Polyamine metabolism, lignin degradation and potential applications in
industrial biotechnology
Luaine Bandounas
2011
Pseudomonas and beyond
Polyamine metabolism, lignin degradation and potential applications in
industrial biotechnology
Proefschrift
ter verkrijging van de graad van doctor
aan de Technische Universiteit Delft,
op gezag van de Rector Magnificus prof. ir.K.C.A.M. Luyben,
voorzitter van het College voor Promoties,
in het openbaar te verdedigen op dinsdag 8 november 2011 om 12:30 uur
door
Luaine BANDOUNAS
Master of Science
In
Molecular Cell Biology and Bioinformatics
Dit proefschrift is goedgekeurd door de promoter:
Prof. dr. J.H. de Winde
Copromotor
Dr.ir. H.J. Ruijssenaars
Samenstelling promotiecommissie:
Rector Magnificus Voorzitter
Prof. dr. J.H. de Winde
Technische Universiteit Delft, promotor
Dr. ir. H.J. Ruijssenaars
BIRD Engineering, copromotor
Prof. dr. I.W.C.E. Arends
Technische Universiteit Delft
Prof. dr. P. Verhaert
Technische Universiteit Delft
Prof. dr. W.R. Hagen
Technische Universiteit Delft
Prof. V. De Lorenzo
Centro Nacional de Biotecnologia
Dr. ir. J. Wery
Dyadic Nederland B.V.
Dr. ir. H.J. Ruijsenaars heeft als begeleider in belangrijke mate aan de
totstandkoming van het proefschrift bijgedragen.
This project is financially supported by TNO, the Netherlands Ministry of Economic
Affairs and the B-Basic partner organizations (
www.b-basic.nl
) through B-Basic, a
public-private NWO-ACTS programme (ACTS = Advanced Chemical Technologies
for Sustainability). This project was carried out within the research programme of
the Kluyver Centre for Genomics of Industrial Fermentation which is part of the
Netherlands Genomics Initiative / Netherlands Organization for Scientific
Research.
Cover image and layout: Luaine Bandounas
ISBN: 978-84-90370-09-1
Table of Contents
Chapter 1
General Introduction 9
Chapter 2 Redundancy in putrescine catabolism in solvent tolerant Pseudomonas putida S12. 32
Chapter 3 Isolation and characterization of novel bacterial strains exhibiting ligninolytic potential. 62
Chapter 4 Decolourization of the lignin-model dye, Azure B by a ligninolytic Bacillus sp. and initial identification of enzymes involved. 92
Chapter 5 Discussion 116 Supplemental data 132 Summary 153 Samenvatting 156 Curriculum Vitae 159 Publications 161 Acknowledgements 162
Chapter 1
General Introduction
Global demand for renewable resources
Due to a growing world population, an average higher life expectancy and an increase in global industrialization, the world’s fossil reserves consumption has steadily been escalating over the last century. The demand for energy is expected to increase over 50 % by 2025 mainly due to worldwide development [1]. Current energy consumption places a huge strain on the environment in terms of the effect of increasing carbon dioxide emissions on the Earth’s climate. The resulting depletion of fossil resources [2, 3] will not only affect the global energy demand, but also the production of chemicals and materials. This necessitates the development of technologies to replace fossil-based resources by renewable resources.
First generation biofuels and chemicals are produced from starches, sugars and vegetable oils which gives rise to ethical and environmental issues [4]. The use of food crops as renewable resource has caused much apprehension and controversy regarding the competition for arable land and increasing food prices [3]. Lignocellulosic biomass, on the other hand, is an extremely promising renewable alternative as it is widespread and readily available to most countries, as well as not threatening the global food or feed supply. Dedicated crops for the production of lignocellulosic biomass may also be grown in combination with food crops, or on non-arable land [4]. Recently it was estimated that in the US approximately 1.3 billion tons of biomass could be sustainably produced annually from forestry and agricultural sources [5]. A conservative estimate of the global biomass production average is approximately 10 dry tons ha-1 year-1, however certain smaller scale field trials have produced significantly more than this amount [1, 6].
Emerging biorefinery industries focus on utilizing biomass as a renewable substrate for biochemicals and bioenergy production [4]. By 2030, the European Union plans to replace one quarter of the EU’s transportation fuels with biofuels and similarly, the United States Department of Energy strives to replace approximately one third of the 2004 liquid fuel
demand with biofuels. In addition, the US government plans to substitute 25% of the industrially produced organic chemicals with biomass-derived alternatives [1, 5].
Fig. 1: Biotechnology vs. traditional fossil resource-based routes for energy, fuel and chemical production [7].
Industrial Biocatalysis
Chemical industries are directing their focus towards developing sustainable processes for the biological production of (fine) chemicals [8]. The main driver for pursuing the bio-based production of chemicals is the shortage of fossil resources. Biotechnology is a potentially important approach for biobased production processes [9] and provides added benefits of wide substrate ranges and high enantio- and regio-selectivity of enzymatic biocatalysts [10, 11]. Biotechnological processes include biocatalysis, biotransformation and fermentation technology. These approaches are usually combined to convert a substrate molecule into the desired end product in a limited number of reaction steps [8, 10].
Biocatalysis involves the use of enzymes or whole cells for the conversion of a substrate into a product of interest. The substrate may either be a (renewable) source of carbohydrates or other fermentable compounds (bioconversion), or a precursor molecule (biotransformation). The product is normally a metabolite or a dead-end product in the production strain [10]. Biocatalytic applications range from food and feed to the production of bulk or fine chemicals [11, 12]. The appropriate biocatalyst is often selected by intensive screening in which enzymes or whole cells are investigated for their abilities to catalyze a specific reaction. Whole cells are of particular interest for reactions requiring multiple reaction steps or specific co-factors. These can be regenerated by metabolically active cells, which is usually less expensive than exogenous addition of co-factors and less complicated than in-vitro regeneration [13]. High throughput screening, metabolic engineering and directed evolution techniques have made huge contributions to the development and implementation of biocatalytic systems [14].
Metabolic engineering strategies
Microorganisms are a vast source of novel and diverse enzymes and metabolic pathways, as well as valuable chemicals or metabolites. They possess a range of metabolic pathways, which serve to provide them with necessary metabolites. Usually, these metabolites are not produced in excess and their formation is strictly regulated [15]. Thus, the performance of the organism commonly has to be improved by metabolic engineering to optimize product formation, eliminate product degradation and enhance substrate import and product export [11].
Various strategies are available for the overproduction of a metabolite or product of interest by a microorganism. These include increasing the amount of precursor; adding, deleting or modifying regulatory genes or sequences; increasing the copy number of genes encoding enzymes involved in bottleneck reactions; and removing unnecessary or competing pathways or reactions [15]. Directed evolution can be utilized to introduce specific gene mutations which influence enzyme activity or functionality [16].
Much information regarding novel metabolic pathways, genes of unknown function and gene regulatory elements, can be obtained from the multitude of microbial genomes that have been completely sequenced. To investigate gene expression patterns and to identify novel metabolic pathways, comparative genomic analysis and microarray technology can be used [17]. These methods can offer information regarding the changes of the metabolic and regulatory networks in microbial cells under various conditions and their response to environmental changes [17].
Solvent tolerant Pseudomonas putida as a whole-cell biocatalyst
Pseudomonas putida is a soil bacterium capable of degrading a wide range of chemical compounds [18]. P. putida KT2440 is the best characterized strain of this species and is considered the workhorse of Pseudomonas research. Much knowledge and understanding has been acquired from its 6.18 Mbp genome sequence [18, 19]. This strain has also been used to study the effects of genome reconstruction. The investigation and alteration of metabolic pathways have lead to increased fluxes towards specific metabolites, resulting in increased product formation [18]. The metabolic diversity of P. putida has been exploited for the production of value-added compounds such as polyhydroxyalkanoates, epoxides, substituted catechols, alcohols and heterocyclic compounds [11, 18]. The catabolic versatility of P. putida KT2440 may be related to the high number of insertion elements present in this strain [18], as many of these elements are related to resistance or accessory functions which bacteria have acquired via gene rearrangements or horizontal gene transfer [20]. Many P. putida strains are able to utilize xenobiotic or toxic compounds which property has been exploited to eliminate environmental pollutants [21].
Our laboratory has genetically engineered P. putida strain S12 to produce a range of aromatic compounds such as cinnamic acid, coumarate, hydroxybenzoate and p-hydroxystyrene. This P. putida strain was specifically selected for the production of toxic hydrophobic chemicals in view of its ability to tolerate organic solvent-like compounds. Its solvent tolerance is based on several adaptation mechanisms: it can modify the composition of the inner and outer membrane, but it also disposes of a solvent efflux
pump that extrudes uncharged lipophilic compounds from the cell [22, 23]. The solvent-tolerance properties of P. putida S12 allow this strain to be cultured in a two-phase (water-solvent) system [24]. In such a two-phase system, very toxic products like p-hydroxystyrene can be extracted in-situ, resulting in reduced product inhibition and improved overall yields [25].
Engineered P. putida S12 produces the aromatic products from model renewables like sugar and glycerol, via the amino acids phenylalanine or tyrosine [26-28]. To enable the production of such aromatic compounds, extensive metabolic engineering was required, usually via a combination of rational, targeted approaches and classical strain improvement techniques [26-30]. Although P. putida S12 has a broad substrate specificity, it is unable to grow on the pentose sugars xylose and arabinose that commonly comprise up to 25 % of the total sugars present in lignocellulosic hydrolysate [31]. Therefore, pathways for the utilization of D-xylose were introduced and, surprisingly, in one case this also resulted in the efficient utilization of L-arabinose [31]. Thus, a P. putida S12 strain was obtained via metabolic engineering and laboratory evolution, which was able to utilize glucose, xylose and arabinose, the three main sugars present in lignocellulosic hydrolysate [31, 32]. Since the carbon substrate is not only utilized for product formation but also to generate reducing equivalents [33], a (cheap) co-substrate may be added to improve the overall yield. A promising auxiliary substrate which can be obtained from biomass (via syngas) is methanol [34]. Methanol is converted by dehydrogenases or oxidases to yield reducing equivalents, thus saving the primary substrate for product formation [35, 36]. A key oxidation intermediate of methanol, however, is the toxic compound formaldehyde, which must be efficiently metabolized to prevent accumulation. P. putida S12 was genetically engineered to efficiently metabolize formaldehyde to allow the use of methanol as an auxiliary substrate, which considerably increased the biomass yield on the primary substrate glucose [33].
Thus, P. putida S12 has been engineered at multiple levels to construct a useful platform organism for the production of chemicals from renewable, biobased feedstocks. Several of the improved P. putida S12 strains have furthermore been studied at the systems level in
order to identify the crucial changes underlying their improved performance. The analyses were based on the P. putida KT2440 genome sequence as a reference standard [19], in view of the high level of conservation of catabolic and biosynthetic pathways among different strains of P. putida [18, 37, 38]. A microarray-based genomotyping study furthermore demonstrated that P. putida S12 was genetically most similar to P. putida KT2440, directly after P. putida DSM3931 (which is a subculture of the mt-2 parent of strain KT2440): 86.9% of the genes with assigned functions in KT2440 were also identified in P. putida S12 [39]. Due to this high level of genetic conservation, KT2440-based microarrays could be used successfully for comparative transcriptome and proteome studies on P. putida S12 [18, 39-43].
Lignocellulosic biomass as sustainable feedstock for the production of bio-based chemicals and fuels.
The conversion of lignocellulosic biomass such as grasses, crop residues, sawdust, wood chips, pulp and paper waste has been highlighted as an important sustainable and renewable alternative means for energy, fuel and chemicals production [3, 44]. Lignocellulosic biomass is an abundant renewable resource; large amounts are available from the agricultural, forestry, food, pulp and paper industries, as well as in the form of municipal solid waste [2]. To date, biobased feedstocks are used in many applications such as plastics, solvents and lubricants. Current bio-based processes, however, usually rely on purified feedstocks e.g carbohydrates. Therefore purification and separation of components is extremely important, which presents a huge challenge especially for lignocellulosic feedstocks [1].
Recently, important progress has been made with respect to the utilization of the sugar fraction of lignocellulosic biomass for bio-based chemicals and fuels production. However, the potential for utilizing the recalcitrant lignin fraction as a renewable feedstock still needs to be explored. At present, lignin is mainly incinerated for heat and power generation, although in view of its structural richness it could also be suited as feedstock for the production of value-added chemicals such as substituted aromatics [1]. Far more
value could be gained from lignocellulosic biomass if the lignin fraction could be converted into products of interest by efficient, controlled depolymerization reactions. At present, only a small percentage (1 - 2 %) of lignin is isolated from spent pulping liquors and utilised in specialty products, amounting to 1 million tons per year worldwide [45-47]. The availability of efficient enzymes for controlled depolymerization of lignin could greatly increase the extent to which this compound can be utilized. The vast microbial diversity should therefore be studied extensively for ligninolytic capacities, in order to obtain an enzymatic toolkit for lignin valorisation.
Table 1: Content of cellulose, hemicellulose and lignin commonly found in agricultural waste and residues [48-51].
Lignocellulosic biomass Cellulose (%) Hemicellulose (%) Lignin (%)
Newspaper 40-55 25-40 18-30 Paper 85-99 0 0-15 Corn cobs 45 35 15 Nut shells 25-30 25-30 30-40 Grasses 25-40 35-50 10-30 Hardwood stems 40-55 24-40 18-25 Softwood stems 45-50 25-35 25-35 Wheat straw 30 50 15 Lignocellulose composition
Lignocellulosic materials such as wood, consist of three main components: cellulose, hemicellulose and lignin. These constituents occur in varying degrees depending on the source. Furthermore, small amounts of pectin, ash and proteins are present [2]. Cellulose comprises approximately 45 % of the dry weight of wood and consists of D-glucose subunits linked by β-1,4 glycosidic bonds [52]. Adjacent cellulose chains interact via hydrogen bonds, Van der Waal’s forces and hydrophobic interactions to form crystalline microfibrils, which are covered by hemicellulose and lignin (Fig. 3) [2]. The second most abundant component of lignocellulose, comprising approximately 25 – 30 % of the dry wood weight, is hemicellulose [52, 53]. This carbohydrate polymer comprises various
polysaccharides consisting of numerous monomers linked by β-1,4- and sometimes β -1,3-glycosidic bonds [52].
Fig. 2: General overview of renewable lignocellulosic biomass pretreatment and conversion to fuels and chemicals.
Lignin is a recalcitrant aromatic polymer comprising 10 – 25 % of lignocellulosic biomass [56]. It consists mainly of three hydroxycinnamyl-derived alcohol monomers: p-coumaryl (p-hydroxyphenyl alcohol), coniferyl (guaiacyl propanol) and sinapyl (syringyl) alcohols
(Fig. 4), which differ in degree of methoxylation [57]. Approximately half of the lignin structure is composed of β-Ο-4-linked ethers (arylglycerol-β-aryl), followed by phenylcoumarans, resinols and other minor subunits [58]. Since the aryl ethers are more difficult to oxidize than the 10% phenolic content of lignin [58], degradation of the β-Ο-4 substructure is considered a crucial step in lignin degradation [59]. Besides the β-aryl ether (β-O-4) linkage, which can relatively easily be cleaved chemically, more chemically resistant C-C bond linkages are present, such as β-1, β-5, β-β, 5-5 and 5-O-4 (Fig. 5) [57, 60].
Fig. 3: A simplified representation of a plant macrofibril consisting of bundles of cellulose microfibrils encased in hemicellulose and lignin [54, 55].
Hardwood lignin mainly consists of
guaiacyl and syringyl subunits, while softwood, which usually has a higher lignin content, predominantly contains guaiacyl subunits with more resistant linkages (β-5, 5-5 and 5-O-4) due to the available C5 position [52, 61]. Lignin serves to reinforce plant stems offering stability and strength, make the plant’s vascular tissue waterproof to allow water transport, and provide protection against pathogens [61, 62].
Pretreatment of lignocellulosic biomass
The major challenge for efficient utilization of biomass for the production of biofuels or value-added chemicals is its conversion to fermentable sugars. The initial steps usually involve the destruction of various interactions between lignin and hemicellulose to make the cellulose fibrils accessible to hydrolysing chemicals or enzymes [64]. This is generally accomplished by physico-chemical pretreatment, most commonly heating under acidic conditions [56, 65]. Cellulose and hemicellulose are then depolymerized by chemical or enzymatic hydrolysis to monomeric sugars, which can be fermented to ethanol or other products [56]. Table 2 presents an overview of various lignocellulosic pretreatment methods, as well as their effect on lignocellulose.
One of the problems arising from some of the various lignocellulosic pretreatment methods is the release or formation of (toxic) inhibitors such as furfural, 5-hydroxymethylfurfural, aromatics and organic acids which may affect the overall microbial fermentation yield [65]. The production of such toxic by-products could be prevented by replacing the thermochemical pretreatment with a biological pretreatment [52], e.g., by incubating lignocellulosic biomass with white-rot fungi. Such a pretreatment has been reported to be comparable to an alkaline pretreatment in efficiency, allowing enzymatic hydrolysis of both hemicellulose and cellulose to fermentable sugars [44], to an even higher final glucose concentration [66]. Biological pretreatment of lignocellulose requires less energy input and milder conditions, but the rate of hydrolysis is usually quite low [3].
Table 2: Examples of lignocellulose pretreatment methods
Biomass pretreatment
method Result Reference
Physico-chemical pretreatment methods
Steam explosion (high temperature and rapidly
reduced pressure)
Degrades hemicellulose and lignin [3, 68]
Ammonia fibre expansion (AFEX)
Hemicellulose remains untouched, but cells walls more receptive to enzyme
hydrolysis
[5]
CO2 explosion Effective for pretreatment of cellulose [69]
Thermochemical pretreatment (H2SO4 at
140-200°C)
Makes cellulose in cell walls more
accessible to saccharifying enzymes [5]
(Bio-)Chemical pretreatment methods
Acid pretreatment (Sulfuric, nitric or hydrochloric acid)
Removes hemicellulosic components,
exposing cellulose for enzymatic digestion [70]
Alkali pretreatment Removes lignin and uronic acid
substitutions on hemicellulose [71]
Peroxide pretreatment Oxidative delignification and reduction of
cellulose crystallinity [72] Ozone pretreatment Reduces lignin content [70]
Organosolv process Breaks internal lignin and hemicellulose
bonds [3, 73]
Biological treatment with white-rot fungi or ligninolytic microorganisms
Delignification liberates cellulose and
hemicellulose from lignin [52]
Physical pretreatment methods
Mechanical treatment (chipping, grinding, milling)
Reduces cellulose crystallinity [3, 74]
Pyrolysis, also thermochemical
pretreatment
Lignin biodegradation
The key challenge for the bioconversion of lignin into high-value products usually lies in the recalcitrant and complex nature of the lignin polymer. However, specialized organisms have evolved that are capable of depolymerizing lignin, such as white rot and soft rot fungi. These micoorganisms attack lignin and cellulose, while brown rot fungi specifically attack cellulose [3]. Lignin degradation by white-rot fungi involves numerous, mostly oxidative reactions. In addition, demethoxylation or demethylation reactions take place, as well as propyl side-chain cleavage between Cα and Cβ [75]. Brown rot fungi can degrade cellulose and hemicellulose, and at advanced stages of decay, a slightly modified lignin residue is left behind [76]. To date, white rot basidiomycetes are the most efficient lignin degraders due to their extracellular oxidative enzymes, collectively called ligninases [2, 61]. Usually, these ligninolytic enzymes cannot penetrate the intact wood structure. Therefore, mediator molecules such as activated oxygen species are often required to transfer the oxidizing potential of the enzyme to the lignin substrate. These, usually small, molecules can diffuse into the structure and initiate the degradation process [77]. Generally, lignin is depolymerized by the combined efforts of extracellular laccases, lignin peroxidases (LiP’s), manganese peroxidases (MnP’s), secondary secreted metabolites and reactive oxygen species [78]. The oxidoreductase enzymes transfer electrons from a substrate to an electron acceptor, which is hydrogen peroxide in the case of peroxidases and oxygen for laccases and tyrosinases [79]. Due to the large, highly complex and diverse structure of lignin, microbial depolymerization generally occurs extracellularly, after which the smaller resulting monomers are mineralized intracellularly [5, 78].
Laccases oxidize complex substrates such as lignin, which makes them particularly interesting for diverse industrial applications [80, 81]. Laccases belong to the multi-copper oxidase family [55, 79] and have been widely studied in fungi, although various bacterial laccases have also been described. Several potential bacterial laccases have been identified in Bacillus species, such as Bacillus halodurans [82], Bacillus sphaericus and Bacillus subtilis, either or not associated with spores [79]. Similarly, laccase-like enzymes were identified in E. coli (yacK), Pseudomonas putida (cumA), Streptomyces griseus (epoA)
and Xanthomonas campestris (copA) [79, 83]. Laccases are found both intracellularly, as well as extracellularly as spore coat proteins. Although their functions are quite different in various organisms, they commonly serve to catalyze either polymerization or depolymerization reactions [81].
Peroxidases are peroxide-dependent oxidative enzymes, capable of oxidizing many organic and inorganic compounds. Prokaryotic peroxidases are usually located intracellularly, while fungal peroxidases are extracellular [84]. Peroxidases can react with manganese (Mn2+) ions, methoxylated aromatics and phenolic compounds [85]. Lignin peroxidase (LiP) can directly oxidize lignin, 1,2,4,5-tetramethoxybenzene, phenolic compounds, anilines and certain non-phenolic structures. Although manganese peroxidases (MnP) are also strong oxidizing enzymes, they are not able to directly oxidize non-phenolic lignin-related structures [58]. MnP transfers its oxidizing potential to Mn3+ which diffuses into the cell wall and attacks lignin [58]. The main difference between LiP’s and MnPs is that LiP’s are not very efficient at oxidizing the non-phenolic lignin compounds, probably due to their inability to penetrate the pores in lignocellulose. MnPs produce stronger oxidizing agents which do penetrate the substrate, but the overall yields are quite low [58]. A versatile peroxidase (VP) has been described in certain fungi, which is able to oxidize phenolic compounds by a combination of catalytic activities similar to Lip, MnP and plant or microbial peroxidases [61, 86]. Glyoxal oxidases, aryl-alcohol oxidases, aryl-alcohol dehydrogenases and quinone reductases have also been reported to be involved in lignin degradation [61]. Fenton-based OH radical-producing reactions could also be involved in the degradation of phenolic and non-phenolic lignin related compounds [87].
In addition to fungi, certain soil bacteria such as Nocardia and Rhodococcus, have been reported to degrade lignin [88]. Bacterial enzymes that potentially play a role in lignin degradation could be laccases, monooxygenases, multiple ring-cleaving dioxygenases and phenol oxidases [89]. The Gram-negative Sphingobium sp. SYK-6 (formerly known as Sphingomonas paucimobilis SYK-6) has the ability to degrade various lignin monoaryls and biaryls, such as vanillin, vanillate, syringaldehyde, syringate, phenylcoumarane and diarylpropane arising from lignin degradation [90, 91]. Sphingobium sp. SYK-6 is also able
to cleave the most abundant intermolecular linkage in lignin, the β-aryl ether linkage [90, 91]. The biphenyl component of lignin, which comprises approximately 10% of the structure depending on the source of lignin, can be degraded by Pseudomonas, Sphingomonas, Burkholderia, Rhodococcus, Bacillus, Ralstonia, Acinetobacter, Comamonas and Achromobacter genera [92], as well as Pandoraea pnomenusa B-356 [93].
With regard to the potential application of ligninolytic enzymes for lignin valorization, bacterial enzymes may not need the (expensive) mediators commonly required by fungal depolymerizing enzymes in industrial applications. Furthermore, many of the fungal ligninolytic enzymes such as LiPs, MNPs and VPs contain a heme prosthetic group [94]. In an industrial environment, it may not be feasible to utilize these enzymes, as the prosthetic group may be difficult to incorporate or they may require complicated reactivation procedures [95]. Thus, the identification of enzymes involved in bacterial lignin degradation would possibly provide important alternative systems for lignin conversion to products of interest. Recent insight suggests that bacterial enzymes may be more specific, exhibit higher thermostability and may be more suited for alkaline pH conditions than those present in fungi [56, 82, 96]. Ligninolytic bacteria or their enzymes may be exploited in industrial applications for the controlled depolymerization of waste lignin, instead of it being incinerated for heat.
Bacterial metabolism of lignin-related and lignin-derived compounds
Many bacteria may not be able to degrade lignin, but some are capable of utilizing lignin-derived compounds released by other lignin-degraders, such as fungi. Bacteria employ O-demethylation systems in the utilization of lignin-derived compounds, such as vanillin demethylase, syringate demethylase and tetrahydrofolate-dependent aromatic O-demethylase, in conjunction with C1-metabolism [96]. Pseudomonads are not known as very efficient lignin degraders, but some are able to efficiently degrade several low molecular weight lignin model compounds, such as anisoin, benzoin, biphenyls and chlorobiphenyls [97]. An important catabolic route for lignin-related aromatic compounds is the β-ketoadipate pathway which has been identified in various bacterial genera, such
as Bacillus, Pseudomonas, Rhodococcus, Streptomyces and Alcaligenes [98]. β-Ketoadipate pathway enzymes convert lignin-derived aromatic compounds to protocatechuate or catechol. These are then converted to β-ketoadipate, followed by a two-step conversion to tricarboxylic acid intermediates [99, 100]. The decarboxylation of lignin-related aromatic compounds like ferulic acid and p-coumaric acid has been described for several microorganisms, including Bacillus species [101].
Many ligninolytic organisms are also able to utilize or degrade lignin model compounds such as lignin mimicking dyes. Synthetic industrial dyes can be applied as effective lignin model compounds to screen for the presence of ligninolytic enzymes, as certain enzymes active towards lignin are also capable of decolourizing these recalcitrant dyes [102]. Several Bacillus sp. isolates have been reported to possess ligninolytic and industrial dye degrading capabilities [89, 103-106]. It appears that certain bacteria have various enzyme systems to deal with lignin-derived compounds, although extensive research is required to identify and characterize these systems.
Scope and outline of this thesis
This research was carried out within the framework of the B-Basic (Bio-Based Sustainable Industrial Chemistry) programme and focused on the genetic engineering of microorganisms for the production of bio-based chemicals from renewable resources, as well as the identification of novel microorganisms and enzymes involved in lignin degradation for eventual application in lignin valorisation.
Chapter 1 provides a general overview of the use of microorganisms, especially the solvent-tolerant Pseudomonas putida S12 for the production of chemicals from renewable resources. The use of lignocellulose as a sustainable alternative to fossil resources is discussed, including the composition and pretreatment of lignocellulose. An overview is presented of enzyme systems and microorganisms reported to be involved in the degradation or utilization of lignin.
Chapter 2 discusses the polyamine metabolism of P. putida S12. Of particular interest is the polyamine putrescine, which has been associated with general stress conditions and may play a role in the solvent stress response of P. putida S12. The metabolic pathway of putrescine was unknown in P. putida S12; and it was therefore investigated in order to gain a better understanding of the role of polyamines in (solvent) stress.
Although Pseudomonas putida S12 has a broad substrate affinity, it cannot degrade or utilize lignin efficiently. Therefore, it was necessary to investigate other bacteria as potential sources of novel ligninolytic enzymes for waste lignin valorisation. Chapter 3 describes the isolation and characterization of three soil isolates enriched on Kraft lignin. The ligninolytic potential of Pandoraea norimbergensis LD001, Pseudomonas sp. LD002 and Bacillus sp. LD003 was evaluated by investigating their ability to degrade lignin model dyes and to grow on Kraft lignin or aromatic monomers as sole carbon source.
The Bacillus sp. LD003 that demonstrated ligninolytic potential, as described in Chapter 3, also displayed a particular affinity for decolourizing the dye Azure B, which is frequently
used to assess lignin-degrading activity. Bacillus sp. LD003 was therefore considered to be a promising source of novel ligninolytic enzymes. Chapter 4 describes the isolation and identification of enzyme(s) involved in Azure B decolourization in Bacillus sp. LD003.
Chapter 5 provides a summary of the findings described in Chapters 2 – 4. The importance of regulating and maintaining polyamine homeostasis in the solvent-tolerant Pseudomonas putida S12 is discussed, as well as the challenges involved in the screening of ligninolytic microorganisms and the bioconversion of lignin into products of interest.
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Chapter 2
Redundancy in putrescine catabolism in solvent tolerant Pseudomonas
putida S12.
This chapter was published as:
Bandounas L, Ballerstedt H, de Winde JH, Ruijssenaars HJ. Redundancy in putrescine catabolism in solvent tolerant Pseudomonas putida S12. Journal of Biotechnology, 2011, 154 (1):1-10
.
Abstract
Pseudomonas putida S12 is a promising platform organism for the biological production of substituted aromatic compounds due to its extreme tolerance towards toxic chemicals. Solvent or aromatic stress tolerance may be due to membrane modifications and efflux pumps; however in general, polyamines have also been implicated in stressed cells. Previous transcriptomics results of P. putida strains producing an aromatic compound, or being exposed to the solvent toluene, indicated differentially expressed genes involved in polyamine transport and metabolism. Therefore, the metabolism of the polyamine, putrescine was investigated in P. putida S12, as no putrescine degradation pathways have been described for this strain. Via transcriptome analysis various, often redundant, putrescine-induced genes were identified as being potentially involved in putrescine catabolism via oxidative deamination and transamination. A series of knockout mutants were constructed in which up to six of these genes were sequentially deleted, and although putrescine degradation was affected in some of these mutants, complete elimination of putrescine degradation in P. putida S12 was not achieved. Evidence was found for the presence of an alternative pathway for putrescine degradation involving γ-glutamylation. The occurrence of multiple putrescine degradation routes in the solvent-tolerant P. putida S12 is indicative of the importance of controlling polyamine homeostasis, as well as of the high metabolic flexibility exhibited by this microorganism.
Introduction
Pseudomonas putida S12 displays exceptional tolerance towards a range of toxic organic solvents at concentrations which are usually lethal to other microorganisms [1-3]. Solvent-tolerant P. putida strains dispose of special mechanisms to cope with hydrophobic toxic compounds, which include modifications to the inner and outer membrane, as well as active extrusion of solvents through membrane-associated efflux pumps [4, 5]. Due to the robust nature and extreme tolerance towards various types of chemicals, including aromatic compounds [6-8], P. putida S12 was considered a suitable host for high-level aromatics production. For this reason, P. putida S12 was engineered for the production of substituted aromatic compounds such as phenol, t-cinnamate, coumarate, p-hydroxybenzoate and p-hydroxystyrene from renewable feedstock [9-14].
Polyamines like putrescine are found in many organisms [15], acting as signalling and regulatory molecules in response to stress conditions caused by reactive oxygen species (ROS), heat, UV, acid and osmotic pressure [16, 17]. They enhance the expression of genes encoding transcription factors such as Cya, FecI, Fis [18], as well as RpoS which has been implicated in stress tolerance of Pseudomonas fluorescens [19]. Polyamines also play a critical role in outer membrane function, affecting both the production and the functionality of outer membrane porins such as OmpF and OmpC in E. coli. By blocking such porins, polyamines decrease membrane permeability and protect the cell, e.g., against acidic stress [20, 21].
The membrane damage brought about by solvent exposure is typically associated with the formation of ROS [22-24]. In view of the involvement of polyamines in oxidative stress and outer membrane functioning, these compounds may be expected to contribute to solvent tolerance, in addition to established mechanisms such as efflux pumps and membrane modification systems. Recently, novel indications for such a role of polyamines, particularly putrescine, were found in a transcriptome analysis of a p-hydroxybenzoate producing P. putida S12 strain. This study revealed a number of genes presumably involved in putrescine transport and metabolism that were differentially expressed in this
strain compared to a non-producing control strain [25]. Also in other studies, polyamine-associated genes were found to be differentially expressed in solvent-exposed P. putida strains [3, 26].
In view of the apparent relationship between putrescine and solvent stress response, information on putrescine metabolism in P. putida S12 is of importance to gain a better understanding and control of the solvent tolerance mechanisms. Although several pathways have been described in literature [15, 16, 27, 28], it is unknown which of these are present or relevant in P. putida S12. The present study describes genes and pathways involved in putrescine degradation by P. putida S12, identified via transcriptome analysis.
Materials and Methods
Media and cultivation
The plasmids and strains used are listed in Table 1a and 1b. The growth media used were Luria broth (LB) [29] or phosphate buffered mineral salts medium (MM) [30]. Unless otherwise stated, 20 mM glucose was used as the carbon source in the mineral salts media (MMG). If putrescine was added as nitrogen source, (NH4)2SO4 was omitted from MMG. Cultures were grown in 20 ml liquid medium in 100-ml Erlenmeyer shake flasks at 180 rpm in a rotary shaker. Antibiotics were added to solid or liquid media as required in the following concentrations: for E. coli 10 μg/L tetracycline or 10 μg/L gentamicin; for P. putida S12 30 μg/L tetracycline (Tc) or 25 μg/L gentamicin (Gm). P. putida was cultivated at 30 °C and E. coli at 37 °C. Cultures were inoculated with cells from an overnight culture, to a starting OD600 of 0,05 - 0,1.
DNA techniques
Genomic DNA was isolated using the FastDNA kit (Q-Biogene) in combination with a FastPrep FP120 Homogenizer (Thermo Scientific). The QIAprep spin miniprep kit (QIAGEN) was used to isolate plasmid DNA. A Gene Pulser electroporation device (BioRad) was used
to introduce plasmid DNA into electrocompetent cells. DNA concentrations were determined using a ND-1000 spectrophotometer (Nanodrop). The QIAEXII gel extraction kit (QIAGEN) was used to isolate agarose-trapped DNA fragments. Polymerase Chain Reactions (PCRs) were performed using Accuprime Pfx polymerase (Invitrogen) as specified by the manufacturer. Enzymes from Fermentas GmbH were used for DNA digestions and ligations according to the manufacturer’s instructions. Oligonucleotide synthesis and DNA sequence analysis were performed by Eurofins MWG Operon. Nucleotide sequences were analysed using the Basic Local Alignment Search Tool (BLAST) [31].
Table 1a: Plasmids used in this study.
Plasmids Characteristics Reference
pJQ200SK P15A ori sacB RP4 Gmr (pBluescriptSK); suicide vector [34]
pJNNcre(t)
pJN under control of nagR-pNagAa, for expression of the Cre-recombinase used in the Cre-loxP site specific recombination system [35]
Unpublished
pGEM-T Easy Apr, used for cloning PCR fragments Promega
pJQ∆hpd-lox-tetA Used as source for the tetA- marker flanked by loxP
sites (tetA-lox) [36]
pJQspuC-lox-tetA pJQ200SK containing aminotransferase (PP5182)
interrupted by tetA-lox This study
pJQaptA-lox-tetA pJQ200SK containing β-alanine pyruvate
aminotransferase (PP0596) interrupted by tetA-lox This study
pJQox1-lox-tetA pJQ200SK containing oxidoreductase (PP3146)
interrupted by tetA-lox This study
pJQgabT-lox-tetA pJQ200SK containing 4-aminobutyrate
aminotransferase (PP0214) interrupted by tetA-lox This study
pJQox2-lox-tetA pJQ200SK containing oxidoreductase (PP4548)
interrupted by tetA-lox This study
pJQpuuA-lox-tetA pJQ200SK containing glutamate—putrescine ligase
Table 1b: Strains used in this study.
Bacterial strain Alternate
name*
Properties # Reference
P. putida S12 - Wild-type (ATCC 700801) [30]
P. putida S12∆spuC KO-1 Gene encoding class III aminotransferase (PP5182) deleted This study
P. putida S12∆aptA - Gene encoding β-alanine pyruvate aminotransferase (PP0596)
deleted
This study
P. putida S12∆gabT - Gene encoding putative 4-aminobutyrate aminotransferase
(PP0214) deleted
This study
P. putida S12∆ox1 - Gene encoding oxidoreductase (PP3146) deleted This study
P. putida S12∆spuC∆gabT KO-2 Gene encoding putative 4-aminobutyrate aminotransferase
(PP0214) deleted in S12∆spuC
This study
P. putida S12∆spuC∆aptA - Gene encoding β-alanine pyruvate aminotransferase (PP0596)
deleted in S12∆spuC
This study
P. putida S12∆ox1∆ox2 - Gene encoding oxidoreductase (PP4548) deleted in S12∆ox1 This study
P. putida S12∆spuC∆gabT∆aptA KO-3 Gene encoding β-alanine pyruvate aminotransferase (PP0596)
deleted in S12∆spuC∆gabT
This study
P. putida S12∆spuC∆gabT∆aptA∆ox1 KO-4 Gene encoding oxidoreductase (PP3146) deleted in
S12∆spuC∆gabT∆aptA
This study P. putida
S12∆spuC∆gabT∆aptA∆ox1∆ox2
KO-5 Gene encoding oxidoreductase (PP4548) deleted in
S12∆spuC∆gabT∆aptA∆ox1
This study P. putida
S12∆spuC∆gabT∆aptA∆ox1∆ox2∆puuA
KO-6 Gene encoding glutamate—putrescine ligase (PP5299) deleted in
S12∆spuC∆gabT∆aptA∆ox1∆ox2
This study
E. coli DH5α - General cloning strain Invitrogen
*
Simplified strain name used throughout text. # PP locus tags taken from the P. putida KT2440 genome database [32, 33] were used to denote
Construction of knockout mutants
The suicide vector pJQ200SK [34] was used as the backbone for constructing knockout plasmids for targeted gene disruption [37]. The target genes were amplified in two fragments, with a minimum size of 600 basepairs (bp) by polymerase chain reaction (PCR) using the primers listed in Table 2. The primers were designed such that 50 -100 bp were omitted from the centre of the gene. The suicide vector and the gene fragments were digested with the appropriate restriction enzymes, ligated and transformed into E. coli. The plasmids were isolated and linearized at the unique restriction site between the cloned target gene fragments. Subsequently, a tetracycline resistance gene (tetA) flanked by 2 loxP sites, isolated from pJQ∆hpd-lox-tetA [36], was ligated into the linearized plasmid after treatment with bacterial alkaline phosphatase (BAP, Invitrogen).
After transformation to E. coli, the knockout plasmids were isolated, verified by restriction analysis and transformed to P. putida S12 for targeted gene disruption as previously described [36, 38]. P. putida S12 disruption mutants were plated on LB + Tc agar plates. For double cross-over recombinant selection, P. putida S12 transformants were scored for tetracycline resistant (TcR) and gentamicin sensitive (GmS) phenotype. Successful replacement of the original gene with the tet-loxP disrupted copy was confirmed by PCR and/or sequence analysis. The tetA marker was cured from selected double cross-over knockout mutants to enable a subsequent round of targeted gene disruption. For this purpose, plasmid pJNNcre(t) (Table 1a) was transformed to selected knockout mutants which were selected on LB + Gm agar plates. These GmR colonies were incubated in LB + Gm until growth was visible, after which 0,1 mM salicylic acid was added for Cre induction. After overnight incubation, dilutions were plated onto LB agar and the following day, loss of the Tc antibiotic marker was screened for on LB agar and LB + Tc agar plates. To cure the tetracycline sensitive (TcS) colonies from pJNNcre(t), these were grown overnight in liquid LB medium. Loss of pJNNcre(t) was confirmed by screening for gentamicin sensitive (GmS) phenotype and loss of the tetA marker was confirmed by PCR.
Microarray analysis
Three 1-L Erlenmeyer shake flasks containing 150 ml MM + glucose (10 mM), were inoculated with P. putida S12 from an overnight culture to a starting OD600 of 0.06. At mid-exponential phase, 1 mM putrescine (Putrescine dihydrochloride, Sigma-Aldrich) was added to each flask. Samples (1 ml) were drawn after 0, 15 and 30 minutes of putrescine addition. Immediately after sampling, the samples were quenched and handled as previously described [36]. The cDNA samples were analysed on high-density, custom-made P. putida KT2440-genome based microarrays (Affymetrix) with additional probe sets of known sequences of P. putida S12 and related strains [39, 40]. The arrays were hybridized and scanned according to modified manufacturer’s protocols [36, 41, 42]. GeneSpring GX Software (version 7.3.1) and the GC RMA algorithm was used for data analysis [42]. After normalization, a 1-way ANOVA (P value cut-off of 0.05) was used to select genes which changed significantly in conditions t = 15 min vs. t = 0, t = 30 min vs. t = 0 and t = 30 min vs. t = 15 min. Genes which were more than 2-fold up-regulated were considered for further analysis.
Analytical methods
Putrescine was analysed by ion-exchange chromatography (DIONEX ICS-3000). A CSRS ultra II suppressor was used in combination with an IonPac CS17 column (2 x 250 mm) and an IonPac CG17 (2 x 50 mm) guard column all at 30 ºC. Methane sulphonic acid (MSA, 10 mM) was used as the eluent, at a flow rate of 0,4 ml/min. The cell density was measured at 600 nm (OD600) with a µQuant MQX200 universal microplate spectrophotometer (Bio-tek), using flat-bottom 96-well microplates (Greiner). Cell dry weight (CDW) was calculated from the OD600 value, assuming that an OD600 of 1 is equivalent to 0,47 g/L CDW [43].
Table 2: Primers used for PCR amplification of genes targeted for disruption.
Primer Sequence (5’ → 3’) # Details
MW1 5’-gcgggatccttattgaatcgcctcaagggtcag-3’ Upstream spuC1 (PP5182), BamHI MW2 5’-gcgtctagaaaggtgaaggagatcctcgcc-3’ Downstream spuC1 (PP5182), XbaI MW3 5’-gcgtctagatcggcgatgaaggccgcgac-3’ Upstream spuC2 (PP5182), XbaI MW4 5’-gcggcggccgcatgagcgtcaacaacccgcaaacc-3’ Downstream spuC2 (PP5182), NotI MW5 5’-gcgggatccgatgtagtccgaccagttatag-3’ Upstream spuC1(PP5182) , BamHI MW6 5’-gcggcggccgcgccgccagcctactgtgtgg-3’ Downstream spuC2 (PP5182) , NotI LB09 5’-gcgggatcctgagaatgcccttcgg-3’ Upstream aptA1 (PP0596), BamHI LB10 5’-gcgtctagaccgaagggttacctgaagcg-3’ Downstream aptA1 (PP0596), XbaI LB11 5’-gcgtctagaagcgcgataccgccc-3’ Upstream aptA2 (PP0596), XbaI LB12 5’-gcggcggccgccgtctatctgctgg-3’ Downstream aptA2 (PP0596), NotI LBA13 5’-gcgggatccgcccagaagctggccgctgc-3’ Upstream gabT1 (PP0214), BamHI LBA14 5’-gcgtctagatcggcgtcgttcttgaaaatgcgc-3’ Downstream gabT1 (PP0214), XbaI LBA15 5’-gcgtctagagaccagcacggcatcctgc-3’ Upstream gabT2 (PP0214), XbaI LBA16-2 5’-gcggcggccgcatcagcaccgccgtgacttgc-3’ Downstream gabT2 (PP0214), NotI LB17 5’-gcgggatccccgacatccacatcctggt-3’ Upstream ox1-1 (PP3146), BamHI LB18 5’-gcgtctagagcagctatgtggtggcca-3’ Downstream ox1-1 (PP3146), XbaI LB19 5’-gcgtctagagtgcagcgtcaacc-3’ Upstream ox1-2 (PP3146), XbaI LB20 5’-gcggcgccgcgaccgtatcgactcggg-3’ Downstream ox1-2 (PP3146), NotI LBA33 5’-gcgggatccggccggcaccggcatgc-3’ Upstream ox2-1 (PP4548), BamHI LB34 5’-gcgtctagatgaggtcgcaatcgatgccatag-3’ Downstream ox2-1 (PP4548), XbaI LB27 5’-gcgtctagacagcgccgcggcatgggc-3’ Upstream ox2-2 (PP4548), XbaI LB36 5’-gcggcggccgcgagacgcggccctgggattt-3’ Downstream ox2-2 (PP4548), NotI MW67 5'-gcgtcagcgcagcttctcgagca-3' Downstream ox2 gene (PP4548) LB132F 5’-gcgactagtacatgctgctgcactggtag -3’ Upstream puuA1 (PP5299), BcuI LB132R 5’-gcgtctagaaggtacacctgggtcgactg- 3’ Downstream puuA1 (PP5299), XbaI LB133F 5’-gcgtctagagtcgcctacgaccatgaaat- 3’ Upstream puuA2 (PP5299), XbaI LB133R 5’-gcggcggccgcatgcggctcgatatttgaag- 3’ Downstream puuA2 (PP5299), NotI
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