Acetone activation by the strictly anaerobic bacterium Desulfococcus biacutus
Dissertation submitted for the degree of Doctor of Natural Sciences
Presented by Olga Brígida Gutiérrez Acosta
at the University of Konstanz
Faculty of Sciences
Department of Biology
Date of the oral examination: 06.december.2013 First supervisor: Prof. Dr. Bernhard Schink Second supervisor: Prof. Dr. Jörg Hartig Third supervisor: Prof. Dr. Alasdair Cook
ACKNOWLEDGEMENTS
I express my sincere thanks to Prof. Bernhard Schink for supervising me during my doctoral research work. Thanks for giving me the opportunity to develop my doctoral thesis in your working group, for guiding me, for sharing ideas, and for all discussions.
I thank the members of my thesis committee Prof. Alasdair Cook, Prof. Jörg Hartig and Prof. Bernhard Schink, for their participation in my evaluations as a member of the graduate school Konstanz Research School Chemical Biology. Thank you for the pleasant discussions and valuable comments.
I also thank Prof. Peter Kroneck for the valuable discussions about my work, and for the personal support. I am also thankful to Dr. David Schleheck, Dr. Felix Ten Brink and Dr. Diliana Simeonova for their help and discussion.
Thanks to all members and participants of the DFG priority program 1319, for recognizing the scientific advance made on this theme during my doctoral research. I especially thank Prof. Bernard T. Golding for the valuable discussions about my work.
I am grateful to all members of the AG Schink for the help and collaborations during my doctoral research and for the coffee and cake time.
My warm thanks to all my friends for the great moments during my stay in Konstanz, with special thanks to Natalia Charlina and to Jennifer Ignatious for the sincere and lovely friendship.
My most profound thank to my parents, brothers, and all the new members of the Gutierrez Acosta family for their support in this personal achievement. Immensely thanks to my husband Norman Hardt for his encouragement, professional support and companionship.
AGRADECIMIENTOS
Por la realización de este trabajo de tesis doctoral, quiero expresar mis más sinceros agradecimientos a mi supervisor de tesis, el Prof. Bernhard Schink. Gracias por darme la oportunidad de desarrollar mi tesis en su grupo de trabajo y por guiarme acertadamente durante la realización de la misma.
Agradezco a los miembros de mi comité de tesis Prof. Alasdair Cook, Prof. Jörg Hartig y Prof. Bernhard Schink, por haber participado como tales durante mis evaluaciones como miembro de la escuela de graduados Konstanz Research School Chemical Biology. Gracias por las gratas discusiones y sugerencias que me ayudaron a tener un mejor desempeño en mi trabajo.
Quiero agradecer también al Prof. Peter Kroneck por las discusiones sobre mi trabajo, y por el apoyo personal. Así mismo agradezco al Dr. David Schleheck, Dr. Felix Ten Brink y Dr. Diliana Simeonova por sus sugerencias y apoyo en la realización de mi trabajo.
Gracias a los miembros y participantes del programa científico alemán de prioridad DFG 1319 por reconocer el avance científico en el tema, logrado durante mi tesis doctoral. Agradezco especialmente al Prof. Bernard T. Golding por sus apreciadas discusiones sobre mi trabajo.
A todos los miembros del grupo de trabajo del Prof. Schink, por su apoyo y colaboración durante mi estancia doctoral y por todos los momentos de café.
Gracias a todos mi amigos que me brindaron gratos momentos durante mi estancia en Konstanz. En especial muchas gracias a Natalia Charlina y Jennifer Ignatious por brindarme una amistad sincera.
Mis más profundos agradecimientos van hacia mis padres, mis hermanos y todos los nuevos integrantes de la familia Gutiérrez Acosta por apoyarme en este logro más en mi vida. Agradezco inmensamente a mi esposo Norman Hardt por el apoyo brindado tanto personal como profesional y por su compañía.
I
Table of Contents
CHAPTER 1 General Introduction……………...... …………...………………1 1.1 Important aspects about acetone…………………………………….……1 1.2 Acetone degradation by aerobic bacteria………………………………….1 1.3 Acetone degradation by nitrate-reducing bacteria…………………………2 1.4 Acetone degradation by sulfate-reducing bacteria (SRB)…………………..3 1.5 Acetone degradation by fermenting bacteria………………………………4 1.6 Energetic considerations of acetone degradation………………………….5 1.7 SRB and genome sequences………………………………………………6 1.8 Hypothesis………………………………………………………………..6 1.9 Aim of the thesis………………………………………………………….7 CHAPTER 2 Carbonylation as a key reaction in anaerobic acetone activation by Desulfococcus biacutus………………………………...………...8 2.1 Abstract…………………………………………………………………...8 2.2 Introduction………………………………………………………………9 2.3 Experimental procedures………………………………………………...11 2.4 Results…………………………………………………………………...16 2.5 Discussion……………………………………………………………….25 Acknowledgements………………………………………………………..28 CHAPTER 3 ATP and thiamine pyrophosphate dependence of acetone degradation by the sulfate-reducing bacterium Desulfococcus biacutus monitored by a fluorogenic ATP analogue……………..29 3.1 Abstract…………………………………………………………………29 3.2 Introduction……………………………………………………………..30 3.3 Materials and Methods………………………..…………………………32 3.4 Results and Discussion…………………………………………………..34 3.5 Conclusions………………………………………………………………39 Acknowledgements………………………………………………………40 II
CHAPTER 4 Acetone utilization under sulfate-reducing conditions: draft genome sequence of Desulfococcus biacutus and a proteomic survey of acetone-inducible proteins…………………..…...…..…41 4.1 Abstract…………………………………………………………………41 4.2 Introduction……………………………………………………………..41 4.3 Materials and Methods…………………………………………………..43 4.4 Results…………………………………………………………………...46 4.5 Discussion……………………………………………………………….67 Acknowledgements…………………..………………………………….69 CHAPTER 5 General Discussion……………...…………………………………70 5.1 Activation of acetone by Desulfococcus biacutus…………………………….70 5.2 Comparison with other hydrocarbon activation………………………….74 5.3 Future research…………………………………………………………..76 SUMMARY…………………………………………………………………………...77 ZUSAMMENFASSUNG…………………………………………………………….78 RECORD OF ACHIEVEMENT……………………………………………...... 79 ABGRENZUNG DER EIGENLEISTUNG……………………………………….80 REFERENCES…………………………………………………………….………...81 SUPPLEMENTARY DATA………………………………………………………...91 SCIENTIFIC CONTRIBUTIONS LIST…………………………………………..95
CHAPTER 1
General Introduction
1.1 Important aspects about acetone
Acetone production is an important process in synthetic chemistry. Due to its physicochemical properties, acetone represents one the most used solvents in industry. Acetone is used mainly as an intermediate in the synthesis of methacrylate, bisphenol A, diacetone alcohol, and some other compounds (Sifniades et al., 2011). Acetone regulation was exempted by the EPA in 1995. It is neither considered as hazardous pollutant in the Clean Air Act, nor as a priority pollutant in the Clean Water Act (Sifniades et al., 2011). However, exposure to high concentration of acetone vapors causes eye irritation and narcosis. In addition, acetone has been reported as a toxic compound (Singh et al., 1994). Acetone gets into the environment also by bacterial fermentations, for example by several Clostridium species (Duerre et al., 1992; Han et al., 2011; Lépiz-Aguilar et al., 2013). It is also one of the three ketone bodies that are formed in the human body and are excreted by diabetic mammalians (Asagoe et al., 1968; Kalapos, 1997; Laffel, 1999). Therefore, the metabolic pathways by which acetone is degraded under different environments are of special interest.
1.2 Acetone degradation by aerobic bacteria
Acetone can be degraded by some aerobic bacteria and mammals (Bondoc et al., 1999; Koop and Casazza, 1985; Park et al., 1995) via oxygen-dependent hydroxylation to acetol mediated by cytochrome P450, as it was shown with Mycobacterium smegmatis (Landau and Brunengraber, 1987; Taylor et al., 1980). However, an enzyme converting acetone to acetol has not been described yet. The first report on different ways of methyl ketones
1
Chapter 1. General Introduction
degradation was made for hydrocarbon-utilizing bacteria (Lukins and Foster, 1963). The mechanism of acetone activation has been under discussion for several years. Evidence of a carboxylation reaction was shown with the photosynthetic bacterium
Rhodopseudomonas gelatinosa (Siegel, 1950; Siegel, 1954). Requirement of CO2 as a co- substrate for acetone degradation was observed for first time with a methanogenic co- culture (Platen and Schink, 1987), with Thiosphaera pantotropha which was able to grow aerobically as well as anaerobically (Bonnet-Smits et al., 1988), and with Rhodobacter capsulatus and other phototrophs (Birks and Kelly, 1997). The need for CO2 suggested a carboxylation of the methyl group of acetone, forming acetoacetate. A CO2- and ATP- dependent activation of acetone was also observed with cell-free extracts of the aerobic bacterium Xanthobacter autotrophicus strain Py2 (Sluis and Ensign, 1997). The carboxylation of acetone was also investigated with Rhodococcus rhodochrous, but in this case the activity was not stimulated with ATP, but it depended on the presence of other nucleotides like GTP, ITP, CTP and UTP (Clark and Ensign, 1999). Acetone carboxylase was first purified from Xanthobacter autotrophicus strain Py2 (Sluis and Ensign, 1997). Its further characterization and comparison with the carboxylase of the phototrophic bacterium Rhodobacter capsulatus showed that they are identical in subunit composition (α2β2γ2 multimers of 85-, 78-, and 20-kDa subunits) and in kinetic properties (Sluis et al., 2002). Electron paramagnetic resonance (EPR) spectra of cell extracts of acetone-grown cells of R. capsulatus showed that in this bacterium the acetone carboxylase was manganese-dependent (Boyd et al., 2004).
1.3 Acetone degradation by nitrate-reducing bacteria
Nitrate-dependent acetone oxidation was shown with Thiosphaera pantotropha (Paracoccus pantotrophus) as mentioned above. The degradation of acetone was also studied with the denitrifying bacterium strain BunN, which essentially required CO2 for growth with acetone (Platen and Schink, 1989). Therefore, it was concluded that the degradation of acetone by strain BunN occurs similar to T. pantotropha via carboxylation reaction. This initial reaction was proven with cell-free extracts of strain BunN, in which a decarboxylation of acetoacetate occurred in the presence of ADP and MgCl2, and was specific for acetone-grown cells (Platen and Schink, 1990). Since a carboxylation reaction
2
Chapter 1. General Introduction
could not be proven, a different approach to support the carboxylation reaction was done with labeling experiments. Cell-free extracts of strain BunN catalyzed the exchange of
14CO2 into acetoacetate in an ADP-dependent reaction (Janssen and Schink, 1995a). Acetoacetate was then converted to a labeled acetoacetyl-CoA by succinyl-CoA: acetoacetate CoA transferase. Those experiments led to the conclusion that the activating reaction occurred in an ATP-dependent carboxylation of acetone. The carboxylating enzymes were analyzed with the nitrate reducers Alicycliphilus denitrificans K601, Paracoccus denitrificans, and Paracoccus pantotrophus (Dullius et al., 2011). The activity of carboxylase from the enriched enzymes was dependent on ATP, and could be detected also with butanone. However, the carboxylation reaction was also detected with UTP, ITP, and GTP. The enriched fractions were analyzed by SDS-PAGE where it was observed molecular weights similar to the aerobic acetone carboxylases. Sequence analysis of the enriched fractions resulted in similar subunit composition (85.3-, 78.3-, and 19.6-KDa) as it was found for the acetone carboxylases of X. autotrophicus and R. capsulatus. Recently, an ATP-dependent carboxylation of acetone was again shown with the nitrate reducer Aromatoleum aromaticum (Schühle and Heider, 2012) which was reported before to be able to carboxylate acetophenone to benzoylacetate during anaerobic ethylbenzene degradation (Jobst et al., 2010). In both carboxylation reactions the presence of ATP was needed. At the moment it is well established that aerobic and nitrate-reducing bacteria employ similar carboxylating mechanisms to activate acetone, and that the carboxylating enzymes are molecularly highly similar.
1.4 Acetone degradation by sulfate-reducing bacteria (SRB)
Under sulfate reducing conditions the degradation of acetone has been studied with the Gram-negative bacterium Desulfococcus biacutus (Janssen and Schnik, 1995; Platen et al., 1990), and with Desulfobacterium cetonicum (Janssen and Schink, 1995b). The degradation of acetone by SRB appeared to depend on the presence of CO2 as well. However, neither activity of acetone carboxylase was detected in cell-free extracts, nor acetoacetate- decarboxylating activity could be found. These results indicated that acetoacetate is not an intermediate in the activation of acetone by SRB. Formation of a free intermediate was excluded, based on the absence of CoA transferase or CoA ligase activity in cell-free
3
Chapter 1. General Introduction
extracts of acetone-grown cells. The presence of acetoacetyl-CoA thiolase suggested that acetone may be activated in a reaction that probably leads directly to acetoacetyl-CoA. The possibility of a carbonylation reaction leading to a 3-hydroxybutyryl-CoA derivative was studied with D. biacutus, but without any success (Dullius, 2011). Since D. biacutus oxidizes acetyl residues through the Wood-Ljungdahl pathway, it possesses CO dehydrogenase activity. Therefore, this bacterium is able to convert CO2 to CO and employ this as a co-substrate in acetone activation. Studies in cell-free extracts of acetone-grown cells did not show activity of 3-hydroxybutyryl-CoA dehydrogenase. Moreover, 3-hydroxybutyrate was never found as the product of acetone activation. These results indicated that 3-hydroxybutyrate is not an intermediate in the metabolism of acetone. From all these studies it was concluded that the activation of acetone in SRB, or at least in D. biacutus may occur through a mechanism different from the carboxylation reaction described for aerobic and nitrate-reducing bacteria.
1.5 Acetone degradation by fermenting bacteria
Acetone can also be degraded in the absence of electron acceptors (Platen et al., 1994; Platen and Schink, 1987; Symons and Buswell, 1933; Wikén, 1940). An enrichment culture (WoAct) that was obtained from anoxic sediment was able to degrade acetone anaerobically (Platen and Schink, 1987). Microscopy studies showed that one of the cooperation partners was a filamentous bacterium highly similar to Methanothrix sp (Methanosaeta sp). Inhibition experiments with streptomycin to avoid acetone degradation, and with acetylene and bromoethanesulfonate to inhibit acetate degradation, confirmed that acetone was converted to acetate which was further degraded to methane and CO2. Further studies with the enrichment culture WoAct indicated that acetone degradation by the fermenting bacterium does not necessarily depend on acetate removal by the methanogenic partner (Platen et al., 1994). However, acetone degradation was impeded when the concentration of acetate reached 10 mM. Up to now it is clear that acetone can be degraded in syntrophy, but the mechanism of acetone activation in this process has not been studied in detail yet.
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Chapter 1. General Introduction
1.6 Energetic considerations of acetone degradation
Acetone is activated by aerobic and nitrate-reducing bacteria by carboxylation in a CO2- and ATP-dependent reaction to form acetoacetate. According to equation 1 (Eq. 1), activation of acetone via carboxylation is an endergonic reaction that requires less than one ATP.
+ → − + + ∆ -1 (Eq. 1) CH 3COCH 3 CO2 CH 3COCH 2COO H ( G0´ = +17.1 kJ mol )
In the proposed mechanism, the γ and β phosphodiester bonds of ATP need to be hydrolyzed during the reaction; one is invested for the enolization of acetone (to the form of phosphoenolacetone), and the other one is used to carboxylate the enolized acetone to acetoacetate. After carboxylation, acetoacetate is activated to acetoacetyl-CoA with the investment of one further ATP. In total, two ATP equivalents are invested for acetone activation and one more for further acetoacetate degradation. This energy expenditure can be afforded by aerobic and nitrate-reducing bacteria because the subsequent oxidation of the acetyl moieties releases sufficient energy. Therefore, growth with acetone by those bacteria is possible not affected despite such energy expensive activation reaction. Acetone degradation by SRB is energetically more difficult. According to equation 2, the degradation of acetone coupled to the reduction of sulfate yields less than two ATP.
+ 2− + + → + + ∆ (Eq. 2) CH 3COCH 3 2SO4 4H 3CO aq)(2 2H S aq)(2 3H 2O ( G0´ = - 115.3 kJ mol-1)
Oxidation of the acetyl residue of acetyl-CoA through the CO dehydrogenase (“Wood- Ljungdahl”) pathway can form only about one ATP equivalent per acetyl residue. Thus, acetone degradation through the carboxylation reaction described above (eq. 1) could not be supported through the subsequent oxidation of the acetyl residues. Therefore, a different mechanism for CO2-dependent acetone activation has to be postulated for these bacteria.
5
Chapter 1. General Introduction
1.7 SRB and genome sequences
SRB are prokaryotes that are able to perform dissimilatory sulfate reduction. Those prokaryotic microorganisms including bacteria and archaea are organized in five different phylogenetic lineages (Thauer et al., 2007): the mesophilic deltaproteobacteria, the Gram- positive bacteria, the thermophilic Gram-negative bacteria, the Euryarchaeota, and theThermodesulfobiaceae. SRB are ubiquitous in nature, therefore, they can be found in various anoxic environments where sulfate is present, like soil, sediments, marine and freshwater, for example. Depending on their ability to oxidize the C2 carbon unit of acetyl-CoA to CO2, they can be grouped as complete oxidizers or incomplete oxidizers. There are two known ways for the oxidization of the acetyl residues; the first one is through the tricarboxylic acid cycle, and the second one is through the carbon monoxide dehydrogenase (CODH) pathway.
The completed genomes of several SBR are already available. Among them are found: Archaeoglobus sulfaticallidus strain PM70-1 (Stokke et al., 2013), the marine deltaproteobacterium Desulfobacula toluolica Tol2 which is able to degrade aromatic compounds (Wohlbrand et al., 2013), Desulfobacca acetoxidans strain ASRB2 (Goker et al., 2011), Strain NaphS2 which can grow anaerobically on naphthalene (DiDonato et al., 2010), Desulfobacterium autotrophicum strain HRM2 able to grow on fatty acids (Strittmatter et al., 2009), for example. Despite the sequenced genomes of SRB, the genome of an acetone-degrader sulfate reducing microorganism has not been reported until now.
1.8 Hypothesis
Since the degradation of acetone by D. biacutus theoretically cannot be driven by the same carboxylation mechanism that is employed by aerobic and nitrate reducers, a novel mechanism must be involved for the activation under strictly anoxic conditions. The involvement of a carbonylation reaction possibly leading to an aldehyde derivative is postulated for this bacterium.
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Chapter 1. General Introduction
1.9 Aim of the thesis
In the present work the mechanism of activation of acetone by the sulfate-reducing bacterium Desulfococcus biacutus is to be elucidated. It is desirable to find the product of the activation of acetone, and confirm the possible involvement of a carbonylation reaction. Whether the initial reaction is ATP-dependent is also part of the objectives of this work. To give a better understanding on the reaction mechanism, the enzymes that are involved in the strictly anaerobic acetone degradation by D. biacutus need to be identified. Furthermore, obtaining the genome sequence and genome annotation of D. biacutus representing the first acetone-degrading sulfate-reducing bacterium is one of the main targets of the thesis.
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CHAPTER 2
Carbonylation as a key reaction in anaerobic acetone activation by Desulfococcus biacutus
Olga B. Gutiérrez Acosta, Norman Hardt and Bernhard Schink Published in Applied and Environmental Microbiology (2013) 79(20):6228-6235
2.1 Abstract
Acetone is activated by aerobic and nitrate-reducing bacteria via an ATP-dependent carboxylation reaction to form acetoacetate as the first reaction product. In the activation of acetone by sulfate-reducing bacteria, acetoacetate has not been found as an intermediate. Here, we present evidence of a carbonylation reaction as the initial step in the activation of acetone by the strictly anaerobic sulfate reducer Desulfococcus biacutus. In cell suspension experiments, CO was found to be a far better co-substrate for acetone activation than CO2. The hypothetical reaction product acetoacetaldehyde is extremely reactive and could not be identified as a free intermediate. However, acetoacetaldehyde dinitrophenylhydrazone was detected by mass spectrometry in cell-free extract experiments as a reaction product of acetone, CO, and dinitrophenylhydrazine. In a similar assay, 2-amino-4-methylpyrimidine was formed as product of a reaction between acetoacetaldehyde and guanidine. The reaction depended on ATP as a co-substrate. Moreover, activity of aldehyde dehydrogenase (CoA acylating) tested with the putative physiological substrate was found at specific activity of 153 ± 36 mU mg-1 protein, and was specifically induced in cell-free extracts of acetone-grown cells. Moreover, acetoacetyl-CoA was detected (by mass spectrometry) after the carbonylation reaction as the subsequent intermediate after acetoacetaldehyde is formed. These results together
8
Chapter 2. Introduction
provide evidence that acetoacetaldehyde is an intermediate in the activation of acetone by sulfate-reducing bacteria.
2.2 Introduction
Acetone is produced by bacterial fermentations, for example by several Clostridium species (Han et al., 2011). It is also produced in chemistry as a solvent and as an intermediate in synthetic chemical industry. Aerobic degradation of methyl ketones was first observed with hydrocarbon-utilizing bacteria (Lukins and Foster, 1963). Acetone is degraded by some aerobic bacteria (Taylor et al., 1980) and mammalian liver cells via oxygenase- dependent hydroxylation to acetol (Landau and Brunengraber, 1987). Carboxylation of acetone to acetoacetate as a means of acetone activation was first proposed for a methanogenic enrichment culture (Platen and Schink, 1987). Requirement of CO2 as a co-substrate for acetone degradation was also observed with the nitrate reducer Thiosphaera pantotropha (Bonnet-Smits et al., 1988) and with Rhodobacter capsulatus and other phototrophs (Birks and Kelly, 1997). The reaction was studied with the nitrate-reducing strain Bun N under anoxic conditions, and it was concluded that acetoacetate was formed in an ATP-dependent carboxylation of acetone (Platen and Schink, 1989; Platen and Schink, 1990).
Attempts to measure an in vitro carboxylation of acetone at that time were unsuccessful.
However, exchange of radioactively labeled CO2 with the carboxyl group of acetoacetate was catalyzed by cell-free extracts of strain Bun N (Janssen and Schink, 1995a). A similar
CO2- and ATP-dependent activation reaction was observed with the aerobic bacterium Xanthobacter autotrophicus strain Py2 (Sluis et al., 1996). A comparison between the acetone carboxylase of strain Py2 and the carboxylase of the phototrophic bacterium Rhodobacter capsulatus showed that they are identical in subunit composition (α2β2γ2 multimers of 85-, 78-, and 20-kDa subunits) and in kinetic properties (Sluis and Ensign, 1997; Sluis et al., 2002). A similar subunit composition was found recently with the acetone carboxylase of the nitrate reducer Aromatoleum aromaticum (Schühle and Heider, 2012), and with the
9
Chapter 2. Introduction
acetone carboxylases of Alicycliphilus denitrificans, Paracoccus denitrificans, and Paracoccus pantotrophus (Dullius et al., 2011). Thus, it appears well established that aerobic and nitrate-reducing bacteria activate acetone by an ATP-dependent carboxylation reaction. Because the γ and β phosphodiester bonds of ATP need to be hydrolyzed during the reaction, two ATP equivalents are invested into a reaction that theoretically would require less than one ATP (acetone + CO2 ‰ acetoacetate- + H+ ;∆G0’ = +17.1 kJ mol-1). At least one further ATP is required for acetoacetate activation to acetoacetyl-CoA. This energy expenditure can be afforded by aerobic and nitrate-reducing bacteria because the subsequent oxidation of the acetyl moieties releases sufficient energy.
Acetone degradation by sulfate-reducing bacteria (SRB) is energetically more difficult. Oxidation of the acetyl residue of acetyl-CoA through the CO dehydrogenase (“Wood- Ljungdahl”) pathway can form only about one ATP equivalent per acetyl residue. Thus, acetone degradation through the carboxylation reaction described above could not be supported through the subsequent oxidation of the acetyl residues. Therefore, a different mechanism for CO2-dependent acetone activation has to be postulated for these bacteria. Acetone degradation was studied with the sulfate-reducing bacteria Desulfococcus biacutus and Desulfobacterium cetonicum (Janssen and Schink, 1995b; Janssen and Schnik, 1995). No acetone-carboxylating or acetoacetate-decarboxylating activity could be found in cell-free extracts of these bacteria. There was high acetoacetyl-CoA thiolase activity present in acetone-grown cells, but no activity of an acetoacetate-activating CoA transferase or CoA ligase. Moreover, these bacteria excreted acetate at a 1:1 ratio during growth on butyrate or 3-hydroxybutyrate, but did not accumulate acetate during growth on acetone. From these results we concluded that acetoacetate is not a free intermediate in acetone metabolism, and that activation of acetone may lead directly to an activated acetoacetyl residue, e.g., acetoacetyl-CoA (Janssen and Schink, 1995b).
Since both sulfate reducers oxidize acetyl residues through the Wood-Ljungdahl pathway, they have CO dehydrogenase activity. Therefore, they could convert CO2 to CO and employ this as a co-substrate in acetone activation, to form acetoacetaldehyde rather than
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Chapter 2. Introduction
acetoacetate as a reaction product. In the present study, we elucidated this hypothesis with D. biacutus and found strong evidence for this novel type of reaction.
2.3 Experimental procedures
Bacterial growth conditions
Desulfococcus biacutus strain KMRActS was grown in freshwater mineral medium as described before (Janssen and Schink, 1995b; Widdel and Pfennig, 1981). The medium was reduced with 1 mM sulfide, buffered with CO2/bicarbonate, and adjusted to a final pH of 7.2. Cells were grown in 1 l flasks with medium supplemented with 5 mM acetone or 5 mM butyrate as sole carbon source, and 10 mM sulfate as the electron acceptor.
Cultures were incubated under a strictly anoxic N2/CO2 (80/20) atmosphere at 30°C in the dark.
Cell suspension experiments
Cells were harvested in the late exponential growth phase at an optical density (OD 600) of 0.3. All experiments with cell extract and cell suspensions were done under strictly anoxic conditions inside an anoxic glove box. Cells were centrifuged at 6,000 x g at 10°C. The pellet was washed at least twice with 50 mM potassium phosphate (KP) buffer, pH 7.2, supplemented with 3 mM dithioerythritol as reducing agent. Cells were re-suspended in the same buffer with the addition of NaCl (1.0 g * l-1) plus MgCl2 * 6 H2O (0.6 g * l-1).
Cell suspensions with a final OD 600 of 12 were prepared in 5 ml flasks containing KP buffer with 5 mM acetone and 10 mM sulfate. The sulfate-reducing activity was measured at different time intervals for several hours. The gas phase was either N2/CO (90/10),
N2/CO2 (80/20), or N2.
Preparation of cell-free extracts
Cells were harvested as described above, however, at 4°C. The cell pellet was re- suspended in the KP buffer described above, containing 0.5 mg DNase ml-1 and 1 mg ml-1 of complete protease inhibitor cocktail (Complete Mini, EDTA-free protease
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Chapter 2. Experimental procedures
inhibitor cocktail tablets, Roche Diagnostics GmbH, Mannheim, Germany). Cells were disrupted by passing them two times through a cooled French pressure cell at 100 MPa. Cell debris and unopened cells were removed by centrifugation at 27,000 x g for 20 min at 4°C.
Carbon monoxide dehydrogenase (CODH) assay
Activity of CO dehydrogenase was measured at 30°C with a photometer 100-40 (Hitachi, Tokyo, Japan). Cell-free extracts of acetone-grown cells were used for enzyme assays. Enzyme activity was tested in the already described KP buffer with the addition of 2 mM benzyl viologen (BV) as the electron acceptor. The activity was tested in cuvettes previously flushed with CO, or by addition of CO to the complete reaction mixture. The effect of CODH inhibition by potassium cyanide (KCN) was checked with a final concentration of 3 and 5 mM KCN. Reduction of BV was followed at 578 nm (ε 578 nm = 8.65 mM-1cm-1). One unit was defined as 1 µmol of BV reduced per min.
Aldehyde dehydrogenase (CoA acylating) assay
Activity of aldehyde dehydrogenase was measured in anoxic cuvettes in the same Hitachi photometer. Cell-free extracts of acetone-grown cells were used for enzyme assays; control experiments with cell-free extracts prepared from butyrate-grown cells were run under the same conditions. Enzyme activity was followed in 50 mM KP buffer, pH 7.2, supplemented with 3 mM dithioerythritol as described before, with the addition of 2 mM coenzyme A (CoA) and 5 mM NAD+ as the electron acceptor. The reaction was started by addition of 2 mM acetaldehyde, or by addition of 20 µl of acetoacetaldehyde- containing solution (see below). NADH formation was followed at 340 nm (ε340 nm = 6.292 mM-1·cm-1). Control assays were run with boiled cell-free extracts. One unit was defined as 1 µmol of NAD reduced per min. Preparation of acetoacetaldehyde solution was done as follows: 20 µl (9.96 mg) of acetylacetaldehyde dimethyl acetal (4,4- dimethoxy-2-butanone, ALDRICH Chemistry, SIGMA-ALDRICH) was mixed with 40 µl of 37% HCl in 2 ml KP buffer, and stirred for 20 min. The reaction mix was diluted with four volumes of the same KP buffer (200 µl in 1 ml), and from this final mixture 20 to 25 µl was added to the cuvette for assay of acetoacetaldehyde dehydrogenase.
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Chapter 2. Experimental procedures
Activation of acetone in cell-free extract
Cell-free extracts of D. biacutus cells grown with acetone were used for enzyme assays; control experiments were run with extracts of butyrate-grown cells. All assays were carried out under strictly anoxic conditions at 30°C. Activation of acetone was tested in a total volume of 4 ml with 5 mM acetone, 5 mM ATP, and CO (10% in the headspace) as co-substrate. The reaction mix was incubated under mild stirring for at least 3 h, and samples were taken at different time intervals with syringes previously flushed with N2. Increment of carbonyl groups was quantified with 2,4-dinitrophenyl hydrazine (DNPH) or by derivatization of the reaction product with guanidine hydrochloride to form 2- amino-4-metylpyrimidine (see methods section below). The acetone activation reaction was also tested in the presence of 5 mM potassium cyanide (KCN), an inhibitor of CO oxidation by CO dehydrogenase (Wang et al., 2013). In a further reaction setup, the same reaction mix received in addition 2 mM CoA and 5 mM NAD+. Samples of 250 µl were taken at different time intervals and acidified with 50 µl of 3 M HCl, followed by centrifugation at 10,000 x g for 10 min. The supernatant was mixed with acetonitrile (50:50) and used for the assay of acetoacetyl-CoA by electrospray ionization mass spectrometry (ESI-MS). Authentic acetoacetyl-CoA (Sigma) was used as reference.
Preparation of acetoacetaldehyde for derivatization with DNPH
Acetoacetaldehyde was prepared by chemical deprotection of acetylacetaldehyde dimethyl acetal. 9.96 mg of the protected compound was mixed with 100 µl of 1.25 M HCl in methanol, and 800 µl of acetonitrile. The reaction mix was stirred under N2 atmosphere at room temperature, and was monitored by thin layer chromatography (TLC).
Chemical synthesis of DNPH derivatives
Dinitrophenylhydrazine (DNPH; 10 mg) was dissolved in 10 ml acetonitrile or ethyl acetate, and stirred for approx. 30 min under N2 atmosphere until a clear red solution was obtained. The product of the acetoacetaldehyde dimethylacetal deprotection reaction was transferred immediately into the DNPH solution and kept at room temperature while stirring for 60 min. The reaction was followed by TLC. The product of the derivatization was purified by column chromatography using a mixture of 95% dichloromethane and
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Chapter 2. Experimental procedures
5% methanol as eluent. The product was analysed with ESI-MS and proton nuclear magnetic resonance (1H NMR) spectroscopy.
Isolation and characterization of DNPH derivatives from the enzyme reaction
The product of acetone activation was derivatized with DNPH. For that purpose, 300 µl of each sample taken from the reaction mix was introduced slowly into 300 µl of freshly prepared DNPH solution, and mixed for 1 h. DNPH derivatives were extracted by mixing a defined volume of the derivatization reaction mix with ethyl acetate. Derivatives were detected by high pressure liquid chromatography (HPLC), UV-spectrophotometry, and TLC. The main spot observed was scraped from the TLC plate and dissolved in dichloromethane. Further analysis using ESI-MS and 1H NMR spectroscopy was performed to characterize the derivatization product.
Derivatization with guanidine
The product of acetone activation was also derivatized with guanidine hydrochloride. The reaction conditions were set according to a procedure proposed before (Haley and Maitland, 1951). 300 µl of each sample taken from the reaction mix was introduced slowly into 500 µl of an aqueous 0.5 M guanidine hydrochloride solution (pH 9.0). The reaction was stirred for at least 24 h at 30°C. The reaction product was analysed with RP- HPLC and compared with a 2-amino-4-methylpyrimidine reference compound (ALDRICH Chemistry, SIGMA-ALDRICH).
Analytical methods
DNPH solution for quantification of carbonyl groups was prepared as follows: 0.1 g of DNPH (60% w/w) was slowly introduced into a solution of 2 M HCl. The solution was stirred for 2 h at room temperature and passed through a cellulose acetate membrane filter (Whatman ® OE 66, 0.2 µm). This solution was prepared freshly every time that it was required, as well as the standards for calibration curves. For determination of carbonyl compounds, samples from the acetone activation assay mix were slowly introduced into 100 µl of DNPH solution with further addition of 500 µl water and 100 µl 10 M NaOH. Samples were mixed for approx. 1 h, and subsequently the absorbance
14
Chapter 2. Experimental procedures
of the hydrazone derivative was measured spectrophotometrically at 540 nm and 360 nm. Standards of acetone dinitrophenylhydrazone were prepared the same way, and used for quantification of carbonyl groups. Analysis of 2-amino-4-methyl pyrimidine was performed by reversed-phase HPLC. A Shimadzu HPLC system equipped with UV/Vis diode array detector was used. For analysis, 50 µl samples were injected on a C-18 reverse-phase column (Grom-Sil 120 ODS, 5 µm, 150 x 4.6 mm; Grom). Eluents contained 10 mM Na2HPO4/KH2PO4 buffer, pH 7.0 (A) and acetonitrile (B), at a flow rate of 0.8 ml min−1. The elution cycle proceeded as follows: 20% B for 2 min, then a linear increase to 90% B within 9 min, and return to 20% B within 1 min, followed by an equilibration step at 20% B for 6 min. The DNPH derivative was detected with the same HPLC system using the following elution cycle: 10% B in the first minute, then a linear increase to 90 % B within 55 min, and a final 5 min equilibration at 10% B. Sulfide formed in the cell suspension experiments was quantified with the methylene blue method (Cline, 1969). Protein content of cell-free extracts was determined with the bicinchoninic acid assay (BCA protein assay kit, Thermo, Scientific). TLC was done on silica plates (silica gel 60, Merck). After drying the samples under air for 2 min, the run started with a mobile phase of 95% dichloromethane plus 5% methanol. Spots were visualized by UV light and I2 vapour. Mass spectrometric analysis was performed with an ESI source (ESI-IT: Bruker Esquire 3000 plus) in the positive and negative ion mode under the following fixed instrument settings: spray ion voltage, 1000 V; nebulizer, 13 psi; gas flow, 7 l min-1; capillary temperature, 300°C. For NMR analysis a Bruker Avance III 400 MHz spectrometer and Bruker AVIII 600 MHz spectrometer were used. 1H chemical shifts are reported relative to the residual solvent peak and are given in ppm (δ). Spectra were measured at approx. 17°C and processed using MestReNOVA (v5.3.1).
Chemicals
Most chemicals were of analytical grade and purchased from Acros, Fluka, Sigma, Merck or Aldrich, and were used without any further purification. Dry solvents were purchased from Fluka; solvents for column chromatography were either distilled from technical grade (dichloromethane) or purchased as for chromatography grade (ethyl acetate, and methanol).
15
Chapter 2. Results
2.4 Results
Acetone degradation in cell suspensions
As a first approach to examine the hypothesis of possible acetone carbonylation, we checked for acetone-dependent sulfate reduction in suspensions of intact cells of D. biacutus with CO, CO2 or N2 in the gas phase (Fig. 1A). Sulfide formation was measured as an indicator of acetone degradation. Fig. 1A shows that the highest activity and the highest extent of sulfide formation were detected with CO in the gas phase. With CO as co-substrate, sulfide was formed to a concentration of 8.5 mM after 3 h of reaction. With
CO2 only about one fourth of this activity was observed, and nearly no sulfide was produced in the absence of either CO, CO2, or acetone (Fig. 1A).
Table 1. Activity of CO dehydrogenase measured in acetone-grown cell extracts of Desulfococcus biacutus
Growth condition Sp act Approx. (mU/mg protein) % activity Without KCN 882 ± 191 100
With KCN (3 mM) 121 ± 16 14
With KCN (5 mM), 10-min preincubation 24 ± 2 3
With KCN (5 mM), 20-min preincubation 13 ± 2 1
Acetone activation in cell-free extracts
Activation of acetone with CO was tested in cell-free extracts. Since we expected formation of acetoacetaldehyde as reaction product the activity was measured by quantifying keto and aldehyde groups with DNPH. In the presence of acetone, CO and ATP, cell-free extracts of D. biacutus catalyzed the formation of ketone equivalents as it is shown in Fig. 1B. Inhibition of CODH with KCN was checked before testing the
16
Chapter 2. Results
carbonylation reaction. According to the results in Table 1, CODH is strongly inhibited after 20 min of incubation with 5 mM KCN. Therefore, to prevent a possible oxidation of CO by CODH, KCN was added to the acetone activation reaction mix to a final concentration of 5 mM and this mix was pre-incubated for 20 min before addition of acetone. Figure 1B shows that the presence of KCN did not affect the acetone activation reaction that was measured with quantification of carbonyl groups. The reaction was
stimulated by the presence of NH4+ ions or, less efficiently, by K+ ions.
Figure 1. Acetone degradation in cell suspensions and in cell-free extracts of Desulfococcus biacutus. (A) Sulfide production in cell suspension experiments. Acetone and sulfate were added at
concentrations of 5 and 10 mM, respectively, and CO and CO2 were present at initial concentration of 10 and 20% (v/v), respectively. (B) Formation of ketone equivalents measured with DNPH during acetone activation in cell-free extracts. Acetone and ATP were added at initial concentration of 5 mM each one, and CO was present at an initial concentration of 10% in the headspace. Inhibition of CO dehydrogenase was performed using 5 mM KCN. Before the addition of acetone, both samples were pre-incubated for 20 min with or without KCN.
17
Chapter 2. Results
Carbonylation of acetone was also tested in the same reaction system containing acetone, CO, and ATP with the addition of CoA, and NAD+. The formation of acetoacetyl-CoA was analysed by ESI-MS. In the mass spectrum shown in Fig. 2, a specific peak signal at m/z 343.6 (I) was assigned to a fraction of acetoacetyl-CoA, after the loss of 507.0 Da (II). Compared to an acetoacetyl-CoA standard, the same loss of the 507.0 Da (II) fraction was observed. The minor deviation of the mass analysis (0.5 mass units) is due to a calibration error of the ESI-MS system. The cleavage at one of the phosphorus-oxygen bonds produces the lost fraction which corresponds to 3’-Phospho-ADP. This loss has been observed to be a common phenomenon of acyl-CoA compounds (Haynes et al., 2008; Magnes et al., 2005; Norwood et al., 1990).
Figure 2. ESI-MS of acetoacetyl-CoA that was formed after the acetone activation reaction. Activation of acetone was tested in a total volume of 4 ml with 5 mM acetone, 5 mM ATP, and CO (10% in the headspace) as co-substrate, supplemented with 2 mM CoA, and 5 mM NAD+. The CoA derivative was detected in the negative mode. The signal at m/z 343.6 (indicated as I) was assigned to a fraction of acetoacetyl-CoA, after the loss of a fractal of 507.0 Da, corresponding to 3’-Phospho-ADP (indicated as II). The signal at m/z 765.6 belongs to CoA (indicated as III).
18
Chapter 2. Results
Identification of products from deprotection of acetoacetaldehyde dimethylacetal
The hypothetical reaction intermediate of acetone carbonylation, acetoacetaldehyde, is known to be highly reactive (Dolgikh et al., 1964; Price, 1992) and is not commercially available. The commercially available acetoacetaldehyde dimethylacetal could be easily deprotected in acidic solution and converted to acetoacetaldehyde. While TLC analysis indicated that only one compound was formed, in the ESI mass spectrum of this deprotection reaction (Fig. 3A), a specific peak at m/z 205.2 (IV) was observed, and was attributed to the trimer of acetoacetaldehyde; a second peak at m/z 273.2 (V) was attributed to the tetramer of acetoacetaldehyde, and a third one at m/z 291.3 (VI) belonging to the tetramer of acetoacetaldehyde plus water was also observed. With 1H NMR analysis we confirmed that one of the produced compounds was triacetylbenzene (see 1H NMR analysis data). Trimerization of acetoacetaldehyde to triacetylbenzene has been observed before (Dolgikh et al., 1964; Price, 1992). Therefore, we conclude that acetoacetaldehyde was produced during the deprotection reaction and reacted with itself to form triacetylbenzene and also the tetrameric derivative. Attempts to derivatize acetoacetaldehyde with DNPH in a two-step process including deprotection and derivatization or, alternatively, in a continuous reaction, led to compounds with the specific signals of the trimer and tetramer of acetoacetaldehyde, and presented a mass spectrum pattern that was highly similar to that produced without addition of DNPH (Fig. 3B), indicating that the DNPH adduct was not formed under these conditions.
19
Chapter 2. Results
Figure 3. Identification of products of acetoacetaldehyde dimethyl acetal deprotection reaction. (A) ESI-MS spectrum of the chemically deprotected acetoacetaldehyde dimethyl acetal. (B) ESI- MS spectrum after derivatization of the reaction products with DNPH. Products of acetoacetaldehyde cyclization were identified as follows: compound IV correspond to triacetylbenzene (trimmer of acetoacetaldehyde), compound V to the tetramer of acetoacetaldehyde, and compound VI represents compound V plus water.
Identification of the DNPH derivative formed during the acetone activation reaction
In order to identify the hypothetical intermediate acetoacetaldehyde, the product of acetone metabolism in cell-free extracts was derivatized with DNPH as described above. Samples taken from the reaction mix for acetone activation in cell-free extracts were reacted with DNPH. In the ESI-MS analysis the acetoacetaldehyde-DNPH derivative ion [M+Na]+ could be observed at m/z 288.9 (VII) after 180 min of enzyme reaction (Fig. 4A). All attempts to purify the acetoacetaldehyde DNPH derivative failed because the compound proved to be unstable during the isolation process. Interestingly, after column
20
Chapter 2. Results
chromatography (silica column; see materials and methods) we observed in the mass spectrum the presence of the tetramer of acetoacetaldehyde peak signal at m/z 273.2 and the remains of acetoacetaldehyde-DNPH at m/z 289.2 (Fig. 4B). Unfortunately, the acetoacetaldehyde DNPH derivative that was formed after the acetone activation reaction could not be isolated for analysis by 1H NMR spectroscopy.
Figure 4. Identification of reaction products of acetone activation by cell-free extracts. (A) ESI- MS spectrum of DNPH derivative from enzyme reaction product. Compound VII was attributed to the acetoacetaldehyde DNPH. (B) ESI-MS spectrum of DNPH derivative after column chromatography. Compound V represents the tetramer of acetoacetaldehyde, and compound VII was assigned to the acetoacetaldehyde DNPH. Spectra were measured in the positive mode.
NMR analysis
The products formed during deprotection of acetoacetaldehyde dimethylacetal were analysed by 1H NMR, and resulted in the following chemical shifts: 1H NMR (400 MHz, DMSO) δ 8.62 (s, 3H), 2.72 (s, 9H). This analysis showed the presence of
21
Chapter 2. Results
triacetylbenzene. The spectrum of commercial triacetylbenzene (TCI Europe) was checked for comparison, and resulted in the following chemical shifts: 1H NMR (400 MHz, DMSO) δ 8.63 (s, 3H), 2.72 (s, 9H).
Derivatization with guanidine
Another strategy to trap the hypothetical acetoacetaldehyde was an N-C-N condensation reaction to form 2-amino-4-methylpyrimidine (Haley and Maitland, 1951). We employed this type of condensation reaction as well to derivatize the acetoacetaldehyde hypothetically formed in the enzymatic acetone activation reaction. Samples taken from the reaction mix were reacted with guanidine hydrochloride, and the products were analysed by RP-HPLC. The analysis showed that this product appeared exactly at the same retention time (3.65 min) as a commercial reference of 2-amino-4- methylpyrimidine, and that it presented also the same UV-absorption spectra, thus indicating that the expected reaction between acetoacetaldehyde and guanidine had occurred (Fig. 5; A and B). The formed pyrimidine could be detected only if guanidine was present at high excess. Due to the high background of excess guanidine, it was not possible to follow this reaction by ESI-MS and NMR spectroscopy. However, since we detected 2-amino-4-methylpyrimidine, it is highly probable that acetoacetaldehyde was produced during the reaction of acetone with CO, and this conclusion is also supported by the detection of the acetoacetaldehyde DNPH derivative by mass spectrometry and by the detection of acetoacetyl-CoA after the carbonylation reaction. Formation of 2-amino- 4-methylpyrimidine in the assay system described above required ATP as a co-substrate. In the absence of ATP, no such product was formed (Fig. 6). Dependence on ATP was also confirmed in the test system using DNPH as trapping agent. The described reactions were observed only with cell-free extracts of acetone-grown cells. Control experiments with extracts of butyrate-grown cells did not produce DNPH or guanidine-reactive products.
22
Chapter 2. Results
Figure 5. Identification of the reaction product of acetone activation after derivatization with guanidine. (A) HPLC analysis of the 2-amino-4-methylpyrimidine reference compound and of the product formed during the reaction between the intermediate (acetoacetaldehyde) and guanidine. (B) Absorption spectra of the commercial 2-amino-4-methylpyrimidine and the product formed after the reaction between acetoacetaldehyde and guanidine.
Figure 6. Dependence of acetoacetaldehyde formation from acetone and CO on the presence of ATP as a co-substrate. The formation of 2-amino-4-methylpyrimidine was quantified as the product of the reaction between acetoacetaldehyde and guanidine. Acetone and ATP were added at initial concentrations of 5 mM. CO was present at an initial concentration of 10% in the headspace. Quantification of 2-amino-4-methylpyrimidine was done by measuring the
absorption at 290 nm.
23
Chapter 2. Results
Aldehyde dehydrogenase (CoA acylating) activity
Activity of aldehyde dehydrogenase was measured in cell-free extract of D. biacutus with acetaldehyde, and with acetoacetaldehyde that was prepared by deprotection of acetoacetaldehyde dimethylacetal. Previous experiments showed the instability of acetoacetaldehyde, nevertheless, this compound must be an intermediate before the trimer and tetramer are formed. Therefore this acidic solution was added also as a substrate for aldehyde dehydrogenase with the addition of CoA, and NAD+ as the electron acceptor. The activity was detected in cell-free extract of acetone-grown cells at 18 ± 3 mU mg-1 protein with acetaldehyde, and 153 ± 36 mU mg-1 protein with the acetoacetaldehyde preparation (Table 2).Addition of CoA caused an increase of the activity from 30 to 100%. A control assay with extract of butyrate-grown cells indicated that aldehyde dehydrogenase is specifically induced during the metabolism of acetone.
The activity with acetaldehyde increased 5 fold after addition of 20 mM NH4+ to the reaction mix.
Table 2. Aldehyde dehydrogenase (CoA acylating) activity measured in cell-free extracts of Desulfococcus biacutus.
Specific activity (mU per mg protein)
ªWith With acetaldehyde acetoacetaldehyde
+ + (-) NH4 (+) NH4
Acetone-grown cell-free extract 5 ± 0.5 18 ± 3 153 ± 36
Butyrate-grown cell-free extract *ND 1± 0.2 20 ± 7
* ND, Not detected ª Ammonium addition did not stimulate the reaction with acetoacetaldehyde.
24
Chapter 2. Discussion
2.5 Discussion
In the present study, degradation of acetone under strictly anoxic conditions was investigated with the sulfate-reducing bacterium Desulfococcus biacutus. Based on our experimental results, we propose that acetone is activated by carbonylation with CO to form acetoacetaldehyde, rather than by carboxylation to acetoacetate as described for aerobic or nitrate-reducing bacteria. CO proved to be a far better co-substrate for acetone degradation than CO2 in cell suspension experiments. Since D. biacutus can reduce CO2 to CO by its carbon monoxide dehydrogenase enzyme (Janssen and Schink, 1995b) CO is available as a co-substrate for this activation reaction.
Derivatization of carbonyl compounds with DNPH has been used in the quantification of aldehydes and ketones (Cirera-Domenech et al., 2013; Huang et al., 2007; Uchiyama et al., 2011; Van den Bergh et al., 2004). In our study, the increase of ketone equivalents measured with DNPH in cell-free extracts indicated that CO and acetone were condensed to an aldehyde molecule.
A simultaneous experiment in which CO dehydrogenase was inhibited by KCN gave a similar increase of the ketone equivalents, suggesting that CO is the real co-substrate for acetone activating reaction, rather than being oxidised to CO2 by CO dehydrogenase. This result is supported by cell suspension experiments. In control assays with chemically prepared acetoacetaldehyde, no reaction with DNPH was observed. Obviously, the formed acetoacetaldehyde had undergone a cyclization reaction to form a trimer as observed before (Dolgikh et al., 1964; Price, 1992). The formed 1,3,5-triacetylbenzene which was identified by ESI-MS and 1H NMR spectroscopy, further reacted to form the tetrameric compound. Interestingly, after the enzymatic acetone activation reaction the DNPH-acetoacetaldehyde derivative was detected by mass spectrometry with a peak at m/z 288.9, which strongly suggests that acetoacetaldehyde was indeed produced in the enzymatic activating reaction. While trying to purify this derivative we could detect the tetramer of acetoacetaldehyde (m/z 273.2) and minor amounts of the DNPH- acetoacetaldehyde derivative (m/z 289.2), indicating that the derivative might have
25
Chapter 2. Discussion
disintegrated, perhaps due to the acidic conditions that were used during the chromatographic separation process.
Formation of acetoacetaldehyde from acetone and CO is also supported by the formation of 2-amino-4-methylpyrimidine with guanidine as co-substrate, a reaction which is very specific for the detection of 1,3-dioxo aliphatics.
A further strong indication of the formation of an aldehyde as a first reaction product in acetone activation is the presence of aldehyde dehydrogenase activity. This activity was found only in extracts of acetone-grown cells, and was substantially higher when the putative physiological substrate was added, thus indicating that an aldehyde is formed specifically during degradation of acetone, and it is highly probable that this aldehyde is our hypothetical acetoacetaldehyde. Moreover, the detection of acetoacetyl-CoA after activation of acetone in the presence of CO, ATP, CoA and NAD+ supports again the formation of acetoacetaldehyde as intermediate.
The acetone-carbonylating activity was stimulated by monovalent cations such as NH4+ or K+. The activating enzyme differs from the ketone carboxylases employed by aerobic and nitrate-reducing bacteria, which depend on the presence of divalent cations such as Mg2+ and Mn2+ (Boyd et al., 2004; Jobst et al., 2010). However, the acetone-carbonylating enzyme activity of D. biacutus was stimulated by NH4+ ions, similar to the acetone carboxylases of Cupriavidus metallidurans strain CH34 and Xanthobacter autotrophicus strain Py2 (Rosier et al., 2012; Sluis et al., 2002). Neither genomic nor proteomic analysis of acetone-grown cells of D. biacutus provided any indication of acetone carboxylases similar to those described for aerobic or nitrate-reducing acetone oxidizers (unpublished results from our lab).
The observed conversion of acetone with CO to acetoacetaldehyde required ATP as a co-substrate. Perhaps this ATP is needed to stabilize the enol tautomer of acetone in the form of acetone enolphosphate. This compound is the real substrate of carboxylation by the acetone carboxylases described in the past (Boyd and Ensign, 2005; Schühle and Heider, 2012) and may as well be the real acceptor of CO in the carbonylation reaction proposed here. Since the reaction product acetoacetaldehyde is extremely reactive it
26
Chapter 2. Discussion
appears plausible that it is not released free into the cytoplasm but is immediately oxidized further to acetoacetyl-CoA, perhaps in a multienzyme complex. After all, acetone activation and conversion to acetoacetyl-CoA through this new carbonylation pathway (Fig. 7) would require a minimum of only one ATP equivalent rather than three as in the well-described carboxylation pathway, and would therefore be much better suited for bacteria operating at a small energy budget, such as sulfate-reducing bacteria. Thus, acetone activation is another example to demonstrate that strict anaerobes such as sulfate reducers use strategies in the degradation of comparably stable compounds that are basically different from those employed by nitrate reducers, as studies with various aromatic compounds have shown in the past (Philipp and Schink, 2012). The biochemistry of the novel acetone carbonylation reaction will be subject to further studies in our lab.
Figure 7. Acetone activation mechanism by aerobic and nitrate-reducing bacteria and the proposed novel activation by carbonylation in sulfate-reducing bacteria.
27
Chapter 2. Acknowledgments
Acknowledgments
We thank Prof. B. T. Golding (School of Chemistry, Newcastle University), Prof. Peter Kroneck (Konstanz University), and Tobias Strittmatter (Konstanz University) for valuable discussions. We thank Ines Joachim and Martin Ehrle for practical support, Antje Wiese for media preparation, and the Konstanz Research School Chemical Biology (KoRS-CB) for the fellowship granted to Olga Brígida Gutiérrez Acosta. This work was supported by the Deutsche Forschungsgemeinschaft (DFG) through the SPP 1319 priority program.
28
CHAPTER 3
ATP and thiamine pyrophosphate dependence of acetone degradation by the sulfate-reducing bacterium Desulfococcus biacutus monitored by a fluorogenic ATP analogue
Olga B. Gutiérrez Acosta, Norman Hardt, Stephan M. Hacker, Tobias Strittmatter, Bernhard Schink, and Andreas Marx. Submitted
3.1 Abstract
Acetone can be degraded by aerobic and anaerobic microorganisms. Studies with the strictly anaerobic sulfate-reducing bacterium Desulfococcus biacutus indicate that acetone degradation by these bacteria starts with an ATP-dependent carbonylation reaction leading to acetoacetaldehyde as first reaction product. The reaction represents the second example of a carbonylation in the biochemistry of strictly anaerobic bacteria, but the exact mechanism and dependence on cofactors is still unclear. Here we present the development of a novel Förster-resonance energy transfer-based ATP probe 1 which allows to follow the consumption of ATP in the carbonylation reaction in cell-free extracts of D. biacutus in real time. The fluorogenic ATP analogue acts as activity probe to follow nucleotide-dependent enzymatic reactions. Upon processing of the probe by an ATP-hydrolysing enzyme a specific fluorescence emission is initiated. Using this analytical approach, we found that thiamine pyrophosphate acts as a cofactor to enhance the enzymatic activity in the novel acetone carbonylation reaction.
29
Chapter 3. Introduction
3.2 Introduction
Acetone is an important solvent used in chemical industry (Sifniades et al., 2011). Its microbial degradation has been studied with aerobic and anaerobic bacteria. Both nitrate- reducing and aerobic bacteria activate acetone in an ATP-dependent carboxylation reaction which consumes two ATP equivalents and forms acetoacetate as a first reaction product (Dullius et al., 2011; Schühle and Heider, 2012; Sluis et al., 2002; Sluis et al., 1996). Desulfococcus biacutus was described as a strictly anaerobic chemoorganoheterotrophic bacterium. It can grow with various substrates as carbon source, for example acetone, and uses sulfate as electron acceptor (Platen et al., 1990). Unlike aerobic and nitrate-reducing bacteria, D. biacutus activates acetone not by carboxylation, but uses CO in the activation reaction (Figure 1). The product of this carbonylation reaction is acetoacetaldehyde, as shown recently (Gutierrez Acosta et al., 2013b). The exact mechanism of this novel reaction is still under investigation. Due to the instability of the product acetoacetaldehyde, the activating reaction is difficult to characterize, and therefore the dependence on cofactors could not be assayed so far. Proteomic analysis of acetone-grown cells of D. biacutus indicated that a thiamine pyrophosphate (TPP)-requiring enzyme may be involved in the activating reaction (data not published). TPP is an essential cofactor in several enzymatic reactions, including pyruvate decarboxylase (EC 4.1.1.1) (Kluger, 1987), transketolase (EC 2.2.1.1) (Lindqvist et al., 1992) and pyruvate dehydrogenase (EC 1.2.4.1) (Reed, 1974). The role of TPP in these reactions is to help in the catalysis of carbon-carbon bond-forming and bond- breaking reactions that occur adjacent to a carbonyl group (Agyei-Owusu and Leeper, 2009). Consequently, our aim was to evaluate and understand a possible involvement of TPP in the activation of acetone in cell-free extracts of D. biacutus.
In order to further investigate the novel acetone activation reaction we searched for new techniques. Recently, we introduced a conceptually novel Förster-resonance energy transfer (FRET)-based assay that monitors ATP consumption in real time (Hacker et al., 2013b). In this concept, an ATP analogue is intramolecularly equipped with a fluorescent donor at the γ- or δ-phosphate, respectively, and an acceptor attached to nucleoside part of ATP. In an intact state, the excited donor fluorophore can transfer its energy to the
30
Chapter 3. Introduction
acceptor via FRET. Upon cleavage of the probe by an enzyme, the acceptor dye is spatially separated from the donor dye and the specific fluorescence emission of the donor is restored. By this concept, the activity of enzymes can be detected at high spatiotemporal resolution. In the present study, we describe the rational design and the total synthesis of a novel fluorogenic ATP analogue 1 that was tailored for the analysis of enzymatic reactions of D. biacutus. In order to use this analogue as activity probe towards ATP-dependent enzymes, it is required to ensure its stability in cell extract. Mono γ- modified nucleotides are used usually to be converted by γ-phosphate transferases and to label specific substrates (Bagshaw, 2001; Cole and Yount, 1990; Cremo et al., 1990; Lee et al., 2009; Oiwa et al., 2000; Oiwa et al., 2003; Oiwa et al., 1998; Song et al., 2008; Wang et al., 1999). Most modifications are phosphonyl- (Arzumanov et al., 1996; Green and Pflum, 2007), phosphorester- (Bettendorff et al., 2007; Freeman et al., 2010; Knorre et al., 1976; Korlach et al., 2008; Kruse et al., 1988; Lee et al., 2009; Niyomrattanakit et al., 2011; Sood et al., 2005) or phosphoramide (Green and Pflum, 2007; Green and Pflum, 2009; Knorre et al., 1976; Kruse et al., 1988; Kumar et al., 2005; Lee et al., 2009; Pollack and Auld, 1982; Song et al., 2008; Yarbrough et al., 1979; Zinellu et al., 2010) bonds, respectively. However, Marx et. al.(Hacker et al., 2012) showed recently that phosphorester modifications are stable in a wide pH range and are therefore suited to investigate enzyme reactions. Moreover, we demonstrated that modifications at the C2-position of ATP are tolerated by a broad variety of different ATP consuming enzymes (Hacker et al., 2013a). The synthesis route was developed to be as flexible as possible in order to vary the dyes of the FRET cassettes, as shown previously (Hacker et al., 2013b). However, not all previously synthesised ATP analogues are suitable for studies performed in cell extract due to limited stability of the dyes. According to our investigations sulfoCy3 and eclipse™-quencher turned out to be a FRET cassette that is stable under such conditions. Therefore, we designed and synthesized the novel γ-sulfo-Cy3, C2-eclipse- ATP analogue 1 which opens up the use of ATP activity probes in cell extracts for the first time. The novel probe carries a sulfo-Cy3 dye at the γ-moiety of the triphosphate and an eclipse™-quencher connected to the C2-position of the nucleobase. With this probe 1 in hand we provide a tailor-made nucleotide activity probe to further investigate ATP-dependent reactions, e. g., in the metabolism of D. biacutus. Using this analytical
31
Chapter 3. Introduction
approach, we focused our research on the acetone-activating reaction by D. biacutus and checked for thiamine pyrophosphate (TPP) as a possible cofactor in this unusual novel reaction.
Figure 1. Concept of the quenched ATP probe and biological pathway of interest. (A) Modified ATP probe bearing a fluorescent donor (D) attached to the γ-phosphate group and a quencher (Q) attached to the C2-position of the nucleobase. Due to FRET no fluorescent emission of D can be detected. After cleavage of the ATP probe FRET is no longer possible and direct emission of D can be observed. (B) Reaction scheme of ATP- and CO- dependent acetone activation by Desulfococcus biacutus.
3.3 Materials and Methods
Synthesis of ATP analogue
The description of the organic synthesis of the ATP analogue that was used in this work can be found in Norman Hardt doctoral dissertation, or in the supporting information of the published version of this manuscript.
Cell cultures
The sulfate-reducing bacterium Desulfococcus biacutus strain KMRActS was grown in freshwater mineral medium as described before (Janssen and Schnik, 1995). The medium was reduced with 1 mM sulfide, buffered with CO2/bicarbonate, and adjusted to a final pH of 7.2. Cells were grown in 1 L flasks with medium supplemented with 5 mM acetone or 5 mM butyrate as sole carbon source, and 10 mM sulfate as the electron acceptor.
32
Chapter 3. Materials and Methods
Cultures were incubated under a strictly anoxic N2/CO2 (80/20) atmosphere at 30°C in the dark.
Preparation of cell extract
Cells of D. biacutus were harvested in the late exponential growth phase at an optical density of 0.3 (OD 600). All experiments with cell extract were done under strictly anoxic conditions. Cells were centrifuged at 6,000 x g at 4°C and the pellet was washed at least twice with 50 mM KP buffer, pH 7.2, supplemented with 3 mM dithioerythritol as reducing agent. The cell pellet was suspended in the same buffer plus 0.5 mg DNase mL- 1 and 1 mg mL-1 of complete protease inhibitor (Complete Mini, EDTA-free protease inhibitor cocktail tablets, Roche Diagnostics GmbH). Cells were disrupted by passing them twice through a cooled French pressure cell at 100 MPa. Cell debris and un-opened cells were removed by centrifugation at 27,000 x g for 20 min at 4°C.
Reaction of γ-sulfo-Cy3, C2-eclipse-ATP 1 in cell-free extract
Cell-free extract of D. biacutus prepared after growth on acetone was used to test the carbonylation reaction. All enzyme assays were carried out under strictly anoxic conditions. The buffer solution was the same as described for washing the cells pellet, and contained 1 g L-1 NaCl, 0.6 g L-1 MgCl2 H2O and 3 mM dithioerythritol. The enzyme reaction was tested in Eppendorf tubes sealed with a rubber stopper. Concentrations of substrates were 1 mM acetone, 0.1 mM ATP analog 1, 2 mM TPP and 10% CO in the headspace. The reaction was incubated for 1 hour at 30°C. For the time course experiments, samples were taken at different time intervals with syringes previously flushed with N2. All reactions were stopped with 2 mM EDTA solution. Each reaction condition was run in triplicate.
Fluorescent read out
From each sample, 10 µL aliquots were diluted and uniformly mixed with 90 µL Milli Q water and placed onto a dark 96 well plate for fluorescence measurement. The procedure was run three times for each triplicate.
HPLC analysis of the assay mixtures
33
Chapter 3. Materials and Methods
Protein was precipitated by the addition of 10 µL of 2 M H2SO4, and removed by centrifugation at 10,000 x g. The supernatant was analysed by analytical RP-HPLC as described above. Fractions containing ATP fragments were collected and further characterized by HRMS.
HRMS characterisation of γ-sulfo-Cy3, C2-eclipse-ATP 1 and its fractals after incubation in cell-free extract
Fractions were collected during RP-HPLC separation and analysed by HRMS. Negative probe: HR-ESI-MS 1: found: 827.7091; calculated: 827.7107 (M-2H+,
C69H85ClN13O23P3S22-); deviation: 1.9 ppm. Positive probe: HR-ESI-MS 7: found:
888.2377; calculated: 888.2360 (M-H+, C37H53N3O14P2S2-); deviation: 1.9 ppm.
3.4 Results and Discussion
Synthesis of the ATP analog 1
The synthesis of γ-sulfo-Cy3, C2-eclipse-ATP 1 starts with the linker-modified adenosine triphosphate 2. Nucleotide 2 was synthesized in a seven-step procedure starting from guanosine as reported (Hacker et al., 2013a) (Figure 2). The modification at the γ-position of the triphosphate was introduced by treatment of 2 with 1-bromo-6-azido-hexane in the presence of molecular sieve (MS) to get the doubly linker-modified building block 3 with 70% yield. Afterwards, protection group manipulation was done by ammonolysis to yield compound 4 with the deprotected amino function at 53% yield. Eclipse™-quencher was introduced by treatment of the free amine with the corresponding N- hydroxysuccinimide (NHS) ester of the dye to form 5 with 26% yield. The azide in compound 5 was reduced using tris (2-carboxyethyl)phosphine (TCEP), generating the functional amino group of compound 6 at 57% yield. Finally, sulfoCy3 was introduced analogous to the previous step using NHS ester chemistry, resulting in ATP analogue 1 with 80% yield.
34
Chapter 3. Results and Discussion
Figure 2. Synthesis of γ-sulfo-Cy3, C2-eclipse-ATP 1. Compound 2 was synthesised in a seven step procedure according to Marx et. al. starting from guanosine. a) 1-Azido-6-bromohexane,
DMF, MS, r.t., overnight, 3 in 70% of yield. b) 16% aq NH3, r.t., 2 h, 4 in 53% of yield. c) Eclipse™-quencher-NHS, 0.1 M NaHCO3 (pH 8.7), DMF, r.t., overnight, 5 in 26% of yield. d) TECEP, H2O/MeOH/NEt3, r.t., 4 h, 6 in 57% of yield. e) SulfoCy3-NHS, 0.1 M NaHCO3 (pH 8.7), DMF, r.t., overnight, 1 in 80% of yield.
To evaluate the general concept and optical properties of our synthesized probe 1, fluorescence spectra were measured before and after its complete cleavage. To monitor the efficiency of this process, the cleavage of γ-sulfo-Cy3, C2-eclipse-ATP 1 at α-,β- position was promoted using the snake venom phosphodiesterase (SVPD) of C. adamanteus that is known to efficiently cleave this type of molecules and liberate both dyes of the FRET cassette from each other (Figure 3A). Due to the spatial separation of both dyes FRET is no longer possible, and direct emission of the donor can be detected. Recordings of fluorescence spectra before and after SVPD treatment of probe 1 gave qualitative and quantitative insights into the efficiency of the cleavage process and the validity of the emerging signal (Figure 3B). Moreover, reversed phase high pressure liquide chromatography (RP-HPLC) analytics of the SVPD-digested fluorogenic ATP probe, and the negative control in the absence of SVPD showed a clear shift in retention time and absorption properties, indicating a quantitative conversion of the probe after 30 min of incubation (Figure 3C). Fractions of the RP-HPLC analysis were analyzed by high resolution mass spectrometry (HRMS) to clearly identify the non-cleaved probe 1 and the cleavage products 7 and 8. We could show that upon cleavage of the donor dye from the
35
Chapter 3. Results and Discussion
acceptor the ratio of fluorescence intensity changes dramatically rendering the probe suitable as nucleotide-based activity probe towards ATP consuming enzymes.
Figure 3. Proof of concept of γ-sulfo-Cy3, C2-eclipse-ATP 1. (A) The fluorogenic ATP activity probe 1 is incubated with phosphodiesterase of C. adamanteus (SVPD) that is known to efficiently cleave this type of molecules. (B) Investigation of fluorescence properties via fluorescence emission spectra of the ATP probe 1 without (black) and after digestion (grey); excitation wavelength 532 nm. (C) RP-HPLC analytics of the negative control without SVPD (top) and the SVPD digested ATP probe 1 (bottom) were performed according to the general procedures. After 30 min of incubation quantitative conversion is detected. Fractals of negative and positive control were identified by HRMS
Use of the ATP analogue 1 in cell-free extracts of D. biacutus
Activation of acetone in D. biacutus proceeds in an ATP-dependent carbonylation reaction to acetoacetaldehyde (Gutierrez Acosta et al., 2013b). This metabolic intermediate is highly unstable (Dolgikh et al., 1964; Price, 1992), and could be identified only as the dinitrophenylhydrazone derivative, or after derivatization with guanidine leading to 2-amino-4-methylpyrimidine. Unlike nitrate-reducing and aerobic bacteria, D. biacutus cannot invest two ATP equivalents in this activation reaction as nitrate-reducing
36
Chapter 3. Results and Discussion
and aerobic bacteria do (Dullius et al., 2011; Schühle and Heider, 2012; Sluis et al., 2002), since their overall energy budget is much tighter. Nevertheless, acetone activation by sulphate reducers does require some energy input by ATP hydrolysis in the initial step. Accordingly, we showed recently that the carbonylation reaction in D. biacutus is ATP- dependent (Gutierrez Acosta et al., 2013b).
In order to gain further insights into the reaction mechanism and the process of ATP consumption we employed ATP analogue 1 in the acetone-activating reaction, and checked for a possible requirement of cofactors that may help the activating mechanism, specifically TPP. Proteomic analysis had indicated that one of the enzymes specifically expressed after growth with acetone could bind TPP (data not published), thus, we were specifically interested in a possible involvement of TPP as cofactor in the metabolism of acetone. The activating reaction was evaluated under strictly anoxic conditions in cell-free extracts of acetone-grown cells, in the presence and absence of TPP. According to the concept of the novel ATP probe, fluorescence emission of the donor sulfoCy3 can be measured after cleavage of the ATP analogue 1. Indeed we found an increase of the fluorescent signal of the hydrolysed ATP analogue 1 by the acetone-activating reaction (Figure 4). Addition of TPP to the reaction mixture increased the hydrolysis rate of ATP analogue 1 up to two fold. This indicates that TPP acts as cofactor in the enzymatic activation of acetone in D. biacutus. The activity depended on acetone, CO, and TPP. The ATP analogue 1 was used to a small extent also by other ATP-cleaving enzyme(s) present in the cell-free extract as the background hydrolytic activity measured in the absence of acetone indicates. In the control assays under N2, no significant activity could be detected, thus confirming that CO is undoubtedly required for the activation of acetone (Figure 4 A and B).
RP-HPLC analysis of the reaction mixtures showed clear spectral properties and retention times in line with the SVPD experiments. The control reaction in the absence of CO gave a clear signal at 21 min, representing the non-cleaved probe 1. Cleavage of 1 promoted by TPP, CO, acetone, and CCE gave a signal at 16 min, representing the cleavage product 7 (Figure 4C). All reaction products were verified by HRMS (see supporting information).
37
Chapter 3. Results and Discussion
Figure 4. Reaction of ATP analogue 1 in cell-free extract of D. biacutus. All experiments were performed under strictly anoxic conditions at 30°C. Concentrations of the substrates were 1 mM acetone, 0.5 mM ATP probe 1, 2 mM TPP and 10% CO in the headspace. (A) Time course experiments of the hydrolysis of ATP probe 1 in the acetone activation reaction. (B) Activity of hydrolysis of ATP probe 1 after 60 min of enzymatic reaction. (C) RP-HPLC analysis after 60 min of the negative control (top) gave intact ATP probe with retention time of 21 min; positive control (bottom) showed fractal 7 with retention time of 16 min, and typical spectral properties. Fractals of negative and positive control were further identified by HRMS.
According to our findings, TPP assists the acetone degrading enzymes in D. biacutus, probably by acting as a cofactor in the activating reaction. However, the mechanism in which TPP is involved in acetone activation is still unclear. It could participate in a similar manner as in pyruvate decarboxylase: pyruvate acetaldehydetransferase. In a similar way, TPP could help to stabilize the unstable intermediate acetoacetaldehyde through formation of an acetohydroxyethyl-TPP intermediate. At this step, addition of CoAS- would form a CoA anion radical that would enable the release of acetoacetyl-CoA. Reactive aldehydes are highly toxic because of their ability to form Michael type adducts with thiol groups of proteins, inhibiting in this way crucial cellular processes (Esterbauer et al., 1975; Schauenstein, 1967). Formation of an acetohydroxyethyl-TPP intermediate
38
Chapter 3. Results and Discussion
would be a suitable way to prevent damage to the bacterial cell by the reactive acetoacetaldehyde.
Another way of involvement of TPP in our reaction may be similar to its hypothetical role in phosphoketolase (EC 4.1.2.9). The proposed mechanism for this enzyme reaction involves binding of a phosphorylated ketone compound with TPP to form an aldehyde. In a similar way acetone could be activated with ATP to form phospho-enolacetone which then binds to TPP. The hydroxyethyl-TPP intermediate can proceed further as in pyruvate oxidase (EC 1.2.2.2) or as in pyruvate oxidoreductase (EC 1.2.7.1) which involves a transfer of the acetyl group to CoA (Wahl and Orme-Johnson, 1987). The reaction mechanism is further investigated in our lab. With regard to the stability of the ATP analog 1, the abiotic control showed that this probe was stable under the reducing conditions that were applied in the reaction mix, thus supporting the value of this novel probe. With the use of the new ATP analogue we also showed that the applicability of this probe may be extended for understanding mechanisms of further reactions in anaerobic biochemical processes.
3.5 Conclusions
In the present study, we present a new doubly dye-labeled dark-quenched nucleotide- based activity probe 1 that is applicable to study ATP hydrolysis in cell extracts. The synthesized probe clearly demonstrates the potential of the concept of FRET- based activity probe by generating an easily detectable fluorescence signal. With this probe we enlarge the toolbox of nucleotide-based activity probes to a broader field of ATP- requiring processes. This probe might not only be of value for basic research but may open up novel avenues for subsequent applications in complex biological processes.
In particular we showed the application of this probe regarding the novel acetone activation reaction that was proposed to occur in the sulfate-reducing bacterium Desulfococcus biacutus. Addition of TPP revealed that this cofactor enhances the rate of ATP hydrolysis during acetone activation. We showed that the ATP analogue 1 was an
39
Chapter 3. Conclusions
adequate tool to successfully confirm that the activation of acetone by D. biacutus proceeds in an ATP-dependent carbonylation reaction as we have shown before in a different approach.
Acknowledgements
We gratefully acknowledge funding by the Deutsche Forschungsgemeinschaft within the SFB 969 and the SPP 1319 priority program, the Studienstiftung des Deutschen Volkes for a stipend to S. M. H. and the Konstanz Research School Chemical Biology for fellowships granted to O. B. G. A. and N. H. We thank Antje Wiese for preparation of bacterial growth media.
40
CHAPTER 4
Acetone utilization under sulfate-reducing conditions: draft genome sequence of Desulfococcus biacutus and a proteomic survey of acetone-inducible proteins
Olga B. Gutiérrez Acosta, David Schleheck and Bernhard Schink To be submitted
4.1 Abstract
The sulfate-reducing bacterium Desulfococcus biacutus activates acetone with CO as co- substrate. The mechanism of acetone activation is of great interest because it represents a novel type of activation under strictly anoxic conditions. In this study, the genome sequencing of Desulfococcus biacutus representing the first genome report of an acetone- utilizing sulfate reducer was established. The annotated genome signifies an important tool to investigate the genes and operons involved in the metabolism of acetone and some other compounds of interest. By using the annotated genome in combination with proteomic analyses we could identify the genes and proteins that are up-regulated during acetone metabolism by D. biacutus. Interestingly, a thiamine pyrophosphate requiring enzyme resulted to be highly induced during growth with acetone and is possibly involved in the activating reaction. Moreover, a B12- dependent enzyme and proteins that are involved in redox processes were also induced with acetone.
4.2 Introduction
It is well known that aerobic and nitrate-reducing bacteria can utilize acetone for growth through activation of acetone by a carboxylation reaction to form acetoacetate as the first
41
Chapter 4. Introduction
reaction product (Dullius et al., 2011; Schühle and Heider, 2012; Sluis et al., 1996). Under these conditions, those bacteria can gain sufficient energy from the respiration of acetone in order to sustain also the energy input required for the activation of acetone in addition to the energy required for growth. What is not fully known is how bacteria perform the pathway for utilization of acetone under strictly anoxic, sulfate-reducing conditions. For theoretical considerations, under sulfate-reducing conditions there appears to be insufficient energy derived from acetone respiration, in order to support growth and the endergonic activation of acetone. Dissimilatory sulfate reduction is performed by microorganisms that include bacteria and archaea of five different phylogenetic lineages (Thauer et al., 2007): The mesophilic deltaproteobacteria, the Gram-positive bacteria, the thermophilic Gram-negative bacteria, the Euryarchaeota, and theThermodesulfobiaceae. Sulfate reducing microorganisms (SRM) are ubiquitous in nature, therefore, they can be found in different anoxic environments where sulfate is present, e.g., in soil of terrestrial habitats and sediments of marine and freshwater habitats. Depending on their ability to oxidize the C2-carbon unit of acetyl-CoA to CO2, they can be grouped as complete oxidizers and incomplete oxidizers. There are two known ways for the oxidization of the acetyl residues. The first one is through the tricarboxylic acid cycle, and the second one is through the carbon monoxide dehydrogenase (CODH) pathway.
The completed genomes of several SRM have readily been made available in the recent years. For example, those include the deltaproteobacterium strain NaphS2, which is able to utilize naphthalene (DiDonato et al., 2010), and the marine deltaproteobacterium Desulfobacula toluolica Tol2 that utilizes aromatic compounds (Wohlbrand et al., 2013), Desulfobacterium autotrophicum HRM2 that utilizes fatty acids (Strittmatter et al., 2009), Desulfatibacillum alkenivorans AK-01 that utilizes hexadecane (Callaghan et al., 2012), Desulfotalea psychrophila that utilizes lactate and alcohols (Rabus et al., 2004), as well as of Desulfotomaculum acetoxidans (Spring et al., 2009), Archaeoglobus fulgidus (Klenk et al., 1997), Desulfarculus baarsii strain 2st14T (Sun et al., 2010), and Desulfococcus oleovorans (Joint Genome Institute; project id: 4002948).
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Chapter 4. Introduction
Up to date only two sulfate-reducing, acetone-utilizing organisms are known, one is Desulfobacterium cetonicum (Janssen and Schink, 1995b), and the second is the Gram- negative deltaproteobacterium Desulfocccus biacutus strain KMRActS (Platen et al., 1990) which has been isolated from an anaerobic sludge digestor of a waste water treatment plant. It has been proposed that these bacteria activate acetone in a carbonylation reaction to form acetoacetaldehyde in a CO- and ATP- dependent reaction (Gutierrez Acosta et al., 2013b). However, the reaction has not yet been completely understood, and the corresponding enzyme(s) remained undefined. Here, we present the draft genome sequence and annotation of the first acetone-degrading sulfate-reducing bacterium, deltaproteobacterium Desulfocccus biacutus KMRActS. The annotated genome information was used in a differential proteomics approach for the identification of genes that are specifically expressed in D. biacutus during growth with acetone, in comparison to growth with butyrate.
4.3 Materials and Methods
Bacterial growth conditions
The sulfate-reducing bacterium Desulfococcus biacutus strain KMRActS was grown in freshwater mineral medium (Gutierrez Acosta et al., 2013b; Janssen and Schnik, 1995). The medium was reduced with a sulfide solution, buffered with bicarbonate, and adjusted to a final pH of 7.2. The strain was grown in 1 L flasks with medium supplemented with 5 mM acetone or 5 mM butyrate as sole carbon source, and 10 mM sulfate as the electron acceptor. The cultures were incubated under a strictly anoxic gas phase N2/CO2 (80/20) at 30°C in the dark.
DNA extraction
The DNA was extracted using the CTAB Protocol for bacterial DNA extraction of the DoE-JGI, (http://my.jgi.doe.gov/general/index.html). The DNA pellet was re- suspended in sterile water containing RNAse (99µL water + 1 µL RNAse (10 mg mL-1).
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Chapter 4. Materials and Methods
Cell-free extract preparation
Cells of D. biacutus were harvested in the late exponential growth phase at an optical density of 0.3 (OD 600). Cells were centrifuged at 6,000 x g at 4°C and the pellet was washed at least three times with 30 mM Tris-HCl buffer, pH 7.2, supplemented with 0.5 mg DNase mL-1 and 1 mg mL-1 of complete protease inhibitor cocktail (Complete Mini, EDTA-free protease inhibitor cocktail tablets, Roche Diagnostics GmbH, Mannheim, Germany). Cells were disrupted by passing them through a cooled French pressure cell at 100 MPa. Cell debris and un-opened cells were removed by centrifugation at 27, 000 X g for 20 min at 4°C. Cell-free extract of soluble and membrane fractions were obtained by ultracentrifugation at 50, 000 X g for 60 min.
SDS-polyacrylamide gel electrophoresis (SDS-PAGE)
Two-dimensional SDS-PAGE was performed using the BIO-RAD/IEF System. The first dimension was done by focusing the soluble protein according to its isoelectric point. Samples were pretreated for removal of salts by passing freshly prepared soluble protein by gravity through a SephadexTM G-25 column (illustra NAP-25 Columns; GE Healthcare). Ultrapure water (Milli Q; Millipore System) was used for column equilibration and protein elution. After desalting, samples of 1 mg protein were precipitated overnight at -20°C by adding 5 volumes of ice-cold acetone. Protein was then collected by centrifugation (13,000 x g, 10 min, 4°C), and the protein pellet was left to dry under a hood for 30 min. Previous to the first separation, the protein pellet was rehydrated by addition of 300 µL rehydration buffer (ReadyStripTM IPG Strip Instruction Manual, BIO-RAD). The solubilized protein was loaded into a rehydration-equilibration tray (PROTEAN® IEF focusing tray) as described in the IPG Strip Instruction Manual. Accordingly, an IPG strip (17 cm length, linear pH gradient range 5 - 8) was placed gel side down onto the protein sample, and stored overnight at room temperature to allow all the protein to load into the IPG strip, and for its rehydration. Finally, the first electrophoresis could be done using a PROTEAN® IEF Cell (BIO-RAD). Isoelectric focusing started with a maximal current of 50 µA per strip at 20°C for 1 h with a maximal voltage of 1000 V (desalting), followed by a voltage ramp (rapid) to a maximal voltage of
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Chapter 4. Materials and Methods
10,000 V during 3 h, and an additional focusing at 10,000 V until a total of 40,000 Volt- hours (Vh) was reached. Afterwards, each strip was equilibrated in SDS-equilibration buffers I and II (with DTT and iodoacetamide, respectively) as described in the ReadyStripTM IPG Strip Instruction Manual. Once the strips were equilibrated, they were placed onto an SDS-PAGE gel for the second separation. Strips were covered with an overlay of Tris-Glycin-SDS buffer solidified with agarose (0.5 %) before the second electrophoresis run. The second separation was done on a gradient SDS-PAGE gel according to Laemmli (Laemmli, 1970). Gradient gels contained 5 - 18% polyacrylamide in the resolving gel and 4% polyacrylamide in the stacking gel, and were cast as large gels (17 cm x 20 cm, Protean II xi, BIO-RAD). The second electrophoresis was running at 15 mA until the front entered the resolving gel, and further at 25 mA per gel during 15 h. The gel chamber was connected to a cooling system to keep the electrophoresis run at 8°C. Gels were stained with colloidal Coomassie blue.
Membrane proteins were obtained as described in the cells extract preparation section
(see above), and were solubilized with 200 mM NaH2PO4 buffer, pH 6.0, containing 150 mM NaCl and 10% SDS (w/v). Protein samples were mixed 1:2 with loading buffer (0.0625 M Tris-HCl, pH 6.8, 2% (w/v) SDS, 25% glycerol (v/v), 0.01% (w/v) bromophenol blue and 5% β-mercaptoethanol) and heated at 100°C for 5 min prior to loading. The full electrophoresis procedure was applied to protein of acetone-grown-cells and compared with that of butyrate-grown-cells. Protein bands were excised to be analyzed by peptide fingerprinting-mass spectrometry.
Peptide fingerprinting-mass spectrometry and database searching
Protein bands (or spots) of interest were excised from gels and submitted to peptide- fingerprinting-mass spectrometry at the Proteomics Facility of the University of Konstanz (www.proteomics-facility.uni-konstanz.de) to identify the corresponding genes. The MASCOT engine (Matrix Science, London, UK) was used to match each peptide fingerprint against an IMG database of all predicted protein sequences of the annotated genomes of Desulfococcus biacutus strain KMRAcS (IMG database, Progenus database; GATC biotech.). Mass fingerprints were also matched against the external EMBL and
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Chapter 4. Materials and Methods
NCBI databases. Our standard parameters for searching and scoring were set as follows: One missed cleavage site allowed. Fixed modifications: carbamidomethyl Cys. Variable modification: N-term. pyro-Glu, N-term. Gln, Met-oxidation. Peptide charge: 2+, 3+, 4+. Peptide tolerance: 1.0 Da. MS/MS tolerance: 0.8 Da. If not stated otherwise (see Results), a minimal score of 200 and/or minimal sequence coverage of 30% was set as cut-off for low-scoring hits.
4.4 Results
Draft-genome sequencing and annotation
Total genomic DNA of Desulfocccus biacutus KMRActS was extracted using the CTAB Protocol for bacterial DNA extraction of the DoE-JGI (See materials and methods) and submitted to whole-genome shotgun sequencing using the Illumina technology (GATC, Konstanz). Sequencing and assembly resulted in a genome sequence with a total size of 5.2 Mb distributed over 159 individual contigs. From the annotation of the contigs via the IMG pipeline of the DoE-JGI, a total of 4773 genes were predicted (Table 1). Of these, 98.64% are protein-coding genes, and 73.75% are genes that code for proteins with predicted functions; the genes attributed to encode transmembrane proteins represent 23.63% of the total genes (Table 1).
Table 1. Genome statistics from the IMG annotation
Number % of Total DNA, total number of bases 5242029 100.00% DNA coding number of bases 4646037 88.63% DNA G+C number of bases 3055509 58.29% 1
DNA scaffolds 159 100.00% CRISPR Count 8
Genes total number 4773 100.00%
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Chapter 4. Results
Protein coding genes 4708 98.64% RNA genes 65 1.36% rRNA genes 3 0.06% 5S rRNA 1 0.02% 16S rRNA 1 0.02% 23S rRNA 1 0.02% tRNA genes 51 1.07% Other RNA genes 11 0.23% Protein coding genes with function prediction 3520 73.75% without function prediction 1188 24.89% not connected to SwissProt Protein Product 4708 98.64% Protein coding genes with enzymes 1028 21.54% w/o enzymes but with candidate KO based enzymes 136 2.85% Protein coding genes connected to Transporter Classification 606 12.70% Protein coding genes connected to KEGG pathways 1131 23.70% not connected to KEGG pathways 3577 74.94% Protein coding genes connected to KEGG Orthology (KO) 2033 42.59% not connected to KEGG Orthology (KO) 2675 56.04% Protein coding genes connected to MetaCyc pathways 988 20.70% not connected to MetaCyc pathways 3720 77.94% Protein coding genes with COGs3 3512 73.58% with KOGs3 1486 31.13% with Pfam3 3707 77.67% with TIGRfam3 1307 27.38% with IMG Terms 724 15.17% with IMG Pathways 254 5.32% with IMG Parts List 228 4.78% in paralog clusters 1410 29.54% in Chromosomal Cassette 4773 100.00% Number of Chromosomal Cassettes 543 - Fused Protein coding genes 24 0.50% Protein coding genes coding signal peptides 243 5.09% Protein coding genes coding transmembrane proteins 1128 23.63%
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Chapter 4. Results
COG clusters 1738 0.01% KOG clusters 708 0.00% Pfam clusters 2005 0.01% TIGRfam clusters 1021 0.01%
The genome of D. biacutus was also annotated with the Progenus pipeline (Progenus SA. GATC, Konstanz), and the inferred protein EC number distribution is shown in Table 2. The distribution of the protein EC number is obtained for the top-level and subclass EC numbers.
Table 2. Top level EC number occurrences
Oxidoreductases (1) 291 Transferases (2) 517 Hydrolases (3) 315 Lyases (4) 93 Isomerases (5) 62 Ligases (6) 100
Proteomic approach
The annotated draft-genome sequence of D. biacutus was used to generate a reference database for peptide fingerprinting-mass spectrometry (PF-MS), in order to allow for the identification of proteins that are specifically expressed in D. biacutus cells during growth with acetone in comparison to cells grown with butyrate. The analysis of the protein expression patterns was done in a gel-based approach, by 1D- and 2D-polyacrylamide gel electrophoresis (PAGE), where the soluble proteins and the membrane proteins were analyzed separately. Therefore, the cells were disrupted, the cell-debris removed by centrifugation, and the membrane fragments collected by ultra-centrifugation; the obtained supernatant represented the soluble protein fraction. Soluble proteins were
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Chapter 4. Results
separated by 2D-PAGE (Fig. 1) and membrane proteins were solubilized with SDS and separated by 1D-PAGE (Fig. 2). Protein bands or spots of interest were excised and submitted to PF-MS. The obtained identifications for all analyzed soluble proteins are shown in Table 4, and these for the analyzed membrane proteins in Table 5. The spots /bands excised and identified by peptide fingerprinting-mass spectrometry were sorted into proteins that appeared either exclusively in extracts of acetone-grown cells, and in proteins that appeared to be common to acetone- and butyrate-grown cell extracts (excised from the acetone-gel). In total, 36 spots were excised (as indicated in Fig. 1AB) and identified by PF-MS (Table 4).
Identified soluble proteins that appeared to be specifically expressed during growth with acetone
Several genes identified for spots of apparently acetone-inducible, soluble proteins were found to be located in the same gene cluster (Table 3). The protein spots were identified as follows.
The prominent spot AS_3 validly identified the locus tag DebiaDRAFT_04566, which is annotated to encode a thiamine pyrophosphate (TPP)-requiring enzyme (COG0028) [acetolactate synthase / pyruvate dehydrogenase (cytochrome) / glyoxylate carboligase / phosphonopyruvate decarboxylase]. This putative enzyme catalyzes with TPP the cleavage/fusion of substrates at a carbon-carbon bond that connects a carbonyl group to an adjacent reactive group. The protein sequence shows homology to a predicted TPP binding domain protein (Dvul_0101) of Desulfovibrio vulgaris DP4, and to predicted TPP binding domain protein (Dole_1326) in Desulfococcus oleovorans. In D. vulgaris DP4, the gene it is located in a cluster with genes that encode for an aldehyde dehydrogenase (Dvul_0100), FAD- dependent pyridine-nucleotide-disulfide oxidoreductase (Dvul_0102), and for a ferredoxin-dependent glutamate synthase (Dvul_0104). In Desulfococcus oleovorans, the gene is encoded next to a gene for an alcohol dehydrogenase GroES domain protein (Dole_ 1324).
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Chapter 4. Results
Two other genes that are clustered with the TPP-enzyme (Scheme 1) gene (DebiaDRAFT_04566) were attributed to other proteins that appeared to be specifically induced in acetone- grown cells. One of these is DebiaDRAFT_04571, attributed to spot AS_17, annotated to encode a protein of the family of short-chain dehydrogenases/ reductases with different specificities. These types of dehydrogenases possess at least 2 domains. The first binds to NAD+ or NADP+, and the second determines the substrate specificity and catalysis. Another identified gene in the cluster is DebiaDRAFT_04573, for spot AS_25, which is annotated to encode a C-terminal-domain/subunit (cobalamin- binding) of a methylmalonyl-CoA mutase enzyme complex. This enzyme complex performs different types of reaction at the cobalt-carbon bond, such as the interconversion of methyl-malonyl-CoA to succinyl-CoA. Importantly, the corresponding N-terminal domain (large subunit) of the methylmalonyl-CoA mutase is encoded directly upstream (DebiaDRAFT_04574) and was co-identified as an apparently acetone- inducible protein, represented by spot AS_ 2.
Scheme 1. Gene cluster that is encoded in D. biacutus which involves genes that are up-regulated during the growth on acetone. The corresponding spot numbers of the acetone-induced proteins are marked in red.
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Chapter 4. Results
A further acetone-inducible protein, spot AS_12, was identified as DebiaDRAFT_04514, predicted as represent a Zn-dependent dehydrogenases. In the direct proximity of this gene, another gene, DebiaDRAFT_04510, was attributed by acetone-inducible spot AS_19, annotated to encode a ‘putative redox-active protein’ (PF09719). Notably, this gene cluster appears not to be conserved in any other bacterial genome available in the IMG database; downstream are two genes for benzoyl-CoA reductase subunits (Scheme 2).
A further prominent spot, AS_32, validly identified DebiaDRAFT_01796, predicted as desulfoferredoxin, and was detected only during the growth with acetone. The protein functions by interacting non-covalently with iron (Fe) ions, and has oxidoreductase activity. The gene is encoded in a cluster that contains genes that encode for cytochrome bd-type quinol oxidase subunit 1 (DebiaDRAFT_01799), cytochrome d oxidase subunit 2 (cydB) (DebiaDRAFT_01800), multimeric flavodoxin subunit WrbA (DebiaDRAFT_01801), multimeric flavodoxin subunit WrbA (DebiaDRAFT_01801), rubredoxin (DebiaDRAFT_01802), for an uncharacterized flavoprotein (DebiaDRAFT_01803), ferredoxin (DebiaDRAFT_01804), and DsrE/DsrF-like family (DebiaDRAFT_01805). This complex is similar to the one found in Desulfovibrio alaskensis G20 which also contains desulfoferrodoxin. This protein contains two types of iron: an Fe4-S4 site very similar to that found in desulfoferrodoxin of Desulfovibrio gigas, and an octahedral coordinated high-spin ferrous site most probably with nitrogen/oxygen- containing ligands. From the genes belonging to this complex, only the predicted desulfoferredoxin gene was upregulated during the growth on acetone. The function of the full complex is not clear, however, it may be involved in the protection of the bacterial cells against oxidative stress by removing superoxide radicals, for example. But, the specificity for the degradation of acetone must be elucidated.
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Chapter 4. Results
Scheme 2. Second gene cluster that is encoded in D. biacutus which involves genes that are up- regulated during the growth on acetone. The corresponding spot numbers of the acetone- induced proteins are marked in red.
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Chapter 4. Results
Figure 1. 2D- PAGE separation of the proteins that are expressed during growth with (A) acetone and with (B) butyrate. The amount of protein that was used for each condition was one mg. M: molecular marker.
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Chapter 4. Results
Table 3. Neighborhood from identified cluster in D biacutus.
Neighborhood Gene Product Name COG ID COG Name Gene ID Viewer Centered on this Gene COG1028 Dehydrogenases with different specificities 2512528356 DebiaDRAFT_04571 ª Dehydrogenases with different specificities (related to short-chain alcohol (related to short-chain alcohol dehydrogenases) dehydrogenases)
COG1703 Putative periplasmic protein kinase ArgK 2512528357 DebiaDRAFT_04572 LAO/AO transport system ATPase and related GTPases of G3E family
COG2185 Methylmalonyl-CoA mutase, C-terminal 2512528358 DebiaDRAFT_04573 ª Methylmalonyl-CoA mutase C-terminal domain/subunit (cobalamin-binding) domain
COG1884 Methylmalonyl-CoA mutase, N-terminal 2512528359 DebiaDRAFT_04574 ª Methylmalonyl-CoA mutase, N-terminal domain/subunit domain/subunit
COG1180 Pyruvate-formate lyase-activating enzyme 2512528349 DebiaDRAFT_04564 Pyruvate-formate lyase-activating enzyme
COG0028 Thiamine pyrophosphate-requiring 2512528351 DebiaDRAFT_04566 ª Thiamine pyrophosphate-requiring enzymes [acetolactate synthase, pyruvate enzymes [acetolactate synthase, pyruvate
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Chapter 4. Results
dehydrogenase (cytochrome), glyoxylate dehydrogenase (cytochrome), glyoxylate carboligase, phosphonopyruvate carboligase, phosphonopyruvate decarboxylase] decarboxylase]
COG1309 Transcriptional regulator 2512528348 DebiaDRAFT_04563 Transcriptional regulator
COG0491 Zn-dependent hydrolases, including 2512528354 DebiaDRAFT_04569 Zn-dependent hydrolases, including glyoxylases glyoxylases
COG0491 Zn-dependent hydrolases, including 2512528355 DebiaDRAFT_04570 Zn-dependent hydrolases, including glyoxylases glyoxylases
ª Proteins that were found to be specific induced during the growth in acetone.
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Table 4. Proteins identified by peptide fingerprinting-mass spectrometry from protein spots excised from 2D-gels of soluble proteins in D. biacutus (see Fig. 1). Distributed as acetone-induced proteins, and proteins found in both, acetone and butyrate grown cells. The identification was done using the annotated genome of D. biacutus.
Spot Obs. Obs. IMG Locus tag Annotation name Predic. Predic. Score Cov. Id. mass pI Mass pI (kDa) (KDa) (%)
Identified protein spots that were observed specifically in acetone-grown cells:
AS_1 170 8 DebiaDRAFT_02387 Formyltetrahydrofolate synthetase 64.155 7.93 1045 55
AS_3 70 7 DebiaDRAFT_04566 Thiamine pyrophosphate-requiring enzymes 77.949 6.18 652 40%
AS_4 70 7.5 DebiaDRAFT_03619 Adenosine phosphosulphate reductase, alpha subunit 74.097 6.67 1595 51%
AS_12 40 6.6 DebiaDRAFT_04514 Threonine dehydrogenase and related Zn-dependent 38.760 5.76 970 61% dehydrogenases
DebiaDRAFT_04509 Acetyl-CoA acetyltransferases 42.070 5.85 200 31%
AS_17 23 6.6 DebiaDRAFT_04571 Dehydrogenases with different specificities (related to short- 28.158 6.01 1944 81%
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Chapter 4. Results
chain alcohol dehydrogenases)
AS_19 20 5.7 DebiaDRAFT_04510 Putative redox-active protein (C_GCAxxG_C_C) 21.016 5.28 1035 75%
AS_25 15 6.3 DebiaDRAFT_04573 Methylmalonyl-CoA mutase C-terminal domain 15.081 5.53 1315 77%
AS_32 13 6.7 DebiaDRAFT_01796 Desulfoferrodoxin 13.812 6.07 449 79%
Identified protein spots that were observed in both, acetone and butyrate-grown cells:
AS_2 70 5.4 DebiaDRAFT_03619 Adenosine phosphosulphate reductase, alpha subunit 74.097 6.67 576 40%
DebiaDRAFT_04566 Thiamine pyrophosphate-requiring enzymes 77.949 6.18 364 30%
DebiaDRAFT_04574 Methylmalonyl-CoA mutase, N-terminal domain/subunit 65.254 5.09 209 35%
AS_5 60 8 DebiaDRAFT_02387 Formyltetrahydrofolate synthetase 64.155 7.93 1027 68%
DebiaDRAFT_03619 Adenosine phosphosulphate reductase, alpha subunit 74.097 6.67 343 24%
DebiaDRAFT_03447 NAD(P)H-nitrite reductase 61.335 8.32 314 26%
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Chapter 4. Results
AS_6 50 6.7 DebiaDRAFT_03586 ATP sulphurylase 47.262 6.29 1338 55%
AS_7 45 7 DebiaDRAFT_03586 ATP sulphurylase 47.262 6.29 1454 60%
AS_8 45 5.5 DebiaDRAFT_04385 Sulfite reductase, dissimilatory-type alpha subunit 50.255 5.20 814 75%
AS_9 38 5.4 DebiaDRAFT_00156 ABC-type branched-chain amino acid transport systems, 42.533 5.49 1367 56% periplasmic component
AS_10 40 5.9 DebiaDRAFT_02798 ABC-type branched-chain amino acid transport systems, 40.870 6.36 1844 58% periplasmic component
AS_11 40 6.1 DebiaDRAFT_01292 ABC-type branched-chain amino acid transport systems, 40.412 6.20 1514 74% periplasmic component
DebiaDRAFT_03292 Acyl-CoA dehydrogenases 42.984 5.54 326 35%
AS_13 40 7.7 DebiaDRAFT_04384 sulfite reductase, dissimilatory-type beta subunit 43.581 6.82 682 59%
AS_14 33 5.4 DebiaDRAFT_01640 Pterin binding enzyme 34.293 4.83 771 58%
DebiaDRAFT_04718 Electron transfer flavoprotein, alpha subunit 33.377 4.83 138 36%
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Chapter 4. Results
AS_15 33 6 DebiaDRAFT_02722 ABC-type amino acid transport/signal transduction systems, 36.649 6.63 1168 78% periplasmic component/domain
DebiaDRAFT_01640 Pterin binding enzyme. 34.293 4.83 191 30%
AS_16 33 7 DebiaDRAFT_00168 Malate dehydrogenase, NAD-dependent 32.746 6.15 1411 80%
DebiaDRAFT_02722 ABC-type amino acid transport/signal transduction systems, 36.649 6.63 245 54% periplasmic component/domain
AS_18 25 5.5 DebiaDRAFT_04513 Enoyl-CoA hydratase/carnithine racemase 27.721 5.05 847 81%
DebiaDRAFT_04490 ABC-type amino acid transport/signal transduction systems, 31.051 6.00 730 65% periplasmic component/domain
DebiaDRAFT_04571 Dehydrogenases with different specificities (related to short- 28.158 6.01 518 55% chain alcohol dehydrogenases)
DebiaDRAFT_01784 Enoyl-CoA hydratase/carnithine racemase 27.681 5.15 467 57%
DebiaDRAFT_03805 Short-chain dehydrogenases of various substrate specificities 28.885 5.22 417 66%
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Chapter 4. Results
AS_20 24 5.8 DebiaDRAFT_02892 ABC-type amino acid transport/signal transduction systems, 26.762 6.63 653 56% periplasmic component/domain
DebiaDRAFT_04510 Putative redox-active protein (C_GCAxxG_C_C) 21.016 5.28 403 46%
AS_21 23 6.2 DebiaDRAFT_02997 Molecular chaperone (small heat shock protein) 21.430 5.53 582 70%
AS_22 17 5.7 DebiaDRAFT_02783 Peroxiredoxin 18.629 5.28 942 78%
AS_23 15 5.4 DebiaDRAFT_03620 Adenosine phosphosulphate reductase, beta subunit 16.689 4.88 535 83%
AS_24 15 5.7 DebiaDRAFT_03620 Adenosine phosphosulphate reductase, beta subunit 16.689 4.88 340 79%
AS_26 17 6.5 DebiaDRAFT_04190 Rubrerythrin 19.936 5.80 832 64%
DebiaDRAFT_04573 Methylmalonyl-CoA mutase C-terminal domain 15.081 5.53 416 47%
AS_27 20 7.1 DebiaDRAFT_03710 Nitroreductase 19.975 6.31 471 50%
DebiaDRAFT_04190 Rubrerythrin 19.936 5.80 333 48%
AS_28 20 7.6 DebiaDRAFT_00527 Putative methyltransferase, YaeB/AF_0241 family 20.571 6.92 445 42%
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Chapter 4. Results
AS_29 14 5.4 DebiaDRAFT_04565 Hypothetical protein 14.555 4.94 260 76%
DebiaDRAFT_01155 Response regulator containing CheY-like receiver domain 13.673 4.73 235 37% and AraC-type DNA-binding domain
AS_30 13 5.4 DebiaDRAFT_04571 Dehydrogenases with different specificities (related to short- 28.158 6.01 137 12% chain alcohol dehydrogenases)
AS_31 13.5 6.1 DebiaDRAFT_01099 Uncharacterized conserved protein
AS_33 12 8 DebiaDRAFT_01088 RNA-binding proteins (RRM domain) 9.962 9.26 263 47%
DebiaDRAFT_01796 Desulfoferrodoxin 13.812 6.07 176 20%
AS_34 9 5.6 DebiaDRAFT_03397 Cold shock proteins 7.181 5.18 279 68%
AS_35 8 5.6 DebiaDRAFT_03129 Uncharacterized conserved protein 7.726 6.23 617 50%
AS_36 7 6.7 DebiaDRAFT_03129 Uncharacterized conserved protein 7.726 6.23 635 58%
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Table 5. Proteins identified by peptide fingerprinting-mass spectrometry from protein spots excised from 1D-gels of solubilized membrane fraction proteins in D. biacutus (see Fig. 2). Distributed as acetone-induced proteins, and proteins found in both, acetone and butyrate grown cells. The identification was done using the annotated genome of D. biacutus.
Spot Obs. IMG Locus tag Annotation name Predic. Score Id. mass Mass (kDa) (KDa)
Identified protein spots that were observed specifically in acetone-grown cells:
AM_2 70 DebiaDRAFT_04566 Thiamine pyrophosphate-requiring enzymes 77.949 282
AM_10 25 DebiaDRAFT_03042 Predicted NADH:ubiquinone oxidoreductase, subunit 24.120 159 RnfG
Identified protein spots that were observed in both, acetone and butyrate-grown cells:
Pyruvate:ferredoxin (flavodoxin) oxidoreductase, AM_O 135 DebiaDRAFT_04339 132.673 335 homodimeric CO dehydrogenase/CO-methylating acetyl-CoA synthase AM_1 120 DebiaDRAFT_01638 81.809 588 complex, beta subunit
AM_4 55 DebiaDRAFT_03347 Proton translocating ATP synthase, F1 alpha subunit 54.965 811
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Chapter 4. Results
AM_5 50 DebiaDRAFT_03345 ATP synthase, F1 beta subunit 51.052 1146
AM_6 48 DebiaDRAFT_00583 Hypothetical protein 48.300 700
AM_7 38 DebiaDRAFT_03345 ATP synthase, F1 beta subunit 51.052 204
AM_9 36 DebiaDRAFT_01843 5,10-methylenetetrahydrofolate reductase 34.998 155
AM_11 18 DebiaDRAFT_03348 ATP synthase, F1 delta subunit 20.265 607
AM_12 14 DebiaDRAFT_03344 ATP synthase, F1 epsilon subunit (delta in mitochondria) 15.117 447
AM_13 11 DebiaDRAFT_03135 Bacterial nucleoid DNA-binding protein 9.797 181
AM_14 10 DebiaDRAFT_04571 Dehydrogenases with different specificities (related to short- 28.158 127 chain alcohol dehydrogenases)
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Chapter 4. Results
Identified membrane proteins that appeared to be specifically expressed during growth with acetone
Among the membrane proteins, band AM_2 appeared as an acetone-induced protein and was attributed to DebiaDRAFT_04566, which is the TPP-requiring enzyme family protein that was also found among the acetone-induced proteins in the soluble protein fraction (see above). Notably, the generation of the membrane fragments during the cell preparation involved four washing steps. Hence, it is unlikely that the washing protocol to separate the membrane fraction from soluble fraction was not sufficient. Moreover, from comparing the 2D and 1D -PAGE analyses, we conclude that the TPP-requiring enzyme was represented in rather low amounts in the soluble protein fraction, compared to the prominent protein band observed in the membrane fraction. Indeed, in the membrane fraction it is most clearly visible that this protein is specifically induced during growth with acetone. Thus, it seems as if this protein is associated to the membrane. The band AM_1 was identified as the beta subunit of CO dehydrogenase/CO- methylating acetyl-CoA synthase complex (DebiaDRAFT_01638). This protein was visible also for butyrate-grown cells, but seemed to be increasingly expressed during growth on acetone. This subunit is encoded in a gene cluster together with the CO dehydrogenase/acetyl-CoA synthase delta subunit (corrinoid Fe-S protein) (DebiaDRAFT_01636), the carbon-monoxide dehydrogenase catalytic subunit (DebiaDRAFT_01637), the CO dehydrogenase/acetyl-CoA synthase gamma subunit (corrinoid Fe-S protein) (DebiaDRAFT_01639), pterin binding enzyme (DebiaDRAFT_01640), phosphoenolpyruvate synthase/pyruvate phosphate dikinase (DebiaDRAFT_01642), and glyceraldehyde-3-phosphate dehydrogenase/erythrose-4- phosphate dehydrogenase (DebiaDRAFT_01643).
An Acetyl-CoA carboxylase, carboxyltransferase component (alpha and beta subunits), spot AM_3, (DebiaDRAFT_00010) was visible also for butyrate-grown cells, but seemed to be increasingly expressed during growth on acetone. The spot AM_0, identified as pyruvate: ferredoxin (flavodoxin) oxidoreductase, homodimeric, could be slightly seen in the acetone induced proteins
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Chapter 4. Results
(DebiaDRAFT_04339). Interestingly, a predicted NADH:ubiquinone oxidoreductase subunit RnfG is upregulated during growth with acetone, band AM_10, DebiaDRAFT_03042.
Figure 2. SDS-PAGE separation of the acetone and butyrate membrane proteins. Linie 1, acetone cell extract (25 µg); Linie 2, acetone cell extract (50 µg); Linie 3, butyrate soluble protein (75 µg); Line 4, acetone soluble protein (75 µg); Line 5, butyrate membrane protein (75 µg); Line 6, acetone membrane protein (75 µg). M, molecular protein marker.
Identified proteins attributed to sulfate reduction and ATP synthesis
For the soluble protein fraction, the prominent, apparently acetone-inducible spot AS_4 validly identified DebiaDRAFT_03619, which is predicted to encode the alpha-subunit of adenosine phosphosulphate reductase (AprA). This gene is located in a cluster with adenosine phosphosulphate reductase beta-subunit gene (DebiaDRAFT _03620), and also this gene was identified, for both acetone and butyrate grown cells (Spots AS_23 and
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Chapter 4. Results
AS_24). The gene cluster contains also the gene for the subunit A of heterodisulfide reductase and related polyferredoxins (HdrA) (DebiaDRAFT _03617). Similar gene cluster can be found in the genomes of Desulfovibrio vulgaris DP4, Desulfovibrio vulgaris Hildenborough, and Desulfococcus oleovorans Hxd3.
The spot AS_8 identified DebiaDRAFT_04385, which most likely encodes a dissimilatory-type sulfite reductase alpha-subunit gene. The gene is co-located in a cluster with the gene for dissimilatory-type sulfite reductase beta–subunit (DebiaDRAFT_04384), which was also identified by PF-MS (spot AS_13). In the same cluster are genes located that code for dissimilatory sulfite reductase D (DebiaDRAFT _04383) and for NADH: flavin oxidoreductases of the Old Yellow Enzyme family (DebiaDRAFT_04379). This cluster is also found in Desulfococcus oleovorans Hxd3, and Desulfovibrio vulgaris Hildenborough, and DP4 strains, and in Desulfovibrio alaskensis G20.
For the membrane proteins, all subunits of ATP synthase were found in both, acetone and butyrate cells, as follows: band AM_4, DebiaDRAFT_03347, proton translocating ATP synthase, F1 alpha subunit; bands AM_5 and AM_7, DebiaDRAFT_03345, ATP synthase, F1 beta subunit; band AM_8, DebiaDRAFT_03346 ATP synthase, F1 gamma subunit; band AM_11, DebiaDRAFT_03348 ATP synthase, F1 delta subunit; band AM_12, DebiaDRAFT_03344 ATP synthase, F1 epsilon subunit. All genes belonging to this ATP synthase are arranged in a gene cluster within the genome of D. biacutus.
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Chapter 4. Discussion
4.5 Discussion
The degradation pathway for acetone in sulfate-reducing bacteria is of great interest due to energetic considerations, as described in the introduction. In this study, we established a draft genome sequencing of the acetone-utilizing sulfate reducer Desulfococcus biacutus, as an important tool to investigate its genes and operons involved in the metabolism of acetone and some other compounds of interest, as well as to identify and compare complexes that are possibly involved in its energy conservation. We furthermore evaluated its proteome in respect to the degradation of acetone, by using proteomic analyses in combination with the established draft genome sequence. The results from the proteomic analysis support the notion that Desulfococcus biacutus, uses a different mechanism for the activation of acetone, rather than carboxylation as observed in aerobic or nitrate reducing condition (see Introduction). The survey of protein expressed during growth with acetone gave no indication for an expression of an acetone carboxylase. However, in the genome of D. biacutus, there are two candidate genes for N-methylhydantoinase A / acetone carboxylase beta subunit genes (hyuA) (DebiaDRAFT_01948, DebiaDRAFT_02725), but these gene were not identified in the proteomic approach. Furthermore, the alpha (acxB) and gamma (acxC) subunits of N- methylhydantoinase A / acetone carboxylase were not found in the genome of Desulfococcus biacutus. Furthermore, the putative beta subunit gene (hyuA) of a possible acetone carboxylase in D. biacutus, is localized in a gene cluster together with ABC aminoacid transporter genes. Hence, the metabolic function of this subunit (hyuA) is more likely that of a hydantoinase that takes part in the production of L- or D- aminoacids, rather than being involved in the activation of acetone in D. biacutus.
The detection of a thiamine pyrophosphate (TPP)-requiring enzyme that was found only in cell extracts of acetone-grown cells, suggests that TPP might be involved as a cofactor in an enzymatic reaction that occurs only when D. biacutus grows with acetone as carbon source. It has been shown that TPP participates in carbon-carbon bond-forming or bond-breaking reactions, especially for those substrates that contain a carbonyl group,
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Chapter 4. Discussion
such as in acetone (Kluger, 1987; Kochetov and Izotova, 1972; Kondo and Ishimoto, 1974; Lindqvist et al., 1992; Pierce et al., 2010; Reed, 1974). For the acetone-activating mechanism that has been proposed before for D. biacutus, acetone is possibly carbonylated to form an aldehyde derivative (acetoacetaldehyde) in an ATP- dependent reaction (Gutierrez Acosta et al., 2013b). For this novel activating mechanism, the involvement of a TPP-requiring enzyme could be a suitable way to keep the highly reactive aldehyde in a safety position, since its release could cause severe damages to the bacterial cell. Preliminary experiments with cell extracts of acetone-grown cells of D. biacutus demonstrated that the addition of TPP enhances the hydrolysis rate of an ATP analogue in the presence of acetone and CO (Gutierrez Acosta et al., 2013a). It is, however, not clear whether a radical reaction is involved in the mechanism to generate the aldehyde, or if TPP perhaps serves only to stabilize the aldehyde and make it ready for the next enzyme reaction. In this case, it might be possible that a TPP-aldehyde intermediate is formed in a way so that CoASH can attack the intermediate to form acetoacetyl-CoA. The genes co-encoded with the gene for the TPP-binding enzyme in D. biacutus and in other sulfate-reducing deltaproteobacteria like D. vulgaris and D. oleovorans, i.e., aldehyde and/or alcohol dehydrogenase genes, also support the notion of possible role of these genes in reactions involving aldehydes. Importantly, one of the genes in the clusters with the TPP-enzyme gene (DebiaDRAFT_04566), dehydrogenase DebiaDRAFT_04571 that possesses a NAD+ binding domain, was found to be specifically induced during growth with acetone. This enzyme might represent a NAD+- dependent aldehyde dehydrogenase, responsible for the conversion of the aldehyde formed during the activation of acetone in the already mentioned mechanism (Gutierrez Acosta et al., 2013b).
Interesting is also the observation that in the TPP-enzyme gene cluster, there is a gene coding for a B12-binding domain protein (DebiaDRAFT_04573), and that this proteins was also observed solely in extracts of acetone-grown cells. Hence, the possibility of a cobalamin-dependent reaction cannot be ruled out. Interestingly, preliminary experiments showed an increase of the Co2+ signal in the presence of acetone, CO and
CoB12 (adenosylcobalamin), in cell-free extracts (Fig. 1 and 2 supplementary data).
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Chapter 4. Discussion
Whether a radical reaction is involved in the activation mechanism is not clear yet. It could be that this activity was due to a reaction with the corrinoid iron-sulfur protein that is involved in the Wood-Ljundal pathway, annotated in D. biacutus as CO dehydrogenase/acetyl-CoA synthase gamma subunit, corrinoid Fe-S protein (DebiaDRAFT_01637). Furthermore, cobalamin-dependent reactions occur at the lower reduction potential in the cell, especially in the reduction of Co2+ to Co+ (E0´ = -523 mV). In order couple a reaction with cobalamine the presence of another system that allows electron transfer is required. Within the genome of D. biacutus, an Rnf system operon is present (subunits RnfABCDGE). In fact, among the proteins from the membrane fraction, an acetone-inducible protein was detected and identified as predicted NADH: ubiquinone oxidoreductase subunit RnfG (DebiaDRAFT_03042). However, its possible participation in energy conservation in D. biacutus needs to be clarified.
Acknowledgements
We gratefully acknowledge funding by the Deutsche Forschungsgemeinschaft within the SPP 1319 priority program, and the Konstanz Research School Chemical Biology for the fellowship granted to Olga Gutiérrez.
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Chapter 5. General Discussion
CHAPTER 5
General Discussion
The present work gives an overview of the degradation of acetone by the sulfate-reducing bacterium Desulfococcus biacutus. It is well established that aerobic acetone degradation, as well as nitrate-dependent degradation of acetone, is mediated by acetone carboxylases (Dullius et al., 2011; Schühle and Heider, 2012; Sluis and Ensign, 1997; Sluis et al., 2002).
In this CO2 and ATP-dependent carboxylating mechanism an input of two ATP equivalents is required for the conversion of acetone to acetoacetate (Boyd and Ensign, 2005; Sluis et al., 1996). Acetone carboxylation energetically represents for aerobes and nitrate reducers a type of activation without energy limitation. Sulfate reducers like D. biacutus theoretically are not able to degrade acetone by the same carboxylating mechanism. Therefore, a different activation process was postulated for the activation of acetone by SRB, and is going to be described in more detail through the course of this chapter.
5.1 Activation of acetone by Desulfococcus biacutus
For investigation of the reaction mechanism that is involved in the activation of acetone by SRB, D. biacutus has been used as a model microorganism. It has been shown that acetone was degraded only if CO2 was provided in the gas phase. But, a carboxylating mechanism leading to acetoacetate could not be proven (Janssen and Schink, 1995b; Janssen and Schnik, 1995; Platen et al., 1990). As an alternative, a carbonylation reaction leading to acetoacetaldehyde was hypothesized in this work.
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Chapter 5. General Discussion
Suspension experiments with D. biacutus cells with CO or CO2 in the gas phase resulted in higher acetone degradation rates when CO was supplied. Moreover, experiments where CO dehydrogenase was inhibited by KCN as it was suggested before (Wang et al., 2013) indicated that CO is involved in the reaction as co-substrate and not as an electron donor. Further experiments in cell extract showed that the addition of KCN did not affect the degradation of acetone, because the activating reaction that was measured by quantifying carbonyl groups proceeds highly similar as without KCN. The increment in keto groups that was quantified with 2,4-DNPH in cell extracts gave an indication that acetone was converted to acetoacetaldehyde as it was hypothesized. After the reaction with acetone, CO, and ATP, the product acetoacetaldehyde could be trapped as DNPH derivative and analyzed by ESI-MS (Gutierrez Acosta et al., 2013b). This indicated that acetone and CO condensed in an ATP-dependent reaction. Coupling of the reaction with ATP hydrolysis was also proven in a derivatization reaction with guanidine to form 2- amino-4-methylpyrimidine. Since in the enzymatic reaction the pyrimidine could be formed only after a condensation reaction between guanidine and acetoacetaldehyde the presence of this metabolite as an intermediate of the activation of acetone was confirmed.
Activity of aldehyde dehydrogenase was found to be induced in acetone-grown cell extracts. This activity (153 ± 36 mU per mg of protein) was considerably higher when the putative physiological substrate acetoacetaldehyde was added to start the reaction (Gutierrez Acosta et al., 2013b). In comparison, the addition of acetaldehyde represented only a small fraction of the activity (18 ± 3 mU per mg of protein). This experiment clearly shows that in the metabolism of acetone by D. biacutus an aldehyde derivative is formed, being acetoacetaldehyde the most probable candidate for this derivative.
More evidence was provided with the detection of the CoA derivative as the further intermediate in the metabolism of acetone by D. biacutus. In a similar reaction with acetone, CO and ATP, the addition of CoA led to the formation of acetoacetyl-CoA as it was shown by ESI-MS (Gutierrez Acosta et al., 2013b).
In comparison to the activating mechanism in aerobic bacteria, the formation of acetoacetaldehyde as the product of the activation under strictly anoxic conditions would
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Chapter 5. General Discussion
be an energetically suitable way to avoid the investment of three ATP equivalents for conversion of acetone to acetoacetate and its subsequent activation to acetoacetyl-CoA. In the mechanism that is proposed here the conversion of acetoacetaldehyde to acetoacetyl-CoA does not require input of energy. Nonetheless, it is assumed that there must exist a smart and safe way to prevent the release of acetoacetaldehyde because otherwise it could be toxic for the bacterial cell. In that regard, it may be speculated that in the activating mechanism the involvement of an enzyme complex would be a good strategy for the activation of acetone and further conversion to the CoA derivative. This idea is also suggested from proteomic and genomic data.
The genome of D. biacutus that was sequenced and annotated during the course of this work supports the hypothesis that under sulfate-reducing conditions the activation of acetone does not occur through carboxylation. According to the genome annotation, D. biacutus does not have any operon similar to that which is induced in the aerobic degradation of acetone. In that case, the arrangement of the acx A, acx B, and acx C are localized next to each other (Schühle and Heider, 2012). On the contrary, in D. biacutus only a putative beta subunit of the acetone carboxylase can be found in the genome, however, is not induced during growth with acetone. Since it is likely that acetone carboxylases and hydantoinases have a common ancestor gene, it is possibly that the catalytic role of this subunit that is annotated as hydantoinase/carboxylase is more as that of a hydantoinase, therefore involved in the production of amino acids (Altenbuchner et al., 2001).
In addition, the proteins which are up-regulated during the metabolism of acetone are arranged in a cluster that is presumably specific for D. biacutus. Those proteins include a thiamine-pyrophosphate-requiring enzyme which is clustered with a NAD+ or NADP+ binding dehydrogenase and a B12-dependent protein (Table 3). It is still unclear how a TPP-binding enzyme can be involved in the degradation of acetone. Nevertheless, initial experiments indicated that the activation of acetone is stimulated by TPP, as it was shown by measuring the hydrolysis rate of an ATP analogue in cell-free extracts (Gutiérrez-Acosta, et al., 2013). At the moment, the role of TPP in enzymatic reactions is not fully understood, however, it is known that this cofactor assists in reactions that
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Chapter 5. General Discussion
occur adjacent to carbonyl groups (Ferreira et al., 2008; Mansoorabadi et al., 2006; Pierce et al., 2010) as it is the case also with our reaction system. It is conceivable that TPP is required to form a TPP-intermediate with acetone or its phosphorylated form which is afterwards carbonylated to form acetoacetaldehyde. If this is the case, this TPP-aldehyde intermediate would then allow the ligation of CoA to its further conversion to acetoacetyl-CoA. A different possibility of the reaction mechanism with TPP (after discussion with Prof. B. T. Golding) might be a direct ligation of the TPP-carbanion to CO, which then will react with phospho-enolacetone to form acetoacetaldehyde (Fig. 1). Even so, this is only an idea on the involvement of TPP in the activating mechanism that must be studied further in detail.
Figure 1. Proposed reaction mechanism of the possible role of thiamine pyrophosphate in the strictly anaerobic activation of acetone by Desulfococcus biacutus.
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Chapter 5. General Discussion
The products of ATP hydrolysis are presumably AMP +PPi (Gutiérrez and Schink, supplementary data). This may indicate that in the activation of acetone by D. biacutus indeed two ATP equivalents have to be invested, but there might also be a pyrophosphatase involved. Indeed, in the genome a proton-translocating pyrophosphatase was found to be localized. The hydrolysis of PPi is an exergonic process with a standard free energy change of -22 kJ mol-1 (Thauer et al., 1977), and therefore could be coupled to the endergonic activation of acetone. Thus, such a type of pyrophosphatase could help in energy conservation in this energy-limited microorganism.
The carbonylation of acetone could also be mediated by a B12-dependent protein, as suggested from the proteomic analysis and also from experimental results. However, changes in the spectrum of the CoB12 (adenosylcobalamine) depended on acetone and
CO, but not on ATP. Therefore, the specific role of a B12-dependent reaction is still questionable. If a B12-dependent enzyme may be involved, then the reverse of the carbonylation reaction that is proposed for D. biacutus would perhaps resemble the decarbonylation reaction of an aldehyde mediated by a cobalt- porphyrin enzyme (Dennis and Kolattukudy, 1992).
5.2 Comparison with other hydrocarbon activation
Hydrocarbons are apolar compounds with low reactivity at room temperature. In comparison to aerobic activation, little is known about the strictly anaerobic activation of hydrocarbons. However, the initial activating reaction is of particular biochemical interest due to the high energy barrier for dissociation of the C-H bond. Degradation of hydrocarbons starts with activation by introducing a functional group. As in the case of acetone activation, hydrocarbons are activated by various mechanisms, depending on the presence of oxygen (Baboshin and Golovleva, 2012; Díaz et al., 2013), nitrate (Callaghan et al., 2009; Rabus et al., 2002; Rabus and Widdel, 1995), sulfate (Aeckersberg et al., 1991;
Aeckersberg et al., 1998; Meckenstock and Mouttaki, 2011) or only CO2 as electron acceptor (Anderson and Lovley, 2000; Zengler et al., 1999). Different from acetone activation, the enzymes that are involved in hydrocarbon activation under aerobic
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Chapter 5. General Discussion
conditions are different from those that are employed by nitrate reducers. For instance, aerobic degradation of alkanes is mediated by oxygenases (mono- or di-oxygenases) which form reactive oxygen species in their active sites (via reduction of O2 to a peroxide state) in order to attack the hydrocarbon. Under nitrate-reducing conditions, as well as under sulfate-reducing conditions, it has been shown that activation of alkanes is initiated by addition to fumarate, yielding alkylsuccinates in a reaction that is analogous to toluene activation (Kropp et al., 2000; Rabus et al., 2001). The activating enzymes termed alkylsuccinyl synthase (Ass), benzylsuccinyl synthase (Bss) are quite similar to the 2- methylnaphtalene activating enzyme (Nms) which also performs addition to fumarate (Selesi et al., 2010). Those enzymes catalyze the activation through a glycyl radical formation (Eklund and Fontecave, 1999; Selmer et al., 2005).
A different activation mechanism was proposed with the sulfate-reducing bacterium Desulfococcus oleovorans Hxd3. This bacterium activates long-chain alkanes (C12-C20) via carboxylation at the C3 carbon, yielding a fatty acid that is one carbon shorter than the parent alkane, as it was shown with hexadecane. However the carboxylating mechanism is so far unknown. For this bacterium, earlier discussions suggested an energy-driven carbonylation at the C-terminal methyl group of the alkane, yielding an aldehyde as product (Aeckersberg et al., 1998), but no convincing evidence for this type of reaction was shown. Similar to that assumption, we have recently shown evidence that acetone is activated in an energy-driven carbonylation to yield an aldehyde (Gutierrez Acosta et al., 2013b) as it was already discussed above with the sulfate-reducer Desulfococcus biacutus.
In the case of the nitrate-reducing strain HdN1, a different type of alkane activation was proposed. Strain HdN1 grows with alkanes as organic substrates and uses nitrate as - electron acceptor. It has all genes for a common reduction of NO3 to N2, however, does not grow with N2O as the only electron acceptor (Zedelius et al., 2011). Experimental evidence suggested the involvement of a possible intermediate that may be formed - - during the reduction of NO3 or NO2 , thus representing a novel alkane-activating reaction in which the electron acceptor might be involved.
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Chapter 5. General Discussion
Finally, it can be summarized that the initial step of hydrocarbon activation depends on the presence of the electron acceptor, and of the type of hydrocarbon. In that regard, it can be expected that with respect to hydrocarbon activation still different reactions are to be discovered. In fact, one should consider the possibility of CO as a co-reactant, similar to acetone activation, for the initial step in the degradation of other hydrocarbons under strictly anoxic conditions. The carbonylation pathway that is proposed in this thesis would require an energy investment of less than 1 ATP equivalent and therefore represents a suitable way for bacteria which have to operate with low energy gain, as sulfate reducers do. Thus, acetone activation is another example to demonstrate that strict anaerobes such as sulfate reducers use strategies in the degradation of comparably stable compounds that are basically different from those employed by nitrate reducers, as studies with various aromatic compounds have shown in the past (Philipp and Schink, 2012).
5.3 Future research
The mechanism of activation of acetone under sulfate-reducing conditions has been better understood. However, there are several questions that need to be answer. For instance, the enzymes that are involved in the activating reaction need to be confirmed. There are several indications of the involvement of a thiamine-dependent enzyme in the activation, nevertheless the enzyme needs to be purified and further characterized. Whether an enzyme complex is taking part in the reaction needs also to be clarified.
From genomic and proteomic data a B12-dependent enzyme was found to be induced in acetone-grown cell-free extracts, but further experiments to clarify its specificity for the metabolism of acetone are required. Therefore, the possibility of a cobalamin dependent reaction remains unclear. It is also not clear which are products of the hydrolysis of ATP, because preliminary experiments showed that AMP is the product of the reaction but it has to be confirmed. It would be also interesting to know whether a proton translocating pyrophosphatase is taking part in energy conservation.
76
SUMMARY
In the context of this dissertation the activation of acetone under strictly anoxic conditions was studied. For that purpose the sulfate-reducing microorganism Desulfococcus biacutus has been used. It is concluded that the activation of acetone by D. biacutus occurs in a novel activating mechanism with CO as cosubstrate. This ATP-dependent carbonylation of acetone leads to the formation of acetoacetaldehyde as intermediate. The sequenced and annotated genome of D. biacutus that is part of this work allowed a better understanding of the activation of acetone, and it opened the doors to further investigations on this topic. For example, with genomic and proteomic analysis we found a TPP-dependent enzyme which might be involved in the activation reaction in the degradation of acetone. Moreover, initial experiments showed that TPP is a cofactor in the initial reaction as it was shown with the use of an ATP analogue. There is also the possibility that a B12 enzyme participates in the acetone degradation pathway. Finally, we conclude that strictly anaerobic activation of ketones and hydrocarbons differs highly from the activation in the presence of oxygen, and that the discovery of this novel acetone activation might help in further investigations of hydrocarbon activation.
77
ZUSAMMENFASSUNG
Im Rahmen dieser Arbeit wurde die Aktivierung von Aceton unter strikt anoxischen Bedingungen untersucht. Für diese Studien wurde der sulfat-reduzierende Mikroorganismus Desulfococcus biacutus verwendet. Es wurde bestätigt, dass die Aktivierung von Aceton durch D. biacutus nach einem neuartigen Aktivierungsmechanismus, der CO als Substrat verwendet, geschieht. Diese ATP-abhängige Carbonylierungsreaktion von Aceton führt zur Bildung eines Acetoacetaldehyds als Intermediat. Das im Rahmen dieser Arbeit sequenzierte und annotierte Genom von D. biacutus, ermöglichte weitere Einblicke und ein besseres Verständnis der Acetonaktivierung und öffnete die Tür für weitere Errungenschaften auf diesem Gebiet. Es konnte zum Beispiel mittels genomischer und proteomischer Analyse ein TPP-abhängiges Enzym gefunden werden, welches in der Aktivierungsreaktion innerhalb des Metabolismus von Aceton involviert sein könnte. Darüber hinaus konnte mittels einer neuartigen ATP-Aktivitätssonde gezeigt werden, dass TPP ein Kofaktor in der initiierenden Reaktion darstellt. Es besteht weiterhin die
Möglichkeit, dass ein B12 Enzym an der Acetonabbaureaktion teilnimmt. Abschließend konnte gezeigt werden, dass sich die strikt anaerobe Aktivierung von Ketonen und Kohlenwasserstoffen stark von der Aktivierung in Anwesenheit von Sauerstoff unterscheidet und dass die Entdeckung dieser neuartigen Acetonaktivierung für zukünftige Untersuchungen auf dem Gebiet der Kohlenwasserstoffaktivierung sich als hilfreich erweisen könnte.
78
RECORD OF ACHIEVEMENT
Unless stated otherwise all experiments in this work were done and analyzed by myself. All experiments were mostly developed and planned by myself, and with help of my supervisor Prof. Bernhard Schink. The chapters of this dissertation unless stated otherwise were written by myself and corrected by Prof. Bernhard Schink and by the corresponding coauthors. The synthesis of the ATP analogue that was used in chapter 3 was done by Norman Hardt under the supervision of Prof. Andreas Marx, as well as the Figures 2 and 3 that involve the synthesis and proof of concept of the ATP analogue. Chapter 3 was written in equal contribution by Norman Hardt and by myself. Revisions and correction were done by our respective supervisors. The lab work, DNA extraction and all process for the genome sequencing and annotation that is mentioned in chapter 4 was only done by myself with the supervision of Dr. David Schleheck and Prof. Bernhard Schink.
79
ABGRENZUNG DER EIGENLEISTUNG
Falls nicht anders angegeben, wurden alle Experimente dieser Arbeit von mir selbst durchgeführt und analysiert. Alle Experimente wurden hauptsächlich von mir selbst unter der Aufsicht von meines Doktorvaters Prof. Bernhard Schink entwickelt und geplant. Die Kapitel dieser Arbeit, soweit nicht anders angegeben, wurden von mir selbst geschrieben und von Prof. Bernhard Schink korrigiert. Die Synthese des in Kapitel 3 verwendeten ATP-Analogs, sowie die Abbildungen 2 und 3, welche die Synthese und den Nachweis des Konzepts des ATP-Analog beinhalten, wurden von Norman Hardt durchgeführt und angefertigt. Kapitel 3 wurde mit gleicher Beitragsleistung von Norman Hardt und mir angefertigt. Berichtigungen und Korrekturen wurden von den Doktorvätern Prof. Andreas Marx und Prof. Bernhard Schink durchgeführt. Die in Kapitel 4 erwähnte Laborarbeit, DNA Extraktion und alle Schritte der Genomsequenzierung und –annotation wurden nur von mir selbst unter Hilfestellung von Dr. David Schleheck und Prof. Bernhard Schink durchgeführt.
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References
REFERENCES
Aeckersberg, F., F. Bak, and F. Widdel. 1991. Anaerobic oxidation of saturated hydrocarbons to CO2 by a new type of sulfate-reducing bacterium. Archives of microbiology. 156:5-14. Aeckersberg, F., F.A. Rainey, and F. Widdel. 1998. Growth, natural relationships, cellular fatty acids and metabolic adaptation of sulfate-reducing bacteria that utilize long- chain alkanes under anoxic conditions. Archives of microbiology. 170:361-369. Agyei-Owusu, K., and F.J. Leeper. 2009. Thiamin diphosphate in biological chemistry: analogues of thiamin diphosphate in studies of enzymes and riboswitches. The FEBS journal. 276:2905-2916. Altenbuchner, J., M. Siemann-Herzberg, and C. Syldatk. 2001. Hydantoinases and related enzymes as biocatalysts for the synthesis of unnatural chiral amino acids. Current Opinion in Biotechnology. 12:559-563. Anderson, R.T., and D.R. Lovley. 2000. Hexadecane decay by methanogenesis. Nature. 404:722-723. Arzumanov, A.A., D.G. Semizarov, L.S. Victorova, N.B. Dyatkina, and A.A. Krayevsky. 1996. Gamma-phosphate-substituted 2'-deoxynucleoside 5'-triphosphates as substrates for DNA polymerases. The Journal of biological chemistry. 271:24389-24394. Asagoe, Y., S. Nakamoto, F. Kitamuro, and M. Shimao. 1968. Studies on the metabolism of acetone bodies in diabetic rabbits. Yonago acta medica. 12:69-74. Baboshin, M.A., and L.A. Golovleva. 2012. Biodegradation of polycyclic aromatic hydrocarbons (PAH) by aerobic bacteria and its kinetics aspects. Mikrobiologiia. 81:695-706. Bagshaw, C. 2001. ATP analogues at a glance. Journal of cell science. 114:459-460. Bettendorff, L., B. Wirtzfeld, A.F. Makarchikov, G. Mazzucchelli, M. Frederich, T. Gigliobianco, M. Gangolf, E. De Pauw, L. Angenot, and P. Wins. 2007. Discovery of a natural thiamine adenine nucleotide. Nature chemical biology. 3:211-212. Birks, S.J., and D.J. Kelly. 1997. Assay and properties of acetone carboxylase, a novel enzyme involved in acetone-dependent growth and CO2 fixation in Rhodobacter capsulatus and other photosynthetic and denitrifying bacteria. Microbiology. 143:755- 766. Bondoc, F.Y., Z. Bao, W.Y. Hu, F.J. Gonzalez, Y. Wang, C.S. Yang, and J.Y. Hong. 1999. Acetone catabolism by cytochrome P450 2E1: studies with CYP2E1-null mice. Biochemical pharmacology. 58:461-463. Bonnet-Smits, E.M., L.A. Robertson, J.P. Van Dijken, E. Senior, and J.G. Kuenen. 1988. Carbon dioxide fixation as the initial step in the metabolism of acetone by Thiosphaera pantotropha. Journal of General Microbiology. 134:2281-2289.
81
References
Boyd, J.M., H. Ellsworth, and S.A. Ensign. 2004. Bacterial acetone carboxylase is a manganese-dependent metalloenzyme. The Journal of biological chemistry. 279:46644- 46651. Boyd, J.M., and S.A. Ensign. 2005. ATP-dependent enolization of acetone by acetone carboxylase from Rhodobacter capsulatus. Biochemistry. 44:8543-8553. Callaghan, A.V., B.E. Morris, I.A. Pereira, M.J. McInerney, R.N. Austin, J.T. Groves, J.J. Kukor, J.M. Suflita, L.Y. Young, G.J. Zylstra, and B. Wawrik. 2012. The genome sequence of Desulfatibacillum alkenivorans AK-01: a blueprint for anaerobic alkane oxidation. Environmental microbiology. 14:101-113. Callaghan, A.V., M. Tierney, C.D. Phelps, and L.Y. Young. 2009. Anaerobic biodegradation of n-hexadecane by a nitrate-reducing consortium. Applied and environmental microbiology. 75:1339-1344. Cirera-Domenech, E., R. Estrada-Tejedor, F. Broto-Puig, J. Teixido, M. Gassiot-Matas, L. Comellas, J.L. Lliberia, A. Mendez, S. Paz-Estivill, and M.R. Delgado-Ortiz. 2013. Quantitative structure-retention relationships applied to liquid chromatography gradient elution method for the determination of carbonyl-2,4- dinitrophenylhydrazone compounds. Journal of chromatography. A. 1276:65-77. Clark, D.D., and S.A. Ensign. 1999. Evidence for an inducible nucleotide-dependent acetone carboxylase in Rhodococcus rhodochrous B276. Journal of bacteriology. 181:2752- 2758. Cline, J.D. 1969. Spectrophotometric determination of hydrogen sulfide in natural waters. Limnol. Oceanogr. 14:454-458. Cole, D.G., and R.G. Yount. 1990. Photolabeling of the 6 and 10 S conformations of gizzard myosin with 3'(2')-O-(4-Benzoyl)benzoyl-ATP. Proline 324 is near the active site. The Journal of biological chemistry. 265:22537-22546. Cremo, C.R., J.M. Neuron, and R.G. Yount. 1990. Interaction of myosin subfragment 1 with fluorescent ribose-modified nucleotides. A comparison of vanadate trapping and SH1-SH2 cross-linking. Biochemistry. 29:3309-3319. Dennis, M., and P.E. Kolattukudy. 1992. A cobalt-porphyrin enzyme converts a fatty aldehyde to a hydrocarbon and CO. Proceedings of the National Academy of Sciences of the United States of America. 89:5306-5310. Díaz, E., J.I. Jiménez, and J. Nogales. 2013. Aerobic degradation of aromatic compounds. Current Opinion in Biotechnology. 24:431-442. DiDonato, R.J., Jr., N.D. Young, J.E. Butler, K.J. Chin, K.K. Hixson, P. Mouser, M.S. Lipton, R. DeBoy, and B.A. Methe. 2010. Genome sequence of the deltaproteobacterial strain NaphS2 and analysis of differential gene expression during anaerobic growth on naphthalene. PloS one. 5:e14072. Dolgikh, A.N., A.V. Bogdanova, G.I. Plotnikova, T.M. Ushakova, and M.F. Shostakovskii. 1964. Butadiyne derivatives. Reaction of 1-buten-3-ynil sulfides with water. Communication 10. Zelinskii Institute of Organic Chemistry, Academy of
82
References
Sciences, USSR. Translated from Izvestiya Akademii Nauk SSSR, Seriya Khimicheskaya. No. 1:127-132. Duerre, P., H. Bahl, and G. Gottschalk. 1992. ChemInform Abstract Acetone-Butanol Fermentation: basis of a modern biotechnological process?, ChemInform Volume 23, Issue 37. ChemInform. 23. Dullius, C.H., C.Y. Chen, and B. Schink. 2011. Nitrate-dependent degradation of acetone by Alicycliphilus and Paracoccus strains and comparison of acetone carboxylase enzymes. Applied and environmental microbiology. 77:6821-6825. Eklund, H., and M. Fontecave. 1999. Glycyl radical enzymes: a conservative structural basis for radicals. Structure. 7:R257-262. Esterbauer, H., H. Zollner, and N. Scholz. 1975. Reaction of glutathione with conjugated carbonyls. Zeitschrift fur Naturforschung. Section C: Biosciences. 30:466-473. Ferreira, L.M., M.M.B. Marques, P.M.C. Glória, H.T. Chaves, J.-P.P. Franco, I. Mourato, J.-R.T. Antunes, H.S. Rzepa, A.M. Lobo, and S. Prabhakar. 2008. Reaction of aromatic nitroso compounds with chemical models of ‘thiamine active aldehyde’. Tetrahedron. 64:7759-7770. Freeman, R., T. Finder, R. Gill, and I. Willner. 2010. Probing protein kinase (CK2) and alkaline phosphatase with CdSe/ZnS quantum dots. Nano letters. 10:2192-2196. Goker, M., H. Teshima, A. Lapidus, M. Nolan, S. Lucas, N. Hammon, S. Deshpande, J.F. Cheng, R. Tapia, C. Han, L. Goodwin, S. Pitluck, M. Huntemann, K. Liolios, N. Ivanova, I. Pagani, K. Mavromatis, G. Ovchinikova, A. Pati, A. Chen, K. Palaniappan, M. Land, L. Hauser, E.M. Brambilla, M. Rohde, S. Spring, J.C. Detter, T. Woyke, J. Bristow, J.A. Eisen, V. Markowitz, P. Hugenholtz, N.C. Kyrpides, and H.P. Klenk. 2011. Complete genome sequence of the acetate- degrading sulfate reducer Desulfobacca acetoxidans type strain (ASRB2). Standards in genomic sciences. 4:393-401. Green, K.D., and M.K. Pflum. 2007. Kinase-catalyzed biotinylation for phosphoprotein detection. J Am Chem Soc. 129:10-11. Green, K.D., and M.K. Pflum. 2009. Exploring kinase cosubstrate promiscuity: monitoring kinase activity through dansylation. Chembiochem : a European journal of chemical biology. 10:234-237. Gutierrez Acosta, O.B., N. Hardt, S.M. Hacker, T. Strittmatter, B. Schink, and A. Marx. 2013a. ATP and thiamine pyrophosphate dependence of acetone degradation by the sulfate-reducing bacterium Desulfococcus biacutus monitored by a fluorogenic ATP analogue. Submitted. Gutierrez Acosta, O.B., N. Hardt, and B. Schink. 2013b. Carbonylation as a key reaction in anaerobic acetone activation by Desulfococcus biacutus. Applied and environmental microbiology. 79:6228-6235. Hacker, S.M., N. Hardt, A. Buntru, D. Pagliarini, M. Mockel, T.U. Mayer, M. Scheffner, C.R. Hauck, and A. Marx. 2013a. Fingerprinting differential active site constraints of ATPases. Chemical Science. 4:1588-1596.
83
References
Hacker, S.M., M. Mex, and A. Marx. 2012. Synthesis and stability of phosphate modified ATP analogues. The Journal of organic chemistry. 77:10450-10454. Hacker, S.M., D. Pagliarini, T. Tischer, N. Hardt, D. Schneider, M. Mex, T.U. Mayer, M. Scheffner, and A. Marx. 2013b. Fluorogenic ATP analogues for online monitoring of ATP consumption: observing ubiquitin activation in real time. Angewandte Chemie International Edition. Haley, C.A., and P. Maitland. 1951. Organic reactions in aqueous solution at room temperature. Part I. The influence of pH on condensations involving the linking of carbon to nitrogen and of carbon to carbon. J. Chem. Soc. 691:3155-3174. Han, B., V. Gopalan, and T.C. Ezeji. 2011. Acetone production in solventogenic Clostridium species: new insights from non-enzymatic decarboxylation of acetoacetate. Applied microbiology and biotechnology. 91:565-576. Haynes, C.A., J.C. Allegood, K. Sims, E.W. Wang, M.C. Sullards, and A.H. Merrill, Jr. 2008. Quantitation of fatty acyl-coenzyme As in mammalian cells by liquid chromatography-electrospray ionization tandem mass spectrometry. Journal of lipid research. 49:1113-1125. Huang, X.H., H.S. Ip, and J.Z. Yu. 2007. Determination of trace amounts of formaldehyde in acetone. Analytica chimica acta. 604:134-138.
Janssen, P.H., and B. Schink. 1995a. 14CO2 exchange with acetoacetate catalyzed by dialyzed cell-free extracts of the bacterial strain BunN grown with acetone and nitrate. European journal of biochemistry / FEBS. 228:677-682. Janssen, P.H., and B. Schink. 1995b. Metabolic pathways and energetics of the acetone- oxidizing, sulfate-reducing bacterium, Desulfobacterium cetonicum. Archives of microbiology. 163:188-194. Janssen, P.H., and B. Schnik. 1995. Catabolic and anabolic enzyme activities and energetics of acetone metabolism of the sulfate-reducing bacterium Desulfococcus biacutus. Journal of bacteriology. 177:277-282. Jobst, B., K. Schühle, U. Linne, and J. Heider. 2010. ATP-dependent carboxylation of acetophenone by a novel type of carboxylase. Journal of bacteriology. 192:1387-1394. Kalapos, M.P. 1997. Acetone metabolism: biochemistry, toxicology and clinical implications. Orvosi hetilap. 138:1187-1193. Klenk, H.P., R.A. Clayton, J.F. Tomb, O. White, K.E. Nelson, K.A. Ketchum, R.J. Dodson, M. Gwinn, E.K. Hickey, J.D. Peterson, D.L. Richardson, A.R. Kerlavage, D.E. Graham, N.C. Kyrpides, R.D. Fleischmann, J. Quackenbush, N.H. Lee, G.G. Sutton, S. Gill, E.F. Kirkness, B.A. Dougherty, K. McKenney, M.D. Adams, B. Loftus, S. Peterson, C.I. Reich, L.K. McNeil, J.H. Badger, A. Glodek, L. Zhou, R. Overbeek, J.D. Gocayne, J.F. Weidman, L. McDonald, T. Utterback, M.D. Cotton, T. Spriggs, P. Artiach, B.P. Kaine, S.M. Sykes, P.W. Sadow, K.P. D'Andrea, C. Bowman, C. Fujii, S.A. Garland, T.M. Mason, G.J. Olsen, C.M. Fraser, H.O. Smith, C.R. Woese, and J.C. Venter. 1997. The complete
84
References
genome sequence of the hyperthermophilic, sulphate-reducing archaeon Archaeoglobus fulgidus. Nature. 390:364-370. Kluger, R. 1987. Thiamin diphosphate: a mechanistic update on enzymic and nonenzymic catalysis of decarboxylation. Chemical Reviews. 87:863-876. Knorre, D.G., V.A. Kurbatov, and V.V. Samukov. 1976. General method for the synthesis of ATP gamma-derivatives. FEBS letters. 70:105-108. Kochetov, G.A., and A.E. Izotova. 1972. Interaction of thiamine pyrophosphate and transketolase. Doklady Akademii nauk SSSR. 205:986-988. Kondo, H., and M. Ishimoto. 1974. Requirement for thiamine pyrophosphate and magnesium for sulfoacetaldehyde sulfo-lyase activity. Journal of biochemistry. 76:229- 231. Koop, D.R., and J.P. Casazza. 1985. Identification of ethanol-inducible P-450 isozyme 3a as the acetone and acetol monooxygenase of rabbit microsomes. The Journal of biological chemistry. 260:13607-13612. Korlach, J., A. Bibillo, J. Wegener, P. Peluso, T.T. Pham, I. Park, S. Clark, G.A. Otto, and S.W. Turner. 2008. Long, processive enzymatic DNA synthesis using 100% dye- labeled terminal phosphate-linked nucleotides. Nucleosides, nucleotides & nucleic acids. 27:1072-1083. Kropp, K.G., I.A. Davidova, and J.M. Suflita. 2000. Anaerobic oxidation of n-dodecane by an addition reaction in a sulfate-reducing bacterial enrichment culture. Applied and environmental microbiology. 66:5393-5398. Kruse, C.H., K.G. Holden, P.H. Offen, M.L. Pritchard, J.A. Feild, D.J. Rieman, P.E. Bender, B. Ferguson, R.G. Greig, and G. Poste. 1988. Synthesis and evaluation of multisubstrate inhibitors of an oncogene-encoded tyrosine-specific protein kinase. 2. Journal of medicinal chemistry. 31:1768-1772. Kumar, S., A. Sood, J. Wegener, P.J. Finn, S. Nampalli, J.R. Nelson, A. Sekher, P. Mitsis, J. Macklin, and C.W. Fuller. 2005. Terminal phosphate labeled nucleotides: synthesis, applications, and linker effect on incorporation by DNA polymerases. Nucleosides, nucleotides & nucleic acids. 24:401-408. Laemmli, U.K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature. 227:680-685. Laffel, L. 1999. Ketone bodies: a review of physiology, pathophysiology and application of monitoring to diabetes. Diabetes/metabolism research and reviews. 15:412-426. Landau, B.R., and H. Brunengraber. 1987. The role of acetone in the conversion of fat to carbohydrate. Trends in Biochemical Sciences. 12:113-114. Lee, S.E., L.M. Elphick, A.A. Anderson, L. Bonnac, E.S. Child, D.J. Mann, and V. Gouverneur. 2009. Synthesis and reactivity of novel gamma-phosphate modified ATP analogues. Bioorganic & medicinal chemistry letters. 19:3804-3807. Lépiz-Aguilar, L., C.E. Rodríguez-Rodríguez, M.L. Arias, and G. Lutz. 2013. Acetone- Butanol-Ethanol (ABE) Production in fermentation of enzymatically hydrolyzed
85
References
cassava flour by Clostridium beijerinckii BA101 and solvent separation. Journal of microbiology and biotechnology. 23:1092-1098. Lindqvist, Y., G. Schneider, U. Ermler, and M. Sundstrom. 1992. Three-dimensional structure of transketolase, a thiamine diphosphate dependent enzyme, at 2.5 A resolution. The EMBO journal. 11:2373-2379. Lukins, H.B., and J.W. Foster. 1963. Methyl ketone metabolism in hydrocarbon-utilizing mycobacteria. Journal of bacteriology. 85:1074-1087. Magnes, C., F.M. Sinner, W. Regittnig, and T.R. Pieber. 2005. LC/MS/MS method for quantitative determination of long-chain fatty acyl-CoAs. Analytical chemistry. 77:2889-2894. Mansoorabadi, S.O., J. Seravalli, C. Furdui, V. Krymov, G.J. Gerfen, T.P. Begley, J. Melnick, S.W. Ragsdale, and G.H. Reed. 2006. EPR spectroscopic and computational characterization of the hydroxyethylidene-thiamine pyrophosphate radical intermediate of pyruvate:ferredoxin oxidoreductase. Biochemistry. 45:7122- 7131. Meckenstock, R.U., and H. Mouttaki. 2011. Anaerobic degradation of non-substituted aromatic hydrocarbons. Current Opinion in Biotechnology. 22:406-414. Niyomrattanakit, P., S.N. Abas, C.C. Lim, D. Beer, P.Y. Shi, and Y.L. Chen. 2011. A fluorescence-based alkaline phosphatase-coupled polymerase assay for identification of inhibitors of dengue virus RNA-dependent RNA polymerase. Journal of biomolecular screening. 16:201-210. Norwood, D.L., C.A. Bus, and D.S. Millington. 1990. Combined high-performance liquid chromatographic-continuous-flow fast atom bombardment mass spectrometric analysis of acylcoenzyme A compounds. Journal of chromatography. 527:289-301. Oiwa, K., J.F. Eccleston, M. Anson, M. Kikumoto, C.T. Davis, G.P. Reid, M.A. Ferenczi, J.E. Corrie, A. Yamada, H. Nakayama, and D.R. Trentham. 2000. Comparative single-molecule and ensemble myosin enzymology: sulfoindocyanine ATP and ADP derivatives. Biophysical journal. 78:3048-3071. Oiwa, K., D.M. Jameson, J.C. Croney, C.T. Davis, J.F. Eccleston, and M. Anson. 2003. The 2'-O- and 3'-O-Cy3-EDA-ATP(ADP) complexes with myosin subfragment-1 are spectroscopically distinct. Biophysical journal. 84:634-642. Oiwa, K., T. Yamaga, and A. Yamada. 1998. Direct observation of a central bare zone in a native thick filament isolated from the anterior byssus retractor muscle of Mytilus edulis using fluorescent ATP analogue. Journal of biochemistry. 123:614-618. Park, B.C., Y. Liu, D.R. Averill, Jr., M.I. Jackman, and D.W. Rosenberg. 1995. Cytochemical localization of the acetone-inducible cytochrome P-450 isoform, CYP2E1, in murine colonic epithelium. Pharmacology. 50:339-347. Philipp, B., and B. Schink. 2012. Different strategies in anaerobic biodegradation of aromatic compounds: nitrate reducers versus strict anaerobes. Environmental Microbiology Reports. 4:469-478.
86
References
Pierce, E., D.F. Becker, and S.W. Ragsdale. 2010. Identification and characterization of oxalate oxidoreductase, a novel thiamine pyrophosphate-dependent 2-oxoacid oxidoreductase that enables anaerobic growth on oxalate. The Journal of biological chemistry. 285:40515-40524. Platen, H., P.H. Janssen, and B. Schink. 1994. Fermentative degradation of acetone by an enrichment culture in membrane-separated culture devices and in cell suspensions. FEMS microbiology letters. 122:27-32. Platen, H., and B. Schink. 1987. Methanogenic degradation of acetone by an enrichment culture. Archives of microbiology. 149:136-141. Platen, H., and B. Schink. 1989. Anaerobic degradation of acetone and higher ketones via carboxylation by newly isolated denitrifying bacteria. Journal of general microbiology 135:883-891. Platen, H., and B. Schink. 1990. Enzymes involved in anaerobic degradation of acetone by a denitrifying bacterium. Biodegradation. 1:243-251. Platen, H., A. Temmes, and B. Schink. 1990. Anaerobic degradation of acetone by Desulfococcus biacutus spec. nov. Archives of microbiology. 154:355-361. Pollack, S.E., and D.S. Auld. 1982. Fluorescent nucleotide triphosphate substrates for snake venom phosphodiesterase. Analytical biochemistry. 127:81-88. Price, E.E. 1992. Atomic Form. Special reference to the configuration of the carbon atom. Benzene and some of its derivatives. Chapter III. . Longmans Green Company, London. Rabus, R., M. Kube, A. Beck, F. Widdel, and R. Reinhardt. 2002. Genes involved in the anaerobic degradation of ethylbenzene in a denitrifying bacterium, strain EbN1. Archives of microbiology. 178:506-516. Rabus, R., A. Ruepp, T. Frickey, T. Rattei, B. Fartmann, M. Stark, M. Bauer, A. Zibat, T. Lombardot, I. Becker, J. Amann, K. Gellner, H. Teeling, W.D. Leuschner, F.O. Glockner, A.N. Lupas, R. Amann, and H.P. Klenk. 2004. The genome of Desulfotalea psychrophila, a sulfate-reducing bacterium from permanently cold Arctic sediments. Environmental microbiology. 6:887-902. Rabus, R., and F. Widdel. 1995. Anaerobic degradation of ethylbenzene and other aromatic hydrocarbons by new denitrifying bacteria. Archives of microbiology. 163:96- 103. Rabus, R., H. Wilkes, A. Behrends, A. Armstroff, T. Fischer, A.J. Pierik, and F. Widdel. 2001. Anaerobic initial reaction of n-alkanes in a denitrifying bacterium: evidence for (1-methylpentyl)succinate as initial product and for involvement of an organic radical in n-hexane metabolism. Journal of bacteriology. 183:1707-1715. Reed, L. 1974. Multienzyme Complexes. Acc. Chem. Res. 7:40-46. Rosier, C., N. Leys, C. Henoumont, M. Mergeay, and R. Wattiez. 2012. Purification and characterization of the acetone carboxylase of Cupriavidus metallidurans strain CH34. Applied and environmental microbiology. 78:4516-4518.
87
References
Schauenstein, E. 1967. Autoxidation of polyunsaturated esters in water: chemical structure and biological activity of the products. Journal of lipid research. 8:417-428. Schühle, K., and J. Heider. 2012. Acetone and butanone metabolism of the denitrifying bacterium "Aromatoleum aromaticum" demonstrates novel biochemical properties of an ATP-dependent aliphatic ketone carboxylase. Journal of bacteriology. 194:131-141. Selesi, D., N. Jehmlich, M. von Bergen, F. Schmidt, T. Rattei, P. Tischler, T. Lueders, and R.U. Meckenstock. 2010. Combined genomic and proteomic approaches identify gene clusters involved in anaerobic 2-methylnaphthalene degradation in the sulfate-reducing enrichment culture N47. Journal of bacteriology. 192:295-306. Selmer, T., A.J. Pierik, and J. Heider. 2005. New glycyl radical enzymes catalysing key metabolic steps in anaerobic bacteria. Biological chemistry. 386:981-988. Siegel, J.M. 1950. The metabolism of acetone by the photosynthetic bacterium Rhodopseudomonas gelatinosa. Journal of bacteriology. 60:595-606. Siegel, J.M. 1954. The photosynthetic metabolism of acetone by Rhodopseudomonas gelatinosa. The Journal of biological chemistry. 208:205-216. Sifniades, S., A.B. Levy, and H. Bahl. 2011. Acetone. In Ullmann's Encyclopedia of Industrial Chemistry. Wiley-VCH Verlag GmbH & Co. KGaA. Singh, H.B., D. O'Hara, D. Herlth, W. Sachse, D.R. Blake, J.D. Bradshaw, M. Kanakidou, and P.J. Crutzen. 1994. Acetone in the atmosphere: Distribution, sources, and sinks. Journal of Geophysical Research: Atmospheres. 99:1805-1819. Sluis, M.K., and S.A. Ensign. 1997. Purification and characterization of acetone carboxylase from Xanthobacter strain Py2. Proceedings of the National Academy of Sciences of the United States of America. 94:8456-8461. Sluis, M.K., R.A. Larsen, J.G. Krum, R. Anderson, W.W. Metcalf, and S.A. Ensign. 2002. Biochemical, molecular, and genetic analyses of the acetone carboxylases from Xanthobacter autotrophicus strain Py2 and Rhodobacter capsulatus strain B10. Journal of bacteriology. 184:2969-2977. Sluis, M.K., F.J. Small, J.R. Allen, and S.A. Ensign. 1996. Involvement of an ATP- dependent carboxylase in a CO2-dependent pathway of acetone metabolism by Xanthobacter strain Py2. Journal of bacteriology. 178:4020-4026. Song, H., K. Kerman, and H.B. Kraatz. 2008. Electrochemical detection of kinase- catalyzed phosphorylation using ferrocene-conjugated ATP. Chemical communications:502-504. Sood, A., S. Kumar, S. Nampalli, J.R. Nelson, J. Macklin, and C.W. Fuller. 2005. Terminal phosphate-labeled nucleotides with improved substrate properties for homogeneous nucleic acid assays. J Am Chem Soc. 127:2394-2395. Spring, S., A. Lapidus, M. Schroder, D. Gleim, D. Sims, L. Meincke, T. Glavina Del Rio, H. Tice, A. Copeland, J.F. Cheng, S. Lucas, F. Chen, M. Nolan, D. Bruce, L. Goodwin, S. Pitluck, N. Ivanova, K. Mavromatis, N. Mikhailova, A. Pati, A. Chen, K. Palaniappan, M. Land, L. Hauser, Y.J. Chang, C.D. Jeffries, P. Chain, E. Saunders, T. Brettin, J.C. Detter, M. Goker, J. Bristow, J.A. Eisen, V. Markowitz,
88
References
P. Hugenholtz, N.C. Kyrpides, H.P. Klenk, and C. Han. 2009. Complete genome sequence of Desulfotomaculum acetoxidans type strain (5575). Standards in genomic sciences. 1:242-253. Stokke, R., W.P. Hocking, B.O. Steinsbu, and I.H. Steen. 2013. Complete genome sequence of the thermophilic and facultatively chemolithoautotrophic sulfate reducer Archaeoglobus sulfaticallidus Strain PM70-1T. Genome announcements. 1. Strittmatter, A.W., H. Liesegang, R. Rabus, I. Decker, J. Amann, S. Andres, A. Henne, W.F. Fricke, R. Martinez-Arias, D. Bartels, A. Goesmann, L. Krause, A. Puhler, H.P. Klenk, M. Richter, M. Schuler, F.O. Glockner, A. Meyerdierks, G. Gottschalk, and R. Amann. 2009. Genome sequence of Desulfobacterium autotrophicum HRM2, a marine sulfate reducer oxidizing organic carbon completely to carbon dioxide. Environmental microbiology. 11:1038-1055. Sun, H., S. Spring, A. Lapidus, K. Davenport, T.G. Del Rio, H. Tice, M. Nolan, A. Copeland, J.F. Cheng, S. Lucas, R. Tapia, L. Goodwin, S. Pitluck, N. Ivanova, I. Pagani, K. Mavromatis, G. Ovchinnikova, A. Pati, A. Chen, K. Palaniappan, L. Hauser, Y.J. Chang, C.D. Jeffries, J.C. Detter, C. Han, M. Rohde, E. Brambilla, M. Goker, T. Woyke, J. Bristow, J.A. Eisen, V. Markowitz, P. Hugenholtz, N.C. Kyrpides, H.P. Klenk, and M. Land. 2010. Complete genome sequence of Desulfarculus baarsii type strain (2st14). Standards in genomic sciences. 3:276-284. Symons, G.E., and A.M. Buswell. 1933. The Methane fermentation of carbohydrates 1,2. Journal of the American Chemical Society. 55:2028-2036. Taylor, D.G., P.W. Trudgill, R.E. Cripps, and P.R. Harris. 1980. The microbial metabolism of acetone. Journal of General Microbiology. 118:159-170. Thauer, R.K., K. Jungermann, and K. Decker. 1977. Energy conservation in chemotrophic anaerobic bacteria. Bacteriological reviews. 41:100-180. Thauer, R.K., E. Stackebrandt, and W.A. Hamilton. 2007. Energy metabolism and phylogenetic diversity of sulphate-reducing bacteria. Cambridge University Press. Uchiyama, S., Y. Inaba, and N. Kunugita. 2011. Derivatization of carbonyl compounds with 2,4-dinitrophenylhydrazine and their subsequent determination by high- performance liquid chromatography. Journal of chromatography. B, Analytical technologies in the biomedical and life sciences. 879:1282-1289. Van den Bergh, V., H. Coeckelberghs, H. Vankerckhoven, F. Compernolle, and C. Vinckier. 2004. Study of the carbonyl products of terpene/OH radical reactions: detection of the 2,4-DNPH derivatives by HPLC-MS. Analytical and bioanalytical chemistry. 379:484-494. Wahl, R.C., and W.H. Orme-Johnson. 1987. Clostridial pyruvate oxidoreductase and the pyruvate-oxidizing enzyme specific to nitrogen fixation in Klebsiella pneumoniae are similar enzymes. The Journal of biological chemistry. 262:10489-10496. Wang, D., Y. Luo, R. Cooke, J. Grammer, E. Pate, and R.G. Yount. 1999. Synthesis of a spin-labeled photoaffinity ATP analogue, and its use to specifically photolabel
89
References
myosin cross-bridges in skeletal muscle fibers. Journal of muscle research and cell motility. 20:743-753. Wang, V.C., M. Can, E. Pierce, S.W. Ragsdale, and F.A. Armstrong. 2013. A unified electrocatalytic description of the action of inhibitors of nickel carbon monoxide dehydrogenase. J Am Chem Soc. 135:2198-2206. Widdel, F., and N. Pfennig. 1981. Studies on dissimilatory sulfate-reducing bacteria that decompose fatty acids. I. Isolation of new sulfate-reducing bacteria enriched with acetate from saline environments. Description of Desulfobacter postgatei gen. nov., sp. nov. Archives of microbiology. 129:395-400. Wikén, T. 1940. Untersuchungen über Methangärung und die dabei wirksamen Bakterien. Archiv. Mikrobiol. 11:312-317. Wohlbrand, L., J.H. Jacob, M. Kube, M. Mussmann, R. Jarling, A. Beck, R. Amann, H. Wilkes, R. Reinhardt, and R. Rabus. 2013. Complete genome, catabolic sub- proteomes and key-metabolites of Desulfobacula toluolica Tol2, a marine, aromatic compound-degrading, sulfate-reducing bacterium. Environmental microbiology. 15:1334-1355. Yarbrough, L.R., J.G. Schlageck, and M. Baughman. 1979. Synthesis and properties of fluorescent nucleotide substrates for DNA-dependent RNA polymerases. The Journal of biological chemistry. 254:12069-12073. Zedelius, J., R. Rabus, O. Grundmann, I. Werner, D. Brodkorb, F. Schreiber, P. Ehrenreich, A. Behrends, H. Wilkes, M. Kube, R. Reinhardt, and F. Widdel. 2011. Alkane degradation under anoxic conditions by a nitrate-reducing bacterium with possible involvement of the electron acceptor in substrate activation. Environ Microbiol Rep. 3:125-135. Zengler, K., H.H. Richnow, R. Rossello-Mora, W. Michaelis, and F. Widdel. 1999. Methane formation from long-chain alkanes by anaerobic microorganisms. Nature. 401:266-269. Zinellu, A., V. Pasciu, S. Sotgia, B. Scanu, F. Berlinguer, G. Leoni, S. Succu, I. Cossu, E.S. Passino, S. Naitana, L. Deiana, and C. Carru. 2010. Capillary electrophoresis with laser-induced fluorescence detection for ATP quantification in spermatozoa and oocytes. Analytical and bioanalytical chemistry. 398:2109-2116.
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Supplementary data
Supplementary data
Enzyme reactions in cell-free extracts
Cells of D. biacutus were harvested in the late exponential growth phase at an optical density of 0.3 (OD 600), and cell-free extracts were prepared as described in the experimental procedure in chapter 2 (Gutierrez Acosta et al., 2013b).
Cobalamin dependent reaction in cell-free extracts
Dependence of cobalamin was done with acetone-grown cell-free extracts of D. biacutus under strictly anoxic conditions, and in the absence of light. The reaction set up included 100 µg of protein, 100 µM cobalamin (5-deoxyadenosylcobalamin), 5 µM Ti-NTA, 2mM ATP, 2mM acetone, and CO. Control experiments without acetone, without CO or dead cell extracts were also performed. A change in the state of the cobalamin was detected in the presence of acetone and CO (Fig. 1 and 2). The peak signal at 475 nm corresponding to the Cobalt II of the cobalamin increased with the addition of acetone and CO, however did not depend on the presence of ATP.
Products of the ATP hydrolysis during the acetone activating reaction in cell-free extracts
The hydrolysis of 3mM ATP was measured in the presence of 2mM TPP, 3mM acetone, CO and cell-free extracts (Fig. 3 and 4). Samples were taken at different reaction time intervals and measured in a HIC-ZILLIC column. The results from this preliminary experiments indicated that AMP is the product of the ATP-dependent activation of acetone.
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Supplementary data
0.7
0.6
CO+ATP+CE+CoB12 0.5
0.4 CO+ATP+CE+CoB12+acetone 5m
0.3 CO+ATP+CE+CoB12+acetone 15m
0.2 CO+ATP+CE+CoB12+acetone 30m Abs
0.1
0 200 300 400 500 600 700 -0.1
-0.2
-0.3 wavelenght (nm)
Figure 1. Reaction with cobalamine in cell-free extract of Desulfococcus biacutus. Increase of the peak at 475 nm (Co II) was detected with the addition of acetone and CO.
92
Supplementary data
Figure 2. Reaction with cobalamine in cell-free extract of Desulfococcus biacutus. Increase of the peak at 475 nm (Co II) was specific for acetone and CO as it is indicated with controls without acetone and without CO.
93
Supplementary data
Figure 3. ATP consumption during the activation of acteone in acetone- grown cell-free extracts.
uV 1500000 Data1:5mM ATP_001.lcd PDA Ch1:254nm,4nm(1,00) Data2:7-0_001.lcd PDA Ch1:254nm,4nm(1,00) Data3:7-15_002.lcd PDA Ch1:254nm,4nm(1,00) Data4:7-30_003.lcd PDA Ch1:254nm,4nm(1,00) 1250000 Data5:7-60_004.lcd PDA Ch1:254nm,4nm(1,00) Data6:7-120_005.lcd PDA Ch1:254nm,4nm(1,00)
1000000
750000
500000
250000
0
22,75 23,00 23,25 23,50 23,75 24,00 24,25 24,50 24,75 min
Figure 4. ATP consumption during the acetone activation with acetone- grown cell-free extracts. AMP appears at 23 min of retention time, ADP appears at 23.8 min approx. and ATP can be seen at 24.7 min approx.
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SCIENTIFIC CONTRIBUTIONS LIST
Articles 1. Olga B. Gutiérrez Acosta, Norman Hardt and Bernhard Schink (2013). Carbonylation as a key reaction in anaerobic acetone activation by Desulfococcus biacutus. Applied and environmental microbiology. 79:6228-6235. 2. Gutierrez Acosta, O.B., Hardt, N., Hacker, S.M., Strittmatter, T., Schink, B. and A. Marx (2013). ATP and thiamine pyrophosphate dependence of acetone degradation by the sulfate-reducing bacterium Desulfococcus biacutus monitored by a fluorogenic ATP analogue. Submitted. 3. Olga B. Gutiérrez Acosta, David Schleheck and Bernhard Schink (2013). Acetone utilization under sulfate-reducing conditions: draft genome sequence of Desulfococcus biacutus and a proteomic survey of acetone-inducible proteins. To be submitted.
Attended meetings and workshops 1. Deutsche Forschungsgemeinschaft (DFG) SPP 1319 priority program. Second workshop in Biological transformations of hydrocarbons without oxygen: from the molecular to the global scale. Mar. 09-11 2011, Herrsching/ Ammersee
Oral presentation: Anaerobic activation of acetone by nitrate-reducing and sulfate- reducing bacteria.
2 Deutsche Forschungsgemeinschaft (DFG) SPP 1319 priority program. Third workshop in biological transformations of hydrocarbons without oxygen: from the molecular to the global scale. Sept. 24-26 2012, Windenreuther Hof/ Freiburg
Oral presentation: Activation of acetone by strictly anaerobic bacteria.
3 Vereinigung für Allgemeine und Angewandte Mikrobiologie (VAAM). Annual meeting, Mar. 10-13 2013, Bremen, Germany
Poster presentation: Evidence of a carbonylation reaction in the activation of acetone under strictly anoxic conditions.
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