Microbial desulfonation pathways for natural and pharmacologically relevant C 3-sulfonates

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Microbial desulfonation pathways for natural and pharmacologically

relevant C ₃ -sulfonates

Dissertation

zur Erlangung des akademischen Grades des Doktors der Naturwissenschaften (Dr. rer. nat.)

an der Universität Konstanz Fachbereich Biologie

vorgelegt von

Jutta Mayer

Tag der mündlichen Prüfung: 18. Februar 2011 1. Referent: Prof. Dr. Alasdair M. Cook

2. Referent: Prof. Dr. Bernhard Schink

3. Referent: Prof. Dr. Jörg S. Hartig (Prüfungsvorsitzender)

Konstanzer Online-Publikations-System (KOPS) URL: http://nbn-resolving.de/urn:nbn:de:bsz:352-123990

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Mayer, J., T. Huhn, M. Habeck, K. Denger, K. Hollemeyer and A. M. Cook (2010).

2,3-Dihydroxypropane-1-sulfonate degraded by Cupriavidus pinatu bonensis JMP134:

purification of dihydroxypropanesulfonate 3-dehydrogenase. Microbiology 156: 1556-1564.

Mayer, J. and A. M. Cook (2009). Homotaurine metabolized to 3-sulfopropanoate in Cupriavidus necator H16: enzymes and genes in a patchwork pathway. J Bacteriol 191: 6052- 6058.

Denger, K., J. Mayer, M. Buhmann, S. Weinitschke, T. H. M. Smits and A. M. Cook (2009). Bifurcated degradative pathway of 3-sulfolactate in Roseovarius nubinhibens ISM via sulfoacetaldehyde acetyltransferase and (R)-cysteate sulfolyase. J Bacteriol 191: 5648-5656.

Mayer, J., K. Denger, K. Kaspar, K. Hollemeyer, T. H. M. Smits, T. Huhn and A. M. Cook (2008). Assimilation of homotaurine-nitrogen by Burkholderia sp. and excretion of sulfopropanoate. FEMS Microbiol Lett 279: 77-82.

Further publications:

Denger, K., J. Mayer, K. Hollemeyer and A. M. Cook (2008). Amphoteric surfactant N-oleoyl-N-methyltaurine utilized by Pseudomonas alca ligenes with excretion of N-methyltaurine. FEMS Microbiol Lett 288: 112-117.

Mayer, J., K. Denger, T. H. M. Smits, K. Hollemeyer, U. Groth and A. M. Cook (2006).

N-Acetyltaurine dissimilated via taurine by Delftia acidovorans NAT. Arch Microbiol 186:

61-67.

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An dieser Stelle möchte ich mich herzlich bei Prof. Dr. Alasdair M. Cook bedanken:

… dafür, dass er es mir ermöglicht hat, die Doktorarbeit in seiner Gruppe unter besten Bedingungen anzufertigen,

… für das spannende und vielseitige Thema, das er mir überlassen hat,

… für seine stete Hilfsbereitschaft bei Problemen und ganz besonders für die große wissenschaftliche Freiheit, die ich in seiner Arbeitsgruppe erfahren habe.

Prof. Dr. Bernhard Schink danke ich für die Übernahme des Koreferats, Prof. Dr. Jörg S. Hartig für die Übernahme des Prüfungsvorsitzes. Außerdem möchte ich mich bei beiden für die Betreuung im Rahmen der Graduiertenschule bedanken.

Mein herzlicher Dank geht an alle Mitglieder der Arbeitsgruppe, Karin Denger, Sabine Lehmann und Dr. David Schleheck, sowie an die früheren Gruppenmitglieder Dr. Zdeněk Krejčík und Dr. Sonja Weinitschke, für die Hilfsbereitschaft, die zahlreichen Diskussionen und die tolle Atmosphäre im Labor. Es ist schön, mit Freunden zusammenarbeiten zu dürfen!

Besonders danken möchte ich David und Karin für die kritische Durchsicht meiner Arbeit.

Herzlich danken möchte ich weiterhin…

… Dr. Thomas Huhn für die Synthese und Charakterisierung von kostbaren Sulfonaten, sowie für die Beratung unserer Chemikerinnen Katrin Kaspar und Birgit Fredrich. Auch ihnen sei herzlich gedankt!

… Dr. Theo H. M. Smits für die Einführung in molekularbiologische und bioinformatische Methoden und sein Mitwirken bei der Mittelbeschaffung für mein Projekt.

… Dr. Klaus Hollemeyer für die zahlreichen MALDI-TOF-MS Messungen.

… den Arbeitsgruppen Schink und Mendgen für die Mitbenutzung von Geräten und Chemikalien, sowie für die gute Nachbarschaft.

… Prof. Dr. Andreas Marx, der mir während meines Studiums, insbesondere während Bachelor- und Masterarbeit, ein motivierender Mentor war.

… der Graduiertenschule „Konstanz Research School Chemical Biology“ für die Möglichkeit, in zahlreichen Vorträgen und Kursen über den Tellerrand zu schauen, sowie andere Doktoranden und deren Forschung kennen zu lernen.

… dem Zukunftskolleg der Universität Konstanz für die finanzielle Förderung und die interessanten Einblicke in andere Disziplinen.

Auch meiner Familie möchte ich an dieser Stelle für ihre unermüdliche moralische und finanzielle Unterstützung während Studium und Promotion danken!

Mein herzlichster Dank ist an meinen Freund Markus Wieland gerichtet: Du hast mich immer unterstützt, mir Zuversicht gespendet, und an mich geglaubt!

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SUMMARY

Homotaurine (3-aminopropanesulfonate), cysteate (2-amino-3-sulfopropanoate), 3-sulfolactate (2-hydroxy-3-sulfopropanoate), 2,3-dihydroxypropane-1-sulfonate (DHPS) and 3-sulfopropanoate are widespread, naturally occuring C₃-sulfonates. To our knowledge, organosulfonates are degraded solely by microorganisms, which are capable of cleaving the chemically stable C-sulfonate bond. The elucidation of degradative pathways in aerobic bacteria utilizing the above mentioned C3-sulfonates as sole sources of carbon and energy or, if possible, as sole sources of nitrogen, was the aim of this study.

Isolates utilizing homotaurine as sole carbon and energy source or as sole nitrogen source were easily obtained. The assimilation of homotaurine-nitrogen was studied with an isolate identified as Burkholderia sp. strain N-APS2. The organism excreted 3-sulfopropanoate during growth with homotaurine-nitrogen, and expressed an inducible homotaurine:2-oxoglutarate aminotransferase. The same phenomena were observed during work with the genome- sequenced Cupriavidus n ecator H16, which revealed the involvement of genes and enzymes from both GABA and sulfonate metabolism: GABA permease (GabP) for homotaurine-uptake, GABA transaminase (GabT) for its deamination to 3-sulfopropanal, and succinate- semialdehyde dehydrogenase (GabD1) for the oxidation of the latter to 3-sulfopropanoate, whose excretion was attributed to the sulfite/sulfonate exporter TauE.

The assimilation of cysteate-nitrogen by C. necator H16 was also found to involve an initial transamination reaction. A cysteate:2-oxoglutarate aminotransferase (Coa), which might be an aspartate aminotransferase (Aoa) according to its substrate spectrum, yielded 3-sulfopyruvate.

Traces of the latter were found in the growth medium, together with putative 3-sulfolactate, whose formation from 3-sulfopyruvate was attributed to the activity of a putative (S)-sulfolactate dehydrogenase (SlcC). Again, TauE might be responsible for organosulfonate export; the identity of a transporter for cysteate uptake, however, is still unknown.

The utilization of 3-sulfolactate as a source of carbon and energy in the genome-sequenced Roseovarius nubin hibens ISM was found to involve a largely inducible, bifurcated pathway, which allowed the desulfonation via sulfoacetaldehyde acetyltransferase (Xsc) and (R)-cysteate sulfo-lyase (CuyA). A putative tripartite tricarboxylate transporter (TTT; SlcHFG) was responsible for uptake of sulfolactate, which was oxidized to 3-sulfopyruvate by a membrane- bound sulfolactate dehydrogenase (SlcD). 3-Sulfopyruvate, the point of bifurcation, was transaminated to cysteate in one branch, or decarboxylated to sulfoacetaldehyde in the other branch. The decarboxylating enzyme, 3-sulfopyruvate decarboxylase (ComDE) is known from the coenzyme M biosynthetic pathway.

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In this study, 3-Sulfolactate was discovered as a central intermediate in the degradation of several C₃-sulfonates, such as DHPS, homotaurine and 3-sulfopropanoate. In the dissimilation of racemic DHPS, three inducible DHPS dehydrogenases (HpsNOP) acted as a racemase (HpsOP) and oxidized (R)-DHPS to (R)-sulfolacate (HpsN). These enzymes were studied in R.

nubinhibens ISM, which degraded the resulting sulfolactate via the bifurcated pathway described above, and in Cupriavidus pinatubonensis JMP134, which used (R)-sulfolactate sulfo- lyase (SuyAB) for desulfonation. Transporter candidates were available in both organisms: a putative tripartite ATP-independent periplasmic (TRAP) transporter in the marine R.

nubinhibens, and a major facilitator superfamily (MFS) transporter in the terrestrial C.

pinatubonensis.

The utilization of homotaurine as a source of carbon and energy was studied in R. nubinhibens, whose substrate spectrum was found to include not only 3-sulfolacate and DHPS, but also homotaurine and 3-sulfopropanoate. As in the nitrogen-assimilatory pathway, transamination and oxidation were observed, but a different transporter (HtaABCD) and aminotransferase (HtaE) became apparent. Two more novel enzymes were found for the further degradation of 3-sulfopropanoate: 3-sulfopropanoate dehydrogenase (SpuBCDA), which yielded 3-sulfopropenoate, and 3-sulfopropenoate dehydratase (SpuIJ), whose predicted product was 3-sulfolactate. Activities of Xsc and CuyA confirmed the presence of the bifurcated pathway for 3-sulfolactate degradation (see above).

Work on the dissimilation of homotaurine revealed nine organisms which encoded the spu-gene cluster for 3-sulfopropanoate degradation. Rhodobacter sphaeroides 2.4.1, which grew with 3-sulfopropanoate, was presumed to degrade it via Xsc; there were, however, no gene candidates for the sulfopyruvate decarboxylase ComDE. Instead, we found a gene encoding for a putative novel 3-sulfopyruvate decarboxylase (SpuE), which is currently under investigation.

The pathways discovered in this thesis were interpreted to represent final steps in the biodegradation of sulfoquinovose, the polar head group of the ubiquitous plant sulfolipid, which is an important intermediate of the biological sulfur cycle. This hypothesis was supported by bioinformatic analyses, which revealed the widespread occurrence of the above pathways within genome-sequenced bacteria.

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ZUSAMMENFASSUNG

Homotaurin (3-Aminopropansulfonat), Cysteat (2-Amino-3-sulfopropanoat), 3-Sulfolactat (2-Hydroxy-3-sulfopropanoat), 2,3-Dihydroxypropan-1-sulfonat (DHPS) und 3-Sulfopropanoat sind weitverbreitete, natürlich vorkommende C₃-Sulfonate. Der Abbau von solchen Organosulfonaten erfolgt nach heutigem Wissensstand ausschließlich durch Mikroorganismen, da nur diese in der Lage sind, die chemisch stabile Kohlenstoff-Sulfonat-Bindung zu spalten.

Ziel dieser Arbeit war die Aufklärung von Abbauwegen für die oben genannten C3-Sulfonate in aeroben Bakterien, welche das jeweilige Substrat als Kohlenstoff- und Energiequelle, oder, falls möglich, als Stickstoffquelle verwenden.

Die erfolgreiche Anreicherung von Homotaurin-verwertenden Kulturen zeigte, dass das Substrat sowohl als Kohlenstoff- wie auch als Stickstoffquelle für Bakterien dienen kann. Die Assimilation von Homotaurin-Stickstoff wurde anhand eines Isolates, welches als Burkholderia sp. identifiziert wurde, untersucht. Während des Wachstums mit Homotaurin-Stickstoff schied der Organismus 3-Sulfopropanoat aus, und exprimierte eine induzierbare Homotaurin:2-Oxo- glutarat-Aminotransferase. Diese Beobachtungen wurden auch mit Cupriavidus necator H16 gemacht, dessen Genom vollständig sequenziert ist. Weiterführende Arbeiten mit Stamm H16 zeigten die Beteiligung von Enzymen und Genen aus dem GABA- und dem Sulfonat- Stoffwechsel: GABA-Permease (GabP) für die Aufnahme von Homotaurin in die Zelle, GABA- Transaminase (GabT) für die Deaminierung von Homotaurin zu 3-Sulfopropanal und Succinat- Semialdehyd-Dehydrogenase (GabD1) für die Oxidation von 3-Sulfopropanal zu 3-Sulfopropanoat. Die Ausscheidung des Oxidationsprodukts wurde dem Sulfit-/Sulfonat- Exporter TauE zugeschrieben.

Eine Transaminierung war auch die erste Reaktion bei der Assimilation von Cysteat-Stickstoff in C. necator H16. Diese wurde von einer Cysteat:2-Oxoglutarat-Aminotransferase (Coa) katalysiert, deren Produkt als 3-Sulfopyruvat identifiziert wurde. Die Aktivität eines 3-Sulfopyurvat-reduzierenden Enzyms, vermutlich (S)-Sulfolactat-Dehydrogenase (SlcC), konnte nachgewiesen werden; weiterhin wurden Spuren von 3-Sulfopyruvat im Kulturüberstand gefunden, zusammen mit bisher nur provisorisch identifiziertem 3-Sulfolactat. Deren Ausscheidung wurde ebenfalls TauE zugeschrieben, während der Transporter für Cysteat immer noch unbekannt ist.

Die Dissimilation von 3-Sulfolactat wurde in Roseovarius nubin hibens ISM, dessen Genomsequenz bekannt ist, untersucht. Dabei wurde ein gegabelter Abbauweg entdeckt, der R.

nubinhibens eine Desulfonierung mittels Sulfoacetaldehyd-Acetyltransferase (Xsc) sowie (R)-Cysteat-Sulfolyase (CuyA) erlaubt. Ein Transporter der TTT (tripartite tricarboxylate transporter)-Familie (SlcHFG) war vermutlich für die Aufnahme von 3-Sulfolactat verantwortlich, welches durch eine membrangebundene Sulfolactat-Dehydrogenase (SlcD) zu

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3-Sulfopyruvat oxidiert wurde. Dieses wurde einerseits zu Cysteat transaminiert, andererseits zu Sulfoacetaldehyd decarboxyliert, und stellt somit den Gabelungspunkt des Abbauweges dar.

Das decarboxylierende Enzym, 3-Sulfopyruvat-Decarboxylase (ComDE), ist aus dem Biosyntheseweg für Coenzym M bekannt.

In der vorliegenden Arbeit wurde 3-Sulfolactat als zentrales Intermediat im Abbau von Homotaurin, DHPS und 3-Sulfopropanoat identifiziert. Bei der Dissimilation von racemischem DHPS wurden drei induzierbare Dehydrogenasen (HpsNOP) entdeckt, die als DHPS-Racemase wirkten (HpsOP) und (R)-DHPS zu (R)-Sulfolactat oxidierten (HpsN). Diese Enzyme wurden in R. nubinh ibens ISM und Cupriavidus necator JMP134 charakterisiert, die das entstandene 3-Sulfolactat über den oben beschriebenen gegabelten Abbauweg bzw. über die (R)-Sulfolactat- Sulfolyase (SuyAB) verwerteten. In beiden Organismen waren Gene vorhanden, die vermutlich DHPS-Transporter kodieren: ein TRAP (tripartite ATP-independent periplasmic)-Transporter in R. nubinhibens und ein MFS (major facilitator superfamily)-Transporter in C. necator.

Die Dissimilation von Homotaurin wurde in R. nubin hibens ISM untersucht, dessen Substratspektrum nicht nur 3-Sulfolactat und DHPS, sondern auch Homotaurin und 3-Sulfopropanoat umfasste. Wie bei der Assimilation von Homotaurin-Stickstoff wurde auch hier eine Transaminierungs- und eine Oxidationsreaktion beobachtet, allerdings konnte die Beteiligung eines anderen Transporters (HtaABCD) und einer Pyruvat-gekoppelten Aminotransferase (HtaE) festgestellt werden. Zwei weitere, bisher unbekannte Enzyme wurden für die Umwandlung des Intermediats 3-Sulfopropanoat zu 3-Sulfolactat entdeckt:

3-Sulfopropanoat-Dehydrogenase (SpuBCDA), deren Produkt als 3-Sulfopropenoat identifiziert wurde, und 3-Sulfopropenoat-Hydratase (SpuIJ), die letzteres vermutlich zu 3-Sulfolactat hydratisiert. Die Aktivitäten von Xsc und CuyA wurden als Bestätigung für den weiteren Abbau des intrazellulär gebildeten 3-Sulfolactats über den gegabelten Abbauweg (s. o.) betrachtet.

Bioinformatische Analysen ergaben, dass neun Organismen ein komplettes spu-Gen-Cluster für die Dissimilation von 3-Sulfopropanoat besitzen. Das Bakterium Rhodobacter sphaeroides 2.4.1, welches 3-Sulfopropanoat als Kohlenstoff- und Energiequelle verwertet, nutzt für die Desulfonierung wahrscheinlich Xsc. Es wurde aber kein Gen, das für die 3-Sulfopyruvat- Decarboxylase ComDE kodiert, gefunden. Stattdessen wurde ein Gen entdeckt, welches vermutlich eine neuartige 3-Sulfopyruvat-Decarboxylase (SpuE) kodiert. Diese Hypothese wird derzeit weiter untersucht.

Die in dieser Doktorarbeit entdeckten Abbauwege wurden als abschließende Schritte im Abbau von Sulfoquinovose interpretiert. Sulfoquinovose ist der polare Teil des ubiquitär vorkommenden pflanzlichen Sulfolipids, das ein wichtiges Intermediat des biologischen Schwefelkreislaufs darstellt. Diese Hypothese wurde durch bioinformatische Analysen unterstützt, die zeigten, dass die oben beschriebenen Abbauwege unter den Bakterien mit sequenzierten Genomen sehr weit verbreitet sind.

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CHAPTER 1: GENERAL INTRODUCTION

1.1 Organosulfonates

1.1.1 General characteristics

Organosulfonates are widespread - probably ubiquitous - compounds of both natural and anthropogenic origin. They are present in organisms of all three domains, where they have diverse and in some cases still unknown functions. Natural organosulfonates occur in terrestrial (e.g. Huxtable 1986; Autry and Fitzgerald 1990) as well as in marine environments (e.g.

Shibuya et al. 1963; Vairavamurthy et al. 1994) and in the atmosphere (Baker et al. 1991), and xenobiotic sulfonates, which are in use e.g. as laundry detergents, pharmaceuticals or dyes, are discharged in waste waters (e.g. Reemtsma et al. 2006).

Organosulfonates are characterized by a –SO₃^- moiety, which is covalently linked to an oxygen atom (R-O-SO3-), a nitrogen atom (R-N-SO3-) or a carbon atom (R-C-SO3-). Correspondingly, the compounds are then termed O-sulfonates (sulfate esters), N-sulfonates or C-sulfonates. The latter group is relevant for this work, and is meant whenever the term “sulfonate” is used in this text.

Sulfate esters and N-sulfonates can be hydrolyzed enzymatically, whereas the carbon-sulfur bond in the C-sulfonates is highly stable and resistant also to chemical hydrolysis (Wagner and Reid 1931). This indicates the need for a different, non-hydrolytic biochemistry in desulfonation reactions of C-sulfonates (Cook et al. 1999). Bacteria and fungi play an outstanding role in the metabolism of these compounds, because – to our current knowledge – they are the only organisms which can cleave the C-sulfonate bond (Huxtable 1992).

The oxidation state of the sulfur atom in organosulfonates has been reported as +4 (Huxtable 1992; Seitz et al. 1993), but data from X-ray absorption spectroscopy with several sulfonates provided firm evidence for +5 (Vairavamurthy et al. 1993).

The sulfonate group is a strong acid, with pKa values comparable to those of mineral acids (Huxtable 1992). Thus, organosulfonates are negatively charged over the physiological pH range and require transport across membranes (Huxtable 1992; Graham et al. 2002).

1.1.2 Naturally occuring organosulfonates

The smallest organosulfonate is methanesulfonoate (Fig. 1). It is formed by oxidation of dimethylsulfide in the atmosphere, and comes back to earth by wet and dry depostion.

Dimethylsulfide from algal and cyanobacterial origin is emitted from the oceans in a scale of

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several million tons per year, so methanesulfonate is a relevant part of the biogeochemical sulfur cycle (Baker et al. 1991; Kelly and Murrell 1999). The oceans, or rather their inhabitants, offer a variety of organosulfonates (Christophersen and Anthoni 1986): numerous marine invertebrates and fish use taurine (2-aminoethanesulfonate; Fig. 1) and N-methyltaurine as osmolytes (Huxtable 1992; Yin et al. 2000). Isethionate (2-hydroxyethanesulfonate; Fig. 1) occurs in the squid giant axon (Koechlin 1954). At least 20 sulfonated compounds were discovered in marine and freshwater algae, including e.g. (mono- and di-methylated) taurine(s) (Lindberg 1955), isethionate (Holst et al. 1994), sulfoacetate, homotaurine (3-aminopropanesulfonate), 3-sulfolactate (2-hydroxy-3-sulfopropanoate), cysteate (2-amino-3-sulfopropanoate) (see all in Fig. 1), and also more complex molecules such as sulfoquinovose (SQ, 6-deoxy-6-sulfoglucose; Fig. 1) (Shibuya et al. 1963; Miyazawa et al. 1970) or the diatom sulfolipid 1-deoxyceramide-1-sulfonate (Anderson et al. 1978).

Sulfoquinovose is the polar head group of the plant and algal sulfolipid (sulfoquinovosyl diacylglycerol, SQDG) (Benson et al. 1959; Daniel et al. 1961; Benson 1963). About one third of the plant sulfur is in SQ, which is biosynthesized annualy in amounts of 10¹⁰ tons – so again here, we have a major component of the biological sulfur cycle (Harwood and Nicholls 1979).

Organosulfonates are also present in the archaeal and bacterial kingdoms: coenzyme M (CoM, 2-mercaptoethanesulfonate; Fig. 1) is an essential cofactor in methanogenesis (McBride and Wolfe 1971) and alkene degradation (Allen et al. 1999). Capnine (2-amio-3-hydroxy-15- methylhexadecane-1-sulfonate) is a sulfolipid in the genus of the gliding bacteria Cytophaga (Godchaux III and Leadbetter 1984). Petrobactin sulfonate is a siderophore of the oil-degrading Marinobacter hydrocarbo noclasticus, an example for a naturally occuring aromatic organosulfonate (Hickford et al. 2004). These are rare, compared to the long list of known aliphatic sulfonates. Further examples are the echinosulfonates A-C, isolated from a marine sponge (Ovenden and Capon 1999), and aeruginosin B (Fig. 1), a red pigment from Pseudomonas aeruginosa (Herbert and Holliman 1964).

Organosulfonates occur free or derivatized. Taurine, for example, has several physiological functions in mammals (Huxtable 1992), taurocholate (Fig. 1) is an emulsifier in digestion (Gowda et al. 2009), N-acetyltaurine is a component of spiders’ webs (Vollrath et al. 1990), and the methylated taurine derivatives were detected in algae (see above) (Lindberg 1955).

Considering the vast abundance of organosulfonates in living organisms, their deposition in soils and sediments is not surprising. In forest soils, sulfonate sulfur is a major form of organic sulfur, irrespective of depth, and contributes up to 50 % of total sulfur (Autry and Fitzgerald 1990). Soil, peat and riverine humic acids are dominated by sulfonates (Morra et al. 1997), and even in reducing marine sediments, 20-40 % of the total organic sulfur is found in the oxidation state +5 (Vairavamurthy et al. 1994; Vairavamurthy et al. 1997). The large pool of sulfonate

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sulfur in soils is interpreted such, that the deposition rate must exceed the mineralization rate, which is in general expected to be rapid and exclusively biological (Autry and Fitzgerald 1990).

The organosulfonates which accumulate in marine sediments in depths down to 100 m are hypothesized to contribute to kerogenesis (Vairavamurthy et al. 1994).

Fig. 1. Naturally occurring and xenobiotic organosulfonates. 2-C10-LAS, one example of linear alkylbenzene sulfonates; arseno sugar, 2-hydroxy-3-sulfopropyl-5-deoxy-5-(dimethylarsenoso)- furanoside; ^a, xenobiotic organosulfonates.

1.1.3 Xenobiotic organosulfonates

Life without industrially produced organosulfonates would be far less comfortable than it is today: we would miss biodegradable laundry detergents, our clothes would be less bright and colourful, the foam from our hair and body shampoos would be less mild, and some pharmaceuticals would not exist to alleviate certain diseases.

Linear alkylbenzensulfonates (LAS; Fig. 1) are the major surfactants worldwide, and their annual use, mainly as laundry detergents, amounts to 430,000 tons in Western Europe (Jensen et al. 2007). Sulfonated derivatives of stilbene are also components of laundry formulations and act as fluorescent optical brighteners (McPherson and Omelczenko 1980). Frequently used textile, food and drug dyes are sulfonated azo dyes, such as Orange II (Fig. 1) or Ponceau BS (Chen et al. 2009).

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The C₃-compounds propanesulfonate and propane-1,3-disulfonate (Fig. 1) are mentioned here as representatives of aliphatic xenobiotic sulfonates. The former is in laboratory use as a component of HPLC mobile phases, the latter was in clinical testing as an orphan drug for the treatment of amyloid A amyloidosis (Kiacta^TM, formerly Fibrillex^TM; eprodisate), but the application for its accreditation was withdrawn in 2008 (Renders 2008).

1.2 C

3

-Sulfonates

1.2.1 Homotaurine

Homotaurine (Fig. 1) is a natural amino sulfonic acid which was discovered in 1970 in extracts of the marine red alga Grateloupia livida (Miyazawa et al. 1970) and later identified in several other red algal species (Ito et al. 1977).

As a homologue of taurine (see Chapter 1.3.1) and an analogue of the neurotransmitter 4-aminobutyrate (γ-aminobutyrate, GABA), homotaurine aroused the interest of the pharmaceutical industry and reached phase III clinical trials as a drug candidate (Alzhemed™;

tramiprosate) for the treatment of Alzheimer’s disease. In animal experiments it reduced the formation of amyloid fibrils and thus the deposition of amyloid plaques, which are one of the current explanations for brain dysfunction in Alzheimer’s disease (Aisen 2005). The results of the clinical trials, however, were inconclusive and the studies were stopped at that point.

Instead, the pharmaceutical company sells the compound now as active ingredient of a

“nutraceutical”, a dietary supplement, which is advertized to protect brain function (Swanoski 2009). Homotaurine was also considered for the treatment of hemorrhagic stroke (then named Cerebril™), but again here clinical testing was cancelled after phase II (annual information form 2007 of Neurochem Inc.). The N-acetylated derivative of homotaurine (Campral™;

acamprosate) is an accredited pharmaceutical to facilitate abstinence of detoxified alcoholics, with a daily dosage of 2 g (Scott et al. 2005). Furthermore, homotaurine and its derivatives are included in basic investigations for several other neurological, muscular, cardiovascular and anti-oxidative functions (Gupta et al. 2005). The surfactant 3-[(3-cholamidopropyl)- dimethylammonio]-1-propanesulfonate (CHAPS) and the “Good buffers” cyclohexyl- aminopropanesulfonate (CAPS) and 3-(N-morpholino)-propanesulfonate (MOPS) (Good and Izawa 1972) are further homotaurine derivatives, which are commonly used in laboratories.

Thus, homotaurine and its derivatives can be expected in the environment, especially in the oceans and in waste water.

As an amino sulfonic acid, homotaurine is a potential source of carbon, nitrogen and sulfur for bacteria. The utilization of homotaurine as sole source of sulfur by Corynebacterium glutamicum and Escherichia coli was reported by other groups, focussing on transport and desulfonation (see Chapter 1.4.1) (Eichhorn et al. 2000; Koch et al. 2005).

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Homotaurine was also studied as an alternative substrate for several enzymes. Taurine:pyruvate aminotransferase (Tpa, EC 2.6.1.77) from Bilophila wadswo rthia, catalyzing the initial deamination in taurine degradation, was found to act on homotaurine (Laue and Cook 2000a), as was 4-aminobutyrate transaminase (EC 2.6.1.19) from GABA-grown Pseudomonas fluorescens (De Gracia and Jollès-Bergeret 1973). The latter enzyme is also highly active with homohypotaurine, the corresponding sulfinate.

Furthermore, homotaurine is a substrate for tauropine dehydrogenases of several marine invertebrates (Kanno et al. 1996; 1997; Kanno et al. 1998) and of a marine red alga (Sato et al.

1993) – where homotaurine is available as a natural substrate (e.g. Ito et al. 1977).

1.2.2 (R)-Cysteate

(R)-Cysteate (L-Cysteat) (Fig. 1) is another widespread amino organosulfonate, whose natural occurence was first discovered in wool, where it was formed by weathering of cystine (dicysteine) residues (Consden et al. 1946). Later it was detected as an intermediate in the sulfur assimilation of the freshwater algae Chlorella and Scenedesmus (Shibuya et al. 1963), and also in several marine algae, together with the related compounds cysteinolate (2-amino- 3-hydroxypropanesulfonate) and its isomer 2-hydroxy-3-aminopropanesulfonate (Ito 1969;

Miyazawa et al. 1970; Ito et al. 1977). Besides other sulfonates, cysteate was also found in spiders’ webs (Fischer and Brander 1960).

The compound was also shown to be involved in bacterial sulfur metabolism: in Bacillus subtilis it is formed during sporulation as a precursor of 3-sulfolactate (Koshikawa et al. 1981), which is a major component of mature endospores (Bonsen et al. 1969). In Staphylococcus aureus strains it was found as an intermediate during the assimilation of inorganic sulfur sources, together with homocysteate and sulfopyruvate (Seltmann and Voigt 1977). In the genus of the gliding bacteria Cytophaga, derivatized cysteate was discovered in the sulfolipids capnine (Fig. 1) and its N-acylated variants (Godchaux III and Leadbetter 1984), and it was shown to be a precursor in their biosynthesis (White 1984). This group of sulfolipids, termed capnoids, were not only found in Cytophaga spp., but also in closely related organisms, such as Capnocytophaga, Flavobacterium or Flexibacter spp., and are major constituents of their cell envelopes (Godchaux III and Leadbetter 1980; 1983). Also the sulfolipid composition of the non-photosynthetic diatom Nitzschia alba contains significant amounts of similar sulfolipids (Anderson et al. 1978).

Only recently, an alternative branch of the CoM biosynthetic pathway was discovered in Methanosarcinales and Methanomicrobiales, which involves an (R)-cysteate synthase. It forms cysteate by β-elimination of phosphate from phosphoserine with subsequent addition of sulfite to the α,β-unsaturated intermediate deyhdroalanine (see Chapter 1.3.2) (Graham et al. 2009).

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In mammals, cysteate is present in the brain (10 nmol/g wet weight) (Ida and Kuriyama 1983), in plasma and urine (see Weinstein and Griffith 1988). Its origins there are supposed to be dietary – it can be formed during food processing from cyst(e)ine residues of proteins (Chang et al. 1982) – or biosynthetic. The latter case, however, is controversial: some reports show a formation of cysteate from cysteine or from 3’-phosphoadenosin-5’-phosphosulfate (PAPS) and dehydroalanine, whereas others contradict these findings or doubt their role in vivo (see Weinstein and Griffith 1988).

Wherever it comes from naturally, (R)-cysteate is in equilibrium with 3-sulfopyruvate and 3-(R)-sulfolactate in mammals. As analogues of (S)-aspartate (L-aspartate), oxaloacetate and (S)-malate (L-malate), they are interconverted by aspararate aminotransferase (EC 2.6.1.1) and malate dehydrogenase (EC 1.1.1.37). The depletion of this combined pool occurs by the irreversible decarboxylation of cysteate to taurine, or the excretion of 3-sulfolactate (Weinstein and Griffith 1988).

Cysteate and cysteine sulfinate were hypothesized to act as neurotransmitters (Thompson and Kilpatrick 1996), and evidence supporting physiological functions in neuronal activity has been collated (e.g. Croucher et al. 2001).

It remains to be noted, that in some of our research publications a mistake in the nomenclature of cysteate appeared (Denger et al. 2009; Denger and Cook 2010; Mayer et al. 2010): we wrongly designated L-cysteate as (S)-cysteate, which is in fact in the (R)-configuration. The same applies to L-cysteate sulfo-lyase, which is (R)-cysteate sulfo-lyase.

1.2.3 3-Sulfolactate

3-Sulfolactate (Fig. 1) is a widespread natural organosulfonate occurring in archea, bacteria, algae, plants and animals. In archea, it is an intermediate in CoM biosynthesis (see Chapter 1.3.2) (White 1986). In bacteria, it is a major component of endospores, where it accounts for 5 % of the dry weight (Bonsen et al. 1969). It was shown to be formed from cysteate (Koshikawa et al. 1981), and alternatively hypothesized to originate from phosphoenolpyruvate, catalyzed by B. subtilis’ homologues of the archaeal ComA and ComB (see Chapter 1.3.2) (Graham and White 2002). 3-Sulfolactate was observed as an intermediate in the sulfur assimilation in green algae (Shibuya et al. 1963). It also accumulated during the metabolism of glyceryl SQ in green plants, and was thus attributed to be an intermediate in the sulfoglycolytic pathway hypothesized by A. A. Benson (Lee and Benson 1972). A metabolism for sulfosugars comparable to the glycolytic (Embden-Meyerhoff-Parnas) pathway is also proposed to exist in bacteria, which are able to grow with SQ as sole source of carbon (Roy et al. 2003). Again here, 3-sulfolactate was observed, however, in this case as a minor end product.

In mammals, 3-sulfolactate is in an equilibrium with 3-sulfopyruvate and cysteate, and malate dehydrogenase is involved in their interconversion (see above) (Weinstein and Griffith 1988).

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In microorganisms, the oxidation of 3-sulfolactate to 3-sulfopyruvate was observed in the archaeal CoM biosynthesis (Graham and White 2002), and in several bacterial degradative pathways, and at least three types of sulfolactate oxidoreductases were found. The best- characterized is ComC (EC 1.1.1.272) from Methanocaldococcus jannaschii, which plays a role in the biosynthesis of CoM. It is specific for the (R)-enantiomer of 3-sulfolactate (Graupner et al. 2000a; Irimia et al. 2004). Only recently ComC was shown to be involved in bacterial sulfolactate degradation by Chromohalobacter salexigens . Together with SlcC, an (S)-sulfolactate dehydrogenase (EC 1.1.1.-), it effects a racemase, enabling the degradation of both enantiomers of racemic 3-sulfolactate (Denger and Cook 2010). A third sulfolactate oxidoreductase is the membrane-bound SlcD (EC 1.1.99.-), which we identified in sulfolactate degradation in Roseovarius nubinhibens ISM (see Chapter 5).

1.2.4 3-Sulfopropanoate

3-Sulfopropanoate (Fig. 1) is rarely mentioned in the literature, and besides a report about its chemical synthesis (Kharasch and Brown 1940), there is only the postulation that it might be the product of subsequent transamination and oxidation of homotaurine by GABA aminotransferase (see Chapter 1.2.1) and putative succinate semialdehyde dehydrogenase (EC 1.2.1.16) (De Gracia and Jollès-Bergeret 1973).

1.2.5 2,3-Dihydroxypropane-1-sulfonate

2,3-Dihydroxypropane-1-sulfonate (DHPS, sulfopropanediol) (Fig. 1) was identified in 1966 in the fresh-water diatom Navicula pelliculosa (Busby 1966). Previously, it was tentatively identified as an intermediate in algal sulfur metabolism (Shibuya et al. 1963). Few years later, it was established as ubiquitous in diatoms and detected in the plant kingdom (Lee and Benson 1972). Then it was attributed to be involved in the hypothesized sulfoglycolytic pathway in plants (Benson and Lee 1972; Lee and Benson 1972).

The need to investigate the biodegradability of DHPS as a representative of sulfonates of plant origin was recognized in 1967. H. L. Martelli showed its utilization as sole source of carbon and sulfur by aquatic bacteria. Thereby, she found excess ³⁵S from the radiolabelled substrate accumulating as sulfate and an unknown degradation product (Martelli 1967). This was not investigated any further, but several decades later, DHPS was detected as a transient excretion product in the bacterial degradation of SQ. As with 3-sulfolactate, DHPS was then considered to be part of bacterial sulfoglycolysis (Roy et al. 2003).

A noteworthy sugar derivative of DHPS is 2-hydroxy-3-sulfopropyl-5-deoxy-5-(dimethylarsenoso)-furanoside (Fig. 1). It was discovered in the brown alga Ecklonia radiata, which

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accumulates arsenic in high concentrations, and is hypothesized to play a role in the cycling of arsenic in marine ecosystems (Edmonds and Francesconi 1981).

1.3 Further relevant organosulfonates

1.3.1 Taurine

Discovery and general characteristics

During the past 180 years, a large variety of free or derivatized organosulfonates was discovered in nature, and a remarkable lineup of sulfonated compounds was evolved by mankind. The first sulfonate was isolated from ox bile in 1827 by two German chemists (Tiedemann and Gmelin 1827). At that time, the compound’s –SO3- moiety remained undiscovered, but its nitrogen content and similarities to the amino acid asparagine were observed. Thus, Tiedemann and Gmelin termed their isolate “Gallen-Asparagin” (bile-asparagine) and described it as stable to air, heat, acids and bases (Tiedemann and Gmelin 1827). Few years later, it appears in the literature with a new name, which is well-known today: taurine (Demarçay 1838).

Taurine (2-aminoethanesulfonate, Fig. 1) is one of the most prominent and best-characterized organosulfonates, and probably the best example to reflect the ubiquity and the versatility of this class of compounds.

Taurine was detected in high concentrations (µmol/g wet weight) in the animal kingdom (with the exception of protozoans); in the human body it is present in considerable amounts of 1 g per kg body weight (Huxtable 1992). Further rich sources of taurine and its N-(di-)methylated derivatives are red algae (e.g. Lindberg 1955; Shibuya et al. 1963), and the viscid droplets of spiders’ webs, where it is found in molar concentrations together with its N-acetyl derivative and isethionate (2-hydroxyethanesulfonate) (Vollrath et al. 1990; Townley et al. 2006).

In mammals, taurine is a major organic solute present in many tissues such as brain, heart and muscles, kidney, liver, spleen, retina and blood cells (Jacobsen and Smith 1968; Wright et al.

1986). It has functions e.g. in developmental processes, in photoreceptors, in the modulation of neuronal excitability and osmoregulation (Huxtable 1992). Besides the dietary uptake, many animals can biosynthesize taurine (see below) (Stipanuk 2004), but they can not metabolize it any further, as they are not able to cleave the carbon-sulfur bond. Thus, conjugation to form emulsifiers for digestion (taurocholate, Fig. 1) (Gowda et al. 2009) and/or excretion in e.g.

urine, faeces or tears, are the only possibilities to remove excess taurine (Huxtable 1992). The released taurine is readily degraded by bacteria, which was found in 1926 (den Dooren de Jong 1926), and later studied in detail (e.g. Kondo et al. 1971; Ruff et al. 2003).

The industrial production of taurine as additive for “energy” drinks (Clauson et al. 2008) and commercial cat food (Morris et al. 1990), and of taurine derivatives for hair and body care products (e.g. Johnson 1978), is large.

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Taurine biosynthesis

Two mechanisms are known for the biosynthetic introduction of the sulfonate moiety. The nucleophilic addition of sulfite at (transiently formed) unsaturated carbon bonds takes place in the biosynthesis of SQ (see Chapter 1.3.3) (Mulichak et al. 1999; Sanda et al. 2001) and in the two routes to CoM (via 2-phospho-3-sulfolactate or cysteate) (Graham et al. 2002; Graham et al. 2009). The main mechanism in taurine biosynthesis is the oxidation of thiol groups, however, a reaction involving sulfite was also observed (Chapeville and Fromageot 1957;

Tolosa et al. 1969).

In addition to dietary uptake, man biosynthesizes about 200 to 400 µmol of taurine per day (Irving et al. 1986). With exceptions among omnivores and carnivores (e.g. cats), mammals are able to produce sufficient taurine (Huxtable 1992). Two closely related biosynthetic pathways are established in mammals, which involve (i) the oxidation of the sulfur-containing amino acid (R)-cysteine (L-cysteine) (Huxtable 1986; Stipanuk 2004), and (ii) the oxidation of cysteamine (Huxtable 1986; Dominy Jr. et al. 2007) by two different thiol dioxygenases (Fig. 2).

(R)-Cysteine is oxidized by cysteine dioxygenase (EC 1.13.11.20) to cysteine sulfinate, which is decarboxylated to hypotaurine (2-aminoethanesulfinate) by cysteine sulfinate decarboxylase (EC 4.1.1.29) (Fig. 2). Both enzymes are predominantly located in the liver, which is the main taurine production site (Huxtable 1986; Stipanuk 2004). Hypotaurine is possibly oxidized to taurine by another hepatic enzyme (Brand et al. 1998): NAD⁺-dependent hypotaurine dehydrogenase activity (EC 1.8.1.3) was observed in rat liver homogenates (Sumizu 1962), a reaction which has never been confirmed.

The oxidation of cysteamine to hypotaurine by cysteamine dioxygenase (EC 1.13.11.19) is the second pathway of taurine biosynthesis in mammals (Dominy Jr. et al. 2007) (Fig. 2).

Cysteamine is ubiquitous in every cell, as it is constitutively formed in coenzyme A degradation (Robishaw and Neely 1985). Cysteamine dioxygenase and cysteine dioxygenase are the only known mammalian thiol dioxygenases, and each of them is highly specific for its substrate, cysteamine or cystein, respectively (Chai et al. 2005; Dominy Jr. et al. 2007). In contrast to the localization of cysteine dioxygenase in liver tissue, cysteamine dioxygenase is ubiquitously expressed, with the highest levels in brain, heart and skeletal muscle (Dominy Jr. et al. 2007).

Apart from the apparently different spatial distribution of the two taurine biosynthetic pathways, their relative contribution to the taurine pool remains unclear (Stipanuk et al. 2006).

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Fig. 2. Taurine biosynthetic pathways: the two mammalian alternatives via cysteine sulfinate and cysteamine are established pathways (Robishaw and Neely 1985; Stipanuk 2004; Dominy Jr. et al. 2007), whereas algal taurine formation from sulfoquinovose is hypothesized from the sulfonated intermediates observed (e.g. Shibuya et al. 1963). The route via cysteine lyase was exclusively found in the yolk sac of chicken embryos (Chapeville and Fromageot 1957; Tolosa et al. 1969).

In contrast, taurine biosynthesis via cysteate is possibly the major pathway in the yolk sac of chicken embryos (Chapeville and Fromageot 1957). Again here, cysteine is the precursor, however, its thiol group is not oxidized by cysteine dioxygenase, but exchanged for sulfite by cysteine lyase (EC 4.1.1.10) (Tolosa et al. 1969; Braunstein and Goryachenkova 1976). The products of this pyridoxal 5’-phosphate (PLP)-dependent β-replacement reaction are hydrogen sulfide and cysteate. The latter is decarboxylated to taurine by a cysteate-specific enzyme – cysteine sulfinate is not a substrate for this decarboxylase (Chapeville and Fromageot 1957).

Until today reports about cysteine lyase in other organisms are limited to localization studies in

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several vertebrate embryos (Fischer et al. 1982), and the physiological relevance of this biosynthetic pathway is doubted (Stipanuk 2004).

The origin of the considerable amounts of taurine and its derivatives in marine and freshwater algae (e.g. Lindberg 1955; Shibuya et al. 1963; Ito et al. 1977) is unknown. Cysteine can be expected as a precursor, but so far none of the relevant enzymes mentioned above was found in the plant kingdom. The only sulfonation mechanism known in plants is the addition of sulfite to an unsaturated UDP-glucose derivative, which yields SQ (e.g. Mulichak et al. 1999). The metabolism of the latter via the hypothesized sulfoglycolytic pathway (see below) would generate a range of sulfonated C3-compounds, many of which were already detected in algae (e.g. Shibuya et al. 1963). 3-Sulfolactate is one example, and its conversion to taurine via 3-sulfopyruvate and cysteate, which was also identified in algae (Shibuya et al. 1963; Ito 1969), seems feasible (Fig. 2).

1.3.2 Coenzyme M and its biosynthesis

Coenzyme M (CoM, 2-mercaptoethanesulfonate; Fig.1) is another important representative of the C₂-sulfonates. It was discovered in 1971 as one of the seven coenzymes involved in methanogenesis and is the smallest organic cofactor (MW 142 g/mol) that is known today. “M”

stands for “methyl transfer”, as CoM is the terminal methyl carrier which releases methane (McBride and Wolfe 1971). For almost three decades CoM was considered to be unique to methanogenic archea, but in 1999 it was found as a coenzyme in bacterial alkene degradation.

Here, CoM assists with the carboxylation of the alkyl epoxide, which results from the initial monooxygenation of the alkene substrate (Allen et al. 1999).

Furthermore, human medicine benefits from CoM (“Mesna”), which has mucolytic and cytoprotective effects during chemotherapy (e.g. Shaw and Graham 1987; Fowler et al. 2009).

The biosynthesis of CoM is well studied (e.g. Graham and White 2002; Graham et al. 2009), and involves several sulfonates and enzymes which are relevant for this work. The elucidation of the CoM biosynthetic pathway(s) (Fig. 3) started in R. H. White’s laboratory in 1985. First radiolabelling experiments allowed White to deduce that the sulfonate moiety is introduced by a nucleophilic addition of sulfite to phosphoenolpyruvate, and that the thiol group originates from cysteine (White 1985; 1986). Further, 3-sulfolactate, 3-sulfopyruvate and sulfoacetaldehyde were detected as intermediates (White 1986), and one by one, the enzymes ComA, ComB, ComC and ComDE could be established in Methanocaldococcus (formerly Methanococcus) jannaschii (Graham and White 2002). (2R)-Phospho-3-sulfolactate synthase (ComA; EC 4.4.1.19) was found to catalyze the proposed initial Michael addition, which is one of the few known sulfonation mechanisms (Graham et al. 2002). (R)-Sulfolactate dehydrogenase (ComC;

EC 1.1.1.272), which oxidizes the (R)-enantiomer of 3-sulfolactate and other L-2-hydroxy- carboxylic acids such as (S)-malate and (S)-lactate (Graupner et al. 2000a; Graham and White

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2002), is not only involved in a biosynthetic process, but also in one of the bacterial biodegradative pathways for racemic 3-sulfolactate (Denger and Cook 2010). The next step in CoM biosynthesis is the decarboxylation of 3-sulfopyruvate by 3-sulfopyruvate decarboxylase (ComDE; EC 4.1.1.79) (Graupner et al. 2000b), a reaction that enables the transition from C₃- to C2-sulfonate metabolism.

The further reactions that introduce the thiol group of CoM are not yet understood (Graham and White 2002; Graham et al. 2009). Cysteine is supposed to be involved, but it is unclear, if it directly reacts with sulfoacetaldehyde to form a thiazolidine intermediate, or if it supplies sulfur to a biosynthetic enzyme (White 1986; Graham and White 2002).

Methanococcales, Methanobacteriales and Methanopyrales (class I methanogens) have homologues of the comABCDE genes, whereas Methanomicrobiales and Methanosarcinales (class II methanogens) have no homologs of comABC genes – still the latter are able to produce CoM (Graham et al. 2009). Only recently, a paralogue of threonine synthase from Methanosarcina acetivorans was discovered to catalyze the PLP-dependent formation of cysteate from (S)-phosphoserine (L-phosphoserine) and sulfite (Graham et al. 2009). The hypothesized reaction mechanism proceeds via β-elimination of phosphate from (S)-phosphoserine and the subsequent β-addition of sulfite to intermediate dehydroalanine. This enzyme would enable an alternative route for CoM synthesis (Fig. 3B), together with the deamination of cysteate, which is attributed to (S)-aspartate aminotransferase (Weinstein and Griffith 1988; Helgadóttir et al. 2007).

The bacterium Xanthobacter autotrophicus, which uses the coenzyme in propylene degradation (Allen et al. 1999), has several gene candidates for ComA, but not for the further enzymes of Graham’s pathway(s). Thus, at least a third CoM biosynthetic pathway can be expected (Graham et al. 2009).

Fig. 3. Proposed coenzyme M biosynthetic pathways in class I (A) and class II methanogens (B) adapted from Graham and co-workers (Graham et al. 2009). CS, cysteate synthase; 2OG, 2-oxoglutarate; AspAT, aspartate aminotransferase; for further abbreviations see main text.

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1.3.3 Sulfoquinovosyl diacylglycerol and sulfoquinovose

The plant sulfolipid: Sulfoquinovosyl diacylglycerol

In 1959 A. A. Benson and co-workers discovered a sulfolipid in a large range of higher plants and photosynthetic microorganisms (Benson et al. 1959). The identity of the polar headgroup was recognized two years later as sulfoquinovose (SQ; 6-deoxy-6-sulfoglucose) (Daniel et al.

1961), and thus the sulfolipid could be established as sulfoquinovosyl diacylglycerol (SQDG;

Fig. 4) (Benson 1963). The fatty acid components are predominantly linolenate and palmitate, but linoleic, oleic and stearic residues were also observed (Benson 1963; O'Brien and Benson 1964). The sulfolipid is present in photosynthetic tissues of all plants and algae, in photosynthetic bacteria and some flagellates (Benson 1963; Harwood and Nicholls 1979). These findings implied that SQDG is directly involved in photosynthesis (Barber 1986), but this hypothesis was later abandoned (Benning et al. 1993; Güler et al. 1996). The sulfolipid was also found in a few non-photosynthetic bacteria, e.g. Sinorhizobium (formerly Rhizobium) meliloti (Cedergren and Hollingsworth 1994), in the non-photosynthetic diatom Navicula alba (Anderson et al. 1978), in germ cells of a sea urchin (Isono et al. 1967) and in chlorophyll-free plant tissues (Benson 1963), however, in lower concentrations. Exceptions among phototroph organisms which lack SQDG are several bacterial species, the cyanobacterium Gloeobacter violaceus is one example (Selstam and Campbell 1996). This supports the idea that the sulfolipid function is not in photosynthesis, at least not directly or exclusively. Today SQDG is attributed to stabilize chloroplast structure and function, and to substitute for phospholipids during phosphate limitation in e.g. Arabidopsis (Yu and Benning 2003) or bacteria, such as Rhodobacter sphaeroides (Benning et al. 1993), and to serve as an internal sulfur source for protein biosynthesis during sulfur starvation in a green alga (Sugimoto et al. 2007). Furthermore SQDG is involved in the salt stress response in the halophyte Crithmum maritinum (Ben Hamed et al. 2005).

Compared to other organisms, plants represent a large reservoir of sulfur. In leaf tissues, organic sulfur is almost exclusively distributed among protein and sulfolipid, which contain comparable amounts of sulfur. SQDG concentrations observed in photosynthetic plant tissues are 1 to 6 mM (Benson 1963), the proportion of SQDG in membrane lipids of several algae range from 30 to 50 mol% (Heinz 1993). The enormous annual biosynthesis of SQDG (3.6 x 10¹⁰ tons) and its periodic deposition to the soil by deciduous plants underlines the importance of this compound in the sulfur cycle (Harwood and Nicholls 1979).

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Biosynthesis of sulfoquinovose and the sulfolipid

The structure of the plant sulfolipid was established soon after its discovery (Benson 1963), but the elucidation of its biosynthesis was a long quest of almost 50 years (Benning 2007). Three major pathways contribute to the biosynthesis of SQDG: carbohydrate metabolism, sulfur assimilation, and fatty acid biosynthesis – and again here the central question is the sulfonation reaction (Benning 1998).

Several hypotheses were developed, beginning with A. A. Benson’s first idea, that the sulfonate group could arise from the peroxidation of a disulfide link (Benson et al. 1959), followed by his suggestion of a gluconeogenetic pathway for the formation of SQ (e.g. Benson and Shibuya 1961; Benson 1963). This hypothesis was based on and strengthened by the results of

35S-radiolabelling experiments with various algae (e.g. Shibuya et al. 1963; Busby 1966). These experiments revealed several sulfonated analogues of glycolytic intermediates and derivatives thereof, such as sulfolactate, sulfolactaldehyde, dihydroxypropanesulfonate, cysteate, cysteinolate and SQ (see Fig. 5). For the initial reaction Davies and co-workers proposed the sulfonation of phosphoenolpyruvate (Davies et al. 1966) – a reaction which is known today to occur in CoM biosynthesis (Graham et al. 2002).

Results from studies with leaves of a higher plant (the groundnut, Arachis hypogaea), however, did not support the hypothesized sulfogluconeogenetic pathway, as the compounds listed above were formed in negligible amounts only (Gupta and Sastry 1988). The discovery of a nucleoside diphosphate sulfosugar in the ³⁵S-labelled algae fostered a theory, which proposed a sulfonation reaction at a nucleotide anhydrosugar derivative (Shibuya et al. 1963). This latter hypothesis pointed to the right direction: about 40 years later, a uridine 5’-diphosphate (UDP)-SQ synthase (SQD1; EC 3.13.1.1) was identified and characterized in Arabidopsis thali ana (Sanda et al.

2001). The enzyme forms UDP-SQ from UDP-glucose and sulfite (Fig. 4), which originates from the sulfate reduction pathway. Its mechanism is supposed to involve a tightly bound NAD⁺, which assists to form a UDP-4-ketoglucose-5-ene intermediate, at which the addition of sulfite takes places (Fig. 4, grey box) (Pugh et al. 1995; Essigmann et al. 1999; Mulichak et al.

1999). The gene encoding UDP-SQ synthase [SQD1; sqdB in (cyano)bacteria] is highly conserved in photosynthetic organisms (Benning 2007). The last step in sulfolipid biosynthesis is catalyzed by UDP-SQ:diacylglycerol sulfoquinovosyl transferase (SQD2, SqdX in cyanobacteria), which links the UPD-activated sulfosugar to the diacylglycerol moiety (Fig. 4) (Seifert and Heinz 1992). This enzyme is located in the inner chloroplast membrane and could not be purified so far (Tietje and Heinz 1998). Work with a SQD2 knock-out mutant, however, confirmed its function and indicated the relevance of the sulfolipid in phosphate starvation (Yu et al. 2002).

Only recently a UDP-glucose pyrophosphorylase (UGP3; EC 2.7.7.9) involved in sulfolipid biosynthesis was identified in Arabidopsis chloroplasts (Okazaki et al. 2009). This finding

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solved the remaining question of UDP-glucose supply within the chloroplast, where SQD1 and SQD2 are located (Benning 2007). UPG3 catalyzes the activation of glucose 1-phosphate with uridine 5’-triphosphate (UTP) to UDP-glucose (Fig. 4). It is supposed to be conserved in plants, which supports its relevance in sulfolipid biosynthesis (Okazaki et al. 2009).

Biodiversity in SQDG synthesis is, however, indicated in bacteria: while cyanobacteria use the pathway as described above, Rhodobacter sphaeroides seems to have an additional route to form the sulfolipid via sulfoquinovosyl dihydroxyacetone (Rossak et al. 1997; Benning 2007).

Fig. 4. Biosynthetic pathway for sulfoquinovosyl diacylglycerol. The mechanism of UDP-SQ synthase is depicted in the grey box. Sulfite is supposed to derive from the sulfate reduction pathway in the chloroplast. R1 and R2 represent acyl chains of different length and degree of saturation. Adapted from:

(Seifert and Heinz 1992; Benning 1998; Essigmann et al. 1999; Mulichak et al. 1999; Sanda et al. 2001;

Okazaki et al. 2009).

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Degradation of SQ and the sulfolipid: the sulfoglycolytic pathway(s)

Considering the vast amounts of sulfolipid biosynthesized each year, the elucidation of (a) degradative pathway(s) for the sulfolipid, or SQ respectively, is of great importance in order to close a significant gap in the biological sulfur cycle. Still, little is known on this field, but all available results support the hypothesis of SQ degradation by a sulfoglycolytic pathway (Fig. 5) (Benson and Shibuya 1961; Benson and Lee 1972). SQDG catabolism in plants begins with deacylation by an acyl hydrolase which forms sulfoquinovosyl glycerol (Burns et al. 1977;

1980). The latter, although it is the α-anomer, was shown to be hydrolyzed by a β-galactosidase to glycerol and SQ (Shibuya and Benson 1961). The formation of SQ, sulfolactaldehyde and sulfolactate from radiolabelled sulfoquinovosyl glycerol was observed in leaves of alfalfa (Medicago sativa ) (Lee and Benson 1972). As analogues of glucose 6-phosphate, glyceraldehyde 3-phosphate and 3-phosphoglycerate (see Fig. 5), these intermediates support the idea of sulfoglycolysis in plants, as do the earlier observations of sulfonic acid analogues of glycolytic intermediates in algae (see above) (Shibuya et al. 1963; Busby 1966).

To our present knowledge, desulfonation reactions are attributed to microorganisms. In fact, forest soil communities degrade SQ to sulfate and sulfate esters (Strickland and Fitzgerald 1983), and several bacterial isolates (Klebsiella sp., Agrobacterium sp., and several Pseudomonas spp.) utilize the sulfosugar as sole source of carbon and energy (Roy et al. 2000;

Roy et al. 2003). Some of these isolates thereby excrete sulfate (Roy et al. 2000; Roy et al.

2003), others accumulate dihydroxypropanesulfonate (transiently) and 3-sulfolactate (Roy et al.

2003), or a so far unidentified sulfonate (Roy et al. 2000). The degradation of methyl-SQ by a Flavobacterium sp. yielded intracellular cysteate and sulfoacetate, as well as sulfate, which was excreted (Martelli and Benson 1964). Again here, the occurrence of C₃- and C₂-sulfonate intermediates indicates a sulfoglycolytic pathway for SQ degradation in bacteria. Roy and co- workers furthermore initiated assays of enzymes involved in this proposed bacterial pathway (Roy et al. 2003): crude extracts of SQ-grown Agrobacterium sp. strain ABR2 and Klebsiella sp. strain ABR11 contained an apparent sulfoquinovose kinase activity (possibly 6-deoxy- 6-sulfofructokinase), when fructose 6-phosphate or glucose 6-phosphate was replaced with SQ in the phosphofructokinase assay. The presence of this enzyme activty indicates degradation of SQ by a sulfoglycolytic, i.e. an Emden-Meyerhof-Parnas type, pathway. Extracts of Klebsiella sp. strain ABR11 furthermore showed an exclusively NAD⁺-dependent sulfoquinovose dehydrogenase activity, which was clearly distinguishable from the exclusively NADP⁺- dependent glucose 6-phosphate dehydrogenase activity. Roy et al. interpreted the presence of SQ dehydrogenase activity such, that Klebsiella sp. additionally has an Entner-Doudoroff type pathway for the degradation of SQ (Fig. 6) (Roy et al. 2003). The initial oxidative reaction might, however, also enable a route analogous to the phosphoketolase pathway (Fig. 6) (Cook et al., manuscript in preparation). Both options would generate 3-sulfolactaldehyde from SQ.

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So far, no speculations about yields of ATP and reduction equivalents were made. Considering that SQ does not need the initial activation by glucokinase in the preparatory phase, and assuming that the putative formation of 3-sulfolactate from 1-phospho-3-sulfolactate generates one ATP, the net output of energy would be the same as for glucose in all three pathways mentioned above.

The hypothesized sulfoglycolytic pathway in plants and algae (and its putative alternative routes in bacteria) are supposed to be sources of several C3-sulfonates relevant for this work.

3-Sulfolactate (Shibuya et al. 1963; Lee and Benson 1972; Roy et al. 2003), 2,3-dihydroxypropanesulfonate (Busby 1966; Benson and Lee 1972), and possibly cysteate (Shibuya et al. 1963) and homotaurine (Cook et al. , manuscript in preparation) can be considered as metabolites of SQ (see Fig. 5).

Fig. 5. The hypothesized sulfoglycolytic (Embden-Meyerhof-Parnas type) pathway adapted from Roy and co-workers (Roy et al. 2003) (upper line and columns 2-4) and C3-sulfonates which are presumed to derive from it (column 1). Column 4 shows the energy-yielding phase of the Embden-Meyerhof-Parnas pathway [adapted from a textbook (Kröger 1999)], which is also the presumed fate of DHAP released from 6-deoxy-6-sulfofructose 1-phosphate. The sulfoglycolytic pathway is presumed to exist in plants and algae (e.g. Shibuya et al. 1963; Lee and Benson 1972) and bacteria (Roy et al. 2003), but the desulfonation reactions are attributed exclusively to bacteria (Roy et al. 2003; Denger et al. 2009; Mayer et al. 2010; see Chapters 5 and 6). The bacterial degradation of dihydroxypropanesulfonate and 3-sulfolactate will be presented in Chapters 5 and 6. DHAP, dihydroxyacetone phosphate; GAP, glyceraldehyde 3-phosphate; 1,3-bis-PG, 1,3-bisphosphoglycerate; 3-PG, 3-phosphoglycerate.

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1.4 Utilization of C

₃

-sulfonates by bacteria

1.4.1 Homotaurine as a substrate for bacterial growth

Reports about the utilization of homotaurine as a nutrient for microorganisms are rare: (i) the nitrate-reducer Paracoccus pantotro phus NKNCYSA utilizes homotaurine under anoxic conditions as sole electron and carbon source, thereby excreting ammonia, sulfate and presumably CO₂ (Mikosch et al. 1999), and (ii) E. coli and C. glut amicum assimilate homotaurine sulfur under conditions of sulfate and cysteine starvation (Eichhorn et al. 2000;

Koch et al. 2005). Two members of the ATP-binding cassette (ABC) transporter superfamily, TauABC and SsuABC (Ssu for sulfonate-sulfur utilization), are responsible for homotaurine uptake under sulfur-limited conditions (see Chapter 1.6.1) (Eichhorn et al. 2000). Desulfonation is attributed to TauD, a 2-oxoglutarate-dependent taurine-dioxygenase, (Eichhorn et al. 1997) and SsuD, an FMNH₂-dependent alkanesulfonate monooxygenase (see Chapter 1.4.4) (Eichhorn et al. 1999; Koch et al. 2005).

Fig. 6. SQ degradation via the hypothesized Entner-Doudoroff type pathway (Roy et al. 2003) and the presumed alternative route (Cook et al. , manuscript in preparation), which is analogous to the phosphoketolase pathway. Evidence for a sulfoquinovose dehydrogenase is available (Roy et al. 2003).

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1.4.2 Cysteate: a versatile substrate

Containing carbon, nitrogen and sulfur, cysteate is a potential source of these three macroelements for microorganisms. And indeed, Stapley and Starkey showed in 1970 that cysteate can serve as sole source of carbon and energy, nitrogen and sulfur for aerobic bacterial isolates (Stapley and Starkey 1970). They observed the excretion of excess ammonia, sulfite and sulfate, and expected the involvement of an adaptive enzyme system. The elucidation of degradative pathways started about 30 years later, when cysteate oxidation was investigated in P. pantotrophus NKNCYSA. The organism utilizes (R)-cysteate as sole source of carbon and energy in respirations with either nitrate or molecular oxygen (Mikosch et al. 1999; Rein et al.

2005). The core of the pathway is the desulfonation reaction, which is catalyzed by an Fe²⁺- dependent altronate deydratase-like enzyme, 3-sulfolactate sulfo-lyase (SuyAB; EC 4.4.1.24).

SuyAB was shown to yield pyruvate and sulfite from 3-sulfolactate (Fig. 7A) (Rein et al. 2005), which was later specified as (R)-sulfolactate (Denger and Cook 2010). SuyAB is incuced upon growth with cysteate and consists of 8- and 42-kDa subunits (Rein et al. 2005). (R)-sulfolactate originates from the conversion of cysteate to 3-sulfopyruvate, and reduction of the latter by constitutively expressed enzymes (Fig. 7A). It is not clear, if these reactions are catalyzed by aspartate aminotransferase (EC 2.6.1.1) and malate dehydrogenase (EC 1.1.1.37), or if distinct cysteate:2-oxoglutarate aminotransferase (EC 2.6.1.-) and (R)-sulfolactate dehydrogenase (e.g.

ComC) exist. The identity of the transporter for cysteate also remained unclear. An inducible sulfite dehydrogenase (EC 1.8.2.1) oxidizes sulfite released by SuyAB to sulfate (Fig. 7A), whose excretion is attributed to SuyZ, a putative sulfate exporter.

Work with Ruegeria (formerly Silicibacter) pomeroyi DSS-3^T revealed another pathway of aerobic cysteate dissimilation (Fig. 7B), which seems more widespread (Denger et al. 2006b):

cysteate is directly desulfonated by the PLP-dependent (R)-cysteate sulfo-lyase (CuyA; EC 4.4.1.25). The enzyme, which is inducible upon growth with cysteate, catalyzes the formation of pyruvate, sulfite and ammonium from (R)-cysteate. A major portion of the released sulfite is excreted directly, only a minor portion is oxidized to sulfate, although R. pomeroyi expresses a sulfite dehydrogenase during growth with cysteate. CuyZ, an orthologue of the putative sulfate exporter SuyZ, is attributed to transport sulfite out of the cell. The genes encoding CuyA and CuyZ are located in an operon (cuyAZ), which is assumed to be controlled by the LysR-type transcriptional regulator CuyR (Denger et al. 2006b). Again here, cysteate uptake is mediated by an unknown transporter, whereas the excretion of the ammonium ion is supposed to occur via an ortholog of the AmtB (ammonia/methylammonia) transporter (Khademi et al. 2004).

Besides its utilization in aerobic or nitrate-reducing metabolisms, cysteate can also act as an electron acceptor itself (Lie et al. 1996; Laue et al. 1997b), and in fermentations it was found to serve as a sulfur source (Chien et al. 1995) or as a carbon source, with the dismutation of the sulfonate moiety to sulfide and sulfate (Laue et al. 1997a).

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Fig. 7. Dissimilatory pathways of (R)-Cysteate and (racemic) 3-sulfolactate in P. p antotrophus NKNCYSA (A) and in R. pomeroyi DSS-3 (B), and of racemic 3-sulfolactate in C. salexigens DSM 3043 (C). 1, undefined (R)-cysteate transporter; 2, (R)-cysteate:2-oxoglutarate aminotransferase [possibly (S)-aspartate:2-oxoglutarate aminotransferase]; 3, (S)-glutamate dehydrogenase; 4, (R)-sulfolactate dehydrogenase [possibly (S)-malate dehydrogenase]; 5, undefined sulfite dehydrogenase; 6, undefined ammonium exporter (possibly AmtB); 7, putative sulfate exporter; 8, undefined sulfolactate transporter;

9, undefined sulfite exporter. Further abbreviations: see main text. Adapted from (Cook et al. 2006) (A, B) and (Denger and Cook 2010) (C).

Microbial desulfonation pathways for natural and pharmacologically relevant C 3-sulfonates