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ScienceDirect Exploring bacterial lignin degradation Margaret E Brown1 and Michelle CY Chang1,2 Plant biomass represents a renewable carbon feedstock that could potentially be used to replace a significant level of petroleum-derived chemicals. One major challenge in its utilization is that the majority of this carbon is trapped in the recalcitrant structural polymers of the plant cell wall. Deconstruction of lignin is a key step in the processing of biomass to useful monomers but remains challenging. Microbial systems can provide molecular information on lignin depolymerization as they have evolved to break lignin down using metalloenzyme-dependent radical pathways. Both fungi and bacteria have been observed to metabolize lignin; however, their differential reactivity with this substrate indicates that they may utilize different chemical strategies for its breakdown. This review will discuss recent advances in studying bacterial lignin degradation as an approach to exploring greater diversity in the environment. Addresses 1 Department of Chemistry, University of California, Berkeley, CA 947201460, USA 2 Department of Molecular and Cell Biology, University of California, Berkeley, CA 94720-1460, USA Corresponding author: Chang, Michelle CY ([email protected])
Current Opinion in Chemical Biology 2014, 19:1–7 This review comes from a themed issue on Bioinorganic chemistry
Lignin is a highly complex aromatic heteropolymer whose biological role in plants is to increase cell wall integrity and resistance to attack by pathogens. It is made from the enzyme-mediated radical polymerization of three different phenolic monomers that differ from each other by the number of methoxy substituents present on the aromatic ring (n = 0–2, Figure 1) [3,6,7]. Upon radical initiation, the oxidizing equivalent can delocalize around the monomer until a crosslink is formed with a neighboring radical on the growing polymer, leading to the various C–C and C–O bonding motifs that have been characterized in lignin structures (Figure 1) [6]. As the phenolic and b-hydrogen are shared between all three monomers, these sites participate in a large number of the linkages present in lignin. Indeed, the b-aryl ether (or b-O-4) linkage has been estimated to represent up to 80% of the bonding motifs found in lignin [6,8]. The pattern of methoxy group substitution on each monomer influences lignin structure as they protect these positions from radical coupling reactions. As such, the monomer composition in individual plants controls lignin structure and the hydroxyphenyl (H):guaiacyl (G):syringyl (S) ratios can be genetically manipulated in various ways to improve lignin properties for downstream pulping processes or animal forage [4,6,8,9]
Edited by Elizabeth M Nolan and Mitsuhiko Shionoya For a complete overview see the Issue and the Editorial Available online 25th December 2013 1367-5931/$ – see front matter, # 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.cbpa.2013.11.015
Introduction Plant biomass offers one of the few renewable carbon feedstocks that is produced on a large enough scale to displace a significant volume of petroleum-derived chemicals. In comparison to traditional fossilized sources, plant biomass presents a major challenge for downstream processing as the majority of the carbon is derived from the plant cell wall in the form of recalcitrant structural biopolymers rather than residing in the more easily accessible sugar, starch, and oil fractions [1,2]. In particular, there is an intense interest in understanding the degradation of cellulose and hemicellulose as they directly yield fermentable sugars upon hydrolysis. However, a molecular understanding of lignin deconstruction is also essential since it can comprise up to 30% of plant biomass and also must be removed before the cellulose and hemicellulose can be accessed [3,4,5]. www.sciencedirect.com
Despite the naturally evolved recalcitrance of lignin, select microbes have discovered enzymatic approaches to its depolymerization. The most active microbes with respect to lignin degradation identified to date are fungi, such as those belonging to the white-rot or brown-rot families that decompose wood [10–12]. Lignin peroxidase (LiP) was the first lignolytic enzyme to be isolated from Phanerochaete chrysosporium and was found to contain a heme cofactor that is competent to oxidize unusually high potential sites, such as aromatic rings [13–15]. In addition to LiP, fungi also utilize other secreted metalloenzymes to break down lignin, including the heme-containing manganese peroxidases (MnP) [16,17] and versatile peroxidases (VP) [18] as well as multicopper-dependent laccases [12]. Taken together, the current model for microbial lignin degradation invokes the oxidative combustion of lignin mediated by a broad range of small molecule oxidants produced by these metalloenzymes, such as the vetratryl alcohol cation radical [15] and various Mn(III) coordination complexes [16,17] or those produced in secondary radical cascades [11,19] (Figure 2a). These diffusible mediators, rather than the enzymes themselves, are thought to react directly with lignin to generate radical sites within the substrate and initiate a cascade of bond scission reactions that ultimately leads to Current Opinion in Chemical Biology 2014, 19:1–7
2 Bioinorganic chemistry
Figure 1
lignin monomers HO
OH
OH
OH
HO
5
β O 4
OMe
OMe
O OMe
MeO
OMe
OMe
OH
OH
OH
p-coumaryl alcohol
coniferyl alcohol
sinapyl alcohol
γ OH α
6 5 R
1
4 O•
OMe
MeO O
p-hydroxyphenyl
guaiacyl
syringyl
O
O
phenylcoumaran
O
β-5 (α-O-4)
OMe
R
O
resinol
O
5 MeO
OMe
O
OMe
β
β-O-4
R = H, OMe O
β
β-aryl ether
2 3
HO
OMe
4 O α
O
O
β
β
OMe
β-β (γ-O-α)
5
MeO
5 O4 O
monomer oxidation
OMe
biphenyl ether
biphenyl
5-O-4
5-5
common lignin bonding motifs
lignin units
Current Opinion in Chemical Biology
Lignin structure. Lignin is constructed from three major hydroxycinnamyl alcohol monomer units which are crosslinked to form the corresponding units in lignin (left). Lignification involves oxidation of the monomer to the phenoxyl radical, followed by various radical-radical coupling reactions with the growing polymer, resulting in a broad range of lignin crosslinks (right). These structures are among the major structural units characterized to date (adapted from [6]).
Figure 2
(a)
(c)
OH
HO
• OOH OMe
Mn(III)-L
O
O2, H2O
O
HO OMe
OMe •
OMe
OMe
OMe
OMe OMe
OMe
R
• O2−
• OH
•
−
OH OMe O
O•
(b)
+
OMe
•
• OH
• CO2
HO
O +
OMe
VA•+
HO
HO
O
OH
OMe
O
H2O H2O2 O
VA•+
VA
O
FeIV
LiP
Fe
LiP
VA•+ lignin
generation of oxidant
diffusion
lignin oxidation
bond scission
Current Opinion in Chemical Biology
Microbial degradation of lignin. (a) Various small molecule oxidants generated directly from LiP, MnP, and laccases, as well as secondary oxidants produced by radical cascades. Oxidizing species are represented in red. (b) The current model for lignin degradation involves enzymatic generation of the radical mediator, which can then diffuse to the lignin substrate and transfer the oxidizing equivalent to the polymer. Upon formation of a ligninbased radical, bond scission reactions will ensue that lead to depolymerization. (c) Proposed reactions of model dimers leading to formation of observed products in fungal ligninases (adapted from [11]). Current Opinion in Chemical Biology 2014, 19:1–7
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Exploring bacterial lignin degradation Brown and Chang 3
its decomposition to smaller aromatic compounds, CO2, and water (Figure 2b,c). Given the complexity of the lignin substrate, the identity and properties of the full enzyme and mediator system involved in lignin metabolism remains to be fully elucidated. In this regard, exploration of new biological diversity may allow us to discover alternative molecular approaches involved in lignin metabolism in the environment. For example, microbes often demonstrate species selectivity with regard to the metabolism of various plants, which may possibly be related to differences in lignin structure and bonding. Compared to fungi, bacteria are much less well characterized with respect to their lignin metabolism but are known to exhibit different reactivity based on differences in the resulting product profile. In this context, we will discuss recent advances in studying lignin degradation in bacteria and efforts to expand the scope of our understanding of environmental microbial processes. Bacterial lignin metabolism
While much focus has been placed on studying lignin degradation by fungi because of their high activity, various genera of bacteria have also been observed to metabolize lignin and shown to be competent to release 14 C-CO2 from labeled lignin [20–25]. While ensuing in vitro [26] and in vivo [27,28] studies have shown that bacteria also have the capacity to catabolize non-phenolic compounds, the relationship of these activities with respect to lignin degradation remains unclear. Many soil bacteria, especially actinomycetes, have been reported to react with lignin to both solubilize it and produce a high molecular weight metabolite termed acidprecipitable polymeric lignin (APPL) [22,29]. Although the metabolism of lignin is not as complete compared to fungal systems, it is clear that bacteria can react with lignin and possibly produce smaller aromatics that can be imported into the cell for aromatic catabolism, which is also widespread in soil bacteria [30–33]. The first molecular information on bacterial APPL formation was reported with the isolation of a secreted bacterial heme peroxidase from a gram-positive bacterium, Streptomyces viridosporus T7A [26,34], which indicated that bacteria also likely possess a set of extracellular oxidative enzymes involved in lignin metabolism. Initial studies showed that the T7A peroxidase was also biochemically competent for the degradation of non-phenolics [26] but was not as oxidizing as fungal peroxidases [35]. Since advances in DNA sequencing have revolutionalized de novo bacterial genome sequencing, many of these environmental hosts have become amenable toward genome-level studies. As a result, the oxidative systems of lignin-reactive bacteria can be identified for further characterization. Upon screening of several different soil www.sciencedirect.com
bacteria and ranking both APPL formation and extracellular peroxidase activity, Amycolatopsis sp. 75iv2 (formerly Streptomyces setonii and Streptomyces griseus 75vi2) was identified as highly active with respect to both assays [36], which was consistent with previous literature reports on its physiology [37]. Using the Illumina platform, a rapid de novo genome for A. sp. 75iv2 was assembled and revealed the presence of a large number of genes encoding oxidative enzymes, such as heme peroxidases, laccases, and cytochrome P450s, as well as several predicted genes for production of extracellular peroxide [36,38]. This genome assembly also enabled proteomic analysis of the secreted protein fraction where biomass-degrading enzymes reside. Further functional studies showed that two heme-containing proteins were consistently found to be abundant in the secretome of A. sp. 75iv2 and were subsequently named Amyco1 and Amyco2 [36]. On the basis of sequence homology, the functions of Amyco1 and Amyco2 could be assigned as a bifunctional catalase-peroxidase and catalase, respectively, which was confirmed by in vitro biochemical experiments. Further studies showed that Amyco1 was competent to form the typical two-electron (compound I) and one-electron (compound II) oxidized states found in peroxidases and that it could degrade phenolic lignin model compounds [36]. More recently, A. sp. 75iv2 has been found to be able to grow on lignin as a sole carbon source and studies to further characterize the molecular basis for this phenotype are underway. Bacterial heme peroxidases
At the same time, new phenotypic screens have been designed to help identify novel lignolytic bacterial species. Using a set of fluorescent and colorimetric substrate probes, several previously uncharacterized bacteria were also found to demonstrate the ability to react with lignin [39]. Studies of knockout strains of one of these genera, Rhodoccocus jostii RHA1, implicated dye-decolorizing peroxidases (DyP) in this behavior [40]. DyPs belong to a small but distinct family of heme peroxidases, which contain a distal aspartate residue in place of a histidine [41]. In addition, they have often been found to be responsible for the extracellular oxidation and decomposition of dyes and other aromatic xenobiotics in fungi [42–44]. They separate into four major clades with the smaller and less active bacterial enzymes belonging to the A-clade and B-clade and the larger and more active fungal enzymes residing to the D-clade. C-type DyPs are prokaryotic in origin but cluster more closely with the fungal D-clade and appear to share their high activity with respect to oxidation of typical DyP substrates [45]. Of the two DyPs found in the R. jostii RHA1 genome, the B-type DypB was found to be responsible for the observed reactivity with lignin and was shown to be biochemically competent for the degradation of Current Opinion in Chemical Biology 2014, 19:1–7
4 Bioinorganic chemistry
non-phenolic lignin compounds [40]. While DypB did not contain a canonical secretion sequence, biochemical experiments suggest that it could be exported from the cell through a different mechanism [46]. It was also observed to have low MnP activity (kcat/KM = 16 M 1 s 1) 1) that could amplify its ability to degrade lignin-based compounds. Additional structural and biochemical studies have also been carried out on DypB [47,48] and have aided the engineering of the Mn binding pocket for a 14-fold improvement in the MnP activity [49]. While not as widely distributed in bacterial genomes as other heme peroxidases or cytochromes, genes encoding DyPs are found to be prevalent in actinomycetes. Interestingly, A. sp. 75iv2 contains three DyPs (DyP1–3), one of which is a typical bacterial A-type DyP (DyP3) and two of which belong to the C-clade (DyP1-2). The presence of two C-type DyPs was unexpected as they are observed less frequently but are often clustered with other genes encoding biomass-degrading enzymes in actinomycetes [50]. In particular, DyP2 belonged to a particularly divergent and sparsely populated branch within the C-clade and was thus selected for additional biochemical and structural characterization. In vitro assays with various substrates showed that DyP2 had high peroxidase and MnP activity (kcat/KM = 105–106 M 1 s 1) near the same magnitude as fungal enzymes, including those involved in lignin degradation [50]. In addition, DyP2 was competent to oxidize higher potential substrates such as Reactive Black 5. Another interesting observation was that the presence of Mn(II) enabled a secondary oxygenase activity that expanded the substrate scope of DyP2 to include the degradation of non-phenolic lignin model compounds [50]. The high MnP and secondary Mn oxygenase activity appears to be related to the relatively high affinity of DyP2 for Mn(II) that was supported by the discovery of a metal-binding site in the crystal structure that was localized 15 A˚ from the heme site.
Given the breadth of bacterial enzymes and pathways that functionalize or degrade aromatics, this discovery may indicate that many bacteria may interact differently with lignin as a substrate for use as an aromatic carbon source rather than carry out its rapid degradation to access the sugar-based cellulose and hemicellulose substrates as is believed to be the case in many fungal systems. Discovery of new lignin-reactive bacteria and study of microbial consortia
Certainly the exploration of greater microbial biodiversity in new environments will help to extend our understanding of the scope of lignin metabolism in microbes. Scaledup screening studies have identified many new bacterial species that appear to be much more active than previously characterized strains [52]. Indeed, the study of these new microbes may reveal the presence of new mechanisms for lignin reactivity especially when they originate from unusual environments where active biomass degradation occurs. For instance, greater availability of microbial metagenomes from sources where biomass is rapidly degraded — such as wood-feeding insects [53–55], the rumen of cows [56], or active soils [57] — will allow us to assess greater diversity at the genetic level. In particular, environments where lignin degradation occurs under anaerobic or microaerobic conditions [58,59] may be especially interesting as canonical lignin degradation involves peroxide- or oxygen-dependent enzymes. Indeed, insufficient knowledge of the suite of genes involved in lignin modification creates roadblocks in functional annotation of metagenomes by sequence homology alone [53]. However, continuing physiological [60] and structural studies [61] could help to identify new chemical transformations on lignin as a result of microbial metabolism. In addition, isolation or comparative genomics studies of microbes from environments such as rainforest soil [62] or forest areas [63,64,65] also serve to improve our understanding of the function of various members of microbial consortia.
Other bacterial enzymes involved in lignin processing
In general, the heme peroxidases from these lignin-reactive bacteria have been found to be less oxidatively powerful compared to the fungal enzymes involved in lignin degradation. However, it is possible that heme peroxidases do not constitute the major lignolytic enzymes in bacterial systems. Beyond the canonical enzymes used by fungi, bacteria also contain a suite of oxidative enzymes that could modify lignin for breakdown by hydroxylation or demethylation, such as cytochrome P450s, non-heme iron enzymes, or Mn- and Cu-containing oxidases. In one interesting example, a gene encoding a non-heme iron dioxygenase fused to a carbohydrate binding domain (CBD) was found from a cellulolytic streptoycetes isolated from wood-boring insects [51]. Biochemical studies of SACTE_2871 showed that was competent to bind lignin polymers as well as catalyze O2-dependent ring-opening of catechols. Current Opinion in Chemical Biology 2014, 19:1–7
Conclusions Lignin is a complex substrate that is known to require a suite of oxidative enzymes and assorted small molecule co-factors for its degradation. By exploring greater diversity of the organisms that can react with and metabolize lignin, we may be able to gain new insight into strategies for biomass deconstruction. Bacterial systems have been found to be less oxidatively powerful compared to lignolytic fungal systems to date but may provide a rich source for elucidating new accessory enzymes that could act synergistically with the major oxidative enzymes to activate and uncap various sites, similar to what has been involved in cellulose degradation. For example, hydroxylation or demethylation of various sites could serve to generate new chemical handles for further degradation. In addition, new studies showing the existence of oxidative enzymes that are capable of associating with lignin www.sciencedirect.com
Exploring bacterial lignin degradation Brown and Chang 5
could indicate that this process is not completely mediated through secondary small molecule mediators. In this regard, the existence of direct interactions between the substrate and lignin processing enzymes may also begin to explain differential reactivity profiles that are observed. Taken together, work in this area will continue to help bring more molecular detail into understanding how lignin degradation occurs in the environment.
Acknowledgements Research in our laboratory in this area is funded by the generous support of the University of California, Berkeley and the Energy Biosciences Institute.
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51. Bianchetti CM, Harmann CH, Takasuka TE, Hura GL, Dyer K, Fox BG: Fusion of dioxygenase and lignin-binding domains in a novel secreted enzyme from cellulolytic Streptomyces sp. SirexAA-E. J Biol Chem 2013, 288:18574-18587. An unusual fusion between a ring dioxygenase and lignin binding domain was found in a streptomycete symbiont isolated from a wood-boring insect. This report is the first evidence that an enzyme involved in aromatic degradation can possibly associate directly with biomass. 52. Taylor CR, Hardiman EM, Ahmad M, Sainsbury PD, Norris PR, Bugg TDH: Isolation of bacterial strains able to metabolize lignin from screening of environmental samples. J Appl Microbiol 2012, 113:521-530. Scaled-up screening of bacterial strains isolated from woodland soils and enriched by growth on lignocellulose led to the identification of many isolates with high lignin degrading activity. 53. Warnecke F, Luginbuhl P, Ivanova N, Ghassemian M, Richardson TH, Stege JT, Cayouette M, McHardy AC, Djordjevic G, Aboushadi N et al.: Metagenomic and functional analysis of hindgut microbiota of a wood-feeding higher termite. Nature 2007, 450:560-565. 54. Shi W, Syrenne R, Sun J-Z, Yuan JS: Molecular approaches to study the insect gut symbiotic microbiota at the ‘omics’ age. Insect Sci 2010, 17:199-219. 55. Boucias DG, Cai Y, Sun Y, Lietze V-U, Sen R, Raychoudhury R, Scharf ME: The hindgut lumen prokaryotic microbiota of the termite Reticulitermes flavipes and its responses to dietary lignocellulose composition. Mol Ecol 2013, 22:1836-1853. 56. Hess M, Sczyrba A, Egan R, Kim T-W, Chokhawala H, Schroth G, Luo S, Clark DS, Chen F, Zhang T et al.: Metagenomic discovery of biomass-degrading genes and genomes from cow rumen. Science 2011, 331:463-467. The authors sequence and analyze the metagenome of the cow rumen and focus on cataloguing genes involved in biomass degradation. 57. Daniel R: The metagenomics of soil. Nat Rev Microbiol 2005, 3:470-478. 58. Kato K, Kozaki S, Sakuranaga M: Degradation of lignin compounds by bacteria from termite guts. Biotechnol Lett 1998, 20:459-462. 59. Kajikawa H, Kudo H, Kondo T, Jodai K, Honda Y, Kuwahara M, Watanabe T: Degradation of benzyl ether bonds of lignin by ruminal microbes. FEMS Microbiol Lett 2000, 187:15-20. 60. Geib SM, Filley TR, Hatcher PG, Hoover K, Carlson JE, JimenezGasco MdM, Nakagawa-Izumi A, Sleighter RL, Tien M: Lignin degradation in wood-feeding insects. Proc Natl Acad Sci U S A 2008, 105:12932-12937. 61. Mansfield SD, Kim H, Lu F, Ralph J: Whole plant cell wall characterization using solution-state 2D NMR. Nat Protoc 2012, 7:1579-1589. This protocol describes methods for 2D-NMR characterization of native lignins. 62. Huang X-F, Santhanam N, Badri DV, Hunter WJ, Manter DK, Decker SR, Vivanco JM, Reardon KF: Isolation and characterization of lignin-degrading bacteria from rainforest soils. Biotechnol Bioeng 2013, 110:1616-1626. 63. Martinez D, Challacombe J, Morgenstern I, Hibbett D, Schmoll M, Kubicek CP, Ferreira P, Ruiz-Duenas FJ, Martinez AT, Kersten P et al.: Genome, transcriptome, and secretome analysis of wood decay fungus Postia placenta supports unique www.sciencedirect.com
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mechanisms of lignocellulose conversion. Proc Natl Acad Sci U S A 2009, 106:1954-1959. 64. Eastwood DC, Floudas D, Binder M, Majcherczyk A, Schneider P, Aerts A, Asiegbu FO, Baker SE, Barry K, Bendiksby M et al.: The plant cell wall-decomposing machinery underlies the functional diversity of forest fungi. Science 2011, 333:762-765. A large-scale comparative genomics study indicates that fungi have evolved different machinery for lignin breakdown based on their ecological niche.
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65. Fernandez-Fueyo E, Ruiz-Duen˜as FJ, Ferreira P, Floudas D, Hibbett DS, Canessa P, Larrondo LF, James TY, Seelenfreund D, Lobos S et al.: Comparative genomics of Ceriporiopsis subvermispora and Phanerochaete chrysosporium provide insight into selective ligninolysis. Proc Natl Acad Sci U S A 2012, 109:5458-5463. A comparative genomics study of two white rot fungi reveals differences in a strain that simultaneously degrades lignin and cellulose (P. chrysosporium) compared to one that solely depolymerizes lignin (C. subvermispora).
Current Opinion in Chemical Biology 2014, 19:1–7
ScienceDirect Exploring bacterial lignin degradation Margaret E Brown1 and Michelle CY Chang1,2 Plant biomass represents a renewable carbon feedstock that could potentially be used to replace a significant level of petroleum-derived chemicals. One major challenge in its utilization is that the majority of this carbon is trapped in the recalcitrant structural polymers of the plant cell wall. Deconstruction of lignin is a key step in the processing of biomass to useful monomers but remains challenging. Microbial systems can provide molecular information on lignin depolymerization as they have evolved to break lignin down using metalloenzyme-dependent radical pathways. Both fungi and bacteria have been observed to metabolize lignin; however, their differential reactivity with this substrate indicates that they may utilize different chemical strategies for its breakdown. This review will discuss recent advances in studying bacterial lignin degradation as an approach to exploring greater diversity in the environment. Addresses 1 Department of Chemistry, University of California, Berkeley, CA 947201460, USA 2 Department of Molecular and Cell Biology, University of California, Berkeley, CA 94720-1460, USA Corresponding author: Chang, Michelle CY ([email protected])
Current Opinion in Chemical Biology 2014, 19:1–7 This review comes from a themed issue on Bioinorganic chemistry
Lignin is a highly complex aromatic heteropolymer whose biological role in plants is to increase cell wall integrity and resistance to attack by pathogens. It is made from the enzyme-mediated radical polymerization of three different phenolic monomers that differ from each other by the number of methoxy substituents present on the aromatic ring (n = 0–2, Figure 1) [3,6,7]. Upon radical initiation, the oxidizing equivalent can delocalize around the monomer until a crosslink is formed with a neighboring radical on the growing polymer, leading to the various C–C and C–O bonding motifs that have been characterized in lignin structures (Figure 1) [6]. As the phenolic and b-hydrogen are shared between all three monomers, these sites participate in a large number of the linkages present in lignin. Indeed, the b-aryl ether (or b-O-4) linkage has been estimated to represent up to 80% of the bonding motifs found in lignin [6,8]. The pattern of methoxy group substitution on each monomer influences lignin structure as they protect these positions from radical coupling reactions. As such, the monomer composition in individual plants controls lignin structure and the hydroxyphenyl (H):guaiacyl (G):syringyl (S) ratios can be genetically manipulated in various ways to improve lignin properties for downstream pulping processes or animal forage [4,6,8,9]
Edited by Elizabeth M Nolan and Mitsuhiko Shionoya For a complete overview see the Issue and the Editorial Available online 25th December 2013 1367-5931/$ – see front matter, # 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.cbpa.2013.11.015
Introduction Plant biomass offers one of the few renewable carbon feedstocks that is produced on a large enough scale to displace a significant volume of petroleum-derived chemicals. In comparison to traditional fossilized sources, plant biomass presents a major challenge for downstream processing as the majority of the carbon is derived from the plant cell wall in the form of recalcitrant structural biopolymers rather than residing in the more easily accessible sugar, starch, and oil fractions [1,2]. In particular, there is an intense interest in understanding the degradation of cellulose and hemicellulose as they directly yield fermentable sugars upon hydrolysis. However, a molecular understanding of lignin deconstruction is also essential since it can comprise up to 30% of plant biomass and also must be removed before the cellulose and hemicellulose can be accessed [3,4,5]. www.sciencedirect.com
Despite the naturally evolved recalcitrance of lignin, select microbes have discovered enzymatic approaches to its depolymerization. The most active microbes with respect to lignin degradation identified to date are fungi, such as those belonging to the white-rot or brown-rot families that decompose wood [10–12]. Lignin peroxidase (LiP) was the first lignolytic enzyme to be isolated from Phanerochaete chrysosporium and was found to contain a heme cofactor that is competent to oxidize unusually high potential sites, such as aromatic rings [13–15]. In addition to LiP, fungi also utilize other secreted metalloenzymes to break down lignin, including the heme-containing manganese peroxidases (MnP) [16,17] and versatile peroxidases (VP) [18] as well as multicopper-dependent laccases [12]. Taken together, the current model for microbial lignin degradation invokes the oxidative combustion of lignin mediated by a broad range of small molecule oxidants produced by these metalloenzymes, such as the vetratryl alcohol cation radical [15] and various Mn(III) coordination complexes [16,17] or those produced in secondary radical cascades [11,19] (Figure 2a). These diffusible mediators, rather than the enzymes themselves, are thought to react directly with lignin to generate radical sites within the substrate and initiate a cascade of bond scission reactions that ultimately leads to Current Opinion in Chemical Biology 2014, 19:1–7
2 Bioinorganic chemistry
Figure 1
lignin monomers HO
OH
OH
OH
HO
5
β O 4
OMe
OMe
O OMe
MeO
OMe
OMe
OH
OH
OH
p-coumaryl alcohol
coniferyl alcohol
sinapyl alcohol
γ OH α
6 5 R
1
4 O•
OMe
MeO O
p-hydroxyphenyl
guaiacyl
syringyl
O
O
phenylcoumaran
O
β-5 (α-O-4)
OMe
R
O
resinol
O
5 MeO
OMe
O
OMe
β
β-O-4
R = H, OMe O
β
β-aryl ether
2 3
HO
OMe
4 O α
O
O
β
β
OMe
β-β (γ-O-α)
5
MeO
5 O4 O
monomer oxidation
OMe
biphenyl ether
biphenyl
5-O-4
5-5
common lignin bonding motifs
lignin units
Current Opinion in Chemical Biology
Lignin structure. Lignin is constructed from three major hydroxycinnamyl alcohol monomer units which are crosslinked to form the corresponding units in lignin (left). Lignification involves oxidation of the monomer to the phenoxyl radical, followed by various radical-radical coupling reactions with the growing polymer, resulting in a broad range of lignin crosslinks (right). These structures are among the major structural units characterized to date (adapted from [6]).
Figure 2
(a)
(c)
OH
HO
• OOH OMe
Mn(III)-L
O
O2, H2O
O
HO OMe
OMe •
OMe
OMe
OMe
OMe OMe
OMe
R
• O2−
• OH
•
−
OH OMe O
O•
(b)
+
OMe
•
• OH
• CO2
HO
O +
OMe
VA•+
HO
HO
O
OH
OMe
O
H2O H2O2 O
VA•+
VA
O
FeIV
LiP
Fe
LiP
VA•+ lignin
generation of oxidant
diffusion
lignin oxidation
bond scission
Current Opinion in Chemical Biology
Microbial degradation of lignin. (a) Various small molecule oxidants generated directly from LiP, MnP, and laccases, as well as secondary oxidants produced by radical cascades. Oxidizing species are represented in red. (b) The current model for lignin degradation involves enzymatic generation of the radical mediator, which can then diffuse to the lignin substrate and transfer the oxidizing equivalent to the polymer. Upon formation of a ligninbased radical, bond scission reactions will ensue that lead to depolymerization. (c) Proposed reactions of model dimers leading to formation of observed products in fungal ligninases (adapted from [11]). Current Opinion in Chemical Biology 2014, 19:1–7
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Exploring bacterial lignin degradation Brown and Chang 3
its decomposition to smaller aromatic compounds, CO2, and water (Figure 2b,c). Given the complexity of the lignin substrate, the identity and properties of the full enzyme and mediator system involved in lignin metabolism remains to be fully elucidated. In this regard, exploration of new biological diversity may allow us to discover alternative molecular approaches involved in lignin metabolism in the environment. For example, microbes often demonstrate species selectivity with regard to the metabolism of various plants, which may possibly be related to differences in lignin structure and bonding. Compared to fungi, bacteria are much less well characterized with respect to their lignin metabolism but are known to exhibit different reactivity based on differences in the resulting product profile. In this context, we will discuss recent advances in studying lignin degradation in bacteria and efforts to expand the scope of our understanding of environmental microbial processes. Bacterial lignin metabolism
While much focus has been placed on studying lignin degradation by fungi because of their high activity, various genera of bacteria have also been observed to metabolize lignin and shown to be competent to release 14 C-CO2 from labeled lignin [20–25]. While ensuing in vitro [26] and in vivo [27,28] studies have shown that bacteria also have the capacity to catabolize non-phenolic compounds, the relationship of these activities with respect to lignin degradation remains unclear. Many soil bacteria, especially actinomycetes, have been reported to react with lignin to both solubilize it and produce a high molecular weight metabolite termed acidprecipitable polymeric lignin (APPL) [22,29]. Although the metabolism of lignin is not as complete compared to fungal systems, it is clear that bacteria can react with lignin and possibly produce smaller aromatics that can be imported into the cell for aromatic catabolism, which is also widespread in soil bacteria [30–33]. The first molecular information on bacterial APPL formation was reported with the isolation of a secreted bacterial heme peroxidase from a gram-positive bacterium, Streptomyces viridosporus T7A [26,34], which indicated that bacteria also likely possess a set of extracellular oxidative enzymes involved in lignin metabolism. Initial studies showed that the T7A peroxidase was also biochemically competent for the degradation of non-phenolics [26] but was not as oxidizing as fungal peroxidases [35]. Since advances in DNA sequencing have revolutionalized de novo bacterial genome sequencing, many of these environmental hosts have become amenable toward genome-level studies. As a result, the oxidative systems of lignin-reactive bacteria can be identified for further characterization. Upon screening of several different soil www.sciencedirect.com
bacteria and ranking both APPL formation and extracellular peroxidase activity, Amycolatopsis sp. 75iv2 (formerly Streptomyces setonii and Streptomyces griseus 75vi2) was identified as highly active with respect to both assays [36], which was consistent with previous literature reports on its physiology [37]. Using the Illumina platform, a rapid de novo genome for A. sp. 75iv2 was assembled and revealed the presence of a large number of genes encoding oxidative enzymes, such as heme peroxidases, laccases, and cytochrome P450s, as well as several predicted genes for production of extracellular peroxide [36,38]. This genome assembly also enabled proteomic analysis of the secreted protein fraction where biomass-degrading enzymes reside. Further functional studies showed that two heme-containing proteins were consistently found to be abundant in the secretome of A. sp. 75iv2 and were subsequently named Amyco1 and Amyco2 [36]. On the basis of sequence homology, the functions of Amyco1 and Amyco2 could be assigned as a bifunctional catalase-peroxidase and catalase, respectively, which was confirmed by in vitro biochemical experiments. Further studies showed that Amyco1 was competent to form the typical two-electron (compound I) and one-electron (compound II) oxidized states found in peroxidases and that it could degrade phenolic lignin model compounds [36]. More recently, A. sp. 75iv2 has been found to be able to grow on lignin as a sole carbon source and studies to further characterize the molecular basis for this phenotype are underway. Bacterial heme peroxidases
At the same time, new phenotypic screens have been designed to help identify novel lignolytic bacterial species. Using a set of fluorescent and colorimetric substrate probes, several previously uncharacterized bacteria were also found to demonstrate the ability to react with lignin [39]. Studies of knockout strains of one of these genera, Rhodoccocus jostii RHA1, implicated dye-decolorizing peroxidases (DyP) in this behavior [40]. DyPs belong to a small but distinct family of heme peroxidases, which contain a distal aspartate residue in place of a histidine [41]. In addition, they have often been found to be responsible for the extracellular oxidation and decomposition of dyes and other aromatic xenobiotics in fungi [42–44]. They separate into four major clades with the smaller and less active bacterial enzymes belonging to the A-clade and B-clade and the larger and more active fungal enzymes residing to the D-clade. C-type DyPs are prokaryotic in origin but cluster more closely with the fungal D-clade and appear to share their high activity with respect to oxidation of typical DyP substrates [45]. Of the two DyPs found in the R. jostii RHA1 genome, the B-type DypB was found to be responsible for the observed reactivity with lignin and was shown to be biochemically competent for the degradation of Current Opinion in Chemical Biology 2014, 19:1–7
4 Bioinorganic chemistry
non-phenolic lignin compounds [40]. While DypB did not contain a canonical secretion sequence, biochemical experiments suggest that it could be exported from the cell through a different mechanism [46]. It was also observed to have low MnP activity (kcat/KM = 16 M 1 s 1) 1) that could amplify its ability to degrade lignin-based compounds. Additional structural and biochemical studies have also been carried out on DypB [47,48] and have aided the engineering of the Mn binding pocket for a 14-fold improvement in the MnP activity [49]. While not as widely distributed in bacterial genomes as other heme peroxidases or cytochromes, genes encoding DyPs are found to be prevalent in actinomycetes. Interestingly, A. sp. 75iv2 contains three DyPs (DyP1–3), one of which is a typical bacterial A-type DyP (DyP3) and two of which belong to the C-clade (DyP1-2). The presence of two C-type DyPs was unexpected as they are observed less frequently but are often clustered with other genes encoding biomass-degrading enzymes in actinomycetes [50]. In particular, DyP2 belonged to a particularly divergent and sparsely populated branch within the C-clade and was thus selected for additional biochemical and structural characterization. In vitro assays with various substrates showed that DyP2 had high peroxidase and MnP activity (kcat/KM = 105–106 M 1 s 1) near the same magnitude as fungal enzymes, including those involved in lignin degradation [50]. In addition, DyP2 was competent to oxidize higher potential substrates such as Reactive Black 5. Another interesting observation was that the presence of Mn(II) enabled a secondary oxygenase activity that expanded the substrate scope of DyP2 to include the degradation of non-phenolic lignin model compounds [50]. The high MnP and secondary Mn oxygenase activity appears to be related to the relatively high affinity of DyP2 for Mn(II) that was supported by the discovery of a metal-binding site in the crystal structure that was localized 15 A˚ from the heme site.
Given the breadth of bacterial enzymes and pathways that functionalize or degrade aromatics, this discovery may indicate that many bacteria may interact differently with lignin as a substrate for use as an aromatic carbon source rather than carry out its rapid degradation to access the sugar-based cellulose and hemicellulose substrates as is believed to be the case in many fungal systems. Discovery of new lignin-reactive bacteria and study of microbial consortia
Certainly the exploration of greater microbial biodiversity in new environments will help to extend our understanding of the scope of lignin metabolism in microbes. Scaledup screening studies have identified many new bacterial species that appear to be much more active than previously characterized strains [52]. Indeed, the study of these new microbes may reveal the presence of new mechanisms for lignin reactivity especially when they originate from unusual environments where active biomass degradation occurs. For instance, greater availability of microbial metagenomes from sources where biomass is rapidly degraded — such as wood-feeding insects [53–55], the rumen of cows [56], or active soils [57] — will allow us to assess greater diversity at the genetic level. In particular, environments where lignin degradation occurs under anaerobic or microaerobic conditions [58,59] may be especially interesting as canonical lignin degradation involves peroxide- or oxygen-dependent enzymes. Indeed, insufficient knowledge of the suite of genes involved in lignin modification creates roadblocks in functional annotation of metagenomes by sequence homology alone [53]. However, continuing physiological [60] and structural studies [61] could help to identify new chemical transformations on lignin as a result of microbial metabolism. In addition, isolation or comparative genomics studies of microbes from environments such as rainforest soil [62] or forest areas [63,64,65] also serve to improve our understanding of the function of various members of microbial consortia.
Other bacterial enzymes involved in lignin processing
In general, the heme peroxidases from these lignin-reactive bacteria have been found to be less oxidatively powerful compared to the fungal enzymes involved in lignin degradation. However, it is possible that heme peroxidases do not constitute the major lignolytic enzymes in bacterial systems. Beyond the canonical enzymes used by fungi, bacteria also contain a suite of oxidative enzymes that could modify lignin for breakdown by hydroxylation or demethylation, such as cytochrome P450s, non-heme iron enzymes, or Mn- and Cu-containing oxidases. In one interesting example, a gene encoding a non-heme iron dioxygenase fused to a carbohydrate binding domain (CBD) was found from a cellulolytic streptoycetes isolated from wood-boring insects [51]. Biochemical studies of SACTE_2871 showed that was competent to bind lignin polymers as well as catalyze O2-dependent ring-opening of catechols. Current Opinion in Chemical Biology 2014, 19:1–7
Conclusions Lignin is a complex substrate that is known to require a suite of oxidative enzymes and assorted small molecule co-factors for its degradation. By exploring greater diversity of the organisms that can react with and metabolize lignin, we may be able to gain new insight into strategies for biomass deconstruction. Bacterial systems have been found to be less oxidatively powerful compared to lignolytic fungal systems to date but may provide a rich source for elucidating new accessory enzymes that could act synergistically with the major oxidative enzymes to activate and uncap various sites, similar to what has been involved in cellulose degradation. For example, hydroxylation or demethylation of various sites could serve to generate new chemical handles for further degradation. In addition, new studies showing the existence of oxidative enzymes that are capable of associating with lignin www.sciencedirect.com
Exploring bacterial lignin degradation Brown and Chang 5
could indicate that this process is not completely mediated through secondary small molecule mediators. In this regard, the existence of direct interactions between the substrate and lignin processing enzymes may also begin to explain differential reactivity profiles that are observed. Taken together, work in this area will continue to help bring more molecular detail into understanding how lignin degradation occurs in the environment.
Acknowledgements Research in our laboratory in this area is funded by the generous support of the University of California, Berkeley and the Energy Biosciences Institute.
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