Nothofagus spp. wood (Southern Beech), chemically consistent with modern angiosperm wood in its lignocellulosic structure (Bird et al., 2014), was obtained from the Yallourn coal seam, as described in Tilston et al. (2016). Sugarcane (Saccharum officinarum) bagasse (fibrous matter remaining after crushing of sugarcane) was obtained from Mossman Sugar Mill, Australia, representing a cellulose-rich material. Macroalgae (Cladophora vagabunda) was collected at the Townsville Barramundi Fish Farm in Kelso, Townsville Australia. This sample represents a non-woody material with lower carbon concentration and surface area than wood or bagasse, but high in nitrogen and inorganic nutrients (Bird et al., 2011; McBeath et al., 2015). All three materials are important potential sources of biochar for carbon sequestration and/ or agricultural amendment. Finally, a commercial wood waste biochar, prepared at 550°C, was obtained from BEST Energies Ltd., Australia.

The Nothofagus spp. wood, S. officinarum bagasse and C. vagabunda algae were dried at 105°C, chipped and sieved, with the >2mm fraction retained for pyrolysis, and converted to PyC under a 3.5 L min⁻¹ nitrogen flow for 1 h. Three temperatures of 305°C, 414°C, and 512°C were used, as in Bird et al. (2011). This is the temperature range over which the most significant changes in PyC chemistry are observed (Ascough et al., 2008). Following pyrolysis, samples were lightly crushed and sieved to 0.5–2mm. Together with the BEST biochar, this yielded 10 sample types for field exposure.

Samples were exposed to local environmental conditions using a litterbag approach (see below) for 1 year at the Daintree Rainforest Observatory, Queensland, Australia. The field location is in a UNESCO World Heritage Site at Cape Tribulation, 140 km north of Cairns (16.103°S; 145.447°E; 70 m asl; Figure 1). Mean monthly temperature is 22–28°C (annual mean 25°C), and annual rainfall is 3,500mm, with a pronounced December to March wet season. This site was chosen to maximize the rate of PyC-environment interactions, plus no historical burning is recorded, meaning natural PyC abundance in the soil is very low.

Vegetation is an evergreen mesophyll to notophyll “tall forest” (Torello-Raventos et al., 2013) producing broadleaf litter to 5–10cm depth in the dry season, which rapidly decays in the wet season. Soils are developed on colluvium from metamorphic and granitic mountains, forming Acidic, Dystrophic, Brown Dermosols (Deckers et al., 1998; Isbell, 2002) or Haplic Cambisols (Hyperdystic, Alumic, Skeletic; Deckers et al., 1998), with a dark grayish brown silty loam to silty clay loam upper A-horizon. The profile contains 20–50% cobbles and stones, and the organic-rich (3.7% C, 0.3% N) soil is mildly acid (pH 5.5–6.5) with the <2mm faction (0–10cm) comprising 28% clay, 54% silt and 19% sand.

For each aliquot c.5 g of dry sample was placed within a 125μm aperture nylon mesh bag, sealed and pegged to the soil surface, which had been cleared of leaf litter. The litterbag approach is well-established in the soil science literature, although potential drawbacks must also be considered, for example partial inaccessibility of litter to decomposers (St John, 1980). Four manipulations (soil chemical environments) were used and samples were duplicated for each of these (i.e., 20 samples per manipulation). The bags were covered with polyester shade cloth and enclosed in a wire mesh cage to exclude foraging wildlife. The treatments comprised:

NL—Samples laid directly on the soil surface without any litter cover;

The exact pH at the sample site was not determined, but surface soil and leachate water after 1 year of environmental exposure shows that treatments L and NL had a local pH of 5.6–6.5, and treatments L-LM and NL-LM had a local pH of 6.6–8.0. After recovery samples were gently washed free of loosely adhering soil particles, dried at 105°C and weighed. Both freshly prepared samples and exposed material were analyzed as described below.

Carbon (%C) and nitrogen (%N) concentration was measured using a Costech elemental analyser (EA) (Milan, Italy) fitted with a zero-blank auto-sampler. External reproducibility for %C and %N was better than 0.5%. The stable carbon isotope ratio (expressed as δ¹³C) of the samples was measured using a ThermoFinnigan Delta^plus XL isotope ratio mass spectrometer (Thermo Finnigan GmbH, Bremen, FRG), linked to the EA via a ConFlo III. Analytical precision was ±0.2‰ for δ¹³C and ±0.3‰ for δ¹⁵N. Oxygen (%O) and hydrogen (%H) concentration was measured using a Thermo Finnigan high temperature conversion elemental analyser (TC/EA) (Thermo Finnigan GmbH, Bremen, FRG). External reproducibility for %O and %H was better than 0.7%. The ash concentration of the samples was determined by loss-on-ignition of 1 g at 900°C in ceramic crucibles in a muffle furnace where reproducibility was better than 1.0%.

Hydrogen pyrolysis (HyPy; Ascough et al., 2009, 2010b; Meredith et al., 2012) was used to release the < 7 ring polycyclic aromatic hydrocarbons (PAHs, the non-stable PyC fraction), from all samples. Briefly, samples were loaded with an aqueous/methanol solution of ammonium dioxydithiomolybdate [(NH₄)₂MoO₂S₂], and pyrolysed by resistive heating to a final temperature of 550°C, all under a hydrogen pressure of 15 MPa and a sweep gas flow of 5 L min⁻¹ (Meredith et al., 2012).

HyPy products were trapped on cooled silica, and desorbed with 10 ml aliquots of n-hexane and dichloromethane (DCM). Eluents were combined and GC–MS analyses in full scan mode (m/z 50–450) were performed on a Varian CP-3800 gas chromatograph equipped with a VF-1MS fused silica capillary column (50m × 0.25mm i.d., 0.25mm thickness) with helium as the carrier gas, interfaced to a Varian 1200 mass spectrometer (EI mode, 70 eV). The abundances of individual PAHs were quantified using the mass chromatograms of the molecular ion of each compound, following the addition of squalane (Aldrich) and 1-1 binapthyl (Acros Organics) as internal standards, assuming a response factor for each compound of 1 (Meredith et al., 2013). The PAHs comprise those that are solvent extractable (“free”) and those covalently bonded to the biochar released by HyPy.

Solid-state ¹³C Nuclear Magnetic Resonance Spectroscopy, using cross-polarization magic angle spinning (¹³C-CPMAS NMR) was used to identify polyaromatic vs. aliphatic and O-alkyl carbons. A 400 MHz Varian VNMRS instrument was used, operating at 100.56 MHz for ¹³C using a 4 mm magic-angle spinning probe with a zirconium oxide rotor and Teflon end caps. Spectra were referenced to external, neat tetramethylsilane, and for cross polarization, typical acquisition conditions were a 1 s recycle delay, 1 ms contact time and a sample spin-rate of 12 kHz. A variable amplitude 1H spin-lock pulse was used for the cross polarization step. The pulse sequence incorporated a spin-echo to suppress background signal from the Vespel spinner housing; this has resulted in a small amount of signal in some spectra at ~111 ppm.

Weights of samples and their carbon concentration were accurately determined before and after exposure in the field, making it possible to quantify changes in sample mass and carbon concentration during exposure of the samples to the environment. The proportional mass difference in samples before (M_T0) and after exposure (M_T1) is obtained by:

The proportional change in carbon concentration (mg C), ash concentration, oxygen, hydrogen, and nitrogen are obtained by substituting values before (X_T0) and after exposure (X_T1) into Equation (1).

The potential for significant differences between mass and C/O/H/N for samples was assessed after data was scrutinized for normality using a D'Agostino-Pearson normality test. Subsequently, samples were compared using a Two-way ANOVA followed by Tukey's multiple comparisons test, performed using GraphPad Prism version 7.02 for Windows, GraphPad Software, La Jolla California USA, www.graphpad.com. A p-value of < 0.05 indicated a significant difference.

Full results of measurements for all samples are presented in Supplementary Table S1, and summary data is presented in Table 1. The mass and ash concentration changes following field exposure for Nothofagus spp. wood biochar samples are presented in Bird et al. (2017) (Table 1), but are discussed here in comparison to other samples. Nothofagus wood spp., S. officinarum bagasse, and BEST biochar contained little ash, with most samples below the limit of detection before and after environmental exposure (Table 1). The exception is C. vagabunda algae biochar, which contained 28–38% ash by weight, increasing with production temperature. A large amount (65–83%) of this ash was lost during exposure (Table 2).

The mass of BEST biochar changed little with environmental exposure, and (excluding treatment NL), Nothofagus spp. wood mass increased by <10% (Table 2) (NB: isolated reductions in mass of the latter samples are outweighed by the increases). On the other hand, large mass increases of up to +36% occur in treatment NL for Nothofagus spp. wood and S. officinarum bagasse biochar shows changes of −9 to 41% (Table 2). C. vagabunda algae biochar again behaved differently, as its mass decreased by −41 to −52% (Table 2). This was consistently different to both the Nothofagus spp. wood and S. officinarum bagasse biochar (p ≤ 0.005). There were statistically significant differences at 95% confidence between sample types, and treatments. An interaction between sample type and treatment (p ≤ 0.05) was only observed for the 300°C samples, where mass gains in Nothofagus spp. wood biochar were greater than for other materials. At higher temperatures, there was no difference between the overall mass increases for Nothofagus spp. wood and S. officinarum bagasse biochar. C. vagabunda algae biochar showed no significant effect of treatment type at 95% confidence, and there was no clear difference in this between different temperatures, indicating that the mass losses in this sample type were consistent for all samples after 1 year. For the 300°C and 400°C Nothofagus spp. wood and S. officinarum bagasse biochar, treatment NL was associated with higher mass losses than other treatments (p ≤ 0.05). At 500°C this effect was only evident for the S. officinarum bagasse biochar. Overall, C. vagabunda algae biochar showed a very different pattern of change to other sample types, and differences between treatments were most pronounced for samples produced at lower temperatures.

Full results of measurements for all samples are presented in Supplementary Table S1, and summary data is presented in Table 1. For Nothofagus spp. wood, S. officinarum bagasse, and BEST biochar before environmental exposure, %C, concentrations clearly increased and %O and %H concentrations decreased as production temperature increased (Table 1), consistent with previous work (e.g., Ascough et al., 2008). C. vagabunda algae is different; although O is progressively lost at higher production temperatures, C and H show little change with production temperature after 300°C. Only C. vagabunda contained measureable quantities of nitrogen, which increased with initial charring at 300°C, thereafter remaining unchanged at production temperatures up to 500°C (Table 1).

For samples after environmental exposure, in all treatments except NL (exposed on the surface without litter cover) changes in carbon concentration relative to before exposure ranged from −1 to 33% in Nothofagus spp. wood, most S. officinarum bagasse, and BEST biochar (Table 2). Again, C. vagabunda algae is different; after environmental exposure C concentration decreases markedly, between −40 and −50%. There was no interaction between treatment and material type at any temperature (where p ≥ 0.1 for the possibility of significant difference). There was no significant difference between carbon concentration increases in Nothofagus spp. wood, and S. officinarum bagasse except for treatment NL in 300°C samples (p = 0.05), where C increases in Nothofagus spp., wood, exceeded S. officinarum bagasse. Both were, however, consistently different to C. vagabunda biochar (p ≤ 0.01). For at least some Nothofagus spp., wood, and S. officinarum bagasse samples prepared at 300 and 400°C, treatment NL showed higher C increases than other treatments (p ≤ 0.05). This effect was not apparent in the 500°C samples).

After environmental exposure, oxygen concentration rises in Nothofagus spp., S. officinarum, and BEST biochar, in contrast with large decreases in oxygen in C. vagabunda algae biochar (p ≤ 0.05). There are apparent differences between Nothofagus spp., and S. officinarum biochar prepared at 400 and 500°C, where oxygen concentration increases are greater in S. officinarum biochar (e.g., 55% vs. 100% change in treatment NL). For C. vagabunda algae biochar there is no clear effect of treatment upon sample oxygen content, whereas for Nothofagus spp., and S. officinarum biochar at all temperatures, oxygen increases in treatment NL are greatest. Oxygen increases for samples except C. vagabunda algae biochar appear somewhat larger in samples prepared at higher temperatures, although this correlation could not be tested.

After exposure, changes in hydrogen concentration broadly mirror those observed for %O, where the %H changes in C. vagabunda algae biochar (c.40% loss of starting amount) are significantly different to the overall H gains in other samples (p ≤ 0.001). Changes in other samples are not significantly different from each other at any production temperature, and there is no clear effect of production temperature upon results for any specific material. For C. vagabunda algae biochar H change was not clearly different between treatments, whereas for other samples, increases in H were consistently greater in treatment NL. The net effect of %C, %O, and %H changes is that the pattern in O/C and H/C in all samples (i.e., a decrease with production temperature) is still clearly apparent (Figure 2).

There is little difference in %N in C. vagabunda algae biochar following environmental exposure; the majority of measured values do no change by greater than the limit of analytical uncertainty of ±0.5%, and there is no significant difference with production temperature or treatment (p ≥ 0.01 for the possibility of a relationship).

Full results of measurements for all samples are presented in Supplementary Table S1, and summary data is presented in Table 1. Decreases in Nothofagus spp. wood and S. officinarum δ¹³C during charring are due to loss of cellulose (Ascough et al., 2008). Conversely, C. vagabunda algae shows a small increase in δ¹³C of ~1‰ above 400°C, this is greater than the analytical uncertainty of ±0.2‰ (Table 2). For the vast majority of samples, δ¹³C following environmental exposure did not vary in excess of normal analytical uncertainty. The exception is an increase of up to 1.1% in C. vagabunda algae biochar produced at 400°C following exposure. Only C. vagabunda contained sufficient nitrogen to measure δ¹⁵N values; these increased with charring to 300°C but were then constant to 500°C. The δ¹⁵N of all C. vagabunda algae biochar decreased after environmental exposure by up to 0.5‰; this is greater than the analytical uncertainty of ±0.3‰ with no correlation to production temperature or treatment.

The HyPy products consist of the <7 ring polycyclic aromatic hydrocarbons (PAHs) in the samples and comprise both those covalently bound to the biochar and present in the free (solvent extractable) state. The measured concentrations of PAHs in the samples are low overall and probably represent only a fraction of the labile biochar carbon (Table 3). Along with pyrene and phenanthrene a range of compounds known to result from incomplete combustion of biomass are detected (Simoneit, 2002).

In freshly-produced Nothofagus spp. biochar, PAH concentration decreased with production temperature (Table 3); this trend was reversed following exposure. Extractable PAHs in Nothofagus spp. PyC are more abundant than in biochar produced from the other starting materials. In fresh S. officinarum bagasse biochar the total weight of PAHs increased with production temperature (Table 3), as does the ratio of 3+4:5+6 ring PAHs. Following exposure there was a slight increase in PAH abundance in all S. officinarum bagasse samples, which was largest in the 300°C S. officinarum. The pattern in ratio of 3+4:5+6 ring PAHs remains the same, although there is an anomalously high ratio (6.25) for 400°C samples in treatment NL.

In fresh C. vagabunda algae biochar PAH concentration also increased with production temperature. One difference from other samples is that naphthalene does not feature in the spectra of C. vagabunda algae samples either before or after environmental exposure. After exposure only the PAH concentration of the 400°C samples increases, particularly pyrene and fluoranthene. In BEST biochar the total content of PAHs is similar to that in Nothofagus spp. samples, although the absolute concentrations remain small. There is no effect of treatment upon PAH concentration.

The changes over 300–500°C for lignocellulosic biomass comprise loss of cellulosic carbons at 60–105 ppm, lignin at 55 ppm, and aliphatic carbon (at 10–40 ppm), and increasing dominance of aromatic carbons at c. 129 ppm (Figure 3). In S. officinarum bagasse the process of aromatization is slower than in Nothofagus spp. wood, with peaks for lignin and aliphatic carbons at 400°C. Although C. vagabunda algae is substantially different to the other feedstocks (e.g., higher lipid concentration at 173 ppm), this sample follows the overall charring pattern above, and is dominantly aromatic at 500°C.

After environmental exposure there are no clear differences attributable to treatment. Instead, production temperature has the biggest effect upon chemical changes. For samples produced at 300°C, cellulosic carbons are removed in preference to lignin, and many samples are oxidized, with peaks for carboxylic carbons at c.170 ppm, and signal at 150 ppm for oxygenated aromatics. Aromatic carbons increasingly dominate the spectra. Aliphatic signal is reduced, demonstrating the reduction in proportion of aliphatic carbons in the sample, relative to starting material. Remaining aliphatic carbons may be lipids, which are known to resist oxidation (Knicker, 2010). The aromatic peak in samples shifts closer to 130 ppm and narrows after exposure, meaning that the range of different forms of polyaromatic domains is reduced, possibly due to loss of smaller aromatics (Figure 3).

Production temperature is known to dictate the aromaticity of biochar/PyC, and determine the extent to which non-aromatic material (including oxygen and hydrogen) is removed from the sample (e.g., Ascough et al., 2008; Krull et al., 2009). A finding from this research, however, is that lignocellulosic and non-lignocellulosic biomasses have very different trajectories of chemical change during pyrolysis. In lignocellulosic biomass there is basically a linear relationship between temperature and carbon increase/ oxygen and hydrogen decrease, and aromaticity. At temperatures of c.500°C, samples become increasingly homogeneous. This is likely to have the effect that trajectories of environmental alteration for a specific environment are similar for different types lignocellulosic biomass (where the starting composition is similar) produced at a consistent temperature. This is borne out by the results for Nothofagus spp. and BEST wood biochars presented below. A difference is that the rate of aromatization in S. officinarum bagasse samples appears slower at lower temperatures than in Nothofagus spp. wood biochar or BEST Biochar. We posit that this is the basis for the different patterns of environmental alteration we observe for these samples.

During production of C. vagabunda algae biochar there appears to be a chemical factor that limits aromatization over the 300–500°C window. This could be the effect of chemical reactions involving the high ash concentration during pyrolysis (McBeath et al., 2015), meaning that the potential for environmental alteration of C. vagabunda algae biochar (i.e., non-lignocellulosic material) is greater, compared to lignocellulosic biomass, such as wood, for a given temperature. This is consistent with observations that non-woody biochar is less stable than plant-based material produced at an identical temperature (e.g., Singh et al., 2012). It is also important to consider that there may be physical differences between these types of materials that affect their survival potential in the environment, such as porosity, and micro/ macroscopic cracking. However, the physical structure of the biochars was not assessed in this study, so in the following discussion we focus on chemical factors.

The process of PyC production forms PAHs (e.g., Ballentine et al., 1996; Keiluweit et al., 2012), thought to form a proportion of the “semi-labile” fraction in PyC (McBeath et al., 2015). In Nothofagus spp. wood the highest temperature biochar has the lowest concentration of PAHs, indicating more extensive aromatization, which would raise the carbon content of the biochar, as observed in the results presented here. This would mean that the potential for environmental alteration is lower for a woody biomass such as the Nothofagus spp. biochar sample produced at the same temperature as S. officinarum bagasse and C. vagabunda algae samples, which display lower C contents The potential for alteration/ degradation appears linked to degree of aromaticity (Harvey et al., 2012) and O:C ratio (Spokas, 2010), meaning for the samples analyzed here, predicted alteration potential increases in the order BEST biochar< Nothofagus spp. Wood< S. officinarum bagasse< C. vagabunda algae. These predictions are confirmed by the results observed after environmental exposure of the samples.

The results of the study, when considered in total, were consistent with the research hypothesis that material prepared at higher temperatures is less subject to alteration in the environment, supporting previous work (e.g., Zimmerman, 2010). For example, mass losses only displayed an interaction between sample type and treatment (p ≤ 0.05) for samples prepared at 300°C, and treatment effects for both mass losses and changes in C concentration were stronger at lower temperatures. However, the effects we observed were relatively subtle, although also manifest in the ¹³C-MAS-NMR, and PAH results, as explained below. We posit that these effects are predominantly due to a decrease in less labile carbon vs. resistant polyaromatic domains in higher-temperature biochars. Despite this, some small changes in the ¹³C-MAS-NMR spectra of even samples produced at 500°C are observed, as the aromatic peak at c.130 ppm was slightly narrowed for all feedstock types and treatments. It is possible this reflects loss of a small proportion of the range of aromatic domains present in the samples before environmental exposure, most probably a fraction of the semi-labile component (sensu Bird et al., 2015). This is supported by the results of PAH measurement, where the greatest reduction in PAH concentration is seen for the lowest temperature biochar. The mechanism by which this could be achieved remains unclear, although growing evidence demonstrates microbial action resulting in degradation of indigenous biochar carbon over timescales on the order of a year (Ameloot et al., 2013; Farrell et al., 2013; Tilston et al., 2016; Bird et al., 2017). Increasing aromatization also increases resistance to oxidative processes, known to be one of the main agents of PyC alteration and degradation. This is shown in the O/C ratios where despite the loss of many oxygen-containing O-alkyl structures, this is more than offset by the additional oxygen introduced to the structure of lower temperature (300–400°C) biochars after 1 year.

Differences in biochar chemistry after environmental exposure attributed to specific feedstock are relatively low between Nothofagus spp. wood PyC, and S. officinarum bagasse biochar, and although changes appear somewhat smaller in BEST Biochar, for all these sample types, changes in mass, C/O/H/N, and other characteristics are in the same direction. Changes in C. vagabunda algae biochar for these parameters are always in the opposite direction, however, and are consistently larger than those in other samples. The results therefore support the hypothesis that, although different types of woody biomass show relatively small differences in change with environmental exposure, at least at a gross scale, non-woody biomass is more susceptible to alteration than woody biomass (c.f. Hilscher et al., 2009; McBeath et al., 2014). This appears largely due to the removal of ash that is readily solubilised and/or physically migrated away from the sample. This ash contains essential plant nutrients (e.g., Na, Ca, Cl, Mg, K), and so this type of biochar may have a beneficial effect for soil fertility (e.g., Novak et al., 2009), although this will be over a restricted timescale. This type of biochar contains more labile carbon than the other samples, particularly aliphatic material indigenous to the sample and sorbed to its structure (Middelburg et al., 1997), and the C concentration decrease observed in the samples, potentially due to loss of labile C, is so large that any introduction of exogenous carbon, such as that observed in other samples (e.g., via soil microorganisms c.f. Steinbeiss et al., 2009; Bird et al., 2017), is not visible. It should be noted that along with complete degradation to CO₂, there is the possibility that some C is removed by solubilization and leaching during environmental exposure (Major et al., 2010; Abiven et al., 2011; Bird et al., 2017).

C. vagabunda algae biochar is the only sample in which δ¹⁵N and %N can be observed, although there is no effect of treatment or production temperature. There is a slight increase in δ¹⁵N overall, although this is very close to the limit of analytical precision. This may indicate that microbial action has removed lighter N isotopes via fractionation during decomposition (bonds involving ¹⁴N being broken more easily), possibly through decomposition of “black nitrogen” (Knicker, 2010; McBeath et al., 2015).

There is a clear effect of depositional environment on biochar alteration even after the relatively short timescale of this study. However, this is not consistent with the hypothesis that material in alkaline environments is more subject to alteration. Although for C. vagabunda algae biochar there is no effect from different treatments upon chemical changes, for other samples, increases in mass, and C/O/H concentrations are far more extensive in the material under the NL treatment. Mass increases due to ash and carbon increases are much greater in all samples under this treatment; the mass increase indicates the potential for a significant proportion of soil mineral ingress (c.f. Bird et al., 2008). Over time in the environment, it is evident that mineral material can strongly associate with both external and internal surfaces of PyC (Lehmann et al., 2011; Jaafar et al., 2015). This can be the result of abiotic processes such as water movement and biotic processes such as the movement of soil macrofauna. This interpretation is further supported by the increases in ash content in many samples, a phenomenon previously observed by Bird et al. (2014). An increase in sample carbon could result from removal of non-carbon elements (e.g., H), due to consumption by microbes. This labile carbon removal has been observed in incubations of the Nothofagus spp. wood charcoal samples (Bird et al., 2017). Conversely, increases in sample carbon concentration and mass could result from the ingress of exogenous carbon, e.g., fungi, soil biota, humic acids (Steinbeiss et al., 2009; Lehmann et al., 2011). For example, Mohan et al. (2014), and others demonstrate the sorption of organic compounds from the depositional environment onto the structure of PyC. Fungal ingress to samples equivalent to some of the material in this study has observational support in field (Bird et al., 2014), and laboratory (Ascough et al., 2010c) studies. The concept of an increase in mass due to introduction of exogenous carbon for the NL treatments is supported by the findings of Bird et al. (2017), using equivalent material to the Nothofagus spp. biochar described in the present study. Here, an increase in radiocarbon concentration of ~0.5–1 pMC clearly demonstrates the addition of exogenous carbon to the radiocarbon-dead biochar. Using δ¹³C measurements, the same study indicated a 5–10% contribution of exogenous carbon to the lower temperature biochars. This is also found in our results, where a decrease in δ¹³C occurs overall, albeit to a lower extent than observed for the sample group analyzed by Bird et al. (2017) after 3 years. In this case, the larger %C increases observed in NL may be due to the higher oxygen availability for these samples on the soil surface, leading to unhindered access for microbiota, leading to decomposition of the more labile fraction of the biochar structure (containing non-C elements), raising the %C content as polyaromatic material, dominated by C remains behind. This use of biochar carbon by microbiota is consistent with the findings of Bird et al. (2017) for biochars inoculated with microbial material after drying following environmental exposure. Respiration of CO₂ from these samples was linked to both use of biochar carbon and some of the exogenous carbon from the deposition environment, using δ¹³C measurements. This would explain why mass and %C increases on the same scale are not observed in the NL-LM treatment, even though this was also on the soil surface without litter. Here the limestone covering could physically (through reduced oxygen availability), or chemically (through increased pH) retard the ingress of microorganisms.

The ingress of exogenous carbon is supported by the δ¹³C measurements, as the sample materials all have higher δ¹³C values, by up to 15.4‰, than the surrounding leaf litter (−29‰). The δ¹³C are small, but in the negative direction. Based on the observations of Bird et al. (2017), this ingress is limited to 1–2% carbon; therefore as the carbon increase for the NL treatment is so large, these low changes in δ¹³C indicates use of indigenous sample carbon by microbiota inhabiting biochar surfaces.

Chemical changes in the samples following environmental exposure include oxidation of the biochar structure, which is most complete for samples under the NL treatment, where significant increases in oxygen concentration are observed. The results of the ¹³C-MAS-NMR measurement suggest that direct oxygenation of aromatic structures may be even more important than carboxylation in biochar/ PyC alteration, as signal at 150 ppm for oxygenated aromatics (Freitas et al., 1999) is present, while carboxylic peaks are small. Similarly, changes in the amounts of PAHs in samples after exposure indicate removal of pre-existing small-ring aromatics, and possibly removal of material that has been formed by cleavage from the larger aromatic skeleton of the biochar. However, the fact that these PAHs are extractable from the samples after environmental exposure points to these being sorbed to the sample structure, slowing their biodegradation and/or removal from site of formation after being cleaved from initially larger aromatic structures (c.f. Manilal and Alexander, 1991).

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.