3School of Materials Science and Engineering, University of New South
Wales, Kensington NSW 2052, Australia. 4Nanjing Agricultural University, Nanjing 210095, China. 5Ithaka Institute for Carbon Strategies, Ancienne Eglise
9, Arbaz 1974, Switzerland. 6Department of Soil Science and Plant Nutrition, WG Climate Change Research for Special Crops, Hochschule Geisenheim
University, von-Lade Str. 1, Geisenheim 65366, Germany. 7Department of Soil and Crop Sciences and Department of Chemistry, Colorado State University,
Fort Collins, CO 80523, USA. 8Department of Molecular, Cellular, and Biomedical Sciences, University of New Hampshire, Durham, NH 03824, USA.
9National High Magnetic Field Laboratory, Florida State University, 1800 East Paul Dirac Drive, Tallahassee, FL 32310-4005, USA. 10 Austrian Cooperative
Research, Centre for Electron Microscopy and Nanoanalysis, Steyrergasse 17, Graz 8010, Austria. 11 BayCEER Analytics, University of Bayreuth, Bayreuth
95440, Germany. 12 Dipartimento di Scienze Agrarie e Forestali, Università degli Studi di Palermo, v.le delle Scienze ed. 4, Palermo 90128, Italy. 13 Institute of
Energy Engineering, Chair for Energy Process Engineering and Conversion Technologies for Renewable Energies, Technische Universität Berlin, Fasanenstraße
89, Berlin 10623, Germany. 14 Environmental Mineralogy and Chemistry, Center for Applied Geoscience, University of Tuebingen, Sigwartstrasse 10,
Tuebingen 72076, Germany. 15 Department for Civil, Environmental, and Geo-Engineering, University of Minnesota, 500 Pillsbury Drive S.E, Minneapolis, MN
55455-0116, USA. 16 BioTechonology Institute, 140 Gortner Labs, 1479 Gortner Avenue, St. Paul, MN 55108-6106, USA.
17
Present address: Environmental
Analytics, Agroscope, Reckenholzstraße 191, 8046 Zurich, Switzerland. Correspondence and requests for materials should be addressed to
A.K. (email: andreas.kappler@uni-tuebingen.de)
NATURE COMMUNIC ATIONS |8: 1089 |DOI: 10.1038/s4146 7-017-01123-0 |www.nature.com/naturecommunications 1
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Biochar is the product of O
2
-limited thermal treatment of
biomass (pyrolysis) and is used in agriculture as a livestock
feed supplement, compost additive and soil amendment as
well as for manure treatment1. It is applied to improve animal,
plant and soil health, to reduce nutrient losses by volatilization or
leaching, to prevent soil erosion, and to improve soil water
retention, soil carbon content and the long-term fertility of
agricultural soils2,3. Biochar is very recalcitrant and can con-
tribute to climate change mitigation by the sequestration of stable
carbon4and reduction of agricultural emissions of CO
2
,N
2
O and
CH
4
5.
Positive impacts of biochar on (agro)ecosystems are often
explained by the porosity and sorption capacity3, redox
properties6,7, and liming capacity of biochar8and by biochar's
influence on soil structure9,10, water holding capacity11 and
nutrient transformations in soil2. However, despite these insights
into biochar functioning in soil, a detailed mechanistic under-
standing of how biochar influences plant growth is still lacking.
Biochar was shown to increase crop yield by up to 400% com-
pared to fully fertilized controls12, while the overall mean of
biochar-induced yield increases is 18% according to meta-
analyses13. However, many studies used high biochar application
amounts of >10 t ha−1, which is not economically feasible. Recent
research suggested that biochar should be combined with organic
amendments to increase soil fertility even when biochar is applied
at low (0.5–2t ha
−1) biochar application rates14,15. Co-com-
posting, which consists of mixing biochar with manure or other
compost feedstock with high contents of both nutrients and labile
organic carbon before starting an aerobic composting process,
was shown to enhance the agronomic performance of biochar as a
soil amendment16. The co-composted biochar handpicked from
the final biochar-amended compost was shown to promote plant
growth beyond the combination of pristine biochar with either
mineral fertilizer or mature non-biochar-amended compost. This
phenomenon was explained by co-composted biochar’s slow
release of essential plant nutrients like nitrate and phosphate16.
However, despite evidence for surface oxidation, sorption of
organic molecules and increase in carboxyl groups on biochar
surfaces17,18, the underlying mechanisms and the impact of co-
composting on the biochar microstructure remain unclear. A
mechanistic understanding of how co-composting impacts bio-
char’s reactivity and function is a prerequisite for large-scale and
global beneficial use of biochar in agriculture, additionally for
understanding the formation of extremely fertile Amazonian and
African Dark Earths that received anthropogenic input of both
nutrient-rich organic matter and pyrogenic carbon centuries
ago19,20, and eventually for the development of low-cost biochar-
based fertilizers that can promote high crop yields with com-
parably small application doses of biochar21,22. In addition,
mechanistic insights into biochar co-composting will help to
improve our current understanding of how biochar ages in soil23.
On the surface of co-composted biochar, we identified an
organic coating by applying a suite of spectro(micro)scopical
techniques and argue that this coating controls the interaction of
co-composted biochar with water and nutrients. We found that a
similar coating was present in non-composted but soil-aged
biochar. This study demonstrates why co-composted or organi-
cally amended biochar helps optimize its agronomic performance.
Results
Extraordinary nutrient retention by co-composted biochar.We
manually picked co-composted biochar particles (BC
comp
) after
60 days of aerobic composting plus 6 months of storage and
analyzed them without any further sample preparation24.BC
comp
contained 2.0 ±0.1 g NO
3
−-N kg−1, of which only 43% could be
extracted by conventional extraction (Fig. 1), clearly demon-
strating the potential of co-composted biochar as a slow release
fertilizer. We investigated both pristine (fresh, no post-
production treatment, no detectable nitrate) biochar (BC
prist
)
and BC
comp
with a suite of spectro(micro)scopic and mass-
spectrometric techniques to identify the underlying molecular
mechanism(s) for the remarkable retention of anionic and
cationic nutrients by co-composted biochar16 and to evaluate the
impact of co-composting on the long-term stability of biochar,
i.e., its carbon sequestration potential.
Spectromicroscopy and mass spectrometry. First, we used13C
solid-state nuclear magnetic resonance (NMR) spectroscopy, X-
ray photoelectron spectroscopy (XPS) and scanning electron
microscopy (SEM) to identify co-composting-induced changes in
biochar carbon speciation on the bulk level and structural and
chemical changes of the biochar surface. NMR spectroscopy did
not reveal significant differences in C speciation between BC
prist
and BC
comp
on the bulk level (Supplementary Fig. 1). In line with
this, bulk XPS, i.e., XPS of biochar particles powdered and sieved
to 100 µm, only showed a minor increase in oxidized carbon
species (C–O, C =O, COO, Fig. 2a-b). However, XPS applied on
whole particles revealed a considerable increase in the O/C ratio
and increase in N content by co-composting (Fig. 2a-b). Since
XPS is a surface sensitive technique that provides information on
the outermost ~10 nm this also suggests that only the biochar
surface, not the bulk biochar particles, is altered during co-
composting. SEM revealed that this surface alteration of BC
comp
is
not homogeneous, but that there were regions of degraded bio-
char pores (Fig. 2c) and hotspots of partially and completely
coated biochar surfaces (Fig. 2d, e). Elemental mapping using
energy dispersive X-ray spectroscopy (EDS) indicated hetero-
geneous contents of Fe, Al and Si in the coating (Fig. 2f).
We characterized the coating in situ with mass spectrometry
and spectroscopic techniques across a range of both spatial and
energy resolutions to identify the chemical and physical proper-
ties of the surface modification. Desorption atmospheric pressure
photoionization Fourier-transformed ion-cyclotron resonance
mass spectrometry (DAPPI FT-ICR MS) applied to biochar
particles provides precise information on the elemental
0
1
2
Pristine biochar Co-composted
biochar
1.0 (1 h) 2.0 (1 h) 3.0 (1 h)
4.0 (20 h) 5.0 (48 h) 6.0 (96 h)
NO
3
–
-N (g N kg
–1
)
Fig. 1 Biochar nitrate content based on repeated extractions with 2 M KCl.
Each segment of the stacked bar represents one step of six consecutive
extractions with KCl: the first, second and third extraction took 1 h each, the
fourth took 20 h, the fifth 48 h, and the sixth took 95 h. Error bar represents
1 S.E. of the total extraction carried out in triplicates
ARTICLE NATURE COMMUNICATIONS | DOI: 10.1038/s41467-017-01123-0
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composition of compounds available for desorption on the
biochar surface. According to DAPPI FT-ICR MS, co-composting
increased the relative abundance of CHON (C
W
H
X
O
y
N
z
) class
compounds and decreased the relative abundance of CHO
(C
x
H
y
O
z
) class compounds (Fig. 3). Furthermore, co-
composting increased the abundance-weighted N/C ratio of all
formulas assigned to the compounds that desorbed from the
biochar surface at atmospheric pressure from 0.008 to 0.029, and
increased the abundance-weighted O/C ratio of such formulas
from 0.388 to 0.618. Within the CHON class, the van Krevelen
diagram (Supplementary Fig. 2) suggests an increase in the O/C
ratio of the desorbed compounds, consistent with an actual
increase in the abundance-weighted O/C ratio from 0.278 to
0.566. To accurately identify carbon functional groups and spatial
distribution of N in ultra-thin sections of BC
comp
, we used
scanning transmission X-ray microscopy (STXM) which has a
spot size of 30–50 nm. We could clearly demonstrate the
heterogeneous distribution of nitrogen suggesting that there are
hotspots of nutrient retention on the outer surface and inside
pores of the biochar, rather than a homogenous enrichment and
these hotspots showed a higher absorption by carboxylic moieties
(Fig. 4). However, beyond this change, there are neither
considerable differences in carbon speciation between bulk co-
composted biochar and these hotspots nor between pristine and
co-composted biochar (Supplementary Fig. 3).
Scanning transmission electron microscopy (STEM) equipped
with electron energy loss spectroscopy (EELS) allows sub-micron
scale resolutions on ultra-thin sections of the biochars and
information on the carbon and nitrogen speciation in precisely
(nm-range) defined regions. With STEM, we identified a meso-
porous organic coating of up to 200 nm thickness on a substantial
share of the investigated surfaces of BC
comp
(Fig. 5and
Supplementary Fig. 4). STEM samples were sputter-coated with
gold on a three-dimensional (3D) moving sample holder prior to
Al Si Fe
0%
20%
40%
60%
80%
100%
Pristine
particle
Composted
particle
Pristine
powder
Composted
powder
C O N Al Si Ca F
0%
20%
40%
60%
80%
100%
Pristine
particle
Composted
particle
Pristine
powder
Composted
powder
C–C, C=C, C–H C–O C=O COOMg
ab
cd
ef
Fig. 2 Identification of biochar surface modifications induced by co-composting. aElemental composition of the surface (particle) and the bulk (powder)
biochar according to X-ray photoelectron spectroscopy (XPS). bCarbon-containing functional groups according to region scans (higher energy resolution)
of the same samples. c–fScanning electron micrographs of co-composted biochar with ca region with degraded, but still exposed original biochar surfaces
showing typical porous structure (scale bar, 20 µm); da region of biochar surface partially covered with organic coating (scale bar, 10 µm); ea region of
biochar surface completely covered with organic coating (scale bar, 600 µm). fmap overlay of eshowing the heterogeneous distribution of Al (green), Si
(blue) and Fe (red) within the organic coating as quantified by energy-dispersive X-ray spectroscopy
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sample preparation to clearly differentiate the sample-epoxy resin
border. The organic coating was thicker on pore surfaces than on
exposed, outer surfaces, where it was mostly limited to 10–20 nm
non-porous spots (Supplementary Fig. 4D). The coating was
completely absent in BC
prist
(Supplementary Fig. 5). EELS
revealed a higher N content of the coating compared to the
carbonaceous biochar matrix and minor alterations of the C
speciation (Fig. 5b–e) including peak shifts of the aromatic C =C
1s–π* transition by 0.5 eV from co-composted biochar matrix to
the coating (Fig. 5c, d). Additionally, we found nanometer-sized
hot-spots of silica and other elements in both pristine and co-
composted biochar (Supplementary Fig. 5B). Similar hotspots of
calcium were identified on the surface of the co-composted
biochar, right under the coating (Fig. 5f, g). A similar
phenomenon was described for pyrogenic organic matter isolated
from Terra Preta25.
Additional STEM on crushed BC
comp
revealed the formation of
complex organo-mineral associations involving Ca and redox-
active Fe (Supplementary Fig. 6). Similar structures have been
suggested as hotspots for redox reactions in the liquid and
gaseous phase of soils and suggest that the organic coating is not
only a porous matrix for nutrient retention, but also a potential
location for nutrient transformations26,27.
To obtain more detailed information on the identity and
properties of this organic coating, the material was characterized
ex situ by first pre-washing the BC
prist
and BC
comp
with water to
remove loosely attached compost organic matter followed by
extraction of the biochar particles with 0.05 M NaOH28 to isolate
the organic coating for further analysis after filtration to <0.45
µm. After this procedure, the co-composted biochar lost its
brownish color and is almost completely black again (Supple-
mentary Fig. 7). Ion chromatography and elemental analysis of
the extracted coating revealed an increased content of N and C,
especially nitrate, organic carbon, carbonate, Ca and K in BC
comp
compared to extracts obtained from BC
prist
(Supplementary
Table 4). The1H solution NMR spectrum of the BC
prist
extract
comprised only few defined peaks, while the spectrum of BC
comp
extracts was dominated by broad peaks that did not allow detailed
evaluation (Supplementary Fig. 10). The complexity of the
spectrum showed that the formation of the organic coating in
the co-composted biochar cannot solely be explained by
oxidation of the original biochar carbon but entirely new
species/entities of carbon were evidently introduced into the
biochar porous system during co-composting, which is supported
by data obtained by spectrofluorometry (Supplementary Fig. 11).
This compost derived organic carbon have had to be sorbed on
the biochar because if it just had been weakly associated with the
biochar, it would have been removed already by the first washing
step (10 min with water), while analysis was performed on the
eluate of the third washing step. In line with that, Fourier-
transformed infrared (FTIR) spectroscopy of freeze-dried BC
comp
extracts revealed similar bands as found in both a powdered
BC
prist
and powdered compost reference (Fig. 6a). This suggests
that biochar nano-particles were associated with compost organic
matter to form the porous coating. Biochar nano-particles can
form during co-composting by physical disintegration, potentially
by graphitic sheet expansion, when biochar sorbs water from the
compost29. Quantification of the electron exchange capacity
(EEC), i.e., the sum of electron accepting and donating capacity6,
revealed that the EEC per mmol of carbon of BC
comp
eluates is
higher than the EEC both of BC
prist
and of compost eluates
(Fig. 6b). Thus, the chemistry of the coating is different from both
of these compounds. Our data suggests that the reaction of
compost organic matter with biochar created more reactive
carbon moieties that then formed the organic coating. Although,
looking at EDC/EEC ratios, eluate of co-composted biochar
presents similar ratio than compost, indicating that the co-
0
20
60
40
(%) relative abundance
CHN
Pristine
Co-composted
CHO CHON CHOS HC
Fig. 3 In situ characterization of the organic coating by DAPPI FT-ICR MS.
Relative abundance of C, H, O, N and S bearing compound classes on the
surface of pristine and co-composted biochar according to desorption
atmospheric pressure photoionization Fourier-transformed ion-cyclotron
resonance mass spectrometry (DAPPI FT-ICR MS)
3
2
Absorption (arbitrary units)
1
0
280 290 300 310
(eV)
ab
c
Fig. 4 Scanning transmission X-ray microscopy of co-composted biochar. a
Average image on linear absorbance scale (optical density) calculated from
a stack across the C1s absorption edge (278–330 eV) of an ultra-thin
section of co-composted biochar. Scale bar, 500 nm. bC1s stack with
overlay of regions of interest for downstream data evaluation: green—
regions on or directly adjacent to the two biochar particles with relatively
high N/C ratio (based on X-ray absorption ratio), red—manually defined
region of the particle center, blue—all regions in the analyzed area with high
N/C ratio, includes the green marked region. Same scale as a.cX-ray
absorption spectra extracted from the regions defined in bwith the
respective colors, the blue spectrum again includes the green region.
Spectra reveal a relative increase in absorption at the 1s-pi* transition
(288.6 eV; R-COOH; CO62) from particle center (red) to particle
associated high N/C (green) to all areas with high N/C (blue). In-depth
analysis of this and two other regions of interest is displayed in
Supplementary Fig. 3B, C. Comparison of with pristine biochar is displayed
in Supplementary Fig. 3A
ARTICLE NATURE COMMUNICATIONS | DOI: 10.1038/s41467-017-01123-0
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composted biochar presents the same redox state than the
compost. From an absolute point of view, pristine biochar shows
lower EEC which is increased by co-composting which could be
attributed to conformational re-arrangements of carbon moieties
taking place during the sorption compost organic matter onto
pristine biochar, making the redox-active moieties more exposed
to the electrochemical techniques. Alternatively, biochar might
have preferentially sorbed compounds with a higher EEC than the
average EEC of the compost. However, there was no clear
evidence of fractionation of the organic matter as consequence of
the sorption, as suggested by fluorescence emission spectroscopy
(Supplementary Fig. 11).
Porosity and interaction with water. To identify the mechanism
responsible for the retention of polar nutrients such as nitrate by
the observed porous organic coating of biochar surfaces, we used
fast field cycling NMR (FFC NMR) relaxometry quantifying the
strength of biochar interactions with water30. We found that co-
composting changed the distribution of relaxation times (Fig. 7
1
43
2
5
5
15
20
10
Energy loss (eV)
CO
134
6
7
Energy loss (eV)
300
(1)
(1) (2) (3) (4)
(5)
Biochar
Energy loss (eV)
Energy loss (eV)
50
40
30
20
10
10
20
400 405 410
300 320 340
0
20
40
60
80
100
120
100 nm50 nm
200 nm
30
40
50
60
70
0
285 290 295
1
2
3
4
2
(2)
N
Electrons (104)
Electrons (104)
350 400 450 500 550
Organic coating
Biochar
Organic coating
Counts (103)
Electrons (103)
ac f
db
e
g
Fig. 5 Scanning transmission electron microscopy and electron energy loss spectroscopy of organic coating on co-composted biochar. Micrographs were
obtained from different ultra-thin sections of the same biochar particle. a,bSTEM HAADF micrograph and EELS spectra of an ultra-thin section of co-
composted biochar revealing an organic coating. Position 1: biochar with N and O below detection limit of EELS; 2: epoxy resin; 3: empty hole (resin did not
penetrate this pore); 4: gold that was sputtered onto the biochar particle before embedding to identify the sample surface; 5: organic coating, porous
appearance, contains N and O according to EELS spectra shown in b; 6 and 7: Location of EELS sum spectra shown in b.bEELS spectra revealing only minor
differences in C speciation between biochar (red) and organic coating (blue), but considerably increased content of N in the coating. Letters indicate
absorption edge (K shell) of C, N and O. cCloseup on a region of thick (~120 nm) organic coating, probably located inside a biochar pore (no gold coating
detectable). dC-K near edge EELS spectrum of biochar matrix and coating as marked in cshowing a peak shift of the C =C1s–π* transition (1) of aromatic
carbon by ~0.5 eV (284.9–285.5 eV). C–C 2s-2pz σ* transition (2) was not altered (291.2 eV). eN-K near edge EELS spectrum of coating as marked in c
showing (1) imine N 1s–π* transition at 399–400 eV; (2) amide N 1s–3p/π* transition at 401.3 eV; (3) nitro N 1s–π* transition at 403.6 eV; (4) corresponds
to HC ≡N* transition at 406.8 eV with σ* resonance position at 420.8 eV63.fOrganic coating formed at a biochar surface that is rich in Ca hotspots of
bright appearance in the HAADF micrograph. Coating is shearing off as an artefact of the mechanical force applied during sample preparation, most likely
by the microtome, which indicates a rather plastic nature of the coating compared to the biochar that broke (lower part of the micrograph). Thickness of
coating varies between ~20 and 50 nm and the presence of gold indicates a semi-exposed position in the original biochar particle. gEELS spectra from
regions indicated in fshowing the peak shift of the C =C1s–π* transition and the presence of Ca in the region of the bright spot in the outermost region of
the biochar
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and Supplementary Fig. 12), i.e., time needed for protons to
recover the longitudinal component after magnetization31. The
different relaxation times indicate the strengths of interactions of
water molecules with biochar pore surface as well as with pore
water-dissolved or suspended compounds30,32. Lower relaxation
times in BC
comp
suggest the presence of non-homogeneously
distributed H-bond donor/acceptor groups. Thus, water is better
anchored to the surfaces of BC
comp
and thus less mobile, which
can explain the retention of water-dissolved nutrients, such as
nitrate. Liquid chromatography organic carbon detection (LC-
OCD, Supplementary Tables 5–7, Supplementary Fig. 17 and
Supplementary Discussion) of aqueous biochar extracts revealed a
high content of humic-substance-like organic molecules, that
might act as surfactants that facilitate this process. In addition,
the organic coating can shrink pore sizes thereby reducing water
mobility31. This is supported by gas adsorption measurements
that showed that co-composting led to ~18% reduction in specific
surface area (SSA) according to CO
2
adsorption, especially in the
pore size range of 0.65–0.85 nm, and ~64% SSA reduction to N
2
adsorption, especially in the pore size range of 1.25–1.45 nm
(Fig. 8, Supplementary Fig. 13 and Supplementary Discussion).
Reference materials:
Freeze dried washing solution
Co-composted
biochar eluate
Pristine biochar
eluate
Compost
Pristine biochar
powder
1,400
NaOH
cm
–1
0
20
20
40
40
60
EAC
mmol e
–
(mol C)
–1
Compost Pristine
biochar
Co-composted
biochar
Absorption (arbitrary units)
EDC
mmol e
–
(mol C)
–1
900 400
a
b
Fig. 6 Ex situ analysis of the organic coating extracted from the biochar
with 0.05 M NaOH. aFourier-transformed infrared (FTIR) spectra of
freeze-dried eluates of pristine and co-composted biochar. FTIR spectra of
pure compost, pristine biochar and NaOH were measured as a reference. b
Electron accepting (blue) and electron donating (red) capacity (EAC and
EDC) of biochar eluates normalized to their carbon content. Error bars
represent one standard deviation of at least five replicates
100
80
60
Relaxation time distribution (a.u.)
40
20
0
1
1.5
12
25
55
85
Pristine biochar
Co-composted biochar
10 100 1,000
Longitudinal relaxation time (T1, ms)
Fig. 7 Analysis of biochar surface hydrophilicity by fast field cycling nuclear
magnetic resonance relaxometry: Relaxograms, i.e., distribution of
relaxation times, of pristine and co-composted biochar. The shorter the T
1
value, the better water molecules are anchored to the biochar surface.
Conversely, the longer the longitudinal relaxation time, the weaker are the
interactions between water and the surface of the biochar system
0
100
200
300
400
500
600
BET
DFT total
DFT < 2 nm
DFT > 2nm
BJH > 3 nm
Pristine
Co-comp
Pristine washed
Co-comp washed
SSA (m2 g–1)
DFT CO2 total
N2 adsorption
CO2 adsorption
Fig. 8 Specific surface area (SSA) according to N
2
and CO
2
gas adsorption.
Total SSA according to the Brunauer, Emmett and Teller method, total SSA
according to the quenched-solid density functional theory (QSDFT)
method, SSA due to micropores according to the QSDFT method, SSA for
mesopores up to 34 nm according to the QSDFT method, SSA for pores
bigger than 3 nm according to the Barrett, Joyner and Halenda method and
SSA of micropores with CO
2
adsorption and applying the non local density
function theory method of pristine and co-composted biochar, pristine
biochar after washing and co-composted biochar after washing
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Thus, among micropores, the larger, more accessible pores were
preferentially clogged. Pores >3 nm were not affected; however,
pore clogging might be underestimated due to new mesopores of
the organic coating increasing total porosity. In essence, CO
2
gas
adsorption reveals that the reduction of SSA by composting is less
severe than reported previously33.
Collectively, our data showed that co-composting results in the
formation of a heterogeneous, meso-porous and hydrophilic
organic coating on biochar surfaces including inner pore surfaces.
The coating was identified as a composite of the original compost
organic matter and biochar nano-particles and it is enriched in N
(especially NO
3
−), K, Ca, Si, Fe and Al. It has a higher N/C ratio
than the biochar matrix, a higher redox activity, and adds another
porous component to the porous biochar. However, bulk carbon
speciation of the biochar were not altered by co-composting. This
suggests that co-composting of biochar does not affect its long-
term stability in soil, however, this needs further evidence from
incubation studies.
Our data provide, for the first time, mechanistic evidence on a
molecular scale as to why the combination of biochar and organic
amendments yields optimal agronomic benefits by biochar
amendment. The enhanced interaction of co-composted biochar
with water (hydrophilicity of organic coating decreases biochar’s
hydrophobicity) is most likely responsible for its improved
nutrient retention and therefore its superior agronomic perfor-
mance16. Co-composting of biochar with nutrient-rich feedstock
or other ways of treating nutrient-rich waste streams with biochar
(e.g., liquid manure) is therefore key to increase biochars’
agronomic value.
Organic coating on soil-aged biochars. We additionally found
that the organic coating is not restricted to the biochar example
presented in this study. STEM also revealed an organic coating in
another co-composted wood biochar (Supplementary Fig. 14) and
in a soil-aged biochar (BC
soil
, Supplementary Fig. 15). Basic
biochar properties of BC
soil
were similar to those of BC
prist
and
BC
comp
(similar feedstock, same process and pyrolysis condi-
tions), but it was produced in 2012 and amended to a field
experiment after soaking in a fertilizer solution. The organic
coating on BC
soil
has slightly different properties (lower electron
accepting capacity, no evidence for incorporation of biochar
nano-particles, different interaction with water, Supplementary
Fig. 16) compared to BC
comp
. Nevertheless, the evidence of
organic coatings on both the composted and the soil-aged biochar
strongly suggests that the organic coating of natural and
anthropogenic (biochar) pyrogenic carbon is likely a ubiquitous
phenomenon and corresponds to the finding that biochar stabi-
lizes rhizodeposits34. This could explain the previously reported
nitrate-retention capacity of biochar particles aged for 2.5 years in
afield experiment35. Hence it is reasonable to assume that similar
coatings may also have formed both on anthropogenic pyrogenic
carbon in Amazonian and African Dark Earths soils and as well
on natural pyrogenic carbon in soils, which was recently shown to
account for on average 13.7% of the global soil organic carbon
pool36.
Discussion
Our study revealed that the formation of an organic coating on
biochar, that was observed on all biochar particles investigated in
this study, is the dominant molecular mechanism of biochar
alteration by co-composting and soil ageing. This mechanism
replaces the paradigm of surface oxidation of native biochar
carbon as the dominant process of ageing. It might partially
explain biochar’s longevity in soil4, as the organic coating might
protect aromatic biochar surfaces to some extent from further
(oxidative) degradation. Surface oxidation might play a certain
role, as biochar particles are not completely covered by the
organic coating (Fig. 2c, d). Moreover, functionalization of bio-
char surfaces by oxidation might facilitate the formation of the
organic coating, as oxidized carbon moieties facilitate the inter-
action of biochar with organic molecules by hydrogen bonds18.
However, our study clearly shows that increased abundance of
oxidized carbon moieties like carboxylic groups or increased O/C
ratios of biochar should not be simply assigned to surface oxi-
dation, when the data is obtained from bulk analysis.
Future research needs look at the mechanisms of the formation
of the organic coating. Although we presented evidence through
LC-OCD that the organic coating consists of humic-like sub-
stances, microbial activity could play an important role in the
formation of the organic coating. It needs to be clarified if the
increase in EEC is the result a chemical alteration of organic
matter or the result of preferential sorption of organic matter with
a high EEC.
We conclude that the combination of biochar with non-
pyrogenic organic matter (e.g., manure) is the key strategy to
develop carbon-fertilizer carriers that are effective at low appli-
cation rates21,22. Annual application of such biochar-based fer-
tilizers could sequester carbon and mitigate global warming5—a
global mitigation strategy that could be implemented due to its
economic value for farmers, without the necessity for further
subsidies.
Methods
Materials. Biochar was produced from mixed woody waste materials via slow
pyrolysis (700 °C) by Swiss Biochar (Belmont-sur-Lausanne, VD, Switzerland) with
a Pyreg reactor37 and was characterized by Eurofins Umwelt Ost GmbH, Hals-
brücke-Tuttendorf, Germany, as requested by the European Biochar Certificate1.
Results are displayed in Supplementary Tables 1–3. This biochar was part of the
COST Action-TD1107 biochar ring trial38 and both pristine/non-co-composted
and co-composted aliquots from the same composting experiment were subject to
previous studies39.
Biochar was co-composted at the Ithaka Institute at St. Léonard, VS,
Switzerland, from August to October 2014. Biochar was mixed into compost
feedstock composed of cow, horse and poultry manure and green waste at a rate of
4.3% (dry matter w/w). Compost was managed aerobically as described by
Kammann et al.24 resulting in temperatures above 60 °C for more than 2 weeks.
Compost feedstock was composted without biochar amendment as a control. After
frost-protected storage during winter, biochar was picked manually from the
mature compost in spring 2015. For biochar picking, 50 kg of biochar-amended
compost were stored for analysis in a plastic bag. For each analytical procedure
(extraction, characterization), a subsample of compost was collected as a composite
sample from this bag. 10–30 char particles were separated from this individual
subsample and then prepared for analysis. The actual number of biochar particles
finally involved in analysis varied strongly between the different techniques. E.g.
only 3 particles of each treatment could by analyzed by STEM and 2 for STXM,
while extraction or powder-based analysis (gas adsorption) where based on at least
20 particles.
The same type of biochar was already purchased from Swiss Biochar in 2012
(separate analysis in Supplementary Tables 1–3). It was soaked in a diluted
commercial fertilizer solution overnight (1:1 w/v, 1.25 g N kg−1biochar) and
incorporated at a rate of 60 Mg ha−1into the upper 15 cm of a Terric Anthrosol
(top- and subsoil of a Cambisol mixed by construction activities) at the Tuebingen-
Sand field site (lat. 48.5342, long. 9.0711). The plot was cropped with Emmer wheat
(Triticum dicoccon) in 2012 and winter vetch (Vicia villosa) in 2013 and green
fallow thereafter. Biochar was manually picked from soil samples obtained from the
upper 15 cm of three sampling sites in spring 2015.
Serial extraction of nitrate. Nitrate was repeatedly extracted from both pristine
and co-composted biochar 1:10 (w/v) with 2 M KCl in 50 ml Falcon tubes (n=3)
on a roller shaker (ROLLER 10 digital, IKA, Staufen, Germany) at 50 r.p.m. at
room temperature (22 ±3 °C). After each extraction step, the extractant was dec-
anted through a 0.5 mm sieve. The volume of the decanted extractant was mea-
sured to correct the data for the residual extractant (2 M KCl) that cannot be
removed by decanting (residual =water content of biochar + added extractant—
decanted extractant) which carries a small quantity of already extracted nitrate
from one to the subsequent extraction. Fresh 2 M KCl was added to the biochar at
the same volume. Each sample was extracted six times: three times for 1 h, for 18 h,
for 48 h and for 96 h, respectively, resulting in a total extraction time of 167 ±1h.
Nitrate was quantified using a continuous flow analyzer (SEAL Analytical,
NATURE COMMUNICATIONS | DOI: 10.1038/s41467-017-01123-0 ARTICLE
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Content courtesy of Springer Nature, terms of use apply. Rights reserved
Germany) after reduction of the nitrate to nitrite with hydrazine, which was pre-
pared according to SEAL’s advice for soil extracts. Nitrite was quantified by UV–vis
absorption at 550 nm after reaction with N-1-naphtyl-ethylendiamin. The SEAL
system is equipped with a dialysis membrane that removes any extraneous
microparticles to prevent side reactions or additional absorbance. Standards were
prepared in 2 M KCl to provide the same matrix.
13C solid-state nuclear magnetic resonance spectroscopy. Solid-state NMR
spectra were collected using a Bruker (Billerica, USA) Avance III 600 MHz spec-
trometer equipped with a 4.0 mm magic-angle spinning Efree triple resonance
HCN probe. 22–29 mg of each sample were packed into 50 µl 4.0 mm zirconia
rotors. 1D-13C-spectra were acquired using standard Bruker cross-polarization
(CP) pulse sequence at 10 or 12 kHz sample rotation frequency and at 27.0 °C VT
inlet gas temperature. 24,576 scans were signal averaged for each spectrum with 3 s
recycle delay.13C polarization was achieved using a ramped-amplitude1H-13CCP
pulse for 2.0 ms, and the spectra were acquired for 15 ms under 81kHz two-phase-
modulated (TPPM)1H decoupling.13C chemical shifts were referenced externally to
2,2-dimethyl-2-silapentane-5-sulfonate sodium salt using the adamantane methy-
lene peak at 40.48 ppm40. All data were processed with applying a 30 Hz expo-
nential line broadening using Bruker Topspin 3.5.
Gas adsorption.N
2
and CO
2
adsorption were combined to characterize the SSA
and pore volume (PV) of the biochar samples. To this end, the adsorption and
desorption isotherms for both gases were measured at 77 K and 273 K, respectively.
Before the measurements, the char samples were pretreated to clean the char
surface from other adsorbed species (degassing) by heating them in vacuum at 120
°C during 6 h. The samples were also milled to reduce the influence of transport
limitations on the measurements, especially in the case of N
2
adsorption, due to the
low temperatures. The N
2
isotherms were determined measuring the amount of N
2
adsorbed/desorbed as function of the N
2
relative pressure P/P
0
in the measurement
station, where P is the pressure of N
2
in the measurement station and P
0
is the N
2
saturation pressure at the measurement temperature (~77 K). The relative pressure
P/P
0
ranged from ~0.005 to ~0.995. In the case of CO
2
the relative pressure ranged
from ~0.00005 to ~0.028. For both degassing and adsorption/desorption mea-
surements the gas sorption system Nova 2000 provided by Quantachrome
Instruments (Boynton Beach, FL, USA) was used. Several methods were applied to
the isotherms in order to derive pore surfaces areas and volumes, which are
included in the NovaWin software (Quantachrome Instruments). In the case of N
2
adsorption, the Brunauer, Emmett and Teller (BET)41 method was used to
determine the total internal SSA of the samples. The DFT (density functional
theory)42 method was applied to the adsorption isotherm to characterize the pore
size distribution, in particular, the QSDFT (quenched solid-state functional theory)
method, considering slit/cylindrical pores. With this method, micropores and
mesopores up to ~34 nm in pore width could be measured. The Barrett, Joyner and
Halenda (BJH)43 method was applied to characterize mesopores bigger than ~3 nm
and macropores up to the detectable limit of N
2
adsorption. The total pore volume
was determined from the volume of N
2
adsorbed at a relative pressure (P/P
0
)≈1,
assuming that at this point the pores are filled with liquid N
2
. For CO
2
adsorption,
the nonlinear DFT (NLDFT) was applied to determine the SSA and volume due to
micropores.
X-ray photoelectron spectroscopy. XPS uses X-rays to excite atoms of a sample
and analyzes the energy of the emitted photoelectrons, which provides information
on both elemental composition and chemical bonds of the outermost 10 nm44. XPS
was performed on an ESCALAB250Xi (Thermo Scientific, UK) using mono-
chromated Al K α(1486.68 eV, 150 W) at a spot size of 500 µm under high-vacuum
conditions (<2×10−9mbar) and a photoelectron takeoff angle of 90° was used. It
was calibrated with Au (Au 4f7 =83.96 eV), Ag (Ag 3d5 =368.21 eV) and Cu (Cu
2p3 =932.62 eV). Pass energy was 100 eV for survey scans and 20 eV for region
scans. Biochar particles were gently washed with DI water three times, dried at 40 °
C and were measured both as a particle and as a powder after grinding in an agate
mortar.
Scanning electron microscopy. In SEM, a focused beam of accelerated electrons
scans over the samples and creates a multitude of characteristic secondary emis-
sions. We used secondary electrons, which provide predominantly topographic
information of the sample, and EDS. The energy of the X-rays is characteristic for
the atom that was excited by a primary electron (beam electron) and thus provides
information about the elemental content of the outermost hundreds of nm to ~2
µm of the sample, depending on the composition of the sample and the energy of
the incident electron beam45. SEM micrographs and X-ray spectra were obtained
on a Zeiss Sigma SEM with Bruker EDS at a working distance of 8–9 mm and an
acceleration voltage of 10–15 kV. Individual biochar particles were mounted on
SEM stubs with conductive carbon paint. Sputter coating was not necessary.
Scanning transmission X-ray microscopy. Ultra-thin-sections for STXM were
prepared by cutting pieces of biochar (no pretreatment, no drying) under cryo-
conditions in an ultramicrotome Leica UCT, gluing the sample with a droplet of
2.3 M saccharose and catching the ultra-thin-sections with saccharose, too. They
were placed on formvar-coated 200 mesh copper grids. Grids were carefully rinsed
with water to remove the saccharose and mounted on a STXM sample plate.
Despite the rinsing, saccharose was still present on all samples. Therefore, a
reference spectrum of pure saccharose was used to quantitatively map saccharose
by spectral decomposition in the samples. Ultra-thin-sections were obtained from
both center and the edge of pristine biochar particles as well as the edge of co-
composted biochar particles. Measurements were conducted at the Canadian Light
Source beamline 10ID-1.
Analytical scanning transmission electron microscopy. Pieces of biochar were
covered in gold all-around in the Leica EM ACE 600 (Leica Microsystems, Vienna,
Austria; 45 mA, 8,0 × 10−3mbar, working distance 50 mm) using a rotating device
to enable 3D coating. This resulted in a minimum of 25-nm-thick gold layer on the
biochar. These nuggets were embedded in Spezifix40 (Struers, Willich, Germany)
and after hardening carefully trimmed with a trim 90 blade (Diatome, Switzerland).
The preparation of 50 nm slices was done at room temperature with the ultra-
sonic-knife (Ultra Sonic, Diatome) for reducing compression and allowing best
structure preservation in the ultramicrotome Leica EM UC6 (Leica Microsystems).
The slices were transferred from the water with a perfect loop (Diatome) on a 200
mesh grid. STEM investigations were performed on a monochromated probe
corrected FEI Titan G2 60–300 (STEM) microscope with an X-FEG Schottky field-
emission electron source operated at 60 kV. The microscope is equipped with a FEI
Super-X detector (Chemi-STEM technology), consisting of four separate silicon
drift detectors46 (120 mm acquisition area) and a Dual EELS –Gatan Imaging
Filter (GIF) Quantum47. The microscope was run in monochromated mode with a
spatial resolution of 0.1 nm and an energy resolution at full-width at half-
maximum of 0.17 eV. Two different detectors: high angular annular dark field
(HAADF) and annular dark field (ADF), a beam current of approximate 120 pA
and acquisition times ranging from 20 to 30 μs/pixel were used for micrographs
acquisition. Analytical investigations involved electron energy loss (EELS) and X-
ray (EDX) spectroscopy in STEM mode by line scans and spectrum imaging,
which, however, were acquired with different acquisition times and pixel sizes
depending on the investigated area. Energy loss near edge structure (ELNES) of
carbon-, nitrogen- and oxygen- K ionization edges were analyzed considering the
specificfinger prints and energy shifts of different types of bonding which might be
present in the sample, with graphite as a reference. Elemental quantification has
been performed by using the k-factor method45,48. Sample thickness in different
areas was determined by EELS48. However, for the mean free path of the sample
with the major constituent carbon a calculated value of 76.6 nm (Malis formula48)
was used. The images and spectra were recorded by a Gatan Digiscan unit and
Digital Micrograph software and were corrected for dark current and gain
variations.
Desorption atmospheric pressure photoionization FT-ICR MS. A modified
ThermoFisher LCQ atmospheric pressure photoionization (APPI) source (Ther-
moFisher Corp., Bremen, Germany) was used for all Fourier-transformed ion
cyclotron resonance mass spectrometry (FT-ICR MS) experiments49. Pieces of
pristine and co-composted biochar were held by tweezers ∼1 mm from the exit of
the heated ceramic nebulization tube and ~10 mm from the FT-ICR MS inlet. Gas-
phase neutrals were produced through a combination of thermal and chemical
desorption50, and ionized by dopant-assisted APPI51. Nitrogen was used as
nebulizer gas at 100 psi with toluene as dopant at a flow rate of 50 μl min−1. The
temperature of the heated nebulizer gas/solvent plume ranged from 100 to 500 °C
depending upon the sample. Mass spectra were acquired by a custom-built FT-ICR
MS based on a passively shielded 9.4 T horizontal 200 mm bore diameter super-
conducting solenoid magnet (Oxford Corp., Oxford Mead, UK) operated at room
temperature52. A modular ICR data station (Predator) facilitated instrument
control, data acquisition, and data analysis53. Positive ions were accumulated in an
external quadrupole for 50–500 ms, and passed through an rf-only quadrupole into
an octopole equipped with tilted wire extraction electrodes for improved ion
extraction and transmission54. Helium gas was introduced into the octopole to
collisionally cool the ions before transfer through rf-only quadrupoles (total length
127 cm) into a seven-segment open cylindrical ICR cell with capacitively-coupled
excitation electrodes based on the Tolmachev configuration55. Approximately
25–50 time-domain acquisitions were co-added, Hanning-apodized, and zero-filled
once prior to fast Fourier transform and magnitude calculation. ICR frequencies
were converted to m/z values by the quadrupolar electric trapping potential
approximation56. Spectra were internally calibrated from abundant homologous
alkylation series (compounds that differ in elemental composition by integer
multiples of CH
2
). Mass spectral peaks with signal magnitude greater than six times
the baseline root-mean-square (r.m.s.) noise level were assigned elemental com-
position values (C
0–100
H
0–200
N
0–5
O
0–15
S
0–2
) with PetroOrg software (Y.E. Corilo;
Florida State University; all rights reserved), subject to the following constraints for
chemical feasibility: mass error <1 p.p.m.; the computed ring and double bond
equivalent must be an integer value ≥0; neutral formulas containing an odd
number of nitrogen atoms must have an odd nominal mass, and neutral formulas
containing zero or an even number of nitrogen atoms must have an even nominal
mass (nitrogen rule); 2 ≤H≤(2 C + 2); 0.33 ≤H/C ≤2.25; 0 ≤O≤(C + 2); O/C
<1.2; N/C <0.5; S/C <0.257.
ARTICLE NATURE COMMUNICATIONS | DOI: 10.1038/s41467-017-01123-0
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Content courtesy of Springer Nature, terms of use apply. Rights reserved
DAPPI FT-ICR MS was performed both on original biochar samples and on
washed biochar as described below. However, data on the washed samples
was more comprehensive and better calibrated, i.e., more detected masses could be
assigned to formulas. In the original co-composted biochar, 6,063 detected
masses were not assigned formulas, while only 4,350 could not be assigned after
washing.
Biochar washing for liquid extraction of the organic coating. Biochar particles
were washed in four steps to remove any compost induced surface alterations and,
at the same time, to obtain a suspension of the co-composting derived material that
forms the coating of the biochar as identified with STEM. 15 mL DI water were
added to 2.5 g moist biochar in a Falcon tube (step 1). After 5 min on a roller
shaker at 50 rpm (Roller 10 digital, IKA, Staufen, Germany), water was decanted
through a 0.5 mm mesh sieve and 15 ml of DI water were added to repeat this
procedure (step 2). After decanting, 15 ml of 0.05 M NaOH solution were added as
suggested by Tsechansky and Graber28 and Falcon tubes were placed on the shaker
for 90 min (step 3). After decanting, 15 ml of DI water were added for 20 min to
remove residual NaOH (step 4). Separate washing procedures were conducted
for fresh and co-composted biochar. Afterwards, the biochar was dried at
40 °C for 72 h for DAPPI FT-ICR MS, SEM and STEM as described above. After
washing, co-composted biochar was almost black again (Supplementary Fig. 7).
Washed biochar was analyzed using SEM and STEM to verify the removal
of the coating. SEM showed that the original biochar surface structure was
widely exposed again, while few regions still showed surface alterations
(Supplementary Fig. 8). In STEM we could not identify a coating after washing,
but granular aggregations of Ca and O were present on the biochar surface
(Supplementary Fig. 9).
Washing solution of step 3 (0.05 M NaOH) was analyzed by FTIR,1H NMR,
excitation–emission matrix (EEM) fluorescence spectroscopy, ion chromatography,
and for dissolved organic and inorganic carbon (DOC/DIC) and electron accepting
and donating capacity (EAC/EDC). Before analysis, resulting washing solutions
were filtered through a 0.45 µm syringe filter. For FTIR, washing solutions
and original 0.05 M NaOH solution were freeze-dried. For1H NMR, washing
procedure was repeated and 0.05 M NaOH was prepared with D
2
O instead of H
2
O.
Major anions and cations were quantified by ion chromatography (IC, DX-120,
Dionex, Sunnyvale, CA, USA). DOC + DIC was quantified on a Vario cube
(Elementar, Hanau, Germany), DOC was quantified on the same device after
acidifying the samples to pH 1 with HCl. DIC was calculated as [DOC + DIC]—
[DOC].
Excitation–emission matrix. EEM fluorescence spectra of washing solution were
analyzed using a Fluoromax4 (Horiba, Jobin-Yvon) spectrofluorometer. EEM
fluorescence spectra were recorded over a range of excitation wavelength (300–500
nm) and emission wavelength (400–600 nm) relevant for natural organic matter58.
Electron accepting and donating capacity. The redox properties, i.e., electron
accepting and donating capacities (EAC/EDC) of biochar washing solutions were
determined by mediated electrochemical reduction (MER) and oxidation (MEO)
following established procedures6,59. In brief, the electrochemical system consists
of a glassy carbon cell (Sigradur G, HTW, Germany) as working electrode, a Ag/
AgCl as reference electrode (Bioanalytical systems Inc., USA) and a platinum wire
(0.5 mm, 99.9 %, Sigma-Aldrich Co., USA) attached to a platinum gauze (52 mesh,
99.9 %, Sigma-Aldrich Co., USA) as auxiliary electrode. The applied potential was
measured against Ag/AgCl electrode but is reported against standard hydrogen
electrode (EH =−0.49 V in MER and EH =+ 0.61 V in MEO). Electron transfer
mediators were used during the measurements to ensure a fast electron transfer
between the electrodes and the analyte59; diquat-dibromide monohydrate (DQ,
Sigma-Aldrich Co., USA) was used for MER and 2,2′-azino-bis(3-ethylben-
zothiazoline-6-sulfonic acid) diammonium salt (ABTS, Sigma-Aldrich Co., USA)
for MEO. EAC and EDC are calculated from the measured reductive and oxidative
currents considering normalization to carbon content (µmol e-/mmol C)6. The
total EEC is the sum of the individual accepting and donating capacities (EEC =
EAC + EDC). As an additional reference, dissolved organic matter was extracted
from non-biochar-amended control compost in analogy to the biochar washing
procedure (0.05 M NaOH, 1.5 h, same shaker).
Fourier-transformed infrared spectroscopy. Fourier-transformed infrared
(FTIR) Spectroscopy provides information on bonding modes in organic molecules
by absorption of infrared radiation, which depends the vibrational response of the
functional groups60. FTIR absorbance spectra of KBr pellets prepared with 0.2%
biochar, 0.4% freeze-dried washing solutions or 0.4% freeze-dried control compost
were measured with a Vertex 80 v (Bruker) with 128 scans. A KBr pellet without
sample was used for background measurements.
1H solution nuclear magnetic resonance spectroscopy. 2 ml of the pristine and
the co-composted biochar washing solutions were lyophilized (freeze-dried)
overnight. The dry samples were rehydrated up to 500 µl using 100% D
2
O. NMR
spectra were collected using a Bruker Avance III 600 MHz spectrometer equipped
with a Bruker 5 mm SmartProbeTM.1H 1D spectra were acquired for 2 s with a 20 s
recycle delay and 1024 scans using standard Bruker pulse sequence with water
suppression. All spectra were processed with applying a 0.30 Hz exponential line
broadening using Bruker Topspin 3.5.
Fast field cycling nuclear magnetic resonance relaxometry. 1 g of each biochar
sample was suspended in 3 g of MilliQ grade water (resistivity of 18.2 MΩcm at
298 K). Milli-Q water was produced by a Milli-Q Advantage A10 Ultrapure Water
Purification System (Millipore Corporation, Massachusetts, USA). The suspensions
were allowed to sediment overnight prior to the relaxometry investigations.
The samples were put in the probe of a Stelar SpinMaster FFC-2000 fast field
cycling relaxometer (Stelar s.r.l., Mede, PV−Italy) and analyzed at 25 °C. The
basic theory about FFC NMR relaxometry and the sequence applied for the
experiments reported in this study have been already summarized by Conte and
Alonzo31. In brief, non-polarized (NP) and polarized (PP) sequences were
applied. In the NP sequence, a relaxation field (B
RLX
) was changed between proton
Larmor frequency (ω
L
) values ranging in the interval 0.01–30 MHz. Each B
RLX
was
applied for a period τarrayed with 32 logarithmic spaced time sets, each of them
adjusted at every relaxation field in order to optimize the sampling of the
decay/recovery curves. At the end of each B
RLX
field a1H 90° pulse was applied into
an acquisition magnetic field (B
ACQ
) held for a fixed time at the ω
L
value
of 7.2 MHz. The 1H 90° pulse was needed to make magnetization observable and
the free induction decay acquirable. A time domain of 100 µs sampled with 1000
points was applied. Field-switching time was 3 ms, while spectrometer dead time
was 15 µs. For all of the experiments a recycle delay of 20 s was used. In the PP
sequence, a polarization field (B
POL
) set at the ω
L
of 9 MHz was applied prior to
each B
RLX
field. The period of time during which B
POL
was applied (referred to as
polarization time or T
POL
) corresponded to five times the T
1
estimated at this
frequency.
The crossover field between NP and PP sequences was approximately retrieved
when the relaxation field intensity was half of that of the polarization field31.
FFC NMR relaxometry data elaboration. The longitudinal relaxation time (T
1
)
values of the observed nuclei were obtained for each B
RLX
by changing the τvalues
as reported above. The relationship between signal intensity and τis modeled by
Eq. 1:
IτðÞ¼I0exp τ=T1
ðÞ½
k:ð1Þ
Here, I(τ) is the 1H signal intensity at each fixed B
RLX
,I
0
is the 1H signal
intensity at the thermal equilibrium, T
1
is the average proton spin-lattice relaxation
time and kis a heterogeneity parameter related to the stretching of the decay
process. This function, which accounts for the large sample heterogeneity resulting
in a multi-exponential behavior of the decay/recovery curves31, can be considered
as a superposition of exponential contributions, thereby describing the likely
physical picture of some distribution in T
1
. Eq. 1has the advantage that it is able to
handle a wide range of behaviors within a single model. For this reason,
assumptions about the number of exponentials to be used in modeling nuclear
magnetic dispersion (NMRD) data are not necessary. The NMRD profiles resulting
from the elaboration of the decay/recovery curves are reported in Supplementary
Fig. 13. The data acquired at the relaxation field of 0.01 MHz were transformed by
applying the Uniform PENalty regularization (UPEN) algorithm31 which allowed
the achievement of the distribution of the longitudinal relaxation times (also
referred to as relaxograms).
Liquid chromatography organic carbon detection. 1 g of pristine, co-composted
and soil-aged biochar was added to 10 ml of distilled water at 50 °C for 24 h and
regularly stirred, and then centrifuged and filtered (10,170 g, 5 min, 0.45 µm) to
separate the solid and liquid phases. The supernatant solutions were analyzed by
Liquid Chromatography –Organic Carbon Detection LC-OCD model 8 (DOC
Labor, Germany). Details of the measurement procedure have been described in
full by Huber et al61. In this study, a Toyopearl TSK HW50S LC column was used
with a phosphate buffered mobile phase of pH 6.4 at a flow rate of 1.1 ml min-1.
Injection volumes were 1,000 µl. The supernatant solutions are also analyzed to
determine total organic nitrogen and carbon and total inorganic carbon using a
thermocatalytic high-temperature oxidation process (Multi N/C 2100, Analytic
Jena, Germany).
The chromatographic eluent subdivides into six sub-fractions, biopolymers,
humics, building blocks, low molecular-weight neutrals and hydrophobic organic
carbon (Supplementary Table 5), as suggested by Huber et al.61
Data availability. The data sets generated during and/or analyzed during the
current study are available from the corresponding author on reasonable request.
Received: 16 March 2017 Accepted: 15 August 2017
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Acknowledgements
We thank E. Struve, K. Mayer and P. Riede for laboratory support, G. Müller for support
at Canadian Light Source, P. Ingio and F. Zeitvogel for support during STXM data
evaluation, H. Chen for support at NHFML, W. Gerber for photographs, D. Mitchell for
input on EELS and A. Flicker and M. Nowak for FTIR access and support. N.H. was
financially supported by a BMBF PhD scholarship provided by the Rosa Luxemburg
Foundation, Berlin, Germany. T.B. and R.B.Y. were supported by the Agriculture and
Food Research Initiative Competitive (grant no. 2013-67019-21359) from the USDA
National Institute of Food and Agriculture and a National Science Foundation SusChEM
Award (grant no. EAR1451494). The co-composted biochar was obtained from a com-
post that was produced during a ‘Short Term Scientific Mission’(STSM) granted to NH
by the EU COST Action TD1107 ‘Biochar as option for sustainable resource manage-
ment’. Access to the ASTEM Titan3 60–300 at TU Graz was enabled by European Union
through the EC Grant ESTEEM2 (20141212‐Hagemann and 20150703-Hagemann).
Research described in this work was performed at the Canadian Light Source, which is
supported by the Natural Sciences and Engineering Research Council of Canada, the
National Research Council Canada, the Canadian Institutes of Health Research, the
Province of Saskatchewan, Western Economic Diversification Canada and the University
of Saskatchewan. A portion of this work was performed at the National High Magnetic
Field Laboratory, which is supported by the National Science Foundation Division of
Materials Research through DMR 11-54790, the State of Florida and Florida State
University. We thank E. Graber and K. Spokas for valuable discussions.
Author contributions
The study was jointly designed by N.H., S.J., H.-P.S., S.B. and A.K. N.H. and H.-P.S.
performed the composting experiment. J.H. performed the soil-aging experiment. N.H.
performed extraction experiments and coordinated all other analysis. N.H. and A.K.
wrote the manuscript, which was greatly improved by C.I.K., T.B. and H.-P.S. Spectro-
scopical and spectrometric analysis were performed as follows: SEM: S.J. and N.H.;
STEM: S.J., M.A., N.H. and C.M.; STXM: M.O.; NMR relaxometry: P.C.; NMR spec-
troscopy: K.V. and K.W.E.; gas adsorption: A.D.-A.; FT-ICR-MS: R.B.Y. and A.M.; EEC:
E.S., S.O.; LC-OCD: S.J. and S.T.
Additional information
Supplementary Information accompanies this paper at 10.1038/s41467-017-01123-0.
Competing interests: The authors declare no competing financial interests.
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