Long-term litter decomposition controlled by manganese redox cycling

Marco Keiluweit; Peter Nico; Mark E. Harmon; Jingdong Mao; Jennifer Pett-Ridge; Markus Kleber

doi:10.1073/pnas.1508945112

Edited by François M. M. Morel, Princeton University, Princeton, NJ, and approved August 18, 2015 (received for review May 8, 2015)

Significance

The rate-controlling mechanisms of litter decomposition are of fundamental importance for ecosystem nutrient cycling, productivity, and net carbon (C) balance. Current C cycling models rely primarily on climatic factors and lignin content as the main predictors of litter decomposition rates. Here, we show how the ability of the integrated plant–soil system to promote active redox cycling of manganese (Mn) regulates litter decomposition. Our work suggests that incorporating the coupling of litter decomposition and other elemental cycles, such as the Mn cycle, into conceptual and numerical models may significantly improve our mechanistic understanding and predictions of C cycling in terrestrial ecosystems.

Abstract

Litter decomposition is a keystone ecosystem process impacting nutrient cycling and productivity, soil properties, and the terrestrial carbon (C) balance, but the factors regulating decomposition rate are still poorly understood. Traditional models assume that the rate is controlled by litter quality, relying on parameters such as lignin content as predictors. However, a strong correlation has been observed between the manganese (Mn) content of litter and decomposition rates across a variety of forest ecosystems. Here, we show that long-term litter decomposition in forest ecosystems is tightly coupled to Mn redox cycling. Over 7 years of litter decomposition, microbial transformation of litter was paralleled by variations in Mn oxidation state and concentration. A detailed chemical imaging analysis of the litter revealed that fungi recruit and redistribute unreactive Mn²⁺ provided by fresh plant litter to produce oxidative Mn³⁺ species at sites of active decay, with Mn eventually accumulating as insoluble Mn^3+/4+ oxides. Formation of reactive Mn³⁺ species coincided with the generation of aromatic oxidation products, providing direct proof of the previously posited role of Mn³⁺-based oxidizers in the breakdown of litter. Our results suggest that the litter-decomposing machinery at our coniferous forest site depends on the ability of plants and microbes to supply, accumulate, and regenerate short-lived Mn³⁺ species in the litter layer. This observation indicates that biogeochemical constraints on bioavailability, mobility, and reactivity of Mn in the plant–soil system may have a profound impact on litter decomposition rates.

Decomposition of above-ground plant detritus (litter) is a fundamental process regulating the release of nutrients for plant growth and the formation of soil organic matter (SOM) in forest ecosystems (1). Litter decomposition regulates the proportion of litter-derived carbon (C) that is either retained in the system as SOM or lost as CO₂ (2), thereby influencing net C storage in soils. Although even small decomposition rate increases may accelerate climate change by virtue of increasing CO₂ emissions from soils (3), uncertainty persists over the rate-controlling mechanisms (4, 5).

Litter decomposition rates are strongly influenced by climatic factors (e.g., temperature and moisture) and have long been linked to litter chemistry, specifically lignin content (6). Lignin—an aromatic biopolymer—often makes up between 15% and 40% of the litter mass and is concentrated in cell walls (7). It encrusts cellulose microfibrils to form protective physical units (“ligno–cellulose complexes”) that are embedded in a matrix of hemicellulose. Conventional thinking suggests that the relatively high initial litter decomposition rates are due to the preferential use of soluble and readily accessible polysaccharides and hemicelluloses relative to lignin components. In later stages, decomposition slows when litter has become enriched in lignin and any remaining celluloses and hemicelluloses remain enclosed and protected in ligno–cellulose complexes (8). However, more recent research suggests that lignin readily decomposes in the presence of dissolved organic substrates (5). This effect was attributed to dissolved and readily assimilable substrates providing decomposer organisms with the extra energy required for the cometabolic breakdown of lignin. This observation suggests that litter decomposition may be controlled not only by litter chemistry, but also by the availability of key resources for the efficient microbial breakdown of individual biopolymers.

One such key resource seems to be manganese (Mn). Numerous studies (9 ⇓⇓⇓⇓–14) have measured a strong positive relationship between Mn content of litter and litter decomposition (expressed as % mass loss) in boreal, temperate, and semiarid forest ecosystems. Among all other parameters tested (water soluble C, lignin, N, P, K, Ca, and Mg contents), Mn content best predicted litter mass loss, leading Berg et al. (10) to conclude that “Mn concentration is the single main factor” governing litter decomposition in these forest biomes. However, efforts to experimentally validate the effect of Mn availability on decomposition rates have produced ambiguous results (15) and have been hampered by the lack of explicit consideration of Mn redox cycling in the soil system.

Microbial Mn oxidation has long been implicated in the enzymatic degradation of lignin (16). Culture studies with lignin-decomposing fungi showed that trivalent Mn³⁺ is generated during the oxidation of lignin model compounds (17). It has been proposed that Mn³⁺ is formed via the oxidation of soluble Mn²⁺ by fungal exo-enzymes, such as Mn peroxidase or phenol oxidases (e.g., laccase). When stabilized in solution by chelating ligands, these soluble Mn³⁺–ligand complexes were suggested to act as diffusible and potent oxidants of lignin analogs (16). More recently, it was discovered not only that fungi can oxidize Mn enzymatically, but that a diverse range of heterotrophic bacteria isolated from terrestrial and aquatic systems can do so as well (18, 19). Despite the widespread physiological potential for microbial Mn oxidation, it is unclear to what extent the natural Mn redox cycle is coupled to organic matter degradation in forest ecosystems.

Circumstantial evidence suggests that microbial oxidation may be the main driver of Mn cycling in forest litter layers. Foliar litter is a major source of Mn compared with other plant litter and mineral soils (20). Mn in live foliage is naturally present in its reduced, mobile form (i.e., Mn²⁺), facilitating its supply to photosystem II and other enzymatic systems (21). Foliar litter becomes enriched in Mn relative to fresh plant material and further accumulates Mn during the decomposition process (9, 22). Mn in dead foliage (i.e., litter) was found to be bound in organic complexes (23), but it is unclear whether Mn is present as unreactive Mn²⁺ or the more reactive Mn³⁺ form. A recent spectroscopic investigation showed that foliar tissue contains predominantly Mn²⁺ that is gradually oxidized to Mn^3+/4+-oxides in the soil upon decomposition (24). This observation led Herndon et al. (24) to suggest that Mn stored in leaves is solubilized upon litter fall but rapidly immobilized as Mn^3+/4+ oxides after aging in the soil. Others have observed Mn-rich black precipitates on needle litter colonized by fungi (25), resembling Mn^3+/4+ oxides as observed after fungal Mn oxidation in model systems (26, 27).

Here, we investigated the impact of biotic Mn redox cycling on long-term litter decomposition in forest ecosystems, taking advantage of a field experiment conducted in an old-growth Douglas-fir forest in the Oregon Cascades. During this 7-y experiment, each year’s litterfall was spatially separated by nylon mesh placed on the forest floor. The mesh created a series of six well-confined litter layers (a “littercake”) containing Douglas-fir needles at increasing decomposition stage. Our approach was to resolve Mn transformation and litter decomposition within the forest ecosystem at two different scales—across the whole soil profile and in microenvironments on needles colonized by fungi. Across the litter layer, we hypothesized that Mn oxidation covaries with the breakdown of aromatic litter components (i.e., lignins and tannins). Changes in Mn chemistry were identified by selective extractions and X-ray absorption near edge structure (XANES) spectroscopy, whereas the alterations in the molecular composition of litter was determined using a combination of Fourier-transform infrared (FTIR) spectroscopy, NMR, and laser desorption postionization mass spectrometry (LDPI-MS). Within microenvironments on needle surfaces, we anticipated that oxidative degradation of aromatic compounds occurs where fungi produce Mn³⁺. To spatially resolve Mn oxidation state and chemical transformations in these microenvironments, we applied a multimodal chemical imaging approach that coupled microscale X-ray fluorescence (µXRF)/X-ray absorption spectroscopy (XAS) with µFTIR imaging.

Results

Mn Transformations.

With increasing litter age, Mn gradually accumulated and became more oxidized. Total Mn concentrations in decomposing needle litter progressively increased and, on dry-weight basis, became higher than in fresh needles or the underlying mineral horizons (Table S1). The contribution of pyrophosphate-extractable Mn (Mn_PYRO, Mn in soluble and organically complexed pools) increases most rapidly in the first three layers, before reaching a plateau in layer 4 (Table S1).

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Table S1.

Chemical transformations of litter over 6 y of decomposition

Concurrently, Mn XANES absorption maxima shifted from energies associated with Mn²⁺-dominated phases to Mn³⁺- and Mn⁴⁺-rich phases as litter age increased (Fig. 1). The average oxidation state of Mn calculated from these shifts progressively increased from +2.07 in fresh needles to +2.74 in layer 6 (Fig. 1). The relative amount of Mn³⁺ species increased most rapidly in layers 1 and 2 whereas that of Mn⁴⁺ species increased strongly from layers 3–6. Needles in the underlying O horizons showed another slight increase in average oxidation state (+2.83). Mn oxidation state (+3.1 and +3.0) and, consequently, the relative contributions of Mn³⁺ and Mn⁴⁺ were found to be highest in the mineral soil.

Fig. 1.

Change in Mn speciation in Douglas-fir needle litter (L) over 6 years of decomposition. Mn K-edge XANES spectra with approximate positions of absorption maxima for Mn²⁺, Mn³⁺, and Mn⁴⁺ forms (50). Pie chart Insets show the relative amounts of Mn²⁺, Mn³⁺, and Mn⁴⁺ with the adjacent number representing the average oxidation state as determined by linear combination fitting (50). Fresh needle litter (FL) and the underlying organic (O) and mineral (A and B) horizons were included for comparison.

Litter Decomposition.

Overall, we observed a strong relationship between Mn redox transformations and litter decomposition. The increase of Mn in soluble and organically complexed pools (Mn_PYRO) was significantly correlated with microbial processing of the litter. The C/N ratio, which might be expected to decrease with increased microbial processing, decreased most rapidly from layers 1–3 (Table S1). This decrease in C/N was correlated with increases in Mn_PYRO (R² = 0.76, P < 0.05) (see Fig. 3A), but not with changes in organically complexed Al, Ca, or Fe (P > 0.05). Similarly, the FTIR absorbance of amide groups, associated with microbial protein and chitin, increased (Fig. 2A) and is positively correlated with Mn_PYRO (R² = 0.94, P < 0.05) (Fig. 3B).

Fig. 2.

Chemical transformations of Douglas-fir needles over 6 y of decomposition. (A) FTIR spectra of needle litter. Absorbance of both (i) amide I and II groups associated with bacterial/fungal protein and chitin increased and (ii) C-O-C groups of polysaccharides increased with litter age whereas (iii) absorbance of COO-stretch of carboxylates and the ar-C-C stretch of aromatic lignin decreased. (B) ¹³C-NMR spectra of needle litter. Gray lines show spectra obtained by unselective CP/TOSS experiments including all carbons, whereas black lines show spectra obtained by selective CP/TOSS-DD sensitive to nonprotonated carbons and mobile carbons (e.g., -CH₃ and long-chain -(CH₂)_n-). (C) Synchrotron-LDPI mass spectra of needle litter. Fragments with m/z of 340, 370, 396, 399, 400, 406, and 414 showed ionization energies of ≤8.5 eV and were thus assigned to aromatic structures. Detailed description of peak assignments can be found in SI Materials and Methods. ∑ar⁺ denotes the sum of normalized intensities of peaks that increase with litter age (m/z = 340 and 370) whereas ∑ar⁻ represents the sum of normalized intensities of peaks decreasing over time (m/z = 396, 399, 400, 406, and 414). Mass spectra presented here were collected with a photon energy of 9.5 eV and were chosen because they showed less fragmentation and greater signal-to-noise ratio.

Fig. 3.

Covariation of litter and Mn transformations in Douglas-fir needles over 6 y of decomposition. (A) Changes in the C/N ratio across the litter layer in relation to pyrophosphate-extractable Mn (Mn_PYRO), here used as a proxy for bioavailable Mn. (B) Changes in the FTIR absorbance of amide I and II groups as a proxy for microbial protein/chitin abundance relative to variations in average Mn oxidation state. (C) Changes in litter decomposition (expressed by the ratio of saccharide/aromatic carbon absorbance in the FTIR spectra) in relation to Mn oxidation state. (D) Changes in litter decomposition (as indicated by the ratio of saccharide/aromatic carbon in ¹³C NMR spectra) in relation to Mn oxidation state. (E) Lignin transformations (expressed by the ratio of C_ar-O/C_ar-C in the ¹³C NMR spectra) in relation to average Mn oxidation state. (F) Changes in lignin transformations (expressed by the relative ratio in signal intensity of LDPI-detected aromatic mass fragments shown in Fig. 2C) in relation to average Mn oxidation state.

Differences in the litter’s molecular composition correspond with increases in Mn oxidation state across the decomposition sequence. FTIR spectra show that the relative absorbance of regions associated with saccharide and aromatic functional groups (Fig. 2A) is significantly correlated with Mn oxidation state (R² = 0.79, P < 0.05) (Fig. 3C). Similarly, relative changes in the abundance of saccharide and aromatic moieties in the ¹³C NMR spectra (Fig. 2B) are well correlated with Mn oxidation (R² = 0.98, P < 0.05) (Fig. 3D).

To evaluate a link between the chemical alteration of aromatic litter components and Mn transformations, we gathered ¹³C NMR and synchrotron-LDPI mass spectrometry data. Two different types of aromatic carbons are usually identified in ¹³C NMR spectra: protonated (C_ar-H) and quaternary nonoxygenated (C_ar-C) at 142–110 ppm, or oxygenated aromatic ring carbons (C_Ar-O) at 162–142 ppm (28). The abundance of oxygenated relative to nonoxygenated aromatic ring carbons (C_ar-O/C_ar-R, with R = C or H) in ¹³C NMR spectra, used here as a proxy for ring oxidation, was strongly correlated with changes in Mn_PYRO (R² = 0.92, P < 0.05) (Fig. 3E). Synchrotron-LDPI is a soft-ionization mass spectrometry technique that is particularly sensitive for lignin-derived compounds (29, 30). Inspection of the resulting mass spectra showed that one set of peaks at lower mass-to-charge (m/z) ratios (∑ar⁺; m/z = 340–370) gains intensity with increasing litter age whereas the intensity of a second set of peaks at higher m/z ratios (∑ar⁻, m/z = 396–414) declines (Fig. 2C). To confirm that these mass fragments (i.e., m/z = 340, 370, 396, 399, 400, 406, and 414) correspond to aromatic structures, we determined ionization energies (IEs) for each of the fragments in a second LDPI experiment as previously described (31, 32) and detailed in SI Materials and Methods. This method exploits the fact that aromatic ring structures have lower ionization energies (≤8.5 eV) than other organic moieties (>9 eV) (29, 30). Because ionization energies of these mass fragments ranged from 7.3 to 8.5 eV (Table S2), we attributed these peaks to aromatic moieties. The observed gradual transition from one set of aromatic compounds to the other (∑ar⁺/∑ar⁻) therefore points to a structural change in aromatic litter components, which showed a strong correlation with Mn oxidation state (Fig. 3F) (R² = 0.88, P < 0.05).

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Table S2.

Ionization energies for fragments associated with most prominent mass peaks and their evolution over 6 y of litter decomposition

Mn Form and Distribution on Decomposing Needle Surfaces.

Because Mn chemistry and molecular composition of the litter changed more rapidly in the initial stages of decomposition, we chose needles from layer 1 for detailed imaging analysis. These needles were colonized by fungi forming dense hyphal networks with distinct dark patches hypothesized to be Mn^3+/4+ oxides (Fig. S1A). Elemental maps of resin-embedded cross-sections from three needles showed that Mn concentrated in diffuse patches on the surface, associated either with individual hyphae or larger particles (Fig. S1B). Other major elements (e.g., Ca, Fe, and Si) were randomly distributed. Mn XANES spectra collected from diffuse patches and discrete particles on the needle surface showed a greater abundance of Mn³⁺ and Mn⁴⁺ and, consequently, a higher average oxidation state than those extracted from individual hyphae and the needle tissue (Fig. S1C).

Fig. S1.

µ-XRF/XAS analysis of whole needle cross-sections taken from the top litter layer (year 1). (A) Photographs showing individual needles colonized by fungal hyphae associated with dark patches. (B) False color XRF maps of resin-embedded needle cross-sections showing the concentration of Mn (green) dense patches, hyphae and particles in contrast to Ca (blue) and Si (red). Color intensity is directly related to elemental concentrations. The apparent 3D appearance of the map is due to the penetration of X-rays ∼100–200 µm into the resin-embedded cross-sections. (C) Mn XANES spectra of the regions highlighted in B, illustrating the more oxidized state of Mn in diffuse patches and particles associated with the needle surfaces. Pie charts show the relative amounts of Mn²⁺, Mn³⁺, and Mn⁴⁺ and the number denoting the average valence state as determined by linear combination fitting (50).

Manganese Chemistry at the Hyphae–Epidermis Interface.

Photographs of two cryo-sectioned decomposing needles show dark hyphae colonizing the epidermis region (Fig. 4A). Along this interface, µFTIR chemical imaging visually separates needle tissue rich in aromatic structures from amide-rich fungal materials (Fig. 4B). Elemental maps of this interface showed that Mn accumulates where fungal hyphae are in direct contact with the needle epidermis (Fig. 4C). We collected Mn XANES spectra from hotspots (points 3–5, Fig. 4C) along this interface (Fig. 4D). Spectral deconvolution indicated that the hotspots associated with fungi on the needle surface were more oxidized (points 3–5; average oxidation state, 2.6–2.7) than Mn in the needle interior (points 1–2; average oxidation state, 2.1–2.2) (Fig. 4C). Interestingly, these hotspots of oxidized Mn were colocalized with carbonyl groups (1,750-1,680 cm⁻¹) as shown in the µFTIR absorbance map (Fig. 4B).

Fig. 4.

Mn transformations at the hyphae–epidermis interface of two decomposing needles. (A) Photographs of needle thin sections showing fungal hyphae (FH) colonizing the outer walls of epidermis cells (ED), partially fungal-infected hypodermis cells (HD), and mesophyll tissue (MP). The white line delineates the boundary between fungal hyphae and the needle epidermis. (B) Corresponding µFTIR heat maps showing the distribution of aromatic, amide, and carbonyl functional groups. Heat maps were generated using the integrated absorbance of spectral regions given in the figure. (C) Corresponding Mn distribution maps of the same region of interest generated by µXRF. (D) Mn XANES spectra collected at locations along the hyphae–epidermis interface (points 1 and 2 in C) and the needle’s mesophyll tissue (points 4 and 5). Pie charts show relative amounts of Mn²⁺, Mn³⁺, and Mn⁴⁺ at each location, with numbers indicating the average oxidation state. (E) µFTIR spectra extracted from transect across the hyphae epidermis interface, shown as red line in B). Note that absorbance at ∼1,700 and ∼1,610 cm⁻¹ at the interface (B) increases relative to (C) infected epidermis regions and a fresh needle epidermis. (Scale bars: 30 µm.)

To further investigate the potential chemical alterations due to oxidative Mn species at the site of direct contact between fungal hyphae and the needle surface, we extracted µFTIR spectra along a transect across this interface (red line, Fig. 4B). Spectra taken from fungal tissue, the fungi–epidermis interface (contact zone), the needle epidermis, and mesophyll are presented in Fig. 4E. The spectrum extracted from the infected epidermis (C) showed a lower absorbance of saccharide bands (1,180–950 cm⁻¹), as well as enhanced absorbance of bands corresponding to carbonyl (∼1,700 cm⁻¹) and aromatic C = C groups (∼1,600 cm⁻¹) compared with uninfected epidermis regions (Fig. 4E).

Discussion

We initially hypothesized that litter-decomposing fungi repurpose Mn²⁺ naturally present in litter to produce reactive Mn³⁺ species at the site of oxidative needle degradation. Our results suggest a strong relationship between biotic Mn oxidation and litter decomposition both at macroscales (litter layer) and microscales (fungal needle colonization), the causality of which is discussed in the following.

Across the Douglas-fir litter layer examined here, Mn was progressively transformed. Total Mn increased rapidly in years 1–3, followed by a slower increase in years 4–6, a trend consistent with Mn accumulation in litter layers observed elsewhere (9, 22). Similarly, the amount of pyrophosphate-extractable Mn (Mn_PYRO) and the proportion of Mn³⁺ as evidenced by XANES increased most rapidly within the first 3 y whereas solid Mn⁴⁺ phases did not occur until years 4–6 (Table S1 and Fig. 1). Pyrophosphate is expected to complex and extract dissolved and/or organically complexed Mn (33), which we consider a proxy for bioavailable Mn, but does not differentiate between unreactive Mn²⁺ and oxidative Mn³⁺ species. On the other hand, XANES spectroscopy can detect Mn³⁺, without providing unambiguous information about its physical state. Combined, however, our extraction and spectroscopy results suggest the transformation of litter-borne Mn into bioavailable Mn²⁺ and reactive Mn³⁺ forms in the initial stages of decomposition. In later stages, increasing Mn oxidation resulted in Mn^3+/4+ forms similar to those observed in solid Mn oxide precipitates (24, 26, 27).

This rapid formation of bioavailable and reactive Mn forms in the first years of decomposition is accompanied by increased microbial processing of the litter. Accumulation of bioavailable and reactive Mn correlated with decreasing C/N ratios and stronger contributions of amide functionalities in the decomposing litter (Fig. 3 A and B). N content (34), fungal biomass (35), and visible fungal colonization of needle litter (36 ⇓–38) commonly rise within the first 2 y of decomposition. Within a similar time frame, maxima in MnP activity can be observed (35). Our results support the hypothesis that Mn mobilization and the formation of reactive Mn phases are caused by successive microbial colonization of the litter.

Further, the strong relationship between Mn cycling and chemical transformation of litter points to a direct involvement of Mn in the decomposition process. Mn oxidation is strongly related to the relative loss of saccharides and the transformation of aromatic litter components (Fig. 3). Oxidative transformations of aromatic compounds occurred across the litter layer, reflected in changes in abundance of aromatic compounds (Fig. 2C) and increases in oxygenated aromatic ring carbons (Fig. 2 A and B). These transformations were significantly correlated with Mn mobilization and oxidation (Fig. 3 E and F), suggesting that decomposing fungi actively cycle Mn for the purpose of using oxidized Mn species in the breakdown of aromatic structures.

Our imaging analysis can be reconciled as evidence that fungi actively promote Mn transport and oxidation during litter colonization. The high concentrations of Mn found in single hyphae and dense hyphal networks (Fig. S1B) imply biotic accumulation and transport of Mn in the litter layer. Greater contribution of Mn³⁺ and Mn⁴⁺ forms to overall Mn concentrations in hyphae further indicate active biotic Mn oxidation. In addition to Mn associated with fungal hyphae, we observed larger, Mn-rich particles consisting predominantly of Mn^3+/4+ forms on needle surfaces. A significant fraction of hyphae found on needle surfaces can be assumed dead (36 ⇓–38), and Blanchette (25) showed dead wood-colonizing hyphae covered in Mn precipitates. These observations suggest that, when the fungal supply of Mn³⁺-stabilizing chelators such as oxalic acid ceases upon cell death, excess Mn³⁺ disproportionates and precipitates as Mn^3+/4+ oxides on dead fungal residues. These coprecipitates may then accumulate as larger aggregates observed here and elsewhere (24).

Imaging further showed that reactive Mn³⁺ occurs in hotspots at the interface between hyphae and needle epidermis (Fig. 4C). These hotspots showed a mixture of Mn²⁺ and reactive Mn³⁺ species and almost no signs of Mn^3+/4+ oxide accumulation (Fig. 4A). If microbially produced Mn³⁺ engages in oxidation reactions with litter components at this interface, it is reduced back to Mn²⁺ in the process. In this scenario, no Mn^3+/4+ oxides accumulate, and fungi continuously reoxidize Mn²⁺ to form Mn³⁺. The fact that we find comparable amounts of Mn²⁺ and Mn³⁺ is therefore consistent with active cycling of the Mn^2+/3+ couple and its involvement in oxidation reactions.

Evidence for chemical alterations of saccharide and aromatic components at infected sites with high Mn³⁺ concentrations reveal a direct involvement of Mn in litter decomposition. First, decreasing absorbance of saccharides in spectra taken from the contact zone between hyphae and epidermis (Fig. 4E) is in good agreement with bulk saccharide loss across the litter layers (Table S1) and preferential (hemi)cellulose removal from the site of attack. Second, increased absorbance of bands arising from conjugated C=O and C=C bonds can be attributed to aromatic decomposition products released during the oxidative breakdown of plant material (39, 40). Removal of saccharides and concurrent oxidative alteration of aromatic lignin structures is a widely noted fungal decay pattern, where hyphae generate oxidized trenches in cell walls to liberate and release (hemi)cellulose components without invading the cell interior (41).

In summary, our study reveals the mechanistic link between microbial Mn cycling and the transformations of organic compounds during litter decomposition (Fig. 5A). Collectively, our bulk and microscale results show that litter-decomposing fungi first recruit and accumulate reduced Mn²⁺ in the litter layer, transform it into oxidative Mn³⁺ forms at the site of oxidative litter decomposition, and later accumulate it as Mn^3+/4+ oxide precipitates (Fig. 5B). Moreover, we find that Mn oxidation—specifically the formation of reactive Mn³⁺ species—in this ecosystem is not incidental, but tightly coupled to the oxidative degradation of aromatic structures in litter. These findings provide a mechanistic basis for the highly significant “Mn-dependence” (9 ⇓–11) of litter decomposition across a wide variety of forest ecosystems. Our results suggest that this relationship largely rests on the ability of decomposer organisms to recruit Mn in the litter layer and oxidize it at the site of litter decay. This insight demonstrates that Mn bioavailability and oxidation rate should be recognized as major determinants of litter decomposition in forest ecosystems. Furthermore, it provides strong support for a new ecological concept of litter decomposition: That the ability of decomposer organisms to degrade litter is, in large part, controlled by availability of, and access to, resources critical for the biochemical breakdown of litter.

Fig. 5.

Macro- and microscale coupling of C and Mn cycles in forest ecosystems. (A) Mn cycling through the entire plant–soil system and its link to litter decomposition and SOM formation. (B) Microscale Mn cycling by litter-decomposing fungi and its role in the oxidative breakdown of lignin or other aromatic litter components.

Our results support the hypothesis that plant–soil systems in forest biomes have coevolved to optimize the “cell-wall degrading machinery” (42), thereby maximizing litter decomposition (and thus the recycling of nutrients) by ensuring the availability of key resources such as Mn (Fig. 5A). Because aspects of global change also impact ecosystem Mn fluxes, bioavailability in soils (43, 44), plant uptake, and foliar litter concentrations (45), the tight coupling we demonstrate between Mn cycling and litter decomposition suggests that further research on regulators of ecosystem Mn fluxes is warranted.

SI Materials and Methods

Litter Decomposition Study.

This study took advantage of an ongoing litter decomposition experiment at the H. J. Andrews Experimental Forest, Oregon, United States. About 90% of the area’s annual precipitation falls from October to April, with the wettest period in December and peak drought conditions occurring in July. Within the HJA, the study was conducted at a site in watershed no. 8, at an elevation of 982 m and slope aspect of 223 degrees. Mean annual temperature at the nearby headquarters averaged 8.8 °C, and annual precipitation was 2,200 mm during 1974–2003 (46).

A rolling litter decomposition study spanning the years 2005–2011 was conducted at this site to examine the temporal and spatial variation in litter decomposition across individual layers. Nylon mesh panels (1-mm netting, 60 × 60-cm frames) were placed on the litter surface at this site annually. They were stacked over time to delineate annual litter fall from that of the subsequent year without restricting process intensities by artificial closure. Upon harvest at the end of the dry season in November 2011, the whole littercake was transferred on a supporting sheet and placed in a sealed plastic container. Recent needles and underlying O, A, and B horizon material were also collected and placed in amber vials and zip lock bags. Samples were transported at 4 °C and immediately returned to the laboratory. In the laboratory, individual layers of the littercake were separated, and decaying needles were manually separated from other forms of litter, such as cones and twigs. Soils were pushed through a 2-mm sieve and mixed thoroughly. Needle litter from each individual layer and the underlying soil horizons were then subsampled for laboratory analyses. Any variability reported included effects of subsampling and laboratory procedures. One subsample from each litter layer and mineral soil horizon was processed at field moisture for wet-chemical extractions and microscopic analyses. For spectroscopic analyses, another subsample was dried in an anaerobic glove box at room temperature in the dark to prevent changes in Mn oxidation state. A third subsample was oven-dried (50 °C, overnight) and ground for litter chemistry analyses.

Manganese Chemistry.

Total Mn (as well as Fe, Ca, and Al) content was quantified with X-ray fluorescence spectrometry (XEPOS HE XRF spectrometer; SPECTRO Analytical Instruments). To determine the amount of bioavailable Mn, we additionally quantified soluble and organically complexed Mn (Mn_pyro) using an Na-pyrophosphate extraction (49). Briefly, an aliquot of field-moist needle litter or soil equivalent to 200 mg of oven-dry sample was combined with 14 mL of 0.1 M Na-pyrophosphate solution at pH 10 in a 15-mL polypropylene centrifuge tube, agitated in the dark using a reciprocal shaker for 8 h, and centrifuged at 17,000 × g for 15 min. After centrifugation, the supernatant was removed, filtered through 0.22-μm syringe filters, and stabilized in 1% high-purity HNO₃. Concentrations of Mn (as well as Fe, Ca, and Al) in the extract were determined on a Perkin-Elmer SCIEX Elan DRC II inductively coupled plasma mass spectrometer (ICP-MS).

Mn oxidation state was determined using Mn XANES. Anaerobically dried litter and soil samples were hand-ground in the glove box, densely packed in poly(tetrafluoroethylene) sample holders, and sealed with X-ray transparent Kapton tape. Mn XANES spectra were recorded at the wiggler beamline 4-3 at the Stanford Synchrotron Radiation Lightsource (SSRL) with an Si(110) ϕ = 0 double-crystal monochromator, and fluorescence yield was collected with a Stern–Heard ion chamber detector (51). Background subtraction, normalization, and fitting of the spline function were performed on the collected spectra using the Athena software package (52). The average oxidation state of Mn was derived from individual spectra using a linear-combination fitting procedure (50). The procedure uses 17 pure valence Mn²⁺, Mn³⁺, and Mn⁴⁺ reference standards available (open-source, web link available in ref. 48), and fitting results are summarized in Table S3. In Manceau et al. (50), this method yielded the correct average valences to within about 0.04 valence units, and the correct fractions of the Mn²⁺, Mn³⁺, and Mn⁴⁺ states to within 4.4–4.6% when applied to the collection of mixed-valence Mn minerals. Based on the deviation between the sum of the three fractions from unity (Table S3), we estimate the error in our study to be <8% for bulk XANES and <11% for µXANES.

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Table S3.

Fractional and average valence states of Mn obtained from a linear combination fit of Mn XANES spectra as described in ref. 50

Litter Decomposition State.

Total C and N contents were determined using a Europa Scientific 20/20 isotope ratio mass spectrometer. Changes in the organic composition of the decomposing litter were determined using FTIR spectroscopy. FTIR spectra of the samples pressed in KBr pellets were recorded from 4,000 to 650 cm⁻¹ with a resolution of 4 cm⁻¹ on a Thermo Nicolet NEXUS 670 FTIR spectrometer (Thermo Fisher Scientific). For each sample, 512 scans were collected in transmission mode and averaged before Fourier transform processing. Three pellets were prepared and analyzed for each sample, and the resulting spectra were averaged. The OMNIC software package (Thermo Fisher Scientific) was used on the fingerprint region of the spectra (1,800–850 cm⁻¹) to perform baseline correction, normalization, and peak deconvolution on the spectra.

¹³C-NMR experiments were performed on a Bruker Avance 400 spectrometer at 100 MHz (400 MHz ¹H frequency). All experiments were performed with 4-mm sample rotors in a double-resonance probe head. Semiquantitative structural information on all carbons in the sample was obtained by ¹³C cross polarization/total sideband suppression (CP/TOSS) as described in ref. 28. The NMR experiments were conducted at a spinning speed of 5 kHz and a CP time of 1 ms, with a ¹H 90° pulse length of 4 µs and a recycle delay of 1 s. Four-pulse total suppression of sidebands was used before detection, and two-pulse phase-modulated decoupling was applied for optimum resolution. The corresponding subspectrum, with signals of nonprotonated carbons and carbons of mobile groups such as rotating CH₃, was obtained by ¹³C CP/TOSS combined with 40-ms dipolar dephasing (DD). The spectra and subspectra obtained with CP/TOSS and CP/TOSS with DD were assigned to different carbon functional groups following previous literature (53 ⇓⇓–56).

To identify molecular changes in aromatic components of the litter (lignins and tannins), synchrotron-based laser desorption post ionization (LDPI) mass spectrometry of fresh needles and litter layers was performed on a modified time-of-flight secondary ion mass spectrometer (TOF.SIMS V; IonTOF) coupled to a synchrotron UV light port at beamline 9.0.2 of the Advanced Light Source, Lawrence Berkeley National Laboratory (29, 31). Before analysis, 3 mg of ground needle material was suspended in high-purity (99.9%) MeOH, pipetted on silicon substrates (Wafer World), and dried in a clean hood under constant N₂ gas flow. The desorption laser was calibrated to emit 8.5-ns pulses focused to a spot diameter of ∼30 μm, and its power was reduced until no laser-induced fragmentation could be detected in the background (∼0.7 MW⋅cm⁻²) while achieving sufficient desorption efficiency. Under ultra-high vacuum, the sample was scanned with the laser with 2 mm⋅s⁻¹ over a 20-mm distance to avoid sample damage. To optimize the signal-to-noise ratio (S/N), mass spectra collected during each scan were the sum of ∼16,000 laser shots on the surface. We performed three replicate scans for each sample, and mass spectra are presented as averages normalized to the total intensity.

Zimmermann and coworkers (29, 30) showed that aromatic ring structures with m/z ratios greater than 100 have significantly lower IE (<8.5 eV) than other environmentally relevant compound classes (>9 for alcohols, alkanes, and alkenes). Using synchrotron-LDPI, we exploited this fact for selective detection of aromatic analytes within the mixture of organic compounds found in litter. To confirm that the most prominent mass peaks (e.g., m/z = 340, 370, 396, 399, 400, 406, and 414) in our mass spectra correspond to aromatic structures, we determined ionization energies (IEs) for each of the mass peaks in a second LDPI experiment as previously described (31, 32). Briefly, IEs were determined by varying the incident UV photon energy (i.e., the ionization energy) between 7.5 and 12.5 eV in 1-eV increments, and we then collected the resulting mass spectra. Normalized relative intensity of the mass peak of interest was then plotted against the photon energy used for each scan (Fig. S2). Using a linear fit, the intercept with the ordinate (photon energy) represents the IE of a given mass peak (listed in Table S2). Peaks at m/z ratios of 340, 368, 396, 399, 400, 406, and 412 showed IEs of ≤8.5 eV and were thus attributed to aromatic moieties.

Fig. S2.

Signal intensity of mass peaks (m/z ratios ranging from 340 to 414) plotted against incident photon energies (ranging from 8.5 to 13.5 eV with 1 eV increments). The ordinate intercept of the linear fit approximates the ionization energy (IE) for the respective mass fragment. Mass peaks with ionization energies ≤ 8.5 eV (dotted line) are identified as aromatic moieties. In this case, all fits intercept the ordinate at energies equal or below 8.5 eV.

Chemical Imaging Analyses.

Needles from all layers were visibly colonized by fungal hyphae, frequently concentrated around dark infections of the surface. To determine Mn distribution and oxidation state associated with these fungal infections, elemental maps and Mn XANES spectra of cross-sectioned needle litter were obtained using X-ray fluorescence mapping and X-ray absorption spectroscopy (µXRF/XAS). To this end, individual needles taken from the top layer were embedded in epoxy (Spurr; TedPella) and cured. Cross-sections were obtained by cutting the resin block and polishing the exposed surface using sand paper and diamond paste. µXRF maps and µXANES spectra were collected at the hard X-ray microprobe beamline 10.3.2 at the Advanced Light Source (ALS) at Lawrence Berkeley National Laboratory. µXRF maps were acquired by continuously raster scanning the sample with a 5 × 5-µm beam in 5-µm steps and a 100-ms count time. Maps were collected with the incident photon energy set at 7,000 and 10,000 eV, and the fluorescent peaks for K, Ca, Ti, Si, Cr, Mn, Fe, Ni, Cu, and Zn were obtained at each energy. The fluorescence yield was normalized by I₀ and the dwell time. Several regions of interest were selected from the µXRF maps for Mn K-edge µXANES analysis. Fluorescence yield was recorded in QuickXANES mode (30 single spectra) to acquire data with minimal beam damage per spot. QuickXANES spectra were checked for beam damage, and, if necessary, spectra showing evidence for beam reduction were excluded from further analysis. In all cases, a minimum of five QuickXANES spectra were averaged for each region of interest, and the resulting spectra were calibrated and normalized as described for bulk Mn XANES spectra.

To obtain high-resolution maps of the functional group chemistry at the hyphae–needle interface, thin sections of infected needles from the top layer were also prepared using a cryostat (Leica 1950 Cyrostat; Leica Instruments) without the use of carbon-based resins. Before sectioning, needles were surrounded by MilliQ H₂O and flash frozen in liquid N₂. The specimens were then removed from the tube and transferred on a chuck mount that was allowed to equilibrate to cutting temperature in the cryostat for 30 min. To attain 5-µm sections, the cryostat was set to −18 °C chamber temperature and −16 °C mount temperature. A 70-µm curl guard was adjusted as needed to achieve flat, nondeformed sections. Flat sections were transferred to gold-coated (IR reflective) microscope slides, and needle surfaces infected with fungi were identified using a light microscope. High-resolution infrared maps of these locations were acquired using synchrotron FTIR (SR-FTIR) spectromicroscopy at ALS beamline 1.4.3 (Lawrence Berkeley National Laboratory). Maps were collected on a Nicolet Nic-Plan IR microscope (Thermo Fisher Scientific) equipped with a liquid-N₂ cooled mercury-cadmium telluride (MCT) detector connected to the synchrotron light source. All maps were collected by point-by-point raster scanning of the thin sections using a 32× Reflachromat Nicolet objective with a 2-µm spot size. For each spot, 64 scans were collected from 4,000 to 650 cm⁻¹ at a resolution of 4 cm⁻¹. After completion of the SR-FTIR analysis, µXRF maps of the same regions were collected with a 3 × 3-µm beam in 2-µm steps and a count time of 100 ms. µXANES spectra at the Mn K-edge of regions of interest identified in the µXRF maps were acquired as outlined above.

Data Analysis.

All statistical analyses were performed using OriginPro (OriginLab). Reported SEs are based on three analytical replicates.

Materials and Methods

Litter Decomposition Study.

This study took advantage of an ongoing litter decomposition experiment at the H. J. Andrews Experimental Forest (HJA), Oregon, United States. About 90% of the area’s annual precipitation falls from October to April, with the wettest period in December and peak drought conditions occurring in July. Within the HJA, the study was conducted at a site in watershed no. 8, at an elevation of 982 m and slope aspect of 223 degrees. Mean annual temperature at the nearby headquarters averaged 8.8 °C, and annual precipitation was 2,200 mm during 1974–2003 (46). The soil underlying a dense cover of old-growth Douglas fir (Pseudotsuga menziesii) is an Andic Dystrudept and shows abundant patches of dense ectomycorrhizal mats (47), which can result in peroxidase activities 28- to 126-times greater than that of nonmat soils (48). We therefore expected particularly rapid Mn cycling at this site.

A “rolling” litter decomposition study spanning the years 2005–2011 was conducted at this site to examine the temporal and spatial variation in litter decomposition across individual layers. Nylon mesh panels (1-mm netting, 60 × 60-cm frames) were placed on the litter surface at this site annually. They were stacked over time to delineate each year’s litter fall from the next without confining processes by artificial closure. Upon harvest at the end of the dry season in November 2011, the whole littercake was transferred on a supporting sheet and placed in a sealed plastic container. Fresh needles and underlying O, A, and B horizon material were also collected and transported anaerobically in capped amber vials containing dry ice. Samples were transported at 4 °C and immediately returned to the laboratory. In the laboratory, individual 60 × 60-cm litter layers were separated. Decaying needles were manually isolated from other forms of litter such as cones and twigs, and mixed. Soils were pushed through a 2-mm sieve and mixed thoroughly. Needle litter and soil samples were then stored for further analyses as described in SI Materials and Methods.

Manganese Chemistry.

Total Mn (as well as Fe, Ca, and Al) content was quantified with X-ray fluorescence spectrometry (XEPOS HE XRF spectrometer; SPECTRO Analytical Instruments). To determine the amount of bioavailable Mn, we additionally conducted Na-pyrophosphate extractions (49), with extractable Mn (Mn_PYRO) taken to originate predominantly from soluble and organically complexed Mn (33). Extracted Mn concentrations were measured using inductively coupled plasma mass spectrometry (ICP-MS). Mn oxidation state was determined using Mn XANES (50). Litter and soil samples were dried and hand-ground in an anaerobic glove box and sealed with X-ray transparent Kapton tape. Mn XANES spectra were recorded at the wiggler beamline 4–3 at the Stanford Synchrotron Radiation Lightsource (SSRL) (51).

Litter Decomposition State.

Total C and N content was determined using a Europa Scientific 20/20 isotope ratio mass spectrometer. Changes in the organic composition of the decomposing litter were determined using FTIR spectroscopy. FTIR spectra of the samples pressed in KBr pellets were recorded from 4,000 to 650 cm⁻¹ with a resolution of 4 cm⁻¹ on a Thermo Nicolet NEXUS 670 FTIR spectrometer (Thermo Fisher Scientific).

Chemical Imaging Analyses.

To obtain high-resolution maps of the functional group chemistry at the hyphae–needle interface, thin sections (<5 µm) of infected needles from the top layer were also prepared using a cryostat (Leica 1950 Cyrostat; Leica Instruments) without the use of carbon-based resins and transferred to gold-coated (IR reflective) microscope slides. High-resolution infrared maps of these locations were acquired using synchrotron FTIR (µFTIR) spectromicroscopy at ALS beamline 1.4.3. After completion of the µFTIR analysis, µXRF maps of the same regions and Mn µXANES spectra of selected points within that region were collected at ALS beamline 10.3.2. Further details on sample preparation, analytical procedures, and data processing can be found in SI Materials and Methods.

Data Analysis.

All statistical analyses were performed using OriginPro (OriginLab Corp.). Reported SEs are based on three analytical replicates.

Acknowledgments

We thank J. Sexton for setting up the decomposition study and M. Sarginci for sample processing. We thank M. Marcus and H. Bechtel for help and support at Advanced Light Source beamlines 10.3.2 and 1.4.3, respectively, and E. Nelson for assistance at Stanford Synchrotron Radiation Lightsource beamline 4-3. M. Keiluweit acknowledges funding through a Lawrence Scholar Fellowship awarded by the Lawrence Livermore National Laboratory (LLNL). Funding for M.E.H. and the long-term litter decomposition experiment was provided by a National Science Foundation grant to the H. J. Andrews Long-Term Ecological Research Program (Grant DEB-0823380). Analytical work was performed under the auspices of the US Department of Energy (DOE) by LLNL under Contract DE-AC52-07NA27344. Funding was provided by LLNL Laboratory Directed Research and Development Award 10-ERD-021 “Microbes and Minerals: Imaging C Stabilization” (to J.P.-R., P.N., and M. Kleber), and the work of P.N. was supported by Lawrence Berkeley National Laboratory Award IC006762 as sub-award from LLNL and DOE-Biological and Environmental Research Sustainable Systems scientific focus area. M. Kleber acknowledges support through a research fellowship from the Institute of Soil Landscape Research at the Zentrum für Agrarlandschaftsforschung. Use of the Advanced Light Source is supported by the Director, Office of Science, Office of Basic Energy Sciences, US DOE under Contract DE-AC02-05CH11231. Use of SSRL at the Stanford Linear Accelerator Center National Accelerator Laboratory is supported by the US Department of Energy, Office of Science, Office of Basic Energy Sciences under Contract DE-AC02-76SF00515.

Footnotes

↵¹Present address: Stockbridge School of Agriculture, University of Massachusetts, Amherst, MA 01003.
↵²To whom correspondence should be addressed. Email: keiluweit{at}umass.edu.

Author contributions: M. Keiluweit, P.N., M.E.H., J.P.-R., and M. Kleber designed research; M. Keiluweit performed research; J.M. contributed new reagents/analytic tools; M. Keiluweit analyzed data; and M. Keiluweit, P.N., J.P.-R., and M. Kleber wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1508945112/-/DCSupplemental.

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Submit

[1] ↵
Hättenschwiler S,
Tiunov AV,
Scheu S
(2005) Biodiversity and litter decomposition in terrestrial ecosystems. Annu Rev Ecol Evol Syst 36(1):191–218
.
OpenUrl CrossRef

[2] Hättenschwiler S,

[3] Tiunov AV,

[4] Scheu S

[5] ↵
Prescott CE
(2010) Litter decomposition: What controls it and how can we alter it to sequester more carbon in forest soils? Biogeochemistry 101(1-3):133–149
.
OpenUrl CrossRef

[6] Prescott CE

[7] ↵
Chapin FS,
Matson PA,
Vitousek PM
(2011) Principles of Terrestrial Ecosystem Ecology (Springer, New York)
.

[8] Chapin FS,

[9] Matson PA,

[10] Vitousek PM

[11] ↵
Cotrufo MF,
Wallenstein MD,
Boot CM,
Denef K,
Paul E
(2013) The microbial efficiency-matrix stabilization (MEMS) framework integrates plant litter decomposition with soil organic matter stabilization: Do labile plant inputs form stable soil organic matter? Glob Change Biol 19(4):988–995
.
OpenUrl CrossRef

[12] Cotrufo MF,

[13] Wallenstein MD,

[14] Boot CM,

[15] Denef K,

[16] Paul E

[17] ↵
Klotzbücher T,
Kaiser K,
Guggenberger G,
Gatzek C,
Kalbitz K
(2011) A new conceptual model for the fate of lignin in decomposing plant litter. Ecology 92(5):1052–1062
.
OpenUrl CrossRef PubMed

[18] Klotzbücher T,

[19] Kaiser K,

[20] Guggenberger G,

[21] Gatzek C,

[22] Kalbitz K

[23] ↵
Meentemeyer V
(1978) Macroclimate and lignin control of litter decomposition rates. Ecology 59(3):465–472
.
OpenUrl CrossRef

[24] Meentemeyer V

[25] ↵
Turner S,
Gallois P,
Brown D
(2007) Tracheary element differentiation. Annu Rev Plant Biol 58(1):407–433
.
OpenUrl CrossRef PubMed

[26] Turner S,

[27] Gallois P,

[28] Brown D

[29] ↵
Berg B,
McClaugherty C
(2007) Plant Litter: Decomposition, Humus Formation, Carbon Sequestration (Springer, Berlin)
.

[30] Berg B,

[31] McClaugherty C

[32] ↵
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Long-term litter decomposition controlled by manganese redox cycling

Significance

Abstract

Results

Mn Transformations.

Litter Decomposition.

Mn Form and Distribution on Decomposing Needle Surfaces.

Manganese Chemistry at the Hyphae–Epidermis Interface.

Discussion

SI Materials and Methods

Litter Decomposition Study.

Manganese Chemistry.

Litter Decomposition State.

Chemical Imaging Analyses.

Data Analysis.

Materials and Methods

Litter Decomposition Study.

Manganese Chemistry.

Litter Decomposition State.

Chemical Imaging Analyses.

Data Analysis.

Acknowledgments

Footnotes

References

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Main menu

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Search

Significance

Abstract

Results

Mn Transformations.

Litter Decomposition.

Mn Form and Distribution on Decomposing Needle Surfaces.

Manganese Chemistry at the Hyphae–Epidermis Interface.

Discussion

SI Materials and Methods

Litter Decomposition Study.

Manganese Chemistry.

Litter Decomposition State.

Chemical Imaging Analyses.

Data Analysis.

Materials and Methods

Litter Decomposition Study.

Manganese Chemistry.

Litter Decomposition State.

Chemical Imaging Analyses.

Data Analysis.

Acknowledgments

Footnotes

References

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