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- Materials and Methods
- Supporting Information
Microbial decomposition of plant litter and soil organic matter sustains ecosystem productivity by cycling carbon, nitrogen, and other nutrients. Decomposition, in turn, is primarily controlled by climate and plant litter chemistry. The climate that sustains plant growth and decomposition is rapidly changing; Earth’s average surface temperature is projected to increase by 1.4–5.8°C by the end of this century (IPCC, 2007). At the same time, rainfall events are expected to become less frequent and more intense, resulting in longer, more frequent periods of drought. These changes could directly affect ecosystem nutrient cycling by affecting the chemical composition and thus the decomposability of litter produced. As the efflux of CO2 through microbial decomposition of organic matter is a significant component of the global carbon cycle (Davidson et al., 2000), the climate-induced change in litter chemistry could alter the global carbon budget as well.
Polyphenols, which are synthesized through the phenylpropanoid pathway, represent a diverse and the most abundant class of plant secondary compounds. They are involved in many ecological and physiological functions in plants, including defenses against pathogens and herbivores, lignification, pigmentation, pollination and plant–plant interactions (Dixon et al., 2005). Tannins constitute the second most abundant polyphenolics in vascular plant species after lignin, and are characterized by their capacity to interact with proteins. Structurally, tannins can be divided into two major classes – condensed tannins (CTs) and hydolyzable tannins (HTs; Kraus et al., 2003a). Condensed tannins or proanthocyanidins are produced by both gymnosperms and angiosperms, and are polymeric flavanoids commonly linked by a C–C interflavan bond at C-4–C-8 or C-4–C-6 between the flavan-3-ol monomers (Fig. 1a). Proanthocyanidins can be further subdivided based on their B-ring hydroxylation pattern, with procyanidins having a di-hydroxy and prodelphinidins a tri-hydroxy B-ring. Evolutionarily, hydrolyzable tannins are more advanced than condensed tannins, and are limited to relatively advanced dicotyledonous plant families (Kraus et al., 2003a). Hydrolyzable tannins are complex esters of gallic acid with glucose (Fig. 1b), and based on the presence and absence of intramolecular C–C coupling between the galloyl groups, they are further divided as gallotannins and ellagitannins, respectively (Fig. 1b,c).
In plants, the tannin concentration is highly variable among species, while within species it varies with the age and tissues (Schweitzer et al., 2008). Environmental factors such as nutrient availability, drought, pH, herbivory, ozone and CO2 concentration also affect tannin concentrations (Herms & Mattson, 1992; Bussotti et al., 1998; Kraus et al., 2003b; Jaakola & Hohtola, 2010; Lindroth, 2010). Tannins may account for up to 25% of foliar DW (Kraus et al., 2003b), and because of their limited resorption during senescence, tannins may undergo further concentration in senesced tissues, thus forming a major fraction of leaf C input to soils. Owing to their protein complexation capacity, tannins can directly influence ecosystem processes such as litter decomposition and nutrient cycling by lowering the catalytic efficacy of enzymes, as well as reducing the substrate suitability for proteolytic enzymes (Kraus et al., 2003b). Tannins act as multidentate ligands binding to proteins through their phenolic groups and/or hydrophobic regions. These interactions with proteins are mostly noncovalent, and HTs, because of their high hydrophobicity, are thought to precipitate proteins through hydrophobic interactions, whereas hydrogen bonding (between phenolic hydroxyl and peptide carbonyl groups) dominates in CT–protein complexes (Hagerman et al., 1998).
This biological activity of tannins depends more strongly on their structure than on their abundance (Kraus et al., 2003a). The reactivity of tannins is affected by the concentration of condensed vs hydrolyzable tannins, the hydroxylation pattern of the B-ring, extent of polymerization, type of cross-linkage between monomeric units, substitution pattern of the A-ring, and cis vs trans confirmation at C-2–C-3 (Kraus et al., 2003a). Thus, the influence of tannin quantity on nitrogen mineralization is not straightforward. Many studies have found a negative relationship between the amount of tannin and soil N mineralization (Kraus et al., 2004; Nierop et al., 2006), whereas others have reported no effect (McCarty & Bremner, 1986; Schimel et al., 1996) or a positive relationship (Kanerva et al., 2006; Kanerva & Smolander, 2008). High-molecular-weight tannins cause enzyme/substrate precipitation (Bradley et al., 2000), whereas low-molecular-weight tannins directly affect microbial metabolism through toxicity (Schimel et al., 1996; Fierer et al., 2001). Some low-molecular-weight tannins are also rapidly utilized by microbes as a C source, which results in microbial immobilization of N (Kraus et al., 2004). Thus, depending on the tannin quality the decrease in N mineralization could be caused by various mechanisms. As the biological function of tannin is strictly structure-dependent, it is difficult to generalize about how a given quantity of tannins will influence ecosystem processes (Kraus et al., 2003a,b, 2004; Hernes & Hedges, 2004; Nierop et al., 2006).
Although tannin quantities are known to respond to environmental stimuli, very little is known about how climate change will affect tannin structure, and consequently biological reactivity. Few studies have looked at the effects of changes in CO2 concentration on litter chemistry. Liu et al. (2005, 2009) found no effect of elevated CO2 on concentrations of soluble sugars, phenolic and condensed tannins in Aspen, and elevated CO2 had little effect on the decomposability of the litter produced (Norby et al., 2001). Even in the absence of quantitative changes, climatic stress, by altering the phenylpropanoid pathway, could cause structural variations in tannins. Also, the response of plants to CO2 has been shown to be limited by N availability (Norby et al., 2010), and soil moisture was reported to be the most important factor controlling soil carbon dynamics in a constructed old-field experiment also manipulating temperature and CO2 (Garten et al., 2009).
The main objectives of this study were to ascertain the influence of predicted warming and altered precipitation on the composition and structural chemistry of tannins, and to quantify the corresponding changes in their biological reactivity. We hypothesized that increased climatic stress resulting from either drought or increased temperatures, by modifying the phenylpropanoid pathway, would induce the production of polyphenols that are structurally different from the constitutive polyphenols. We expected that the differences in structural chemistry would make these stress-induced polyphenols more reactive. Understanding how warming and altered precipitation can affect tannin composition and structure will help us predict how litter decomposition and nutrient dynamics may change in a warmer world.
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- Materials and Methods
- Supporting Information
Balanced availability of nutrients is required for optimal plant growth; changes in the nutrient acquisition ability of a plant or mineralization rates in the soil can affect the metabolite composition of the plants. In this study, warming and precipitation change caused both qualitative and quantitative changes in the chemistry of Acer leaf litter. Tannins in leaf litter from the most water-stressed treatment (drought + warming) underwent structural changes, causing them to be more readily extractable and five times more biologically reactive. Few other studies have examined how the structural chemistry of tannins in leaf litter responds to predicted climatic changes. The dramatic changes we observed could be ecologically significant.
Using FTIR, we found that warming and precipitation altered Acer leaf litter chemistry (Fig. 4). Together, drought and warming caused the greatest water stress for plants, and resulted in higher relative concentrations of cellulose and plant defense compounds, such as flavonoids, cutin, and waxes. Similarly, warming increased abundance of waxes, cutin, and polyphenols across all precipitation treatments. Thus, warmer, more water-stressed conditions led to a greater abundance of protective compounds in leaf litter. Polyphenols, including flavonoids, regulate oxidative stress in plants (Hernandez & Van Breusegem, 2010) and act as an herbivore deterrent (Degabriel et al., 2009), whereas leaf waxes can reduce water loss (Aharoni et al., 2004).
Tannin abundance and quality
Acer leaf litter contained a mixture of HTs and CTs (Fig. 5; Bate-Smith, 1978), with CTs predominating across all climate treatments (Fig. 5). The climate treatments affected leaf tannin profiles of Acer, most dramatically in the drought + warming treatment, which caused a doubling of total tannin content, to 30% of leaf DW (Fig. 6a). This change in tannin profile could result from changes in the phenylpropanoid pathway that would help the plant to mitigate water stress. Under nutrient deficiency or drought, plant growth can slow before C assimilation declines (Herms & Mattson, 1992). Under these conditions, flavonoid biosynthesis has been proposed as a pathway for excess energy dissipation and carbon diversion by plants (Hernandez & Van Breusegem, 2010), thus reducing the production of reactive oxygen species. Further, because of their H-atom transfer or one-electron transfer mechanism, flavonoids can quench reactive oxygen species, protecting plant tissues from peroxidation damage (Leopoldini et al., 2004). Flavonoids also protect against herbivory as a result of their protein complexation and pro-oxidant capacities, and so help the plant defend previously acquired resources (Herms & Mattson, 1992; Wright et al., 2010). We observed a decrease in leaf litter CTs in the treatments providing optimal growth conditions (warmed and wet; Fig. 5c), suggesting lower investment in these defense compounds under the best growth conditions.
Warming and precipitation change affected the quality of CT; the litter of foliage exposed to drought + warming produced CT with a high relative abundance of procyanidins (dihydroxy B-ring) that were less polymerized (Table 2). The hydroxylation pattern of the B-ring is primarily governed by the activity of various plant enzymes, including flavonoid 3′-hydroxylase and flavonoid 3′, 5′-hydroxylase (Menting et al., 1994). Owing to the temperature regulation of enzyme induction and activity, cooler temperatures favor flavonoids with higher degrees of hydroxylation (prodelphinidins, Jaakola & Hohtola, 2010), which could in turn result in the observed higher prodelphinidin content of unwarmed treatments. The condensation of monomeric flavan-3-ols, though less understood, is also under strict enzymatic control. Polyphenol oxidases catalyze the conversion of flavan-3-ols to their respective quinones, which are then converted to carbocations. Oligomeric proanthocyanidins are formed following a nucleophilic attack by C-6 or C-8 of catechins on these carbocations (Dixon et al., 2005). In plants, the down-regulation of polyphenol oxidases is associated with an increased drought tolerance (Thipyapong et al., 2004). Given their significant role in proanthocyanidin biosynthesis, this down-regulation of polyphenol oxidases could contribute to the lower degree of polymerization under climatic stress. Changes in condensation patterns can also occur post-enzymatically, when mild acid conditions could result in cleavage of interflavanic bonds, resulting in a reduction in the degree of polymerization (Vidal et al., 2002). The cellular pH of higher plants can become more acidic at higher temperature (Aducci et al., 1982), which could thus reduce the degree of polymerization of CTs in the warmed treatments (Table 2). Further, a reduction of CT polymerization could explain, in part, the low yield of purified tannins from warmed treatments in acid-butanol assay (Table 2). During the oxidative acid-butanol depolymerization of CT, only the flavones at the extender position yield colored anthocyanidins – not the terminal catechin units. Hence, CT that predominantly consists of shorter chains will result in lower color yield, because the terminal units are not detected (Kraus et al., 2003a).
In the drought + warming treatment, the total quantity of CT increased but the amount of CT sequestered to fiber did not increase (data not shown), resulting in a reduced fraction of bound CT (Fig. 5d). In green leaves, tannins are confined to vacuoles (Marles et al., 2003) and sequestered away from the normal cell metabolism. However, during leaf senescence, following vacuole collapse the tannins could interact with cell wall components and could be irreversibly bound. Interaction of tannins with cell walls can occur during the active growth stage as well (Gagne et al., 2006). The smaller fraction of bound CT in the drought + warming treatment (Fig. 5d) could be partially explained by their lower degree of polymerization (Table 2), which would in turn decrease their interaction with cell wall components (Bindon et al., 2010). Phenolic hydroxyl groups of tannins could associate with carboxylic groups of cellulose through hydrogen bonding. The affinity of such associations has been shown to decrease with an increase in temperature because of preferential sorption of tannins to highly energetic sites of cellulose (Espinosajimenez et al., 1987) and this could potentially reduce tannin sequestration in warmed treatments. Also, the CT in droughted + warmed treatments had a higher proportion of less reactive dihydroxy B-rings that have a lesser propensity for forming B-ring quinones (Nierop et al., 2006). This would decrease the association of stressed tannins with cell-wall components. Litter from unwarmed treatments had relatively high contents of pectin (Fig. 4), which is a major component in cell walls that could sequester CT (Bindon et al., 2010). This could explain the high abundance of fiber-bound tannins in unwarmed treatments (Fig. 5d). Unlike CT, leaf mesophyll cell walls are the primary sites of synthesis and deposition of HTs (Grundhofer et al., 2001), and this difference in subcellular localization could result in a greater proportion of HTs being readily extractable.
Biological reactivity of litter tannins
The drought + warming treatment resulted in a doubling of CTs and HTs in litter (Fig. 5), but a fivefold increase in glucosidase complexation capacity of the litter (Fig. 6d), highlighting the importance of tannin quality in determining interactions with proteins. The protein complexation capacity of tannins is governed by the structure of both tannins and proteins. The main attributes of tannins that govern their complexation capacities are their molecular size (higher molecular size, resulting from better cross-linking, results in higher complexation capacity), conformational flexibility (higher flexibility, by providing better accessibility to the binding sites, results in better complexation capacity) and water solubility (higher hydrophobicity, by decreasing the hydration shell around molecules, will result in higher complexation; Spencer et al., 1988).
In our study, despite the lower degree of polymerization of the CT fraction (Table 2), the highest protein complexation capacities were found in litter from the drought treatment, whether unwarmed or warmed (Fig. 6c,d). This could be explained by two mechanisms. First, there is a trade-off between the degree of polymerization and the conformational freedom of a molecule. CTs with a higher degree of polymerization would experience structural rigidity as a result of restricted rotation about the repeating 4–8 or 4–6 interflavan bonds (Fletcher et al., 1976; Spencer et al., 1988). Thus, polymerization beyond a threshold level could negatively affect the conformational freedom of tannins (possible steric hindrance of 3′, 4′-dihydroxyphenyl groups; Haslam, 1974), thereby decreasing their binding capacity to proteins. Second, water-stressed treatments had the highest concentration of HTs during purification (Table 2). This could be attributed to the lower degree of polymerization of CT in these treatments (Table 2), which could result in lower binding of these compounds to the resins and subsequent loss during the methanol-wash step. The preferential concentration of HT during purification indirectly indicates a higher degree of polymerization of HT fractions in the stressed treatments, which could result in their greater influence in protein complexation capacity.
Acer produces a mixture of HTs and CTs (Bate-Smith, 1978), and the concentration of the extractable forms in the drought + warming treatment was approximately double that of the control treatment (Fig. 6b). In our study, the tannic acid precipitated more protein on a per-weight basis than Acer tannin from any of the treatments. Based on the NMR spectra and the acid-butanol reaction, the commercial tannic acid used in this study did not have any detectable CT. A higher protein complexation capacity of tannic acid that far exceeds the complexation capacity of pure CT samples, and samples with a mixture of CTs and HTs have been reported before (Kraus et al., 2003a). Depending on its structural flexibility, HT may have increased the protein complexation ability of our samples as much or more than CT, since structural rigidity could hinder the protein interactions of highly polymerized CT.
The Acer litter consistently had higher glucosidase complexation capacity than BSA complexation capacity (Fig. 6c,d). This could be explained by structural differences between the two proteins that could contribute to their reactivity. BSA is a globular protein whose structure, which can be approximated as prolate to oblate spheroids (Jachimska et al., 2008) with limited structural flexibility, causes steric hindrance to protein binding. Glucosidase has an extended random coil confirmation, which increases accessibility to its peptide linkages and thus offers more binding surface to tannins. Similar differences in the reactivity of tannins to protein quality have been reported before. For example, Deaville et al. (2007) reported that the binding affinity of conformationally restrained ellagitannins to flexible gelatin was four times greater than to structurally rigid BSA. Thus, the fivefold increase in β-glucosidase precipitation capacity of litter shown in our study could be partially indicative of the structural flexibility of tannins from climatic stress treatments. Although CT with a low degree of polymerization is theoretically considered to be more labile than its long-chain counterpart, recent studies have shown that short- and long-chained CTs have similar recalcitrance (Kraus et al., 2004). Our study demonstrates that warming and altered precipitation can alter the composition and structure of Acer tannins, with implications for their reactivity.
Although climatic changes eventually restructure plant communities, responses of individual species can strongly influence ecosystem functioning in the intermediate term. Where plant communities change slowly, changes in litter chemistry of individual forest species may affect biogeochemical cycling more strongly than changes in species composition. To our knowledge, this is the first study to examine the changes in leaf litter chemistry of plants exposed to simulated warming and precipitation changes. Tannin quantities have previously been shown to respond to environmental stimuli. However, our results, by profiling the subclasses of tannins, depict this variation at a finer level and identify consequences of these changes for biological reactivity. This study clearly documents variations in allocation to foliar compounds and strong changes in the quality and reactivity of tannins induced by climatic stresses. Our results in Acer suggest that tannins produced under climatic stress can have higher enzyme complexation capacities, and hence the quantity alone cannot be taken as a benchmark for predicting the ecosystem properties of tannins. Higher reactivity could inhibit litter decomposition by immobilizing the microbial enzymes that catalyze the catabolic reactions during decomposition, and by protecting protein substrates from proteolytic enzymes. Similarly, the higher mobility and reactivity of stress-induced tannins could affect the decomposability of resident soil organic matter. This would mean decreased nitrogen availability to plants, potentially weakening net terrestrial carbon uptake. If these mechanisms operate similarly across many species, this mechanism could have widespread consequences for nutrient cycling and soil carbon sequestration, potentially providing an important medium-term feedback to climate change.