The chemical biology of dimethylsulfoniopropionate.

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Cite this: Org. Biomol. Chem., 2015, 13, 1954

Received 14th November 2014, Accepted 18th December 2014 DOI: 10.1039/c4ob02407a www.rsc.org/obc

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The chemical biology of dimethylsulfoniopropionate† Jeroen S. Dickschat,*a,b Patrick Rabea,b and Christian A. Citrona,b Dimethylsulfoniopropionate is a highly abundant sulfur metabolite in marine ecosystems. Its biosynthesis by different organisms including plants, marine algae and dinoflagellates is discussed. Furthermore, the accumulated knowledge about bacterial uptake systems and its climatically relevant degradation by marine bacteria to methanethiol or dimethylsulfide is presented. Finally, uptake and degradation of synthetic DMSP analogs are addressed.

Introduction Dimethylsulfoniopropionate (DMSP, Scheme 1) was first isolated by Challenger and Simpson from the red algae Polysiphonia fastigiata and P. nigrescens1 and is today known to be

a Kekulé-Institut für Organische Chemie und Biochemie, Rheinische FriedrichWilhelms-Universität Bonn, Gerhard-Domagk-Straße 1, 53121 Bonn, Germany. E-mail: [email protected]; Fax: +49 228 735813; Tel: +49 228 735797 b Institut für Organische Chemie, Technische Universität Braunschweig, Hagenring 30, 38106 Braunschweig, Germany † Electronic supplementary information (ESI) available: Tabulated accession numbers of genes involved in DMSP demethylation (Table 1) and cleavage of DMSP (Table 2). See DOI: 10.1039/c4ob02407a

Jeroen S. Dickschat studied Chemistry in Braunschweig. During his PhD with Prof. Stefan Schulz he worked on bacterial volatiles as a fellow of the Fonds der Chemischen Industrie. In 2005 he moved to Saarland University for a postdoctoral stay with Prof. Rolf Müller, followed by a postdoc at Cambridge University with Prof. Peter Leadlay as fellow of the Deutsche Akademie der Naturforscher LeopolJeroen S. Dickschat dina. In 2008 he was awarded an Emmy Noether fellowship which offered the opportunity to return to Braunschweig for his independent career. In 2013 he was an interim professor at the University of Marburg and since 2014 he is professor at the University of Bonn. His research interests include the synthesis, structure elucidation, biosynthesis and function of microbial natural products.

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Scheme 1

Structure of dimethylsulfoniopropionate (DMSP).

produced by various marine and estuarine organisms. The estimated annual rate of total production by marine organisms is ca. 109 tons (for comparison: the estimated weight of the Cheops pyramid is ca. 7.5 × 109 tons).2 DMSP is made by phytoplankton, macroalgae and a few angiosperms, and the knowledge about the diverse biosynthetic pathways via which these enormous amounts of DMSP are made is presented in

Patrick Rabe

Patrick Rabe studied chemistry at the University of Braunschweig. He received the FritzWagner Award of the Helmholtz Centre for Infection Research for his master thesis in 2013. In 2012 he started his PhD with Prof. Jeroen Dickschat working on mechanistic studies of bacterial terpene cyclases founded with a scholarship by the Beilstein Institute for the Advancement of Chemical Sciences.

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the first section of this review. One of the most important functions of DMSP in marine and estuarine ecosystems is its osmoregulation in plants and phytoplankton. Early studies with the coccolithophore Hymenomonas carterae showed that depending on the environmental salt concentration the intracellular DMSP can reach levels of up to 0.3 mol L−1 to counteract the external osmotic pressure.3 Upon lysis by grazing of microzooplankton or due to viral infection the intracellular DMSP can be released.4 These processes are responsible for the secretion of large quantities of DMSP into the environment. Marine bacteria that are to our current knowledge not able to produce DMSP can make use of this dissolved DMSP and benefit from its osmoprotective properties after uptake via their transport systems, discussed in detail in the second section of this review. In marine species such as the cyanobacterium Trichodesnium sp. MD 14–50 this may be of ecological relevance,5 but osmoprotection by DMSP works also in saltstressed cultures of Klebsiella pneumoniae or Escherichia coli that do not get in contact with DMSP in their natural habitats.6,7 As is also discussed in this review, recent work has demonstrated that DMSP transporters can even channel various synthetic DMSP derivatives. Not very surprisingly, if marine microorganisms encounter the compound in their natural habitat and ingest DMSP, they incorporate its sulfur into amino acids and live on it,8 which underlines the role of DMSP as one of the most important sulfur and carbon sources for many marine bacteria.9,10 More astonishingly, some bacteria seem to be completely adapted to DMSP. It represents the only ecologically relevant sulfur source these bacteria are able to use, while genetic information for prominent pathways such as sulfate reduction were lost.11 As will be discussed in this review the ability of marine bacteria to degrade DMSP as well as the various underlying pathways are well investigated, although there are hints that there is more to be discovered. Finally, the accumulated knowledge

Christian A. Citron was born in 1985 in Braunschweig, Germany. He graduated from the Technical University of Braunschweig in 2010 with a master’s thesis about stereochemical aspects of terpene biosynthesis and volatile metabolites from streptomycetes with Prof. Jeroen Dickschat. Afterwards, he continued his work in the Dickschat group for a PhD thesis in the same field which he finished in 2013. His Christian A. Citron current research as post-doctoral fellow in the Dickschat group focuses on the elucidation of biosynthetic pathways to terpenes in various bacteria and fungi using feeding experiments with isotopically labelled precursor molecules.


Review

about the degradation of synthetic DMSP derivatives will be presented, revealing some substrate flexibility of DMSP degrading enzymes. A few other excellent reviews on the physiology10,12 and ecology13 of DMSP have recently been published.

Biosynthesis of DMSP Various marine and estuarine organisms from nearly all kingdoms of life have assimilated the required genetic information or evolved the biosynthetic capacity to make DMSP. Its production is long known from costal plants,14 green algae,15 red algae,1 diatoms,16 coccolithophores,16 and dinoflagellates,17 and most recently it was discovered that even corals can produce DMSP.18 This chapter summarises the knowledge about the biosynthesis in plants and the various types of “algae”, a poorly defined term under which green and red algae, coccolithophores, diatoms and dinoflagellates are summarised, and corals. For clarity, the various types of “algae” will be briefly introduced here. Green algae belong together with the higher plants (embryophytes) to the viridiplantae ( plantae sensu stricto), while red algae (rhodophyta) are archaeplastida ( plantae sensu lato), a taxon that contains both viridiplantae and rhodophyta. As we will see, the biosynthesis of DMSP in plants and the closely related green algae is different, while the taxonomically more distinct green and red algae use the same pathway. Other unicellular eukaryotes that are often referred to as “algae” include coccolithophores, diatoms, and dinoflagellates. These organisms belong to the chromalveolata, that are taxonomically distinct from archaeplastida. Interestingly, there are hints that some chromalveolata (coccolithophores and diatoms) use the same pathway towards DMSP as green and red algae, while dinoflagellates have evolved a third unique pathway. The three well described biosynthetic pathways to DMSP in plants, green algae and dinoflagellates must have evolved independently. It is, however, unknown whether some taxonomically distinct organisms such as green algae and diatoms use the same pathway due to a horizontal gene transfer event or due to its independent evolution. Most of our knowledge about DMSP biosynthesis in the different types of organisms is based on detailed feeding studies with isotopically labelled biosynthetic precursors and the identification of pathway intermediates in culture extracts, while the acquired information about the genes and enzymes for DMSP biosynthesis is scarce. Biosynthesis of DMSP in plants The biosynthetic pathway to DMSP has been investigated in the plants Wollastonia biflora (Compositae) and Spartina alterniflora (Gramineae), revealing that DMSP biosynthesis in plants starts from methionine. The intermediates of the plant pathway are not only different to those of the pathway in algae (vide infra), but interestingly, also the central steps of the pathways in the two investigated plant species are different, while the first and the last steps are the same (Scheme 2). In both

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Review

Scheme 2 Biosynthesis of DMSP in the plants Wollastonia biflora and Spartina alterniflora.

plant species the pathway is initiated by methylation of L-methionine to S-methyl-L-methionine (1).14,19 In W. biflora this is followed by a pyridoxal 5′-phosphate (PLP) dependent transamination-decarboxylation sequence that directly yields 3-(dimethylsulfonio)propionaldehyde (4).20 The likely intermediate, 4-(dimethylsulfonio)-2-oxobutyrate (3), was shown to be instable and has therefore never been observed.20 In contrast, in S. alterniflora 1 is first converted into 3-(dimethylsulfonio)propylamine (2) by decarboxylation and then by oxidative deamination to 4.19,21 In both plants a final oxidation step yields DMSP.19,22 The first intermediate of the plant pathway was identified by Hanson and coworkers in 1994 in pulse-chase experiments with leaf discs of W. biflora and [U-14C]methionine as a probe. In these experiments the radioactively labelled amino acid was injected in one portion to leaf discs of the plant ( pulse), and, after a lag time, its unlabelled counterpart was amended (chase). During the course of the experiment the content of labelling in 1 first strongly increased and then slowly decreased, which is the typical behaviour of a pathway intermediate. In contrast, no such observation was made for 3-(methylthio)propionic acid that was considered as an alternative intermediate.15 Furthermore, [13CH3,C2H3]-1 was incorporated as one intact unit into DMSP, indicating that 1 is a direct precursor of DMSP, and not unspecific distribution of labelling from [13CH3,C2H3]-1 via the C1 pool is responsible for

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its occurrence in DMSP.15 The responsible enzyme for the formation of 1 from L-methionine, S-adenosyl-L-methionine:Lmethionine S-methyltransferase, was purified from leaves of W. biflora and shown to be a homotetramer composed of 115 kDa subunits. The enzyme was able to methylate L-selenomethionine and L-ethionine, but not L-cysteine or S-methyl-Lcysteine.23 The last pathway intermediate in W. biflora, 4, was established in a pulse-chase feeding experiment with [35S]-1, and by feeding of [methyl-14C]-4 that was incorporated into DMSP with high rates.22 The first steps of the pathway in S. alterniflora were also investigated by feeding of radioactively labelled precursors. Labelling from [35S]methionine showed up in 1, demonstrating that the first step towards DMSP is the same in Compositae and Gramineae.19 Feeding of [35S]-1 and [35S]-2 to S. alterniflora resulted in the occurrence of labelling in the small pool of 4 that rapidly turned over to DMSP, showing that the last step of the DMSP pathway is also the same as in W. biflora.19 The central steps of the pathway in W. biflora were investigated by feeding of [methyl-2H3,15N]-1. The experiment revealed that the amino group of 1 is lost by transamination, and not by deamination, during DMSP assembly, based on the following arguments. While the labelled methyl group appeared in DMSP, the nitrogen label occurred mainly in glutamic acid, but not in the amide nitrogen of glutamine. During transamination the amino group is transferred via PLP to 2-oxoglutarate, yielding glutamic acid, while a deamination would liberate ammonia, which would re-enter the amino acid pool via glutamine.20 This pointed to a pathway via instable 3. The central steps from 1 to 4 of the DMSP pathway in S. alterniflora deviate from the respective steps in W. biflora. In S. alterniflora 2 was shown to be an intermediate towards DMSP by pulse-chase experiments using [35S]-1 together with feeding of [35S]-2. These experiments demonstrated that 1 first undergoes decarboxylation to 2.19 The occurrence of labelling from [35S]-1 or [35S]-2 in 4 suggested this aldehyde is the latest intermediate towards DMSP.19 In a subsequent study an enzyme activity was observed in leaf extracts from S. alternifolia that specifically converted the L-enantiomer, but not the 14 D-enantiomer of [U- C]-1 in a PLP dependent reaction into equimolar amounts of [U-14C]-2 and 14CO2.21 A radioassay with [35S]-2 and leaf extracts revealed O2-dependent amine oxidase activity for its conversion via 4 into DMSP, whereas a PLP dependent transaminase activity as observed in W. biflora was clearly ruled out.21 Using total enzyme extracts from chloroplasts and cytosol fractions in incubation experiments with pathway intermediates showed that in W. biflora the methylation step occurs in the cytosol of the plant cells, while the other steps of the biosynthetic pathway take place in the chloroplasts.24 Biosynthesis of DMSP in green and red algae First experiments on the biosynthesis of DMSP in green algae were performed by Greene in 1962.15 These experiments by feeding of radioactively labelled methionine isotopomers to Ulva lactuca showed that the sulfur atom and the whole carbon


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backbone of DMSP are derived from this amino acid.15 The formation of DMSP from methionine in the red alga Chondria coerulescens was shown later by similar experiments.25 A feeding study with both enantiomers of [methyl-14C]methionine demonstrated that the L-amino acid is efficiently incorporated, while minor incorporation of the D-amino acid was attributed to enzymatic epimerisation of D-methionine.26 Unfortunately, the nature of the biosynthetic intermediates remained elusive from these studies, but involvement of deamination, decarboxylation, oxidation and methylation was already suggested by Greene.15 In the last decade of the twentieth century, Hanson and coworkers have addressed the algal DMSP pathway to study its intermediates in detail by feeding of [35S]methionine to the green alga Ulva intestinalis ( previously classified as Enteromorpha intestinalis) and analysis of culture extracts for the presence of 35S-enriched compounds by mass spectrometry. Based on the identified metabolites and their labelling kinetics a pathway starting from L-methionine was suggested (Scheme 3).27 First, L-methionine undergoes a transamination to yield 4-(methylthio)-2-oxobutanoic acid (5) which is subsequently reduced to 2-hydroxy-4-(methylthio)butanoic acid (6). Its S-methylation gives 4-(dimethylsulfonio)-2-hydroxybutanoate (7) which is transformed into DMSP by oxidative decarboxylation. Alternative pathway intermediates such as S-methyl-L-methionine and 3-dimethylsulfoniopropionaldehyde that are relevant to the pathway in higher plants were ruled out, because no significant incorporation of labelling from S-methyl-[35S]-L-methionine or no labelling kinetics as expected for a pathway intermediate was observed.27 The transamination mechanism for the formation of 5 was further supported by feeding of [15N]methionine, resulting in the uptake of labelling in glutamic acid, but not in the amide nitrogen of glutamine.27 In vitro experiments with cell free extracts demonstrated furthermore, that the transamination of L-methionine to 5 is 2-oxoglutarate-dependent, while the reduction to 6 requires NAD(P)H.28 Incubation experiments with both enantiomers of 6 and 7 and cell free extracts of U. intestinalis showed a preferred turnover of the (R)-enantio-

Scheme 3

Biosynthetic pathway to DMSP in green and red algae.


Review

mers, thus establishing the absolute configurations of these pathway intermediates.28 S-Adenosyl-L-methionine (SAM) was identified as the methyl group donor for the methylation to 7.28 The enzyme for this step, S-adenosyl-L-methionine:2hydroxy-4-(methylthio)butanoic acid S-methyltransferase, has a decreased activity under sulfur limiting conditions in Ulva pertusa.29 Finally, the oxidative decarboxylation from 7 to DMSP was shown to be O2-dependent by growing U. intestinalis in an 18O2 atmosphere, resulting in significant uptake of 18O labelling in DMSP.27 Biosynthesis of DMSP in diatoms and coccolithophores Hanson showed in feeding experiments with the coccolithophore Emiliania huxleyi and the diatom Melosira nummuloides that the radioactive labelling from [35S]methionine appeared in 7, the intermediate of the pathway in green algae.27 Based on this finding it was hypothesised that coccolithophores and diatoms may use the same pathway as green algae. Subsequent work was performed with the sea-ice diatom Fragilariopsis cylindrus.30 This organism was grown under high salinity conditions in order to upregulate the expression of genes for DMSP biosynthesis and to find the respective enzymes in total enzyme extracts by two-dimensional protein gel electrophoresis. For comparison, a total enzyme extract obtained at moderate salinity levels was used. Enzymes that were overexpressed under high salinity conditions were selected for MALDI-TOF analysis, and those unknown enzymes that showed high homology to amino transferases, NADPH-dependent reductases, methyl transferases and O2-dependent decarboxylases were suggested as candidate enzymes for the four steps of the DMSP biosynthetic pathway as it is known from green algae (Scheme 3). The same study also provided information about the encoding genes for these enzymes.30 Further characterisation of the algal genes and enzymes putatively involved in the biosynthesis of DMSP in diatoms has not been reported yet. Biosynthesis of DMSP in dinoflagellates A third pathway towards DMSP is known from dinoflagellates. Uchida and co-workers investigated the dinoflagellate Crypthecodinium cohnii by feeding of various 14C- and 35S-labelled isotopomers of methionine.17 The labelling from all carbons and the sulfur atom of methionine, apart from the 14C-labelling at the carboxylic acid group, was incorporated into DMSP. The addition of the plant pathway intermediate 1 in the feeding experiments with radioactively labelled methionine did not suppress the uptake of labelling into DMSP, indicating that 1 is no intermediate in dinoflagellates and the pathway must be different to the pathway in plants (Scheme 2).17 In subsequent work a L-methionine decarboxylase was purified from total enzyme extracts of C. cohnii.31 The enzyme catalyses the first step of the DMSP pathway in dinoflagellates (Scheme 4), i.e. conversion of L-methionine to 3-(methylthio)propylamine (8). The C. cohnii L-methionine decarboxylase is a homodimer of 100 kDa subunits and is PLP dependent. Incubation experiments with [1-14C]methionine resulted in the release of 14 CO2.32

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Scheme 4


Biosynthesis of DMSP in dinoflagellates.

Biosynthesis of DMSP in corals DMSP biosynthesis was recently reported from the two corals Acropora millepora and Acropora tenius.18 The occurrence of DMSP in corals was previously attributed to a production by the symbiont Symbiodinium, but investigations of algal-free juvenile corals clearly demonstrated a heat stress dependent DMSP production by the marine invertebrates. Two orthologues of Fragilariopsis cylindrus genes30 for DMSP biosynthesis were found in the coral genomes, coding for a NAD(P)H dependent reductase and a SAM dependent methyl transferase. Based on this finding it was proposed that the biosynthesis of DMSP in corals may operate as in green algae, diatoms and coccolithophores (Scheme 2).18

DMSP transport As described in the previous chapter, DMSP can be generated by a variety of marine and estuarine organisms including plants, green and red algae, coccolithophores, diatoms, dinoflagellates, and animals. Thus, DMSP is present in all marine ecosystems and can be used for various purposes not only by the producing organisms, but also by other species that live in the same habitat as the DMSP producers. DMSP is an efficient osmoprotectant that reduces or increases the intracellular turgor by accumulation or efflux of this compatible solute to maintain the physical integrity of the cell.33 Possibly due to the different bioavailabilities of sulfur and nitrogen in marine and terrestrial environments DMSP is the preferred compatible solute for marine organisms, while terrestrial species use nitrogen compounds such as glycine betaine (10), choline (11), carnitine (12), or ectoin (13) instead (Scheme 5).34 This

Scheme 5 Structures of naturally occurring nitrogen-containing compatible solutes.

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chapter will summarise the accumulated knowledge about DMSP transport, i.e. the variety of transport proteins, their mechanisms for DMSP efflux and uptake, and their distribution in living organisms. DMSP uptake by phytoplankton has been demonstrated using doubly labelled [13C22H6]DMSP35 and is known from various bacteria including Roseobacter and SAR11 clade bacteria,36,37 cyanobacteria such as Synechococcus and Prochlorococcus,38,39 bacilli (Bacillus subtilis, Listeria monocytogenes),40,41 enterobacteria (Escherichia coli),7 rhizobia (Ensifer meliloti),33 and pseudomonads (Pseudomonas doudoroffii).42 Bacteria from the SAR11 clade even depend on the uptake of reduced sulfur compounds such as DMSP, because they lack the assimilatory sulfate reduction genes.11 Diatoms,39,43 dinoflagellates,43 and green algae29 also actively sequester DMSP from their environment. As shown by competition experiments with radioactively labelled 10 and DMSP marine bacteria can assimilate DMSP via transporters for nitrogen-containing compatible solutes such as 10.44 Two transport systems are mainly responsible for the active uptake of 10 and related nitrogen compounds, the ABC (ATP binding casette) transporters45 and the BCCT (betaine choline carnitine transport) proteins.46 The enzymologies and mechanisms of action of these two transport systems are fundamentally different. All prokaryotic ABC transporters have three elements in common, i.e. a transmembrane protein (TMP), a nucleotide binding protein (NBP), and a substrate binding protein (SBP, Fig. 1).47 Two of these components form a heterotetrameric structure comprised of two TMPs and two NBPs.48,49 The SBP for capture and delivery of the transported molecule interacts with this heterotetramer of the ABC transport system. ATP binding to the NBP and its hydrolysis to ADP and phosphate deliver the energy to cause a reversible conformational switch in the TMPs that channels the substrate through the membrane.48,50 Two of the best studied systems are the multicomponent ABC transporters ProU in Escherichia coli and OpuA in Bacillus subtilis.51,52 Unfortunately, there is no unified nomenclature for the ABC transporters in different organisms. The subunits of the E. coli ABC transporter are termed ProV (NBP), ProW (TMP), and ProX (SBP), while the respective

Fig. 1 Mechanism of ABC transporters. The substrate (S) is bound by a substrate binding protein (SBP, left). Binding of adenosin triphosphate (ATP) to the nucleotide binding protein (NBP) causes a conformational switch in the NBP and the transmembrane protein (TMP) that opens a channel for the substrate (middle). After channeling of the substrate adenosin diphosphate (ADP) and phosphate (Pi) are released and the channel closes again (right).


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subunits of the B. subtilis ABC transporter are called OpuAA, OpuAB, and OpuAC. As will be discussed in detail in the subsequent chapter DMSP can be catabolised by the action of various enzymes, one of which is the DMSP hydrolase DddD that liberates dimethyl sulfide.53 In the α-proteobacteria Burkholderia ambifaria AMMD, Rhizobium sp. NGR234, Hoeflea phototrophica DFL-43 and Rhodobacterales bacterium KLH11 the dddD gene is located in close proximity to genes that code for the constituents of ABC transport systems, which is suggestive of the involvement of these ABC transporters in DMSP uptake.34 Direct evidence was obtained by expression of the ABC transporter genes potABCD from B. ambifaria together with the dddD gene from Rhizobium sp. NGR234 in the E. coli mutant strain MKH13 that is impaired in the uptake of glycine betaine due to disruption of several genes for transport systems.54 In the presence of DMSP the liberation of dimethyl sulfide was detected, which was not the case in the relevant control experiments. This clearly demonstrated that DMSP is taken up by B. ambifaria via the respective ABC transport system.34 In B. subtilis, two ABC transport systems, OpuA and OpuC, are capable of channeling DMSP through the cell membrane, albeit with different efficiency. As revealed by disruption of the genes for all other transport systems, OpuA has only weak affinity towards DMSP, while OpuC is an excellent importer of DMSP, its naturally occurring selenium analog dimethylseleniopropionate (14, DMSeP, Scheme 6), and the synthetic derivatives dimethyltelluriopropionate (15, DMTeP), ethylmethylsulfoniopropionate (16, EMSP), diethylsulfoniopropionate (17, DESP), isopropylmethylsulfoniopropionate (18, IPSP), and tetramethylenesulfoniopropionate (19, TMSP). Only the uptake of methylpropylsulfoniopropionate (20, MPSP) proved to be less efficient.40 Molecular modelling studies based on the crystal structures of the substrate binding proteins OpuAC55,56 and OpuCC57 revealed a higher substrate flexibility of the OpuCC protein. The cationic portion of a short-chain substrate such as 10 is bound to an aromatic cage of four tyrosine residues by cation–π-interactions, while the carboxylate

Review

group is recognised by the active site residues Glu-19 and Thr74. The cationic moiety of substrates with a longer chain such as DMSP is likewise bound to the aromatic cage, whereas the binding mode of the carboxylate function is altered to binding to Glu-19 and Ser-51, similarly to the situation in the OpuCC:12 complex.57 Like ABC transporters the BCCT system is also ubiquitous in all kinds of organisms. While the first discovered BCC transporters from Escherichia coli,58 Bacillus subtilis59 and Corynebacterium glutamicum60 were found to import glycine betaine, choline and carnitine (10–12, Scheme 5) and were therefore named BCCT,61 later studies revealed that several other molecules including DMSP and ectoine (13) are also accepted.46 The nomenclature of BCCTs is also rather confusing, e. g. the proteins from C. glutamicum, E. coli, and B. subtilis are called BetP, CaiT, and OpuD, respectively. The amino acid sequences of BCC transporters are of poor homology, which makes their identification by bioinformatics difficult. The crystal structures of BetP from C. glutamicum62 and CaiT from E. coli63 show that BCC transporters are asymmetric homotrimers with a terminal domain that protrudes into the cytoplasm. As in ABC transporters the primary substrate binding motifs of BetP and CaiT are composed of aromatic residues that are involved in cation– π-interactions.62,63 Mechanistically, these transmembrane proteins act either as symporters62 or antiporters,63 i.e. they act as transporters of two different solutes in the same or opposite directions. The E. coli BCCT system is able to mediate the transport of DMSP.7 Ecologically more important is the finding that a specific BCC transporter for channeling of DMSP is present in the marine bacterium Marinomonas MWYL1 (DddT). In this strain the DMSP transporter is encoded within the dddTBCR operon that is located next to the dddD gene involved in DMSP catabolism (vide infra, ddd = DMSP-dependent dimethyl sulfide). A homolog of this BCCT type enzyme has also been found in Sagittula stellata E37.53 Also in Halomonas the dddT gene is clustered with other ddd genes for DMSP degradation. The function of the DddT protein was confirmed by investigation of a dddT knockout mutant which failed to grow on DMSP. Furthermore, expression of the dddT gene in a betaine uptake defective E. coli mutant enabled DMSP influx.64 Finally, dddT homologs occur in two isolates (Psychrobacter and Pseudomonas) from the gut microbiome of Atlantic Herring (Clupea harengus) where the gene is again clustered with other ddd genes.65

Degradation of DMSP

Scheme 6 Structures of the naturally occurring dimethylseleniopropionate (14) and synthetic DMSP analogs.


DMSP that is either actively exported by marine phytoplankton, or liberated via lysis by viruses or by zooplankton, is mainly catabolised by marine bacteria via two competing pathways, the demethylation pathway and the cleavage pathway.66 The responsible genes encoding the enzymes for DMSP catabolism are very widespread in marine bacteria, but homologs also occur in terrestrial bacteria and in a few cases even in

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fungi. However, the presence of these genes in terrestrial organisms may not be relevant for DMSP catabolism, since DMSP is not present in terrestrial habitats, and may thus have other functions. While the demethylation pathway yields water-soluble methanethiol (MeSH) from DMSP, the cleavage pathway produces the climatically relevant volatile dimethyl sulfide (DMS) via DMSP lysis or hydrolysis. Therefore, it is of large interest how the two pathways are distributed in the environment and how they are differentially regulated in bacteria such as Ruegeria pomeroyi that possess the genes for both pathways.67 Carbon supply, temperature and UV-A dose have been suggested as important external factors that control a “bacterial switch” mechanism between DMSP demethylation and lysis.68 Although it is difficult to estimate, only a minor fraction of the global DMSP may be cleaved to the climatically relevant volatile DMS.69–71 In this chapter the accumulated knowledge about both pathways will be discussed in detail. DMSP demethylation pathway Although the production of MeSH from DMSP by marine bacteria is a long known phenomenon,72,73 the DMSP demethylation pathway was only recently fully elucidated in the laboratories of Moran (Scheme 7). Along this pathway DMSP is first demethylated by the methyl transferase DmdA that transfers a methyl group to tetrahydrofolate (FH4) to yield 3-methylmercaptopropionic acid (21).74 This free acid is subsequently converted by the coenzyme A ligase DmdB into the corresponding coenzyme A ester 22 via activation with ATP. The dehydrogenase DmdC catalyses its FAD-dependent oxidation to methylthioacryloyl-CoA (23), followed by the addition of water by the enoyl-CoA hydratase DmdD to yield 24. This hemithioacetal immediately liberates MeSH to give the coenzyme A thioester of malonyl semialdehyde (25) that upon hydrolysis of the thioester decarboxylates to acetaldehyde (26) and carbon dioxide.75 Recently, it was suggested that methanethiol can be

Scheme 7 DMSP degradation via the demethylation pathway. The mechanism of the first step by DmdA is shown in detail in Scheme 8.

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reduced to methane which accumulates in surface waters based on a microcosm experiment in which methane production was observed when arctic surface water, colonised by different DMSP degrading bacteria, was supplemented with DMSP.76 The dmdA gene was first identified in Ruegeria pomeroyi DSS-3 using a transposon mutagenesis approach.74 Homologs of this gene can be found in many marine bacteria, e. g. from the Roseobacter clade (Table 1 of ESI†), the SAR11 clade, and γ-proteobacteria,75 and even in coral-associated phages that may be involved in spreading of the gene in marine ecosystems.77 The purified DmdA enzymes from Pelagibacter ubique HTCC1062 and Ruegeria pomeroyi DSS-3 showed specific demethylase activity with DMSP.78 All other three genes of the demethylation pathway, dmdB, dmdC and dmdD, were identified in a subsequent study in R. pomeroyi by bioactivity guided purification of enzymes from total enzyme extracts, MALDITOF analysis of the purified proteins, and heterologous expression and knockout of the coding genes.79 The DmdB enzyme was identified as a major regulatory point in DMSP demethylation. A phylogenetic analysis and biochemical characterisation of DmdB from various organisms revealed that the DmdB enzymes separate into two functionally equivalent clades.80 The genes dmdB and dmdC are not only present in marine bacteria, but can also be found in many terrestrial bacteria such as β- and γ-proteobacteria. Furthermore, dmdB and dmdC homologs occur in bacteria that do not possess a dmdA gene, and it was shown for a few representatives including the typical soil bacterium Myxococcus xanthus that growth on 21 resulted in production of MeSH. Moran suggested79 that in these organisms 21 may arise from a side reaction of the salvage pathway that converts 5′-methylthioadenosin into methionine.81 In contrast, the dmdD gene is not very widespread, but it was shown that cell extracts of the dmdD-negative bacterium Ruegeria lacuscaerulensis possessed DmdD activity, suggesting that a non-orthologous isofunctional enzyme may be encoded in this strain.79 As is evident from the summarised data in Table 1 of ESI,† among the Roseobacter clade bacteria all four dmd genes of the demethylation pathway are only present in R. pomeroyi DSS-3, Leisingera nanhaiensis DSM 24252, Phaeobacter arcticus DSM 23566, Phaeobacter inhibens DSM 17395, and the two Sulfitobacter mediterraneus strains 1FIGIMAR09 and KCTC 32188. The crystal structures of DmdA from Pelagibacter ubique apo and in complex with the substrate DMSP or the cofactor FH4 were reported in 2012.82 The cofactor is bound to the active site by hydrogen bonds to the highly conserved S122, E204 and Y206 residues (Scheme 8) and by π-stacking to Y95, while DMSP shows specific interactions with R11, Y32, W197 and S246 (not shown). A superimposition of the obtained crystallographic data of the DmdA:DMSP and DmdA:FH4 complexes provided insight into a concerted mechanism of the demethylation reaction with a methyl transfer from DMSP to FH4 that proceeds with a concomittant proton transfer via water to an unidentified general base. This mechanism resembles the proposed mechanism of SAM-dependent N-methyltransferases


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Overlay of the structural data from the wildtype enzyme and of the E121A mutant in complex with 23 gave insights into the enzyme mechanism (Scheme 9). The substrate methylthioacryloyl-CoA (23) binds with its carbonyl oxygen to an oxyanion hole. A water molecule is activated by E121 and attacks to the Michael acceptor of 23, while protonation at the substrate’s α-carbon is mediated by E141. Elimination of MeSH from the resulting hemithioacetal 24 yields the coenzyme A thioester of malonyl semialdehyde (25). Subsequently, 25 may be attacked by the carboxylate of E141 to give rise to an anhydride intermediate that spontaneously hydrolyses to malonic semialdehyde ( pathway a). Alternatively, E141 may activate water for direct hydrolysis of the CoA thioester 25 to yield malonic semialdehyde ( pathway b). Its spontaneous decarboxylation results in the final product 26. DMSP cleavage pathway

Scheme 8 Mechanism of methyl transfer from DMSP to the cofactor FH4 by DmdA.

such as human phenylethanolamine N-methyltransferase or the protein arginine methyltransferase PRMT3.83,84 Crystal structures of DmdD from Ruegeria pomeroyi DSS-3 were also recently reported.85 The enzyme crystallises as a hexamer (dimer of trimers) revealing a crystallographic C3 axis.


As described in the previous chapter, DMSP can be degraded by many marine bacteria via the demethylation pathway to yield MeSH. Alternatively, marine bacteria can also degrade DMSP to the poorly water soluble volatile DMS that is released in large quantities from the oceans into the atmosphere. Several enzymes for the degradation of DMSP to DMS have been described, including the enzymes DddL,86 DddP,87,88 DddQ,89 DddW,90 and DddY91 that all catalyse DMSP lysis to DMS and acrylate (28), while DddD is the only characterised enzyme that cleaves DMSP by hydrolysis to DMS and 3-hydroxypropionate (27, Scheme 10).53 All these enzymes of the cleavage pathway have been discovered by Johnston and coworkers. Interestingly, they differ in their size and amino acid sequence and belong to distinct polypeptide families.10 The first discovered enzyme for DMSP cleavage was the DMSP hydrolase DddD from Marinomonas sp. MWYL1.53 Heterologous expression of the dddD gene in E. coli was shown to depend on the regulatory gene dddR. This gene is part of the dddTBCR operon that is located directly next to the dddD gene and encodes a LysR family transcriptional regulator. Based on its amino acid sequence the DddD enzyme was identified as a type III acyl coenzyme A transferase. The closest characterised homolog was the γ-butyrobetainyl-CoA:carnitine CoA-transferase CaiB from E. coli (26% identity) that transfers coenzyme A from γ-butyrobetainyl-CoA to 12, but fails to catalyse the direct thioesterification of 12 with coenzyme A.92 The homology between DddD and CaiB suggested a mechanism for the cleavage of DMSP by DddD that proceeds via conversion of DMSP into its coenzyme A thioester 29, followed by hydrolysis to 27 (Scheme 10).53 However, the suggested intermediate 29 has never been observed directly, presumably because of its rapid turnover to 27 by DddD. Furthermore, activity of DddD has not yet been established in vitro,10 possibly because it requires similar to CaiB an unidentified coenzyme A thioester as cosubstrate. The DddD homologs from the terrestrial bacteria Sinorhizobium fredii NGR234 ( previously assigned as Rhizobium sp.) and Burkholderia ambifaria (B. cepacia) AMMD were also shown to cleave DMSP by heterologous expression in E. coli.53 Interestingly, the dddD gene

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Scheme 9


Proposed enzyme mechanisms of DmdD.

Scheme 10 Characterised enzymes for DMSP degradation via the cleavage pathway and their products.

also occurs in the two isolates Pseudomonas and Psychrobacter from gut microbiome of the Atlantic Herring (Clupea harengus) that eats copepods that in turn feed on DMSP-producing phytoplankton.65 Finally, the gene was also identified in Oceanimonas doudoroffi DSM 7028.93 Although no dddD homolog is present in Sulfitobacter sp. EE-36 and Roseovarius nubinhibens ISM, both organisms release DMS from DMSP,53,94 suggesting that besides DddD other enzymes for DMSP cleavage may be active in these species.53 This prompted a subsequent study in which the dddL gene was uncovered in Sulfitobacter sp. EE-36.86 Its heterologous expression in E. coli demonstrated that the gene product is a DMSP lyase that converts DMSP into DMS and 28. The DddL enzyme shows no significant homology with any previously characterised enzyme or domain of known function. The following observations suggested that DddL is a transmembrane protein that acts on periplasmic DMSP: First, most of the product 28 was found in the growth medium and not intracellularly, second, bacteria containing the dddL gene show no or only poor growth on DMSP as a single carbon source, and third, the central part of the DddL sequence has high homology to transmembrane domain forming polypeptides.86 A BLAST search revealed that the dddL gene is present in many bacteria of the Rhodobacteraceae family, but not in the α-proteobacterium R. nubinhibens ISM.86 Consequently, Johnston and coworkers investigated this organism in detail and identified the third gene for DMSP cleavage, dddP. Its product DddP belongs to the large family of M24 metallopeptidases and catalyses – like the completely unrelated DddL – the clea-

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vage of DMSP to DMS and 28 in vivo.87 The recombinant dddP gene from R. nubinhibens ISM was also expressed in E. coli and purified, revealing that the enzyme is a homodimer. Incubation experiments with [1-13C]DMSP and [1-14C]DMSP yielded isotopically labelled 28 and DMS as single products, thus giving direct proof for its activity as a DMSP lyase. Although many, but not all metallopeptidases depend on metal ion cofactors such as Co2+,95 the addition of chelators to in vitro reactions of DMSP with DddP did not affect enzyme activity, which is consistent with the finding that purified DddP did not contain any metals.88 Disruption of the dddP gene in R. nubinhibens ISM resulted in a significantly decreased, but not completely abolished DMSP lyase activity, suggesting that there were still more DMSP lyases to discover.74 Two distantly related copies of dddP were found in Oceanimonas doudoroffi DSM 7028.93 Two copies of an additional gene for DMSP breakdown termed dddQ1 and dddQ2 were subsequently discovered in R. nubinhibens ISM.89 These genes are located directly next to each other and are part of an operon spanning ten genes. The sequences of both DddQ enzymes show weak homology to DddL only in the C-terminal halves that are predicted to represent cupin folds, suggesting that DddQ represents a new type of DMSP lyases. The functions of both proteins DddQ1 and DddQ2 were elucidated by heterologous expression of the coding genes in E. coli and conversion of [1-13C]DMSP and [1-14C]DMSP with a cell lysate into DMS and 28 with a corresponding isotopic labelling. In addition, the dddQ gene from R. pomeroyi DSS-3 and three synthesised dddQ genes from a metagenome database96 were shown to encode fully functional DMSP lyases.89 The DMSP lyase DddW was identified from R. pomeroyi DSS-3.90 Expression of the corresponding dddW gene was strongly enhanced during growth in a medium containing DMSP,90,97 and the product of this gene contained a predicted cupin motif as present in the DMSP lyases DddL and DddQ. This prompted further investigations revealing that DddW converts [1-14C]DMSP into DMS and [1-14C]-28. Transcription of the dddW gene in R. pomeroyi DSS-3 is regulated by a DMSPinducible promoter and a LysR type regulator encoded directly upstream of dddW that also responds to DMSP.90 Finally,


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another DMSP lyase named DddY was identified from Alcaligenes faecalis M3A.91 Knockout of the dddY gene abolished DMS production from DMSP, but the mutant grew on DMSP as sole carbon source, suggesting that a second DMSP catabolic pathway is active in this strain. Heterologous expression in E. coli enabled the transformation of [1-13C]DMSP into DMS and [1-13C]-28. The protein is located in the periplasm and is by sequence not related to any of the other characterised DMSP lyases. The fate of the initial products of the DMSP hydrolase DddD, 3-hydroxypropionate (27), and of all different types of DMSP lyases, acrylate (28), was further studied by Johnston and coworkers. A Halomonas HTNK1 was isolated from the macroalga Ulva lactuca by selection on medium with acrylate as sole carbon source. From this strain a gene cluster spanning genes for the BCCT type DMSP transporter DddT, the DMSP hydrolase DddD, and four additional genes termed akuN, akuK, dddA, and dddC was deciphered.64 The gene products AkuN and AkuK showed homology to the E. coli crotonobetainyl-CoA:carnitine CoA transferase CaiB and carnitine hydratase CaiD, respectively, that cooperate in the degradation of carnitine.92 In Halomonas the AkuNK duo acts simultaneously in the conversion of 28 to 27 (Scheme 11). The metabolite 27 is the central intermediate of DMSP catabolism, since it is either

Scheme 11

DMSP degradation via the cleavage pathway.


Review

formed directly by DMSP hydrolysis or via 28 by DMSP lysis. Downstream steps were suggested to proceed via malonate semialdehyde (30) to acetyl-CoA (31) and carbon dioxide under catalysis of the alcohol dehydrogenase DddA and the aldehyde dehydrogenase DddC.64 Recently, the crystal structures of two DMSP cleaving enzymes, DddQ from Ruegeria lacuscaerulensis ITI_1157 and DddP from Roseobacter denitrificans OCh 114, have been solved.98,99 The structural data of DddQ reveal a β-barrel fold that is typical for cupin enzymes. The active site contains a Zn2+ and the highly conserved residues Y120, H123, H125, E129, Y131 and H163. Mutations of all six residues showed their importance for catalytic activity. Four of these residues (H125, E129, Y131 and H163) are involved in Zn2+ binding (Scheme 12). The DddQ:Mes complex with the substrate surrogate 2-(N-morpholino)ethanesulfonic acid (Mes) resulted in a release of Y131 from the metal centre and its conformational rearrangement. A superimposition of the DddQ:Mes structure with the structure of Y131A:DMSP demonstrated that the phenolate oxygen of Y131 is precisely located to abstract a proton from the α-carbon of DMSP, thereby initiating its lysis.98 The crystal structure of DddP revealed that this enzyme is a homodimer with each subunit composed of two domains. Within the active centre a binuclear iron cluster is coordinated by monodentate (D297, E406, H371, D307) and bidentate (D295, E421) ligands. The function of either D337 or Y366 to act as a base for deprotonation at the α-carbon of DMSP was suggested,99 but more experiments are required for deeper insights into the enzyme’s mechanism. The distribution of genes for DMSP cleavage in microorganisms has been summarised previously.10 Since the modern sequencing techniques have delivered a wealth of new information during the past few years, an update will be given here (Table 2 of ESI†). A BLAST search revealed that genes of the DMSP cleavage pathway are most abundant in marine α-proteobacteria, followed by marine γ-proteobacteria. The DMSP hydrolase DddD occurs in many strains of these groups, but also occasionally in terrestrial ( plant associated) α-proteobacteria ( particularly in Rhizobium or closely related genera) and β-proteobacteria (Burkholderia). In contrast, DddL is almost restricted to marine α-proteobacteria, with the exemption of two examples from marine γ-proteobacteria and three from marine δ-proteobacteria. The metallopeptidase DddP is seen both in marine and terrestrial microorganisms. In fact, all bacteria in which it is found belong to the α- and γ-proteobacteria. While DddP is present in various genera of their marine representatives, it is only known from specific genera of terrestrial species, i.e. Mesorhizobium (α-proteobacteria) and Pseudomonas (γ-proteobacteria). Interestingly, DddP is the only DMSP cleavage enzyme also observed in plant associated ascomycete fungi, pointing to an interkingdom horizontal gene transfer.87 Similar to DddL the cupin protein DddQ is restricted to marine α-proteobacteria. Together with the weak sequence homology between DddL and the C-terminal half of DddQ this may point to a common ancestor. The comparably rare DMSP lyase DddW occurs almost exclusively in α-proteo-

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Scheme 12


Mechanism of DMSP lyase DddQ based on structural data.98

bacteria of marine and terrestrial origin, and in one exceptional case in the actinobacterium Geodermatophilus obscurus. Finally, DddY is the only DMSP lyase for which no example in α-proteobacteria is observed. Instead, this enzyme is frequently found in β- and γ-proteobacteria, but in a few cases also in marine δ- and ε-proteobacteria and in one flavobacterium. Many bacteria, particularly α-proteobacteria, contain several types of DMSP cleaving enzymes. The record is held by Ruegeria pomeroyi DSS-3 that contains four enzymes (DddD, DddQ, DddP and DddW). Recently, ddd genes have also been detected in phage genomes pointing to a role of viruses as shuttles in their horizontal transfer.77

Bacterial conversion of DMSP analogs Interestingly, the salt marsh cord grass Spartina alterniflora produces DMSeP (14) in the presence of selenate salts, most likely via the same biosynthetic pathway as for DMSP (Scheme 2).100 Together with the activity of the associated bacterial community to degrade 14 to volatiles this could account for a detoxification mechanisms of selenate contaminated sites. Early investigations also demonstrated that partially purified fractions of cell extracts from Alcaligenes faecalis M3A and Oceanimonas doudoroffi ( previously termed Pseudomonas doudoroffi) exhibiting “DMSP lyase activity” were able to cleave 14 to yield dimethyl selenide (DMSe). Recent feeding experiments were performed with a series of DMSP derivatives in which sulfur was replaced by selenium or tellurium or with alternative sulfonium alkyl substituents (Scheme 13).101–103 These compounds were easily prepared by the HCl catalysed Michael addition of dialkyl chalcogenides to acrylic acid (28). Further feeding studies were performed with isotopically labelled DMSP.101,103 Deuterated [2H6]DMSP was likewise prepared by the addition of [2H6]DMS to 28, while the synthesis of [34S]DMSP required a special approach from 34 S8.103 A particular problem of the synthetic route was that volatile intermediates had to be avoided to prevent losses of the expensive labelled material. Therefore, 34S8 was converted

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Scheme 13

Synthesis of DMSP derivatives.

into K34SCN by treatment with a hot aqueous KCN solution and used in a nucleophilic substitution with tert-butyl 3-bromopropionate (32). The resulting tert-butyl 3-(thiocyanato)propionate (33) was reduced to the corresponding thiol 34 with samarium diiodide. The thiol was sequentially methyl-


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ated with methyl iodide and Meerwein’s salt to the dimethyl sulfonium compound. A final acid catalysed cleavage of the tert-butyl ester yielded the target material [34S]DMSP. The synthetic DMSP derivatives were fed to marine bacteria from the Roseobacter clade and the volatile degradation products were trapped by use of a closed-loop stripping apparatus (CLSA)104 followed by GC/MS analysis.105 Feeding of isotopically labelled [2H6]DMSP to Phaeobacter inhibens ( previously named P. gallaeciensis), Dinoroseobacter shibae and Oceanibulbus indolifex resulted in the uptake of labelling into several MeSH-derived volatiles, thus demonstrating a highly active demethylation pathway in these species. Interestingly, DMSeP was also efficiently demethylated, as indicated by the formation of various MeSeH derived volatiles, whereas no volatiles were detected from DMTeP. Feeding of EMSP resulted in the production mainly of EtSH-derived volatiles, indicating that demethylation is strongly preferred over deethylation in the initial dealkylation step presumably catalysed by DmdA. Accordingly, the production of EtSH-derived volatiles from DESP could be observed, but was poor. Instead, large quantities of diethyl sulfide were found, pointing to an efficient lysis of this compound.101 The formation of [2H6]DMS from [2H6]DMSP could not be observed in this study, because this highly volatile compound was covered by the solvent signal in the GC/MS analysis of CLSA extracts. A subsequent study using the solvent free solid phase microextraction (SPME) revealed the production of minor amounts of [2H6]DMS (10%) and large quantities of [2H6]dimethyl disulfide, the oxidation product of [2H3]MeSH, from [2H6]DMSP in Ruegeria pomeroyi. Furthermore, feeding of [2H6]DMSP to dmdA knockout mutants of R. pomeroyi and P. inhibens reduced, but did not abolish the production of [2H3]MeSH-derived volatiles, suggesting that a second DMSP demethylation pathway must be active in both species. It was also shown in the same study that SPME is a suitable technique for the detection of dimethyl telluride (DMTe) and MeTeH-derived volatiles.102 Recently, it was shown by feeding of [34S]DMSP to P. inhibens and O. indolifex, that the 34S labelling was mainly incorporated into the methylated sulfur atoms of dimethyl trisulfide, but only marginally into the central sulfur, indicating that double demethylation of DMSP is not an important process. During the course of the same study the three DMSP derivatives MPSP, IMSP and TMSP were also fed to P. inhibens and O. indolifex. The feeding experiments with MPSP and IMSP mainly resulted in the lysis products methyl propyl sulfide and isopropyl methyl sulfide, while only traces of volatiles arising via the demethylation pathway were found, suggesting that MPSP and IMSP are no good substrates for DmdA. However, particularly the experiment with O. indolifex is surprising, because the (incomplete) genome sequence does neither show the presence of dmdBCD genes for the late steps of the demethylation pathway nor any gene for DMSP lysis or hydrolysis. TMSP is a derivative that cannot be transformed via the demethylation pathway, and accordingly, only the lysis product tetrahydrothiophene was observed. Surprisingly, also the sulfur compound 1,2-dithian (38) was detected in feeding experiments with TMSP. Its for-


Review

Scheme 14

Suggested formation of 1,2-dithian from TMSP.103

mation is unclear, but was suggested to proceed via ring opening by nucleophilic attack of hydrogen sulfide to 36 followed by lysis to 1,4-butanedithiol (37) and oxidation (Scheme 14).103

Conclusions We have summarised the accumulated knowledge about the biosynthetic pathways to DMSP, which are different in plants, green and red algae, and dinoflagellates. The DMSP made by marine phytoplankton is taken up by marine bacteria for which either ABC or BCC transporters are responsible. The ABC transporter OpuC from B. subtilis was also shown to accept synthetic DMSP derivatives. After DMSP uptake, marine bacteria, particularly α- and γ-proteobacteria, are the main players of DMSP degradation that can proceed either by demethylation and further degradation to MeSH, or via cleavage to DMS and acrylic acid or 3-hydroxypropionic acid. It is also known that DMSP degrading enzymes show a high substrate flexibility and accept many synthetic DMSP analogs. The concert of the DMSP degrading reactions to MeSH or DMS is of utmost global significance. While MeSH is water soluble, stays in the marine environment and is consumed by marine plankton for de novo biosynthesis of amino acids,8 DMS is apolar and volatile and is thus emitted in large amounts of ca. 3 × 107 tons from the oceans into the atmosphere.106 This process closes the biogeochemical sulfur cycle in which sulfate is transported via rivers to the oceans, while the backflow from the oceans to the continents requires atmospheric sulfur volatiles.107 Atmospheric DMS is oxidised to sulfate that acts in the formation of cloud condensation nuclei which influences the local and possibly even the global albedo with a cooling effect on the planet’s climate.108 Even a negative feedback model was suggested in which this cooling effect due to increased DMS emission leads to a slower biomass production and thus a decreased production of DMSP and its catabolites.108,109 A few other interesting aspects of DMSP and its degradation products not discussed in detail here include the chemotactic response of various microbes and even of complex eukaryotic organisms. DMSP itself is a chemoattractant for marine planktonic microbes,110 including DMSP degrading bacteria such as

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Ruegeria (Silicibacter) sp. TM1040 and Alcaligenes faecalis M3A.111,112 The bacterial pathogen Vibrio coralliilyticus also targets its host by responding to the host’s metabolite DMSP. However, the metabolite is not degraded by V. coralliilyticus suggesting that DMSP serves as an “infochemical for host location”.113 Since Roseobacter clade bacteria are abundant in blooms of DMSP producing phytoplankton, bacterial chemotaxis to DMSP plays an important role in the interplay of phytoplankton and bacteria. The DMSP degradation product DMS is a chemoattractant for copepods that feed on DMSP-producing phytoplankton.114 The grazing zooplankton causes the degradation of DMSP to DMS and acrylic acid that act as feeding deterrents towards the protozoan herbivores, thus representing a chemical defence mechanism of the phytoplankton.115 The resulting increased emission of DMS into the atmosphere upon grazing is in turn also recognised by seabirds that eat the grazing krill.116,117 DMSP is the most abundant organic sulfur metabolite in the marine environment, and all these fascinating examples demonstrate that it is also at least to our opinion the most interesting one. The chemical biology of DMSP, especially aspects of its various ecological functions and unsolved questions about the genes, enzymes and mechanisms of its biosynthesis and degradation, will certainly attract a lot of more inspiring future research.

Acknowledgements This work was funded as a project of the Transregional Collaborative Research Centre “Roseobacter” (TRR51) by the Deutsche Forschungsgemeinschaft (DFG) and by the BeilsteinInstitut zur Förderung der Chemischen Wissenschaften with a PhD scholarship (to P.R.).

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