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Chemical and structural properties of carbonaceous products obtained by pyrolysis and hydrothermal carbonisation of corn stover.


Carbon (C) sequestration via conversion of biomass (short-term biodegradable carbon) into a more durable form (e.g. charcoal) has created a great deal of interest within the framework of global climate change (Seifritz 1993; Titirici et al. 2007; Lehmann and Joseph 2009). Coalification has occurred naturally as a process mitigating high concentrations of atmospheric C[O.sub.2] on a geological time scale. High concentrations of atmospheric C[O.sub.2] have generally been associated with warm periods (Petit et al. 1999; Vimeux et al. 2002) and high rainfall (Zachos et al. 2001). Under such conditions, high biomass growth in low lying areas was favoured, and after subsequent subsidence over long periods, coal was formed. The large-scale use of this geological source of C from the mid 20th Century onwards has been the main cause of increasing concentrations of greenhouse gases (GHG) in the atmosphere. Imitating nature by sequestering carbon through the conversion of biomass into recalcitrant C forms (e.g. charcoal), in order to reverse the present anthropogenic C flux, should therefore not be disregarded.

Much attention is currently focused on obtaining charcoal from slow-pyrolysis processes, with the final aim of adding it to soils as a C sink and to enhance soil properties. In this context, the charred material is denoted 'biochar' (Lehmann et al. 2006; Lehmann and Joseph 2009). Biomass can also be converted into carbonised material by hydrothermal carbonisation (HTC), and the final solid product of this process is termed 'hydrochar' (Titirici et al. 2007; Sevilla and Fuertes 2009a, 2009b; Hu et al. 2010). The latter technique was developed early in the 20th Century with the aim of understanding the mechanisms of coal formation (Bergius and Specht 1913; Berl and Schmidt 1932), as the process of low-temperature HTC is considered to simulate natural coalification at accelerated rates (Hu et al. 2010).

While pyrolysis is based on carbonisation of dry feedstock under [O.sub.2]-free conditions, low-temperature HTC consists of treatment of a mixture of biomass and water at temperatures in the range 170-300[degrees]C, under pressure (Sevilla and Fuertes 2009a, 2009b; Hu et al. 2010). Both processes produce a loss of major constituents (C, H, and O) from the solid phase, with H and O being lost in proportionally greater amounts than C, and such losses are favoured at high temperatures (Amonette and Joseph 2009; Sevilla and Fuertes 2009a). During pyrolysis, condensable gases (e.g. water, hydrocarbons, and tarry vapours) and non-condensable gases (e.g. H2, CO, and C[O.sub.2]) are released, resulting in production of liquid and gas phases, in addition to the solid phase (Antal and Gronli 2003). The HTC process gives rise to water-soluble organic substances (furfural, hydroxymethylfurfural, organic acids, aldehydes, phenols) (Karagoz et al. 2005), gases (e.g. C[O.sub.2]), and a solid product (Sevilla and Fuertes 2009a, 2009b).

Since processes of C condensation during pyrolysis and HTC take place under very different conditions, differences in the solid carbonaceous material are to be expected. Biochar has been reported to have a lower char yield than hydrochar and to be more resistant to microbial decomposition (Hu et al. 2010), although to date, no comparative studies have been carried out. The stability of biochar will, however, be highly dependent on process conditions and, specifically, on the temperature of pyrolysis (Keiluweit et al. 2010). The surfaces of carbonaceous materials produced through HTC have been reported to be very reactive (Hu et al. 2010), and suggested to be dominated by hydroxyl, carbonyl, and carboxylic functional groups (Sevilla and Fuertes 2009a, 2009b), which may therefore improve the chemical reactivity of the charcoal in soils. Conversely, the surfaces of fresh biochars tend to be rather unreactive, although the reactivity increases with ageing (Cheng et al. 2006, 2008) and/or when the charcoal is activated (Downie et al. 2009).

Knowledge of the characteristics of biochar and hydrochar products is therefore crucial if these more durable forms of C are to be applied to soils for agricultural and/or environmental purposes. It is within this context that the main objective of the present study is proposed, i.e. characterisation of the carbonaceous materials produced from corn stover by (i) pyrolysis at 550[degrees]C, and (ii) HTC at 250[degrees]C under pressure. Energy balances, although needed to evaluate the overall C sequestration potential of the technologies, are outside the scope of the present study.

Materials and methods

Raw material

The biomass sample used in the study was collected in the region of Manawatu (New Zealand). Corn stover stocks were initially cut to 25 mm with an electronic chipper and then cut in a 5-mm cross-cutting mill. The material was dried at 60[degrees]C for 24 h before pyrolysis. Cellulose, hemicellulose, and lignin contents were determined using the method based on sequential chemical treatments with neutral detergent, acid detergent, 72% H2504, and ashing (Van Soest 1967). The amounts of cellulose, hemicellulose, and lignin were 38, 35, and 4.9 wt%, respectively.

Preparation of carbonaceous products

Corn stover (200 g) was pyrolysed in a self-purged, gas-fired, rotating drum kiln (made of stainless steel, inner volume 5 L). The raw material was heated to 550[degrees]C (heating rate ~40 [degrees]C/min) for 15 min and then allowed to cool. The resulting solid product was denoted 'biochar'. For HTC, corn stover (22.25 g) was dispersed in water (42 mL) and left for 12 h. The mixture was then transferred to a stainless steel autoclave fitted with a stirring mechanism, heated to 250[degrees]C, and maintained at this temperature at a pressure of 4 MPa for 4 h. The resulting solid product, denoted 'hydrochar', was recovered by filtration, washed with abundant distilled water, and then dried at 110[degrees]C for 4 h.

Characterisation of materials

Chemical characterisation

Elemental analysis (C, H, N) was performed with a TruSpec CHN analyser (LECO Corp., St Joseph, MI). Samples of the chars were combusted at 900[degrees]C for 4 h for determination of ash content. Oxygen content was estimated as follows: O = 100--(C+H+N+ash). The atomic ratios of H/C and O/C are presented on a dry ash-free basis. The effective cation exchange capacity (ECEC) was determined by a modification of the method of Matsue and Wada (1985), according to which the charcoal was equilibrated with 0.01 M Sr[Cl.sub.2], the retained Sr was displaced with 0.5 M HCI solution, and the displaced Sr determined by atomic absorption spectrometry (Perkin Elmer 2380, Norwalk, CT). Prior to this, the biochar sample was rinsed with abundant distilled water to remove ash and thus prevent it from interfering in the determination. Final pH values of the biochar and hydrochar samples after equilibrating with Sr[Cl.sub.2] were 7.54 and 4.90, respectively. The CEC at a buffered pH value of the rinsed samples was determined by the 1 M NH4OAc method (at pH 7) (Chapman 1965). The lime equivalence of the biochar sample was also determined in 5 g of biochar to which 100 mL of 1 M HCl was added. The suspension was back-titrated with 1 M NaOH to pH 7. Results are expressed in kg CaC[O.sub.3]/t biochar. The lime equivalence of the hydrochar was not determined because of the acidic nature of this material.

Contents of acid groups were determined by potentiometric titration (Lopez et al. 2008; Petit et al. 2010). A suspension of each sample was prepared in 0.1 M KN[O.sub.3] and its pH determined. The biochar sample was subsequently acidified with concentrated HCl and thereafter dialysed and lyophilised, to eliminate interfering salts. Hydrochar and biochar suspensions were potentiometrically forward-titrated with KOH or backtitrated with HCl in order to cover the range of pH 3-10. However, as the pH values of both chars were buffered close to 5, the range studied here was pH 5-10. The content of carboxylic groups in the samples was estimated empirically from the Q-pH curves (i.e. the plots of Q--the sample charge determined by establishing the charge balance at each point of the titration curve--against pH), as the value of Q at pH=8, and the content of phenolic groups as twice the difference between Q at pH=10 and pH=8 (Ritchie and Perdue 2003). The Q-pH curves were constructed with data obtained in the potentiometric titrations of the samples.

Other characterisations

For FT-IR determinations, a ground sample was placed onto the Ge window of a Nicolet 5700 FT-IR with an ATR attachment (Onmi sampler nexus). Spectra were obtained over 256 scans with a KBr beam splitter, set at a resolution of 4 [cm.sup.-1], covering the range of 4000-700 [cm.sup.-1]. The reflectance was measured and analysed by using of OMNIC v7.1 software, with Happ-Genzel apodisation and Mertz phase correction.

Solid-state [sup.13]C NMR spectra were acquired with magic angle spinning (MAS) at a [sup.13]C frequency of 100.6 MHz on a Varian Unity INOVA 400 spectrometer. Samples were packed in a 7-mm-diameter cylindrical zirconia rotor with Kel-F end-caps, and spun at 6500 [+ or -] 100 Hz in a Doty Scientific supersonic MAS probe. Cross-polarisation (CP) spectra were acquired using a ramped-amplitude cross-polarisation (CP-ramp) pulse sequence, in which the [sup.1]H spin lock power was varied linearly during the contact time. A 1-ms contact time was used and 4000 transients were collected for each spectrum. Recycle delays (1 s for biochar and 2s for hydrochar) were chosen to be >5x [T.sub.I]H, as determined in preliminary inversion-recovery experiments. Direct polarisation (DP) experiments were carried out using a 90-s recycle delay, and 612 (biochar) and 724 (hydrochar) transients were obtained. For both CP and DP spectra, free induction decays were acquired with a sweep width of 50 kHz; 1216 data points were obtained over an acquisition time of 12 ms. All spectra were zero-filled to 8192 data points and processed with a 100-Hz Lorentzian line broadening and a 0.01-s Gaussian broadening. Chemical shifts were externally referenced to the methyl resonance of hexamethylbenzene at 17.36ppm.

Spin counting experiments were performed using the method of Smemik and Oades (Smemik and Oades 2000a, 2000b). Glycine (AR grade, Ajax Chemicals) was used as an external intensity standard (i.e. the glycine spectrum was obtained separately from those of the samples). For CP spin counting experiments, differences in spin dynamics between the sample and the glycine standard were accounted for by the method of Smernik and Oades (Smemik and Oades 2000a), except that a variable spin lock (VSL) rather than a variable contact time (VCT) experiment was used to determine [T.sub.1p]H (Smemik et al. 2002b). Errors in carbon NMR observabilities ([C.sub.obs] values) are estimated to be [+ or -] 10% in [C.sub.obs]-CP and [+ or -]15% in [C.sub.obs]-DP (Smemik and Oades 2000a).

Scanning electron microscope (SEM) images were obtained with Quanta 200 equipment (FEI, Eindhoven, the Netherlands) after coating the particles with gold with a Bal Tec SCD 500 cool sputting device (Balzers Union, Wallruf, Germany). The Raman spectra were obtained with a Horiva (LabRam HR-800) spectrometer. The source of radiation was a laser operating at a wavelength of 514 nm and a power of 25 mW. Surface analysis of both chars was conducted by X-ray photoelectron spectroscopy (XPS). This was carried out with a Specs spectrometer, with MgK[alpha] (1253.6 eV) radiation emitted from a double anode at 50 W. Binding energies for the high-resolution spectra were calibrated by setting C to 1 s at 284.6 eV. A nonlinear least-squares curve fit, with a Gaussian-Lorentzian mix function and Shirley background subtraction, was used to deconvolute the XPS spectra. Nitrogen gas adsorption was measured with a Micromeritics ASAP 2020 volumetric adsorption system.

Results and discussion

Chemical properties of the carbonised materials

The ash content of the 2 types of char differed widely (Table 1). Pyrolysis produced an increase in the ash content relative to the original feedstock (from 2.8 to 10.8%), whereas a slight decrease was observed after HTC (from 2.8 to 2.1%). This indicates that, while the biochar retains all the mineral matter present in the original feedstock, the hydrochar does not. The latter is attributed to solubilisation of some of the inorganic fraction during the HTC process and is caused by the acidic nature of the aqueous products released. The pH of samples was also very different, 9.89 and 4.70, for the biochar and the hydrochar, respectively. The alkalinity of the biochar provided a liming equivalence of 39.6kg CaC[O.sub.3]/t. The results are consistent with previous reports, in that pyrolysis produces charcoals with pH values generally above neutrality, and generally higher at increasing temperature of pyrolysis (Ueno et al. 2007) and when ash-rich feedstock is used. Acidic pH values have been reported for biochars from pine feedstock pyrolysed at temperatures <400[degrees]C (Calvelo-Pereira et al. 2010).

As expected, the C content in the solid fraction increased after the different carbonisation processes were applied to the corn stover feedstock (C 42.9% wt); the increase was greater with pyrolysis (C 74.3% wt) than with HTC (C 67.8% wt) (Table 1). On the other hand, product yield and the amount of C recovered (percentage of C originally contained in the raw material that is retained in the carbonised sample) were higher in the hydrochar than in the biochar (Table 1), with product yields of 36 and 28% wt, respectively, and recovered C values of 57 and 46% wt, respectively. The same corn stover feedstock, pyrolised at 350 and 400[degrees]C in the same rotator kiln, produced biochars with C concentrations of 64.5 and 69.2% wt (Camps Arbestain 2010). It can therefore be inferred that the recovery of C with HTC was similar to that obtained with pyrolysis at temperatures slightly <400[degrees]C.

Total N content was lower in the hydrochar than in the biochar (0.65 and 0.78%, respectively), while the opposite was observed for H and O (Table 1), suggesting a greater degree of condensation of the latter. This resulted in higher O/C and H/C ratios in the hydrochar than in the biochar (Table 1). This trend was also observed when the hydrochar was compared with the biochars produced at 350 and 400[degrees]C (Camps Arbestain 2010). The variation in the elemental composition of the materials (from biomass to char) was analysed with the aid of the Van Krevelen diagram, as illustrated in Fig. 1 (Van Krevelen 1950), which includes plots of H/C v. O/C atomic ratios. Both processes essentially followed dehydration--decarboxylation reactions, which were more intense with pyrolysis than with HTC. The H/C-O/C ratios for other substances (wood, cellulose, lignite, peat, and different types of coals) were plotted in the same figure for purposes of comparison. The position of the hydrochar on this plot was similar to that of lignite, whereas the biochar occupied a similar position to bituminous coal.


The concentrations of carboxylic and phenolic groups, as estimated by potentiometric titration, were 0.04 and 0.30 mol/kg, respectively, for the biochar, and 0.07 and 0.37 mol/kg for the hydrochar. These data thus indicate that, although present in low amounts, the carboxylic groups in the hydrochar were almost twice as abundant as those in the biochar. However, this does not explain the substantial differences in the O/C atomic ratios of the 2 samples (0.27 in hydrochar and 0.12 in biochar, Table 1). One plausible explanation for the higher O/C ratio in the hydrochar than in the biochar is the presence of stable groups (e.g. ether, quinone, pyrone), which are less reactive than the hydrophilic groups (e.g. hydroxyl, carboxyl, ester). The ECEC values of the biochar and hydrochar samples measured at pH 7.54 and 4.90, respectively, were 193 and 119mmol(+)/kg, and CEC values at pH 7 were 284 and 134mmol(+)/kg. The results thus indicate a higher potential and effective CEC of the biochar than the hydrochar, although the amounts of carboxylic groups in the 2 samples--estimated by acid-base titrations (Table 1)--do not reflect the same trend. The results of XPS and NMR, described below, confirm the greater presence of carboxylic functional groups in the hydrochar than in the biochar. The lack of consistency between the number of acidic functional groups and CEC values may be attributed to the fact that, whereas for the former determination, the biochar was dialysed to eliminate salts, for the latter, the biochar was only rinsed with abundant distilled water. Therefore, the CEC of the biochar may also include a contribution from the inorganic fraction. In addition, the different pH values of the chars may also have contributed to the observed differences in ECEC, as this property is determined by use of an unbuffered solution. Current measurements for the assessment of CEC in chars were originally developed for soils and may not be always applicable for charcoal characterisation. Efforts must be made to adapt these methodologies to the analysis of chars.

XPS, FTIR, [sup.13]C-MAS NMR, and Raman spectra corresponding to the carbonised materials

The oxygen functionalities present at the outer surface of the char particles were analysed by XPS (Watts and Wolstenholme 2003). The C 1s core level spectrum obtained, with the peakfitting of its envelope, is shown in Fig. 2. For both materials, the C 1s envelope included 2 common signals attributed, respectively, to aliphatic/aromatic carbon groups ([CH.sub.x], C-C/ C=C) (284.6 eV) and carboxylic groups, esters, or lactones (-COOR) (289 eV). The former peak was higher for the biochar sample and the latter peak was higher for the hydrochar sample. This is consistent with the presence of a more condensed structure in the biochar and more oxygen-rich functionalities in the hydrochar, as indicated above. In both spectra there was a third signal, corresponding to a hydroxyl group (-C-OR), which appeared at 285.7-286.2 eV in the biochar and at 286.6 eV in the hydrochar. The observed displacement of the latter may be attributed to the presence of hydroxyl groups from linear alcohols or ethers, or phenols with a low degree of condensation. In addition, the biochar sample presented a small fourth peak at 287.2 eV, corresponding to carbonyl groups (>C=O).


The primary differences in the FTIR spectra for the biochar and the hydrochar samples are encountered in the higher number of oxygen bonds in the latter (Fig. 3). Peaks at ~1700, 1210, and 1110 [cm.sup.-1] were observed in the hydrochar but not in the biochar (except for a small band at 1110[cm.sup.-1]). The first peak can be attributed to the stretching of C=O bonds, which are present in ketones, aldehydes, quinone, esters, and carboxylic acid functional groups (Koch et al. 1998; Pradhan and Sandle 1998); the second and third peaks are attributed to the stretching of C-O bonds (carboxyl, ester, and ether groups) and OH deformations of carboxyl-C (Pradhan and Sandle 1998). A band at ~1400[cm.sup.-1] corresponding to C-H bending was identified in both chars (Smith and Chughtai 1995). Vibration bands at 1600 [cm.sup.-1], corresponding to the stretching of C=C groups, were also observed in both samples, reflecting their aromatic nature (Guo and Bustin 1998). The band at 1513 [cm.sup.-1] was only detected in the hydrochar and is attributed to C-H or N-H bending, characteristic of undecomposed litter (Haberhauer et al. 1998). According to those authors, the intensity of the peaks at 1510, 1450, 1370, and 1270 [cm.sup.-1] may serve as a method of assessing the degree of organic matter degradation. All of these peaks were evident in the hydrochar spectrum, but not in the biochar spectrum, which is consistent with the findings of Haberhauer et al. (1998). Finally, the biochar sample produced a band at 875 [cm.sup.-1] attributed to aromatic C-H out-of-plane bending (Ibarra et al. 1996), which indicates the greater degree of aromaticity of this sample.


Solid-state [sup.13]C NMR spectra, acquired by both CP and DP techniques, indicate that the biochar is predominantly aromatic whereas the hydrochar contains a range of C types (Fig. 4). The spectra of the biochar are dominated by a single peak at ~127ppm that can be attributed to C- and H-substituted aromatic C. The next largest peaks in the spectra (marked *) are also related to this peak; they are spinning sidebands (SSBs) that occur when the rate of magic angle spinning is insufficient compared to the chemical shift anisotropy of the signal. The only non-aromatic signals detected for the biochar were therefore 2 small peaks in the alkyl region (more visible in the CP than the DP spectrum) at ~20 ppm and ~40 ppm.

The hydrochar spectra also contain a strong peak in the aromatic region at ~129ppm, along with a sharp peak at ~147 ppm, which can be assigned to O-substituted aromatic C, probably derived from lignin. Two dominant peaks are visible in the alkyl region of the hydrochar spectra. The sharp peak at ~56ppm is probably due to methoxyl C, which is also commonly associated with lignin structures. The broad peak at ~35 ppm is due to C- and H-substituted alkyl C. The small peak at ~30ppm indicates the presence of long-chain polymethylene C, such as found in lipids. However, most of the alkyl C is not of this long-chain type, but rather a complex mixture of short and/or branched chains. In the high ppm region, the hydrochar has a small peak at ~175ppm due to carboxyl C. Most of the signal of the hydrochar at ~190ppm is a SSB due to aromatic C. The peak at ~208 ppm is attributed to ketone and aldehyde C. Overall,-although the biochar spectra reflect the presence of an almost unique aromatic peak, which could be attributed to the presence of char, the hydrochar spectra most closely resembles acid hydrolysis residues and low-grade coals.


The DP spectra were acquired because char carbon is often under-detected in CP spectra (Smemik et al. 2002a, 2002b). We used spin counting (Smemik and Oades 2000a, 2000b) to determine how much potential signal was actually detected ([C.sub.obs]) for both samples and both techniques (CP and DP). Approximately 100% (101% for the biochar and 92% for the hydrochar) of the potential signal was detected using the DP technique, which indicates that these spectra are quantitatively reliable. On the other hand, with CP, only 46% of potential signal was detected for the biochar and 77% for hydrochar. In the case of the biochar, this did not affect the distribution of the signal because almost all of the C was aromatic and equally poorly detected. In the case of the hydrochar, the greater relative size of the alkyl signal in the CP spectrum may be attributed to under-detection of the aromatic signal when CP was used.



The presence of aromatic C was also confirmed by Raman spectroscopy (Fig. 5). These samples exhibit the typical spectra of non-graphitic carbon materials with 2 main peaks: D (around 1361 [cm.sup.-1]) and G (around 1591 [cm.sup.-1]) (Cuesta et al. 1994; Sheng 2007). The vibration mode of the D peak corresponds to a disordered graphitic lattice (graphene layer edges), which for the G peak corresponds to an ideal graphitic lattice. Both peaks therefore reveal the presence of C [sp.sup.2] atoms in benzene or condensed benzene rings in amorphous (partially hydrogenated) carbon, indicating the presence of small aromatic clusters in both samples. As can be observed, both spectra are quite similar.

Overall, the different spectra obtained for each type of char confirmed the results inferred from the elemental analysis of the samples, that is, the greater aromaticity of the biochar sample (produced at 550[degrees]C) than the hydrochar sample, and the greater potential reactivity of the hydrochar relative to the biochar (if the inorganic fraction is excluded). These different properties will have clear implications for the long-term stability of these charcoals in soils. However, care should be taken when generalising from the present results, as the aromaticity of the biochars will depend, to a great extent, on the temperature of pyrolysis (Keiluweit et al. 2010). Biochar produced in our laboratory, from the same corn stover feedstock and at pyrolysis temperatures of 350 and 400[degrees]C, showed a smaller alkyl band in the NMR spectra than that of the hydrochar (Camps Arbestain 2010), thus suggesting that the stability of the hydrochar is lower than that of the biochar produced at 350[degrees]C.

Structural properties of the carbonised materials

Figure 6 shows the SEM images obtained for the corn stover feedstock, and the biochar and hydrochar produced from this material. Both processes caused substantial changes in the surface morphology of the original particles, although they largely retained their macroscopic shape. Major changes were observed at the surface of the hydrochar particles, where sphere-like microparticles adhered to the surface of larger particles (Fig. 6c). These sphere-like microparticles are mainly formed by degradation of the cellulose component during HTC and its subsequent precipitation and growth as spheres (Sevilla and Fuertes 2009a). The diameter of the spherical particles is in the same range as observed by Sevilla and Fuertes (2009b) (0.4-7.0 [micro]m). On the other hand, the lignin component probably suffered only partial degradation (owing to its greater thermal stability), causing a rough texture, so that the original shape of the particles was largely preserved. The biochar sample retained the original macrocellular morphology of the corn stover and, unlike the surface of the hydrochar, that of biochar particles appeared smooth (Fig. 6b).


The nitrogen isotherms obtained at 77 K (Fig. 7) provide the specific surface areas of the samples by applying the BET equation. A type II isotherm, which is typical of non-porous materials, was obtained for both samples. Thus, both samples present relatively small surface areas due to the lack of porosity. Under such circumstances, the specific surface area values calculated only correspond to the external surface. Thus, the surface areas of the BET are 12 and 4[m.sup.2]/g for biochar and hydrochar samples, respectively. However, pyrolysis of biomass at temperatures >400[degrees]C generally leads to rather porous chars and therefore larger surface areas (Keiluweit et al. 2010). This is the case for several wood species, which generate surface areas of up to 350 [m.sup.2]/g after pyrolysis at 500[degrees]C (Lehmann 2007), and for paper mill waste pyrolised at 550[degrees]C, with a final surface area of 115[m.sup.2]/g (Van Zwieten et al. 2009). The porosity of hydrochar, on the other hand, has been reported to be low (Sevilla and Fuertes 2009a, 2009b).


The properties of the chars obtained from 2 distinct carbonisation processes, pyrolysis at 550[degrees]C (to produce biochar) and HTC (to produce hydrochar), were very different. The hydrochar had low ash content and low pH value (4.7), and recovery of C was high, although only about half of the C was aromatic. Hydrochar had high atomic O/C and H/C ratios, mainly attributed to the presence of groups such as ether, quinone, and pyrone, although more reactive groups (e.g. carboxylic, ester, lactones) were also identified and were more abundant than in the biochar. Overall, the characteristics of the hydrochar resembled those of low-grade coals. The biochar, on the other hand, had a higher ash content than the hydrochar and a high pH (~10), which provided the sample with a lime equivalence of ~40kg CaC[O.sub.3]/t. The C recovery was lower than that of the hydrochar, but the C was mostly in the aromatic form, which explains the lower O/C and H/C ratios. The surface area of both samples was very small, although pyrolysis generally produces rather porous chars. The measured CEC values were inconsistent with the concentration of functional groups determined using other techniques (probably due to the presence of ash in the biochar), and suggested the need to adapt CEC methodologies, originally developed for soil measurements, to charcoal characterisation. Both chars could be used as soil amendments for very different requirements. The choice of one or other method of production may be determined by the drying needs of the feedstock (e.g. the drying needs for very wet starting products could be avoided with the HTC technique). Further studies on soil responses and the residence times of the chars (especially the hydrochar) are required in order to pursue long-term C sequestration. In addition, a comparative study of the energy balances is required to ensure that the processes are beneficial in terms of GHG production.



We acknowledge the Manawatu Microscopy and Imaging Centre (MMIC) and Doug Hopcroft for assistance in preparing the samples and operating the SEM images. We are grateful to Kina Hira for assistance in the FTIR measurements. M. S. and J. A. M.-A. acknowledge the assistance of the Spanish MCyT for their award of a Postdoctoral Mobility contract and a Juan de la Cierva contract, respectively. M.C.A. is very grateful for financial support from the Ministry of Agriculture and Forestry of New Zealand. The authors thank the anonymous reviewers for the helpful comments and suggestions provided.

Manuscript received 5 January 2010, accepted 10 May 2010


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A. B. Fuertes (A), M. Camps Arbestain (B,F), M. Sevilla (A), J. A. Macia-Agullo (A), S. Fiol (C), R. Lopez (C), R. J. Smernik (D), W. P. Aitkenhead (B), F. Arce (C), and F. Macias (E)

(A) Instituto Nacional del Carbon (CSIC), Apartado 73, 33080-Oviedo, Spain.

(B)New Zealand Biochar Research Centre, Private Bag 11222, Massey University, Palmerston North 4442, New Zealand.

(C)Departamento de Quimica Fisica, Facultad de Quimica, Universidad de Santiago de Compostela, 15782-Santiago de Compostela, Spain.

(D) School of Agriculture, Food and Wine, The University of Adelaide, Waite Campus, Urrbrae, SA 5064, Australia.

(E) Departamento de Edafologia y Quimica Agricola, Facultad de Biologia, Universidad de Santiago de Compostela, 15782-Santiago de Compostela, Spain.

(F) Corresponding author. Email:
Table 1. Chemical characteristics of biochar and hydrochar products

CEC, Cation exchange capacity (pH 7); ECEC, effective CEC

Sample                  Chemical composition (wt %)

                 C        H        O        N

Corn stover     42.9     5.8      41.7     0.22
Hydrochar       67.8     5.3      16.2     0.65
Biochar         74.3     2.7       8.0     0.78

Sample                  Atomic ratio (A)

                Ash      O/C      H/C

Corn stover     2.8      0.84     1.62
Hydrochar       2.1      0.27     0.94
Biochar        10.8      0.12     0.44

                Yield    Recovered        pH       (mol/kg)
               (%) (B)    (%) (C)     [H.sub.2]O    -COOH

Corn stover      --          --           --         --
Hydrochar        36          57          4.70       0.07
Biochar          28          46          9.89       0.04

                (mol/kg)    ECEC     CEC
               Phenolic     (mmol(+)/kg)

Corn stover        --        --       --
Hydrochar        0.37       119      134
Biochar          0.30       193      284

(A) Dry ash free basis.

(B) Yield expressed as (g product/l00g raw material).

(C) Percentage of carbon originally contained in the raw material
that is retained in the carbonised sample.
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Author:Fuertes, A.B.; Arbestain, M. Camps; Sevilla, M.; Macia-Agullo, J.A.; Fiol, S.; Lopez, R.; Smernik, R
Publication:Australian Journal of Soil Research
Article Type:Report
Geographic Code:8AUST
Date:Sep 1, 2010
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