Article pubs.acs.org/est
Generalized Two-Dimensional Perturbation Correlation Infrared Spectroscopy Reveals Mechanisms for the Development of Surface Charge and Recalcitrance in Plant-Derived Biochars Omar R. Harvey,*,†,‡ Bruce E. Herbert,§ Li-Jung Kuo,∥ and Patrick Louchouarn⊥ †
Water Management and Hydrologic Sciences, Texas A&M University, College Station, Texas 77843, United States Department of Geography and Geology, The University of Southern Mississippi, Hattiesburg, Mississippi 39406, United States § Geology and Geophysics, Texas A&M University, College Station, Texas 77843, United States ∥ Marine Science Laboratory, Pacific Northwest National Laboratory, Sequim, Washington 98382, United States ⊥ Department of Marine Science, Texas A&M University at Galveston, Galveston, Texas 77553, United States ‡
S Supporting Information *
ABSTRACT: Fundamental knowledge of how biochars develop surface-charge and resistance to environmental degradation is crucial to their production for customized applications or understanding their functions in the environment. Two-dimensional perturbation-based correlation infrared spectroscopy (2D-PCIS) was used to study the biochar formation process in three taxonomically different plant biomass, under oxygenlimited conditions along a heat-treatment-temperature gradient (HTT; 200−650 °C). Results from 2D-PCIS pointed to the systematic, HTT-induced defragmenting of lignocellulose H-bonding network and demethylenation/demethylation, oxidation, or dehydroxylation/dehydrogenation of lignocellulose fragments as the primary reactions controlling biochar properties along the HTT gradient. The cleavage of OH...O-type Hbonds, oxidation of free primary hydroxyls to carboxyls (carboxylation; HTT ≤ 500 °C), and their subsequent dehydrogenation/dehydroxylation (HTT > 500 °C) controlled surface charge on the biochars; while the dehydrogenation of methylene groups, which yielded increasingly condensed structures (R−CH2−R →RCH−R →RCR), controlled biochar recalcitrance. Variations in biochar properties across plant biomass type were attributable to taxa-specific transformations. For example, apparent inefficiencies in the cleavage of wood-specific H-bonds, and their subsequent oxidation to carboxyls, lead to lower surface charge in wood biochars (compared to grass biochars). Both nontaxa and taxa-specific transformations highlighted by 2D-PCIS could have significant implications for biochar functioning in fire-impacted or biochar-amended systems.
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INTRODUCTION
chemistry of biochars is commonly studied using Fourier transform infrared (FTIR) spectroscopy. However, significant peak overlaps in key areas of the biochar infrared spectra and subsequent inability to effectively decipher perturbationinduced (e.g., heat-treatment-temperature or HTT) changes in specific bonds limits the suitability of conventional FTIR analysis for mechanistic assessments. One alternative to conventional FTIR analysis, for mechanistic assessments, is generalized two-dimensional perturbation correlation infrared spectroscopy (2D-PCIS).11,12 In addition to providing a better resolution of significant peaks, 2D-PCIS allows for the elucidation of simultaneously- and sequentially occurring processes.12 Despite the widespread use of 2D-PCIS in mechanistic studies on colloids and polymeric materials,12,16,17 no previous
Char/charcoal black carbon produced during natural pyrogenic events (e.g., vegetation fires) or under simulated conditions (hereon referred to as biochars) is receiving significant attention due to increased recognition of their dynamic role in the biogeochemical cycling of carbon, contaminants, and nutrients.1−5 The dynamic nature of biochars is largely attributable to physical and chemical heterogeneity stemming from differences in feedstock chemistry and pyrolysis conditions.4−7 The development of structure−reactivity relationships and a fundamental understanding of how biochars develop their functionalities are therefore crucial to 1) predicting/explaining the behavior of biochars in fire-impacted or biochar-amended systems and 2) designing and selecting optimal biochars for specific environmental applications. Significant progress has been made in elucidating pertinent structure−reactivity relationships.4,8−10 However, a fundamental understanding of the mechanisms controlling the development of biochar functionalities and subsequent structure− reactivity relationships is still lacking. Functional group © 2012 American Chemical Society
Received: Revised: Accepted: Published: 10641
July 24, 2012 September 2, 2012 September 5, 2012 September 5, 2012 dx.doi.org/10.1021/es302971d | Environ. Sci. Technol. 2012, 46, 10641−10650
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studies have used the technique to study the biochar formation process. The advantages of 2D-PCIS for mechanistic assessments are largely attributable to how the spectra are derived and what they represent. In contrast to conventional FTIR analysis, where spectral variations represents the interaction between the infrared beam and the sample, the spectra in 2DPCIS represents the quantitative similarity or dissimilarity between changes in the intensity at vibrations (v1 and v2) across a given external variable/perturbant (T) at a fixed interval (Tmin to Tmax). Therefore, the 2D-PCIS spectrum represents perturbation-induced changes in sample characteristics as T is incrementally changed between Tmin and Tmax and highlights changes in sample characteristics that may not be apparent from conventional FTIR. Mathematically, the 2D-PCIS spectrum, X(v1, v2) can be expressed as X(v1 , v2) = y ̃(v1 , T ) ·y ̃(v2 , T ′)
Table 1. Summary of Noda’s Rules for Interpreting Synchronous (Φ(v1, v2)) and Asynchronous (Ψ(v1, v2)) Cross Peaks in 2D-PCIS Analysis As a Perturbant, T Is Changed Incrementally from Tmin to Tmax if the sign at Ψ(v1, v2) is
+ -
(1)
+
+
-
+
+
-
-
-
interpretation the intensity of v1 and v2 are changing in the same direction, i.e. increasing or decreasing together the intensity of v1 and v2 are changing in opposite directions the change at v1 is occurring predominantly before that at v2 the change at v1 is occurring predominantly after that at v2 the change at v1 is occurring predominantly after that at v2 the change at v1 is occurring predominantly before that at v2
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where ⟨ ⟩ denotes a cross-correlation function designed to compare the dependence of intensity changes, ỹ(v, T), at v1 and v2 on T. To simplify the mathematical manipulation, X(v1, v2) is treated as the complex number function X(v1 , v2) = Φ(v1 , v2) + i Ψ(v1 , v2)
and the sign at Φ(v1, v2) is
EXPERIMENTAL SECTION Biochars and Biochar Characterization. Biochars were prepared under oxygen-limited conditions, from three plant species and HTT ranging from 200 to 650 °C, in increments of 50 °C. A detailed description of the charring procedures is provided in Kuo et al.13 In brief, 5 g of oven-dried plant material (1 cm pieces) was placed in quartz crucibles, covered to create an oxygen-limited environment, and then pyrolyzed at the desired HTT (200−650 °C) for 1 h in a muffle furnace (Lindberg Model 51442/59344). With the exception of grinding and sieving (250 μm), the biochars were used without any further processing. The three plant species represented were honey mesquite (Prosopis glandulosa), cordgrass (Spartina spartinae), and loblolly pine (Pinus taeda), referred to as HM, CG, and PI, respectively. The HM and CG feedstocks were obtained from Laguna Atascosa National Wildlife Refuge (Rio Hondo, TX, USA), while the PI was collected from Bastrop State Park (Bastrop, TX, USA). Taxonomically these materials could be classified as angiosperms (HM, CG) versus gymnosperm (PI); woody (HM, PI) versus nonwoody (CG); C3 (HM, PI) versus C4 (CG); and legumes (HM) versus nonlegumes (CG, PI). Physicochemical properties of the biochars were determined through elemental, X-ray diffraction (XRD), cation exchange capacity (CEC), and biochar recalcitrance (R50) analyses. The carbon, hydrogen, nitrogen, and oxygen (CHNO) content of the biochars has been previously determined in an earlier publication.13 CEC of the biochars was determined based on K+ for Na+ cation exchange at pH 7.3 For XRD analysis (Bruker D8 X-ray diffractometer), samples were mounted as dry powders and scanned over the 4−70 2θ range. Biochar recalcitrance was assessed using the recalcitrance index (R50) proposed by Harvey et al.4 In this approach biochar recalcitrance is calculated as follows: R50 = T50,biochar/T50, graphite, where T50, biochar and T50, graphite are determined by thermogravimetric analysis and represent the temperatures at which 50% of the weight is lost from the biochar and graphite, respectively. Coefficient of variation for CEC and R50 was 500. Dehydrogenation reactions would also explain changes in Iv/ Imax observed in the CH-stretch range of the synchronous 2D correlation spectra (Figure 2b, d, and f). The dehydrogenation of single-bonded R−CHn to yield double-bonded RCHn (e.g., R−CH2−R→ RCH−R) would account for the initial increase in the concentration of RCHn (v = 3050 cm−1) observed with increasing HTT. Further dehydrogenation to form increasingly condensed structures, via aromatization (e.g., RCH−R → RCR), would account for the subsequent decrease in RCHn concentrations in CG and HM biochars produced at HTT > 500 °C. No decrease in RCHn was observed in PI biochars at HTT > 500 °C. The reason for this is not completely clear, but such observation suggested a better preservation of RCHn at higher HTTs in PI biochars compared to CG and HM biochars. The consequence of thermally induced dehydrogenation of CHn on biochar properties was indicative of changes in biochar recalcitrance (as determined by R50 values). For example, R50 of the biochars increased as dehydrogenation (e.g., R−CH2−R→ RCH−R) progressed with increasing HTT (Figure 2b, d, and f). The relationship between HTT, R 50, and CHn concentration also suggested that the further dehydrogenation of RCH−R to RCR was a major reaction accounting for differences in recalcitrance between what Harvey et al.4 referred
primary alcohol and methylene functional groups are present in all three major plant biomacromolecules (hemicellulose, cellulose, and lignin).22 Attributing changes highlighted in Figure 1 to a specific biomacromolecule is therefore very difficult. We used the more generic term “lignocellulose” to collectively refer to the observed changes in the polymeric nature of hemicellulose, cellulose, and lignin of the original plant material. The link between thermal transformation of lignocellulose and biochar formation process was supported by other lines of evidence. For example, XRD analysis showed no evidence for the presence of crystalline cellulosic components in wood or grass biochars produced at HTT ≥ 300 °C (Figure S2).10 A similar transition was also apparent in the OH-stretch region of the synchronous 2D spectra for biochars produced from cellulose and hemicellulose (Beechwood xylan) under similar conditions (Figure S3). Using Struszczyk’s equation,23 the hydrogen-bond strength (EH) for R−CHnOH...O (3400−3385 cm−1) would be on the order of 18 to 19 kJ mol−1 (compared to 0 kJ mol−1 for R−OH). An EH of 18−19 kJ mol−1 is in line with reported values for inter- and intramolecular hydrogen bond strengths in lignocellulose24−27 and is therefore consistent with the fragmentation of lignocelluose in the initial stages of the biochar formation process. The occurrence of the R−CHnOH...O and R−CH2 stretch vibrations on the v1= v2 diagonal of Figure 1 indicates that these bonds change to a greater degree (than free−OH and R CHn) in response to increasing HTT.12 Correspondingly, the negative correlation values for the off-diagonal free−OH and RCHn vibrations indicate that increasing HTT have opposite effects on the concentrations of R−CHnOH...O versus R−OH and R−CH2 versus RCHn in the biochars.12 Plots of changes in relative concentrations (Iv/Imax) of the functional groups with HTT confirmed these observations (Figure 2). For R− CH nOH...O and R−CH 2 functional groups, which are characteristic of unaltered lignocellulosic polymers, Iv/Imax decreased with increasing HTT. In contrast, concentrations of R−OH and RCHn functional groups increased and then decreased with increasing HTT, showing some major shifts in the original plant structure over the HTT range tested. The thermally induced cleavage of H-bonds in lignocellulose to yield free−OH was coincidental with an increase in biochar CEC from less than 20 to maxima of around 35, 60, or 80 cmol kg−1 for PI, HM, and CG biochars, respectively (Figure 2a, c, and e). CEC on these biochars have been linked to deprotonation of carboxylic groups with average pKa of around 4.4, 3.9, and 4.1 for PI, HM, and CG biochars, respectively.3 It would therefore be reasonable to suggest that at least a portion of the free−OH were carboxylic groups produced from primary alcohol (R−CHnOH) groups. The oxidative formation of carboxyls from primary alcohols is well-known.28−31 The presence of thermally produced carboxylic groups on the surfaces of the biochars would be consistent with 1) depolymerization/fragmentation of lignin components to their respective monomeric phenols/alcohols with subsequent thermal oxidation to carboxylic acids13,32−35 and 2) thermal degradation of cellulosic components predominantly via an alternative mechanism to the transglycosylation pathway (which would favor the formation of anhydrosugars36,37). The fragmentation of phenylpropyl units of lignin polymers and their subsequent oxidation can be supported by previously published data, on some of the biochars used in this study, which showed a peak yield of free 10645
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Figure 3. Asynchronous 2D-PCIS spectra in the OH-stretch (3700−3100 cm−1) and CH-stretch (3100−2700 cm−1) analysis windows for (a, b) cordgrass, (c, d) loblolly pine, and (e, f) honey mesquite biochars produced under oxygen-limited conditions, HTD of 1 h, and HTT between 200 and 650 °C. Dark gray, light gray, and white areas indicate positive, zero, and negative correlation values, respectively.
to as Class B (0.5 ≤ R50 < 0.70) and Class C (R50 < 0.5) recalcitrant carbons. According to Harvey et al.,4 in a given environment, a Class B biochars would have a recalcitrance intermediate between soot/graphite and uncharred biomass; while a Class C biochars would have recalcitrance comparable to the uncharred plant biomass. Results from synchronous 2D-PCIS in the CH-stretch region were therefore reflective of the significant role dehydrogenation reactions plays in the development of biochar resisitance to biodegradation.
Asynchronous 2D-PCIS. Asynchronous 2D correlation spectra, Ψ(v1, v2), in the OH-stretch and CH-stretch region are shown in Figure 3. A single sequential correlation, around Ψ(2890, 2940), was observed in the CH-stretch range. Vibrations around 2890 cm−1 were reflective of CH-stretch due to methyl (R−CH3) functional groups, while vibrations around 2940 cm−1 were more in line with methylene (R−CH2) functional groups.18 In lignocellulose materials, although methylene groups are more prevalent in the cellulosic components, methyl groups (mostly as methoxyls, R−OCH3) 10646
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Figure 4. Variation in the (a) O/C and H/C elemental ratios and (b) weight loss for cordgrass (CG), loblolly pine (PI), and honey mesquite (HM) biochars produced under oxygen-limited conditions, HTD of 1 h, and HTT between 200 and 650 °C.
are typically associated with lignin.22 A continuous decrease in Iv/Imax for R−CH2 and R−OCH3 with increasing HTT was consistent with demethylenation and demethylation/demethoxylation (Supporting Information Figure S4b, d, and f). Negative correlation values at Ψ(2890, 2940) suggested that demethylenation reactions occurred predominantly at lower HTT than demethylation/demethoxylation - reflecting the generally lower thermal stability of cellulosic compared to lignin components.22 Figure 3 also indicates that during biochar formation, most (all in the case of CG and PI) asynchronous changes in hydrogen-bonding character occurred in sequence with cleavage of the H-bonded primary alcohol (R−CHnOH...O) functional groups. Of these changes only one [at around Ψ(3400, 3530)] was common to all the plant materials. Other changes were specific to the nature of the plant material. For example, the change at around Ψ(3400, 3630) was unique to the wood biochars (PI and HM); the change at around Ψ(3400, 3200) was unique to the nonlegumes (CG and PI); and the one at Ψ(3400, 3435) was unique to the legume (HM). Asynchronous changes specific to the HM (legume) biochars, but not associated with the R−CHnOH...O vibration, were also apparent at Ψ(3285, 3435) and Ψ(3285, 3365). The specificity of certain vibrations to particular biochars was reflective of variability in the structure, configuration, composition, and, subsequently, H-bonding network of lignocellulose across plant taxa (e.g., grass versus wood or legumes versus nonlegumes). Legume-specific vibrations at 3435, 3365, and 3285 cm−1 also pointed to the involvement of H-bonding via NHn functional groups in the HM biochars.18 Based on Noda’s rules,12 asynchronous changes at 3630, 3530, and 3200 cm−1 occurred predominantly before the mass cleavage of R−CHnOH...O H-bonds at 3400 cm−1. For the legume-specific vibrations, the change at 3285 cm−1 occurred predominantly at higher HTT than those at 3435 and 3365 cm−1. With the exception of that at 3200 cm−1 (EH = 32 kJ mol−1), the sequence of change suggested by Noda’s rules for both legume-specific and other vibrations was consistent with expectations based on H-bond strength where the weaker Hbonds (e.g., 3630−3435 cm−1; EH = 1.4−15 kJ mol−1) were the first to respond to increases in HTT, compared to stronger Hbonds at 3400 cm−1 with EH = 18 mol−1.23 Changes in Iv/Imax for the 3530, 3435, 3365, 3285, and 3200 cm−1 vibrations thus reflect a continuous loss in the hydrogen bonding character with increasing HTT, while changes in Iv/Imax for the vibration at 3630 cm−1 reflect an increase in and subsequent loss of
weakly H-bonded hydroxyls (Supporting Information Figure S3a, c, and e). Asynchronous 2-D spectra were therefore reflective of a systematic defragmenting of the H-bonding network/depolymerization of lignocellulose via HTT-induced weakening/cleavage of H-bonds and dehydroxylation/dehydrogenation reactions. An interesting observation from asynchronous 2D-PCIS was the striking similarity between trends in Iv/Imax at the 3630 cm−1 vibrations and those observed for free−OH in the synchronous OH-stretch spectra. Such similarities suggested that observed differences in charge development between wood and grass biochars could in fact be attributed to differences in the susceptibility to (or efficacy of) HTT-induced depolymerization in the two materials. That is, although HTT-induced depolymerization/H-bond cleavage occurred in both sets of biochars the process was less efficient in wood biochars, under conditions favorable for charge development (HTT 300−450 °C). This would account for the presence of H-bonded OH (albeit, weakly H-bonded OH at 3630 cm−1) in wood biochars compared to only free−OH in grass biochars. Links between 2D-PCIS, the Biochar Formation Process, and Other Physicochemical Properties of Biochars. A consistent feature of changes in Iv/Imax with HTT, for vibrations in synchronous and asynchronous 2DPCIS spectra, was the presence of transition (inflection) points around HTT300−350 °C and HTT450−550 °C (Figures 2 and S4). Similar transition points have been observed using other analytical techniques and were attributed to the complete carbonization of cellulose and lignin, respectively.10,41 Changes in Iv/Imax were consistent with such a link. For example, Iv/Imax for biochars produced around the first transition point (HTT300−350 °C) suggested that these biochars had 65− 85% less H-bonding and R−CHn character than biochars produced at a HTT of 200 °C, while biochars produced around the second transition point (HTT450−550 °C) had 5−25% less H-bonding and R−CHn character than biochars produced around the first transitional point. These changes were in line with the cellulosic (66 to 73%) and lignin (4 to 15%) content of the uncharred plant materials, respectively.13 The transition point at HTT450−550 °C has also been attributed to the onset of aromatization13,42 and was consistent with the almost complete loss in H-bonding/R−CHn character as well as a drastic decrease in free−OH/RCHn character observed around this transition point (Figure 2 and S4). Based on the changes in relative concentration of OH and CHn 10647
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Figure 5. Modified black carbon continuum depicting dominant mechanisms of biochar formation and their impact on biochar elemental composition, cation exchange capacity (CEC), and recalcitrance, as HTT is increased from 200 to 650 °C. Recalcitrance classes C and B have a recalcitrance-index (R50) values of 350 °C. On the other hand, if a high CEC biochar is targeted (e.g., to improve agronomic qualities of soils), a grass feedstock and a HTT of 400−450 °C would be ideal. The results of the study also provide significant insights into the mechanisms controlling observed structure−reactivity or structure−function relationships. For example, the specific link to HTT-induced dehydrogenation of methylene groups (R− CH2−R →RCH−R →RCR) could plausibly explain why 1) carbohydrate-rich biomass (dominated by R−CH2) tend to yield biochars with highly-condensed structures7 and 2) at HTT > 300 °C, cellulose-derived biochars tend to have higher R50 values than grass or wood biochars produced under the same charring conditions.4 A recent study by Kinney et al.46 showed that biochars, produced under oxygen-limited conditions, exhibited their highest water holding capacity and minimum hydrophobicity around HTT350−500. Our observations, of the concomitant peaks in free−OH groups and CEC at the midranges of HTT (300−500 °C), are very consistent with those of Kinney et al.46 and could be used to explain the mechanisms for the development of water holding capacity and hydrophobicity in biochars. Finally, taxa-specific HTT-induced changes such as those highlighted in the present study between grass and wood biochars, in the OH-stretch region of 2D-PCIS spectra, may also provide useful information in understanding how biochars produced from different feedstock may behave in a given environment.
functional groups, as shown in Figures 2 and S4, we believe that the transition point around 300−350 °C is also representative of a shift in biochar functional group chemistry from being dominated by H-bonded OH and R−CHn to being dominated by free−OH and RCHn functional groups. Data on weight loss during the biochar formation process and elemental composition of the resulting biochars are presented in Table S1. Plotting atomic O/C or H/C ratios as a function of HTT revealed a common inflection point, around HTT330 °C (Figure 4a). This inflection point coincided with the HTT300−350 °C transition point observed from Iv/Imax data. Similar inflection points (as observed from Iv/Imax data) at HTT330 and HTT490 °C were also apparent from weight loss data suggesting a link between changes in biochar elemental composition, surface chemistry, and weight loss (Figure 4b). As with Iv/Imax, the largest change in weight loss or atomic ratios (O/C or H/C) were observed in biochars formed between HTT200 and 350 °C. Slopes of piece-wise linear fits in Figure 4a suggested that, on average, for every 100 °C increase in HTT, between HTT200 and 330 °C, every fourth carbon lost an oxygen. This compares to a loss of an oxygen for every 16th C between HTT330 and 650 °C. For the same HTT ranges, the rates of H loss were approximately 2 and 3 times that of O, respectively. A 2:1 loss ratio for H:O was consistent with the evolution of water and pointed to dehydration as a major mechanism responsible for thermal transformations in the biochars.8,13,43,44 Dehydration was attributable to the loss of “chemical” water from lignocellulose33 and was consistent with dehydroxylation-dehydrogenation reactions highlighted by Iv/ Imax in the OH- and CH-stretch regions, respectively. The increase in H:O loss from 2:1 to 3:1 at HTT ≥ 330 °C was indicative of dehydrogenation reactions, which play an increasingly significant role in biochar formation and was attributable to increasingly condensed biochar structure with HTT due to increased conjugation/aromatization of lignocellulose fragments.42,45 Environmental Significance. The inherent variability of biochars, coupled with that of the soils to which they are likely to be applied, suggest that their production and use will need to be customized for specific environmental applications. The fundamental understanding gained through the current study could provide significant insights into the optimization of the biochar production process. For example, Figure 5 suggests that
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ASSOCIATED CONTENT
S Supporting Information *
Additional information (including 2D-PCIS analysis in the fingerprint region). This material is available free of charge via the Internet at http://pubs.acs.org.
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AUTHOR INFORMATION
Corresponding Author
*Phone: (01) 601-266-4529. Fax: (01) 601-266-6219. E-mail:
[email protected]. Notes
The authors declare no competing financial interest. 10648
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ACKNOWLEDGMENTS Financial support for O.R.H. was through the Texas Transportation Institute and the University of Southern Mississippi. The Soil Mineralogy Laboratory at Texas A&M University provided instrument time for XRD analysis. Suggestions from the Associate Editor and three anonymous reviewers improved this manuscript.
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