Producing biochars with enhanced surface activity through alkaline pretreatment of feedstocks.
Biochar is produced by pyrolysis of natural organic materials (e.g. agricultural and forest wastes, municipal waste, or manure) under oxygen-limited conditions. Energy from biomass can also be obtained during the production of biochar (Bridgwater 2003). According to the International Biochar Initiative (www.biochar-international.org), the final use of biochar should be for application to soils to achieve an agronomic and/ or environmental benefit. Due to its predominantly aromatic composition and heterogeneous, condensed macromolecular structure, biochar is chemically and biologically more stable than its original carbon sources. This makes biochar difficult to breakdown, remaining stable in soils potentially for hundreds to thousands of years (Krull et al. 2006). Because of the stability of biochar against chemical and biological degradation, its production represents a promising technology for mitigating greenhouse gas emissions (Lehmann 2007). Process conditions that most affect biochar properties during pyrolysis are the heating rate, final temperature, residence time, and the treatment atmosphere (Emmerich and Luengo 1996).
While fresh biochar generally has a low cation exchange capacity (CEC) (Cheng et al. 2006), biochar weathered in soils develops a high CEC through biotic and abiotic oxidative processes (Lehmann 2007; Cheng et al. 2008). Laboratory enhancement of surface charge on fresh biochars has been achieved after heating (70[degrees]C) for several months (Cheng et al. 2006). However, development of cost-effective biochar products with both agronomic and environmental benefits requires the development of more feasible and environmentally friendly techniques for increasing CEC.
Current knowledge gained from activated carbon studies could have some application to biochar research. Surface charges on activated carbon are generated by physical and chemical methods (Rodriguez-Reinoso and Molina-Sabio 1992). Physical activation involves the use of oxidising gases, such as carbon dioxide or water steam, while chemical activation involves the use of inorganic chemicals, such as zinc chloride, phosphoric acid, and potassium or sodium hydroxides (Linares-Solano et al. 2008). Sodium hydroxide has been reported to promote a better activation of lignocellulosic materials than potassium hydroxide (Lillo-Rodenas et al. 2007), and this occurs predominantly via a non-intercalation mechanism in the former and through an intercalation process in the latter (Raymundo-Pinero et al. 2005; Lozano-Castello et al. 2006; Macia-Agullo et al. 2007).
Bark from pulp mills is a potential feedstock for the production of biochar, because bark is a low-cost by-product that is produced in large quantities (Montane et al. 2005). The most common exotic forest species used in New Zealand pulp mills is radiata pine (Pinus ratiata), followed by Douglas fir (Pseudotsuga menziesii) and eucalyptus (Eucalyptus cinerea). Softwood and hardwood species have different types of lignin, with softwood (e.g. pine) having lower methoxyl content than hardwood (e.g. eucalyptus). The lignin of softwood (gymnosperms) displays a higher proportion of guaiacyl (G) units and no syringyl (S) units, whereas the lignin of hardwoods (angiosperms) has both G and S units, the latter unit being dominant (Lewis and Yamamoto 1990; Liu et al. 2008). These differences are known to affect the likelihood of internal crosslinking, otherwise hampered by methoxyl groups in S units, and result in a variable reactivity of wood towards alkaline reagents (Tsutsumi et al. 1995) and thermal degradation during pyrolysis (Wang et al. 2009).
The aim of this study was to produce biochars from pine and eucalyptus feedstocks with a high surface charge, using alkaline tannery waste as activating agent. The effect of different tannery waste dilution was assessed, together with the potential distinct response of the 2 types of wood. The effectiveness of the different treatments with regard to nutrient retention was also evaluated through an N[H.sub.4.sup.+] sorption-desorption study. It is proposed that the end-use for these activated biochars could be as active filter media for removing nutrients from housed animal and municipal waste streams, and their addition to soils thereafter as slow-release ferfilisers.
Materials and methods
Pine and eucalyptus waste barks were supplied by Carter Holt Harvey NZ Ltd These wastes were not bark sensu stricto, as for eucalyptus, the outer cambium layer was dominant, and for pine, high levels of wood were present. However, these materials will be referred to as 'bark' throughout the text to facilitate reading. The pine bark was chipped with a commercial chipper and the eucalyptus bark peelings were used in the received form. The final particle size of the pine chips was irregular, ranging from 3 to 11 mm in length. In order to minimise processing, no efforts were made to homogenise the size of eucalyptus bark particles. The feedstocks were dried overnight in an oven at 65[degrees]C.
Tannery lime float waste slurry was used as an alkaline treatment of the bark feedstocks. This slurry was produced by the tanning industry as a waste from a washing treatment of livestock skins and hides, in which a concentrated solution of [Na.sub.2]S[O.sub.4] and [Na.sub.2]S is used. The waste consists of hydrolysed proteins and fats, in addition to the original compounds, and is alkaline (pH 13) with a dry matter content of 30%. The composition of the tannery waste is reported in Table 1.
Chemical activation treatments were carried out through impregnation of the feedstocks with either diluted or undiluted alkaline tannery slurry. The first set of treatments involved impregnating 200g (dry weight basis) of both feedstocks with diluted tannery slurry (diluted with distilled water in a 1 : 3 ratio). The impregnation was based on adding to the bark the amount of slurry needed to fill the adsorbent pore volume (Bandosz and Petit 2009); 40 g (dry weight basis) of diluted tannery slurry was needed to impregnate the pine wood (L-PI feedstock) and 60 g (dry weight basis) was required for the eucalyptus bark (L-EU feedstock). The second set of treatments involved impregnating 200g (dry weight basis) of both feedstocks with undiluted tannery slurry (60 g and 40 g for S-EU and S-PI, respectively). Impregnation of feedstocks with both the diluted and undiluted tannery slurry was carried out at room temperature for 5 h, before feedstock/tannery slurry mixtures were dried overnight in an oven at 65[degrees]C and then pyrolysed, as described below. Pine and eucalyptus biochars with no tannery slurry treatment were also produced and are referred to as controls treatments (Ctr-PI and Ctr-EU feedstocks, respectively). Three replicates were made of each treatment.
For each of PI and EU bark feedstocks, 200 g, either with or without alkaline treatment, was pyrolysed at 550[degrees]C by heating at an average heating rate of 28[degrees]C/min for an average time of 20 min, for all treatments, in triplicate, using a gas-fired, 5-L, stainless-steel, rotating drum kiln. When the desired temperature was reached, the kiln was allowed to cool to room temperature. The mixed char and ash residue was removed and another 2 batches of each feedstock were pyrolysed. The 3 replicate mixed char and ash residues of each treatment were pooled into a single sample. Before characterisation of the resulting biochar, subsamples of the mixed char and ash residue were washed repeatedly with deionised water until the electrical conductivity decreased to 50 dS/m. The final pH of the washing solution for the washed biochar materials ranged between 7.0 and 9.5. These washed samples were used for the N[H.sub.4.sup.+] adsorption-desorption experiments and BET measurements. Dominant particle-size was 3-8 mm for the pine biochars and <2 mm for the eucalyptus biochars.
Cellulose, hemicellulose, and lignin content
The cellulose, hemicellulose, and lignin content of the untreated bark feedstocks were determined using the Fibretec System M (Tecator, Hoganas, Sweden). This method is based on sequential chemical treatments with neutral detergent, acid detergent, 72% [H.sub.2]S[O.sub.4], and ashing (Van Soest 1967). The detergent neutral step washes out the cellular content and the residual fraction is referred to as neutral detergent fibres (NDF). With the acid detergent treatment of NDF, cell walls are broken down and the residual fraction is referred to as acid detergent fibres (ADF). Hemicellulose is estimated as NDF-ADF. With a subsequent [H.sub.2]S[O.sub.4] treatment, cell walls are digested, and an acid detergent lignin (ADL) residue is obtained. Cellulose is estimated as ADF-ADL, and ADL is assumed to be mostly lignin.
Total C, H, and N contents of the different biochars were determined using a TmSpec CHNS analyser (LECO Corp. St. Joseph, MI); total O was determined with an EA elemental analyser (Fisons EA-1108-CHNS-O, Fisons Instruments, Milano, Italy). The C-C[O.sub.3] content of biochars was determined using a modification of common static chamber methods (Bundy and Bremner 1972; Tiessen et al. 1983). Organic C was determined as the difference between total C and C-C[O.sub.3]. The ash content was determined by thermo gravimetric analysis on a TA instrument (Alphatech, SDT Q600 manufactured by TA instruments, Newcastle, Australia). The sample was initially heated from room temperature to 900[degrees]C (at a rate of 5[degrees]C/min) under an [N.sub.2] atmosphere and weight loss recorded (data not shown); thereafter, an air current was provided and the ash was determined when there was no further weight change. Ashes do not contain carbonates, which decarboxylate at temperatures <900[degrees]C.
pH and surface acid functional groups
Biochar pH was measured using the methodology of Ahmedna et al. (2000), which involved a 1% (wt/wt) suspension of biochar in deionised water. The suspension was heated in a water bath to -90[degrees]C and stirred for 20 min to allow dissolution of the soluble biochar components. After cooling to room temperature, the pH of the biochar suspension was measured. The content of acid groups on the biochar was determined following potentiometric titration (Lopez et al. 2008; Petit et al. 2010). This method involves the suspension of 0.1 g of sample (previously acidified and subsequently dialysed in cellulose bags, pore size 1000 Da, and lyophilised, to eliminate interfering salts) in 50 mL of 0.1 M KNO3 as the inert electrolyte. Thereafter, biochar suspensions were potentiometrically forward titrated with 0.2M KOH carbonate-free solution or backward titrated with 0.1 M HC1 in order to cover the pH range 3-10. However, pH values of the biochars were buffered close to pH 5; thus with this methodology, functional groups reactive in the pH range 5-10 were estimated. The contents of carboxylic groups in the samples were estimated empirically from the Q (the sample charge) v. pH charge curves as the value of Q at pH 8. The Q-pH curves were obtained from experimental data points of the potentiometric titrations of the samples.
Biochars were ground to 0.1 mm, and 0.5 mg of each sample was placed onto the Ge window of a Nicolet 5700 FTIR with an ATR attachment (OMNI Sampler Nexus). Spectra were obtained over 256 scans with a KBr beam splitter. It was set at a resolution of 4[cm.sup.-1], covering the range of 4000-700 [cm.sup.-1] and with an aperture size of 34cm. The reflectance was measured and analysed using OMNIC v7.1 with Happ-Genzel apodisation and Mertz phase correction. The identification of absorption bands was based on published data and is described in the Appendix.
Solid-state CP MAS [sup.13]C NMR
Solid-state, cross-polarisation magic-angle spinning [sup.13]C nuclear magnetic resonance (CPMAS [sup.13]C NMR) was used to characterise the biochar. All NMR experiments were conducted in the Bruker (Rheinstetten, Germany) AMX 200Mhz horizontal bore magnet. Samples were packed into a 7-mm-i.d. rotor and spun at speeds of 5 kHz in a dual-resonance magnetic angle spinning (MAS) probe from Doty Scientific. The [sup.13]C CP/MAS spectra were acquired with a 1H 90[degrees] pulse for 5.5 [mu], a cross-polarisation contact time of 1000 [mu], an acquisition time of 30 ms, relaxation time of 2 s, and 5000 scans.
X-ray photoelectron spectroscopy
Surface analysis of the eucalyptus samples was conducted using X-ray photoelectron spectroscopy (XPS). This was carried out using a Specs spectrometer, using MgK[alpha] (1253.6 eV) radiation emitted from a double anode at 50 W. Binding energies for the high-resolution spectra (C 1s and N 1s) were calibrated by setting C 1s at 284.6eV. A non-linear least-squares curve fitting with a Gaussian-Lorentzian mix function and Shirley background subtraction was used to deconvolute the XPS spectra.
[FIGURE 1 OMITTED]
Specific surface area (BET)
Nitrogen gas adsorption measurements for BET measurements were performed using a Micromeritics ASAP 2020 volumetric adsorption system. Samples were previously outgassed at 250[degrees]C for 4h.
Scanning electron microscope (SEM)
The surface physical morphology of the samples was examined by Quanta 200 equipment (FEI, Eindhoven, the Netherlands) after coating the particles with gold using a Bal Tec SCD 500 cool sputting device (Balzers Union, Wallruf, Germany).
Ammonium sorption and desorption
A 10-mL volume of N[H.sub.4.sup.+]-N solution, 40 mg N/L as ([N[H.sub.4]).sub.2] S[O.sub.4], was added to 2 g of biochar placed in a 40-mL centrifuge tube and run in triplicate. The N[H.sub.4.sup.+]-N solution concentration used was based on the typical levels present in municipal waste water. The tubes were shaken end-over-end (30 r.p.m.) for 6 h, then centrifuged for 5 min at 7000 r.p.m., and the suspension was filtered (Whatman filter paper 42). This procedure was repeated 4 times with each sample. Ammonium was determined using a Technicon autoanalyser (Technicon, Dublin). The sorbed concentration was estimated by difference between the initial and final concentration of the N[H.sub.4.sup.+]-N. Subsequently, biochars were subjected to desorption of N[H.sub.4.sup.+]-N by adding the samples to 2 separate 10-mL volumes of 2 M KCl solution.
[FIGURE 2 OMITTED]
The significance of results for the adsorption experiment was determined using GraphPad statistical software (GraphPad Prism 5.02 GraphPad Software Inc.). The Student's t-test was used to test for significant differences in the results of N[H.sub.4.sup.+] sorption.
Results and discussion
Composition of the feedstocks used
The cellulose content was higher in the eucalyptus bark (51.3%) than the pine bark (39.7%), whereas the lignin and hemicellulose content were higher in pine bark (26.3 and 17.7%, respectively) than in eucalyptus bark (10.5 and 15.6%, respectively) (Table 2). The NDF values of the original feedstocks were 77 and 84% for eucalyptus and pine samples, respectively, which indicates a greater presence of readily degradable cellular contents (RDCC, estimated as 100--NDF) in the eucalyptus bark (Table 2).
Yield, recovered C, and N, C and N contents
The yield of the different biochars, defined as the ratio of mass of biochar recovered after pyrolysis and the initial mass of the feedstock (or feedstock + tannery slurry) as a percentage, is reported in Table 3. The yield of the 3 pine biochars (Ctr-PI, L-PI, S-PI) ranged from 26 to 27%, whereas the yield for the 3 eucalyptus biochars (Ctr-EU, L-EU, S-EU) ranged from 28 to 32% (Table 3). These yields are considerably greater that the values of ~5% reported by Lillo-Rodenas et al. (2007), which resulted from the chemical activation of eucalyptus feedstocks with NaOH.
As expected from a carbonisation process, the C content of pine increased from 49 to 78%, and that of eucalyptus from 46 to 74% (Table 3). Carbon contents of treated biochars included a C-C[O.sub.3] fraction (<3%) attributed to the formation of carbonates during pyrolysis, as evolved C[O.sub.2] became trapped under alkaline conditions. Organic C contents of the treated biochars were lower than the untreated biochars, with values of 70 and 65% for pine, respectively, and of 56 and 51% for eucalyptus, respectively (Table 3). This lower organic C content could be attributed to the dilution effect caused by the addition of the tannery slurry, as it had a lower C content than the feedstocks (Tables 1, 3). Recovered C ranged between 41 and 43% in the pine biochars and between 42 and 45% in the eucalyptus biochars (Table 3). Biochars treated with diluted tannery slurry had higher H/C ratios than the rest of the samples, this trend being more evident for eucalyptus (data not shown). Pretreating the feedstocks also increased the ash content of the corresponding biochars, as expected (Table 3).
[FIGURE 3 OMITTED]
The carbonisation process of untreated feedstocks increased the concentration of N in the materials from ~0.3% N before pyrolysis, to ~0.6% N, after pyrolysis. The total recovery of N was higher for pine than eucalyptus (57% v. 41% recovery of N, respectively) (Table 3). The tannery waste treatments increased the N content of both pine and eucalyptus feedstocks (up to 1.8% and 3.2%, respectively; Table 3), as expected, due to the richness of N in the raw wastes (Table 1). After pyrolysis, the N contents of the materials either slightly increased or decreased, with 2.0 and 2.1% N for the S-PI and S-EU biochars, respectively. The recovery of N in the treated biochars was lower than the untreated biochars, with values ranging from 16 to 30%, thus indicating a greater N loss from the system, compared with untreated biochars (Table 3). Further research is required to determine the fate of the unrecovered N.
Values of pH and surface charge
The pH values of the biochars ranged from 8.8 for the Ctr-EU biochar up to 10.6 for the S-EU biochar (Table 4). Biochars prepared from alkaline tannery slurry had greater pH values than the corresponding controls. Carboxylic functional groups were more abundant in the Ctr-EU treatment than the Ctr-PI treatment. The tannery slurry treatments promoted the formation of surface charge in all samples under study, but the response to increasing alkaline concentrations was feedstock-dependent. In the biochars from pine wastes, there was an increase of carboxylic functional groups with increasing concentration of tannery slurry (from 0.006 mmol/g in the Ctr-PI treatment to 0.080 mmol/g in the S-PI treatment). In the eucalyptus samples, however, the greatest amount of carboxylic groups was generated with the diluted tannery slurry treatment (L-EU, 0.164 mmol/g), which was the highest value of all biochars in this study. The decrease in functional groups observed in the S-EU treatment could be due to an enhanced degradability of fibrous structure of the eucalyptus bark at high alkalinity. Differences between the 2 feedstocks were mainly attributed to their different types of lignin (David and Shiraishi 2000).
FT-IR, NMR, and XPS spectra
The FT-IR spectra of the 2 original bark feedstocks (Fig. 1a, b) were similar, although differences were evident when the bands and their intensities were compared. In the FT-IR spectrum of pine feedstock some typical features of a G lignin type bark (Faix et al. 1988) could be identified: 1600 [cm.sup.-1] <<1508 [cm.sup.-1] dominant >>1459 [cm.sup.-1]; 1270 [cm.sup.-1] >>1223 [cm.sup.-1]; a maximum at 1140 [cm.sup.-1]; 1031 [cm.sup.-1] >1223 [cm.sup.-1]; 2 separate bands at 856 and 815 [cm.sup.-1]. The eucalyptus bark feedstock had lower intensity bands at 1508 [cm.sup.-1] (attributed to aromatic C=C or amide II N vibrations, found in fresh litter; Haberhauer et al. 1998) and 1270 [cm.sup.-1], whereas that at 1600 [cm.sup.-1] (C=C and/or C-O stretching in conjugated carboxyls; see Appendix) was more prominent. In addition, in the eucalyptus bark, a band at 1329 [cm.sup.-1] was evident, indicating C-O stretching vibrations (in relation to syringyl ring; Wang et al. 2009). A single band appeared at 834 [cm.sup.-1], related to C-H vibrations of syringyl units. Finally, the band at ~1750 [cm.sup.-1] was well defined in both species, although was slightly more pronounced in pine bark, and attributed to C=O stretching (Chiang et al. 2000).
After the tannery slurry treatment, the intensity at 3400 [cm.sup.-1] (H bonded to OH groups; see Appendix) decreased in both bark feedstocks (Fig. 1c, d). The spectra of both treated bark feedstocks displayed narrow bands at 2950 [cm.sup.-1] and 2830 [cm.sup.-1], mainly related to C-H stretching of alkyl structures (Gunzler and Bock 1990), and specifically to methyl (C[H.sub.3]) and methylene (C[H.sub.2]) asymmetric stretching, which were both more intense in the eucalyptus than in the pine biochars. This is consistent with the greater amounts of tannery slurry required by the eucalyptus bark to achieve complete impregnation of its fibres. Both bands were also present in the original bark feedstocks, but the intensity was considerably lower (Fig. 1a, b). As in the original bark feedstocks, a band at 1750 [cm.sup.-1], probably related to C=O stretching in esters (Gunzler and Bock 1990), was observed. The strong bands at 1640 and 1550 [cm.sup.-1] were assigned to protein amide I and protein amide II, respectively (Fig. 1c, d; see Appendix). These were associated with the N compounds present in the tannery slurry, and were more intense in the spectra from the treated eucalyptus. Finally, an increase in the intensity of the band at 1100 [cm.sup.-1] was observed, which was again more evident in the biochar made from eucalyptus. The presence of a band at 1100 [cm.sup.-1] in both treated feedstocks is related to C-O stretching vibrations (O-[H.sub.3] and C-OH groups; Pradhan and Sandle 1999; Sharma et al. 2004).
A band at ~1400 [cm.sup.-1] corresponding to C-H bending (Smith and Chughtai 1995) was identified in the spectra of the 2 untreated biochars (Ctr-PI and Ctr-EU; Fig. 2a, b), this band being more pronounced in the case of the eucalyptus. Vibrations at ~1600 [cm.sup.-1], corresponding to C=C stretching (see Appendix), were also observed in both samples. The eucalyptus biochar showed a band at 875 [cm.sup.-1] attributed to aromatic C-H out-of-plane (Ibarra et al. 1996), which suggests greater aromaticity of this sample. The pine biochar also showed an additional band at 1157 [cm.sup.-1], which could be attributed to the stretching of C-O bonds (carboxyl, ester, and ether groups) and OH deformations of carboxyl-C (Sharma et al. 2004).
The L-EU biochar (Fig. 2d) had an FT-IR spectrum similar to that from Ctr-EU (Fig. 2b), except for the band at 1140 [cm.sup.-1], which was also pronounced in the L-PI biochar (Fig. 2c). This could be attributed to stretching of C-O bonds and OH deformation of carboxyl groups, indicating a higher number of oxygen bonds after this treatment in both biochars. Also, a small band at 712 [cm.sup.-1] was detected in the L-EU biochar (Fig. 2d), which suggested the presence of carbonates, as confirmed by XRD (data not shown). Finally, the S-EU and S-PI biochars showed spectra very similar to the above ones (Fig. 2e, f), the main difference being that the bands at 830 and 715 [cm.sup.-1] were more pronounced. The band at 715 [cm.sup.-1] was attributed to the in-plane deformation vibrations of the planar C[O.sub.3] units (Tatzber et al. 2007).
[FIGURE 4 OMITTED]
The CPMAS [sup.13]C NMR spectra of both pine and eucalyptus bark feedstocks before charring (Fig. 3) were dominated by a signal in the O-alkyl C regions (45-110 ppm), reflecting the dominance of cellulose (Preston et al. 1998; Baldock and Smemick 2002). A methoxyl (also produced by [alpha]-amino groups) carbon signal at 56 ppm and several signals in the aryl C region (110-165 ppm) indicated the presence of lignin and/or that of hydrolysable tannin. Weak signals were observed in the alkyl (0-45 ppm) and carboxyl (165-210 ppm) regions. In contrast, the CP spectra of the Ctr-EU and Ctr-PI biochar treatments were dominated by peaks in the aromatic/unsaturated region (110-165 ppm). Both of these biochars showed signals from the aromatic spinning side bands. The aryl peak was centred at 130.5 ppm for the Ctr-PI and 129.3 ppm for the Ctr-EU treatment. The Ctr-PI biochar had a shoulder at ~155 ppm attributed to O- or N-substitute aryl groups. The CP spectra of the treated biochars, with either diluted (liquid treatment) or undiluted (solid treatment) tannery slurry, showed a similar pattern to that of the control except for the size of the aromatic peak of the biochar from eucalyptus. The eucalyptus biochar had a less intense aryl peak than that of the pine, which could be related to the lower C content of the former (Fig. 3, Table 3). Overall, the results reflect the strong aromaticity of the biochars produced in this experiment (both treated and controls).
The C 1s core level spectra obtained with XPS spectroscopy for Ctr-EU, S-EU, and L-EU biochar samples are shown in Fig. 4 and the normalised area of the peaks corresponding to each O-containing surface group for the different biochars are presented in Table 5. The spectra of the 3 samples displayed signals attributed to the aliphatic/aromatic carbon group (CHx, C-C/ C=C) (284.6 eV), hydroxyl groups (-C-OR) (285.7-286.3 eV), and carboxylic groups, esters, or lactones (-COOR) (289-290 eV). The spectrum of the S-EU treatment had the highest content of hydroxyl groups, whereas that from the L-EU treatment contained the highest carboxylic group proportion. In addition, the spectrum of the latter treatment showed a signal corresponding to carbonyl groups (>C=O) (287.2 eV). Overall, the results indicate the presence of more oxygen-rich functional groups in the L-EU and S-EU biochars than the Ctr-EU biochar, particularly in the diluted alkaline treatment. This finding agrees with the FT-IR spectra and the acidic functional groups determined by titration.
[FIGURE 5 OMITTED]
Structural characteristics of biochars
Adsorption of [N.sub.2] in the Ctr-PI and Ctr-EU biochar samples displayed a type I isotherm, according to the IUPAC classification (data not shown); this indicates that the samples are essentially microporous (pore size <2 nm). The surface area of the Ctr-EU biochar (135 [m.sup.2]/g) was smaller than that of the Ctr-PI biochar (235 [m.sup.2]/g) (Table 4). The treated biochars showed type II isotherms, which are typical of non-porous materials, and the values of specific surface area calculated are only due to the external surface (<9 [m.sup.2]/g, Table 4). A decrease in the surface area of biochars after alkaline activation was also observed in other studies (Chiang et al. 2000). However, this decrease did not imply a reduction in surface charge, as shown above. Finally, the SEM images of untreated biochars (Fig. 5) showed the presence of numerous hollow channels originated from plant cells with honey comb structures. Biochars made from eucalyptus were especially affected by the alkaline tannery slurry treatments, with a collapse of the former structure, rendering a highly disorganised appearance (Fig. 5).
Batch sorption study
Sorption of N[H.sub.4.sup.+]-N after 4 additions of N[H.sub.4.sup.+]-N solutions of 40 mg N/L was significantly (P < 0.05) greater for the Ctr-EU treatment than the Ctr-PI treatment, with average values of 0.036 and 0.025 mmol N/g, respectively (Fig. 6), which represented 61 and 41% of the total amount of N added. Treatment of feedstocks with alkaline tannery slurry increased the N[H.sub.4.sup.+]-N sorption capacity of the resulting biochar, with the increase being highest for the undiluted tannery slurry treatment. The highest sorption was achieved by the S-EU treatment, with a value of 0.047 mmol N/g, which represented 83% of the total amount added. Therefore, sorption did not appear to be directly related to the total amount of surface charge present in the biochar. However, as the amount of N[H.sub.4.sup.+] added was 2-30 times lower than the estimated total amount of carboxylic acid groups, the role of the total amount of surface charge present on N[H.sub.4.sup.+] sorption is difficult to evaluate. The present results also provide evidence of the lack of influence of surface area on N[H.sub.4.sup.+] sorption. Others studies have also observed a reverse correlation between surface area and the sorption of NH3 and N[H.sub.4.sup.+] (Lee and Reucroft 1999; Vassileva et al. 2009).
Ammonium desorption from the different biochars was low, being 14 and 27% of that added, for Ctr-EU and Ctr-PI biochar treatments, and <2% for the treated biochars (Fig. 6). Therefore, the alkaline feedstock treatment increased the N[H.sub.4.sup.+] retention strength of the biochar. This could be attributed to an increase in coulombic forces that prevented a fast desorption. However, other mechanisms could also be involved in N[H.sub.4.sup.+] retention, such as chemisorptions-ammonia fixation (Stevenson 1982) or the role of S-functional groups (Petit et al. 2010). Analyses of N 1 s core level were carried out in 2 samples (Ctr-EU and S-EU) after the sorption experiments. Only the S-EU sample had enough N (atomic percentage of 1.9%) to be detectable by XPS (data not shown), and the latter mainly originated from the tannery waste. After deconvolution, its N 1s core level spectrum provides evidence for 3 species (Raymundo-Pinero et al. 2002): imine-pyridine (398.5 eV), neutral amine (399.4 eV), and pyrrole-pyridone and charged N (400.5 eV). These results do not provide enough evidence to determine the type of mechanism causing N[H.sub.4.sup.+] retention, as the amount added in the sorption experiment was probably too low to be detectable using XPS. However, these results do provide evidence for the presence of non-available forms of N in this treated biochar.
[FIGURE 6 OMITTED]
Pyrolysis of pine and eucalyptus barks to a final temperature of 550[degrees]C produced highly aromatic biochars with large internal microporous surface areas. The final recovery accounted for 43% and 45% of the initial C, respectively, and 57% and 47% of the initial N, respectively. Pre-treament of the barks with alkaline tannery slurry prevented the development of internal microporous surfaces of the biochars. Despite this reduction in internal surface area, the treated biochars had more surface functional groups (particularly carboxyl and carbonyl groups) and greater N[H.sub.4.sup.+] absorption capacity than the untreated chars. Desorption of the absorbed N[H.sub.4.sup.+] was low, especially for treated biochars compared with untreated biochars. Pretreatment of feedstock with alkaline tannery slurry represents a low-cost methodology for surface activation of biochars to improve their N[H.sub.4.sup.+] sink strength for wastewater treatment. The resultant activated biochar has potential as a product for the treatment of waste streams, and once saturated with N[H.sub.4.sup.+], it could also be used as a slow-release fertiliser, which adds value to the original biochar.
Appendix. Assignments of absorption peaks and bands in FTIR spectra, indicating functional groups associated Wavenumber ([cm.sup.-1]) Assignment 3300, 3400 H bonded OH groups (alcohols, phenols, organic acids) 2950 Methyl CH asymmetric C[H.sub.3] 2830, 2855 Methylene symmetric stretch -C[H.sub.2] 1750 C=O stretching 1620 Aromatic and olcifinic C=C, C=0 of bonded conjugated ketones, quinine 1600 C=C, C=0 stretching conjugated to the aromatic ring 1500, 1508 Aromatic ring (C=C) vibrations; amide 11 vibrations 1433, 1459 C-H deformations, asymmetric in C[H.sub.3] and C[H.sub.2] 1400, 1420 Aromatic ring vibrations combined with C-H in plane deformation 1329 Syringyl ring with C-O stretching 1270 Guaiacyl ring with C-O stretching 1223, 1220 C-C plus C-O, C=0 stretching 1220 C-O or C=0 stretching 1133, 1140, 1157 C-O stretching vibrations of C-O-C groups 1100 C-O stretch for O-C[H.sub.3] and C-OH groups 1031 Aromatic CH deformation and C=0 stretching 1010, 1000 C-O-C groups 875, 878 Carbonate ion, C[O.sub.3] 712, 715 Deformation vibrations of planar C[O.sub.3] Wavenumber ([cm.sup.-1]) References 3300, 3400 Chen et al. (2002); Cheng et al. (2006) 2950 Gunzler and Bock (1990); Chiang et al. (2000) 2830, 2855 Gunzler and Bock (1990) 1750 Chiang et al. (2000) 1620 Duggan and Allen (1997) 1600 Guo and Bustin (1998); Wang et al. (2009) 1500, 1508 Sharma et al. (2004); Haberhauer et al. (1998); 1433, 1459 Sharma et al. (2004); 1400, 1420 Smith and Chughtai (1995) 1329 Wang et al. (2009) 1270 Wang et al. (2009) 1223, 1220 Wang et al. (2009) 1220 Wang et al. (2009) 1133, 1140, 1157 Sharma et al. (2004) 1100 Pradhan and Sandie (1999); Sharma et al. (2004) 1031 Wang et al. (2009) 1010, 1000 Arriagada et al. (1997) 875, 878 Chiang et al. (2000) 712, 715 Tatzber et al. (2007)
The authors acknowledge the Manawatu Microscopy and Imaging Centre (MMIC) and Doug Hopcroft for assistance in preparing the samples and operating the SEM images, and Edwin Mercer from Carter Holt Harvey NZ Ltd for supplying the feedstocks. Authors are also grateful to Gonzalo Almendros, Sarah Fiol, Bob Stewart, Mike Bretherton, Anne West, and Ian Furkert for timely guidance and comments. J.A.M.-A. acknowledges the assistance of the Spanish Ministry of Science and Education for its award of a Juan de la Cierva contract. M.C.A. is very grateful for financial support from the Ministery of Agriculture and Forestry of New Zealand. The authors thank the anonymous reviewers for their valuable suggestions and comments on the manuscript.
Manuscript received 5 January 2010, accepted 14 May 2010
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K. Hina (A), P. Bishop (A), M. Camps Arbestain (A,E), R. Calvelo-Pereira (B), J. A. Macia-Agullo (C), J. Hindmarsh (D), J. A. Hanly (A), F. Macias (B), and M. J. Hedley (A)
(A) Institute of Natural Resources, Private Bag 11222, Massey University, Palmerston North 4442, New Zealand.
(B) Departamento de Edafologia y Quimica Agricola, Facultad de Biologia, Universidad de Santiago de Compostela, 15782-Santiago, Spain.
(C) Instituto Nacional del Carbon (CSIC), Apartado 73, 33080-Oviedo, Spain.
(D) Institute of Food, Nutrition and Human Health, Massey University, Palmerston North 4442, New Zealand.
(E) Corresponding author. Email: email@example.com
Table 1. Elemental composition of tannery waste residue Values are total concentration Major Minor elements (g/kg) elements (mg/kg) Al 0.2 As <2.0 B 0.1 Cd <0.10 Ca 12.0 Cr 2.2 Co <0.004 Cu 4.2 Fe 0.3 Pb 0.5 Mg 0.4 Zn 49.0 Mn 0.0 Ni <2.0 Hg <0.001 Mo <0.004 P 1.9 K 4.2 Se <0.02 Na 110.0 Organic C 430.0 N 66.0 S 62.4 Table 2. Values (%) of neutral detergent fibre (NDF), hemicelluloses, cellulose, lignin, and readily degradable cellular contents (RDCC) of pine (PI) and eucalyptus (EU) feedstocks before and after pretreatment with solid, undiluted tannery waste (S) Sample NDF Hemicellulose Cellulose Lignin RDCC PI feedstock 83.7 17.7 39.7 26.3 16.3 S-PI feedstock 63.4 12.8 31.3 19.3 36.6 EU feedstock 77.4 15.6 51.3 10.5 22.6 S-EU feedstock 39.6 7.9 26.6 5.1 60.4 Table 3. Elemental analysis of feedstocks and biochars PI, Pine; EU, eucalyptus; L, S, diluted and undiluted tannery waste; Ctr, control; OC, organic C =total C--C-C032 ; n.a., not analysed/not available Sample Chemical composition (wt, %) OC N H 0 S Ash (A) PI feedstock 48.7 0.28 6.39 46.1 <0.05 0.5 S-PI feedstock 42.6 1.81 6.43 40.1 4.1 7.5 Ctr-PI 77.7 0.61 3.69 16.7 <0.05 4.9 L-PI 70.4 1.33 3.43 15.4 1.0 8.6 S-PI 65.3 1.99 3.11 18.2 1.5 16.9 EU feedstock 45.8 0.33 6.55 48.2 <0.05 4.6 S-EU feedstock 37.5 3.21 6.46 39.2 6.8 14.4 Ctr-EU 73.4 0.56 3.46 12.0 0.2 13.0 L-EU 56.0 1.82 3.11 22.6 2.3 27.8 S-EU 51.4 2.14 2.53 21.2 2.9 30.1 C-C Sample [O.sub.3] Yield Recovery (%) (%) C N S PI feedstock n.a. n.a. n.a. n.a. n.a. S-PI feedstock n.a. n.a. n.a. n.a. n.a. Ctr-PI 0.3 27 43 57 n.a. L-PI 0.8 26 43 19 6 S-PI 1.3 27 41 30 10 EU feedstock n.a. n.a. n.a. n.a. n.a. S-EU feedstock n.a. n.a. n.a. n.a. n.a. Ctr-EU 0.5 28 45 47 n.a. L-EU 1.8 29 43 16 10 S-EU 2.8 32 42 21 13 (A) At 900[degrees]C. Table 4. Values of pH, carboxylic groups, and BET surface area PI, Pine; EU, eucalyptus; L, S, diluted and undiluted tannery waste; Ctr, control Carboxylic BET Samples pH groups ([m.sup.2] (mmol/g) /g) Ctr-PI 9.8 0.006 235.0 L-PI 10.3 0.052 3.3 S-PI 10.5 0.080 9.2 Ctr-EU 8.8 0.037 135.0 L-EU 9.6 0.164 2.2 S-EU 10.6 0.039 2.5 Table 5. Normalised area of the peaks corresponding to each oxygen surface group for the different biochars EU, Eucalyptus; L, S, diluted and undiluted tannery waste; Ctr, control; b.d.l., below detection limit Sample -C-0R >C=O -COOR L-EU 0.146 0.084 0.111 S-EU 0.197 0.057 b.d.l. Ctr-EU 0.163 0.050 b.d.l.
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|Author:||Hina, K.; Bishop, P.; Arbestain, M. Camps; Calvelo-Pereira, R.; Macia-Agullo, J.A.; Hindmarsh, J.; H|
|Publication:||Australian Journal of Soil Research|
|Date:||Sep 1, 2010|
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