Temperature responses of soil organic matter (SOM) remain unclear partly due to its chemical and compositional heterogeneity. In this study, the decomposition of SOM from two grassland soils was investigated in a 1-year laboratory incubation at six different temperatures. SOM was separated into solvent extractable compounds, suberin- and cutin-derived compounds, and lignin-derived monomers by solvent extraction, base hydrolysis, and CuO oxidation, respectively. These SOM components have distinct chemical structures and stabilities and their decomposition patterns over the course of the experiment were fitted with a two-pool exponential decay model. The stability of SOM components was also assessed using geochemical parameters and kinetic parameters derived from model fitting. Compared with the solvent extractable compounds, a low percentage of lignin monomers partitioned into the labile SOM pool. Suberin- and cutin-derived compounds were poorly fitted by the decay model, and their recalcitrance was shown by the geochemical degradation parameter (ω − C16/∑C16), which was observed to stabilize during the incubation. The temperature sensitivity of decomposition, expressed as Q10, was derived from the relationship between temperature and SOM decay rates. SOM components exhibited varying temperature responses and the decomposition of lignin monomers exhibited higher Q10 values than the decomposition of solvent extractable compounds. Our study shows that Q10 values derived from soil respiration measurements may not be reliable indicators of temperature responses of individual SOM components.
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 Soil organic matter (SOM) is an important component of the terrestrial ecosystem and global carbon cycle [Batjes, 1996; Schlesinger and Andrews, 2000]. The acceleration of SOM decomposition with global warming has become one of the major concerns in predicting future climate change. However, SOM decomposition remains unclear in terms of its temperature sensitivity and the decay patterns of heterogeneous SOM components [Melillo et al., 2002; Knorr et al., 2005; Davidson and Janssens, 2006]. Investigations into SOM decomposition have suggested varying and even contrasting responses of SOM components to temperature increases [Fang et al., 2005; Knorr et al., 2005]. According to the Arrhenius theory, the reaction rate (k) of SOM mineralization is a function of the activation energy of SOM components (Ea, J mol−1) within the enzyme-active temperature ranges (∼5–40°C) [Winkler et al., 1996]:
where a is the theoretical rate at Ea = 0, R is the gas constant (8.314 J mol−1 K−1), and T is the absolute temperature (°K). In other words, the temperature sensitivity of SOM mineralization, Q10, defined as the factor by which the reaction rate differs for a temperature interval of 10°C, should increase with increasing Ea or chemical recalcitrance, and decrease with increasing temperature:
However, k and Q10 values derived from the modeling of soil respiration data do not always follow the Arrhenius theory [Giardina and Ryan, 2000; Fang et al., 2005]. A similar mean residence time (the inverse of reaction rate) and a similar temperature sensitivity have been reported for soils with different recalcitrance [Giardina and Ryan, 2000; Fang et al., 2005]. Alternatively, a higher Q10 value has been calculated for the decomposition of recalcitrant SOM when the soil respiration data were fitted with a multipool model [Knorr et al., 2005]. It has been suggested that the single-pool model of soil respiration ignores the heterogeneity of SOM, and hence the k and Q10 values derived from such models are not reliable indicators of the intrinsic kinetic properties of individual SOM components [Davidson and Janssens, 2006]. Even when a multipool soil carbon model is used, SOM is divided into stable and labile pools based on curve fitting of the respiration data rather than the chemical structure of components within SOM. Therefore, each SOM pool consists of a continuum of soil carbon substrates of varying chemical complexity and such an approach may also conceal the kinetic characteristics of individual SOM structures [Davidson and Janssens, 2006]. To better understand the temperature sensitivity of individual SOM components, it is necessary to examine the decomposition of various SOM components with similar chemical properties.
 The stability of SOM components is associated with their intrinsic chemical recalcitrance and their interaction with the soil matrix [Baldock and Skjemstad, 2000]. Macromolecular lipids and aromatic structures are usually considered to be recalcitrant because they are much more resistant to microbial attack in comparison to easily degradable compounds such as proteins and carbohydrates [Melillo et al., 1982; Gleixner et al., 2001; Melillo et al., 2002]. Similarly, chemically bound or mineral-associated SOM is more stable than SOM in a “free” form [Baldock and Skjemstad, 2000]. Based on the chemical form of SOM, components can be separated into solvent extractable compounds (including n-alkanes, n-alkanols, n-alkanoic acids, carbohydrates, and steroids), ester-bound lipids mainly derived from plant cutin (a biopolymer in the epidermis of leaves), suberin (a biopolymer abundant in bark and roots of vascular plants), and waxes, and phenolic monomers that are ether-linked in lignin macromolecules (Figure 1) [Otto et al., 2005]. These SOM components have various stabilities in the natural environment [Feng and Simpson, 2007] and respond differently to environmental changes [Feng et al., 2007]. Generally, lignin monomers are considered to be more resistant to biodegradation due to their aromaticity [Melillo et al., 1982; Gleixner et al., 2001] and suberin- and cutin-derived compounds are more stable than solvent extractable compounds because they are predominantly linked to soil macromolecules by ester bonds [Riederer et al., 1993]. The degradation of solvent extractable compounds, suberin-derived compounds, cutin-derived compounds, and lignin has been extensively explored in sediment studies using geochemical indicators [Goni and Hedges, 1990; Goni et al., 1993]. However, their decomposition patterns in a controlled soil environment are less well understood because studies under natural soil conditions are usually complicated by fresh plant inputs [Otto and Simpson, 2006]. Thus, it is important to investigate the degradation of specific SOM compounds and to assess their decomposition rates and temperature sensitivity. It is especially important to test if structurally recalcitrant SOM components (such as lignin-derived compounds) have higher Q10 values than more readily degradable compounds in the “free” form (solvent extractable compounds).
 Soil incubation studies are useful techniques to investigate the decomposition and mineralization of SOM under controlled environmental conditions when interferences from plant carbon input are limited [Dalias et al., 2001; Bol et al., 2003; Leifeld and Fuhrer, 2005]. This study employs geochemical techniques to examine the decomposition of various SOM components (solvent extractable compounds, suberin-derived compounds, cutin-derived compounds, and lignin monomers) during a 1-year laboratory incubation at six different temperatures. The investigated SOM components have distinct structures and specific sources (such as plant waxes, suberin, cutin, and lignin) and are not considered to be decomposition products of other compounds in the soil. The objectives of this study are: to investigate the temperature dependence of the decomposition of various SOM components and to assess the stability of these SOM components by both geochemical indicators and kinetic modeling. We hypothesize that lignin monomers and suberin- and cutin-derived compounds are more stable than the solvent extractable compounds and that the decomposition of lignin monomers is accelerated to a greater extent by temperature increases (i.e., the Q10 values of lignin monomers are higher than those of the solvent extractable compounds).
2. Material and Methods
2.1. Soil Incubation
 Surface soil samples were collected from two well-drained, pristine grassland soils in western Alberta in late August, 2005. The first soil (Soil E) was sampled from the University of Alberta Ellerslie Research Station, located south of Edmonton, Alberta, and the second (Soil L) was collected from the Agriculture and Agri-Food Canada Research Station near Lethbridge, Alberta. Both soils are typical grassland soils in the Prairie Ecozone of Western Canada which contains large reserves of SOM [Janzen et al., 1998], and have been well characterized in the past [Feng and Simpson, 2007]. They are therefore good candidates for our soil incubation study. The air mean annual temperature (MAT) for Soils E and L is 1.7°C and 5°C, respectively [Janzen et al., 1998]. Details of the sampling site and soil conditions have been described elsewhere [Feng and Simpson, 2007].
 Soils were kept in the dark at 4 ± 1°C for two months after sampling. Soil L had a high abundance of grass roots during the time of sampling, which were removed before incubation (the minimum root diameter was 2 mm). Both soils were passed through a 2-mm sieve, homogenized, and then incubated in 450-ml glass jars (∼350 g dry soil per jar) at six different air temperatures (MAT of the original sites, 2, 4, 8, 12, and 20°C above the MAT, which represented different scenarios of global warming) in the dark. The water content was kept at ∼30% of the soil dry weight (close to field capacity) by weighing and spraying deionized water at soil surfaces twice a week, so that soil moisture did not limit microbial activity. At least five jars of soil were incubated at each temperature, and subsamples (∼50 g) were collected before incubation (Day 0) and randomly from one of the five jars at each incubation temperature on Day 29, 57, 86, 126, 170, 245, and 365, freeze-dried, and ground (<100 μm) thoroughly prior to chemical analyses.
2.2. Microbial Respiration
 Microbial respiration (r), which is equivalent to soil respiration in the absence of plant roots, was measured in triplicate during the incubation on Days 1, 8, 15, 22, 29, 57, 86, 128, 170, 245, and 365, using the alkali absorption method [Winkler et al., 1996]. Respired CO2 was captured by NaOH (1.0 M × 2.0 ml) in small glass vials placed inside the incubation jars. The jars were sealed and left for 24 h and the vials were then removed and capped. Excess NaOH was determined by precipitation with BaCl2 and titration with 0.2 M HCl with phenolphthalein as an indicator [Zhang et al., 2005]. Microbial respiration rates (r) were normalized to the dry weight of the soil samples and expressed in the units of μg CO2 gsoil−1 h−1.
2.3. Chemical Analyses
 Total carbon, inorganic carbon, and total nitrogen contents of Soils E and L were determined in triplicate at the start of the incubation using a Shimadzu TOC 5000 total organic carbon analyzer equipped with a solid sample module capable of analyzing solid samples such as soils and plant materials (Shimadzu Scientific Instruments, Columbia, MD, USA). Because inorganic carbon was not detected, soil organic carbon (OC) content equaled the total carbon content. Soil carbon loss in the 1-year incubation is found to be small relative to the original soil OC content and is consistent with the literature [White et al., 2002], which may well fall within the precision of the soil TOC measurements (∼5%). Therefore, soil carbon loss (%) during the incubation was estimated by the following equation:
where r is the measured microbial respiration rate in the units of μg CO2 gsoil−1 h−1.
 Chemical extractions (solvent extraction, base hydrolysis, and CuO oxidation) were conducted to produce solvent extractable compounds, suberin- and cutin-derived compounds, and lignin monomers, respectively [Otto et al., 2005]. A diagram illustrating the chemical extractions and SOM compositional information obtained from the analyses is shown in Figure 1. Briefly, freeze-dried soil samples (5–10 g) were extracted with 30 ml of dichloromethane, dichloromethane:methanol (1:1; v/v) and methanol, respectively. The combined solvent extractable compounds were filtered through glass fiber filters (Whatman GF/A and GF/F), concentrated by rotary evaporation, and then dried under nitrogen gas in 2-ml glass vials. The air-dried soil residues from solvent extraction (2 g) were then heated at 100°C for 3 h in teflon-lined bombs with 20 ml of 1 M methanolic KOH. The extracts were acidified to pH 1 with 6 M HCl, and the solvents were removed by rotary evaporation. Lipids were recovered from the water phase by liquid–liquid extraction with diethyl ether, concentrated by rotary evaporation, and dried under nitrogen gas in 2-ml glass vials. The base hydrolysis residues were air-dried and further oxidized with copper (II) oxide (CuO) to release lignin-derived phenols. Soil residues (2 g) were extracted with 1 g copper (II) oxide, 100 mg ammonium iron (II) sulfate hexahydrate [Fe(NH4)2(SO4)2·6H2O] and 15 ml of 2 M NaOH in teflon-lined bombs at 170°C for 2.5 h. The extracts were acidified to pH 1 with 6 M HCl, and kept for 1 h at room temperature in the dark to prevent reactions of cinnamic acids. After centrifugation (at 2500 rev min−1 for 30 min), the supernatants were liquid–liquid extracted with diethyl ether. The ether extracts were concentrated by rotary evaporation, transferred to 2-ml glass vials and dried under nitrogen gas.
 The composition and concentration of chemical extracts were analyzed by gas chromatography/mass spectrometry (GC/MS). Extracts from solvent extraction and CuO oxidation were converted to trimethylsilyl (TMS) derivatives by reaction with 90 μl N,O-bis- (trimethylsilyl)trifluoroacetamide (BSTFA) and 10 μl pyridine for 3 h at 70°C before GC/MS analysis. Base hydrolysis products were first methylated by reacting with 600 μl of diazomethane in ether at 37°C for 1 h, evaporated to dryness under nitrogen, and then silylated with BSTFA and pyridine as described above. Oleic acid (C18:1 alkanoic acid) and ergosterol were derivatized in the same method and used as external standards for solvent-extractable n-alkanes, n-alkanols, n-alkanoic acids and soil steroids, respectively. Oleic acid methyl ester was used as external standard for base hydrolysis products, while vanillic acid-TMS was used for CuO oxidation products. GC/MS analysis was performed on an Agilent model 6890N GC coupled to a Hewlett-Packard model 5973 quadrupole mass selective detector. Separation was achieved on a HP5-MS fused silica capillary column (30 m × 0.25 mm i.d., 0.25 μm film thickness). The GC operating conditions were as follows: temperature held at 65°C for 2 min, increased from 65 to 300°C at a rate of 6°C min−1 with final isothermal hold at 300°C for 20 min. Helium was used as the carrier gas. The sample was injected with a 2:1 split ratio and the injector temperature was set at 280°C. The samples (1 μl) were injected with an Agilent 7683 autosampler. The mass spectrometer was operated in the electron impact mode (EI) at 70 eV ionization energy and scanned from 50 to 650 daltons. Data were acquired and processed with the Chemstation G1701DA software. Individual compounds were identified by comparison of mass spectra with literature, NIST and Wiley MS library data, authentic standards, and interpretation of mass spectrometric fragmentation patterns. External quantification standards were used and the response factor was assumed to be 1 for all compound classes. Concentration of individual compound was calculated by comparison of the peak area of the compound to that of the standard in the total ion current (TIC) and was then normalized to the sample OC content.
2.4. Data Analyses
 Enzymatic reactions (such as SOM decomposition by microorganisms) in well-mixed media under equilibrium conditions are usually fitted with a first-order exponential model: g(t) = ∑cie−ki×t, where g(t) is the remaining carbon fitted to observational data, ci is the initial size of carbon “pools” of varying degrees of decomposition, and ki is the decay rate [Schimel and Weintraub, 2003; Davidson and Janssens, 2006]. To simplify the modeling process, a two-pool exponential model was used to fit the decomposition of SOM components in this laboratory incubation:
where C(t) is the concentration of SOM components (mg/g OC) remaining in the soil at time t (days), Cstable and Clabile are the concentrations of stable and labile SOM pools, respectively, and ks and kl are the decay rates of stable and labile SOM pools (day−1). Conceptually, the stable and labile pools of individual SOM components in this study have the same chemical structures but differ in their interactions with minerals or humic substances that may limit their decomposition rate due to physical protection [Sollins et al., 1996; Baldock and Skjemstad, 2000]. Because the resistant SOM components typically have a mean residence time of 20 to 50 years (ks < 1.4 × 10−4 day−1) [Chapin et al., 2002], and are hence not expected to undergo detectable decrease in the 1-year incubation, the model (4) is further simplified to:
 The temperature dependence of the decomposition rate (k) and respiration rate (r) was modeled for the Arrhenius function (equation (1)). Both the a and Ea parameters were allowed to vary for the model fitting of each single class of SOM components. The Q10 value was calculated according to equation (2) for a temperature of 15°C, which is commonly used as the reference temperature [Reichstein et al., 2002]. The decomposition rate (k) of the “labile” SOM pool was also listed for a similar temperature (close to 15°C), i.e., MAT+12°C for Soil E and MAT+8°C for Soil L, to compare the stability of individual SOM components. The model fitting was performed using Origin™ Version 7.0 (Microcal Software, MA, USA) at a confidence level of P ≤ 0.05. The degradation parameters of cutin, suberin, and lignin, and the modeled values of Cstable, Clabile, and k were compared against incubation days or temperature increases using linear regression analysis, and the difference was considered significant at a level of P < 0.05. Due to a high variance associated with the k and Ea values derived from the model fitting, statistical comparisons of the k and Q10 values were not made between different SOM components.
3.1. Microbial Respiration and Soil Carbon and Nitrogen Contents
 Microbial respiration rates (r) were generally higher in Soil L than in Soil E, and r values decreased in a pseudo-exponential mode with incubation time in both soils (Figure 2). In Soil E, r values decreased by more than 40% in the first week of incubation and then slowly decreased to 0.25–0.36 μg CO2 gsoil−1 h−1 at the end of the experiment. In comparison, r values in Soil L decreased sharply at higher temperatures (MAT+12°C and MAT+20°C) in the first week of incubation and decreased much more slowly at lower temperatures (MAT−MAT+8°C) in the first two months of incubation (Figure 2b). Temperature increases significantly enhanced microbial respiration rates during the entire incubation period in that r values measured on the same day of incubation were positively correlated to incubation temperatures (P < 0.05).
 Soil OC content was 4.85% and 2.69% for Soils E and L, respectively. Total nitrogen content was 0.46% for Soil E and 0.28% for Soil L. Both soils had a similar atomic C/N ratio of 11–12 at the start of the incubation. Based on the respiration rate on Day 86 (which was close to the average rate), soil carbon loss during the 1-year incubation was estimated to be 0.08–0.12% in Soil E, which accounted for 1.7–2.5% of the original soil OC content. Similarly, soil carbon loss was about 0.24–0.40% in Soil L, equivalent to 8.9–14.9% of the original OC content. The estimated soil OC loss agrees with the annual carbon loss in fallow cropping soils and those in short-term soil incubation studies [Rasmussen et al., 1998; Reichstein et al., 2000; Leifeld and Fuhrer, 2005]. The size of carbon loss in Soil E was small as compared to its OC content. For comparative purpose, we used 4.85% and 2.69% as the OC content for Soils E and L, respectively, to calculate the OC-normalized concentration of SOM components in the soil.
3.2. Decomposition of Solvent Extractable Compounds
 Based on their chemical structures, the solvent extractable compounds of both soils were grouped into four categories: odd-numbered n-alkanes (in the range of C21–C33), even-numbered n-alkanols (in the range of C16–C30), even-numbered n-alkanoic acids (in the range of C12–C28), and steroids (cholesterol, ergosterol, β-sitosterol, stigmasterol, sitosterone, and campesterol). The composition of n-alkanes, n-alkanols, and n-alkanoic acids reflected a predominant input from plants, i.e., these compounds were primarily derived from plant sources and not from the decomposition of other SOM components. Plant steroids (β-sitosterol, stigmasterol, sitosterone, and campesterol) comprised more than 80% of the steroids detected in both soils with minor inputs of steroids from animals (cholesterol) and fungi (ergosterol [Otto et al., 2005]). Among the identified plant steroids, sitosterone was the degradation product of the precursor sterols (β-sitosterol and stigmasterol) [Otto and Simpson, 2005] and the ratios of precursor sterols (β-sitosterol and stigmasterol) to their degradation product (sitosterone) were 6.5 in Soil E and 2.6 in Soil L at the start of the incubation. Carbohydrates (sucrose, glucose, mannose, and trehalose) were also detected in the solvent extractable compounds of both soils. However, the major carbohydrates have multiple sources such as the fungal input of trehalose and both plant and microbial inputs of glucose [Otto et al., 2005; Feng and Simpson, 2007]. Therefore, carbohydrate distributions are not included here because they have multiple sources and are difficult to interpret within the context of this study.
 The decomposition of solvent extractable compounds was fitted with the first-order exponential decay model (equation (5) and Figure 3). Soil E had a lower concentration of solvent extractable compounds than Soil L, and exhibited a better exponential model fit (Figures 3a–3d) than Soil L (Figures 3e–3h). The exponential decay rate (k) and the size of stable and labile pools (Cstable and Clabile) were derived from model fitting parameters. The concentrations of stable and labile pools of solvent extractable compounds did not differ between different incubation temperatures (P > 0.05) and the average values were taken for each soil (Table 1) to compare the stability of individual classes of compounds. In Soil E, more than 75% of the solvent extractable compounds were classified into the “labile” pool, with soil steroids comprising the highest percentage in the labile fraction (95%). By comparison, 64–85% of the solvent extractable compounds were in the labile pool in Soil L and the concentration of the labile components was much higher than those in Soil E (with the exception of steroids). Solvent extractable compounds in the “labile” pool in both soils had similar decay rates (ranging from 0.010 ± 0.007 to 0.026 ± 0.006 d−1 at a similar temperature, i.e., MAT+12°C for Soil E and MAT+8°C for Soil L; Table 1) except Soil E steroids, which had a faster decay rate of 0.059 ± 0.004 d−1 at MAT+12°C. The decay rates increased with increasing incubation temperature for n-alkanes, n-alkanols, and steroids in Soil E (P < 0.05). The temperature-induced acceleration of decay rates was most pronounced for Soil E steroids and the k value increased from 0.049 ± 0.005 d−1 at MAT to 0.104 ± 0.019 d−1 at MAT+20°C. Temperature dependence was not discernable for the solvent extractable compounds in Soil L and n-alkanoic acids in Soil E (P > 0.05) due to a large variance associated with the k values.
Table 1. Model Fitting Parameters of SOM Components in Grassland Soilsa
Lignin V Units
Lignin S Units
Lignin C Units
Plus or minus standard error.
%labile = Clabile/(Clabile + Cstable) × 100%.
Derived from exponential model fitting, and MAT+12°C for Soil E and MAT+8°C for Soil L are close to 15°C.
3.3. Decomposition of Suberin- and Sutin-Derived Compounds
 Suberin- and cutin-derived compounds were extracted from the soil by base hydrolysis, which cleaves ester bonds that are dominant in both biomolecules [Riederer et al., 1993]. Suberin- and cutin-derived compounds were summarized and calculated based on structural parameters developed by Otto and Simpson . Suberin-derived compounds (∑S) include ω-hydroxyalkanoic acids in the range of C20 − C32, n-alkane-α, ω-dioic acids in the range of C20 − C32, and 9,10-epoxy-α, ω-dioic C18 acid. Cutin-derived compounds (∑C) included midchain hydroxyalkanoic C14, C15, C17 acids, mono- and dihydroxyalkanoic C16 acids and α, ω-dioic acids. Similar to the solvent extractable compounds, these compounds preserved the structures of their original biomolecules and are not decomposition products of other SOM components. These suberin- or cutin-derived compounds have uniform degradation patterns in the environment because microbial decomposition does not discriminate individual compounds from the same source [Riederer et al., 1993; Otto and Simpson, 2006]. Therefore, bulk suberin or cutin can be represented quantitatively by their summed biomarkers (i.e.: ∑S or ∑C).
 The OC-normalized concentrations of suberin- and cutin-derived compounds were plotted versus time (Figure 4). Soil L had a much higher concentration of suberin- and cutin-derived compounds than Soil E during the incubation. The decomposition of suberin-derived compounds in Soil E followed the exponential decay model (Figure 4a) with the decay rates ranging from 0.006 ± 0.003 d−1 at MAT to 0.025 ± 0.007 d−1 at MAT+20°C, and an average Cstable of 1.41 ± 0.15 mg/g OC and a Clabile of 2.10 ± 0.17 mg/g OC. However, suberin-derived compounds in Soil L and cutin-derived compounds in both soils did not fit the exponential decay model well, and hence, fitting parameters were not calculated. The decomposition of suberin- and cutin-derived compounds was alternatively assessed using geochemical degradation parameters, such as ω − C16/∑C16 and ω − C18/∑C18, where ∑C16 or ∑C18 includes ω-hydroxyalkanoic acid, n-alkane-α, ω-dioic acid, and midchain-substituted acids with 16 or 18 carbons, respectively [Goni and Hedges, 1990; Otto and Simpson, 2006]. Both parameters have been reported to increase with progressing cutin degradation in marine sediments [Goni and Hedges, 1990] and with soil depth [Otto and Simpson, 2006; Feng and Simpson, 2007] because cutin acids containing double bonds and more than one hydroxyl group are preferentially degraded compared to ω-hydroxyalkanoic acids. In this study, the ratio of ω − C16/∑C16 stabilized around 0.3 and 0.4 for Soils E and L, respectively (Figures 5a and 5b). However, the ratio of ω − C18/∑C18 fluctuated during incubation with no discernable pattern (Figures 5c and 5d). Suberin-derived compounds degraded faster than cutin-derived compounds in this study, and is evidenced by a decreasing ratio of suberin to cutin with time in both soils (Figures 5e and 5f; suberin/cutin = (∑S + ∑SvC)/(∑C + ∑SvC), where ∑SvC = ω-hydroxyalkanoic C16, C18 acids + di- and trihydroxyalkanoic C18 acids + 9,10-epoxy-ω-hydroxyalkanoic C18 acid + n-alkane-α, ω-dioic C16, C18 acids [Otto and Simpson, 2006]). The suberin/cutin ratios were similar at the start of the incubation study (∼4.4), and declined to 2.0 in Soil E and 2.6 in Soil L by the end of the experiment (Figures 5e and 5f). The degradation of suberin- and cutin-derived compounds in soil has been reported to exhibit the preferential decomposition of midchain hydroxy and epoxy acids (∑Mid, including 7- or 8-hydroxy-1,16-dioic C16 acid, 10,16-dihydroxy C16 acid, 9,10,18-trihydroxy C18 acid, 9,10-dihydroxy-1,18-dioic C18 acid, 9,10-epoxy-18-hydroxy C18 acid) relative to total suberin- and cutin-derived acids (∑SC = ∑S + ∑C + ∑SvC [Otto and Simpson, 2006]). In this study, the ratio of ∑Mid/∑SC increased with incubation time in Soil E (Figure 5g), which changed from 0.1 to around 0.3 after 1 year of incubation. This trend was less prevalent in Soil L (Figure 5h).
3.4. Decomposition of Lignin-Derived Compounds
 Lignin-derived compounds were extracted from the soil by CuO oxidation, which cleaves aryl ether bonds and releases phenolic monomers from the outer part of the lignin biopolymer. Lignin monomers are indicative of lignin composition and degree of oxidation [Hedges and Ertel, 1982; Kogel, 1986; Goni and Hedges, 1992]. Depending on the number and position of methoxy groups on the phenol ring, lignin monomers extracted from both soils were categorized as: vanillyls (V; vanillin, acetovanillone, and vanillic acid), syringyls (S; syringaldehyde, acetosyringone, and syringic acid), and cinnamyls (C; p-coumaric acid, and ferulic acid). The sum of monomers (VSC) decayed exponentially in both soils (Figures 6d–6f). Similar to the results of the solvent extractable compounds, the concentrations of stable and labile VSC did not vary between different incubation temperatures (P > 0.05; Table 1). A slightly lower percentage of VSC was classified into the labile pool (50–69%; Table 1), compared with solvent extractable compounds in both soils. The decay rates of VSC also increased with increasing temperature, but the trend was not significant due to high variance associated with model parameters (Table 1). Nevertheless, VSC in the “labile” pool showed a relatively slow decay rate in both soils (0.002–0.010 d−1 at MAT+12°C for Soil E and MAT+8°C for Soil L; Table 1).
 Lignin degradation was further examined by the acid/aldehyde (Ad/Al) ratios of V and S units, which has been reported to increase with an increasing degree of lignin oxidation in sediments and soils and hence serves as a geochemical indicator of the stage of lignin degradation [Hedges et al., 1988; Opsahl and Benner, 1995; Otto et al., 2005]. The ratio of vanillic acid to vanillin, (Ad/Al)v, increased with incubation time from ∼2.7 to ∼4.2 in Soil E and from ∼1.7 to ∼2.8 in Soil L (Figures 7a and 7b, P < 0.10), but the ratios did not differ at varying incubation temperatures (P > 0.05). Furthermore, the (Ad/Al)s ratio did not change during the first half of the incubation period but increased later in the study (Figures 7c and 7d). Both soils had the highest (Ad/Al)s ratio with the MAT+20°C treatment at the end of the incubation.
3.5. Response of SOM Decomposition and Microbial Respiration to Temperature Changes
 The initial microbial respiration rates in both soils are modeled by the Arrhenius equation (equation (1); R2 = 0.87 for Soil E and R2 = 0.95 for Soil L) for the incubation temperature range. The decay rates (k) of n-alkanes, n-alkanols, and steroids in Soil E and VSC units in both soils were also modeled by the Arrhenius equation (Figures 8b–8d). The quality of model fit was highest for Soil E solvent extractable compounds (0.77 < R2 < 0.87) and VSC (0.68 < R2 < 0.90), and slightly lower for Soil L VSC (R2 = 0.64 for V units, 0.55 for C units, and 0.27 for S units; Figure 8). The model fitting produced largely varied a values: 20–1100 for solvent extractable compounds in Soil E and 100–4,000,000 for VSC in both soils, and no apparent pattern existed for the a values among different classes of compounds. The activation energy (Ea) derived from model fitting is listed in Table 1, and Q10 values were calculated based on equation (2) for a fixed temperature (15°C), which was close to the average incubation temperature in this study and corresponded to the commonly used reference temperature to calculate Q10 in the literature [Reichstein et al., 2002]. VSC in both soils exhibited a higher temperature sensitivity (Q10 values: 1.33–3.45) than the n-alkanes, n-alkanols, and steroids in Soil E (Q10 values: 1.24–1.38), and lignin V units exhibited a much higher Q10 value than S and C units in both soils. VSC in Soil L had lower Q10 values than the corresponding lignin monomers in Soil E. Calculated Q10 values from microbial respiration data were found to be 1.86 for Soil E and 2.49 for Soil L at 15°C. Unfortunately, we were unable to calculate Q10 values for solvent extractable compounds in Soil L and suberin- and cutin-derived compounds in both soils due to poor model fitting. Considering the high variance associated with Ea values, we feel inclined not to do statistical comparisons of the Q10 values between different compounds, and the calculated Q10 values here represent an estimate of the “apparent temperature sensitivities” of various SOM structures partitioned into the labile pool.
4.1. Decompositional Patterns of SOM Components
 During the 1-year incubation, most of the respired CO2 is likely derived from labile SOM that is easily accessible to soil microbes [Leifeld and Fuhrer, 2005], such as soluble carbohydrates, small organic acids, and proteins [Gleixner et al., 2001]. By contrast, the decomposition of the SOM components analyzed in this study (solvent extractable compounds, cutin- and suberin-derived compounds, and lignin monomers) contributes only a small fraction to the soil OC loss (only about 0.01–0.02% OC loss based on the exponential decay curve), and their response to temperature changes may be concealed by the decomposition of the labile SOM components if only soil respiration is measured. It is therefore important to monitor the decomposition of those targeted compounds individually by other techniques.
 Respired CO2 from soil is derived from the mineralization of readily decomposable SOM components. The decomposition process of such compounds is predominantly controlled by microbial enzymatic activities, and hence, SOM decomposition studies that employ soil respiration data are fitted well by the exponential model [Dalias et al., 2001; Fang et al., 2005]. In comparison, the decomposition of individual SOM components is not only regulated by enzyme-catalyzed reactions but also associated with the components' insolubility, molecular architecture, and/or their interaction with minerals or humic substances that may limit the efficacies of enzymatic reactions because of physical protection [Sollins et al., 1996; Baldock and Skjemstad, 2000]. The degradation of SOM components may also be limited at reaction microsites and thus less uniform compared with bulk soil mineralization. The decomposition of individual SOM components in this study did not fit the exponential decay model as well as soil respiration data reported in the literature [Dalias et al., 2001; Fang et al., 2005] and there was a highstandard error associated with k and Ea values derived from the model fitting. The decomposition of Soil E components shows reasonable fits to the decay model (Figures 3 and 6). Soil L data model fitting does not fit as well with the lowest R2 = 0.32 probably due to the high abundance of grass roots present during the time of sampling. Even though the soil was sieved and homogenized before incubation, the fine root debris may have complicated the decay patterns of SOM components through a gradual input of fresh organic matter into the soil over the course of the incubation. Therefore, the fit of the exponential decay model was generally poor in Soil L.
 Suberin- and cutin-derived compounds did not exhibit exponential decay, probably due to their recalcitrance in the soil [Gleixner et al., 2001]. Hence, their decomposition in the soil was indistinguishable during the incubation period. The fluctuation in the abundances of cutin-derived compounds was presumably affected by sample heterogeneity and/or their interactions with mineral surfaces due to selective sorption of polymethylene carbon (the dominant structure in cutin) [Feng et al., 2005], which may provide physical protection against microbial enzymatic attack for cutin-derived compounds [Baldock and Skjemstad, 2000]. The degradation patterns of suberin- and cutin-derived compounds in SOM therefore suggest that the decomposition of specific SOM components is more complicated and may not conform to that of the bulk SOM.
4.2. Recalcitrance of SOM Components
 To assess the stability or recalcitrance of SOM components, the decay rate (k) derived from the decomposition data at the same temperature (MAT+12°C for Soil E and MAT+8°C for Soil L) and the percentage of SOM components in the labile pool were compared (Table 1). The decay rate derived from the exponential model (equation (5)) corresponds to a turnover time of 17–100 d for the solvent extractable compounds and 100–500 d for lignin monomers, and reflects the high decomposability or activity of the “labile” pool of SOM component. The turnover time of the “stable” SOM pool is assumed to be longer than 20 years [Chapin et al., 2002], and therefore the decay rates of the “stable” SOM pool were not modeled due to the limited duration of the incubation. Lignin monomers in the “labile” pool exhibited the lowest decay rates and a slightly lower percentage of VSCs were classified in the labile pool. This kinetic evidence supports the generally assumed slow turnover of lignin compounds in climate models [Gleixner et al., 2001; Davidson and Janssens, 2006]. However, it is difficult to compare the stability of individual lignin monomers (i.e., vanillyl versus syringyl versus cinnamyl) using the kinetic parameters due to a high variance associated with the model results (Table 1). Lignin S and C units have been reported to degrade faster than V units in the environment [Hedges et al., 1988; Opsahl and Benner, 1995; Otto et al., 2005]. Hence, V units are considered to be the most recalcitrant of the lignin monomers. This was corroborated by a faster degradation of S units in comparison to V units during the incubation, where the accelerated degradation of S units at higher temperatures, demonstrated by a higher (Ad/Al)s ratio at MAT+20°C, is more pronounced than that of V units (Figure 7).
 Suberin-derived compounds in Soil E have a slower decay rate and a lower percentage in the labile pool as compared with solvent extractable compounds (Table 1), suggesting that chemically bound soil lipids are more recalcitrant than soil lipids in the “free” form, even when they have similar chemical structures, such as aliphatic acids with 20–32 carbons. The solvent extractable compounds have similar decay rates in both soils (except steroids), but a slightly lower percentage of Soil L lipids are in the labile pool. As mentioned previously, Soil L contained fine root debris, which may have resulted in an underestimation of Soil L decomposition by releasing “fresh” organic matter into the soil. Both soil samples have similar textures and mineralogy [Feng and Simpson, 2007] but Soil L may have more mineral surfaces available for SOM binding due to its low OC content (2.69%) in comparison with Soil E (4.85%). Therefore, a larger fraction of SOM may be associated with minerals in Soil L and partitions into the “stable” pool. Among the solvent extractable compounds, steroids had the highest decay rate in Soil E and the lowest decay rates in Soil L. Cyclic soil lipids (such as steroids) have been observed to be preferentially preserved in soils as compared to aliphatic lipids (such as n-alkanes, n-alkanols, and n-alkanoic acids [Otto and Simpson, 2005]). The contrasting decay rates of steroids in the two soils suggest that environmental factors may play a part in regulating SOM decomposition in different soils, such as the incorporation of cyclic lipids into humic substances [van Bergen et al., 1997] and/or interactions with soil minerals [Ambles et al., 1994; Baldock and Skjemstad, 2000]. We hypothesize that a larger fraction of Soil L steroids may be associated with minerals and hence are protected to a greater extent from degradation in comparison to steroids in Soil E. Alternatively, steroids in Soil L may be more difficult to break down because they are in a higher degradation stage at the start of the incubation as evidenced by the lower ratio of precursor sterols (β-sitosterol and stigmasterol) to their degradation product (sitosterone) in Soil L (2.6) as compared with that in Soil E (6.5).
 Owing to poor fitting of the exponential decay model, cutin-derived compounds are not included in the comparison of kinetic parameters. However, cutin has been reported to be an important, recalcitrant component of SOM [Gleixner et al., 2001]. The cutin degradation parameter (ω − C16/∑C16) in both soils confirms the stability of cutin-derived compounds (Figures 5a and 5b). The fluctuation in the ω − C18/∑C18 ratio (Figures 5c and 5d) was likely due to the preferential degradation of ω-hydroxyalkanoic C18 acid with one double bond [Otto and Simpson, 2006] that was detected together with saturated C18n-alkanoic acid in both soils. Suberin has been reported to be more resistant to degradation than cutin because it has a high content of phenolic units and is embedded in bark and root tissues [Kolattukudy, 1981; Riederer et al., 1993]. However, the aliphatic components of suberin were observed to degrade faster than cutin-derived compounds in our soil incubation study, evidenced by a decreasing ratio of suberin/cutin with time in both soils (Figures 5e and 5f). Because cutin is only derived from aboveground sources [Riederer et al., 1993], cutin-derived compounds may have undergone degradation before they became incorporated into SOM. By contrast, suberin-derived compounds in the soil mainly originate freshly from root tissues. Consequently, cutin-derived compounds may be at a higher stage of degradation than suberin-derived compounds in SOM and hence are more recalcitrant in mineral soils. Therefore, the increasing ratio of ∑Mid/∑SC in Soil E (Figure 5g) likely results from a faster degradation of suberin relative to cutin because the ratio of ∑Mid/∑SC is low in root tissues and high in fresh vegetation that is rich in cutin-derived compounds [Otto and Simpson, 2006]. This observation suggests caution in the interpretation of suberin and cutin degradation data in the soil where suberin stability may be overestimated by a fresh input from root tissues.
4.3. Temperature Sensitivity of SOM Components
 As discussed previously, the degradation of individual SOM components is governed by their interaction with soil minerals and availability to soil microbes as well as their intrinsic structure. A substantial fraction of SOM may be associated with soil minerals and partition into the “stable” pool. Therefore, the Q10 values we calculated here are an estimate of the SOM components' “apparent temperature sensitivities” that reflect the availability of soil substrates to microbial degraders, rather than the “intrinsic temperature sensitivities” [Davidson and Janssens, 2006]. The applicability of our results therefore needs to be tested on a broader scale because the SOM-mineral interactions and microbial communities may differ among different types of soils.
 The temperature sensitivity of decomposition varies greatly among the SOM components in the same soil (Q10 of 1.24–3.45 in Soil E), suggesting heterogeneity in SOM properties and the varying responses of individual SOM compounds to global warming. Lignin monomers exhibited higher Q10 values than the solvent extractable compounds (Table 1), which is consistent with the Arrhenius theory that indicates that the decomposition of more recalcitrant compounds is more sensitive to temperature [Davidson and Janssens, 2006]. In particular, V units that are considered to be the most recalcitrant among lignin monomers showed a much higher Q10 value than S and C units in both soils. VSC in Soil L had lower Q10 values than the corresponding units in Soil E, likely because lignin is at an advanced stage of degradation (in a more recalcitrant form) in Soil E, evidenced by the higher (Ad/Al) ratios of V and S units (Figure 7).
 Soil respiration, which results from mineralization of bulk SOM, resulted in a Q10 value of 1.86 for Soil E and 2.49 for Soil L. Because the temperature sensitivity of soil respiration was measured at the start of the incubation, where labile SOM was presumably more abundant, temperature responses may have been underestimated [Dalias et al., 2001]. Ideally, Q10 values measured at the end of the incubation are better indicators of the temperature response of SOM when most of the labile substrates are exhausted during incubation. Unfortunately, the respiration rates measured at the end of this incubation study are not usable for the calculation of Q10 values because soils incubated in parallel at different temperatures may have varying amounts of labile SOM after the 1-year incubation. However, Q10 values measured sequentially between 15 and 35°C over the course of a long-term incubation (707 d at 25°C) show no significant temporal change [Leifeld and Fuhrer, 2005]. We therefore assume that the Q10 values we calculated represent a fair estimate of the temperature response of soil respiration over the course of the experiment. Nevertheless, Q10 values derived from soil respiration data only represent the average kinetic properties of heterogeneous SOM structures, but not the greatly varied responses to temperature increases for individual SOM components. Because the reaction rate of recalcitrant SOM (such as lignin monomers) is much slower than that of the less stable SOM (such as solvent extractable compounds), changes in k values of recalcitrant SOM are likely to be concealed by the responses of labile SOM when the mineralization of both components is measured simultaneously. Notably, the Q10 values for the refractory lignin units (VSC) are lower (1.33–2.00) than the Q10 value for total respiration (2.49) in Soil L. Again, the calculated “apparent temperature sensitivities” of VSC units reflect the high availability of lignin structures in the “labile” pool to soil organisms rather than their structural recalcitrance.
 The Q10 values derived from bulk SOM mineralization are unrelated to the degradation stage of individual SOM components. In this study, lignin-derived compounds in Soil E were more oxidized (higher (Ad/Al) ratios) and possessed a higher Q10 value than those in Soil L. However, Soil L respiration had a much higher Q10 value (2.49) than Soil E (1.86) probably because recalcitrant SOM such as suberin-derived compounds, cutin-derived compounds, and lignin monomers comprised a smaller fraction of the identified SOM in Soil E (86%) in comparison to Soil L (92%) at the beginning of incubation. Even more labile SOM such as proteins and carbohydrates that were not included in this study may also confound the analysis and the high temperature sensitivity of the recalcitrant SOM pool in Soil E may be concealed by the respiration data. Similar temperature sensitivities of SOM mineralization has been reported for soils with presumably different recalcitrance [Fang et al., 2005]. However, the recalcitrance of SOM is usually assessed by the content of operationally defined SOM fractions, such as water-dissolved carbon, or K2SO4-extracted carbon, which contains a mixture of heterogeneous SOM structures and may not be an accurate indicator of the recalcitrance of SOM.
 We thank Henry Janzen for assistance with selecting and sampling soil L. Funding from the Canadian Foundation for Climate and Atmospheric Sciences (GR-520) is gratefully acknowledged. Leah Nielsen is thanked for conducting part of the chemical extractions. The Natural Sciences and Engineering Research Council (NSERC) of Canada is thanked for support via a University Faculty Award (UFA) to M. Simpson and an undergraduate summer research award (USRA) to L. Nielsen. X. Feng acknowledges funding from the Ontario Graduate Scholarship (OGS) program.