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ScienceDirect Enzymatic conversion of lignin into renewable chemicals Timothy DH Bugg and Rahman Rahmanpour The aromatic heteropolymer lignin is a major component of plant cell walls, and is produced industrially from paper/pulp manufacture and cellulosic bioethanol production. Conversion of lignin into renewable chemicals is a major unsolved problem in the development of a biomass-based biorefinery. The review describes recent developments in the understanding of bacterial enzymes for lignin breakdown, such as DyP peroxidases, bacterial laccases, and beta-etherase enzymes. The use of pathway engineering methods to construct genetically modified microbes to convert lignin to renewable chemicals (e.g. vanillin, adipic acid) via fermentation is discussed, and the search for novel applications for lignin (e.g. carbon fibre). Address Department of Chemistry, University of Warwick, Coventry CV4 7AL, United Kingdom Corresponding author: Bugg, Timothy DH ([email protected])
Current Opinion in Chemical Biology 2015, 29:10–17 This review comes from a themed issue on Energy Edited by Timothy DH Bugg and Michael Resch
http://dx.doi.org/10.1016/j.cbpa.2015.06.009 1367-5931/# 2015 Elsevier Ltd. All rights reserved.
The need to reduce global greenhouse gas emissions has stimulated considerable interest in the generation of new routes to fuels and chemicals from renewable sources, especially from plant biomass [1]. For aromatic chemicals used to make plastics, fine chemicals and materials, the aromatic polymer lignin found in plant cell walls is a readily available but challenging substrate [2,3]. This article will review current research, challenges and prospects for bio-conversion of lignin into renewable chemicals. Since the invention of the oil refinery in the late 19th century, aromatic chemicals have been produced industrially from crude oil, mainly from catalytic reforming of the naphtha fraction, and are used for a range of highvolume industrial applications [2], including plastics from polystyrene (from styrene), polyethylene terephthalate (PET, from para-xylene), and phthalate resins Current Opinion in Chemical Biology 2015, 29:10–17
(from ortho-xylene). An aromatic component of 15–20% is needed in jet fuel, for which renewable sources are needed [4]. As supplies of crude oil dwindle around the world, alternative sustainable sources for these chemicals must be found, for which the aromatic lignin polymer found in plant biomass is an abundant raw material (see Figure 1).
Lignin availability and structural types Lignin is produced on a large scale from the pulp and paper industry [5,6]. The Kraft process involves treatment of biomass with sodium hydroxide and sodium sulphide, generating Kraft lignin (60–100 ktons/yr) as a by-product. The Sulfite process involves treatment with aqueous sulphur dioxide, generating water-soluble lignosulfonates (1 Mtonne/yr). The soda lignin process involves treatment with aqueous sodium hydroxide, generating a sulphur-free soda lignin (5–10 ktons/yr). The Organosolv process uses an ethanol–water extraction method to produce high purity cellulose, hemicellulose and lignin streams from plant biomass, which generates three ktons/yr Organosolv lignin (CIMV, Fr). Potentially even larger amounts of lignin will be produced from the industrial production of cellulosic bioethanol production, which liberates lignin as a low-value byproduct. It has been estimated that the US bioethanol industry alone will generate up to 60 Mtonne/yr lignin by 2022 [2], and large-scale bioethanol production sites at Alagoas in Brazil (82 Ml/yr) and Crescentino in Italy (20 Ml/yr) will also generate lignin-rich product streams. Finding new product streams for lignin is therefore a major unsolved challenge in creating a lignocellulosebased biorefinery. However, it is also important to note that the chemical properties of each type of lignin vary considerably. Depending on the type of chemical pre-treatment used in its production, industrial lignin may be highly ‘condensed’, due to loss of benzylic hydroxyl groups in the b-aryl ether structural unit, forming a benzylic cation that can form new C–C cross-linking bonds. Kraft lignin is often highly condensed, and therefore contains greatly reduced b-O-4 content, and also contains thiol groups arising from the use of sodium sulphide (Na2S) used in the Kraft process [3,5,7]. The Organosolv process uses a milder biomass treatment method, hence Organosolv lignin retains a higher proportion of b-O-4 linkages [8], but to date the Organosolv process has only been commercialised at pilot scale [5]. There is also current interest www.sciencedirect.com
Enzymatic conversion of lignin Bugg and Rahmanpour 11
Figure 1
Oil Refinery
Fractions
Fuels
Chemicals
Petroleum C5 -C8
Petroleum OH
Catalytic reforming
naphtha
Kerosene C6-C16
jet fuel
Diesel oil C8-C21
diesel
CO2H
CO2H
e crude oil cellulose O
OH
Bio-Refinery
O
O HO
cellulosic bioethanol
O
O HO
OH
O OH
O
OHC
n
CH2OH
OMe
Cellulosic biorefinery
Biomass
CHO
OH
OH CHO
O HO OH
O
OMe
vanillin 15 $/kg
?
OMe OH
O
HO
OMe
Pulp/paper manufacture
OMe O
Biorefinery lignin Kraft lignin Lignosulfonate Soda lignin
CO2H
ferulic acid 85 $/kg OMe OH Current Opinion in Chemical Biology
Routes to aromatic chemicals from crude oil (a) and plant biomass (b).
in ionic liquid biomass treatment methods [9], which generate an ionic liquid lignin preparation containing reduced b-O-4 content, and some condensed structures [10].
Challenges in lignin valorisation There has been research into lignin valorisation since the 1980s [11], but there have been very few successful examples of the conversion of lignin into higher value aromatic products, due to a number of challenges, as follows. First, Difficult bond cleavages. The linkages between aryl-C3 units in polymeric lignin are either C–O ether bonds, or C–C bonds, which are not susceptible to hydrolytic cleavage and hence chemically inert, so lignin breakdown requires unusual chemistry or biochemistry. Second, Physical properties. Native lignin from biomass, and more native-like lignins such as Organosolv lignin, are rather insoluble in water and organic solvents, though www.sciencedirect.com
industrial Kraft lignins and lignosulfonates are more water-soluble. Access to the lignin polymer is therefore challenging for both enzyme and chemical catalysts. Third, Heterogeneous structure. Compared with cellulose, the lignin polymer is inherently heterogeneous and irregular in structure, presenting a challenge for enzyme biocatalysts, which are usually highly selective for a particular substrate. Fourth, One lignin is not the same as another lignin. The chemical structure of a lignin preparation is dependent on the method used to prepare the lignin from biomass. In the case of industrial lignins, the lignin structure is often condensed, making it more chemically inert, and the presence of sulfur in Kraft lignins can poison chemical catalysts. Fifth, Repolymerisation vs. depolymerisation. Often the catalysts for lignin depolymerisation will also catalyse the repolymerisation of radical intermediates, resulting in competing repolymerisation (or recondensation) of lignin fragments, generating higher Current Opinion in Chemical Biology 2015, 29:10–17
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molecular weight products. Sixth, Generation of complex mixtures of products. Since lignin is a complex, heterogeneous polymeric substrate, it is not surprising that chemical or bio-catalytic processing of lignin would result in a rather complex mixture of oligomeric fragments and low molecular weight products. Typically, complex mixtures of low molecular weight aromatic products are produced by pyrolytic and chemocatalytic treatment of lignin, hence there is a need for methods to simplify product mixtures by methods such as hydro-deoxygenation, to obtain a smaller number of deoxygenated alkane [3].
Enzymes for lignin breakdown The bio-degradation of lignin by lignin-degrading microbes has been described as ‘enzymatic combustion’, where the oxidising potential of hydrogen peroxide or molecular oxygen by ligninolytic peroxidase enzymes or laccase enzymes respectively is exploited to oxidise aromatic units [11]. White-rot fungi such as Phanerochaete chrysosporium produce an arsenal of extracellular lignin peroxidases, Mn peroxidases and laccases for lignin breakdown, which have been extensively reviewed [12,13]. The development of fungal biocatalysts for large-scale lignin depolymerisation has been hampered by the challenges of fungal protein expression, therefore, in recent years there has been renewed interest in the identification of bacterial biocatalysts for lignin depolymerisation [14]. Bacterial lignin degradation activity has been best characterised in actinobacteria, a- and g-proteobacteria, and in some cases related bacteria have been reported in the gut of wood-infesting termites and beetles [14]. A recent review of lignin-degrading phenotypes and genotypes lists 22 actinobacteria, 10 a-proteobacteria and 11 g-proteobacterial strains with lignin degradation phenotypes, but also seven firmicutes, four b-proteobacteria, one d-proteobacterium, one bacteroides and one archaeal strain [15]. Until recently, the enzymology of bacterial lignin degradation was not well understood [14]. Using a spectroscopic assay method, Ahmad et al. have shown lignin degradation activity in the extracellular fraction of two known aromatic degraders, Rhodococcus jostii RHA1 and Pseudomonas putida [16]. Bioinformatic analysis of unannotated peroxidase genes in the R. jostii RHA1 genome sequence led to the identification of two dye-decolorizing peroxidase (DyP) genes dypA and dypB, with homologues in other lignin-oxidising bacteria [17]. Gene deletion studies revealed that a DdypB mutant shows significant decreased lignin degradation activity, and recombinant DypB catalyses oxidative Ca–Cb cleavage of a b-aryl ether lignin model compound, and also oxidises Mn2+ [17]. The structure of DypB from R. jostii RHA1 shows a ferredoxin-like fold, similar to other DyP structures [18]. Replacement of active site Asn-246 in R. jostii DypB by Ala has been found to increase the kcat for Mn2+ oxidation 80-fold, and a Mn2+ binding site has been identified by Current Opinion in Chemical Biology 2015, 29:10–17
X-ray crystallography, as shown in Figure 2a [19]. Remarkably, the R. jostii RHA1 DypB protein is packaged into a nanocompartment formed by 60 protein subunits encoded by an adjacent encapsulin gene, and encapsulation increases its lignin oxidation activity [20]. The DyPs, discovered nearly a decade ago, form a distinct superfamily of peroxidases, due to their specific primary and tertiary structures and unique reaction characteristics [21]. Phylogenetic analyses have led to the classification of DyPs into four subfamilies (A–D), DyPs from bacteria belong to the A, B and C subfamilies (see Figure 2b), whereas fungal enzymes are found in the D subfamily [22]. A DyP1B enzyme from Gram-negative Pseudomonas fluorescens Pf-5 has been reported to show activity for oxidation of Kraft lignin and Mn2+, and in the presence of Mn2+ releases an oxidised lignin dimer from wheat straw lignocellulose [23]. Heme-dependent lignin-oxidising enzymes have also been identified in soil bacterium Amycolatopsis sp. 75iv2 ATCC 39116 [24]. A Dyp type C peroxidase enzyme has been identified from the same strain which shows Mn2+ oxidation activity with much higher catalytic efficiency than R. jostii DypB, approaching the activity of fungal Mn peroxidase enzymes [25]. Laccase enzymes from fungi have been utilized in different areas such as wood and paper industries, environmental applications and biosensors, and have activity for lignin oxidation [12,13], but few prokaryotic laccases have been discovered. Bioinformatic analysis of >2200 complete and draft bacterial genomes and four metagenomic datasets has identified >1200 putative bacterial genes for laccase-like enzymes, of which 76% showed the existence of putative signal sequences, implying that the encoded enzymes might be exported from the cytoplasm [26]. Bacterial laccases are especially prevalent in actinobacteria [27], but are also found in a-, b- and g-proteobacteria [15]. Laccase enzymes from Streptomyces coelicolor A3(2), Streptomyces lividans TK24, Streptomyces viridosporus T7A, and Amycolatopsis sp. 75iv2 have been characterised kinetically, and were found to catalyse Ca oxidation of lignin model compounds [28]. Deletion of the laccase gene from S. coelicolor A3(2) was found to significantly diminish the ability to form acid-precipitable lignin (APPL), supporting a role in lignin oxidation in vivo, but treatment of ethanosolv lignin in vitro led to a higher molecular weight product, due to competing repolymerisation of lignin fragments [28]. A multicopper oxidase similar to Pseudomonas stutzeri CopA has also been identified from a novel screen of metagenomic DNA libraries using an ermR reporter gene that responds to aromatic lignin degradation products [29]. This multicopper oxidase was found to oxidise a polymeric lignin substrate to generate 2,6-dimethoxybenzene-1,4-diol as a major product [29]. Fungal laccase enzymes have been shown to be effective biocatalysts for delignification of wood and biomass feedstocks, using a suitable laccase mediator www.sciencedirect.com
Enzymatic conversion of lignin Bugg and Rahmanpour 13
Figure 2
(a)
(b)
0.115
0.128 0.136 0.190
0.022 0.019 0.306
A 0.000
Burkholderia terrae Rhodococcus josti RHA1 DyPB ∗ 0 022 Rhodococcus wratislaviensis 0.044 Rhodococcus opacus 0.059 Pseudomonas fluorescens DyP2B 0.057 Pseudomonas chlororaphis O6 0.079 Pseudomonas brassicacearum 0.348 Pseudomonas fluorescens DyPA 0.233 Thermobifida fusca DyPA 0.244 Nocardiopsis dassonvillei 0.163 Rhodococcus RHA1 DyPA ∗ 0.183 Mycobacterium smegmatis 0.165 Amycolatopsis DyP2C 0.038 Amycolatopsis vancoresmycina 0.136 0.037 Amycolatopsis mediterranei 0.216 Rhizobium leguminosarum
0.025
B
0.069
0.078 0.032 0.161
0.025
C
0.239
Pseudomonas fluorescens DyP1B ∗
0.125
0.024
0.220 Current Opinion in Chemical Biology
Bacterial DyP peroxidases. (a) Structure of Rhodococcus jostii RHA1 DypB, showing the location of the Mn2+ binding site (Glu-156, Glu-215, Thr-231, Glu-239, in pink) close to the heme cofactor (in red) and catalytic residues Asp-153 and Arg-244 (in blue). (b). Phylogenetic cladogram for bacterial DyP peroxidases, showing DyP groups A, B, and C; enzymes mentioned in the text are starred.
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Current Opinion in Chemical Biology 2015, 29:10–17
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[30], hence bacterial laccases might have interesting applications for delignification, as well as lignin valorisation. Bacterial b-etherase enzymes have also been recently discovered that catalyse a glutathione-dependent cleavage reaction on lignin model compounds. This class of b-etherase enzymes were previously identified in Sphingobium SYK-6, a strain capable of breakdown of a number of lignin model compounds, and were shown to be dependent on glutathione, whose nucleophilic thiol sidechain was used to cleave the ether linkage of b-aryl ether model compounds [31]. Picart et al. have recently identified four b-etherase enzymes in Novosphingobium strains, which catalyse b-ether cleavage on lignin model compounds, and also show activity towards a fluorescently labelled polymeric lignin substrate [32]. Gall et al. have also characterised b-etherase enzymes from Sphingobium SYK-6 [33] and from two Novosphingobium strains [34], demonstrating that different sub-classes show opposite stereospecificity for the b-aryl ether b-carbon centre: LigE-type enzymes are selective for the R-enantiomer, while LigF-type enzymes are selective for the S-enantiomer [33,34]. Table 1 summarises the properties of the currently known bacterial enzymes for lignin breakdown, only some of which are reported to be active with polymeric lignin, rather than lignin model compounds. It seems likely that a group of oxidative enzymes (and accessory proteins) are needed to break down such a complex substrate, so new genome sequence data and metagenomic data are likely to identify further enzymes. The genome of an Enterobacter lignolyticus strain which shows activity for lignin degradation under anaerobic conditions contains many glutathione S-transferase enzymes which might be related to the LigEF b-etherase enzymes, as well as two
putative laccases [35], while the genome of a Cupriavidus basilensis B-8 strain which oxidises Kraft lignin contains one putative laccase, and a range of aromatic degradation gene clusters [36].
Manipulating lignin degradation pathways to produce fuels and chemicals As an alternative to using enzymes in vitro to convert polymeric lignin into chemicals, pathway engineering could potentially be used to engineer the lignin degradation pathways of a microbe that possesses the metabolic capability to break down lignin, in order to produce useful chemicals. Some recent examples suggest that this may be a practical way forward. While our knowledge of microbial lignin degradation pathways is still incomplete [37], it seems likely that the initial lignin oxidation products are processed via downstream aromatic degradation pathways that are well understood, such as the b-ketoadipate pathway involving intradiol cleavage of protocatechuic acid to 3-carboxymuconic acid, as shown in Figure 3 [38]. Protocatechuic acid and catechol are also metabolised via extradiol cleavage in some bacteria [37,38], hence these metabolites are key branchpoints for aromatic degradation. There are indications that vanillic acid is an intermediate in G unit breakdown by lignin-degrading bacteria, since it has been detected as a metabolite from lignin breakdown [14,37]. Sainsbury et al. have shown deletion of the vanillin dehydrogenase gene on the vanillin degradation pathway in R. jostii RHA1 gave a mutant strain which accumulated vanillin, a high value chemical used in the food/flavour industry, at up to 96 mg/L after six days when grown on minimal media containing 2.5% wheat straw lignocellulose, together with 4-hydroxybenzaldehyde and ferulic acid [39], as shown in Figure 3. Primary metabolism can also be used to generate useful bioproducts:
Table 1 Summary of bacterial enzymes for lignin breakdown. NR, not reported Enzyme
Bacterium
Cofactor
Co-substrate
Low MW substrates
Lignin model compounds
Rhodococcus jostii RHA1 Pseudomonas fluorescens Amycolatopsis sp 75iv2 Streptomyces coelicolor Pseudomonas stutzeri Sphingobium SYK6 Novosphingobium sp.
Heme Fe
H2O2
ABTS, Mn(II)
b-O-4
Heme Fe
H2O2
ABTS, Mn(II)
NR
Heme Fe
H2O2
ABTS, Mn(II)
b-O-4
NR
NR
Cu
O2
ABTS, DMP
b-O-4
Ca oxidation
Ethanosolv
Higher MW
[28]
Cu
O2
ABTS, DMP
b-O-4
NR
HP lignin
Aromatic monomer
[29]
Glutathione
b-O-4
NR
[33]
Glutathione
b-O-4
b-Ether cleavage b-Ether cleavage
Fluorescent lignin
[32]
Substrate DypB
Dyp2 Laccase CopA Etherase
Current Opinion in Chemical Biology 2015, 29:10–17
Polymeric lignin
Reaction Ca–Cb cleavage
Substrate
Products [17]
Kraft lignin Lignocellulose
Ref
Lignin dimer
[23] [25]
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Enzymatic conversion of lignin Bugg and Rahmanpour 15
Figure 3
CO2H
Hemicellulose
OCH3 OH
OH HO
OAr
G type lignin
CHO
CO2H
OCH3 OH
OH
X
OAr
H type lignin
CO2H
OCH3
protocatechuate 3,4-dioxygenase
OH
CO2H
O
OH
CO2H CO2H
OH
OH
OH catechol
H2/cat HO2C
O
CO2H
O triacylglycerol lipids
CO2H
cis,cismuconic acid
X OH
CO2H
CO2H CO2H CO2H
OH
OH
vanillin
CHO
HO
β-ketoadipic acid
protocatechuic acid
OCH3 OH
β-oxidation
TCA cycle
O O
O O
polyhydroxy butyrate
SCoA
C8H17 O O
malonyl CoA
O O n Current Opinion in Chemical Biology
Production of bioproducts from microbial degradation pathways, illustrating schematically the metabolic pathways responsible for accumulation of small molecule products vanillin or cis,cis-muconic acid via gene deletion (in blue), and the accumulation of triacylglycerol lipids or polyhydroxybutyrate via primary metabolism (in green).
Rhodococcus opacus DSM1069 and PD630 are known to accumulate triglyceride lipids from primary metabolism under nitrogen-limiting conditions, and have been shown to grow on minimal media containing ultrasonicated ethanol organosolv lignin as carbon source, generating triglyceride lipids at 4 mg/g us-EOL (or 20 mg/L) after a nine day fermentation [40]. P. putida is a well-known aromatic degrader that also possesses activity for lignin breakdown [16]. Linger et al. have shown that the ability of P. putida to accumulate polyhydroxyalkanoate (PHA) biopolyesters under nitrogen-limiting conditions can be harnessed to convert lignin from alkaline pre-treated liquor (APL) into PHAs in a 48 h fermentation [41]. Pathway engineering has also been used in P. putida to accumulate cis,cis-muconic acid from aromatic degradation, via blockage of the protocatechuate cleavage pathway, and re-routing via catechol cleavage [42]. Using p-coumaric acid as a carbon source, a yield of 13.5 g/L was obtained in a fed-batch bioreactor after 78 h, whereas using alkali-APL a yield of 0.7 g/L was obtained after 24 h [42]. Cis,cis-muconic acid can then be www.sciencedirect.com
converted via catalytic hydrogenation into adipic acid, thereby providing a renewable route to adipic acid from lignin [42]. As we improve our understanding of lignin degradation, it should be possible to use pathway engineering in suitable microbial hosts to generate other useful products from microbial lignin breakdown, hence it is interesting that replacement of the genes encoding the catechol intradiol pathway in P. putida by genes encoding a corresponding extradiol pathway leads to an increase in yield of pyruvate from aromatic substrates [43]. A selective hydroxamic acid inhibitor of protocatechuate 3,4-dioxygenase has also been developed, as an alternative chemical approach to manipulating bacterial lignin degradation [44].
Industrial applications for lignin There is interest in finding new applications for polymeric lignins, as well as depolymerised lignin breakdown products. Jin et al. have used lignin derived from corn bioethanol production to make phenol-formaldehyde adhesive [45], while Ramires et al. have used Organosolv lignin from sugarcane bagasse to prepare phenolic resins Current Opinion in Chemical Biology 2015, 29:10–17
16 Energy
[46]. In principle polymeric lignin could be a template for synthesis of new biomaterials. It has been shown that Kraft lignin can be modified by atom transfer radical polymerisation (ATRP), changing the thermostability and water solubility of the polymer [47]. Polyurethane foams can also be made from either Organosolv or Kraft lignin, with stronger foams being formed from Organosolv lignin [48]. There is interest in the conversion of lignin into carbon fibre, however, there are challenges in generating high quality carbon fibre, notably that highly purified lignin fractions are needed, due to the heterogeneity of most lignin preparations [49]. In a related application, lignin has also been converted into porous carbon, suitable for supercapacitor electrode materials [50]. The high carbon content of lignin makes it potentially interesting for conversion into nanomaterials, and Yiamsawas et al. have recently shown that lignin nanocontainers can be formed from water-soluble lignosulfonates in inverse micro-emulsions [51].
Acknowledgement The authors thank the University of Warwick for funding a Chancellor’s Scholarship to R.R.
References and recommended reading Papers of particular interest, published within the period of review, have been highlighted as: of special interest of outstanding interest
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24. Brown ME, Walker MC, Nakashige TG, Iavarone AT, Chang MCY: Discovery and characterization of heme enzymes from unsequenced bacteria: application to microbial lignin degradation. J Am Chem Soc 2011, 133:18006-18009. 25. Brown ME, Barros T, Chang MCY: Identification and characterization of a multifunctional dye peroxidase from a lignin-reactive bacterium. ACS Chem Biol 2012, 7:2074-2081. The Amycolatopsis sp 75iv2 Dyp2 shows much higher activity for Mn(II) oxidation, approaching the activity for fungal Mn peroxidases. 26. Ausec L, Zakrzewski M, Goesmann A, Schlu¨ter A, Mandic-Mulec I: Bioinformatic analysis reveals high diversity of bacterial genes for laccase-like enzymes. PLoS ONE 2011, 10:25724-25732. 27. Fernandes TAR, da Silveira WB, Passos FML, Zucchi TD: Laccases from actinobacteria — what we have and what to expect. Adv Microbiol 2014, 4:285-296. 28. Majumdar S, Lukk T, Solbiati JO, Bauer S, Nair SK, Cronan JE, Gerlt JA: Roles of small laccases from Streptomyces in lignin degradation. Biochemistry 2014, 53:4047-4058. This paper characterises in detail the activity of some bacterial laccase enzymes, and demonstrates that they have some role in lignin degradation. www.sciencedirect.com
Enzymatic conversion of lignin Bugg and Rahmanpour 17
29. Strachan CR, Singh R, VanInsberghe D, Ievdokymenko K, Budwill K, Mohn WW, Eltis LD, Hallam SJ: Metagenomic scaffolds enable combinatorial lignin transformation. Proc Natl Acad Sci U S A 2014, 111:10143-10148. 30. Gutierrez A, Rencoret J, Cadena EM, Rico A, Barth D, del Rio JC, Martinez AT: Demonstration of laccase-based removal of lignin from wood and non-wood plant feedstocks. Bioresour Technol 2012, 119:114-122. 31. Masai E, Ichimura A, Sato Y, Miyauchi K, Katayama Y, Fukuda M: Roles of enantioselective glutathione S-transferases in cleavage of b-aryl ether. J Bacteriol 2003, 185:1768-1775. 32. Picart P, Mu¨ller C, Mottweiler J, Wiermans L, Bolm C, Dominguez de Maria P, Schallmey A: From gene towards selective biomass valorization: bacterial b-etherases with catalytic activity on lignin-like polymers. ChemSusChem 2014, 7:3164-3171. This paper (and the following reference) demonstrates that b-etherase enzymes could be useful biocatalysts for lignin breakdown. 33. Gall DL, Kim H, Lu F, Donohoe TJ, Noguera DR, Ralph J: Stereochemical features of glutathione-dependent enzymes in the Sphingobium sp. strain SYK-6 b-aryl etherase pathway. J Biol Chem 2014, 289:8656-8667. This paper highlights the stereospecificity of the b-etherase enzyme family. 34. Gall DL, Ralph J, Donohoe TJ, Noguera DR: A group of sequencerelated sphingomonad enzymes catalyzes cleavage of b-aryl ether linkages in lignin b-guaiacyl and b-syringyl ether dimers. Environ Sci Technol 2014, 48:12454-12463. 35. De Angelis KM, D’Haeseleer P, Chivian D, Fortney JL, Khudyakov J, Simmons B, Woo H, Arkin AP, Davenport KW, Goodwin L et al.: Complete genome sequence of Enterobacter lignolyticus SCF1. Stand Genomic Sci 2011, 5:69-85. 36. Shi Y, Chai L, Tang C, Yang Z, Zhang H, Chen R, Chen Y, Zheng Y: Characterization and genomic analysis of kraft lignin biodegradation by the beta-proteobacterium Cupriavidus basilensis B-8. Biotech Biofuels 2013, 6:1-14. 37. Bugg TDH, Ahmad M, Hardiman EM, Rahmanpour R: Pathways for degradation of lignin in bacteria and fungi. Nat Prod Rep 2011, 28:1883-1896. 38. Bugg TDH, Winfield CJ: Enzymatic cleavage of aromatic rings: mechanistic aspects of the catechol dioxygenases and later enzymes of bacterial aromatic degradation pathways. Nat Prod Rep 1998, 15:513-530. 39. Sainsbury PD, Hardiman EM, Ahmad M, Otani H, Seghezzi N, Eltis LD, Bugg TDH: Breaking down lignin to high-value chemicals: the conversion of lignocellulose to vanillin in a gene deletion mutant of Rhodococcus jostii RHA1. ACS Chem Biol 2013, 8:2151-2156. This was the first demonstration that microbial lignin degradation pathways could be manipulated to produce aromatic chemicals from lignin via fermentation.
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40. Kosa M, Ragauskas AJ: Lignin to lipid bioconversion by oleaginous Rhodococci. Green Chem 2013, 15:2070-2074. 41. Linger JG, Vardon DR, Guarneri MT, Karp EM, Hunsinger GB, Franden MA, Johnson CW, Chupka G, Strathmann TJ, Pienkos PT, Beckham GT: Lignin valorization through integrated biological funnelling and chemical catalysis. Proc Natl Acad Sci U S A 2014, 111:12013-12018. This was a nice demonstration of how bacterial degradation of lignin could be harnessed, via primary metabolism, to produce valuable biosynthetic products. 42. Vardon DR, Franden MA, Johnson CW, Karp EM, Guarneri MT, Linger JG, Salm MJ, Strathman TJ, Beckham GT: Adipic acid production from lignin. Energy Environ Sci 2015, 8:617-628. A nice example of pathway engineering in Pseudomonas putida to produce a metabolite that can be converted chemically into adipic acid. 43. Johnson CW, Beckham GT: Aromatic catabolic pathway selection for optimal production of pyruvate and lactate from lignin. Metab Eng 2015, 28:240-247. 44. Sainsbury PD, Mineyeva Y, Mycroft Z, Bugg TDH: Chemical intervention in bacterial lignin degradation pathways: development of selective inhibitors for intradiol and extradiol catechol dioxygenases. Bioorg Chem 2015, 60:102-109. 45. Jin Y, Cheng X, Zheng Z: Preparation and characterization of phenol-formaldehyde adhesives modified with enzymatic hydrolysis lignin. Biores Technol 2010, 101:2046-2048. 46. Ramires EC, Megiatto JD Jr, Gardrat C, Castellan A, Frollini E: Valorization of an industrial organosolv-sugarcane bagasse lignin: characterization and use as a matrix in biobased composites reinforced with sisal fibers. Biotech Bioeng 2010, 107:612-621. 47. Kim YS, Kadla JF: Preparation of a thermoresponsive ligninbased biomaterial through atom transfer radical polymerization. Biomacromolecules 2010, 11:981-988. 48. Pan X, Saddler JN: Effect of replacing polyol by organosolv and kraft lignin on the property and structure of rigid polyurethane foam. Biotech Biofuels 2013, 6:12. 49. Baker DA, Rials TG: Recent advances in low-cost carbon fiber manufacture from lignin. J Appl Polym Sci 2013, 130:713-728. 50. Jeon J-W, Zhang L, Lutkenhaus JL, Laskar DD, Lemmon JP, Choi D, Nandasiri MI, Hashmi A, Xu J, Motkuri RK, Fernandez CA et al.: Controlling porosity in lignin-derived nanoporous carbon for supercapacitor applications. ChemSusChem 2015, 8:428-432. 51. Yiamsawas D, Baier G, Thines E, Landfester K, Wurm FR: Biodegradable lignin nanocontainers. RSC Adv 2014, 4:11661-11663.
Current Opinion in Chemical Biology 2015, 29:10–17
ScienceDirect Enzymatic conversion of lignin into renewable chemicals Timothy DH Bugg and Rahman Rahmanpour The aromatic heteropolymer lignin is a major component of plant cell walls, and is produced industrially from paper/pulp manufacture and cellulosic bioethanol production. Conversion of lignin into renewable chemicals is a major unsolved problem in the development of a biomass-based biorefinery. The review describes recent developments in the understanding of bacterial enzymes for lignin breakdown, such as DyP peroxidases, bacterial laccases, and beta-etherase enzymes. The use of pathway engineering methods to construct genetically modified microbes to convert lignin to renewable chemicals (e.g. vanillin, adipic acid) via fermentation is discussed, and the search for novel applications for lignin (e.g. carbon fibre). Address Department of Chemistry, University of Warwick, Coventry CV4 7AL, United Kingdom Corresponding author: Bugg, Timothy DH ([email protected])
Current Opinion in Chemical Biology 2015, 29:10–17 This review comes from a themed issue on Energy Edited by Timothy DH Bugg and Michael Resch
http://dx.doi.org/10.1016/j.cbpa.2015.06.009 1367-5931/# 2015 Elsevier Ltd. All rights reserved.
The need to reduce global greenhouse gas emissions has stimulated considerable interest in the generation of new routes to fuels and chemicals from renewable sources, especially from plant biomass [1]. For aromatic chemicals used to make plastics, fine chemicals and materials, the aromatic polymer lignin found in plant cell walls is a readily available but challenging substrate [2,3]. This article will review current research, challenges and prospects for bio-conversion of lignin into renewable chemicals. Since the invention of the oil refinery in the late 19th century, aromatic chemicals have been produced industrially from crude oil, mainly from catalytic reforming of the naphtha fraction, and are used for a range of highvolume industrial applications [2], including plastics from polystyrene (from styrene), polyethylene terephthalate (PET, from para-xylene), and phthalate resins Current Opinion in Chemical Biology 2015, 29:10–17
(from ortho-xylene). An aromatic component of 15–20% is needed in jet fuel, for which renewable sources are needed [4]. As supplies of crude oil dwindle around the world, alternative sustainable sources for these chemicals must be found, for which the aromatic lignin polymer found in plant biomass is an abundant raw material (see Figure 1).
Lignin availability and structural types Lignin is produced on a large scale from the pulp and paper industry [5,6]. The Kraft process involves treatment of biomass with sodium hydroxide and sodium sulphide, generating Kraft lignin (60–100 ktons/yr) as a by-product. The Sulfite process involves treatment with aqueous sulphur dioxide, generating water-soluble lignosulfonates (1 Mtonne/yr). The soda lignin process involves treatment with aqueous sodium hydroxide, generating a sulphur-free soda lignin (5–10 ktons/yr). The Organosolv process uses an ethanol–water extraction method to produce high purity cellulose, hemicellulose and lignin streams from plant biomass, which generates three ktons/yr Organosolv lignin (CIMV, Fr). Potentially even larger amounts of lignin will be produced from the industrial production of cellulosic bioethanol production, which liberates lignin as a low-value byproduct. It has been estimated that the US bioethanol industry alone will generate up to 60 Mtonne/yr lignin by 2022 [2], and large-scale bioethanol production sites at Alagoas in Brazil (82 Ml/yr) and Crescentino in Italy (20 Ml/yr) will also generate lignin-rich product streams. Finding new product streams for lignin is therefore a major unsolved challenge in creating a lignocellulosebased biorefinery. However, it is also important to note that the chemical properties of each type of lignin vary considerably. Depending on the type of chemical pre-treatment used in its production, industrial lignin may be highly ‘condensed’, due to loss of benzylic hydroxyl groups in the b-aryl ether structural unit, forming a benzylic cation that can form new C–C cross-linking bonds. Kraft lignin is often highly condensed, and therefore contains greatly reduced b-O-4 content, and also contains thiol groups arising from the use of sodium sulphide (Na2S) used in the Kraft process [3,5,7]. The Organosolv process uses a milder biomass treatment method, hence Organosolv lignin retains a higher proportion of b-O-4 linkages [8], but to date the Organosolv process has only been commercialised at pilot scale [5]. There is also current interest www.sciencedirect.com
Enzymatic conversion of lignin Bugg and Rahmanpour 11
Figure 1
Oil Refinery
Fractions
Fuels
Chemicals
Petroleum C5 -C8
Petroleum OH
Catalytic reforming
naphtha
Kerosene C6-C16
jet fuel
Diesel oil C8-C21
diesel
CO2H
CO2H
e crude oil cellulose O
OH
Bio-Refinery
O
O HO
cellulosic bioethanol
O
O HO
OH
O OH
O
OHC
n
CH2OH
OMe
Cellulosic biorefinery
Biomass
CHO
OH
OH CHO
O HO OH
O
OMe
vanillin 15 $/kg
?
OMe OH
O
HO
OMe
Pulp/paper manufacture
OMe O
Biorefinery lignin Kraft lignin Lignosulfonate Soda lignin
CO2H
ferulic acid 85 $/kg OMe OH Current Opinion in Chemical Biology
Routes to aromatic chemicals from crude oil (a) and plant biomass (b).
in ionic liquid biomass treatment methods [9], which generate an ionic liquid lignin preparation containing reduced b-O-4 content, and some condensed structures [10].
Challenges in lignin valorisation There has been research into lignin valorisation since the 1980s [11], but there have been very few successful examples of the conversion of lignin into higher value aromatic products, due to a number of challenges, as follows. First, Difficult bond cleavages. The linkages between aryl-C3 units in polymeric lignin are either C–O ether bonds, or C–C bonds, which are not susceptible to hydrolytic cleavage and hence chemically inert, so lignin breakdown requires unusual chemistry or biochemistry. Second, Physical properties. Native lignin from biomass, and more native-like lignins such as Organosolv lignin, are rather insoluble in water and organic solvents, though www.sciencedirect.com
industrial Kraft lignins and lignosulfonates are more water-soluble. Access to the lignin polymer is therefore challenging for both enzyme and chemical catalysts. Third, Heterogeneous structure. Compared with cellulose, the lignin polymer is inherently heterogeneous and irregular in structure, presenting a challenge for enzyme biocatalysts, which are usually highly selective for a particular substrate. Fourth, One lignin is not the same as another lignin. The chemical structure of a lignin preparation is dependent on the method used to prepare the lignin from biomass. In the case of industrial lignins, the lignin structure is often condensed, making it more chemically inert, and the presence of sulfur in Kraft lignins can poison chemical catalysts. Fifth, Repolymerisation vs. depolymerisation. Often the catalysts for lignin depolymerisation will also catalyse the repolymerisation of radical intermediates, resulting in competing repolymerisation (or recondensation) of lignin fragments, generating higher Current Opinion in Chemical Biology 2015, 29:10–17
12 Energy
molecular weight products. Sixth, Generation of complex mixtures of products. Since lignin is a complex, heterogeneous polymeric substrate, it is not surprising that chemical or bio-catalytic processing of lignin would result in a rather complex mixture of oligomeric fragments and low molecular weight products. Typically, complex mixtures of low molecular weight aromatic products are produced by pyrolytic and chemocatalytic treatment of lignin, hence there is a need for methods to simplify product mixtures by methods such as hydro-deoxygenation, to obtain a smaller number of deoxygenated alkane [3].
Enzymes for lignin breakdown The bio-degradation of lignin by lignin-degrading microbes has been described as ‘enzymatic combustion’, where the oxidising potential of hydrogen peroxide or molecular oxygen by ligninolytic peroxidase enzymes or laccase enzymes respectively is exploited to oxidise aromatic units [11]. White-rot fungi such as Phanerochaete chrysosporium produce an arsenal of extracellular lignin peroxidases, Mn peroxidases and laccases for lignin breakdown, which have been extensively reviewed [12,13]. The development of fungal biocatalysts for large-scale lignin depolymerisation has been hampered by the challenges of fungal protein expression, therefore, in recent years there has been renewed interest in the identification of bacterial biocatalysts for lignin depolymerisation [14]. Bacterial lignin degradation activity has been best characterised in actinobacteria, a- and g-proteobacteria, and in some cases related bacteria have been reported in the gut of wood-infesting termites and beetles [14]. A recent review of lignin-degrading phenotypes and genotypes lists 22 actinobacteria, 10 a-proteobacteria and 11 g-proteobacterial strains with lignin degradation phenotypes, but also seven firmicutes, four b-proteobacteria, one d-proteobacterium, one bacteroides and one archaeal strain [15]. Until recently, the enzymology of bacterial lignin degradation was not well understood [14]. Using a spectroscopic assay method, Ahmad et al. have shown lignin degradation activity in the extracellular fraction of two known aromatic degraders, Rhodococcus jostii RHA1 and Pseudomonas putida [16]. Bioinformatic analysis of unannotated peroxidase genes in the R. jostii RHA1 genome sequence led to the identification of two dye-decolorizing peroxidase (DyP) genes dypA and dypB, with homologues in other lignin-oxidising bacteria [17]. Gene deletion studies revealed that a DdypB mutant shows significant decreased lignin degradation activity, and recombinant DypB catalyses oxidative Ca–Cb cleavage of a b-aryl ether lignin model compound, and also oxidises Mn2+ [17]. The structure of DypB from R. jostii RHA1 shows a ferredoxin-like fold, similar to other DyP structures [18]. Replacement of active site Asn-246 in R. jostii DypB by Ala has been found to increase the kcat for Mn2+ oxidation 80-fold, and a Mn2+ binding site has been identified by Current Opinion in Chemical Biology 2015, 29:10–17
X-ray crystallography, as shown in Figure 2a [19]. Remarkably, the R. jostii RHA1 DypB protein is packaged into a nanocompartment formed by 60 protein subunits encoded by an adjacent encapsulin gene, and encapsulation increases its lignin oxidation activity [20]. The DyPs, discovered nearly a decade ago, form a distinct superfamily of peroxidases, due to their specific primary and tertiary structures and unique reaction characteristics [21]. Phylogenetic analyses have led to the classification of DyPs into four subfamilies (A–D), DyPs from bacteria belong to the A, B and C subfamilies (see Figure 2b), whereas fungal enzymes are found in the D subfamily [22]. A DyP1B enzyme from Gram-negative Pseudomonas fluorescens Pf-5 has been reported to show activity for oxidation of Kraft lignin and Mn2+, and in the presence of Mn2+ releases an oxidised lignin dimer from wheat straw lignocellulose [23]. Heme-dependent lignin-oxidising enzymes have also been identified in soil bacterium Amycolatopsis sp. 75iv2 ATCC 39116 [24]. A Dyp type C peroxidase enzyme has been identified from the same strain which shows Mn2+ oxidation activity with much higher catalytic efficiency than R. jostii DypB, approaching the activity of fungal Mn peroxidase enzymes [25]. Laccase enzymes from fungi have been utilized in different areas such as wood and paper industries, environmental applications and biosensors, and have activity for lignin oxidation [12,13], but few prokaryotic laccases have been discovered. Bioinformatic analysis of >2200 complete and draft bacterial genomes and four metagenomic datasets has identified >1200 putative bacterial genes for laccase-like enzymes, of which 76% showed the existence of putative signal sequences, implying that the encoded enzymes might be exported from the cytoplasm [26]. Bacterial laccases are especially prevalent in actinobacteria [27], but are also found in a-, b- and g-proteobacteria [15]. Laccase enzymes from Streptomyces coelicolor A3(2), Streptomyces lividans TK24, Streptomyces viridosporus T7A, and Amycolatopsis sp. 75iv2 have been characterised kinetically, and were found to catalyse Ca oxidation of lignin model compounds [28]. Deletion of the laccase gene from S. coelicolor A3(2) was found to significantly diminish the ability to form acid-precipitable lignin (APPL), supporting a role in lignin oxidation in vivo, but treatment of ethanosolv lignin in vitro led to a higher molecular weight product, due to competing repolymerisation of lignin fragments [28]. A multicopper oxidase similar to Pseudomonas stutzeri CopA has also been identified from a novel screen of metagenomic DNA libraries using an ermR reporter gene that responds to aromatic lignin degradation products [29]. This multicopper oxidase was found to oxidise a polymeric lignin substrate to generate 2,6-dimethoxybenzene-1,4-diol as a major product [29]. Fungal laccase enzymes have been shown to be effective biocatalysts for delignification of wood and biomass feedstocks, using a suitable laccase mediator www.sciencedirect.com
Enzymatic conversion of lignin Bugg and Rahmanpour 13
Figure 2
(a)
(b)
0.115
0.128 0.136 0.190
0.022 0.019 0.306
A 0.000
Burkholderia terrae Rhodococcus josti RHA1 DyPB ∗ 0 022 Rhodococcus wratislaviensis 0.044 Rhodococcus opacus 0.059 Pseudomonas fluorescens DyP2B 0.057 Pseudomonas chlororaphis O6 0.079 Pseudomonas brassicacearum 0.348 Pseudomonas fluorescens DyPA 0.233 Thermobifida fusca DyPA 0.244 Nocardiopsis dassonvillei 0.163 Rhodococcus RHA1 DyPA ∗ 0.183 Mycobacterium smegmatis 0.165 Amycolatopsis DyP2C 0.038 Amycolatopsis vancoresmycina 0.136 0.037 Amycolatopsis mediterranei 0.216 Rhizobium leguminosarum
0.025
B
0.069
0.078 0.032 0.161
0.025
C
0.239
Pseudomonas fluorescens DyP1B ∗
0.125
0.024
0.220 Current Opinion in Chemical Biology
Bacterial DyP peroxidases. (a) Structure of Rhodococcus jostii RHA1 DypB, showing the location of the Mn2+ binding site (Glu-156, Glu-215, Thr-231, Glu-239, in pink) close to the heme cofactor (in red) and catalytic residues Asp-153 and Arg-244 (in blue). (b). Phylogenetic cladogram for bacterial DyP peroxidases, showing DyP groups A, B, and C; enzymes mentioned in the text are starred.
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Current Opinion in Chemical Biology 2015, 29:10–17
14 Energy
[30], hence bacterial laccases might have interesting applications for delignification, as well as lignin valorisation. Bacterial b-etherase enzymes have also been recently discovered that catalyse a glutathione-dependent cleavage reaction on lignin model compounds. This class of b-etherase enzymes were previously identified in Sphingobium SYK-6, a strain capable of breakdown of a number of lignin model compounds, and were shown to be dependent on glutathione, whose nucleophilic thiol sidechain was used to cleave the ether linkage of b-aryl ether model compounds [31]. Picart et al. have recently identified four b-etherase enzymes in Novosphingobium strains, which catalyse b-ether cleavage on lignin model compounds, and also show activity towards a fluorescently labelled polymeric lignin substrate [32]. Gall et al. have also characterised b-etherase enzymes from Sphingobium SYK-6 [33] and from two Novosphingobium strains [34], demonstrating that different sub-classes show opposite stereospecificity for the b-aryl ether b-carbon centre: LigE-type enzymes are selective for the R-enantiomer, while LigF-type enzymes are selective for the S-enantiomer [33,34]. Table 1 summarises the properties of the currently known bacterial enzymes for lignin breakdown, only some of which are reported to be active with polymeric lignin, rather than lignin model compounds. It seems likely that a group of oxidative enzymes (and accessory proteins) are needed to break down such a complex substrate, so new genome sequence data and metagenomic data are likely to identify further enzymes. The genome of an Enterobacter lignolyticus strain which shows activity for lignin degradation under anaerobic conditions contains many glutathione S-transferase enzymes which might be related to the LigEF b-etherase enzymes, as well as two
putative laccases [35], while the genome of a Cupriavidus basilensis B-8 strain which oxidises Kraft lignin contains one putative laccase, and a range of aromatic degradation gene clusters [36].
Manipulating lignin degradation pathways to produce fuels and chemicals As an alternative to using enzymes in vitro to convert polymeric lignin into chemicals, pathway engineering could potentially be used to engineer the lignin degradation pathways of a microbe that possesses the metabolic capability to break down lignin, in order to produce useful chemicals. Some recent examples suggest that this may be a practical way forward. While our knowledge of microbial lignin degradation pathways is still incomplete [37], it seems likely that the initial lignin oxidation products are processed via downstream aromatic degradation pathways that are well understood, such as the b-ketoadipate pathway involving intradiol cleavage of protocatechuic acid to 3-carboxymuconic acid, as shown in Figure 3 [38]. Protocatechuic acid and catechol are also metabolised via extradiol cleavage in some bacteria [37,38], hence these metabolites are key branchpoints for aromatic degradation. There are indications that vanillic acid is an intermediate in G unit breakdown by lignin-degrading bacteria, since it has been detected as a metabolite from lignin breakdown [14,37]. Sainsbury et al. have shown deletion of the vanillin dehydrogenase gene on the vanillin degradation pathway in R. jostii RHA1 gave a mutant strain which accumulated vanillin, a high value chemical used in the food/flavour industry, at up to 96 mg/L after six days when grown on minimal media containing 2.5% wheat straw lignocellulose, together with 4-hydroxybenzaldehyde and ferulic acid [39], as shown in Figure 3. Primary metabolism can also be used to generate useful bioproducts:
Table 1 Summary of bacterial enzymes for lignin breakdown. NR, not reported Enzyme
Bacterium
Cofactor
Co-substrate
Low MW substrates
Lignin model compounds
Rhodococcus jostii RHA1 Pseudomonas fluorescens Amycolatopsis sp 75iv2 Streptomyces coelicolor Pseudomonas stutzeri Sphingobium SYK6 Novosphingobium sp.
Heme Fe
H2O2
ABTS, Mn(II)
b-O-4
Heme Fe
H2O2
ABTS, Mn(II)
NR
Heme Fe
H2O2
ABTS, Mn(II)
b-O-4
NR
NR
Cu
O2
ABTS, DMP
b-O-4
Ca oxidation
Ethanosolv
Higher MW
[28]
Cu
O2
ABTS, DMP
b-O-4
NR
HP lignin
Aromatic monomer
[29]
Glutathione
b-O-4
NR
[33]
Glutathione
b-O-4
b-Ether cleavage b-Ether cleavage
Fluorescent lignin
[32]
Substrate DypB
Dyp2 Laccase CopA Etherase
Current Opinion in Chemical Biology 2015, 29:10–17
Polymeric lignin
Reaction Ca–Cb cleavage
Substrate
Products [17]
Kraft lignin Lignocellulose
Ref
Lignin dimer
[23] [25]
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Enzymatic conversion of lignin Bugg and Rahmanpour 15
Figure 3
CO2H
Hemicellulose
OCH3 OH
OH HO
OAr
G type lignin
CHO
CO2H
OCH3 OH
OH
X
OAr
H type lignin
CO2H
OCH3
protocatechuate 3,4-dioxygenase
OH
CO2H
O
OH
CO2H CO2H
OH
OH
OH catechol
H2/cat HO2C
O
CO2H
O triacylglycerol lipids
CO2H
cis,cismuconic acid
X OH
CO2H
CO2H CO2H CO2H
OH
OH
vanillin
CHO
HO
β-ketoadipic acid
protocatechuic acid
OCH3 OH
β-oxidation
TCA cycle
O O
O O
polyhydroxy butyrate
SCoA
C8H17 O O
malonyl CoA
O O n Current Opinion in Chemical Biology
Production of bioproducts from microbial degradation pathways, illustrating schematically the metabolic pathways responsible for accumulation of small molecule products vanillin or cis,cis-muconic acid via gene deletion (in blue), and the accumulation of triacylglycerol lipids or polyhydroxybutyrate via primary metabolism (in green).
Rhodococcus opacus DSM1069 and PD630 are known to accumulate triglyceride lipids from primary metabolism under nitrogen-limiting conditions, and have been shown to grow on minimal media containing ultrasonicated ethanol organosolv lignin as carbon source, generating triglyceride lipids at 4 mg/g us-EOL (or 20 mg/L) after a nine day fermentation [40]. P. putida is a well-known aromatic degrader that also possesses activity for lignin breakdown [16]. Linger et al. have shown that the ability of P. putida to accumulate polyhydroxyalkanoate (PHA) biopolyesters under nitrogen-limiting conditions can be harnessed to convert lignin from alkaline pre-treated liquor (APL) into PHAs in a 48 h fermentation [41]. Pathway engineering has also been used in P. putida to accumulate cis,cis-muconic acid from aromatic degradation, via blockage of the protocatechuate cleavage pathway, and re-routing via catechol cleavage [42]. Using p-coumaric acid as a carbon source, a yield of 13.5 g/L was obtained in a fed-batch bioreactor after 78 h, whereas using alkali-APL a yield of 0.7 g/L was obtained after 24 h [42]. Cis,cis-muconic acid can then be www.sciencedirect.com
converted via catalytic hydrogenation into adipic acid, thereby providing a renewable route to adipic acid from lignin [42]. As we improve our understanding of lignin degradation, it should be possible to use pathway engineering in suitable microbial hosts to generate other useful products from microbial lignin breakdown, hence it is interesting that replacement of the genes encoding the catechol intradiol pathway in P. putida by genes encoding a corresponding extradiol pathway leads to an increase in yield of pyruvate from aromatic substrates [43]. A selective hydroxamic acid inhibitor of protocatechuate 3,4-dioxygenase has also been developed, as an alternative chemical approach to manipulating bacterial lignin degradation [44].
Industrial applications for lignin There is interest in finding new applications for polymeric lignins, as well as depolymerised lignin breakdown products. Jin et al. have used lignin derived from corn bioethanol production to make phenol-formaldehyde adhesive [45], while Ramires et al. have used Organosolv lignin from sugarcane bagasse to prepare phenolic resins Current Opinion in Chemical Biology 2015, 29:10–17
16 Energy
[46]. In principle polymeric lignin could be a template for synthesis of new biomaterials. It has been shown that Kraft lignin can be modified by atom transfer radical polymerisation (ATRP), changing the thermostability and water solubility of the polymer [47]. Polyurethane foams can also be made from either Organosolv or Kraft lignin, with stronger foams being formed from Organosolv lignin [48]. There is interest in the conversion of lignin into carbon fibre, however, there are challenges in generating high quality carbon fibre, notably that highly purified lignin fractions are needed, due to the heterogeneity of most lignin preparations [49]. In a related application, lignin has also been converted into porous carbon, suitable for supercapacitor electrode materials [50]. The high carbon content of lignin makes it potentially interesting for conversion into nanomaterials, and Yiamsawas et al. have recently shown that lignin nanocontainers can be formed from water-soluble lignosulfonates in inverse micro-emulsions [51].
Acknowledgement The authors thank the University of Warwick for funding a Chancellor’s Scholarship to R.R.
References and recommended reading Papers of particular interest, published within the period of review, have been highlighted as: of special interest of outstanding interest
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