Influence of Salinity on Dimethyl Sulfide and Methanethiol Formation and its side Effect on Nitrous Oxide Emissions

INFLUENCE OF SALINITY ON DIMETHYL SULFIDE AND METHANETHIOL FORMATION AND ITS SIDE EFFECT ON NITROUS OXIDE EMISSIONS PAULA LILIANA VILA NOVA SALGADO DISSERTAÇÃO DE MESTRADO EM CIÊNCIAS DO MAR – RECURSOS MARINHOS 2013 PAULA LILIANA VILA NOVA SALGADO INFLUENCE OF SALINITY ON DIMETHYL SULFIDE AND METHANETHIOL FORMATION AND ITS SIDE EFFECT ON NITROUS OXIDE EMISSIONS Dissertação de Candidatura ao grau de Mestre em Ciências do Mar – Recursos Marinhos submetida ao Instituto de Ciências Biomédicas de Abel Salazar da Universidade do Porto. Orientador – Doutora Catarina Maria Pinto Mora Pinto de Magalhães Categoria – Investigadora Afiliação – Centro Interdisciplinar de Investigação Marinha e Ambiental À memória do meu avô, o meu amigo, o meu anjo… v Para ler antes (… cada nova possibilidade que a existência possui, incluindo a menos provável, transforma a existência inteira. Milan Kundera), durante (It is not the strongest of the species that survive, nor the most intelligent, but the one most responsive to change. Charles Darwin) … e depois (Escritas de luz investem pela sombra, mais prodigiosas do que meteoros. A alta cidade irreconhecível cresce sobre o campo. Certo da minha vida e da minha morte, olho os ambiciosos e queria entendê-los. O seu dia é ávido como o laço no ar. A sua noite é a trégua da ira no ferro, rápido ao atacar. Falam de humanidade. A minha humanidade está em sentir que somos vozes da mesma penúria. Falam de pátria. A minha pátria é um ganido de guitarra, alguns retratos e uma velha espada, a clara prece do salgueiral nos entardeceres. O tempo está a viver-me. Mais silencioso do que a minha sombra, cruzo o tropel da sua excitada cobiça. Eles são imprescindíveis, únicos, merecedores do amanhã. O meu nome é alguém e qualquer um. Passo com lentidão, como quem vem de tão longe que não espera chegar. Jorge Luis Borges) vi vii Agradecimentos Sem delongas, aqui vai o meu expansivo obrigado: antes de tudo, à minha orientadora Doutora Catarina Magalhães cuja incansável tarefa de coordenação, orientação e incentivo tornou-se intensivamente profícua e contribuiu grandemente para a elaboração desta dissertação. Os desafios propostos ao longo deste projecto representaram mais do que “resultados estatisticamente significativos”… tornaram-se em caminhos simbolicamente percorridos e terminados, acrescentando uma consciência de crescimento e orgulho pessoal e profissional. Pelo seu sobejar de excelência, profissionalismo, competência e, simultaneamente, amizade: um inefável OBRIGADO, Catarina, por apostares em mim. ao Prof. Dr. Adriano Bordalo do laboratório de Hidrobiologia (ICBAS-UP), por disponibilizar-me o laboratório bem como todo o apoio técnico e logístico ao longo da execução deste trabalho. Adicionalmente, aos colegas de trabalho Catarina Teixeira, pela resolução de problemas e esclarecimento de dúvidas, Ana Machado, pela entusiástica introdução à Microbiologia, Catarina Café, pela partilha de livros e revelação duma genuína amizade, Hugo, pela companhia nos “turnos da noite”, Cláudia, memorável companheira de secretária e SBSR, Evita, por acompanhar-me nas saídas mais preteridas, e à D. Lurdes, pelo “bom dia!” sorridente que concediame todos os dias de manhã. ao director do Mestrado em Ciências do Mar – Recursos Marinhos, Prof. Eduardo Rocha, que mostrou-se sempre disponível para auxiliar-me nas tarefas mais burocráticas. O meu sincero obrigado pela pessoa prestável e expedita que se revelou ao longo de todo o mestrado. aos meus novos colegas do CIIMAR, cuja demonstrativa empatia desde o início da minha mudança para estas instalações criou o ambiente motivador para o término desta dissertação. Um agradecimento especial ao Jacinto, que considero um novo amigo e o meu companheiro dos desabafos e “cervejinha” ao final da tarde. às minhas colegas do laboratório EcoBioTec (CIIMAR), Ana Paula, pela modesta liderança e partilha de gargalhadas, Sandra Ramos, por ser a primeira pessoa ideal a receber-me com um sorriso aquando da minha entrevista, Izabela Reis, amiga possuidora de resposta às minhas perguntas, Maria Monteiro, cuja jovialidade e inocência fazem-me acreditar num mundo melhor, e Tatiana, por transmitir-me energia e contribuir com o seu pensamento positivo em todos os dias de trabalho. A todas um obrigado especial pela boa disposição, ânimo e motivação que proporcionaram e impeliram-me a nunca desistir, especialmente quando sentia o tempo a escapar-se-me. aos meus [velhos e novos] amigos que contribuíram para o meu equilíbrio e partilharam momentos, viagens, noitadas de modo a descontraírem-me da pressão sentida ao longo destes meses… À Mi, Andreia e Paulo, pelos incríveis jantares viii seguidos de karaoke e snooker, ao Carlos e Márcio, que fazem e sempre fizeram rirme como ninguém, às “suecas” Margarida e Vanessa, pelos encontros culturais seguidos de lanches fartos, ao Rui, por oferecer-me a música que inspirou e encorajou-me numa fase difícil, à Maria João, por ser o catalisador na procura pelo meu verdadeiro eu através da dança, e finalmente aos meus amigos da Universidade do Algarve, por ainda fazerem parte da minha vida. à minha família, que demonstraram um incomensurável apoio em todas as decisões da minha vida, aos meus primos/”irmãos”, Nelo e Ricardo, por sentirem e demonstrarem orgulho em mim, à minha avó, que revelou uma clareza de espírito e razão nos momentos em que mais precisei, e à minha tia… estejas onde estiveres o teu olhar e carinho maternal acompanhar-me-ão. à minha prima Lili, que esteve comigo desde o início até o final desta viagem, que foi a minha força nos momentos fracos, que foi o sentido de humor que me animou nas horas mais tristes, que foi a palavra de conforto nas desilusões sentidas… és a minha outra parte! ao meu pai, que possibilitou-me todas as condições para continuar, por ser o sustentáculo das minhas decisões e pelas palavras de apoio e encorajamento desde o início. O teu espírito livre e força fazem parte de mim. à minha mãe… neste momento não tenho palavras para agradecer-te. Tudo o que sou e fiz devo-o a ti, que sempre acreditaste em mim, que ensinaste-me a ver o mundo de olhos fechados, a ouvir a palavra que desperta, a sentir com a alma e com toda a paixão. Cultivaste em mim o gosto pela vida e por tudo o que é belo e etéreo… fizeste com que toda e qualquer experiência na minha vida se tornasse em algo transcendental e incentivaste-me a retirar a essência que toldou o meu ser e o meu caminho. por último, este trabalho foi financiado por Fundos Nacionais através da FCT – Fundação para a Ciência e a Tecnologia no âmbito do projecto PTDC/MAR/098914/2008. Finalmente, a todos aqueles que estiveram presentes no início até/ou no final desta jornada e que acompanharam, incentivaram e inspiraram-me até à derradeira meta mesmo quando eu sentia a perder-me, o meu devoto e sincero obrigado por fazerem parte da minha vida e memórias! Existem momentos inesquecíveis, coisas inexplicáveis e pessoas incomparáveis. Fernando Pessoa xv List of publications The elaboration of this dissertation benefited from the following published or submitted scientific papers and additional conference presentations with a relevant contribution: Magalhães C., Salgado P., Kiene R. P. & Bordalo A. A. (2012) Influence of salinity on dimethyl sulfide and methanethiol formation in estuarine sediments and its side effect on nitrous oxide emissions. Biogeochemistry 110: 75-86 Salgado P., Kiene R. P., Wiebe W. J. & Magalhães C. Salinity as a key regulator of DMSP catabolism pathways in Ruegeria pomeroyi DSS-3. Submitted to Aquatic Microbial Ecology Salgado P., Kiene R. P., Bordalo A. A. & Magalhães C. Influence of salinity on dimethyl sulphide, methanethiol and nitrous oxide oxide emissions. Congresso de Jovens Investigadores de Universidade do Porto (IJUP), Porto, Portugal, February 2011. Salgado P., Bordalo A. A., Kiene R. P. & Magalhães C. Influence of salinity on DMSP and methionine degradation and its regulatory effect on the nitrous oxide reduction step of denitrification. 4th Congress of European Microbiologists FEMS 2011, Geneva, Switzerland, 26-30 June 2011 Salgado P., Kiene R. P., Bordalo A. A. & Magalhães C. Salinity as a key regulator of DMSP catabolism pathways in estuarine sediments and in a Ruegeria pomeroyi pure culture. First EMBO Conference on Aquatic Microbial Ecology: SAME13, Stresa, Italy, 8-13 September 2013 Magalhães C., Salgado P., Buchan A., Machado A., Wiebe W. J., Kiene R. P. How dimethylsulfoniopropionate degradation compounds may control the efficiency of denitrification pathway. First EMBO Conference on Aquatic Microbial Ecology: SAME13, Stresa, Italy, 8-13 September 2013 xvi xvii Table of Contents Agradecimentos ............................................................................................................... vii Resumo ............................................................................................................................ xi Abstract .......................................................................................................................... xiii List of papers ................................................................................................................... xv List of figures .................................................................................................................. xix List of tables .................................................................................................................. xxiii List of abbreviations ....................................................................................................... xxv CHAPTER 1 ..................................................................................................................... 1 Introduction ....................................................................................................................... 1 1.1. Motivation ........................................................................................................... 1 1.2. Background ........................................................................................................ 2 1.2.1. Sulfur Biogeochemistry ................................................................................ 2 1.2.2. Organic sulfur catabolism vs environmental controls ................................... 9 1.2.3. Interactions between sulfur compounds and N2O emissions .......................10 1.3. Goals .................................................................................................................11 CHAPTER 2 ....................................................................................................................12 Material and Methods ......................................................................................................12 2.1. Environmental survey ........................................................................................12 2.1.1. Characterization of sampling area ..............................................................12 2.1.2. Sampling strategy .......................................................................................13 2.1.3. Slurry experiments ......................................................................................14 2.1.4. Water and Sediment characteristics ...........................................................15 2.2. Ruegeria pomeroyi DSS-3 cell cultures experiments .........................................16 2.2.1. Bacteria strain .............................................................................................16 2.2.2. Ruegeria pomeroyi DSS-3 grow conditions ................................................17 2.2.3. Salinity treatments in Ruegeria pomeroyi DSS-3 cell suspensions .............17 2.3. Analytical techniques .........................................................................................17 2.4. Statistical analysis .............................................................................................18 CHAPTER 3 ....................................................................................................................19 The regulatory effect of salinity on dimethyl sulfide and methanethiol formation in estuarine sediments and subsequent interaction on nitrous oxide production ..................19 3.1. Abstract .............................................................................................................19 3.2. Background .......................................................................................................19 3.3. Results ..............................................................................................................21 xviii 3.3.1. Environmental characterization ...................................................................21 3.3.2. DMS vs. MeSH production ..........................................................................23 3.3.3. N2O net accumulation along methionine additions ......................................27 3.4. Discussion .........................................................................................................28 3.4.1. DMS and MeSH production ........................................................................28 3.4.2. Salinity effect on DMS and MeSH potential emissions ................................29 3.4.3. Salinity effect on N2O production during MeSH accumulation .....................31 CHAPTER 4 ....................................................................................................................33 Salinity as a key regulator of DMSP catabolism pathways in Ruegeria pomeroyi DSS-3 .33 4.1. Abstract .............................................................................................................33 4.2. Background .......................................................................................................33 4.3. Results ..............................................................................................................36 4.3.1. R. pomeroyi growing curves at different salinities .......................................36 4.3.2. Salinity effect on R. pomeroyi DSS-3 DMSP cleavage and demethylation catabolism ................................................................................................................36 4.3.3. Side effect of oxic vs. anoxic conditions ......................................................39 4.3.4. Potential chemical production of DMS and MeSH .......................................40 4.4. Discussion .........................................................................................................40 CHAPTER 5 ....................................................................................................................45 Conclusions and future research .....................................................................................45 CHAPTER 6 ....................................................................................................................47 Bibliography .....................................................................................................................47 xix List of figures Figure 1 - Redox sulfur cycle (adapted from Lomans et al. 2002) ..................................... 4 Figure 2 - Representative scheme for the microbial mediated DMSP catabolism and cycling involving the two main biochemical pathways: the demethylation/demethiolation (left) whose final product is acetaldehyde liberating MeSH which can be incorporated into bacterial sulfur-containing amino acids; and cleavage (right) with the hypothetical direct formation of 3HP mediated by lyase DddD, or DMSP can alternatively be converted into acrylate which in turn can be further catalized into 3HP by the AcuN and –K transferases, also releasing DMS with posterior liberation to the atmosphere. DMSPd – dimethylsulfoniopropionate, DMS – dimethyl sulfide, 3HP – 3-hydroxypropionate, MMPA – 3-methiolpropionate, MTA-CoA – methylthioacryloyl-CoA, MeSH – methanethiol. Adapted from Curson et al. (2011b). ............................................................................................... 8 Figure 3 - Ave River estuary and location of sampling sites. ............................................13 Figure 4 - Graphic design of slurry experiments. Replicates of salinity treatments (0, 15, 30 ppt) were incubated under oxic and anoxic conditions and amended with different DMSP and methionine concentrations. DMSPdimethylsulphoniopropionate; Methio – methionine. ......................................................................................................................14 Figure 5 - Net DMS production rates in oxic and anoxic sediment slurries from sites 1 (a), 2 (b), 3 (c) and 4 (d) in the Ave Estuary at three different salinities. For the salinity treatments, sediments from each site were slurried with either Ave River freshwater (0 ppt) or Ave River freshwater amended with different concentrations of sea salts to reach 15 ppt and 30 ppt, respectively. The in situ salinities at each station are given in Table 1. ........................................................................................................................................23 Figure 6 - Net MeSH production rates in oxic and anoxic sediment slurries from sites 1 (a), 2 (b), 3 (c) and 4 (d) in the Ave Estuary at three different salinities. For the salinity treatments, sediments from each site were slurried with either Ave River freshwater (0 ppt) or Ave River freshwater amended with different concentrations of sea salts to reach 15 ppt and 30 ppt, respectively. The in situ salinities at each station are given in Table 1. MeSH accumulation was not detected in any of the treatments in slurries from Station 1, the station with the lowest salinity. ...................................................................................24 xx Figure 7 - The effects of DMSP additions and salinity treatments (0, 15 and 30 ppt) on net DMS accumulation rates in oxic and anoxic sediment slurries from sites 1 (a), 2 (b), 3 (c) and 4 (d) of the Ave Estuary. For the salinity treatments, sediments from each site were slurried with either Ave River freshwater (0 ppt) or Ave River freshwater amended with different concentrations of sea salts to reach 15 ppt and 30 ppt, respectively. .................25 Figure 8 - The effects of methionine additions and salinity treatments (0, 15 and 30 ppt) on net MeSH and N2O accumulation rates in anoxic sediment slurries from sites 1 (a, e), 2 (b, f), 3 (c, g) and 4 (d, h) of the Ave estuary. For the salinity treatments, sediments from each site were slurried with either Ave River freshwater (0 ppt) or Ave River freshwater amended with different concentrations of sea salts to reach 15 and 30 ppt, respectively. 26 Figure 9 - Net N2O production rates in anoxic sediment slurries from sites 1 (a), 2 (b), 3 (c) and 4 (d) in the Ave Estuary at three different salinities. For the salinity treatments, sediments from each site were slurried with either Ave River freshwater (0 ppt) or Ave River freshwater amended with different concentrations of sea salts to reach 15 ppt and 30 ppt, respectively. The in situ salinities at each station are given in Table 1. ...............27 Figure 10 - Growth curves for R. pomeroyi DSS-3 cell culture in MBM (a) and ½ YTSS (b) media in different salinities (10, 20 and 30 ppt). Values were adjusted according to replicates average and the range bars indicate standard deviations. ...............................36 Figure 11 - Net DMS production rates in R. pomeroyi DSS-3 cell suspensions submitted to oxic conditions and simulated gradual salinities in MBM (a) and ½ YTSS (b) media. The salinity treatments (10, 20, 30 ppt) were amended with 10 mM Glucose and different DMSP concentrations. .....................................................................................................37 Figure 12 - Net MeSH production rates in R. pomeroyi DSS-3 cell suspensions submitted to oxic conditions and simulated gradual salinities in MBM (a) and ½ YTSS (b) media. The salinity treatments (10, 20, 30 ppt) were amended with 10 mM Glucose and different DMSP concentrations. .....................................................................................................38 Figure 13 - Net DMS and MeSH accumulation rates for the different salinity treatments (10, 20, 30 ppt) amended with 500 μM DMSP in R. pomeroyi DSS-3 cell suspensions under oxic conditions on MBM (a) and ½ YTSS (b) media with 10 mM Glucose addition. a MeSH production was not detected. ................................................................................38 Figure 14 - Net DMS and MeSH accumulation rates for the different salinity treatments (10, 20, 30 ppt) amended with 500 μM DMSP in R. pomeroyi DSS-3 cell suspensions xxi under anoxic conditions on MBM (a) and ½ YTSS (b) media with 10 mM Glucose addition. ª DMS production was not detected. ................................................................................39 xxii xxiii List of tables Table 1 – Oxidation numbers of Sulfur compounds (adapted from Bentley et al. 2002) .... 3 Table 2 - Physical and chemical characteristics of sediments and overlying water at the sampling sites in the Ave Estuary. ...................................................................................22 xxiv Chapter 1 5 al. 2012). Thus, SO42is an essential compound in several assimilatory and dissimilatory pathways occurring in soils, sediments and water, participating in the sulfur ocean-land transfer (Kwint et al. 1996). Nevertheless, sulfate oxidized sulfur, bio-energetically, represents a disadvantage in what constitutes the biochemical processes of sulfur incorporation into amino acids, requiring a considerable amount of energy on assimilatory reductions (Likens et al. 2002). Therefore, the availability of organic reduced sulfur compounds such as DMSP benefits the above mentioned biochemical transformations through a less energy yield leading, subsequently, to a quick assimilation into bacterial biomass (Kiene et al. 1999). 1.2.1.2. Organic Sulfur Oceanic sulfur flux constitutes one of the most important reservoir for sulfur emissions where the organic compound DMS, among other volatile sulfur gases (e.g. methanethiol, MeSH; carbon disulfide, CS2; hydrogen sulfide, H2S; carbonyl sulfide, COS), stands for the most abundant biogenic fraction that contributes to maintain an atmospheric balance (Charlson et al. 1987, Andreae 1990, Bates et al. 1992, Visscher et al. 2003, Vallina & Simó 2007). In 1972 Lovelock and coauthors emphasized for the first time the importance of the biogeochemical cycle of sulfur through the oceanic DMS emissions to the atmosphere, later estimated to represent a flux of ca. 30% biogenic sulfur, with a contribution of 14% by the Northern Hemisphere and 67% from the Southern Hemisphere (Kloster et al. 2006). After being accepted that DMS was the “missing link” between the ocean and atmosphere sulfur transfer and, consequently, the dominant oceanic sulfur compound emitted to the atmosphere, DMS became the topic of several ecological and biogeochemical studies concerning aquatic systems. Therefore, aquatic environments such as salt marshes/estuarine systems, coastal wetlands and oceans, which emitted high concentrations of volatile organic sulfur, were recognized as important contributors to the global sulfur cycle and, consequently, in shaping earth’s atmosphere and climate (Steudler & Peterson 1984; Kiene & Visscher 1987; Aneja & Cooper 1989). DMS is the most important source of biogenic sulfur derived from DMSP biosynthesized by phytoplankton (among other sources, as described below). DMS has a low residence time and is readily emitted to the atmosphere where it reacts with other species to produce a particulate-phase containing SO42which, in turn, composes most of the cloud condensation nuclei – CCN (Sievert et al. 2007). Hence, an increase in marine DMS Chapter 1 6 emissions would result in a CCN increase and clouds albedo, with a consequent decrease in incoming solar radiation to Earth’s surface (Charlson et al. 1987). However, recent scientific studies have acknowledged the complexity of the CLAW hypothesis and there are several uncertainties relating to this hypothetical climate feedback between DMS and plankton. Quinn and Bates (2011) have reviewed several studies that tried to support the DMS-climate feedback loop theory and verified that CCN has other important sources such as wind-driven sea-salt particles and organic matter and it also depends on particle growth. Besides its influence on global climate, biogenic sulfur compounds play a fundamental role in microorganisms’ metabolism mainly through the assimilation of organic sulfur into biomolecules such as methionine and cysteine (Cooper 1983, Lomans et al. 2002). In fact, inorganic sulfates, which are the most stable and abundant sulfur source in seawater, were considered for a long time an important constituent in the assimilatory pathway by aquatic microorganisms (Jørgensen 1977a, b, Middleton & Lawrence 1977, Widdel & Pfennig 1981, Cuhel et al. 1982, Sievert et al. 2007). However, as abovementioned, this inorganic sulfate yields a high energetic cost for microorganisms due to its sulfur in the most oxidized form (+6; Kiene et al. 1999). Therefore, DMSP, an organic sulfur compound produced in large amounts by numerous species of marine phytoplankton (e.g. Stefels 2000), seaweeds (Reed 1983) and salt marsh grasses (Otte et al. 2004), represents an advantage due to its low and reduced molecular weight (Kiene et al. 1999, Kiene et al. 2000). This organic sulfur compound, also abundant in seawater (an average of < 2.8 nM, Kiene & Slezak 2006), is widely used as a growth substrate by bacterioplankton (Ledyard & Dacey 1994) which rapidly metabolize it and incorporate sulfur into biomass. 1.2.1.3. Organic Sulfur Catabolism The metabolism of organic sulfur represents an important component on the global sulfur cycle. In particular, the catabolism of DMSP which constitutes ca. 24% of the total organic sulfur in surface seawater (Bates et al. 1994) and has now been thoroughly studied namely due to its role as a precursor of DMS and MeSH (Groene 1995, Kiene 1996, Kiene et al. 2000). The DMSP biosynthesis was a process initially approached in higher plants (Greene 1962, Hanson et al. 1994, Trossat et al. 1996) and, later on, elucidated in marine algae (Gage et al. 1997, Summers et al. 1998, Stefels 2000), which confirmed methionine as the precursor protein for DMSP production through different initial routes Chapter 1 7 (transamination vs. methylation) according to the respective organism (Otte et al. 2004). DMSP physiological functions are still yet to be fully defined, however its high intracellular concentrations and compatibility with cell metabolism confer to DMSP a main osmolytic function (Dickson & Kirst 1987, Stefels 2000). Additionally, this solute serves as antioxidant (Sunda et al. 2002, Husband et al. 2012), cryoprotectant (Karsten et al. 1990, Kirst et al. 1991) and herbivore deterrent (Van Alstyne et al. 2001, Strom et al. 2003, Fredrickson & Strom 2009). Also, Stefels (2000) hypothesized an additional buffering capacity of DMSP when reduced sulfur exceeds cell’s ability to convert it into amino acids. More recently, Geng & Belas (2010) suggested DMSP as a trigger molecule under chemotactic response acting in benefit for the phytoplankton-Roseobacter symbiosis. Higher plants and marine algae release particulate DMSP into the extracellular environment by several processes which include viral lysis (Hill et al. 1998), physiological stress (Mulholland & Otte 2002), algal senescence (Yoch 2002 and references therein), zooplankton grazing upon phytoplankton blooms (Wolfe et al. 1997, Archer et al. 2003), exudation (Laroche et al. 1999) and cell lysis (Nguyen et al. 1988, Simó 2004). Consequently, dissolved DMSP (DMSPd) assimilation and its degradation compounds (e.g. DMS, MeSH) production might be enhanced, since this labile organic sulfur compound is now available for marine prokaryotes, representing an important ecological role in the marine carbon and sulfur cycles (Kiene and Bates 1990; Kiene et al. 2000; Kiene and Linn 2000a, b, Yoch 2002). DMSP degradation undergoes a rapid turnover by essentially aerobic microorganisms (Kiene 1990, Taylor & Gilchrist 1991, Visscher et al. 1992, Visscher & Taylor 1994, Zubkov et al. 2001), without excluding its catabolism in alternative anoxic marine communities (Kiene & Taylor 1988, Van Der Maarel et al. 1993, Jonkers et al. 1998). Recent scientific research has focused on two main DMSP degradation pathways – cleavage vs. demethylation/demethiolation (Kiene et al. 2000, Yoch 2002), due to its ecological and biogeochemical relevancy. In fact, cleavage presents itself a considerable enzymatic process resulting in the production of DMS by DMSPd bacterial turnover (Curson et al. 2011b) and posterior liberation of DMS into the atmosphere contributing for the regulation of global climate (Fig. 2). Chapter 1 8 Figure 2 - Representative scheme for the microbial mediated DMSP catabolism and cycling involving the two main biochemical pathways: the demethylation/demethiolation (left) whose final product is acetaldehyde liberating MeSH which can be incorporated into bacterial sulfur-containing amino acids; and cleavage (right) with the hypothetical direct formation of 3HP mediated by lyase DddD, or DMSP can alternatively be converted into acrylate which in turn can be further catalized into 3HP by the AcuN and –K transferases, also releasing DMS with posterior liberation to the atmosphere. DMSPd – dimethylsulfoniopropionate, DMS – dimethyl sulfide, 3HP – 3hydroxypropionate, MMPA – 3-methiolpropionate, MTA-CoA – methylthioacryloyl-CoA, MeSH – methanethiol. Adapted from Curson et al. (2011b). Chapter 1 9 On the other hand, DMSP catabolism can be switched to another biological transformation in which the end product, MeSH, is ca. 75% assimilated by bacteria for subsequent incorporation into amino acids (Kiene & Linn 2000a). This latter process, mediated by marine bacteria, seems to be the preferential enzymatic pathway in seawater (Kiene 1996, Kiene & Linn 2000b), but what controls the “bacterial switch” between the two main pathways of DMSP degradation still remains unanswered and reveals itself as a quest for a better determination of surface ocean DMS concentrations. The progress of genomic and transcriptomic technologies have contributed to unveil some of the complex steps implicated on DMSP catabolism, namely through genes identification (Todd et al. 2007, Todd et al. 2010, Varaljay et al. 2010, Curson et al. 2011a, Todd et al. 2011, Todd et al. 2012, Howard et al. 2011, Moran et al. 2012, Varaljay et al. 2012). 1.2.2. Organic sulfur catabolism vs environmental controls Potential variability in the dynamics of coastal and oceanic ecosystems is likely a consequence of alterations in several ecological features. Specifically, shifts in abiotic factors can lead to redefinitions of chemical and microbiological patterns as well as community composition (Bouvier & del Giorgio 2002, Herlemann et al. 2011, Shah & Shah 2011). Several studies have focused on the influence of different biotic and abiotic factors on phytoplankton and macroalgae DMSP intracellular concentrations (Gage et al. 1997, Sunda et al. 2002, Malmstrom et al. 2004, Pinhassi et al. 2005, Van Alstyne & Puglisi 2007, Kumar et al. 2009, Yang et al. 2011, Husband et al. 2012). Since several algae can accumulate low molecular weight organic solutes, osmotic stress affects the physiology of phytoplankton and macroalgae and their intracellular DMSP accumulation (Kirst 1990). In fact, salinity has been identified as a relevant factor on the physiology of phytoplankton and macroalgae reflecting an increase of intracellular DMSP by the osmotic potential of the cell (Vairavamurthy et al. 1985, Stefels 2000, Yang et al. 2011). While there are relevant studies about the physiological behaviour of phytoplankton and macroalgae and subsequent influence of biotic and abiotic factors on DMSP intracellular concentrations (e.g. Vairavamurthy et al. 1985; Van Alstyne et al. 2003; Stefels et al. 1996, Yang et al. 2011, Husband et al. 2012), relatively few studies have addressed the environmental controls on the pathways of DMSP degradation. It is, however, expected that numerous environmental and biological factors may control DMSP catabolism to DMS (cleavage) versus MeSH (demethylation/demethiolation), with certainly implications on the different production ratios of DMS and MeSH in marine environments. Indeed, there are evidences Chapter 1 10 that the type of microbial species assemblage, seems to be linked to the dynamics of phytoplankton blooms, which then affects DMSP turnover (Jonkers et al. 1998, Zubkov et al. 2001, Pinhassi et al. 2005). Nevertheless, apart from work from the present research team and two other studies that hypothesized salinity as a relevant abiotic factor controlling DMSP catabolism in natural systems (Visscher et al. 2003; Niki et al. 2007, Magalhães et al. 2012), there is still a gap about the environmental controls on the pathways of bacterial degradation of DMSPd to DMS (cleavage) versus MeSH (demethylation/demethiolation). 1.2.3. Interactions between sulfur compounds and N2O emissions Nitrous oxide (N2O) is partially responsible for the destruction of the stratospheric ozone through the reaction with the atomic oxygen, which consequently potentiates the increase of harmful UV-B radiation (Krupa & Kickert 1989, Caldwell & Flint 1994, Nevison & Holland 1997) and was recently identified as the dominant ozone-depleting substance (Ravishankara et al. 2009). Nitrous oxide acts also as a potent greenhouse agent in the atmosphere (300 times more potent than CO2) and represents a particular environmental problem due to its long atmospheric lifetime of 114 years (Dickinson and Cicerone, 1986). Soils and oceans represent the largest sources of N2O emissions, however, anthropogenic sources, such as agriculture, sewage treatments, or combustion of fossil fuel, account for almost two-thirds of the total emissions (Denman et al 2007). Different microbial nitrogen transforming processes, such as microbial nitrification and heterotrophic denitrification, contribute to the formation of N2O and soils represent major sources (Seitzinger 2000, Frame & Casciotti 2010). Therefore, nowadays concern is focused namely on the consequences of climate change aggravated by greenhouse gases and the demand for mitigation measures. Interestingly, a previous study, motivated by the observed production of nitric and nitrous oxides in the redox transition zone of a coastal marine sediment [by denitrifying bacteria] near the sulfide maxima concentration (Sørensen 1978), lead to the analysis and consequent corroboration of a sulfide inhibition on the nitrous oxide reduction pathway of denitrification (Sørensen et al. 1980). This interaction represented a novel quest to better understand the nitrogen and sulfur biogeochemical processes inherent to marine and freshwater environments. Later, Joye & Hollibaugh (1995) also identified a sulfide inhibition of nitrification through experiments with HSadditions within a range to estuarine Chapter 1 11 sediments, enhancing a link between both sulfur and nitrogen cycles. The apparent regulation of the N cycle opened a precedent for researching new and possible sulfur compounds which could influence the nitrogen biogeochemical pathways. In fact, more recently (Magalhães et al. 2012) it was detected a novel interaction between both sulfur and nitrogen biogeochemical cycles, specifically through an interference on the last reductase step of denitrification (reduction of N2O to N2) triggered by a DMSP breakdown product – MeSH (Magalhães et al. 2011). 1.3. Goals The main objective of this study was to investigate the influence of an environmental stressor, salinity, in selecting DMSP degradation routes and to understand how salinity could modulate N2O fluxes through its effect on MeSH and DMS net production. In order to achieve this goal, two chapters (Chapter 3 and 4) are presented with the following detailed objectives: In Chapter 3 we studied the influence of salinity on the DMS vs MeSH net fluxes in sediment microbial communities along the salinity gradient of Ave River estuary; the interaction between sulfur volatiles accumulation (MeSH and DMS) and N2O fluxes, was also evaluated is these natural microbial communities. In Chapter 4 we focused in a more specific evaluation of osmotic pressures onto DMSP catabolism and consequent DMS and MeSH production using a model organism – Ruegeria pomeroyi DSS-3. Here we aim to evaluate the salinity influence on the bacterial preference between the two main DMSP enzymatic pathways – cleavage and demethylation/demethiolation – expressed by the proportion of DMS vs. MeSH net fluxes. This dissertation is integrated in a research project untitled NITROSUL – Novel interaction between marine biogeochemical nitrogen and sulfur cycles: characterization and ecological implications and it represents a follow-up of previous research focusing on the inhibitory interaction between the DMSP degradation products and the last reductase step of denitrification (conversion of N2O to N2). Chapter 2 12 CHAPTER 2 Material and Methods Methodology was organized taking into account the sequence of the experiments performed in order to first evaluate the influence of salinity on DMS and MeSH emissions in estuarine sediments from Ave estuary, and secondly to restrict a more detailed characterization of the salinity effect into DMS and MeSH formation in cell cultures of Ruegeria pomeroyi DSS-3. 2.1. Environmental survey 2.1.1. Characterization of sampling area Ave River is a mesotidal estuary located on the northern coast of Portugal with a 1391 km2 drainage basin, a 40 m3/s average discharge, lithologically composed by granitic (quartz and feldspars) and schist rocks and by silty-clay sediments, with mainly marine origin organic matter (Mil-Homens et al. 2006). Ave River is 94 km (E-W) long, from the spring – Cabreira mountains – till the mouth located at the south of Vila do Conde and its catchment presents a variable annual precipitation between 900 – 3900 mm, which 73% occurs during the wet season (October – March; ARHNorte). The main tributaries are the Este River, on the right edge, and Vizela River, on the left edge, draining 247 km2 and 340 km2, respectively. The Ave estuary is bordered by riparian vegetation dominated by Eucalyptus globulus, Juncus sp., Alnus glutinosa, Populus sp. (Pascoal et al. 2005). Ave River basin covers 15 locations and its estuary is described as one of the most contaminated on the northwest Portuguese region, mainly due to domestic sludge and textile, leather and paper industries which directly discharged their untreated effluents into the streams (Soares et al. 1999; Cunha et al. 2005). Since 1985, several infra-structures were created with the purpose of watershed rehabilitation through e.g. an indicator system of water quality (Oliveira et al. 2005). Also, Ave River basin has been the focus of some heavy metal assessment studies which have corroborated industrial effluent contamination (e.g. Araújo et al. 1998; Soares et al. 1999; Cunha et al. 2005; Mil-Homens Chapter 2 13 et al. 2006). Water column of Ave River estuary ranges a salinity gradient from < 2 ppt upstream, an oligohaline zone, to almost 35 ppt downstream (Da Silva 2011). 2.1.2. Sampling strategy Sampling was performed in September 2010 during low tide, covering the north margin of Ave estuary, at four stations along the riparian zone of Vila do Conde (Fig. 3); the sampling stations were meticulously chosen so that it could be representative of the estuarine salinity gradient. At each site, a total of twenty cores (3 cm diameter and 8 cm long) were collected within 50 cm of each other, and all within of approximately 3 m2. The twenty cores collected from each site were homogenized and used as a composite sample. Also, 0.5 L of sub-superficial water from each sampling station and an additional 5 L sample of freshwater (0 ppt) from an upstream location were collected in acid-cleaned polyethylene bottles. The Ave River freshwater was used for the salinity treatments. All the samples were kept in the dark and transported in refrigerated isothermal containers for posterior processing. Figure 3 - Ave River estuary and location of sampling sites. Additionally, salinity and temperature were measured in situ, at the specific sampling site, with YSI Model 30 probe. At the laboratory, all the water samples were immediately filtered through 0.45 μm pore diameter (Ф) membrane filter (GF/F glass fiber filters, AVE estuary 1 2 3 2 3 4 1 3 4 2 3 4 1 2 3 4 AVE estuary 1 2 3 4Spain AVE estuary 1 2 3 2 3 4 1 3 4 2 3 4 1 2 3 4 AVE estuary 1 2 3 4 AVE estuary 1 2 3 2 3 4 1 3 4 2 3 4 1 2 3 4 AVE estuary 1 2 3 4 AVE estuaryAVE estuaryAVE estuaryAVE estuary 1 2 3 22 33 44 11 33 44 22 33 44 11 22 33 44 AVE estuaryAVE estuaryAVE estuary 1 2 3 4Spain Chapter 2 14 Whatman) and a sub-sample stored at -20ºC for nutrient analysis. Sediment samples were stored at 4ºC and used after no more than 4 days after collection for the slurry experiments and sediment analysis. 2.1.3. Slurry experiments Salinity simulations (15 and 30 ppt) were prepared by, initially, adding artificial sea salts to Ave estuary freshwater (0 ppt) according to Cavanaugh’s formula (1975). This way, it reassures salinity as the only variable in slurry treatments, preserving water chemistry. Sediment slurries consisted in a mix of 5 g of sediment from each sampling site, and 10 ml of overlying water with the different simulated salinities (0, 15 and 30 ppt) in 30 ml serum bottles, as previously described (Magalhães et al. 2011, 2012). For each salinity 3 0 ppt 3 15 ppt 3 30 ppt oxic anoxic 5 µM DMSP 50 µM DMSP 5 µM Methio 50 µM Methio 500 µM Methio 5 µM DMSP 50 µM DMSP 5 µM Methio 50 µM Methio 500 µM Methio Salinity Amendments Figure 4 - Graphic design of slurry experiments. Replicates of salinity treatments (0, 15, 30 ppt) were incubated under oxic and anoxic conditions and amended with different DMSP and methionine concentrations. DMSPdimethylsulphoniopropionate; Methio – methionine. Chapter 3 21 3.3. Results 3.3.1. Environmental characterization Salinity between the sampling sites comprised values between 2.1 ppt and 15.6 ppt, upstream and downstream Ave’s estuary, respectively (Table 2; Fig. 3). The granulometry analysis revealed a tendency towards a sandy sediment (> 2000 µm) downstream’s estuary. Among the inorganic nutrients, NO3revealed a substantial variation (89.2 ± 4.0 to 160.3 ± 12.1 µM; Table 2), with the highest concentration registered at A-1 (160.3 ± 12.1 µM; Table 2), an upstream sampling site. DMSP concentrations in sediment correlated to Chl a ones (r = 0.97, p < 0.05), where A-1 and A-4 presented the highest DMSP values (16.0 ± 2.9 moles L-1 wet sed and 35.5 ± 6.7 moles L-1 wet sed, respectively). Chapter 3 22 Table 2 - Physical and chemical characteristics of sediments and overlying water at the sampling sites in the Ave Estuary. Characteristics of overlying water Characteristics of sediments Dates Sites code Salinity (ppt) N inorganic (µM) NO3NH4+ NO2Chl a (mg g1wet sed) DMSP (µmoles L-1 wet sed) OM (%) Grain Size (% dry weight) < 63µm > 2000 µm 09/21/2010 1 2.1 160.3 ± 12.1 12.2 ± 0.7 6.0 ± 0.1 10.0 ± 2.1 16.0 ± 2.9 2.5 ± 0.2 8.3 15.5 09/21/2010 2 5.2 100.0 ± 3.1 12.5 ± 0.7 4.7 ± 0.0 7.1 ± 0.5 3.8 ± 0.5 2.3 ± 1.0 6.9 28.5 09/21/2010 3 12.4 77.4 ± 1.4 12.4 ± 0.8 3.7 ± 0.0 3.1 ± 0.3 2.6 ± 0.6 1.8 ± 0.2 3.2 20.6 09/21/2010 4 15.6 89.2 ± 4.0 8.1 ± 5.4 4.0 ± 0.1 18.9 ± 2.6 35.5 ± 6.7 1.2 ± 0.0 0.8 33.6 Chapter 3 23 3.3.2. DMS vs. MeSH production Generally, DMS production exceeded MeSH in all sampling sites submitted to different salinity treatments in both oxic and anoxic conditions (Fig. 5, 6). Figure 5 - Net DMS production rates in oxic and anoxic sediment slurries from sites 1 (a), 2 (b), 3 (c) and 4 (d) in the Ave Estuary at three different salinities. For the salinity treatments, sediments from each site were slurried with either Ave River freshwater (0 ppt) or Ave River freshwater amended with different concentrations of sea salts to reach 15 ppt and 30 ppt, respectively. The in situ salinities at each station are given in Table 1. DMS accumulations between incubations with and without oxygen were variable according with to site and the salinity treatment. MeSH accumulations occurred with more extent under anoxic conditions but its net production wasn’t observed in sampling site A-1, upstream Ave’s estuary, regardless of the experimental salinity (Fig. 6a). Chapter 3 24 Figure 6 - Net MeSH production rates in oxic and anoxic sediment slurries from sites 1 (a), 2 (b), 3 (c) and 4 (d) in the Ave Estuary at three different salinities. For the salinity treatments, sediments from each site were slurried with either Ave River freshwater (0 ppt) or Ave River freshwater amended with different concentrations of sea salts to reach 15 ppt and 30 ppt, respectively. The in situ salinities at each station are given in Table 1. MeSH accumulation was not detected in any of the treatments in slurries from Station 1, the station with the lowest salinity. These salinity manipulation experiments revealed an opposite pattern between DMS and MeSH accumulations i.e., while the production of DMS is enhanced in the freshwater treatments (0 ppt) MeSH seems to be utmost expressed at highest salinities (30 ppt). Isolated, all sediment sampling sites demonstrated a consistent salinity effect on DMS net accumulations, regardless of the in situ salinity of the station (Fig. 5, Table 2). Therefore, higher DMS net accumulations were always observed in freshwater treatments (0 ppt) decreasing its production rates towards the highest salinity treatments and this was a coherent pattern in both oxic and anoxic conditions (Fig. 5). In contrast, net MeSH production was considerably lower in oxic conditions and was not detected in majority of the sampling sites (Fig. 6). Nevertheless, anoxic manipulation experiments showed a general net MeSH accumulation, with high rates observed at the highest salinities (30 ppt) and less net effluxes of MeSH in freshwater treatments (0 ppt; Fig. 6). Chapter 3 25 Regarding the substrate amendments (DMSP and methionine), both sulfur volatiles breakdown products (DMS and MeSH) demonstrated the same opposite pattern as determined on the non-amended incubations above mentioned. In all DMSP additions DMS displayed lower production in the highest salinity treatments (30 ppt) and highest net accumulation in the lowest salinities (0 ppt; Fig. 7). This same pattern was found to be coherent for all sediments, collected in different stations, incubated with or without oxygen (Fig. 7). For the methionine amendments, sediment slurries incubated in anoxic conditions confirmed the same opposite pattern previously mentioned (Fig. 8a, b, d); all sampling sites, except for A-3 (Fig. 8c), presented the higher MeSH net accumulation in the highest salinity treatments (30 ppt) and lowest production in 0 ppt treatments. Figure 7 - The effects of DMSP additions and salinity treatments (0, 15 and 30 ppt) on net DMS accumulation rates in oxic and anoxic sediment slurries from sites 1 (a), 2 (b), 3 (c) and 4 (d) of the Ave Estuary. For the salinity treatments, sediments from each site were slurried with either Ave River freshwater (0 ppt) or Ave River freshwater amended with different concentrations of sea salts to reach 15 ppt and 30 ppt, respectively. Chapter 3 26 Figure 8 - The effects of methionine additions and salinity treatments (0, 15 and 30 ppt) on net MeSH and N2O accumulation rates in anoxic sediment slurries from sites 1 (a, e), 2 (b, f), 3 (c, g) and 4 (d, h) of the Ave estuary. For the salinity treatments, sediments from each site were slurried with either Ave River freshwater (0 ppt) or Ave River freshwater amended with different concentrations of sea salts to reach 15 and 30 ppt, respectively. Chapter 3 27 3.3.3. N2O net accumulation along methionine additions For non-amended sediment slurries incubated under anoxic conditions, generally N2O accumulated to a greater extent in the 15 ppt salinity treatments compared to the freshwater (0 ppt) treatment (Fig. 9). Figure 9 - Net N2O production rates in anoxic sediment slurries from sites 1 (a), 2 (b), 3 (c) and 4 (d) in the Ave Estuary at three different salinities. For the salinity treatments, sediments from each site were slurried with either Ave River freshwater (0 ppt) or Ave River freshwater amended with different concentrations of sea salts to reach 15 ppt and 30 ppt, respectively. The in situ salinities at each station are given in Table 1. Also, the higher salinity (30 ppt) treatments tend to reduce potential N2O net accumulation compared to 15 ppt slurries (Fig. 9). In treatments where methionine was added, results showed high N2O accumulations in the intermediate salinity treatments (Fig. 8e, f, g, h), which is in agreement to the results observed in the non-amended sediment slurries (Fig. 9). Chapter 3 28 3.4. Discussion 3.4.1. DMS and MeSH production In natural environments DMSP profiles can vary with seasonality, DMSP-producers composition, phytoplankton blooms (Yoch 2002, Simó et al. 2009). DMSP and its related thiols present higher concentrations associated to sediments than to water column, and Chl a concentrations usually correspond to DMSP (Kiene 1991, Nedwell et al. 1994; Jonkers et al. 1998, Trevena et al. 2000). This correlation is an indication of an algal biomass source for DMSP, an osmolyte mainly synthesized by several photoautotrophic primary producers in marine, estuarine and salt-marsh systems (Karsten et al. 1990, Kirst et al. 1991, Stefels & Van Boekel 1993, Stefels 2000, Van Alstyne et al. 2003, Yang et al. 2011). Results from this study show higher rates of net DMS production at sites where higher DMSP concentration were measured, an observation that is likely due to the fact that DMSP is a major precursor of DMS (Kiene and Bates 1990). In fact, in marine environments DMSP is perceived as the main source of DMS (Stefels & Van Boekel 1993, Van Alstyne & Puglisi 2007), a metabolism which involves different enzymes (Fig. 2; Todd et al. 2009, Todd et al. 2010, Curson et al. 2011b, Moran et al. 2012, Todd et al. 2012). On the other hand, several other studies have addressed alternative sources for DMS formation; DMS can also be produced through a respiratory reduction of DMSO (Griebler 1997, López & Duarte 2004), methylation of MeSH (Kiene & Hines 1995) and anaerobic degradation from methoxylated aromatic compounds under high sulfide concentrations (Bak et al. 1992, Lomans et al. 1997). Even though the present study is unable to differentiate what processes are specifically involved in these estuarine sediments of Ave River, the enzymatic degradation of DMSP represents a relevant biological process, mediated by bacteria, leading to DMS production. Steady-state concentrations and fluxes of DMS and MeSH in the sediment and surface waters depend on complex microbial formation and degradation processes (Kiene 1996, Lomans et al. 1997, Kiene et al. 2000), as well as on complex physicochemical interactions between the different natural elements (Mopper and Taylor 1986). In the sediment slurries from the Ave estuary, the production of DMS always exceeded consumption, resulting in net DMS accumulation. In contrast, MeSH accumulations under oxic conditions were much less important than under anoxic conditions. Actually, MeSH is likely to be subject to oxidation in the presence of O2 (Suylen et al. 1987), but its oxidative metabolism is poorly understood. The absence of MeSH accumulation in oxic treatments has been suggested to be a result of rapid MeSH oxidation to dimethyl disulfide (DMDS) Chapter 3 29 in the presence of oxygen (Visscher et al. 2003) and also, in anaerobic lake sediments, microbial populations can metabolize MeSH into methane and carbon dioxide (Zinder & Brock 1978). Thus, it remains unclear whether lack of production or simply high rates of consumption were responsible for the minimal MeSH accumulations obtained. MeSH accumulation in anoxic treatments was not related to DMSP availability in the sediments suggesting that other pathways of MeSH formation besides DMSP demethylation/demethiolation (Kiene 1996; Kiene et al. 2000) might be important. A multitude of MeSH production pathways may contribute to the MeSH fluxes in our anoxic incubations, including methylation of hydrogen sulfide (Mopper and Taylor 1986, Lomans et al., 1997; 1999), sulfide methylation by methoxylated aromatic compounds (Lomans et al. 2001) or by degradation of sulfur-containing amino acids (Lomans et al. 1997; Lomans et al. 1999). 3.4.2. Salinity effect on DMS and MeSH potential emissions Higher DMS production rates were observed in freshwater treatments (0 ppt), while higher salinity treatments resulted in the lowest rates of DMS accumulation. This pattern of DMS accumulation in the different salinity treatments was found to be coherent in the oxic and anoxic treatments. The role of salinity in regulating DMS flux to the atmosphere has been identified in previous research performed on microbial mats grown in hypersaline ponds with different salinities (Visscher et al. 2003). Higher DMS net accumulation was found in a lower salinity pond (90 ppt) compared with a higher salinity pond (115 ppt) (Visscher et al. 2003). While such high salinities were not tested in our study, our results are in agreement with these previous findings, suggesting that lower salinities stimulated DMS production rates. Salinity fluctuations have a direct impact on intracellular DMSP concentrations of microand macroalgae inhabiting intertidal estuarine environments, where the osmotic function of DMSP has been described (Edwards et al. 1987). Intracellular concentrations of DMSP tend to decrease in response to a decrease in salinity and DMSP biosynthesis increase in higher salinities (Vairavamurthy et al. 1985; Dickson and Kirst, 1987; Edwards et al. 1987). Thus, we hypothesize that the rapid release of DMSP from cells at lower salinities may increase the availability of DMSP to the DMSP-degrading microbial communities, resulting in high levels of DMS accumulation. In fact, Van Bergeijk et al. (2003) described DMSP excretion into the medium by the marine benthic diatom Cylindrotheca closterium after salinity down-shock, which means that there’s an enhancement in extracellular DMSP resulting in high levels of DMS Chapter 3 30 accumulation through the cleavage pathway. Niki et al. (2007) also identified salinity as an important factor controlling the production of DMS in the water column of a coastal bay of Japan. In agreement with the present results, it has been found that low-salinity shock leads to an increase in potential DMS production to the environment (Niki et al. 2007). In addition, DMSP-lyases aren’t exclusive bacterial isozymes and have already been identified in several marine microand macroalgae (De Souza & Yoch 1996, Stefels & Dijkhuizen 1996, Steinke et al. 1996, Steinke et al. 1998), representing a further contribution for DMS formation, besides bacterial production (Niki et al. 2000). In fact, low salinity was found to enhance algal DMSP lyase activity (c.f. Stefels 2000; Steinke et al. 2002), which can increase the relative contribution of algal DMS production compared to bacterial production of DMS. However, optimal performance of the different marine algae lyase activity requires specific chloride concentrations (Steinke & Kirst 1996). These data on net potential DMS productivity do not allow to discriminate between algal vs bacterial DMS production, but in light of the high Chl a levels found, the algal role in DMS production may be relevant in Ave estuarine sediments and should be considered in future work. DMSP additions at different salinity treatments showed a progressive increase in DMS accumulation with the same pattern of salinity effects observed for the non-amended sediments; i.e. higher rates of DMS accumulation in the freshwater treatments. These results together suggest that salinity alone influences the degradation rates of extracellular DMSP and methionine. The inverse relationship between DMS accumulation rate and salinity can be attributed to either enhanced production of DMS from DMSP and methionine or suppressed decomposition of DMS at lower salinities. Salinity also affected net MeSH production but in the opposite sense from that observed with DMS. MeSH accumulation was not detected at all Ave Estuarine sites, but when rates were measurable, higher values were registered for the higher salinity treatments. These results are not in agreement with a previous study (Visscher et al. 2003), which found an increase in MeSH flux resulting from decreased salinity in hypersaline microbial mats. However, the similar regulatory effect of salinity on MeSH accumulation observed in slurries without and with added methionine suggests that sulfur containing amino acids can be potential precursors of MeSH in intertidal Ave Estuary sediments. Other factors may promote MeSH accumulation in these sediments, such as higher sulfide concentrations and lower iron oxide availability (Mopper and Taylor 1986). Higher sulfide levels in more saline sediments would be expected due to the very high sulfate concentrations in seawater (Capone and Kiene, 1988). Chapter 4 37 more evident in the 500 µM DMSP amendment. Thus, the highest DMS accumulations were always registered for the lowest salinity tested in both media and for any given concentration of DMSP. In MBM and with the 500 µM DMSP amendment, 89.1% more DMS was accumulated in the 10 ppt compared with the 20 ppt salinity treatment (Fig. 11a). Additionally, in ½ YTSS medium, amended with 500 µM DMSP, 77.3% lower DMS accumulation was observed in the 20 ppt compared to the 10 ppt salinity treatments and 57.9% less DMS accumulation was registered in the 30 ppt salinity treatment compared to the 20 ppt (Fig. 11b) media at the highest DMSP amendment. In the MBM, MeSH was detected only in the 500 μM DMSP amendment (Fig. 12a), with the highest value occurring in the 30 ppt salinity treatment (568.5 ± 47.9 pmoles h-1 ml-1 cell susp, p <0.001). There was an increase of 90.6% from the 20 ppt to the 30 ppt salinity treatments and no detected production at the lowest salinity tested (10 ppt) (Fig. 12a). Figure 11 - Net DMS production rates in R. pomeroyi DSS-3 cell suspensions submitted to oxic conditions and simulated gradual salinities in MBM (a) and ½ YTSS (b) media. The salinity treatments (10, 20, 30 ppt) were amended with 10 mM Glucose and different DMSP concentrations. Regarding the ½ YTSS medium (Fig. 12b), while it stimulated a high level of MeSH accumulation, smaller increases of MeSH accumulation were observed as salinity increased; MeSH accumulation increased 33.9% between 10 ppt to 20 ppt salinity treatments amended with 500 μM DMSP (p < 0.005). For treatments amended with 50 μM DMSP (p > 0.05; Fig. 12b) there were no significant differences in MeSH accumulation between the different salinities. Interestingly, increasing salinity had the opposite effect on MeSH accumulation, with higher magnitudes of MeSH accumulation in treatments with higher salinities for MBM and ½ YTSS media at the highest DMSP amendment (Fig. 12). a) b) Chapter 4 38 Figure 12 - Net MeSH production rates in R. pomeroyi DSS-3 cell suspensions submitted to oxic conditions and simulated gradual salinities in MBM (a) and ½ YTSS (b) media. The salinity treatments (10, 20, 30 ppt) were amended with 10 mM Glucose and different DMSP concentrations. In MBM medium net DMS accumulation rates were significantly higher than MeSH accumulations in all treatments (Fig. 13a). On the other hand, in ½ YTSS medium MeSH production was higher than that of DMS in almost all the simulated conditions (Fig. 13b) excluding the 10 ppt where DMS net accumulation rates were comparable with 2301.1 ± 101.9 pmoles DMS h-1 ml-1 cell susp. and 2192.0 ± 110.0 pmoles MeSH h-1 ml-1 cell susp. The fact that DMS and MeSH production had opposite production patterns in both media (Fig. 13a, b), revealed that salinity had consistent effects on the different pathways of DMSP degradation in both media. Figure 13 - Net DMS and MeSH accumulation rates for the different salinity treatments (10, 20, 30 ppt) amended with 500 μM DMSP in R. pomeroyi DSS-3 cell suspensions under oxic conditions on MBM (a) and ½ YTSS (b) media with 10 mM Glucose addition. a MeSH production was not detected. a) b) a) b) a Chapter 4 39 4.3.3. Side effect of oxic vs. anoxic conditions Once the cell suspensions were submitted to anoxic conditions (purged with N2 for 15 min), DMSP was still converted to DMS and MeSH in both media but accumulation rates of these sulfur compounds were found to be lower (Fig. 14) than in oxic conditions (Fig. 13). In MBM under anoxic conditions, DMS had higher net accumulation rates in the lowest salinity treatment, and a progressive decrease in higher salinities (p < 0.05; Fig. 14a), as observed under oxic conditions. Under anoxic conditions MeSH accumulation rates were higher in the high salinity treatments, as observed in the oxic conditions, but the differences were not statistically significant (p > 0.05; Fig. 14a). Surprisingly, in anoxic ½ YTSS medium, MeSH accumulation among the salinity treatments differed in the sense that MeSH tended to accumulate less in the highest salinity treatment (30 ppt; Fig. 14b). Also, DMS accumulation was detected only in the 20 ppt salinity treatment and only at a low level (8 pmol DMS h-1 ml cell suspensions-1; Fig. 14b). Thus, these results suggested an oxygen influence in the pattern of salinity regulation of DMSP catabolism when cells are grow in ½ YTSS medium. Figure 14 - Net DMS and MeSH accumulation rates for the different salinity treatments (10, 20, 30 ppt) amended with 500 μM DMSP in R. pomeroyi DSS-3 cell suspensions under anoxic conditions on MBM (a) and ½ YTSS (b) media with 10 mM Glucose addition. ª DMS production was not detected. a) b) a a Chapter 4 40 4.3.4. Potential chemical production of DMS and MeSH There was no detectable DMS and MeSH accumulation in parallel experiments incubated without cells indicating the production of sulfur gases was due to the presence of the cells (data not shown). These trials were prepared according to the aforementioned conditions between 10, 20 and 30 ppt salinities and with 50 μM DMSP and 500 μM DMSP amendments, in both MBM and ½ YTSS media. 4.4. Discussion Methylated sulfur compounds such as DMS and, to a much lesser extent, MeSH comprise an important contribution to the sulfur transfer between aquatic environments and atmosphere (Charlson et al. 1987, Aneja & Cooper 1989, Howard et al. 2006). But their production seems to be controlled by diverse marine bacterioplankton taxa mainly through a “bacterial switch” between the competitive DMSP catabolic pathways cleavage and/or demethylation (Jonkers et al. 1998, Simó 2001, Zubkov et al. 2001, Moran et al. 2012). Consequently, the structure of bacterioplankton communities play a key role in controlling the yield of DMS from DMSP, which can range from 5 to ~100% in the ocean water column (Simó et al. 2000, Zubkov et al. 2001, Pinhassi et al. 2005). In this study, we addressed the importance of salinity as a modulating factor on the magnitude of DMS and MeSH accumulation during DMSP catabolism and tested our hypothesis in a simplified biological model system consisting of a pure culture of R. pomeroyi DSS-3. Our results showed that salinity modulated the prevalence of the two different DMSP degradation routes within R. pomeroyi DSS-3 cultures. Overall, our findings suggested an enhanced MeSH accumulation in higher salinities opposed to higher DMS production in the lowest salinity treatments. These results together suggested that variability in salinities may select the preferential route of DMSP degradation pathway within bacterial communities. The pattern of higher DMS accumulations at lower salinities corresponded to what was observed in estuarine sediment slurries, where the highest net DMS accumulation rates were found in freshwater treatments (Magalhães et al. 2012). Magalhães et al. (2012) hypothesized that lower salinities might increase DMSP release from microand macroalgae cells within the sediment and, therefore, enhance DMS production from more Chapter 4 41 available DMSPd degraded by the diverse microbial community. In fact, Yang et al. (2011) confirmed that within axenic cultures of a DMSP producing diatom, Skeletonema costatum, the liberation of extracellular DMSP increased in lower salinities and, consequently, the increase in DMSP availability had a direct correlation with DMS accumulation. While it’s still unknown if diatoms produce DMSP-lyase, DMS production was already attributed to algal DMSP-lyase activity (Yang et al. 2011; Stefels 2000; Steinke et al. 2002). Previous studies have also reported production of DMS in Skeletonema costatum cultures (Vetter & Sharp 1993, Matrai et al. 1995). Furthermore, high DMSP liberation into the extracellular environment, after a phytoplankton bloom, was reported to be followed by DMS peaks related to high bacterial activity (Levasseur et al. 1996). Therefore, there is a strong connection between DMSPd liberation from phytoplankton cells under lower salinity leading to an increase of available DMSP to the DMSP-degrading microbial communities which, according to their composition, may modulate the levels of DMS and MeSH accumulation (Pinhassi et al. 2005). In agreement, other studies with natural communities suggested that DMS production was amplified in lower salinities (Visscher et al. 2003, Niki et al. 2007, Yang et al. 2011). Visscher et al. (2003) verified that DMS flux from a hypersaline microbial mat increased upon low salinities, even though the lowest salinity treatments tested (85 – 95 ppt) where higher than our salinity range. Additionally, Niki et al. (2007) found that low-salinity shock enhanced DMS production as a result of algal DMSPd lyase stimulation. These previous studies converge to the general hypothesis that salinity does influence the accumulation of DMS in water samples and microbial mats slurries. However, they don’t provide a mechanistic explanation for why salinity influenced DMS production. We believe that our present study demonstrated that salinity indeed influences the relative prevalence of the different DMSPd degradation pathways, by benefiting demethylation/demethiolation over the cleavage catabolic pathway with a significant influence on net accumulation of MeSH vs DMS. It is also well established that the dominant process in ocean waters is demethylation/demethiolation most likely due to the relevant incorporation of MeSH into methionine, by bacterioplankton (Kiene et al. 1999, Simó et al. 2000). This was also supported by the high frequency of DMSP-demethylating cells demonstrated to occur in oceanic metagenomic data (Howard et al. 2008). Our experiments confirmed this trend for high MeSH production in R. pomeroyi DSS-3 cell cultures growth in high salinity treatments. The demethiolation of methylmercaptopropionic acid (MMPA; a transient DMSP degradation compound), leads sequentially to MeSH production, and seems to be favored in the highest salinity treatments (30 ppt) which is in agreement with the previous Chapter 4 42 studies that mentioned a bacterial preference (> 50%) for the MeSH-producing pathway in most high salinity environments (Kiene & Linn 2000b, Kiene et al. 2000, Howard et al. 2008). Moreover, Kiene & Linn (2000b) averaged quantitative 35S-DMSPd partitioning flows by using 35S-DMSP tracer and found that in seawater samples, where salinity was 29, MeSH was the dominant sulfur volatile product (~75%) of initial DMSP metabolism. Additionally, this pathway has been proposed to be the dominant one in ocean waters probably due to an acquired ecological advantage in consequence of shifting ecological conditions such as phytoplankton blooms, which can alter DMSP supply, and benefit dmdA community pool (Howard et al. 2011). In our salinity trials we can corroborate that demethylation/demethiolation is enhanced at higher salinities in R. pomeroyi DSS-3 cell suspensions. MeSH is also a potential degradation product of DMS and can be produced through biological methylation of hydrogen sulfide and conversion of methionine in natural environments (Bak et al. 1992, Lomans et al. 1997, Visscher et al. 2003). Nevertheless, González et al. (1999) did not detected MeSH in R. pomeroyi DSS-3 cell suspensions grown in the presence of DMS, which means that in our cell suspensions the biological formation and subsequent detection of MeSH most likely was derived by demethylation/demethiolation of DMSP. Although, higher MeSH accumulations in the ½ YTSS compared with MBM were registered in all salinity treatments which may be originated from degradation of its organic sulfur compounds, as Bürgmann et al. (2007) noticed on their control samples. The yeast extract-tryptone complex in ½ YTSS medium can provide as abovementioned some conversion of methionine leading to an additional MeSH accumulation. Results from this study revealed that high salinities benefit the MeSH net accumulation through the demethylation/demethiolation route which might be also energetically favorable for marine bacteria and, plus, R. pomeroyi demethylase genes may be enzymatically optimized for these osmotic conditions. Furthermore, the formation of MeSH involves more enzymatic conversion steps than the cleavage route (Curson et al. 2011b, Moran et al. 2012). In low salinities, where bacteria have to prevent at the same time the cellular lysis, the demethylation/demethiolation process might be energetically less costly to provide all sulfur and methyl groups available for microorganisms. On the other hand, the low salinities might inhibit the bacterial sulfur demand which might be a probable cause to favor DMS production through the cleavage pathway (Kiene et al. 2000) and, additionally, the DMSP-derived C and S tend to be “lost by diffusion of DMS through the bacterial cell membrane” (Moran et al. 2012). In our measurements, independently of the salinity treatments, DMS accumulations were favored in detriment of MeSH in MBM. In agreement, other studies have measured very Chapter 4 43 low concentrations of MeSH as a DMSP degradation product due to its rapid turnover through continuous demethylations or conversions (Suylen et al. 1987, Bürgmann et al. 2007, Dickshat et al. 2010, Reisch et al. 2011) what might explain the possible underestimation of net MeSH accumulation rates in our results. The free iron in MBM (from FeSO4) could also bind to MeSH and, therefore, reduce the final MeSH concentrations (Butler et al. 1992). Additionally, R. pomeroyi DSS-3 is able to degrade MeSH up to 40% after some hours of incubation (González et al. 1999) and an excess of available DMSPd might lead to a shifting from demethylation to cleavage, once bacterial sulfur demand is fulfilled, which in turn leads to later DMS liberation (Kiene 1996, González et al. 1999, Kiene & Linn 2000b). While we addressed the importance of salinity as a crucial modulating factor on the magnitude of DMS and MeSH fluxes by R. pomeroyi DSS-3 cell suspensions, it is however expected that salinity is not a single acting factor in regulating DMSP catabolism. Actually, when our incubation conditions were changed to anoxic, previously demonstrated to yield DMS and MeSH via enzymatic cleavage or successive demethylations respectively (Kiene & Taylor 1988), the effect of salinity differed for the ½ YTSS medium. While in MBM medium results suggested that the influence of salinity on DMS and MeSH accumulations is oxygen independent, a shift in the trend of MeSH vs. DMS accumulation with salinity was observed in our anaerobic trials performed with ½ YTSS medium. Probably in ½ YTSS there might be other organic compatible solutes than glucose or its derivatives which might enable microorganisms to cope with salinity stress with less effort and, consequently, DMSP might be left out suspending its inherent catabolic processes in anoxic conditions. On the other hand, MeSH can also result from methoxylated aromatic compounds (Bak et al. 1992) or methylation of H2S (Lomans et al. 1997). Actually, Lizotte et al. (2012) suggested some mediation by substrate availability in the microbial transformations of DMSP, promoting the conversion to DMS. Therefore, if there isn’t limitation of C and S supply and available DMSP is in excess for bacterial sulfur demand, a larger fraction of the DMSP could be degraded to DMS (Kiene et al. 1999). Moreover, if DMSP concentrations are too low it will probably be most or entirely assimilated (Hatton et al. 2012). In the majority of our treatments with lower DMSP addition (50 µM), undetectable DMS and MeSH accumulations were registered. Actually, R. pomeroyi DSS-3 strain is known to rapidly degrade DMSP and incorporating most of the sulfur into stable macromolecules, like proteins, required for bacterial growth (González et al. 1999, Kiene et al. 1999, Kiene & Linn 2000a). At the high biomass of cells in the culture samples, it was likely that 50 µM DMSP represented a low “DMSP availability” relative to sulfur demands, thereby leading to low DMS and MeSH liberation. Chapter 4 44 The present results with different salinity conditions and changes in the presence of O2, confirmed that R. pomeroyi DSS-3 strain is flexible in DMSP metabolism when facing unpredictable growth conditions. These findings are consistent with the conclusion that R. pomeroyi DSS-3 possesses several strategies and characteristics for adaptations in marine environments (Moran et al. 2004, Christie-Oleza et al. 2012). As previous studies found an influence of salinity on the magnitudes of release of DMS and MeSH in natural estuarine sediments along a salinity gradient, the present study confirmed that salinity indeed acts as a modulating factor on the bacterial switch between the alternative DMSP degradation pathways. Thus, abiotic factors might influence the proportion of these DMSP degradation products released to the atmosphere. Further information is needed concerning other factors which might be involved on the control of DMSP degradation products and gene expression. This complement will probably lead to a more reliable determination on the magnitude of the salinity effect on DMSP catabolism and on the subsequent enzymatic processes. Chapter 5 45 CHAPTER 5 Conclusions and future research Shifts on abiotic factors, such as salinity, may indeed induce an ecological adaptation in microbial communities through alterations in their chemical and microbiological patterns. This work reflects a brief perception of the effect of salinity on the dynamics of organic sulfur compounds accumulation by intertidal microbial communities and particularly by one representative model organism of Roseobacter (Ruegeria pomeroyi DSS-3), an ubiquitous group in marine environments and key participants in DMSP assimilation. The obtained results from natural microbial communities revealed that salinity changes lead to an opposite pattern on the magnitudes of DMS and MeSH accumulations with a consequent modulation on the inhibitory interaction between MeSH and nitrous oxide reductase enzyme activity in estuarine sediments. While these findings do not allowed us to discriminate between the processes involved in the DMS and MeSH production, R. pomeroyi DSS-3 experiments gave us some insights on the potential effect of salinity on the two DMSP degradation pathways (cleavage and demethylation/demethiolation). Indeed, our results suggested that variability in salinities select the preferential route of DMSP degradation pathway within bacterial communities. In agreement to what was observed in sediment slurries, suspensions of R. pomeroyi DSS-3 cells showed an enhanced MeSH accumulation in higher salinities opposed to higher DMS production in the lowest salinity treatments. These results together provide a mechanistic explanation for how salinity influenced DMS and MeSH production by demonstrating that salinity indeed influences the relative prevalence of the different DMSPd degradation pathways, with a significant influence on net accumulation of DMS vs. MeSH. Finally, this study represents an important contribution to understand the importance of salinity as a key regulator on DMS, MeSH and N2O emissions to the atmosphere, with potential effects on the global climate balance. Nowadays, with progressive genomic methods it is possible to take a next step forward and expand the knowledge about DMSP biogeochemistry. The use of molecular biology and genetic approaches can give access to information concerning other environmental and biological factors which might be involved on the control of DMSP degradation products and gene expression. In addition, the dissection of DMSP transformations and consequent gene identification on both cleavage and demethylation pathways can reveal a useful insight of DMSP metabolism by allowing to understand if the influence on its enzymatic processes occurs at a genomic or proteomic Chapter 5 46 level. Further research must be developed in order to identify the effects of a possible gene inhibition or induction by salinity. 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